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UvA-DARE (Digital Academic Repository) Encounters with oxygen: Aerobic physiology and HO production of Lactobacillus johnsonii Hertzberger, R.Y. Link to publication Citation for published version (APA): Hertzberger, R. Y. (2014). Encounters with oxygen: Aerobic physiology and HO production of Lactobacillus johnsonii General rights It is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), other than for strictly personal, individual use, unless the work is under an open content license (like Creative Commons). Disclaimer/Complaints regulations If you believe that digital publication of certain material infringes any of your rights or (privacy) interests, please let the Library know, stating your reasons. In case of a legitimate complaint, the Library will make the material inaccessible and/or remove it from the website. Please Ask the Library: http://uba.uva.nl/en/contact, or a letter to: Library of the University of Amsterdam, Secretariat, Singel 425, 1012 WP Amsterdam, The Netherlands. You will be contacted as soon as possible. UvA-DARE is a service provided by the library of the University of Amsterdam (http://dare.uva.nl) Download date: 18 Jun 2017 Encounters with oxygen Aerobic physiology and H2O2 production of Lactobacillus johnsonii Uitnodiging Voor het bijwonen van de openbare verdediging van mijn proefschrift Encounters with oxygen Aerobic physiology and H2O2 production of Lactobacillus johnsonii op vrijdag 4 juli 2014 om 13:00 in de Aula van de Universiteit van Amsterdam, Singel 411, Amsterdam Rosanne Y. Hertzberger [email protected] Encounters with oxygen | Rosanne Y. Hertzberger Aerobic physiology and H2O2 production of Lactobacillus johnsonii Rosanne Y. Hertzberger Paranimfen Johan van Beilen [email protected] Sarah Chorus [email protected] Encounters with oxygen Aerobic physiology and H2O2 production of Lactobacillus johnsonii Rosanne Y. Hertzberger 2014 The work presented in this thesis was carried out in the frame of a PhD-project of NIZO Food Research, funded by Nestlé Research Centre and carried out at the Molecular Microbial Physiology lab, Swammerdam Institute for Life Sciences, University of Amsterdam. The project was part of the Kluyver entre for Genomics of Industrial Fermentation. Chapter 2 and 4 of this thesis have been published: Hertzberger RY, Arents J, Dekker HL, Pridmore RD, Gysler C, Kleerebezem M, Teixeira de Mattos MJ. 2014. H2O2 production in species of the Lactobacillus acidophilus group, a central role for a novel NADH dependent flavin reductase. Appl. Environ. Microbiol. Hertzberger RY, Pridmore RD, Gysler C, Kleerebezem M, Teixeira de Mattos MJ. 2013. Oxygen relieves the CO2 and acetate dependency of Lactobacillus johnsonii NCC 533. PLoS One. 8:e57235. Cover Lactobacillus johnsonii colonies after 9 hours of growth in a 20% O2 and 5% CO2 atmosphere on AnoporeTM inorganic membranes on MRS agar. The colonies were stained with SYTO9 and propidium iodide and photographed through a green filter (showing only live/ SYTO9 stained cells) with a cooled charge-coupled device camera mounted on an Olympus BX-60 fluorescence microscope. Cover design MidasMentink.nl and layout Printing Ipskamp drukkers Encounters with oxygen Aerobic physiology and H2O2 production of Lactobacillus johnsonii academisch proefschrift ter verkrijging van de graad van doctor aan de Universiteit van Amsterdam op gezag van de Rector Magnificus prof. dr. D.C. van den Boom ten overstaan van een door het college voor promoties ingestelde commissie, in het openbaar te verdedigen in de Aula der Universiteit op 4 Juli 2014, te 13:00 uur door Rosanne Yente Hertzberger geboren te Rotterdam Promotiecommissie Promotores Prof. dr. M.J. Teixeira de Mattos Prof. dr. M. Kleerebezem Overige Leden Prof. dr. J.T. Pronk Prof. dr. O.P. Kuipers Prof. dr. J. Hugenholtz Prof. dr. R. Kort Dr. F. Branco dos Santos Dr. E.J. Smid Faculteit der Natuurwetenschappen, Wiskunde en Informatica “Science is the only news; alle else is hearsay and gossip.” Lynn Margulis, condensed from Stewart Brand - Mind, Life and Universe Table of contents Chapter 1: General introduction. Living on the edge: Reactive Oxygen 10 Species production and scavenging in Lactic Acid Bacteria. Chapter 2: Hydrogen peroxide production in species of the Lactobacillus 38 acidophilus group, a central role for a novel NADH dependent flavin reductase. Chapter 3: Transcriptome response in Lactobacillus johnsonii identifies 68 an oxygen induced NADH oxidase that contributes to H2O2 production. Chapter 4: Oxygen relieves the CO2 and acetate dependency of 92 Lactobacillus johnsonii NCC 533 Chapter 5: Genome-wide transcriptome response to CO2 depletion in 114 Lactobacillus johnsonii Chapter 6: General discussion and outlook. 138 References 151 Summary (for a scientific audience) 170 Samenvatting (voor breed publiek) 174 List of Abbreviations 177 Dankwoord 178 About the author 180 Chapter 1 General introduction Living on the edge: Reactive Oxygen Species production and scavenging in Lactic Acid Bacteria. Abstract Lactic acid bacteria (LAB) are generally classified as aerotolerant anaerobes. They grow relatively well in the presence of oxygen but cannot use oxygen as a terminal electron acceptor due to an incomplete electron transfer chain. Oxidases, such as cytochrome oxidase, NADH oxidase, lactate oxidase and pyruvate oxidase play a central role in the aerobic lifestyle of lactic acid bacteria, either protecting against oxidative stress (cytochrome oxidase, H2O-forming NADH oxidase) or aggravating oxidative stress (pyruvate oxidase, lactate oxidase, H2O2-forming NADH oxidase). LAB employ several ROS-scavenging enzymes and molecules to provide resistance against oxidative stress. Here we give an overview of the literature that describes the role of oxidases and ROSscavenging enzymes in aerobic metabolism and oxidative stress resistance of LAB. 10 Introduction “There’s something new here—in the middle of the drop they are lively, going every which way.” Gently, precisely, a little aimlessly, he moved the specimen so that the edge of the drop was under his lens... “But here at 1 the edge they’re not moving, they’re lying round stiff as pokers.” It was so with every specimen he looked at. “Air kills them,” he cried, and was sure he had made a great discovery. – Paul de Kruijf, Microbe Hunters, 1926 – This is Paul de Kruif’s interpretation of what happened when Louis Pasteur first peered through his microscope at a specimen of rancid butter in 1860 (1). Previously, Antonie van Leeuwenhoek in the 17th century and Lazarro Spalanzani in the 18th century had already noted that some of these swarming animalcules live happily in the absence of air. Now Pasteur could add a new category to the list of lifestyles in the microbe world that were dictated by oxygen: air could also be toxic (2). By now, we know that oxygen is not only toxic to these butyric acid producing clostridia that Pasteur was looking at, but that all forms of life that encounter oxygen need some strategy to deal with its hazardous byproducts. The risk is embedded in the chemical makeup of the oxygen molecule. The ground state of the oxygen molecule is the triplet state, which means that the single electron transfer to oxygen is thermodynamically highly favorable, but quantum-mechanically (spin) forbidden. As a consequence it is a relatively inert gas: the building blocks of the cell -lipids, amino acids, nucleotidesare only weak univalent electron donors so their autoxidation is a sluggish reaction. However, oxygen rapidly oxidizes other cellular compounds that are strong univalent electron donors, such as flavins and quinones (3). The products that are formed by these reaction are much stronger univalent electron acceptors than triplet oxygen and are known as reactive oxygen species (ROS, Figure 1.1). They can cause extensive damage through different pathways. Metal centers are especially vulnerable to superoxide. The enzymes that commonly rely on catalytically active [4Fe-4S]-clusters, such as fumarase and aconitase, are quickly inactivated by superoxide (4, 5). The release of free iron metals from damaged centers accelerates further ROS formation: hydrogen peroxide can interact with free ferric ions producing the highly reactive hydroxyl radical. If this so-called Fenton-reaction occurs in the proximity of DNA, it may cause DNA lesions and subsequent cell death. An excellent review on the specific damage that ROS may cause was written by James Imlay (6). 11 flavins, quinones NOX, LOX, POX, flavins COX, NOX was written by James Imlay (6). ·O2-‐ e-‐ H2O2 SOD O2 e-‐ e-‐ ·OH e-‐ Fenton reac@on H2O catalase, peroxidases Figure 1.1; Generation and scavenging of reactive oxygen species. NOX: NADH oxidase, POX: pyruvate oxidase, LOX: lactate oxidase, COX: cytochrome oxidase, SOD: superoxide dismutase. Adjusted from (6) It is therefore not surprising that, on a planet with an atmosphere that consists for more than 20% of oxygen, there is an urgent need for many organisms to invest in mechanisms that eradicate reactive oxygen species from the cells (7). Even species that are dependent on a continuous supply of oxygen for aerobic respiration, require enzymes that continuously scavenge the toxic byproducts. The anti-oxidative arsenal consists of a variety of proteins specialized at this task, including catalase, and superoxide dismutase (SOD) that react with hydrogen peroxide and superoxide radicals, respectively (8). The discoverers of superoxide dismutase, McCord and Fridovich (9, 10) hypothesized that ROS scavengers were the facilitators of an aerobic lifestyle, since their occurrence seemed to correlate with the aerotolerance of bacteria (11). Since then, it was shown that several bacteria that express these proteins indeed rely on their presence for aerotolerance (7, 12, 13). Yet, the physiology of other bacteria indicate that the proposed rule does not apply universally: obligate anaerobes such as the butyric acid producing clostridia that Pasteur had observed, express a variety of ROS-scavengers, including a functional superoxide dismutase (14, 15). Moreover – as McCord and Fridovich already observed for Lactobacillus plantarum - several lactic acid bacteria are aerotolerant but are devoid of a functional SOD or catalase. Such bacterial species challenge our understanding as to how micro-organisms may deal with the consequences of a life in the presence of oxygen. 12 1. Lactic acid bacteria LAB are a phylogenetically diverse group of Gram-positive, non-sporulating, rod- or coccoïd shaped bacteria that have a fermentative metabolism with lactic acid as a major end product. The intimate relationship between LAB and humans is related to 1 the prominent role of these bacteria in food fermentations. They contribute to the taste, texture and shelf life of a wide variety of fermented food products including yoghurt, cheese, butter milk, sour cream, pickles, olives, sauerkraut, sourdough and meat. Besides their presence in these food products, LAB are associated with the mucosal surfaces of the human body. They are amongst the very first microbes that colonize newborns: the lactobacilli that inhabit the vaginal cavity of the mother are microbial pioneers in the neonatal gut (16). Further bacterial transmission from the mother to the infant during breastfeeding also mostly consists of LAB (17, 18). LAB, and in particular streptococci, constitute a considerable part of the upper oro-gastrointestinal tract microbiota (19, 20). In the gastro-intestinal tract, specific LAB species and strains have been proposed to exert a positive effect on the health of the host. Administration of adequate amounts of lactobacilli and bifidobacteria in products designated as probiotics can reduce the severity of antibiotic-associated diarrhea (21, 22), infectious diarrhea (23) and Clostridium difficile –related diarrhea (24). Probiotics can be effective in the prophylactic treatment or necrotizing enterocolitis in preterm infants (25) and they have been proposed to be effective in reducing the risk of atopic dermatitis development in children (26). Several characteristics of LAB were identified that may contribute to these probiotic effects. Firstly, probiotic LAB generally display substantial acid and bile resistance that enables them to survive the hostile environment of the stomach and the upper digestive tract (27). Secondly, they express proteins that allow competitively adherence to intestinal mucus (28), and thirdly, they can strengthen the mucosal barrier and/or immune function (29, 30). 2. LAB and oxygen In terms of their classification with respect to oxygen, LAB are generally referred to as aerotolerant anaerobes. This classification reflects two commonly conserved characteristics in LAB: (i) due to the incompleteness of their electron transport chain, they are unable to use oxygen as a terminal electron acceptor (hence the term “anaerobes”), while (ii) oxygen is also not particularly damaging either, which is 13 illustrated by the relatively good growth of many LAB in aerobic environments (hence the term “aerotolerant”). This tolerance does not mean that LAB are unresponsive to environmental oxygen levels. Oxygen can have a profound effect on growth, metabolism and viability. The presence of oxygen can dictate which pathways are used for ATP generation and which fermentation metabolites are produced (31, 32). The enzymes involved in pyruvate dissipation are particularly influenced by exposure to oxygen (see Figure 1.2). For example, pyruvate formate lyase (PFL) is highly sensitive to oxygen (33) and as a consequence, formate production is completely abolished upon oxygen exposure. The pyruvate oxidase (POX) pathway depends on molecular oxygen as a substrate (34) and facilitates acetate production with additional generation of ATP from central metabolism. Furthermore, oxygen allows the utilization of lactate through the lactate oxidase (LOX) pathway and the regeneration of oxidized reducing equivalents catalyzed by NADH oxidase (NOX) (Figure 1.2) glucose ADP NAD+ ATP NADH pyruvate pfl formate NAD+ pdh NADH acetyl-‐CoA . NADH NAD+ ldh lactate lox H2O2 O2 O 2 pox O2 +C 2 H 2O pat NADH adh NAD+ acetyl-‐P ack ADP ATP ethanol acetate O2 H2O2 / H2O NADH nox NAD+ Figure 1.2; Overview of pyruvate metabolism in LAB (for simplicity the pathways for acetoin/2,3-butanediol were omitted). LDH: lactate dehydrogenase, LOX: Lactate oxidase POX: pyruvate oxidase, PFL: pyruvate formate lyase, PDH: pyruvate dehydrogenase, PAT: phosphate acetyltransferase, ACK: acetate kinase, NOX: NADH oxidase) Besides the additional metabolic reorientation that oxygen may elicit, the generation of hydrogen peroxide (H2O2) and superoxide species in aerobic metabolism, for instance by NOX or POX activity, is a major source of intrinsic oxidative stress in LAB. Especially 14 in obligate homofermentative lactobacilli H2O2 can freely accumulate and may cause growth stagnation and cell death (35-38). At the same time, oxygen can also provide a benefit in terms of additional ATP generation through the POX-pathway, involving activity of POX and acetate kinase (ACK, see Figure 1.2). Accordingly, the degree of 1 aerotolerance encountered in individual LAB species and strains shows considerable variation. Some species may lean more towards “aerotolerant”, such as species in the Leuconostoc genus that can reach approximately 2-fold higher cell densities in aerobic environments as compared to anaerobic environments (39). Other species may lean more towards “anaerobes”, such as Lactobacillus delbrueckii subsp bulgaricus that reaches approximately 2-fold reduced cell densities in aerobic conditions (37). Results presented in this thesis demonstrate that Lactobacillus johnsonii also belongs to this latter group. In the environments where LAB are regularly found, they will frequently encounter molecular oxygen and its detrimental derivatives. The epithelial and mucosal linings of the mouth, vagina and intestine are considered to contain considerable levels of oxygen (40-45). Similarly, LAB used in the preparation of fermented food products are exposed to oxygen during processing, shelf life and consumption (35, 46, 47). This regular oxygen exposure has a considerable impact on their overall metabolism and physiology (48), on their interactions with other bacteria (49) and on their interactions with the host organism (40, 50). In this literature review we explore the aerobic physiology of LAB and in particular their aerotolerance. We consider two aspects that are central Definitions used in this thesis: for the ability of cells to retain ROS Hydrogen peroxide, superoxide, hydroxyl radical. viable and to proliferate in the For simplicity, we restrain ourselves to these three presence of oxygen. Firstly, we compounds (Figure 1.1). will discuss aerobic metabolism Oxidative stress Cellular damage (or the risk of cellular in LAB, which is presumably damage) caused by ROS. mediated by the activities of Oxidative stress resistance: The ability to grow four cytochrome and/or retain viability in the presence of ROS-generating oxidase, POX, LOX and NOX. compounds (e.g. paraquat and plumbagin) or hydrogen Secondly, peroxide. oxidases: we will discuss expression of ROS-scavenging Aerotolerance: The ability to grow and/or retain enzymes and other common viability in the presence of oxygen. anti-oxidative strategies that 15 confer resistance against oxidative stress to LAB. Taken together, these aspects provide insight into the different consequences of oxygen exposure for different LAB species. 3. Cytochrome oxidase and respiration A conserved characteristic amongst LAB is their inability to constitute a functional electron transfer chain that uses molecular oxygen as a terminal electron acceptor. Nevertheless, the requirements for a rudimentary electron transfer chain are present in several LAB, such as NADH dehydrogenase that accepts electrons from reducing equivalents (51) and cytochrome oxidase to catalyze the final electron transfer to oxygen (52). However, with a few exceptions (53, 54) most LAB are unable to synthesize quinones, that are essential for electron shuttling between NADH dehydrogenase and cytochrome oxidase. Furthermore, all LAB lack the essential genes to constitute a complete synthesis pathway for heme (55), an essential and functional prosthetic group of cytochromes and cytochrome oxidases. Therefore, LAB cannot assemble cytochromes and cannot produce functional cytochrome oxidases (52). In several species a respiratory phenotype can be induced by supplementation of hemin, indicated by increased biomass levels, cytochrome synthesis, altered metabolite levels and proton pumping (56-58). Several studies on the respiratory phenotype report a dramatic increase in survival in respiration-permissive environments. The viability loss that is witnessed in certain LAB during aerobic stationary phase can be prevented by the addition of hemin, as was shown in L. lactis (59, 60), L. plantarum and S. agalactiae (57). Moreover, L. plantarum displayed a considerably higher H2O2-tolerance in the presence of hemin (61). Besides the production of additional ATP, hemin addition may also contribute to aerobic robustness through reducing ROS production. The respiratory chain is an important source of reactive oxygen species. Reduced quinone species can spontaneously react with oxygen, resulting in superoxide and H2O2 formation. Enterococcus faecalis is an exception amongst LAB since it is can synthesize demethylmenaquinones. Univalent oxidation of these quinones was shown to be the cause of extracellular superoxide production (62, 63). Activity of cytochrome oxidase through the addition of hemin abolished this superoxide generation. Such an effect of cytochrome oxidase activity on ROS-production was previously observed in Escherichia coli where a flux through the electron transfer chain (by restoring the function of a cytochrome oxidase) could reduce H2O2 production derived from autoxidation of the flavin cofactor of fumarate reductase (64). In L. lactis, aerobic respiration also abolished ROS-production (53), although here 16 the major source of ROS was found to be a H2O forming NADH oxidase (48). We consider however that lowering the NADH/NAD+ balance in L. lactis by shuttling electrons from NADH in an electron transfer chain, would also lower the flux through the H2O forming NADH oxidase and would therefore contribute to lowering superoxide-formation. 1 These studies show that in several LAB, aerobic respiration can increase yield and robustness, and oxidative stress tolerance. Oxidative stress is reduced by diminshing autoxidations of respiratory intermediates and by generating additional ATP. In addition menaquinone and/or hemin has an impact on a variety of physiological aspects, including metabolite profile, pH and redox-state. All these aspects may play a role in the increased oxidative stress tolerance. 4. NADH oxidase One of the consequences of lacking an intact electron transfer chain is that additional redox constraints arise. Glucose to lactate fermentation is redox-neutral: the NADH that is produced through the glycolytic production of pyruvate, is regenerated by the lactate dehydrogenase (LDH) reaction in an equimolar stoichiometry. Pyruvate conversion to acetic acid (through one of the three pathways: PDH, PFL, POX) requires the oxidation of NADH through a different pathway, either by the production of reduced metabolites such as ethanol or acetoin or through the use of molecular oxygen to regenerate NAD+ for example through the activity of NOX (Figure 1.2; (65-67). A second complication in the correct identification of NOX is that other flavoproteins that do not belong to the NOXs may catalyze NADH oxidation as a side activity. Fumarate reductase of Escherichia coli (64, 68) and dihydroorotate dehydrogenase of B. bifidum (69) are examples of proteins with a solvent-exposed flavin moiety that easily autoxidizes, leading to superoxide or H2O2 production. Furthermore, free flavins that are reduced by NADH flavin reductases may spontaneously react with molecular oxygen producing considerable amounts of ROS (6). Reduced flavins can provide an important source of H2O2 in lactobacilli, which is illustrated by the identification of a flavin reductase as the main contributor to H2O2 production by L. johnsonii (chapter 2 of this thesis). Two different NOX enzymes are expressed by LAB: one that catalyzes a four electron transfer producing mainly water and one that catalyzes a two electron transfer producing mainly H2O2. It is not trivial to distinguish between these two forms since LAB may also express an NADH peroxidase (NPR), which in combination with the activity of H2O2-producing NOX results in the same overall reaction as the one catalyzed by the 17 H2O-producing NOX (32, 70) (Table 1.1). In some cases, such as in the eukaryotic DUOX enzyme (71) and bacterial alkyl hydroperoxide reductase (72), the two reactions could even be attributed to a single enzyme. DUOX H2O2 producing and scavenging (NOX and NPR activity) can even be attributed to a single enzyme. Nox-1, a 55 kDa protein requiring exogenous flavin for activity, in Streptococcus mutans displayed a high level of homology with the earlier identified ahpF gene in Salmonella typhimurium (73) that is part of an alkylhydroperoxide reductase (AHPR, encoded by ahpC and ahpF, see also below at 6b). This Nox-1 catalyzes H2O2 producing NADH oxidation but together with the upstream located ahpC gene, it forms an AHPR, which can reduce H2O2 as well as alkyl organic peroxides using NADH (74, 75). A similar feature and DUOX enzymes in eukaryotes (71) Table 1.1; Enzymatic reactions involving reactive oxygen species generally found in LAB. Cytochrome bd oxidase (Q = menaquinone or ubiquinone) Pyruvate oxidase (cofactors, TPP, Mg2+, FAD) Lactate oxidase NADH oxidase (H2O2-forming) NADH oxidase (water-forming) NADH flavin reductase (uses either FAD, FMN or riboflavin as substrate) Catalase (with either hemin or manganese in the active site) Thiol-based peroxidase. R could be substituted for TRX, GT or AHP. 18 NADH peroxidase 1 Superoxide dismutase Fenton reaction Streptococcus mutans also encodes a 50 kDa enzyme Nox-2 that catalyzes the transfer of four electrons to oxygen, producing water. This Nox-2 appears to be essential for regenerating NAD+, since a nox-2 deletion derivative of S. mutans displays hampered growth on mannitol, which is a substrate that generates additional NADH when it is catabolized to pyruvate. Moreover, this deletion mutant also redirected its carbon metabolism towards a higher production of lactate, which is more reduced than the acetate and CO2 combined (76). In other LAB, these type of metabolic effects of NOX are also observed. In L. sanfranciscensis (65, 77) and in L. lactis (66, 77), NOX was shown to exert direct control on the end-product of fermentation. L. lactis shifted from homolactic to mixed acid fermentation upon controlled overexpression of a H2Oforming NOX when growing in the presence of oxygen (66, 74, 77). These examples illustrate that redox balance (NADH/NAD+ ratios) plays a pivotal role in the control of the pyruvate dissipating flux. Lactic acid bacteria may be forced towards lactate formation to sustain redox balance under anaerobic conditions, but can shift towards mixed acid fermentation under aerobic conditions through the alternative electron sink provided by the NOX enzyme activity, with the concomitant benefit of the additional ATP generated through the acetate formation pathway. This metabolic control of redox balance has been exploited in LAB, and in particular in L. lactis to redirect its metabolic flux towards higher production of flavour components like acetoin, diacetyl and acetaldehyde (66, 78) 19 5. Aerobic lactate utilization: pyruvate and lactate oxidase POX plays a prominent role in the metabolism of several well-studied LAB. It catalyzes the oxidative decarboxylation of pyruvate in the presence of inorganic phosphate, releasing acetyl-phosphate, carbon dioxide and H2O2. The enzyme is a homotetramere that contains tightly bound flavin adenine nucleotide (FAD) and uses thiamine pyrophosphate (TPP) and a divalent metal ion as cofactors (79, 80). POX is part of the aerobic lactate utilization pathway. Upon glucose exhaustion in the environment as a consequence of aerobic growth of L. plantarum of L. lactis, the accumulated lactate can be oxidized by the combined activity of an NAD-dependent LDH and POX (81, 82). The resulting acetyl-phosphate can support ATP production through acetate kinase. Deletion of two (poxB and poxF) of this apparently five-fold redundant function in L. plantarum WCFS1 completely abolished aerobic lactate utilization and acetate production in stationary phase, implying that POX (and not PFL or PDH) is the main acetate-producing reaction in this species under these condions (83). A similar aerobic lactate utilization pathway is found in S. pneumoniae, although in this organism the first step is catalyzed by LOX instead of LDH. S. pneumoniae (then referred to as Pneumococcus) Expression of the lox (LOX) and pox (POX) genes in several LAB is repressed in the presence of glucose through CCP-A mediated catabolite control (84-87). This does not appear to be the case in S. pneumoniae where the transcription of these genes (88) nor the reaction they catalyze (lactate utilization, acetate and H2O2 production) were influenced by environmental glucose levels (89). Combined LOX and POX activity results in the accumulation of up to 1 mM of H2O2 during the stationary phase of growth in S. pneumoniae (90) and S. pyogenes (86). Notably, deletion of either the pox or lox gene completely abolished H2O2 production and prevented the dramatic viability loss during the aerobic stationary phase of growth in several LAB (83, 86, 87, 89-91). An exception with respect to the role of POX and LOX in aerobic metabolism is encountered in the species belonging to the L. acidophilus group. The pox-gene is present in these species and an active protein could be purified from L. delbrueckii subsp bulgaricus (34). Furthermore, as we show in this thesis, L. johnsonii can produce sufficient acetate and CO2 to satisfy its growth requirements, in a POX and oxygen dependent manner (chapter 4). However, L. johnsonii and delbrueckii subsp bulgaricus are strictly homofermentative and thus do not produce acetic acid under aerobic conditions, which allows only a limited flux through the POX-pathway. 20 6. ROS scavenging enzymes Many of the above-mentioned enzymes, such as POX, LOX, NFR and NOX, catalyze the transfer of two electrons to oxygen, resulting in H2O2 formation. Consequently, aerobic growth of some LAB is accompanied by the accumulation of substantial amounts (>1 1 mM) of H2O2. In streptococci this H2O2 is predominantly produced by POX and LOX (85, 86, 89, 92). In other bacteria, such as the species belonging to the L. acidophilusgroup, H2O2 is produced in a reaction involving NADH (37, 93, 94). In this thesis we report the identification of the enzyme involved in H2O2 production in L. johnsonii as an NFR and identified a NOX enzyme to contribute to H2O2 production after prolonged aeration (see chapter 2 and 3). Besides the direct involvement of these enzymes, H2O2 may also be generated in spontaneous oxidations of cellular components. Superoxide radicals that are generated during autoxidation of dimethylmenaquinone in E. faecalis (62) and NADH oxidase activity of L. lactis (48) can spontaneously be dismuted to form H2O2. Even in aerobic, respiring bacteria such as E. coli, the deletion of the main H2O2 scavenging enzymes catalase and AHPR leads to H2O2 accumulation, indicating that this is a universal characteristic of bacterial growth in oxygenated environments. (64, 95). Besides the ability to use oxygen for ATP generation, e.g. in aerobic respiration or aerobic lactate utilization, the ability to scavenge the toxic byproducts of oxygen reactions (ROS) is an important factor which contributes to aerotolerance of cells. LAB employ a diverse range of mechanisms to protect against the oxygen radicals that are generated during aerobic metabolism. Below we discuss the enzymes that were shown to contribute to aerotolerance and oxidative stress resistance in LAB by scavenging ROS. An overview of the distribution of genes encoding these ROS-scavenging enzymes in LAB is presented in Table 1.2 and S1.1). 21 22 1 Hemin-dependent catalase (HemCat) from E. faecalis V583 (96) 2 Mn-catalase (MnCat), L. plantarum ATCC 14431 (188) 3 Thioredoxin reductase (TRXR) L. plantarum WCFS1 (113). 4 Thiol peroxidase (TPX) from S. pneumoniae D39 (116) 5 Glutathione reductase (GSHR) L. sanfranciscensis DSM20451 (122) 6 Alkyl hydroperoxide reductase (AHPR), S. mutans NCIB 11723 (74) 7 NADH peroxidase (NPR) from Enterococcus faecalis V583 (117). 8 Manganese-superoxide dismutase (MnSOD) L. lactis MG1363 (48) - - 1 - E. coli K-12 - S. mutans UA159 - - - S. thermophilus LMG 18311 - - - - S. pneumoniae D39 - - S. pyogenes HSC5 - - 1 E. faecalis V583 - - - MnCat2 (accession nr. P60355) - L. sanfranciscensis TMW 1.1304 L. delbrueckii subsp bulgaricus ATCC 11842 L. johnsonii NCC 533 - L. rhamnosus GG - 1 L. casei BL23 L. plantarum WCFS1 L. lactis subsp. cremoris MG1363 HemCat1 ef_1597 2 2 2 2 5 3 3 4 3 2 2 3 2 TrxR3 lp_0761 1 1 - 1 1 1 1 - 1 1 1 1 1 Tpx4 SPD_1464 4 3 1 1 4 3 3 2 3 4 3 4 GshR5 (accession nr. A1YAC0) 5 1 - - - 1 - - 1 1 1 1 1 AhpC6 (accession nr. O66265) - 1 1 1 1 2 2 3 2 5 4 5 5 10 NPR7, ef_1211 2 - - - 1 1 1 1 1 1 - 1 - MnSOD8 llmg_0429 Table 1.2; Distribution of genes encoding antioxidative/ROS-scavenging mechanisms in LAB genomes as found by BlastP (cut-off values: minimal query coverage 60%, maximal e-value 10-10, minimal identitical residues: 20%). The number of genes that fall within the selection criteria are between brackets. For gene annotations and ID see Supplementary table S1.1. a. Catalase LAB are generally referred to as being catalase-negative and indeed, catalase encoding genes are sparsely encountered in LAB genomes. Nevertheless, two different types of catalases are found in LAB. A hemin-dependent catalase is present in the genomes of 1 certain strains of L. plantarum, L. sakei, L. casei and E. faecalis (96, 97). This gene can produce a functional H2O2 scavenging enzyme when cells reside in hemin-containing environments, thus providing H2O2 tolerance under specific conditions (98-100). An alternative catalase that is not depending on hemin supplementation was identified in L. plantarum strain ATCC 14431 (101, 102) and appeared to employ manganese ions in its reactive center (103). The gene encoding MnCAT (accession nr. P60355) is very rare in LAB, and only encountered in a few Pediococcus, Enterococcus and Lactobacillus strains. A L. plantarum strain in which this MnCAT was absent produced H2O2 during aerobic growth indicating that this enzyme is the main H2O2 scavenging activity for these species in hemin-depleted conditions (100). b. Thiol-based peroxidase Whereas catalases are quite scarce in LAB, genes for a second type of H2O2-scavenging enzyme, with an activity that revolves around a catalytically active cysteine residue, are abundantly present on LAB genomes. The nomenclature in literature is quite diverse and elsewhere this group may be referred to as peroxiredoxins, cysteine-based peroxidases, thiol-specific antioxidants or NPRs. Here we will refer to them as thiol-based peroxidases, following Mishra et al (7). We discuss them as a group, since they show significant similarity, both in gene sequence and in reaction mechanism (104). Thiol-based peroxidases catalyze the reduction of H2O2 to water, or organic hydroperoxides to their corresponding alcohol, using NADH or NADPH as the electron donor. Different types of thiol-based peroxidases have been detected in LAB: glutathione /glutathione reductase (GSH/GSHR), thioredoxin /thioredoxin reductase (TRX/TRXR) and alkyl hydroperoxide /alkyl hydroperoxide reductase (AHP/AHPR encoded by ahpC and ahpF, respectively). The reaction mechanism consist of two steps: the smaller polypeptide (either TRX, GSH or AHP) has an active site with two (seleno-)cysteine residues, or in the case of GSH two molecules that each contain a single cysteine residue. One of the cysteine residues reacts with H2O2, leading to the formation of a sulfenic acid that subsequently reacts with the secondary cysteine residue forming a disulfide bond. A dedicated flavoprotein reductase (either TRXR, GSHR or AHPR) can subsequently reduce the disulfide bond. The active site of these reductases also contains cysteine 23 residues and employs NADH or NADPH as an electron donor (7, 105, 106). In the case of thiol peroxidase (TPX), the disulfide bond is reduced by a TRX, which itself is reduced by TRXR. Besides their role in oxidative stress tolerance, these peroxidases protect against a wide variety of environmental stresses that are associated with the formation of reactive oxygen species, such as temperature and low-pH stress (107, 108). This type of NADH related peroxidase activity was shown to be complementary to catalase activity in E. coli (109). However, an important difference is that thiol-based peroxidases require input of electrons from reducing equivalents, whereas catalase does not need a co-substrate besides H2O2. Moreover, thiol-based peroxidases are active at lower ROS concentrations as compared to catalase (7, 74). Below, we discuss the prevalence of four types of thiol-based peroxidases in LAB, (i) thioredoxin/thiol peroxidase, (ii) glutathione reductase, (iii) alkyl hydroperoxide reductase and (iv) NADH peroxidase (Table 1.2 and Table S1.1). Thioredoxin reductase / Thiol peroxidase The genes encoding TRX (trxA) and its corresponding TRXR (trxB) are ubiquitously present in LAB (Table 1.2). TRX is a short polypeptide (103-106 residues) with a conserved cysteine rich catalytic site (CXXC). It is present in all forms of life and in several species it is indispensable. In bacteria, it has been associated with a variety of processes, including gene regulation and signal transduction. TRX can also function as an electron acceptor or hydrogen donor in redox reactions and can contribute to ROS scavenging (110-112). The catalytic cysteine residues of TRX react either directly with H2O2 or indirectly with an oxidized thiol from a so-called peroxiredoxin such as TPX. Oxidized TRX can be reduced by a dedicated TRXR, which has a corresponding CXXC conserved catalytic site. LAB generally have multiple copies of both the TRX encoding trxA gene and the TRXR encoding trxB gene. TRXR is a member of a larger family of pyridine nucleotide-disulfide oxidoreductases that also include GSHR, AHPR and DLD (catalyzing the second reaction in the pyruvate dehydrogenase complex). There are similarities between the members of this group, which complicates correct annotation. This is possibly the reason why not all copies of genes annotated as TRXRs have the CXXC-motif which is deemed essential for its activity (113) (Table S1.1). In several LAB, the TRX/TRXR systems were shown to contribute to oxidative stress resistance and aerotolerance. Mutation analysis of the TRXR encoding trxB gene in L. casei subsp. shirota, in L. plantarum and in L. lactis showed that the TRX/TRXR system 24 is essential for tolerance against disulfide (diamide)- and H2O2-stress (113-115). TRX expression was associated with the induction and repression of a variety of stress proteins in L. plantarum (113) and L. lactis (115). The L. lactis trxB1 gene that contains the canonical CXXC-motif and is predicted to encode a TRXR was not reported to play 1 a role in aerotolerance (115). Nevertheless, the L. lactis genome contains a second TRX homolog, annotated as an AHP (33% identity) and contains the CXXC motif, which may show TRXR activity when the primary TRXR was disrupted in L. lactis. Notably, a TPX that functions in conjunction with the TRX system has been shown to display NADPH peroxidase activity in S. pneumoniae (116), and to contribute to oxidative stress tolerance in E. faecalis (117). Thereby the TRX, TPX system acts as a direct NPR that is involved in aerotolerance. Moreover, this system has also been shown to play a role in gene regulation related to the oxidative stress response in S. pneumoniae (116). Glutathione reductase Almost all LAB lack the enzymatic machinery for GSH synthesis: the GSH synthetase encoding gene from E. coli has no full-length homologs in the Streptococcus, Lactobacillus and Lactococcus genera. However, many LAB can import exogenous GSH and maintain high intracellular levels (108, 118). Furthermore, many species encode the corresponding GSHR, to regenerate reduced GSH. In L. lactis intracellular GSH was estimated to increase to 60 mM when GSH-rich ingredients such as yeast extract were present in the medium. GSH supplementation was correlated with H2O2 tolerance (119) and provided cross-protection against acid stress (120). Similarly, in several LAB the uptake and recycling of GSH is correlated with oxidative stress resistance (118, 121123). Alkyl hydroperoxide reductase In E. coli, the AHPR was observed to be the “first line of defense” against H2O2. The system, encoded by ahpC and ahpF, scavenges H2O2 with high affinity and low-level saturation (Km of 5 µM). It can fully complement catalase deficiency in aerobically growing E. coli (109). As mentioned before, AHPR belongs to the same protein family as TRXR, GSHR and DLD and the corresponding genes are ubiquitously present on LAB genomes. However, copies of the corresponding AHP-encoding ahpC gene are not as prevalent in LAB, indicating that this system is only functional in a subset of LAB (Table 1.2). 25 AHPR was first identified in S. mutans as a H2O2-producing NOX (76, 124). However, the direct reaction with oxygen appeared to be a side-activity. In vivo its main electron acceptor was identified as AHP, whose cysteine residues can be oxidized by H2O2, indicating its function in ROS scavenging (74, 76, 105). Such a role was established in E. faecalis, where deletion derivatives of ahpCF displayed reduced growth and viability in the presence of exogenous H2O2 compared to the wild type (117). However, this role of AHPR is apparently not universally valid, since ahpCF deletion derivatives of S. mutans were not affected in terms of their sensitivity towards H2O2 as compared to the wild type strains (125). Besides these apparently contradictory studies, the role of AHPR in ROS tolerance in LAB has not been studied extensively, and its in vivo function remains to be further elucidated. NADH peroxidase NPR is unique in the class of thiol-based peroxidases since both the reaction between thiol and H2O2 as well as the NADH recycling of the oxidized thiol is catalyzed by a single enzyme. The NPR of E. faecalis, encoded by the npr-gene, is a flavoenzyme with a single active cysteine residue Cys42 (126, 127) that is essential for oxidative stress resistance in this bacterium (117). The NPR shows a remarkable redundancy in LAB. L. plantarum even has 10 genes that show resemblance with high significance (BlastP cut-off values: maximum e-value 10-10, minimum identical residues 20%, minimum query coverage 60%). 6 of these 10 genes are annotated as NOXs and 5 have the conserved active site cysteine residue. One of these NPR homologs (noxV, accession number F9UUC2), displayed H2O-forming NOX activity but its capacity to convert H2O2 was not investigated (128). The high-level of sequence homology between NOX and NPR may suggest that this group of flavoproteins could display substrate promiscuity and thereby play a role in NADH-dependent conversion of both oxygen and H2O2, and possibly also other electron acceptors. c. Superoxide dismutase Superoxide radicals in cells are formed during the autoxidation of cellular components that are strong univalent electron donors, such as quinones and flavins. The one electron transfer reaction between oxygen and flavin leads to the formation of a so-called flavinsemiquinone and a superoxide molecule. The reactivity of the flavinsemiquinone with triplet oxygen and the reactivity of fully reduced flavin with superoxide leads to 26 autocatalytic superoxide formation through redox cycling (129, 130). The reactivity of the superoxide radical towards iron-sulfur clusters is an important factor in oxidative stress damage. Damage to iron-sulfur clusters leads to the inactivation of proteins with such clusters. The release of iron from damaged iron-sulfur clusters accelerates the 1 generation of ROS through the Fenton reaction (68, 131)(see Figure 1.1). Although two superoxide molecules can spontaneously dismutate to form H2O2, the catalysis of this reaction by SOD is in many species an important means to prevent oxidative stress. The presence of a functional SOD in several LAB species is wellestablished (132-137). Interestingly, the main source of superoxide generation in L. lactis was shown to be a supposedly water forming NOX, where micromolar amounts of H2O2 appeared to be exclusively produced via the combined activities of NOX and SOD (48). Whereas SOD in other organisms either uses iron, manganese or copper ions in its catalytic site, only the manganese-form was encountered in LAB. Expression of SOD was shown to contribute to aerotolerance (132, 134) and to H2O2-stress tolerance (125, 138), indicating that superoxide is a central oxygen intermediate contributing to cellular damage during oxygen exposure. 7. Physiological adaptations to protect against oxidative stress Despite the presence of several homologs of thioredoxin reductases, NPRs and glutathione reductases, no NADH-related H2O2 scavenging activity can be detected in the cell extract of the well-studied lactic acid bacteria L. lactis (48) and L. plantarum (139, 140). Analogously, cell extracts of L. plantarum lack SOD activity but the cells show remarkable resistance towards hyperbaric oxygen levels (141). These results indicate that apart from expression of enzymes dedicated to ROS scavenging, lactic acid bacteria employ physiological adaptations to protect against toxic oxygen derivatives. Here, we discuss how LAB utilize metal homeostasis, accumulation of pyruvate and thiol metabolism as a means to reduce ROS-induced damage. a. Intracellular manganese accumulation Apart from its role as a cofactor of catalase and SOD, manganese is accumulated to high intracellular levels in L. plantarum (>20 mM) and functions as a superoxide sink. LAB that show such high intracellular manganese levels (such as L. plantarum and L. 27 casei) are generally more resistant against superoxide-generating compounds such as plumbagin as compared to LAB that do not accumulate high levels of manganese (such as L. lactis and L. acidophilus) (141-143). There are several clues as to how such high intracellular levels of manganese may contribute to superoxide quenching. Bicarbonate-complexed manganese can contribute to H2O2 scavenging, which could indirectly also reduce superoxide toxicity (144). The authors of the original paper describing the phenomenon in L. plantarum suggested that pyrophosphate-complexed manganese could directly react with superoxide in vivo (141). Conversely, manganese accumulation in E. coli, which occurs in mutants lacking the genes for catalase and peroxidase, was effective in protecting against oxidative stress by replacing the reactive Fe2+ ions in metalloproteins and thereby reducing Fenton chemistry (145, 146). Attempts were made to identify the responsible manganese transporters in different LAB. The product of the mnt-gene was identified as a functional Mn2+ uptake system in S. oligofermentans (147) and L. plantarum strain ATCC 14917 (148, 149) but not in strain WCFS1 (150). A recent study reported that in S. oligofermentans, manganese import was regulated upon increasing H2O2 levels and mntA expression substantially contributed to oxidative stress tolerance (147). However, the mechanism remains poorly understood and apart from these studies, no new research has appeared creating a link between regulation of manganese homeostasis and oxidative stress in LAB. b. Regulation of intracellular iron levels Compared to regulation of intracellular manganese levels, controlling intracellular iron may be an even more fundamental physiological adaptation that underlies the aerotolerant nature of LAB. Where other micro-organisms compete with each other to obtain sufficient bioavailable iron sources, LAB seem to have sidestepped this rivalry. Although there are several exceptions (in S. pneumoniae (91) and in heme supplemented growth conditions (151)), the growth requirements of several Lactobacillus species for iron are nearly zero and the intracellular iron levels are very low (152-154). As was mentioned before, such intracellular unbound iron can engage in ROS formation through the Fenton reaction which accelerates the damage to cellular components. These low intracellular iron levels may be a direct consequence of the relatively low numbers of iron sulfur binding gene products in LAB compared to other bacteria such as E. coli. Moreover, several metalloproteins in LAB, such as catalase and SOD, were 28 shown to function with manganese ions instead of ferrous ions (see for instance our previous discussion of catalase and SOD in LAB). Through this reduced iron-dependency of protein function, lactic acid bacteria may avoid the need to maintain high intracellular iron pools, and may thereby be less sensitive to the toxicity of H2O2. 1 Besides the lack of iron-sulfur clusters, LAB also lack copies of the genes encoding bacterioferritin and ferritin, which are the major bacterial iron-storage proteins. The third iron binding protein DPS, which is associated with preventing oxidative stress (155, 156), is present in many LAB and showed to sequester iron and prevent H2O2-induced cell death in streptococci (75, 157, 158). We consider that even with low dependency on iron for protein function, oxidative stress in certain LAB is still correlated with levels of intracellular unbound iron. c. Pyruvate accumulation The metabolic intermediate pyruvate is an important branching point in the metabolism of LAB. Besides its role as central intermediate, pyruvate can be an effective H2O2 scavenger since it reacts non-enzymatically with H2O2, producing CO2 and acetate. The scavenging effect of pyruvate is clearly illustrated by the observation that exogenously provided pyruvate can protect against H2O2-killing in different types of cells (159-161). L. lactis is reported to accumulate substantial amounts of pyruvate (>90 mM) when grown at low growth rates (48, 162). This excretion of pyruvate was found to effectively reduce extracellular H2O2 levels and accompanying oxidative stress (48). It is not clear whether this results from enhanced production rates or from reduced dissipation rates due to stress-induced reduced function of metabolic enzymes (163). d. Cystine metabolism Cystine is a dimer of two cysteine molecules. The supplementation and uptake of cystine by a cystine-binding protein encoded by cyuC was shown to prevent H2O2 accumulation in L. reuteri BR11 (164). The intracellular conversion of cystine into smaller thiols such as H2S is catalyzed by cystathionine-γ-lyase. The gene encoding cystathionine-γ-lyase (cgl) is located in the same operon as cyuC, which also encodes a predicted cystine ABCtransporter (122, 165-167). The aerotolerance and oxidative stress resistance that this operon confers to L. reuteri BR11 indicates that such thiol metabolism may be involved in alternative pathways of oxidative stress resistance in LAB. 29 8. ROS production by LAB Of the abovementioned enzymes and physiological adaptations that are correlated to oxidative stress, only a few (SOD, catalase and AHPR) were shown to contribute to ROS scavenging and prevent the accumulation of the most stable ROS species H2O2. An analysis of the prevalence of these proteins in LAB leads to the conclusion that these proteins are absent in LAB like S. pneumoniae and L. johnsonii. These species only encode proteins such as TRXR, TPX and GSHR (Table 1.2), for which expression was found to be associated with oxidative stress resistance but in vivo ROS-scavenging functionality was not proven. An indication that these proteins are not, or only moderately involved in in vivo ROS scavenging is provided by the relatively high-level of H2O2 production during aerobic growth of LAB encoding these proteins. Hydrogen peroxide production by species of the L. acidophilus group and some streptococci presents an intriguing dilemma since the accumulating H2O2 leads to growth stagnation and cell death. One would assume that this creates a substantial evolutionary pressure for the acquisition and expression of genes encoding ROSscavenging proteins. The expression of either catalase, SOD or AHPR appears as an almost zero-cost option which would allow these species to counteract the damaging side-effects of the presence of oxygen. Nevertheless, despite their lack of ROS scavenging capacities, that are considered essential for aerotolerance in other bacteria, these H2O2 producing species show remarkable aerotolerance, with an initial aerobic growth rate that is comparable to the anaerobic growth rate and growth impairment only occurring when endogenously produced H2O2 reaches millimolar levels. 9. Concluding remarks In this literature review we have provided an overview of the different protein expression and physiological adaptations LAB use to deal with oxygenated environments. We find three overarching themes that play a role in aerotolerance and oxidative stress resistance of LAB. 1 The extent in which LAB can profit from the presence of oxygen. Hemin supplementation induces a respiratory phenotype in several LAB, which results in higher biomass levels and lower oxidative stress. Acetate production through the POX pathway can result in additional ATP generation. Furthermore, acetate and CO2 production through the POX pathway could relieve growth dependencies of some LAB and thereby expand the environmental niches that 30 can be colonized by these species (chapter 4 this thesis). 2 The extent in which a cell generates toxic derivatives during oxygen exposure. Several oxidases such as NOX, POX, LOX and NFR generate ROS which cause 1 oxidative stress. Aerobic lactate utilization in many LAB (through LDH/LOX and POX, facilitated by NAD+ recycling by NOX) can cause cell death due to concomitant H2O2 production. 3 The extent in which a cell is resistant against the toxic derivatives of oxygen (ROS). LAB express several ROS scavengers, such as thiol-based peroxidases, catalase and SOD which are effective in protecting against oxidative stress. Furthermore, regulation of metal homeostasis, such as intracellular accumulation of manganese, and decreasing iron dependency of enzyme functions, leads to a cellular physiology that is more oxidative stress-tolerant. The absence of ROS scavengers are correlated with a considerably lower aerobic biomass levels. We conclude that these cellular mechanisms underlie the general anaerobic aerotolerant nature of LAB. The diversity in phenotypes with respect to oxygen that is encountered within this group of bacteria, can in part be explained by the expression of oxidases and oxygen-dependent pathways, by the presence or absence of ROS scavengers and by physiological adaptations. 10. Lactobacillus johnsonii Lactobacillus johnsonii NCC 533 (previously referred to as Lactobacillus acidophilus La1) is a gram-positive, rod-shaped, non-sporulating, bacteriocin and exopolysaccharideproducing lactic acid bacterium, with a low G-C content (34.6%), belonging to the Lactobacillus genus of the Firmicutes phylum. Within the genomically diverse Lactobacillus genus, L. johnsonii is assigned to the L. acidophilus group on the basis of similarities in DNA and rRNA sequence (168). This group of bacteria has received extensive attention due to their proposed probiotic properties (169-171), their occurrence in the microbiota of the gastro-intestinal tract as well as those encountered in the oral and vaginal cavities in humans (94). Furthermore, species such as L. delbruecki subsp. bulgaricus, L. kefirofaciens and L. helveticus are prominently present in different starter cultures for the production of yoghurt, cheese and kefir. Importantly, to date, none of the species of the L. acidophilus group has been recognized as a potential pathogen in mammals. There has been a great effort in the last two decades to improve taxonomical classification 31 of this group, especially in order to refine the analysis of the enormous amount of human intestine microbiota data that have appeared over the past decade and to support the characterization of novel candidate probiotic strains. Comparative genomics studies revealed a high similarity at DNA, rRNA, protein, as well as metabolic level between the species of the L. acidophilus group (168, 172-175). For many of the species belonging to this group of lactobacilli there are now multiple genome sequences available and the group is continuously expanding due to the identification and sequencing of novel strains and species, which are often isolated from the GI-tract of mammals. L. johnsonii was the second species of the Lactobacillus-genus for which the complete genome sequence was published (176). The sequence indicates that this species has been subject to a process of reductive evolution, displaying a remarkable loss of genes (177, 178). L. johnsonii lacks the genes for the biosynthesis of numerous compounds, including secondary metabolites, amino acids, vitamins, and fatty acids. This is reflected in its fastidious growth requirements and its lack of metabolic versatility. L. johnsonii has been of particular interest to the food industry for its health-supporting properties. It is commercially used as the probiotic ingredient in the LC-1 fermenteddairy products marketed by Nestlé. In this application context, several studies have specifically looked at features related to adherence to the gut epithelium (171), its activity against enterovirulent pathogens in vitro and in vivo (38, 179, 180) and its immunomodulatory properties (181-183). In addition, broader understanding of the physiology and metabolism of L. johnsonii has been of relevance (184-186) to improve its survival and robustness in the harsh conditions encountered in industry and in the upper-gastrointestinal tract. These metabolic characteristics, such as its fastidious growth requirements, may also impact the interactions of L. johnsonii with its host organism. 11. Outline of this thesis L. johnsonii is, like other LAB, an aerotolerant anaerobe. Aerobic growth of this species is accompanied by the production of millimolar amounts of H2O2 (38), which in part results from the absence of several of the ROS-scavenging mechanisms present in other LAB (Table 1.2). This H2O2 production is an intriguing characteristic which L. johnsonii shares with several lactobacilli (94, 187) and streptococci (74, 85, 86). The enzymes involved in H2O2 production in L. johnsonii had previously not been elucidated. One of the primary goals of this thesis was to study the molecular mechanisms underlying 32 H2O2 production in L. johnsonii, e.g. to identify and characterize the enzymes that catalyze the H2O2 producing reactions and decipher the role of H2O2 production in the aerotolerance of L. johnsonii. In more generic terms the thesis also intends to provide a more global understanding of molecular responses of L. johnsonii to the presence of 1 molecular oxygen. In the first two experimental chapters of this thesis, we partially unravel the H2O2 producing reactions and aerobic metabolism in Lactobacillus johnsonii. In chapter 2, we identify and characterize the main H2O2 producing enzyme in L. johnsonii, which belongs to a novel NADH flavin reductase (NFR) enzyme family. In chapter 3 the genome-wide transcriptional response of L. johnsonii to the presence of oxygen is studied, leading to identification of a secondary H2O2 producing enzyme, which belongs to the NADH oxidase enzymes and appears essential for aerotolerance in the absence of NFR. The relationship between specific growth dependencies of L. johnsonii in the presence and absence of oxygen are subsequently studied in chapters 4 and 5. In chapter 4 we demonstrate that oxygen is not only detrimental for L. johnsonii, but can also relieve some of its growth dependencies. Endogenous acetate and CO2 production through the oxygen-dependent POX-reaction overcomes the dependency of this bacterium on exogenous C1 and C2- sources. In chapter 5, the growth dependency of L. johnsonii for CO2 is further characterized by analyzing the genome-wide transcriptional response to CO2 depletion. These analyses indicate that especially the pyrimidine biosynthesis pathway is depending on a CO2 supply. In chapter 6 the experimental results are discussed in the light of our current knowledge of the physiology of these types of lactobacilli. In addition, we discuss the implications of O2 and CO2 metabolism by L. johnsonii and other lactobacilli in the context of their interaction with mammalian host organisms. 33 34 HemCat lp_3578 - - - HemCat (query) L. plantarum WCFS1 L. lactis subsp. cremoris MG1363 L. rhamnosus GG L. casei BL23 E. faecalis V583 HemCat ef_15971 1 TRXR, ef_1338 2 TRXR, ef_2738 3 PNDR, ef_2899 1 TRXR, lcabl_10620 2 TRXR, lcabl_08900 1 TRXR, lgg_00920 2 TRXR, lgg_00810 1 TRXR, llmg_1588 2 AHPR, llmg_0357 3 TRXR, llmg_0776 1 TRXR (query) 2 ferredoxin NAD(P) reductase, lp_2585 TrxR2 lp_0761 AhpC/TSA family protein, ef_2932 TPX, lcabl_08080 TPX, lgg_00728 TPX, llmg_0318 TPX, lp_2323 Tpx3 SPD_1464 1 GSHR, lcabl_23620 2 GSHR, lcabl_27950 3 DLD, lcabl_15390 4 DLD, lcabl_16690 1 GSHR, ef_3270 2 DLD, ef_1356 3 DLD, ef_1661 1 GSHR, lgg_02615 2 DLD, lgg_01323 3 TrxR, lgg_00920 1 GSHR, llmg_1702 2 DLD, llmg_0071 3 PNDR, llmg_2331 4 TRXB1, llmg_1588 1 GSHR, lp_1253 2 GSHR, lp_3267 3 GSHR, lp_1822 4 GSHR, lp_0369 5 DLD, lp_2151 GshR4 (accession nr. A1YAC0) AhpC, ef_2739 AhpC lcabl_26730 AhpC, lgg_02490 AhpC, llmg_0356 - AhpC5 (accession nr. O66265) 1 NPR, lp_2544 2 NOX, lp_3449 3 NPR, lp_1445 4 NOX, lp_0760 5 NOX, lp_1941 6 NOX, lp_0766 7 NOX, pWCFS103_16 8 NOX, lp_1925 9 DLD, lp_2151 10 GSHR, lp_0368 1 NOX, llmg_0408 2 NOX, llmg_1770 3 hypothetical llmg_1249 4 PNDR, llmg_2331 5 DLD, llmg_0071 1 NPR lgg_00491 2 NOX lgg_00325 3 NOX lgg_00212 4 PNDR, lgg_00175 5 DLD, lgg_01323 1 NPR, lcabl_04690 2 NOX lcabl_ 02800 3 NOX lcabl_01750 4 DLD, lcabl_15390 1 NPR (query) 2 NOX, ef_1586 3 coenzyme A disulfide reductase ef_2989 4 PNDR, ef_1932 5 DLD, ef_1356 NPR6 ef_1211 MnSOD, ef_0463 MnSOD, lcabl_20710 - MnSOD (query) - MnSOD7 llmg_0429 Table S1.1; Antioxidative/ROS-scavenging mechanisms in LAB genomes. Cut-off values for BlastP: maximum e-value: 10-10, minimum identical 20%, minimum query coverage 60%). Supplementary materials - - - - S. mutans UA159 L. sanfranciscensis TMW 1.1304 L. delbrueckii subsp bulgaricus ATCC 11842 L. johnsonii NCC 533 TPX, jw_1317 TPX, lj_1153 - Hypoth., lsa_05160 Lipid hydroperoxide peroxidase, smu_924 TPX, stu_0990 TPX (query) - 1 hypoth., lj_0042 2 PNDR, lj_1757 3 TRXR, lj_0852 1 GSHR, jw_3467 2 DLD, jw_0112 3 PNDR, jw_5040 4 PNDR, jw_5551 1 PNDR, ldb_0759 1 hypoth. lsa_2p00270 1 GSHR spd_0685 2 PNDR,spd_1415 3 DLD, spd_1025 1 GSHR, stu_0408 2 DLD, stu_1048 3 PNDR, stu_0557 1 GSHR, smu_140 2 GSHR, smu_838 3 DLD, smu_1424 4 DLD, smu_130 1 GSHR, l897_03320 2 DLD, l897_03940 AhpC, jw_1106 - - - DLD, jw_0112 PNDR, lj_1757 PNDR, ldb_0759 NOX, lsa_05610 1 NOX, smu_1117 2 DLD, smu_1424 AhpC, smu_764 - 1 NOX, spd_1298 2 PNDR, spd_1415 3 DLD, spd_1025 1 NOX, stu_1281 2 PNDR, stu_0557 1 NOX, l897_0435 2 NPR, l897_06900 - AhpC, l897_08790 1 MnSOD, jw_3879 2 SOD [Fe], jw_1648 - - - MnSOD, smu_629 MnSOD stu_0720 MnSOD, spd_0667 MnSOD, l897_05695 1 Hemin-dependent catalase (HemCat) from E. faecalis, (96), 2 Thioredoxin reductase (TRXR) L. plantarum WCFS1 (113), 3 Thiol peroxidase (TPX) from S. Pneumoniae D39 (116), 4 Glutathione reductase (GSHR) L. sanfranciscensis DSM 20451 (122), 5 Alkyl hydroperoxide reductase (AhpC), S. mutans NCIB 11723 (74), 6 NADH peroxidase (NPR) from Enterococcus faecalis V583 (117), 7 Manganese-superoxide dismutase (MnSOD) L. lactis MG1363 (48), DLD = dihydrolipoamide dehydrogenase, PNDR = pyridine nucleotidedisulphide reductases/oxidoreductases, TPX = thiol peroxidase, GSHR= glutathione reductase, TRXR= thioredoxin reductase, NDH = NADH dehydrogenase, AHPR = alkyl hydroperoxide reductase 1 TRXR, jw_0871 2 AHPR, jw_0599 - S. thermophilus LMG 18311 HemCat, jw_1721 1 TRXR, lj_0852 2 hypoth., lj_0501 - S. pneumoniae D39 E. coli K-12 1 TRXR, l897_06810 2 NDH, l897_08795 3 ferredoxin NADP reductase, l897_03465 4 DLD, l897_03940 1 TRXR, spd_1287 2 PNDR, spd_1393 3 PNDR spd_1415 1 TRXR, stu_1650 2 TRXR, stu_1417 3 DLD, stu_1048 1 TRXR, smu_463 2 AHPR, smu_765 3 TRXR, smu_869 4 DLD, smu_130 5 DLD, smu_1424 1 TRXR, lsa_05170 2 ferredoxin NADP reductase, lsa_02530 1 TRXR, ldb_0613 2 TRXR, ldb_1586 - S. pyogenes HSC5 1 35 36 Chapter 2 Hydrogen peroxide production in species of the Lactobacillus acidophilus group, a central role for a novel NADH dependent flavin reductase. Rosanne Hertzberger1,3,4, Jos Arents1, Henk L. Dekker1, R. David Pridmore2,4, Christof Gysler2,4, Michiel Kleerebezem3,5, M. Joost Teixeira de Mattos1. 1) Swammerdam Institute for Life Sciences, University of Amsterdam, Science Park, Amsterdam, The Netherlands 2) Nestlé Research Centre, Vers-chez-les-Blanc, Switzerland 3) NIZO food research, Ede, The Netherlands 4) Kluyver Centre for Genomics of Industrial Fermentation, The Netherlands 5) Host Microbe Interactomics Group, Wageningen University, Wageningen, The Netherlands Published in Applied and Environmental Microbiology, (2014), 80:2229-39 Summary Hydrogen peroxide production is a well-known trait of many bacterial species associated with the human body. In the presence of oxygen, the probiotic lactic acid bacterium Lactobacillus johnsonii NCC 533 excretes up to 1 mM H2O2, inducing growth stagnation and cell death. Disruption of genes commonly assumed to be involved in H2O2 production, e.g. pyruvate oxidase, NADH oxidase and lactate oxidase, did not affect this. Here we describe the purification of a novel NADH-dependent flavin reductase, encoded by two highly-similar genes (LJ_0548 and LJ_0549) that are conserved in lactobacilli belonging to the Lactobacillus acidophilus group. The genes are predicted to encode two 20 kDa proteins containing FMN_red conserved domains. Reductase activity requires FMN, FAD or riboflavin and is specific for NADH, and not NADPH. The Km for FMN is 30 ± 8 µM in accordance with its proposed in vivo role in H2O2 production. Deletion of the encoding genes in L. johnsonii led to 40-fold reduced H2O2 formation, which could only be restored by in trans complementation of both genes. Our work identifies a novel, conserved NADH-dependent flavin reductase that is prominently involved in H2O2 production in L. johnsonii. 38 Introduction Hydrogen peroxide (H2O2) production is a well-known capacity of several bacterial species that are associated with the human body. Some of these H2O2-forming species are opportunistic-pathogens or pathobionts, such as Streptococcus pyogenes (86), Streptococcus mutans (74, 189), and Streptococcus pneumoniae (91, 190). Other H2O2 producing species or strains have been proposed to have probiotic properties, such as Bifidobacterium bifidus (69) and Lactobacillus johnsonii (38), or are prevalent in the commensal vaginal microbiota such as Lactobacillus crispatus, Lactobacillus jensenii and 2 Lactobacillus gasseri (94). Accumulation of H2O2 mainly occurs in species that lack the main H2O2-scavenging enzymes, such as catalase and NADH peroxidase. Analogously, when the genes encoding these enzymes are deleted from the Escherichia coli genome, H2O2 is generated upon oxygenation and accumulates in the extracellular growth medium (95, 191). Hydrogen peroxide is mainly produced in central carbon and energy metabolism by oxidases, including pyruvate oxidase (Pox), lactate oxidase (Lox) and NADH oxidases (Nox) (32). For example, activity of lactate oxidase encoded by S. pyogenes is primarily responsible for H2O2 production in cells that are depleted for glucose (192, 193), whereas H2O2 production by S. pneumoniae is due to pyruvate oxidase activity (91). In several species, H2O2-producing NADH oxidases have been identified: for instance in Thermus thermophilus (194), S. mutans (74) and Amphibacillus xylanus (72). Species that accumulate H2O2 or other reactive oxygen species (ROS) upon exposure to molecular oxygen generally have an energy metabolism that is adapted to anaerobic environments. The proteins that catalyze the low-potential redox reactions in anaerobic energy metabolism, such as fumarate and nitrate respiration, commonly carry lowpotential metal clusters and solvent-exposed flavin cofactors that readily react with oxygen and contribute to the generation of ROS. H2O2 and superoxide (O2-) belong to the strongest oxidant-species and can accelerate the rate of ROS generation, the chemistry of which has been reviewed previously (6). This study focuses on the H2O2 producing species L. johnsonii. This organism is applied as a probiotic supplement in the food industry (176). The strain was isolated from the human intestine, where it interacts with the host epithelium as well as with other microbes (170, 171, 195). The gastro-intestinal (GI) tract is predominantly an anaerobic niche, but the presence of oxygen gradients in the proximity of the mucosal surfaces is well established (40, 41, 196). Hydrogen peroxide derived from species like 39 L. johnsonii may play a role in these environments. Several studies have speculated on the effect that H2O2 may exert on the host as well as on the microbiome. Some authors propose that it can directly damage the epithelium (62, 197) and cause cell death of other bacteria (38, 190). Others suggest that H2O2 accumulation may contribute to the maintenance of a normal and homeostatic microbiota. Especially for the vaginal microbiota, strong evidence exists that women carrying H2O2-producing lactobacilli are less prone to develop bacterial vaginosis (187, 198), which is a very common disease and an independent risk factor for the acquisition of sexually transmitted disease and preterm birth (199, 200). Despite the data that support this hypothesis, the mechanism for the proposed homeostatic effect of H2O2 producing lactobacilli in the microbiota remains largely unknown. It has been suggested that H2O2 can contribute to the anti-inflammatory effect of commensal and probiotic bacteria through its influence on the peroxisome proliferator activated receptor-γ (PPAR-γ) which plays a central role in regulation of intestinal inflammation and homeostasis (201, 202). Expression of PPAR-γ is induced in vivo and in vitro by the presence of L. crispatus and is inhibited by the addition of either catalase or glutathione, pinpointing H2O2 as the responsible factor for the observed induction (50). A recent study on development of type 1 diabetes in rats substantiated the role in immune modulation by bacterially derived H2O2. Here, H2O2 directly affected the activity of indoleamine 2,3-dioxygenase which is an important immune modulator (203). Members of the L. acidophilus group are frequently encountereda the important microbial groups involved in H2O2 production in the vaginal and GI-tract microbiota. This group of lactobacilli encompasses several closely related species (168, 204), including those that are proposed to confer probiotic effects to consumers (L. johnsonii, L. gasseri, and L. acidophilus) as well as several important organisms in food fermentations (Lactobacillus delbruecki subsp. bulgaricus, Lactobacillus kefiranofaciens and Lactobacillus helveticus). Although many studies have reported on the H2O2 production by species of the L. acidophilus group, our understanding of the enzymatic reactions and mechanisms underlying these observations remains limited to the notion that NADH and flavin are involved in the reaction (37, 93) and that it is catalyzed by a protein that is constitutively expressed (94). The enzymes that catalyze the H2O2-generating reactions remain uncharacterized, to date. In this study, we identify a novel NADH-dependent flavin reductase as the primary source 40 for H2O2 in anaerobically grown L. johnsonii NCC 533, a member of the L. acidophilus group, upon exposure to oxygen. The enzyme is encoded by two small consecutive genes that show high similarity and are conserved throughout the L. acidophilus group. Mutation of these genes in L. johnsonii NCC 533 led to a strain that failed to produce H2O2 upon exposure to molecular oxygen. Materials & Methods Bacterial strains and culture conditions 2 L. johnsonii NCC 533 was obtained from the Nestec Culture Collection and cultured in commercial MRS medium (Merck, Whitehouse Station, NJ, USA) (205) at 37°C under static conditions, with minimal headspace for 16 hours. The deletion strains strains NCC 9333, NCC 9334, NCC 9337 and NCC 9359 were precultured in MRS containing 5 µg ml-1 erythromycin, while 5 µg ml-1 chloramphenicol was added for the NCC 9359 strains carrying plasmid pDP1016, pDP1017 or pDP1019. Growth and H2O2 production Cells were grown overnight in closed static tubes at 37°C in LAPTg medium. (20 g L-1 glucose, 10 g L-1 yeast extract, 10 g L-1 bacto peptone 10 g L-1 bacto tryptone plus 1 g L-1 Tween 80). This medium was used instead of the regular MRS medium for lactobacilli, because the meat extract in MRS was found to interfere with the enzymatic assay for H2O2. Cell density was determined by measurement of the optical density at 600 nm. H2O2 concentrations were determined using the phenol red assay (described below). Cell extracts Bacterial cultures were grown in 1 L bottles that were filled to the top with MRS medium, to minimize the headspace volume. Cultures were inoculated with 5 mL overnight precultures in the same medium and incubated for 24 hours at 37°C with continuous stirring. Cells were harvested by centrifugation (5’, 2600 x g, 4°C) and the cell pellets were suspended in 50 mL of 50 mM potassium phosphate buffer (pH 7.0) with 2 mM EDTA and 25 mM NaCl. Lysozyme (Sigma-Aldrich, St. Louis, MO, USA) was added to a final concentration of 1 mg mL-1 and incubated for 30 minutes at 37°C. Subsequently, cells were disrupted by 3 rounds of 1 minute sonication at 100W (Branson Ultrasonics, Danbury, CT, USA) and cooled on ice water. Crude cell debris was removed from the disrupted cell suspension by low-speed centrifugation (5’, 2600 x g, 4°C), followed 41 by ultracentrifugation of the supernatant (60’, 165000 x g, 4°C), generating the cell extract (supernatant) that was used in subsequent purification steps and enzyme assays. Protein concentrations of cell extracts were determined using the MicroBCA (microbicinchoninic acid) assay kit (Thermo Fisher, Scientific Inc., Waltham, MA, USA). Alternatively, 50 mL overnight anaerobic cultures in MRS were centrifuged, suspended in 50 mM potassium phosphate buffer (pH 7.0) and transferred to screw-cap tubes with 100 mg zirconium beads. Cells were disrupted in 3 rounds for 20 seconds and cell debris was removed by centrifugation (10’, 21500 x g, 4°C). Protein concentrations of cell extracts were determined using the MicroBCA assay kit (Thermo Fisher Scientific Inc). These extracts showed a 2-fold lower enzyme activity overall. They were employed for analysis of the activity, Km determination and SDS-gel. Protein purification: ammonium sulfate precipitation, Q column, gel filtration and SDS gel Cell extracts were placed in a beaker at 4°C and ammonium sulfate was added slowly under continuous stirring until reaching intermediate steps (30, 50, 70 and 90%) of saturation. Next, the cell extract was left without stirring on ice for 20 min and subsequently spun down (10’ min, 12000 x g, 4°C). The precipitate was suspended in 50 mM potassium phosphate buffer (pH 7.0) and dialyzed overnight at 4°C against 3 L 20 mM Tris buffer, pH 8.0. Anion-exchange chromatography was carried out with an Äkta FPLC system (GE Healthcare, Little Chalfont, Buckinghamshire, United Kingdom) fitted with a Hi-Trap Q HP 5 mL column (GE Healthcare). As a loading buffer, 20 mM Tris, pH 8.0 was used at a rate of 4 mL min-1 and a linear gradient was applied from 0 to 1 M NaCl in 20 column volumes. The fraction showing NADH oxidase activity (see below) was concentrated to 0.2 mL using a 10K Corning Spin-XUF6 column. This concentrated fraction was subjected to further fractionation by size exclusion chromatography using a Superdex 200 HR 10/30 column (GE Healthcare) in 20 mM Tris buffer (pH 8.0) with 250 mM NaCl at 0.5 mL min-1. For the SDS-gel electrophoresis, cell extracts were boiled for 5 minutes with SDS sample buffer (50 mM Tris-HCl pH 6.8, 100 mM dithiothreitol [DTT], 2% SDS, 0.1% bromophenol blue, 10% glycerol) and loaded on a 15% SDS gel with a 7% stacking gel on a Hoefer system (Thermo Scientific). The amount of sample that was applied was corrected for the variations in OD600 of the culture. The prestained marker Pageruler Plus from Fermentas (Thermo Scientific) was used. Gels were stained with PageBlue 42 Protein Staining Solution (Fermentas Coomassie G-250 dye). Enzyme activity assay NADH dependent flavin reductase activity in crude extract (for preparation, see above) was analyzed by determination of the NADH dissipation, as well as through the determination of the final H2O2 concentration. NADH oxidation was measured by monitoring the absorption at 340 nm at 37°C in a 200 µL reaction mixture with 500 µM NADH or 500 µM NADPH. As a flavin source either 250 µM flavin adenine dinucleotide 2 (FAD) or 25 µM FAD, flavin mononucleotide (FMN) or riboflavin was added. The reaction mixture was buffered by 50 mM potassium phosphate buffer at pH 7.0. For H2O2 measurement, the same reaction mixture in 200 µL volume was used with a lower NADH concentration (250 µM) to prevent oxygen from becoming the limiting substrate. After 10 minutes of incubation, the H2O2 concentration was determined with a phenol red enzymatic assay by transferring 20 µL sample to 180 µL of a reaction mixture containing 5 µg mL-1 horseradish peroxidase (Roche, Penzberg, Germany) and 30 µM phenol red in water. After 5 minutes of reaction, pH was increased by the addition of 10 µL 1M NaOH. Absorption was determined at 620 nm. These enzymatic activity assays were employed to detect protein activity in the fractions that were obtained from the three protein purification steps. Furthermore, the assays were used to determine the enzyme activity level in cell extracts of mutant strains. In the latter instance, 25 µM FMN was used as the flavin source. Enzyme activities are expressed as specific activity per milligram of protein in the cell extract per minute and were measured in triplicate. The specificity constant Km was determined by measuring NADH consumption rate of the cell extract in the presence of various concentrations of FAD, FMN and riboflavin (2.5 – 250 µM). The Km is calculated by fitting a hyperbolic curve (Vmax * Cs / (Km + Cs), with Cs for flavin concentration and optimization for Vmax and Km using the Solver function in Microsoft Excel. (R2 >0.96). Averages and standard deviations are calculated from technical triplicates. Digestion, MS analysis, and protein identification Semipurified protein samples were digested with an in-house protocol using 1 mg of trypsin (modified to prevent autodigestion, Promega, Madison, WI, USA) per 50 mg of 43 protein in a 0.1 M Tris pH 7.5 buffer following alkylation of the cysteine residues with DTT and iodoacetamide. Trypsin digestion was stopped after 16 hours by the addition of 10% trifluoroacetic acid (TFA) to a final concentration of 1%. Tryptic peptides were purified using an 80 µg capacity OMIX tip (Varian, Agilent) and collected in a volume of 30 µL 50% acetonitrile (ACN) - 0.1% TFA. Mass spectrometry analysis of the peptide samples was performed with a Micromass Q-TOF1 (quadrupole time of flight) mass spectrometer (Micromass, Waters, Milford, MA, USA) coupled to a nano-liquid chromatography (nano-LC) system (LC Packings, Dionex, Sunnyvale, CA, USA). The peptides were separated on a nano-analytical column (75 µm i.d., 25 cm length C18 PepMap, Dionex) using a gradient of 0–50% acetonitril and 0.1% formic acid. The LC eluent flow of 300 nL min-1 was directly infused into the Q-TOF1 spectrometer, operating in data-dependent MS and tandem MS (MS/MS) modes. Lowenergy collision-induced dissociation (CID) of selected precursor ions was used to obtain fragmentation spectra of the peptides. After processing the raw data with the Masslynx software (Micromass, Waters) the resulting peaklist (.pkl file) was used to search in the NCBInr database with MASCOT online (Matrix Science, Boston, Ma, USA). The search parameters were: a fixed modification of carbamidomethyl for cysteine, variable modifications of oxidized methionine, trypsin with the allowance of one missed cleavage, peptide and MS/MS tolerance ±0.3 Da and peptide charge state +1. Probability based MASCOT scores were used to evaluate the protein identifications. Construction of L. johnsonii deletion strains An overview of the mutants and plasmids used in this study can be found in table 2.1. An overview of all primers can be found in Supplementary material Table S2.1. The genome sequence of L. johnsonii is deposited in GenBank under accession no. AE017198 (176). 44 Table 2.1 List of strains and plasmids used in this study. Strain Description NCC 533 Lactobacillus johnsonii strain from the Nestec Culture Collection NCC 9333 Δpox (chapter 4) NCC 9334 ΔLJ_1826 (predicted to encode lactate oxidase; Lox) NCC 9337 ΔLJ_1254 and LJ_1255 (predicted to encode NADH oxidase; Nox) NCC 9359 ΔLJ_0548 and LJ_0549 (predicted to encode NADH flavin reductase) pDP749 pDP889 Temperature sensitive, allele exchange plasmid for L. johnsonii NCC 533 (185) pDP749 construct for lox (LJ_1826) deletion pDP902 pDP749 construct for nox (LJ_1254 and LJ_1255) deletion pDP1010 pDP749 construct for LJ_0548 and LJ_0549 deletion pDP794 pDP1016 pNZ124 based expression plasmid with LJ_0045 promoter and LJ_1125 terminator pDP794 with LJ_0548 expression plasmid pDP1017 pDP794 with LJ_0549 expression plasmid pDP1019 pDP794 with LJ_0548 and LJ_0549 expression plasmid 2 The construction of the pox-deletion strain NCC 9333 is described in the Materials & Methods section of chapter 4. The deletion of the gene LJ_1826, predicted to encode a lactate oxidase enzyme, was achieved similarly: the 5’ homology region of the LJ_1826 gene was amplified from L. johnsonii NCC 533 genomic DNA using primers A and B. The 1077 bp amplicon was SacI-BamHI digested and cloned in SacI-BamHI digested pDP749, yielding an intermediate plasmid. The 3’ region of the LJ_1826 gene was amplified using the primers C plus D, the 1170 bp amplicon digested with PstI-KpnI and cloned in the similarly digested intermediate plasmid to yield the lox-deletion plasmid pDP889. Plasmid pDP889 isolated from L. lactis was used to transform NCC 533 (185) and loop-in/loop-out gene replacement was achieved as described previously (206). The deletion was confirmed by PCR analysis and the deletion strain was named NCC 9334. In L. johnsonii NCC 533, NADH oxidase is predicted to be encoded by the genes LJ_1254 and LJ_1255. Deletion of LJ_1254 and LJ_1255 was achieved in the same way as for the LOX-encoding gen: the 1008 bp at the 5’ end region of the LJ_1255 gene was amplified using primers E and F, and the 993 bp at the 3’ end region of the LJ_1254 gene was amplified using primers G plus H. These amplicons were cloned into pDP749 to give plasmid pDP902 and used to produce the nox deletion strain NCC 9337. The deletion of LJ_0548 and LJ_0549 was achieved by amplification of 1062 bp at the 45 5’ end region of the LJ_0548 gene with primers I plus J and the 1098 bp at the 3’ end region of the LJ_0549 gene was amplified using primers K plus L. These were cloned into pDP749 to give plasmid pDP1010 which was used to produce the LJ_0548 and LJ_0549 deletion strain NCC 9359. Construction of the L. johnsonii overexpression strains The L. johnsonii expression plasmid pDP794 was constructed as follows: the predicted bidirectional terminator situated between the LJ_1125 and LJ_1126 genes of NCC 533 was amplified with the primers O and P. This 359 bp amplicon was digested with the restriction enzymes HindIII and XhoI and cloned into similarly digested pNZ124 (207) to yield pNZ124- LJ_1125 trm. The LJ_0045 D-lactate dehydrogenase promoter was amplified using NCC 533 chromosomal DNA as a template using the primers R plus S. This 215 bp amplicon was digested with the restriction enzymes BglII plus SacI and cloned into similarly digested pNZ124- LJ_1125 trm to produce plasmid pDP794. This plasmid, with the promoter region of the LJ_0045 lactate dehydrogenase gene and a bidirectional terminator, was used for overexpression of the LJ_0548 and LJ_0549 genes. For the construction of these overexpression plasmids the following cloning steps were performed: For pDP1016 the gene LJ_0548 was amplified using primers T plus V. The 597 bp amplicon was digested with SphI and HindIII and cloned into SphI plus HindIII digested pDP794 to give plasmid pDP1016. For pDP1017, the LJ_0549 gene was amplified using primers U plus W. The 583 bp amplicon was digested with SphI and HindIII and cloned into SphI plus HindIII digested pDP794 to give plasmid pDP1017. For pDP1019 the genes LJ_0548 and LJ_0549 were amplified using primers T plus W, the 1132 bp amplicon was digested with SphI and HindIII and cloned into SphI plus HindIII digested pDP794 to give plasmid pDP1019. These cloning procedures yielded plasmids on which expression of LJ_0548 and/or LJ_0549 is controlled by the strong ldh promoter. Genetic maps of plasmids pDP1016, pDP1017 and pDP1019 were created using Clone Manager (Supplementary materials Figure S2.1). Growth in batch culture Aerotolerance of NCC 9359 was compared to wildtype L. johnsonii in continuously stirred vessels with 400 mL MRS medium. Batches were sparged with specific gas mixtures containing 5% CO2 and either no oxygen (0% oxygen, anaerobic) or normal 46 oxygen levels (20% oxygen, aerobic). Cultures were grown at 37°C with continuous mixing (ca. 200 rpm) and pH was maintained at 6.5 by automated 4M NaOH titration. Cell densities were determined by measuring the optical density at 600 nm (OD600). Maximum specific growth rate was determined by fitting an exponential trend line through the data points with a minimal R2 of 0.99. Organic acid measurement by HPLC Extracellular metabolite concentrations were determined as described previously (208) 2 using high pressure liquid chromatography (HPLC, LKB and Pharmacia, Oregon City, OR, USA) fitted with a Rezex organic acid analysis column (Phenomenex, Torrance, CA, USA) at 45°C and an RI 1530 refractive index detector (Jasco, Easton, MD, USA). The mobile phase consisted of a 7.2 mM H2SO4 solution. Chromatograms were analyzed using AZUR chromatography software (St. Martin D’Heres, France). Statistical analysis Statistical significance was determined using a Student’s two tailed t-test for unequal or equal variance. An F-test was employed to verify whether variances could be considered equal (p>0.05) or unequal (p<0.05). Results H2O2 accumulation results in premature growth stagnation during aerobic growth of L. johnsonii NCC 533 To assess the growth behavior of L. johnsonii NCC 533 in anaerobic and aerobic conditions, LAPTg medium was inoculated with an overnight culture and incubated at 37°C either in a static tube with minimal headspace (anaerobic) or under continuous shaking with 10 volumes headspace (aerobic). Growth rates of aerobic and anaerobic cultures were similar up to an OD600 of 1.0 (Figure 2.1A). However, aerobic cultures accumulated up to 1 mM H2O2 during growth (Figure 2.1B), leading to growth stagnation at an approximate density of OD600 1.5. This growth stagnation could be completely abolished by the addition of 0.5 mg mL-1 catalase to the medium, which prevented the accumulation of H2O2. These findings show that oxidative stress resulting from endogenous H2O2 production is the main cause for the observed growth arrest of L. johnsonii NCC 533 under aerobic conditions. 47 A B 1,4 4 1,2 1 H2O2 (mM) OD 600 3 2 1 0,8 0,6 0,4 0,2 0 0 2 4 time (h) 6 8 10 0 0 2 4 time (h) 6 8 10 Figure 2.1: Growth and H2O2 concentration of L. johnsonii NCC 533 in LAPTg medium under anaerobic (square symbols), aerobic conditions (circular symbols) or aerobic conditions with 0.5 mg ml-1 catalase added to the medium (triangular symbols). Culture densities were determined by optical density measurement at 600 nm (Panel A) and H2O2 concentrations were determined by the phenol red enzymatic assay (Panel B). The data represent duplicate experiments ± standard error of the mean. H2O2 production is not dependent on predicted pyruvate oxidase, lactate oxidase or NADH oxidase encoding genes. The main contributor to H2O2 production in lactic acid bacteria (LAB) has been proposed to be the oxygen dependent lactate utilization pathway, which oxidizes lactate via pyruvate and acetyl-phosphate to acetate, generating CO2, ATP, NADH and H2O2 (83, 86). The redox balance in this pathway is proposed to be restored by dissipation of the NADH via an NADH-oxidase dependent reaction that generates either H2O2 or water. The oxygen-dependent lactate utilization pathway thereby encompasses three potential H2O2 producing reactions: (i) pyruvate oxidation (catalyzed by the Pox enzyme), (ii) lactate oxidation (catalyzed by the Lox enzyme) and (iii) NADH oxidation (catalyzed by the Nox enzyme). To assess the contribution to the observed H2O2 production of the genes predicted to encode these enzymes in L. johnsonii, mutant derivatives of the wild-type strain were constructed that lack the lactate oxidase encoding gene LJ_1826 (NCC 9334), the pyruvate oxidase encoding gene LJ_1853 (NCC 9333) or the NADH oxidase encoding genes LJ_1254 and LJ_1255 (NCC 9337). The H2O2 production capacity of the mutants was compared to that of the wild-type strain. To this end, all strains were grown anaerobically (static cultures) in LAPTg medium to an OD600 of ~0.7 and subsequently transferred to aerobic conditions (shake flask incubation). H2O2 production was measured after 1 and 2 hours of incubation. Deletion of the predicted pox, lox, or nox genes did not significantly affect the level of H2O2 production after 1 hour in these 48 strains in comparison with the level produced by the parental strain NCC 533 (Figure 2.2, all p-values >0.05). Exposure to oxygen for two hours resulted in small differences between H2O2 levels between the strains: the pox mutant produced less (0.48 mM vs 0.53 mM in the wild type, p<0.05) and the Δnox produced more H2O2 (0.56 mM, p<0.05). It appears justified to conclude that the oxidative lactate-utilization pathway is not responsible for the greater part of the H2O2 production, suggesting that an alternative metabolic conversion may account for the H2O2 production. This suggestion is in agreement with the observation that in the presence of oxygen, no substantial production and excretion of acetate occurs (chapter 4). 2 Previously, it has been suggested that L. delbrueckii, a close relative of L. johnsonii, produces H2O2 via an NADH dependent reaction that is enhanced by the addition of a flavin source (37, 93, 94). To identify the protein and gene involved in such proposed enzymatic reaction in L. johnsonii we initiated its purification, using enzyme activity assays to track the enzyme during purification. 0,7 0,6 mM H2O2 0,5 0,4 0,3 0,2 0,1 0 NCC 533 (wt) NCC 9333 (Δpox) NCC 9334 (Δlox) NCC 9337 (Δnox) Figure 2.2: H2O2 production of L. johnsonii NCC 533 and the derivatives NCC 9333 (Δpox), NCC 9334 (Δlox) and NCC 9337 (Δnox). Anaerobic logarithmic phase cultures were transferred to shake flask and incubated at 37°C. After 1h (open bars) and 2h (closed bars) cells were removed by centrifugation and H2O2 concentrations were determined in the culture medium, using the phenol red / peroxidase enzymatic assay. Data represent average of three independent experiments. Cell extracts of L. johnsonii NCC 533 contain NADH-dependent flavin reductase activity. Cell extracts of L. johnsonii NCC 533 contain NADH consumption activity when a flavin compound is added as a supplement to the assay’s reaction mixture (Figure 2.3A). The activity requires the addition of either FAD, FMN or riboflavin and does not show any activity with NADPH instead of NADH. Addition of 10-fold lower FMN concentration resulted in a significantly lower enzymatic rate (0.21 and 0.12 µmol / mg protein / min, p<.05). Following this observation, we further explored enzyme kinetics with different flavin sources at various concentrations. Michaelis-Menten-like kinetics were observed 49 when the flavin concentration is varied (see Materials & Methods for method of Km calculation), indicating that flavin is a direct substrate for the enzyme. The Km of free flavin does not significantly differ for the various flavins used: 30 µM ± 8 for FMN, 64 µM ± 21 for FAD and 41 µM ± 10 for riboflavin (p<.05). B 0,3 0,25 0,25 0,2 Endpoint mM H2O2 µmol NADH/ mg extract / min A 0,2 0,15 0,1 B 0,05 0 250 µM FAD 250 µM riboflavin 250 µM FMN NADH NADH 25 µM FMN no flavin 0,15 0,1 0,05 NADPH 25 µM FMN 0 25 µM FAD 25 µM Riboflavin 25 µM FMN NADPH, 25 µM FMN Figure 2.3A and B: Typical NADH dependent flavin reductase activity in L. johnsonii cell extract. NADH consumption rates were measured by absorption at 340 nm (Panel A). Endpoint H2O2 concentrations were determined using the phenol red assay (Panel B). Either 500 µM NADH was used, substituted by 500 µM NADPH where indicated (panel A) or 250 µM NADH substituted by 250 µM NADPH where indicated (Panel B). 250 µM or 25 µM of either FAD, FMN or riboflavin was added as a flavin source. Protein concentration in the cell abstracts was determined by the MicroBCA assay. Data represent the average of technical triplicates ± standard deviation and are representative of cell extracts derived in comparable experiments. The reaction is likely to involve a two electron transfer reaction since a considerable amount of H2O2 is formed as an end product, regardless of the flavin form that is added. When the FMN level in the assay is lowered to 25 µM, the H2O2 concentration exceeds the flavin concentration more than five fold (135 µM H2O2 ± 33 µM), indicating that the free flavin is recycled during the reaction (Figure 2.3B). Altogether, these observations allow the classification of the protein(s) responsible for the measured activity as an NADH-dependent flavin reductase. Purification and identification of the NADH-dependent flavin reductase activity. In order to identify the protein(s) responsible for the NADH-dependent flavin reductase activity, cell extract of wild type L. johnsonii NCC 533 was subjected to the following purification steps; (i) ammonium sulfate precipitation, (ii) Q column chromatography and (iii) gel filtration (see Materials & Methods for details). After the ammonium sulfate precipitation, the 50%-70% fractions clearly display most NADH consumption. After the subsequent fractionation of this fraction using anion exchange chromatography, highest activity clearly eluted after 83 an 86 ml (see Supplementary materials, Figure S2.2). These fractions were combined and were subsequently further separated by 50 size exclusion chromatography (Superdex2000). Only a single fraction eluted during this chromatography step that displayed clear H2O2 production in the enzymatic assay (Figure 2.4). The size of the enzymes in this fraction were, based on its elution time, estimated to be ~18kDa. The enzyme(s) in this fraction that showed NADH flavin reductase activity were only partially purified, since multiple bands were apparent when it was loaded on SDS-gel (results not shown). The active fraction obtained was digested with trypsin and analyzed using LC-MS/MS, using the fraction preceding this active fraction (and containing no activity) as a comparative negative-control. In the active fraction 25 L. johnsonii proteins could be assigned with a probability based score of 2 P<0.05. The protein that was predicted with highest probability was the hypothetical protein LJ_0548 to which 8 peptides could be assigned with a total coverage of 63%, indicating a high abundance in the fraction. Three peptides could be assigned to the hypothetical protein LJ_0549 (33% sequence coverage) which is encoded by LJ_0549, the LJ_0548 neighboring gene. Noteworthy, the predicted protein sequences of LJ_0548 (accession number Q74HL7) and LJ_0549 (accession number Q74HL8) both contain a conserved FMN reductase domain, supporting the role of these gene-products in the NADH dependent flavin reductase activity. Furthermore, no peptides belonging to these two proteins were detected in the fraction that did not show any NADH flavin reductase activity but eluted close to the active fraction (negative-control). Taken together, these observations pointed toward the involvement of the LJ_0548 and LJ_0549 genes in the NADH flavin reductase activity of cell extracts of L. johnsonii. 80 0,4 0,35 75 0,25 A 280 0,2 65 0,15 0,1 60 0,05 55 50 H 2O2 (mM) 0,3 70 0 0 5 10 15 20 Volume (ml) 25 30 -0,05 35 Figure 2.4: Size-exclusion chromatogram (Superdex200) of final purification step. Protein concentration is determined by absorption at 280 nm (black line). The eluting proteins were collected in fractions of 1 ml and tested for NADH flavin reductase activity by addition of 500 µM NADH and 250 µM FAD. H2O2 concentration was determined after 10 min (symbols). A mutant derivative of L. johnsonii NCC 533 was constructed that lacks the genes LJ_0548 and LJ_0549 (NCC 9359). The cell extracts of this mutant strain completely 51 lacked flavin dependent NADH reductase activity that was detected in the extracts obtained from the wild-type strain, nor could any H2O2 be detected in the reaction mixture supplemented with the mutant strain extract. These results indicate that the LJ_0548 and/or LJ_0549 encode the NADH dependent flavin reductase activity. The LJ_0548-0549 deletion strain (NCC 9359) was complemented by providing one or both of the deleted genes in trans on a plasmid under expression control of the strong, constitutive D-lactate dehydrogenase gene promoter (LJ_0045; ldhDp). Cell extracts derived from the NCC 9359 strain harboring either the LJ_0548- (pDP1016) or the LJ_0549- (pDP1017) expression plasmid did not show any additional bands on an SDSprotein gel. These cell extracts also did not show any significant NADH-dependent flavin reductase activity. Conversely, the extract derived from the strain harboring the plasmid expressing both the LJ_0548 and LJ_0549 genes (pDP1019) displayed an additional band of 20 kDa on an SDS-gel (Supplementary materials, Figure S2.3) and an NADH consumption rate that was more than 7-fold higher compared to the rate measured in extracts derived from the wild-type strain (Figure 2.5A). The level of H2O2 production driven by the extract derived from the strain overexpressing LJ_0548 and LJ_0549 was comparable to the level produced by the extract from the wild-type, which reflects the maximal level of H2O2 production that can be obtained in this assay as a consequence of the limited amount of NADH provided in the reaction mixture (Figure 2.5B). These results show that the LJ_0548-0549 operon encodes the observed NADH-dependent flavin reductase activity. H2O2 production of the LJ_0548 and LJ_0549 deletion strain Having identified LJ_0548 and LJ_0549 as coding for the enzymes responsible for NADH-dependent flavin reductase activity in the cell extracts of L. johnsonii NCC 533, we studied the in vivo contribution of this activity to the aerobic physiology of this bacterium by comparing the wild type to its nfr-deletion derivative (ΔLJ_0548-LJ_0549, NCC 9359). Maximum specific growth rate of the mutant in shake flask was similar to the wild type and its metabolism remained homolactic (result not shown). However, when exposed to stronger aeration (750 ml/min, 75% N2, 20% O2, 5% CO2), the nfr-deletion derivative displayed a reduced growth rate compared to wild type strain, whereas exposure to the anaerobic gas-mixture equivalent of this regimen (750 ml/min 95% N2, 5% CO2) did not result in a difference between the nfr-mutant and its wild type counterpart (Supplementary material, Figure S2.4). 52 A A B B 0,18 0,16 1 0,14 0,8 0,12 mmol H2O2 µmol NADH/ mg extract / min 1,2 0,6 0,4 2 0,06 0,04 0,2 0 0,1 0,08 0,02 NCC 533 NCC 9359 NCC 9359 +pDP 1016 NCC 9359 +pDP 1017 NCC 9359 +pDP 1019 0 NCC 533 NCC 9359 NCC 9359 +pDP 1016 NCC 9359 +pDP 1017 NCC 9359 +pDP 1019 Figure 2.5: Typical NADH dependent flavin reductase activity in cell extracts of wild-type L. johnsonii NCC 533 and its LJ_0548 - LJ_0549 (NCC 9359) mutant derivative, with and without complementation by plasmid-borne expression of one (pDP1016 and pDP1017) or both (pDP1019) of the deleted genes. Standard assay conditions were employed, containing 500 µM NADH and 25 µM FMN (NADH consumption rates) and 250 µM NADH and 25 µM FMN (to determine H2O2 concentration). NADH consumption rates were measured by absorption at 340 nm (panel A). H2O2 concentrations were determined after 10 minutes of reaction using the phenol red assay (panel B). Protein concentrations in the cell extracts were determined by MicroBCA assay. Data represent average of three technical replicates ± standard deviation and are representative for activity measured in multiple (>3) cell extracts. To test H2O2 production, the wild-type strain (NCC 533) and its LJ_0548-0549 deletion derivatives were grown anaerobically to mid-logarithmic phase (OD600 ~0.8) at 37°C and were then transferred to aerobic (shake-flask) conditions. H2O2 in the spent medium of each of the cultures was assessed after 1 hour of oxygen exposure (Figure 2.6). Notably, the LJ_0548-0549 deletion (strain NCC 9359) resulted in complete loss of the capacity to produce H2O2 under these conditions, while substantial amounts of this reactive oxygen molecule were detected in the NCC 533 culture exposed to the same conditions. Moreover, the NCC 9359 mutant strain that was in trans complemented with plasmid borne expression of either LJ_0548 (pDP1016) or LJ_0549 (pDP1017) did not produce detectable H2O2 levels, whereas the strain complemented with plasmidborne expression of both LJ_0548 and LJ_0549 (pDP1019) displayed a restored H2O2 production capacity, comparable to that observed in the parental strain NCC 533. These data confirm that the L. johnsonii NCC 533 NADH-dependent flavin reductase is encoded by the LJ_0548-0549 cluster and that this activity is the major H2O2 producing system expressed under the conditions employed here. In order to test if LJ_0548 and LJ_0549 play a role in anaerobic fumarate respiration of L. johnsonii, the external metabolite profiles of wild type NCC 533 and NCC 9359 (the 53 LJ_0548 and LJ_0549 deletion derivative) were compared. After 7 hours of anaerobic growth in MRS medium supplemented with 10 mM fumarate, cells were removed by centrifugation and external metabolites were analyzed using HPLC. No change in the concentration of fumarate was observed and no succinate formation was detected. Both aerobic and anaerobic metabolism of the mutant strains remained entirely homolactic (results not shown). 0,25 H2O2 (mM) 0,20 0,15 0,10 0,05 0,00 NCC 533 NCC 9359 NCC 9359 NCC 9359 NCC 9359 + pDP1016 + pDP1017 + pDP1019 Figure 2.6: H2O2 production in L. johnsonii NCC 533 and NCC 9359, with or without complementation of LJ_0548 an LJ_0549 under aerobic conditions. Anaerobic logarithmic phase cultures of wild type L. johnsonii and its deletion derivative were transferred to shake flask and incubated at 37°C. After 1h cells were removed by centrifugation and H2O2 concentrations were determined in the culture medium, using the phenol red assay. Data represent average of three independent experiments ± standard deviation. Sequence analysis of LJ_0548 and LJ_0549, and their conservation among bacteria Using the wealth of genomic sequence availability, the prevalence and context of the LJ_0548 and LJ_0549 single genes as well as the combination of the two consecutive genes were analyzed using diverse genome comparison tools. The LJ_0548 and LJ_0549 genes are predicted to encode proteins of 178 and 184 residues, respectively, that share substantial similarity (40% identity and 59% similarity at amino acid sequence level). The protein domain signature recognition module Interproscan (209) revealed that both genes have a highly conserved FMN reductase domain (PFAM domain 03358), covering the N-terminal 145 residues. In the PANTHER classification system program (210), LJ_0548 was classified as a chromate reductase, a group of enzymes that has been annotated as such due to their potential use in chromate bioremediation (211). The ortholog in Pseudomonas putida has been shown to catalyze the transfer of electrons from NADH to the quinone pool (212). The crystal structure of the E. coli gene annotated as chromate reductase clearly demonstrates the amino acids that constitute the flavin binding site (213). A ClustalW multiple sequence 54 alignment shows that four out of the eleven residues of this binding site are conserved in both LJ_0548 and LJ_0549, i.e. Ser18, Asn20, Glu82 and Ser 117 (Supplementary Materials Figure S2.5). The LJ_0548-0549 locus in L. johnsonii NCC 533 appears to be conserved in all other members of the L. acidophilus group. Examination of the genetic context in more distant species using the STRING module (214) revealed that the closest homologues of LJ_0548 and LJ_0549 are encountered as consecutive genes in several different species (see Figure 2.7), including L. plantarum, and several species belonging to the 2 Streptococcus, Enterococcus and Pediococcus genera. In these examples, the first of these two genes is similar in sequence and size to either LJ_0548 or LJ_0549 and is followed by a second, larger gene, of which the N-terminal residues (~200) are homologues to LJ_0548. This type of arrangement is also present in a more distant species from the Actinobacteriaeae class, Atopobium parvulum, which is a species often found in the human oral cavity(215). In all aforementioned species, the homologues of LJ_0548 and LJ_0549 are annotated as fumarate reductases, NADH dehydrogenases or flavin reductases, but to the best of our knowledge there is no experimental data to support these annotations. 55 Figure 2.7: Genetic context conservation of LJ_0548 and LJ_0549 found using the STRING module. 56 Discussion Our work has identified a novel NADH-dependent FMN reductase in L. johnsonii NCC 533, that is encoded by two adjacent genes (LJ_0548 and LJ_0549) and acts as the major H2O2 producing system in this bacterium. L. johnsonii is a lactic acid bacterium that belongs to the phylogenetically closely related L. acidophilus group (168, 216), which includes several strains of Lactobacillus species that are marketed as probiotic supplements like L. acidophilus, L. johnsonii, L. jensenii, L. crispatus and L. gasseri, but also encompasses the well-known yoghurt bacterium L. delbrueckii. It 2 has been established that these species endure oxidative stress as a consequence of endogenously produced H2O2 (35-37). The enzymes identified here have characteristics that are in agreement with those observed previously in strains of the L. acidophilus group, including that the enzyme reaction consumes NADH, uses flavin as a cofactor, is constitutively expressed and is not produced in response to molecular oxygen (37, 93, 94). In addition, the maximal reaction rate measured in cell extracts of L. johnsonii is in the same order of magnitude as the rate previously reported for L. delbrueckii (37). Apparent Michaelis-Menten kinetics were observed when the concentration of flavin was varied in the enzymatic assay. However, the H2O2 produced in the assay reaches a higher level than the total initial concentration of flavin added to the assay, indicating that free oxidized flavin serves as a substrate in this reaction but the reduced flavin is subsequently oxidized and reused. This is in agreement with the behavior of reduced flavins that spontaneously react with oxygen, yielding H2O2. Despite the recognition of this enzyme activity in various L. acidophilus group species, the molecular characteristics and genetic determinant(s) of this activity have not been described to date. Therefore, our study is the first to identify and characterize this novel enzyme family and its encoding genes. The enzyme is shown to be responsible for the major H2O2 production in an industrially relevant member of the L. acidophilus group, L. johnsonii. This capacity has previously been proposed to influence gut homeostasis and anti-inflammatory activity of this group of organisms (50) and the identification of the responsible genes and the construction of the corresponding deletion strain may accelerate the establishment of this presumed function in vivo, which is assumed to play a role in the bacterium’s protective effect against vaginal disease (187). The Vmax and Km values of the different flavins that were tested in the enzymatic assay did not differ strongly. The Km value for flavins were found in the range of 30-50 µM. Intracellular flavin concentrations in E. coli and Shewanella oneidensis were reported 57 in the order of 0.5 µmol per gram protein (217), which in combination with the “rule-of-thumb” estimate of ~200 g/l as the concentration of intracellular protein in prokaryotes (218) implies that the intracellular flavin concentration would be ~100 µM. The partitioning of flavin bound to protein and available as electron acceptor for the proteins we describe here is unknown. One study on intracellular free FAD concentration in Amphibacillus xylanus finds 13 µM using HPLC (219). Such a flavin concentration in L. johnsonii would be sufficient for its proposed in vivo role. Given the clear confirmation of the in vivo role of Nfr by the physiology of the nfr-deletion mutants, we consider it a valid conclusion that analogous to the previously found concentration in A. xylanus the free flavin levels in L. johnsonii suffice for significant H2O2 production. Surprisingly, the genes that are considered to be responsible for H2O2 production in other LAB were shown to not contribute significantly to this phenotype in L. johnsonii, since the genes predicted to encode lactate, pyruvate or NADH oxidases could be deleted without consequences for the H2O2 production in this species. The homologs of LJ_0548 and LJ_0549 have been annotated as fumarate reductase, NADH dehydrogenase or NAD(P)H dependent FMN reductase. The LJ_0548 and LJ_0549 proteins are predicted to be small flavoproteins that are highly similar (40%). The size of the denatured protein components in the cell extract of the LJ_0548-0549 overexpressing strain on the SDS-gel is 20 kDa, corresponding to the size of the gene product of either LJ_0548 or LJ_0549 as inferred from their gene sequence. Its elution from the gel filtration column suggests that the active protein has a size of ~18 kDa, which would mean that only one of either LJ_0548 or LJ_0549 would be required for activity. However, for complementation of the LJ_0548-0549 deletion strain, both genes appear to be required, whereas in trans complementation with a plasmid harboring either one of the genes failed to result in detectable protein expression (SDS-PAGE) or functional complementation. Although we can conclude that both genes are required to produce the functional enzyme, its exact composition remains unclear. The observation that the deletion derivative NCC 9359 produces small amounts of H2O2 upon prolonged exposure to oxygen indicates that besides the NADH flavin reductase identified here, other H2O2 producing enzymes may exist in this species. Nevertheless, the enzyme identified here appears to be the major contributor to the H2O2 production capacity in this species. Possibly, the additional H2O2 producing reactions involve oxidases, like the aforementioned pyruvate, lactate and NADH oxidase, which may contribute to H2O2 production upon extended oxygen exposure. However, it is unlikely that the conditions of oxygenation used in this study, both in terms of its duration and/or 58 oxygen tension, will be encountered in the GI-tract, which is thought to be the natural habitat of L. johnsonii. We propose therefore that the constitutive flavin reductase is the primary source of H2O2 in an environment where microbes predominantly encounter anaerobic (intestinal lumen) conditions and only sporadically encounter lower and more variable concentrations of oxygen when they are present in closer proximity to the intestine mucosa. In contrast, prolonged exposure to aerobic conditions and/or higher oxygen tensions can occur during industrial processing, which may elicit the activation of alternative H2O2 production reactions as our preliminary observations imply. 2 L. johnsonii has been proposed to have lost numerous genes and pathways during its adaptation to the nutrient-rich environment of the intestinal tract (176). Nevertheless, the newly discovered NADH dependent flavin reductase appears to be constitutively expressed, suggesting that it plays an important role in the lifestyle of L. johnsonii in its natural environment. Since lactate fermentation from glucose is entirely redox-neutral, it is unclear in what metabolic step the NADH is generated that is consumed in the reaction catalyzed by the LJ_0548-549 enzyme. We hypothesize that the additional electrons are generated in the metabolism of one of the many vitamins, peptides and amino acids that are consumed by L. johnsonii in addition to glucose. Although the results presented here do not rule out that this newly identified flavin reductase serves a metabolic purpose in which H2O2 is a side product, we suggest that the production of H2O2 in itself has a biological function. For example, it may contribute to the antimicrobial capacities of L. johnsonii that may be of great importance for the organism to maintain its niche / position within the densely populated microbiota (38). Moreover, H2O2 may serve as a chemical signal in host-microbe interactions, as it has been proposed to influence PPAR-g, one of the major regulators of inflammation in the intestinal epithelium (50). Alternatively, the reduced aerotolerance of the nfrdeletion derivative as compared to its wildtype counterpart suggests that the reaction catalyzed by this enzyme may enable L. johnsonii to prevent or reduce oxidative stress. If the flavins that are reduced by these proteins, form the most readily oxidized parts in the cytoplasm and can effectively capture oxygen, the activity of these flavins may prevent other, more damaging effects of oxygen, such as the direct oxidation of ironsulfur clusters (220, 221) or the formation of semiquinones (212). Also in L. johnsonii, the controlled production of H2O2 may be preferred over the uncontrollable other effects oxygen might exert. The role that this NADH dependent flavin reductase plays in oxidative stress, H2O2 scavenging and aerotolerance of L. johnsonii is the subject of further studies. 59 Acknowledgments This work was supported by Nestlé Research Centre, Vers-chez-les-Blanc, Switzerland. We would like to acknowledge Anne-Cécile Pittet for her technical assistance in construction of the mutants and Filipe Branco dos Santos for his valuable input on the purification of the NADH flavin reductase activity. 60 Supplementary materials Table S2.1: Primers used for construction of L. johnsonii mutant strains (chapter 2). Region ID Sequence LJ_1826 5’ homology 5’ primer A CTACTCCAGAAGAAGTCG LJ_1826 5’ homology 3’ primer B ATATATGGATCCAAGGGTGAAGGACAAAGC LJ_1853 3’ homology 5’ primer C GGGTTTTGCATTCCAGTC LJ_1853 3’ homology 3’ primer D ATATATGGTACCTTGGCCAAAAGTTGGAGC LJ_1254- LJ_1255 5’ homology 5’ E ATATATGAGCTCAATACTCAATGTAAGCGC LJ_1254- LJ_1255 5’ homology 3’ F ATATATGGATCCTGATTGCAGGTCCACCTG LJ_1254- LJ_1255 3’ homology 5’ G ATATATGAATTCTGATTTAGTAGCTGCTGG LJ_1254- LJ_1255 3’ homology 3’ H ATATATGGTACCCAGAAACTATGAAGGCTC LJ_0548- LJ_0549 5’ homology 5’ I ATATATGAGCTCGCTGCAAATGAAGGGCTAGA LJ_0548- LJ_0549 5’ homology 3’ J ATATATGGATCCGCGTTGCTACCTACAATGGC LJ_0548- LJ_0549 3’ homology 5’ K ATATATGAATTCTTTAATTGGTCATGCTGCAG LJ_0548- LJ_0549 3’ homology 3’ L ATATATGGTACCAGCTCGCCTTCACTACGGAG LJ_1125 term 5’ O ATATATAAGCTTTGCCAATGGATAACCAGG LJ_1125 term 3’ P ATATATCTCGAGAATCTCTCTTGGACTTGC LJ_0045 promoter 5’ R ATATATAGATCTCATTATCATAAGGCACCC LJ_0045 promoter 3’ S ATATATGAGCTCGCTAGCGCATGCATTAAACCTCCGTC SphI site before LJ_0548 T ATATATGCATGCAACTCTTTGCCATTGTAGG SphI site before LJ_0549 U ATATATGCATGCAATTACTAGCAATTGTTGG HindIII site after LJ_0548 V ATATATAAGCTTAATTTCATGGGTCGTTCCTC HindIII site after LJ_0549 W ATATATAAGCTTGGTTTTAACTTATTTTTGAGCTTG 2 61 62 ori RepC 2500 3000 Term 2000 3860 bps LJ0548 1000 HindIII 500 1500 pDP1016 3500 XhoI MunI MunI RepA SspI p-LJ0045 CAT MunI BglII PstI NheI SphI NruI BglII StuI NcoI SspI NdeI ori RepC 2500 3000 SalI Term 2000 3867 bps 1000 SspI HindIII SphI MunI EcoRI BglII StuI NcoI EcoRI NcoI p-LJ0045 CAT MunI PstI LJ0549 500 1500 pDP1017 3500 XhoI MunI MunI RepA Figure S2.1: Genetic maps of LJ_0548 and/or LJ_0549 overexpression plasmids. SspI NdeI SalI XhoI MunI MunI ori RepC SspI NdeI HindIII PstI 1000 SspI MunI LJ0548 p-LJ0045 CAT MunI EcoRI NcoI EcoRI LJ0549 2000 4416 bps pDP1019 4000 Term 3000 RepA SalI BglII PstI NheI SphI NruI BglII StuI NcoI NADHconsumptionrate (µM / s) 70 60 50 40 30 2 20 10 30%-50% 70% - 90% 90%+ 50 40 fraction 29 fraction30 30 A 280 1000 900 800 700 600 500 400 300 200 100 0 50%-70% 0 20 10 0 50 100 Eluted volume (ml) 150 -10 NADH consumption rate (uM/s) 0 Figure S2.2: Purification of NADH flavin reductase activity in cell free extract of wild type L. johnsonii. NADH consumption rate (in the presence of 250 µM FAD) is determined in fractions obtained during ammonium sulphate precipitation (panel A). The fraction with highest activity is subsequently used for anion exchange chromatography. All fractions eluting from the Q column (panel B) are tested for NADH flavin reductase activity. On the left axis, absorption at 280 nm is shown which is an indicator of protein concentration. On the right axis, NADH consumption rate is shown in the eluting fractions. Figure S2.3: SDS gel of cell free extracts of L. johnsonii NCC 533 (wt, lane 1), NCC 9359 + pDP 1016 (Δnfr + LJ_0548, lane 2), NCC 9359 + pDP 1017 (Δnfr + LJ_0549, lane 3), NCC 9359 (Δnfr, lane 4), NCC 9359 + pDP 1019 (Δnfr + LJ_0548-LJ_0549, lane 5), PageRulerTM marker (lane 6). 63 1,2 1 µ (h-1) 0,8 0,6 0,4 0,2 0 anaeroob aerobic Figure S2.4: Growth rate of L. johnsonii NCC 533 (grey bars) and its nfr-deletion derivative NCC 9359 (white bars) in MRS-medium in stirred pH controlled sparged with 750 ml/min of N2 + 5% CO2 (anaerobic) or N2+ 20% O2+ 5% CO2 (aerobic). Growth rates were determined as explained in Materials & Methods. Data are average of triplicate experiments ± standard deviation. L. johnsonii LJ_0549 L. johnsonii LJ_0548 E. coli K12 ChrR ----MKLLAIVGTNADFSYNRFLDQFMAKRYKDQAEIEVY-EIADLPRFK 45 ----MKLFAIVGSNADHSYNRDLLNFIKKHFTDRYDIELG-EVKDLPMFK 45 MSEKLQVVTLLGSLRKGSFNGMVARTLPKIAPASMEVNALPSIADIPLYD 50 :::.:::*: . *:* : . : * ::: .: *:* :. L. johnsonii LJ_0549 L. johnsonii LJ_0548 E. coli K12 ChrR KEAQP----DSKVEEFKNKIREADGVIFATPEYDHGIPSALKSAMEWTGS 91 EGVKE----PAAVASFAKKVADADAVLISTPEQQHSVPSSLKSALEWLSS 91 ADVQQEEGFPATVEALAEQIRQADGVVIVTPEYNYSVPGGLKNAIDWLSR 100 .: : * : ::: :**.*:: *** ::.:*..**.*::* . L. johnsonii LJ_0549 L. johnsonii LJ_0548 E. coli K12 ChrR HAQGNADVMKMKPAMVLGTSYGIQGASRAQEEMREILLSPDQSANVLPGN 141 AEHP----FKDKPVVIVGTSVLPQGSARGQSHLKLVLSSPGFGAKVFNGD 137 LPDQP---LAGKPVLIQTSSMGVIGGARCQYHLRQILVFLD--AMVMNKP 145 . : **.:: :* *.:* * .:: :* . * *: L. johnsonii LJ_0549 L. johnsonii LJ_0548 E. coli K12 ChrR EVLIGHAADKFDKNTGDLLDQETIHAIDLAFNNFVKFVEQAQK 184 EFMMGTAPEQFDENGN--LPAGTVKFLDHFFDEFDSFYAEVSK 178 EFMGGVIQNKVDPQTGEVIDQGTLDHLTGQLTAFGEFIQRVKI 188 *.: * ::.* : . : *:. : : * .* ... Figure S2.5: ClustalW2 comparison of LJ_0548 / LJ_0549 to chromate reductase (ChrR) from E. coli. Highlighted in grey are the residues that constitute the flavin binding site, highlighted in yellow are the similarities in these residues in the L. johnsonii genes. 64 2 65 Chapter 3 Transcriptome response in Lactobacillus johnsonii identifies an oxygen induced NADH oxidase that contributes to H2O2 production Summary Oxidative stress due to endogenous hydrogen peroxide production by Lactobacillus species is a well-known issue in the food industry. In this study, the transcriptional response to oxygen was analyzed in Lactobacillus johnsonii, one of the H2O2-producing strains used in the food industry. Aerobic growth conditions led to a more than two-fold repression of 45 gene-specific transcripts as compared to anaerobic growth, whereas transcripts of 6 genes were more than two-fold induced. Among the higher expressed genes were two genes that displayed significant homology to NADH-dependent oxidoreductase (NOX). The transcriptional regulation of the nox promoter by oxygen was verified using a GUS-reporter construct, whereas the nox promoter activity did not appear to respond to other oxidative conditions, e.g. exposure to sublethal levels of H2O2. Experiments in chapter 2 showed that H2O2 production by L. johnsonii largely depends on genes encoding an NADH flavin reductase (NFR). However, here we show that an NFR deficient strain could regain its H2O2 producing capacity upon prolonged oxygen exposure, which was hypothesized to involve the oxygen induced nox locus, of which the transcription appeared to be more strongly induced by oxygen in the NFR-deficient strain as compared to its parental strain. Indeed, deletion of the NOX-encoding locus in the NFR-deficient background, resulted in a strain that could no longer produce H2O2. Moreover, the NFR-NOX deficient strain (nfr-, nox-locus) displayed strongly impaired aerobic growth and oxygenation induced rapid H2O2 independent growth stagnation. We conclude that H2O2 production in L. johnsonii is primarily dependent on NFR but also involves the oxygen-inducible NOX. Moreover, our results imply that the capacity to produce H2O2 plays a prominent role in oxygen tolerance of L. johnsonii. 68 Introduction The lactobacilli belonging to the L. acidophilus group play a key role in the food industry. They are used in cheese fermentation (e.g., L. helveticus), kefir fermentation (e.g., L. kefirofaciens) and in yoghurt fermentation (e.g., L. delbrueckii subspecies bulgaricus). Especially the latter application is of great economic importance and represents a rapidly growing multibillion market. In these dairy fermentations the lactobacilli primarily contribute to the acidification of the milk by the production of lactic acid, but also contribute to the product’s texture and flavor by producing exopolysaccharides, and specific volatile components like acetaldehyde and diacetyl, respectively (222). In addition to these features, several species belonging to the L. acidophilus group are marketed as probiotics that, when consumed in adequate amounts, can convey a health benefit to the consumer (223). 3 For reliable industrial application of lactobacilli, their stress tolerance and functional robustness under industrial conditions is of great importance. To effectively initiate and complete fermentation processes, it is required that the Lactobacillus starter cultures survive the stressful industrial production and processing conditions. Similarly, sustained viability during product-processing and shelf-life of probiotic-containing products is crucial for this product-category to deliver their health benefit to the consumer (224, 225). Thereby, robustness and sustained viability of probiotic cultures under industrial and product conditions is essential to ensure their efficacy. In the life cycle of probiotic lactobacilli, from industrial production to in situ delivery in the intestinal tract, oxidative stress exposure is considered an important cause of viability loss (35). The product environment commonly contains relatively high levels of oxygen (46), in which the lactic acid bacteria that belong to the L. acidophilus group produce significant amounts of H2O2 (37, 38, 94) even at chilled temperatures (36). Lactic acid bacteria are fermentative microorganisms that lack the endogenous capacity for respiration, but many species and strains of this group are relatively aerotolerant and grow well in aerobic environments (see chapter 2). LAB employ several mechanisms for reactive oxygen species (ROS) detoxification to relieve oxidative stress and prevent its corresponding damaging effects on cellular constituents (32). These mechanisms include enzymes that target the ROS hydrogen peroxide (H2O2), such as the heme-requiring catalase expressed by L. sakei (226), and the heme-independent catalase of L. plantarum (100). The latter species is also protected against ROS, including superoxide (O2-) induced cell-damage by its capacity to accumulate high intracellular levels of manganese that 69 can directly scavenge superoxide molecules (141). Superoxide detoxification can also be achieved by the enzyme activity of superoxide dismutase (SOD), which was shown to play a key-role in oxygen tolerance in Lactococcus lactis (188). In addition, several lactic acid bacteria, contain glutathione and thioredoxin reductases (113, 114, 122), peroxidases and/or alkyl hydroperoxide reductases (74, 75, 227), which can effectively remove reactive oxygen species from the cell and maintain reducing conditions in the cell’s cytoplasm. Apart from the thioredoxin reductase system, the lactobacilli belonging to the L. acidophilus group commonly lack these ROS-scavenging activities, which is an important determinant in the H2O2 accumulating capacity that is seen in many species and strains of this group of the lactobacilli (37, 93, 94). A constitutively expressed NADH flavin reductase (NFR), composed of two small (20kDa) flavoproteins, was shown to be the major source of the accumulating H2O2 when anaerobically growing cells were exposed to molecular oxygen. The formation of H2O2 induces premature growth stagnation and cell death in L. johnsonii under aerobic growth conditions (chapter 2) (228). Conversely, L. johnsonii may also benefit from oxygen exposure, which can drive the endogenous pyruvate oxidase-dependent (POX) metabolic pathway that produces C1- and C2- metabolic intermediates that are required for growth. This growth stimulatory effect of oxygen was clearly demonstrated by the observation that L. johnsonii strictly depends on environmental supplies of acetate (C2) and inorganic carbon (C1-) sources when grown under strict anaerobic conditions (chapter 4). Oxygen therefore plays a paradoxical role in the lifestyle of L. johnsonii: its presence results in H2O2 accumulation which threatens viability, but at the same time alleviates some of its environmental growth requirements. Several studies have focused on the transcriptional response of LAB upon their exposure to oxygen, revealing the activation of oxidative stress-related genes, including those encoding ROS-scavenging enzymes, such as the thioredoxin reductase in L. plantarum (113) and the heme-catalase in L. sakei (226). Moreover, aerobic growth of L. lactis activated the expression of genes encoding superoxide dismutase, alkyl hydroperoxide reductase, glutathione reductase and NADH oxidase (noxE), which have all been shown to play a role in oxidative stress tolerance (229). Therefore, the elucidation of transcriptional responses to oxygen exposure can help to decipher the mechanisms by which these LAB can handle oxidative stress generated by oxygen exposure. Notably, the oxygen-induced NADH oxidases of Streptococcus mutans were initially proposed to contribute to its H2O2 production (124), but were in subsequent studies also shown to play a prominent role in aerotolerance. This aerotolerance contribution 70 of the NOX function was associated with the alkyl hydroperoxide reductase activity, which is encoded by the ahpF gene that is genetically linked and co-expressed with the nox1 gene (74, 76). A similar relationship between aerotolerance and H2O2 producing enzymes has also been established in L. johnsonii in which the deficiency of the H2O2 producing NFR resulted in a reduced growth rate under conditions encompassing highlevel oxygen exposure. In this study, we analyze the genome-wide transcriptional response of L. johnsonii to oxygen exposure. Amongst the three most upregulated genes under aerobic conditions we found two genes that display strong homology to NADH oxidoreductases (nox-locus). By means of a GUS-reporter assay it could be confirmed that the gene’s promoter activity is controlled (activated) by the presence of oxygen. Although the nox-locus deletion derivative of L. johnsonii had no apparent phenotype with respect to aerobic growth 3 or H2O2 production, the introduction of this deletion in the L. johnsonii background that lacks NFR led to a strain that has completely lost its capacity to produce H2O2. Moreover, the latter strain appeared to be significantly less aerotolerant as compared to the wild-type or either of the single deletion derivatives (NOX- or NFR-deficient strains). These results indicate that the H2O2 producing activities catalyzed by NFR and NOX are important for aerotolerance of L. johnsonii. Materials & Methods Strains and growth conditions of L. johnsonii Lactobacillus johnsonii NCC 533 was obtained from the Nestec Culture Collection and routinely cultured in MRS medium (205) at 37°C under anaerobic conditions, with minimal headspace (unless indicated otherwise). An overview of all strains used in this study is presented in Table 3.1. Erythromycin and chloramphenicol were supplemented at 5 µg /ml when appropriate. For preculturing of NCC 9360 (nfr-, nox-locus) 500 µg/ ml cysteine was added to sequester oxygen from the growth medium. For anaerobic conditions, cells are grown at 37°C in static 15 ml Falcon tubes with minimal headspace. For aerobic conditions, Erlenmeyers were used that have a headspace volume that is minimally 10-fold larger than the growth medium volume, and were agitated on a rotating incubator at 200 rpm. Cells are inoculated at an OD600 of approximately 0.05 in fresh medium. The regular Lactobacillus medium MRS is used (205) or, where indicated, MRS is substituted for LAPTg (20 g/L glucose, 10 g/L yeast extract,(217), 10 g/L bacto peptone, 10 g/L bacto tryptone plus 1 g/L Tween 80). This 71 medium was used instead of the regular MRS-medium for lactobacilli, because the meat extract in MRS was found to interfere with the enzymatic assay for H2O2. Table 3.1: Strains and plasmids used in this study. Strains and plasmids Genotype NCC 533 Wild type L. johnsonii (176) NCC 9337 EmR, Δnox (LJ_1254-LJ_1255), predicted to encode NADH oxidase Chapter 2 NCC 9359 EmR, Δnfr (LJ0548-LJ0549) encoding NADH flavin reductase Chapter 2 NCC 9360 EmR, Δnox, Δnfr (ΔLJ_1254-LJ_1255, ΔLJ_0548-LJ_0549) This study pDP600 a chloramphenicol-resistant version of pG+host9 containing a complete pBluescript array of unique restriction sites (206) pDP893 pDP600- PpgiA - flpSC - LJ_1125 trm This study pNZ5372 CmR, pNZ273 derivative containing the promoterless gusA gene under control of the LJ_1255 promoter (Pnox) This study pNZ273 CmR, promoter-probe vector, containing a promoterless gusA gene (207) pNZ4040 CmR, pNZ273 derivative containing the promoterless gusA gene under control of the epsB40 promoter (Peps) (240, 241) For transcriptome analysis, cultures were inoculated at an OD of 0.026 ± 0.005 and grown under more tightly controlled conditions in continuously stirred vessels with a 400 ml working volume of MRS medium. The cultures were sparged with specific gas mixtures containing 5% CO2 and either no oxygen (0% oxygen, anaerobic) or normal oxygen levels (20% oxygen, aerobic). Cultures were grown at 37°C with continuous mixing (ca. 200 rpm) and pH was maintained at pH 6.5 by automated 4M NaOH titration. Cell densities were determined by measuring the optical density at 600 nm (OD600). RNA extraction, labelling, hybridization and data analysis Cells were harvested at an OD600 of 0.15 ± .03 from 50 ml of culture by cold centrifugation (5’, 2600xg, 4°C). Cell pellets were resuspended in 0.5 ml ice-cold Tris-EDTA buffer and transferred to screw-cap tubes with 0.5 gram zirconium beads (0.1 mm), 0.25 ml acidic phenol, 30 µl 10% SDS and 30 µl 3 M sodium acetate. After mixing, the samples were 72 immediately frozen in liquid nitrogen and stored at -80°C until further use. Cells were disrupted by bead-beating in 3 rounds of 40 seconds in a Savant FastPrep FP120, with in between cooling on ice. Cell debris was removed by centrifugation (20817 x g, 10’, 4°C) and residual phenol was extracted by addition of ice-cold chloroform followed by centrifugation. RNA was isolated using a High Pure RNA Isolation Kit (Roche Diagnostics, Mannheim, Germany). RNA purity and yield was determined by comparison of absorption at 260 and 280 nanometer (Ultrospec 3000, Pharmacia Biotech, Roosendaal, The Netherlands). RNA quality control was carried out using the RNA 6000 Nano Assay in an Agilent 2100 Bioanalyzer (Agilent technologies, Palo Alto, Ca, USA). The Cyscribe Post-labeling kit was used to synthesize cDNA using 5 µg of total RNA, which was subsequently labeled according to the manufacturer’s protocol (Amersham Biosciences, Amersham, UK). Samples in which the CyDye labeled cDNA 3 concentration was below 24 ng/µl were concentrated prior to cDNA synthesis using a Hetovac VR-1 (Heto Lab Equipment A/S, Birkerod, Denmark). A hybridization scheme was designed that allowed duplicate comparisons between the transcriptome profiles of aerobic and anaerobic grown cultures. 60 Oligomer microarrays (Agilent technologies) were used with 12 ± 2.5 probes per gene and 21841 probes in total (GEO accession number GPL18009). These arrays were employed as previously described (173). In short, two differentially labeled cDNAs (300 ng) were mixed (final-volume 25µl), incubated at 95° C for 3 minutes and subsequently cooled to 68° C. To these mixed cDNAs 25 µl Slidehyb#1 hybridization buffer (Ambion, Austin, USA) and 2X Hi-RPM hybridization buffer (Agilent Technologies) were added and 40 µl of the resulting solution was applied on a 8 * 15K slide preheated at 68°C. Slides were hybridized at 65°C, rotating at 10 rpm for 16 hours in an Agilent hybridization oven (Agilent technologies). Subsequently, slides were washed with wash buffer 1 (Agilent technologies) at room temperature for 1 minute and wash buffer 2 (Agilent technologies) at 37 °C. The slides were dried using nitrogen gas and scanned with a ScanArray Express 4000 scanner (Perkin Elmer, Wellesley, MA). Image analysis and processing were performed using the ImaGene Version 7.5 software (BioDiscovery Inc., Marina Del Rey, CA, USA). The microarrays were scanned at different intensities. For each of the individual microarrays the best scan was selected on the basis of signal distribution (combination of a low number of saturated spots and a low number of low signal spots). The data were normalized using Lowess normalization as available in MicroPrep (230). The data were corrected for inter-slide differences on the basis of total signal intensity per slide using Postprep (230). The median intensity of the 73 different probes per gene was selected as the gene expression intensity. CyberT was used to compare the different transcriptomes, taking into account the duplicates (dye swaps) of each of the conditions (231). This analysis resulted in a gene expression ratio and false discovery rate (FDR) for each gene. Differential gene expression values of expression-ratios with FDR values <0.05 were considered to be statistically significant. All microarray data is MIAME compliant and is available in GEO (accession number GSE52876). Hydrogen peroxide measurements H2O2 concentration was determined by addition of 25 µl cell-free culture media to 175 µl reaction mix containing 5 µg/mL horse radish peroxidase and 30 µM phenol red in water. After 5 minutes of reaction, the pH was increased by the addition of 5 µL 1M NaOH, and absorption was determined at 620 nm and compared to a standard curve (232). Deletion mutants: Construction of NCC 9360 (Δnfr, Δnox) The construction of the L. johnsonii single deletion derivatives ΔLJ_1254-LJ_1255 (Δnox, NCC 9337) and ΔLJ_0548-LJ_0549 (Δnfr, NCC 9359) were described in chapter 2. For this study a double deletion derivative that lacks both LJ_1254-LJ_1255 (nox) and LJ_0548-LJ_0549 (nfr) was constructed. To obtain the double deletion derivative, the ery-cassette was removed from NCC 9359 (Δnfr) by the plasmid-borne expression of the Saccharomyces cerevisiae flpgene (flpSC), using pDP893. The construction of pDP893 employed Lactococcus lactis MG1363 as an intermediate cloning host. It was constructed by amplifying the pgiA promoter (PpgiA) using L. lactis MG1363 genomic DNA as a template, with the primers A (ATATATACTAGTACCCTTAAAAGTGTTAGGAG) (ATATATAAGCTTGAGCTCGCTAGCGCATGCTAATTCCTTTCAATTTCTCGC), and B the resulting amplicon was digested with SpeI and HindIII and ligated in similarly digested pDP600 (Cmr, Emr) (206) generating pDP600-PpgiA. The predicted bi-directional terminator positioned between the LJ_1125 and LJ_1126 genes of NCC 533 (TLJ1125) was amplified using genomic DNA of this L. johnsonii strain as a template and the primers C (ATATATAAGCTTTGCCAATGGATAACCAGG) and D (ATATATCTCGAGAATCTCTCTTGGACTTGC). The resulting amplicon was digested with HindIII and XhoI and cloned into similarly digested pDP600-PpgiA, to yield pDP600- 74 PpgiA-TLJ1125. The S. cerevisiae SC288c 2 micron plasmid (233) was isolated and digested with SphI and XbaI, and the 1.46 kb fragment containing the flp gene was isolated and cloned in similarly digested pDP600-PpgiA-TLJ_1125, yielding plasmid pDP893. This plasmid encodes the flp gene under transcriptional control of the pgiA promoter and is followed by the TLJ1125 terminator. Plasmid pDP893 was isolated from L. lactis MG1363 and used to transform NCC 9359. Transformants were cultured in MRS medium supplemented with chloramphenicol, at 32°C for 5 serial passages (permissive temperature for plasmid replication), then plated to isolate single, chloramphenicol resistant colonies, which were subsequently replica plated to confirm erythromycin sensitivity. The selected colony with the required antibiotic resistance phenotype (Cmr, Emr) was cultured in MRS at 37°C for three serial passages, and subsequently plated to identify single colonies that are chloramphenicol 3 and erythromycin sensitive by replica plating on plates containing these antibiotics. The selected strain was designated NCC 9359-FO (FLP-out). NCC 9359-FO was transformed with pDP902 to achieve the deletion of the nox-locus as described in chapter 2. The deletion was confirmed using primers flanking the target regions. The Δnfr, Δnox derivative of L. johnsonii NCC 533 was designated NCC 9360. Construction of GUS reporter plasmids To study the expression driven by the nox-locus promoter, the region upstream the LJ_1255 gene was amplified using genomic DNA isolated from L. johnsonii NCC 533, and the primers E (ATATTGGATCCCCAGTTGATGAAGTTTTGAAATTCG) and F (CATAAGAATTCCACCATGTTTAAAAGTTACTTTGTCGG). The resulting 184 bp amplicon was digested with EcoRI and BamHI and ligated into similarly digested pNZ273 (207) yielding plasmids pNZ5372, which carries a promoterless copy of the β-glucuronidase encoding gusA gene under control of the LJ_1255 promoter (Pnox). Plasmid constructions were performed in the intermediate cloning host Lactococcus lactis and subsequently transformed to wildtype L. johnsonii (NCC 533) and its nfr-locus deletion derivative (NCC 9359). GUS assay GUS-activity was determined in exponentially growing cultures with OD600 of ~0.3 and ~0.6, taken from the aforementioned batch reactors. Cells were pelleted by centrifugation (2600 * g, 5’, 4°C), washed with 50 mM potassium phosphate buffer pH 7,0 and 75 again pelleted by centrifugation (2600 * g, 5’, 4°C). Cells were resuspended in 50 mM potassium phosphate buffer pH 7,0 with 10 mM ß-mercaptoethanol, 1 mM EDTA and 0,1% Triton X-100 (GUS-buffer). Initial sample volumes were adjusted to normalize the amount of cells at a final OD600 of 4. The cell suspensions were transferred to screw cap tubes with 100 mg 0.1 mm zirconium beads and disrupted by bead-beating in three cycles of 20 seconds. Cell debris was removed by centrifugation (21500 * g, 10’, 4°C) and kept on ice. The protein concentration was determined with a microBCA assay (Thermoscientific). Glucuronidase activity was determined in a 96-wells plate, by adding 100 µl of the cell lysate to 150 µl of the aforementioned GUS-buffer (preheated at 37°C). The reaction was initiated by the addition of 5 µl of the substrate, para-nitro-ß-D phenyl-glucuronide (PNP-gluc) dissolved in 50 mM potassium phosphate buffer pH 7,0. The mixture was kept at 37°C and the rate of increase of absorption at 405 nm was used to calculate the specific activity per mg protein per minute. A molar absorption coefficient of 18000 ml/mmol.cm was used. Results Transcriptome comparison of aerobic and anaerobic L. johnsonii cultures. The whole genome transcriptional response of L. johnsonii to oxygen was determined using microarray based transcriptome analysis, comparing early-logarithmic aerobic and anaerobic growing cultures. RNA was isolated from these cultures and whole genome transcriptome profiles were obtained using oligonucleotide based microarrays (see Materials & Methods section for details). Data analysis focused on genes that were significantly regulated by oxygen (FDR<0.05) and displayed a more than 2-fold expression change between the two conditions. Using these criteria, the expression of 45 genes appeared to be repressed whereas the expression of only 6 genes was induced in aerobic cultures compared to anaerobic cultures (see Table 3.2 and Supplementary materials Table S3.1). Regulation of prophage-related genes Of the 45 downregulated genes, 23 belong to the two L. johnsonii prophages Lj928 and Lj965 (234). It was previously reported that these prophages are transcriptionally silent, and in Northern Blot analysis only transcription of the LJ_1454 gene could be detected (235). The transcriptome data confirmed that the prophage gene expression 76 Table 3.2: Transcriptome comparison of aerobic and anaerobic logarithmic cultures of L. johnsonii. All transcripts that were expressed at least two-fold higher under aerobic conditions, with an FDR<0.05 are included. Gene identities are based on the genome annotation (176) or were determined by BlastP alignment. Gene ID Fold regulation FDR Aerobic / anaerobic Annotation LJ_0814 2,4 2,8E-03 Hypothetical protein LJ_1255 2,3 4,52E-05 possible NADH-dependent oxidoreductase1 LJ_1254 2,3 8,98E-05 Possible NADH-dependent oxidoreductase1 LJ_1454 2,2 2,6E-03 Lj928 prophage protein LJ_1615b 2,1 8,6E-03 hypothetical protein LJ_0480 2,1 1,6E-03 Thioredoxin 3 1 homology ascertained with BlastP; E-value < e-40 with NADH-dependent oxidoreductase of L. salivarius NIAS840 is low (9.7 and 7.6-fold below average expression of all genes for Lj928 and Lj965, respectively). Nevertheless, aerobic growth led to a more than 2-fold repression of 20 of the 44 genes that belong to prophage Lj965 and 26 of the 51 genes belonging to prophage Lj928. Notably, the LJ_1454 gene of prophage LJ928, ranks amongst the 6 genes with the most prominent expression difference between anaerobic and aerobic conditions. Despite the overall low-expression of the prophage associated genes under both conditions tested, these genes appear to be subjected to significant environmental control. Aerobic conditions elicit upregulation of thioredoxin but not of thioredoxin reductase One of the 6 genes expressed at a higher level in the presence of oxygen was the thioredoxin encoding gene LJ_0480 (Trx; 2.1 ± 0.0016). Reduced thioredoxins can participate in numerous redox-reactions in cellular processes, including the scavenging of reactive oxygen species. Besides LJ_0480, the L. johnsonii genome encodes a second copy of this gene (LJ_1665) which was also upregulated under aerobic conditions, albeit with a fold-change of 1.93 ± 0.0028 (fdr<.05), explaining why it was not identified using the selection criteria employed (minimum twofold regulation). Oxidized thioredoxins are regenerated by the activity of thioredoxin reductases (TrxR) (110, 236). For several lactic acid bacteria, transcriptional regulation of these TRX/TRXR systems upon oxygen or H2O2 exposure has been reported (113-115). Four different ORF’s in the L. johnsonii genome have significant homology with the Lactococcus lactis thioredoxin reductase TrxB1 (115), i.e., LJ_0852, LJ_0501, LJ_0042 and LJ_1757. However, only 77 the LJ_0852 encoded protein contains the canonical CXXC motif of the thioredoxin reductase catalytic site (110) and none of these thioredoxin reductase homologues displayed a significant degree of differential expression upon the exposure to molecular oxygen. These observations seem to imply that 3 of the 4 TRXR homologues are unlikely to encode functional thioredoxin reductases (lack of the canonical CXXC motif). The fourth TRXR homologue may be a key-component of the thioredoxin-recycling system although its expression appeared to be unaffected by oxygen. Transcription of genes homologous to NADH oxidase is upregulated by oxygen. Two of the genes that were expressed at a higher level under aerobic conditions (LJ_1254 and LJ_1255 2.28 and 2.32 fold, respectively) encode proteins that display significant homology with NADH oxidases (flavin oxidoreductases) from lactobacilli. To further establish the transcriptional regulation of the NOX encoding locus of L. johnsonii (LJ_1254-1255) the putative promoter region of this locus (PLJ_1255; 184 nucleotides upstream of the LJ_1255 start-codon) was cloned upstream of the promoterless gusA reporter gene in the promoter-probe vector pNZ273 (207), resulting in pNZ5372. GUSactivity in cell free extracts from anaerobically and aerobically growing L. johnsonii NCC 533 harboring pNZ5372 was determined. As a constitutive control a L. johnsonii NCC 533 derivative harboring pNZ4040 was employed. This plasmid is a pNZ273 derivative in which the gusA expression is under control of the constitutive PepsB40 promoter (237). The GUS measurements clearly established that the LJ_1255 promoter was approximately 2- to 2.5- fold induced in aerobic growth conditions compared to anaerobic growth conditions, which is in good agreement with the level of induction of the nox-locus assessed by transcriptome analysis (Table 3.3). In contrast, addition of sublethal amounts of H2O2 to the growth medium (0.2 mM) did not induce the activity of the LJ_1255 promoter significantly (results not shown), indicating that the induction of the nox-locus is induced by exposure to molecular oxygen rather than by oxidative stress. A peculiar genetic organization of the LJ_1254-1255 locus LJ_1254 and LJ_1255 are predicted to encode two overlapping polypeptides of 307 and 86 amino acid residues, respectively (Figure 3.1A). The overlap between the two ORF’s encompasses a 13 bp repeat sequence (Figure 3.1B). The LJ_1255 gene in the L. johnsonii NCC 533 clearly has a stop-codon, suggesting that the LJ_1254-1255 locus of L. johnsonii encodes a disrupted pseudogene. By sequencing transcript-derived cDNA 78 Table 3.3: GUS-activity in mmol / mg protein / min determined in aerobic or anaerobic logarithmic cultures with or without 0.2 mM H2O2. In each experiment, GUS-activity was determined in two samples taken during logarithmic phase (OD ~0.3 and OD ~0.6). Depicted are averages of two independent experiments (totaling 4 samples) ± standard deviation. In case of the positive control L. johnsonii + pNZ4040 and wildtype L. johnsonii two samples of one individual experiment are depicted ± standard error. Aerobic batch Anaerobic batch L. johnsonii NCC 533 + pNZ5372 8.14 ± 1.1 3.9 ± 1.2 L. johnsonii NCC 9359 (Δnfr-locus) + pNZ5372 9.3 ± 0.8 2.5 ± 0.4 L. johnsonii NCC 533 + pNZ4040 21.2 ± 0.4 21.3 ± 4.5 L. johnsonii NCC 533 (wildtype) .08 ± 0.01 .03 ± 0.01 sequences, we confirmed the tandem organization of the two genes and their transcription on a single mRNA molecule (results not shown). We assume that LJ_1254 3 may encode an NADH oxidase, whereas the LJ_1255 encoded short protein is unlikely to be functional as such, but may function in conjunction with the LJ_1254 protein. Within the genetically closely related species of the L. acidophilus group, these two genes are poorly conserved and appear to be randomly distributed in strains belonging to different species. Of the 24 genome sequences available for L. gasseri, one encodes a LJ_1254 homologue, whereas the published genomes of L. crispatus and L. delbrueckii have no regions that are homologous to either of the two genes. Only one of the 11 genomes sequences available for L. acidophilus has a homologue of LJ_1254 with a similar length (LBA_1418 in L. acidophilus NCFM). Upstream of this region a smaller gene is found with homology to LJ_1255 (LBA_1421). However, these two genes are interspersed by two other genes, of which one (LBA_1420) encodes a transposase (See Supplementary materials, Table S3.2 and Figure S3.1). In other LAB, LJ_1254 homologs are found in two published L. lactis genomes, whereas the most intensively studied strain of Lactococcus lactis (MG1363) appears to lack this gene. Several of the published L. plantarum strains have genes with homology to LJ_1254; the most intensively studied strain WCFS1 encompasses two genes with significant similarity to LJ_1254 (lp_1350 and lp_0146). All LJ_1254 homologs found in LAB, apart from the copy in L. acidophilus NCFM have an N-terminal extension of approximately 70-residues (total protein length 369 to 391 amino acid residues), which is homologous with the LJ_1255 encoded protein (Figure 3.1 and Supplementary materials, Table S3.2 an Figure S3.1). 79 Figure 3.1: Schematic representation of the nox-locus (panel A). Panel B shows the overlapping region of LJ_1255 and LJ_1254. Highlighted are the startcodon of LJ_1254, the stopcodon of LJ_1255 and the 13 bp-repeat region. Irrespective of the peculiar genetic organization of the LJ_1254-1255 locus in L. johnsonii NCC 533, we decided to consider both genes as part of the NADH oxidase encoding locus (nox-locus) in further work, since both genes are expressed as a single transcript and are homologous to NADH oxidoreductases in other lactic acid bacteria. Consequently, we deleted both genes present in this locus to study its functionality (see below). H2O2 production by the NFR-deficient strain (NCC 9359) indicates the presence of a secondary H2O2 source. Previously, L. johnsonii has been shown to produce substantial amounts of H2O2 under aerobic conditions (38). H2O2 production in L. johnsonii upon exposure to oxygen was shown to predominantly depend on a novel, constitutively expressed NADH-dependent flavin reductase (NFR) encoded by the LJ_0548-549 genes (nfr-locus, chapter 2). Nevertheless, since in many other bacteria, the activity of NADH oxidases has been identified as the main source of H2O2, a role of the oxygen-induced nox-locus in this process cannot be excluded. To evaluate the role of the nfr- and nox-loci encoded activities in the formation of H2O2 in L. johnsonii, NCC 533 (wild type), NCC 9337 (deletion of nox-locus) and NCC 9359 (deletion of nfr-locus), were cultivated under aerobic growth conditions (aeration was elicited by vigorous shaking at 200 rpm) and growth and H2O2 production were monitored. In agreement with earlier observations, both the wild-type strain and its nox-locus deletion derivative produced H2O2 during the 5-6 hours of aerobic growth (up to 0.36 mM), leading to stagnation of growth. The nfr-locus deletion strain NCC 9359 did not produce any H2O2 during the initial hours of aerobic growth, which is also in agreement with earlier observations. However, prolonged aerobic incubation of the nfr-locus deletion strain resulted in recovery of the H2O2 production capacity (Fig. 3.2B), eventually amounting up to 0.23 mM H2O2 80 after approximately 8-9 hours of aerobic growth. This delayed accumulation of H2O2 in the medium, stagnated the growth of this strain (Figure 3.2A). The observed growth stagnation could in all cases be prevented by addition of 0.5 mg/ml catalase to the growth medium, underpinning that the accumulating H2O2 caused the observed growth stagnation (chapter 2). These observations suggest that besides the main H2O2 producing enzyme NFR, a second system is present in L. johnsonii that is involved in H2O2 production, which is postulated to involve the NOX-encoding genes. To confirm the oxygen exposure mediated activation of the nox promoter in the nfrlocus deficient background pNZ5372 (Pnox-gusA) was introduced into the nfr-locus deletion strain NCC 9359. The pNZ5372 derived GusA expression appeared to be 3.6 fold induced in this strain upon its exposure to oxygen (Table 3.3). This finding not only confirms that the nox-locus is also induced by aeration in this strain, but indicates that 3 the oxygen-mediated regulation in this strain is even slightly enhanced as compared to its parent strain NCC 533. Figure 3.2: Growth and H2O2 production of wild type L. johnsonii NCC 533 (square closed symbols), its Δnoxlocus derivative (LJ_1254-LJ_1255; NCC 9337, circular closed symbols) and its Δnfr-locus (LJ_0548-LJ_0549; NCC 9359 square open symbols). Cultures were incubated at 37°C under continuous shaking at 200 rpm. Culture densities were determined by optical density measurement at 600 nm (panel A) and H2O2 concentrations were enzymatically determined by a phenol red assay (panel B). The data present biological triplicate experiments ± standard deviation. The candidate nox-locus of L. johnsonii contributes to H2O2 production and aerotolerance The data presented above are in agreement with a role of the nox-locus in the recovery of H2O2 production in the NFR-deficient background. To evaluate this, a strain lacking both the nox- (LJ_1254-1255; Δnox-locus) and nfr- (LJ_0548-0549; Δnfr-locus) loci was constructed and designated NCC 9360 (Δnox-, Δnfr-locus; Table 3.1). Growth rate, maximal biomass levels and H2O2 concentration during aerobic growth of this Δnox-, Δnfr-locus double deletion derivative was compared to the wild-type strain and 81 to the single Δnox-locus - and Δnfr -locus deletion derivatives. Notably, in contrast to the wild-type strain or its Δnfr and Δnox single deletion derivatives, NCC 9360 (ΔnoxΔnfr) displayed a strongly reduced aerobic growth rate. Aerobic growth of this strain stagnates at an OD600 of 0.3 (±0.01), whereas the other strains continue to grow up to an OD600 of 2.1 (±0.01). Strikingly, the growth stagnation of NCC 9360 is not accompanied by any detectable H2O2 formation (Table 3.4). Moreover, the premature stagnation of growth of NCC 9360 cannot be prevented by the addition of 0.5 mg/ ml catalase (results not shown), confirming that it is not caused by accumulating H2O2. On the basis of these results, it can be concluded that the combined activities encoded by the nox-and nfr- loci plays an essential role during aerobic growth of L. johnsonii. However, these experiments do not conclusively demonstrate that the nox-locus contributes to H2O2 formation in the NFR deficient L. johnsonii strain, since the lack of detectable H2O2 production in NCC 9360 (ΔnoxΔnfr) may be due its incapacity to grow under aerobic conditions. To evaluate the role of the nox-locus in H2O2 formation in the NFR deficient background, the NCC 9360 (ΔnoxΔnfr) was grown anaerobically to mid-exponential phase (OD600 of 0.25 ± 0.04) and suddenly exposed to oxygen by transferring the culture to shake flasks agitated at 200 rpm. Under these conditions, the L. johnsonii wild-type and its single nox-locus deletion derivative initiate H2O2 production immediately or after a delay (Δnfr strain), eventually leading to growth stagnation (see also above). In contrast, although the NCC 9360 strain (Δnox, Δnfr) continued to grow for at least two generations after the switch to aerobic conditions, it did not produce detectable H2O2 levels (the estimated detection limit of H2O2 detection assay is 0.02 mM) (Figure 3.3). These results indicate that the H2O2 producing capacity detected in the NFR-deficient strain upon prolonged oxygen exposure is depending on the LJ_12541255 nox-locus. Table 3.4: Maximal growth rate, optical density (OD600) and H2O2 concentration determined during aerobic growth of L. johnsonii NCC 533 (wild-type), NCC 9337 (Δnox), NCC 9359 (Δnfr) and NCC 9360 (Δnox, Δnfr) in LAPTg. H2O2 levels and OD600 levels were measured after 6 hours of aerobic growth. Data represent the average of triplicate experiments ± standard deviation. 82 Strain Genotype OD600 µmax (h-1) H2O2 (mM) NCC 533 wild type 2.0 (±0.1) 0.74 (±0.00) 0.36 (±0.01) NCC 9337 Δnox 2.3 (±0.1) 0.85 (±0.08) 0.36 (±0.01) NCC 9359 Δnfr 2.2 (±0.1) 0.72 (±0.02) 0.23 (±0.01) NCC 9360 Δnox, Δnfr 0.3 (±0.03) 0.46 (±0.11) <0.02 Figure 3.3: Anaerobic logarithmic cultures were transferred to aerobic shakeflask conditions at mid-logarithmic stage of growth for L. johnsonii NCC 533 (closed square symbols), NCC 9359 (open square symbols) and NCC 9360 (open circular symbols). H2O2 was determined with an enzymatic phenol red method, with an estimated detection limit of ~0.02 mM. Data represent the average of triplicate experiments ± standard deviation. 3 Discussion H2O2 formation by members of the Lactobacillus acidophilus-group, including L. johnsonii, is considered an important metabolic capacity which has been associated with niche adaptation as well as probiotic functions (38, 50, 94). In chapter 2 we presented experimented showing a novel NADH flavin reductase (NFR), that appears to be conserved among the L. acidophilus-group of lactobacilli and is the dominant enzymatic source of H2O2 synthesis in L. johnsonii upon its initial exposure to oxygen. In the present study we show that a second H2O2-producing enzyme is induced in L. johnsonii upon aerobic growth, which is homologous to the NADH oxidoreductase family proteins (NOX). This oxygen-induced NOX allows the recovery of H2O2 production in the NFR deficient L. johnsonii strain, and a double mutant strain that lacks both the NFR and NOX encoding genes is completely defective in H2O2 production but is also severely impaired in aerobic growth. Our results imply that the H2O2 producing capacity of L. johnsonii is essential for its growth under aerobic conditions. This appears to be in clear contradiction with the proposed toxicity of H2O2. Apparently, complete elimination of the H2O2 producing capacity has detrimental consequences for the strain’s capacity to deal with molecular oxygen exposure and to prevent the oxygen-derived molecular damage, presumably by the formation of ROS. The transcriptome comparison of aerobic and anaerobic cultures confirmed our observations with respect to growth and metabolism of L. johnsonii in the presence of oxygen, presented in chapter 2: apart from peroxide production (which induces 83 premature growth stagnation) and the alleviation of the CO2 and acetate dependencies, the presence of molecular oxygen has little effect on metabolic end-products or maximal growth rate of L. johnsonii (chapter 4). It is important to note that several functions known in other bacteria to be induced in response to oxygen exposure are absent from the L. johnsonii genome, including NADH peroxidase, alkyl hydroperoxidase, superoxide dismutase, glutathione reductase and/or catalase. The thioredoxin / thioredoxin reductase (Trx/TrxR) system is the only system known to play a key-role in oxidative stress tolerance in other lactic acid bacteria that appears to be present in L. johnsonii (113-115). Transcription responses to oxygen exposure in this bacterium included the induction of expression of two copies of the thioredoxin gene. However, the potentially enhanced thioredoxin-mediated ROS-scavenging capacity, is apparently not sufficient to prevent premature growth stagnation elicited by the endogenous production of H2O2. Many factors may play a role in the inability of the thioredoxin system to sufficiently scavenge H2O2 or prevent aerobic growth stagnation. NADPH, which is the electron donor for thioredoxin reductases (111), may be rapidly depleted in LAB that are known to generally maintain low NADPH-pools (67). Furthermore, the discrepancy between oxygen-mediated regulation of thioredoxin (induced) and its complementary thioredoxin reductase (unaffected) may contribute to the limited ROS-scavenging capacity in L. johnsonii. The limited responsiveness to oxygen of L. johnsonii and the failure of the thioredoxin system to prevent oxygen induced growth stagnation may reflect the adaptation of this species to its natural habitat of the mammalian intestinal tract, where oxygen levels are very low and only transient exposure to significant amounts of oxygen is expected to be encountered when bacteria reside closer to the mucosal surface of the intestinal wall. Low level or transient oxygen exposure may actually provide a benefit to L. johnsonii by relieving its requirement for exogenous C-1 and C-2 sources for growth (chapter 4). The nox-locus in L. johnsonii consists of two overlapping ORF’s, i.e., a shorter gene (LJ_1255) encoding the N-terminal region of typical NOX enzymes and a longer gene (LJ_1254) encoding the C-terminal region. This could imply that the nox locus of L. johnsonii encodes a pseudogene. However, the locus is expressed as a single transcript and its transcription is elevated by oxygen exposure of the cells, through the activation of its promoter region located upstream of LJ_1255. Moreover, the functionality and H2O2 producing capacity encoded by the nox-locus is supported by the observed H2O2-negative phenotype of the nox-locus deletion derivative of the L. johnsonii strain that lacks the constitutively expressed nfr-locus, which encodes the predominant 84 peroxide producing enzyme in this species. Our results do not elucidate the exact identity of the active NOX protein. Although a complementary role of the LJ_1255 encoded protein cannot be excluded, it seems likely that the activity is encoded by the larger 3’-gene LJ_1254. This suggestion is supported by the observation that the L. acidophilus NCFM genome encodes a full-length homologue of only LJ_1254, which is not directly preceded by a LJ_1255 homologue. NADH oxidoreductases are known to play a central role in aerobic metabolism of facultative heterofermentative lactic acid bacteria: expression of an NADH oxidase in Lactococcus lactis can induce a mixed acid fermentation, by dissipating the cofactor (NADH) for lactate dehydrogenase activity (66). L. johnsonii, is an obligate homofermentative lactic acid bacterium displaying homolactic pyruvate dissipation (chapter 4). Analogously, L. johnsonii does not produce considerable amounts of 3 metabolic end-products other than lactate, irrespective of the presence of oxygen. Nevertheless, NADH consuming enzymes like NFR and NOX could still influence aerobic metabolism by draining the available NADH pool. This could restrict pyruvate dissipation through the lactate forming, NADH consuming step catalyzed by lactate dehydrogenase. Reduced pyruvate to lactate flux could lead to the accumulation of pyruvate, which has previously been reported for L. lactis (48). The accumulation of pyruvate could provide a chemical scavenger for environmental or endogenously produced H2O2, because pyruvate reacts rapidly with this ROS to form acetate. However, in chapter 4, we present experiments that show that aerobic growth of L. johnsonii does not lead to substantial levels of acetate production, suggesting that the pyruvate accumulation and its subsequent reaction with H2O2 is very limited in this species. Thereby the metabolic effects of NADH oxidation in homofermentative LAB like L. johnsonii is limited and not completely understood to date. In this study we establish that the NFR and NOX enzymes are both involved in the capacity of L. johnsonii to produce H2O2, which eventually leads to a stagnation of growth. In contrast with this toxic effect of H2O2, the enzymes responsible for its production also appear to contribute to aerotolerance in L. johnsonii. This apparent paradox resembles the dualistic activity that has been described for alkyl hydroperoxide reductases (AHP), which catalyzes NADH dependent reduction of H2O2 and organic hydroperoxides such as cumene hydroperoxide. In E. coli this enzyme is part of the oxyR induced oxidative stress response (238) and is the primary H2O2 scavenger (109). It consists of two subunits: a larger flavoprotein (AhpF) and a smaller protein (AhpC) that displays homology to thioredoxin reductases (74, 105). Remarkably, the flavoprotein subunit displayed 85 H2O2:NADH oxidase activity in Amphibacillus xylanus, Sporolactobacillus inulinus and in Streptococcus mutans (72, 76, 239). In the latter species it was shown that addition of the smaller AhpC subunit to the AhpF abolished its H2O2 production activity and induced an alkyl peroxide and H2O2 scavenging activity. (74). Although neither the noxlocus nor nfr-locus encoded proteins display significant homology with these two AHPencoding genes, one could hypothesize that these genes function (or used to function) together with a unidentified secondary protein in a comparable peroxidase complex. This would explain the observation that these genes contribute to both H2O2 production and to aerotolerance. In this study we have identified NOX as a secondary H2O2 source in L. johnsonii. Our results indicate that the mode of H2O2 production by L. johnsonii depends on the environmental conditions. In predominantly anaerobic environments in which only transient exposure to (low levels) of molecular oxygen occurs, the production of H2O2 will strictly depend on the NFR activity encoded by LJ_0548-0549. Such conditions may be expected in the typical habitat in which L. johnsonii is encountered, the intestinal tract of mammals. The intestinal lumen is considered anaerobic, but at closer proximity to the strongly oxygenated intestinal mucosa the intestinal bacteria may become exposed to (low levels of) oxygen (40, 41). Therefore, L. johnsonii would initiate the production of H2O2 when residing in close proximity of the epithelial surfaces, which could play an important role in its proposed effects on PPAR-mediated gene expression control (50). Extended exposure of L. johnsonii to oxygen induces the expression of NOX, encoded by LJ_1254-1255, which contributes to the H2O2 production by this species. Conditions encompassing prolonged oxygen exposure may be encountered by industrially applied L. johnsonii strains (e.g., as probiotic cultures) during production, processing and product shelf-life conditions. This illustrates the relevance of understanding the NFR and NOX dependent pathways for H2O2 production in L. johnsonii in relation to the lifestyle of this microorganism. Such knowledge could enable the adaptation of production and storage conditions for probiotic cultures to ensure higher survival of these bacteria, while it can also be employed to enhance its in vivo health-promoting efficacy which may very well depend on the capacity to produce H2O2 in situ close to the mucosa. 86 Supplementary materials Table S3.1 L. johnsonii genes of which the expression was more than twofold lower in aerobic, as compared to anaerobic batch cultures. Gene ID LJ_0058 LJ_0080 LJ_0081 LJ_0082 LJ_0265 LJ_0295 LJ_0296b LJ_0298 LJ_0299 LJ_0300 LJ_0301 LJ_0302 LJ_0326 LJ_0328 LJ_0388 LJ_0573 LJ_0718 LJ_0719 LJ_0720 LJ_0721 LJ_0849 LJ_0890 regulation 0,41 0,20 0,23 0,23 0,49 0,49 0,38 0,44 0,30 0,34 0,38 0,44 0,36 0,48 0,48 0,39 0,28 0,24 0,24 0,39 0,49 0,50 fdr 5,3E-04 9,8E-11 6,5E-11 6,3E-11 1,8E-03 4,8E-03 1,5E-04 1,4E-03 1,8E-04 8,9E-05 7,2E-03 2,4E-03 4,0E-03 2,7E-02 4,3E-04 4,9E-06 4,3E-05 2,5E-07 7,2E-06 3,4E-03 7,5E-04 7,1E-04 LJ_1117 LJ_1185 0,39 0,50 2,5E-06 2,1E-04 LJ_1187 LJ_1276 LJ_1429 LJ_1434 LJ_1443 LJ_1445 LJ_1446 LJ_1447 LJ_1448 LJ_1449 LJ_1450 LJ_1451 LJ_1452 LJ_1452b LJ_1453 LJ_1455 LJ_1549 LJ_1649 LJ_1763 LJ_1764 0,45 0,45 0,42 0,48 0,44 0,39 0,40 0,45 0,38 0,28 0,40 0,42 0,39 0,31 0,38 0,34 0,48 0,46 0,37 0,32 3,5E-05 4,0E-03 2,3E-03 2,1E-03 9,2E-04 4,2E-05 9,1E-05 4,1E-03 1,8E-04 1,7E-06 3,6E-04 8,1E-04 4,1E-04 4,5E-05 5,4E-04 1,6E-05 9,2E-05 1,9E-04 4,5E-03 5,1E-07 LJ_1766 0,39 3,1E-08 annotation (or BLAST homologies) Bile salt transporter Uncharacterized Glycosyltransferase Endoglucanase Y ABC transporter permease component Lj965 prophage antirepressor Lj965 prophage protein Lj965 prophage protein Lj965 prophage protein Lj965 prophage replication protein Lj965 prophage protein Lj965 prophage antirepressor LJ965 prophage protein LJ965 prophage holin Protein translocase subunit SecA 2 Pheromone response surface protein PrgC Dipeptidase Aminopeptidase C Amino acid permease domain containing protein Amino acid permease-associated protein HPr kinase/phosphorylase ABC transporter permease component, Spermidine/ putrescine UPF0324 membrane protein Carbamoyl-phosphate synthase, pyrimidine-specific, small chain Lipoprotein signal peptidase Carbamoyl-phosphate synthase large chain Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Exodeoxyribonuclease 7 large subunit Dephospho-CoA kinase Carboxymuconolactone decarboxylase Carboxymuconolactone decarboxylase Related Protein Aldehyde-alcohol dehydrogenase 3 87 Table S3.2: Homology of LJ_1254 and LJ_1255 in other lactic acid bacteria as determined by BlastP analysis. Species-strain Gene ID; annotation Length (AA) Homology with LJ_1255 Homology with LJ_1254 L. acidophilus NCFM NADH dependent oxidoreductase LBA_1418 306 No homology Query coverage 84%, Identical: 60% residue 1-265 L. acidophilus NCFM Putative NADH dependent oxidoreductase LBA_1421 57 Query coverage 65% Identical 61% residue 1-56 No homology L. gasseri K7 Uncharacterized protein LK7_09396 383 Query coverage 100% Identical: 92% residue 1-86 Query coverage 100% Identical: 85% residue 77-383 L. salivarius NIAS840 NADH-dependent oxidoreductase NIAS840_00317 382 Query coverage 100% Identical: 84% residue 1-86 Query coverage 97% Identical: 78% residue 83-381 L. lactis subsp. cremoris KW 2 NADH-dependent oxidoreductase KW2_1665 391 Query coverage 96% Identical: 49% residue 4-86 Query coverage 90% Identical: 51% residue 86-367 L. plantarum WCFS1 NADH:flavin oxidoreductase LP_0146 369 Query coverage 100% Identical: 30% residue 1-81 Query coverage 89% Identical: 35% residue 77-354 L. plantarum WCFS1 NADH:flavin oxidoreductase 371 Query coverage 73% Identical: 30% residue 6-69 Query coverage 83% Identical: 34% residue 79-339 Figure S3.3: Genetic context of LJ_1254 and LJ_1255 in other lactic acid bacteria, using BlastP and Microbial Genome Viewer. 88 3 89 Chapter 4 Oxygen relieves the CO2 and acetate dependency of Lactobacillus johnsonii NCC 533 Rosanne Y. Hertzberger1,3,4, R. David Pridmore2,4, Christof Gysler2,4, Michiel Kleerebezem3,5, M. Joost Teixeira de Mattos1 1) Molecular Microbial Physiology, Swammerdam Institute for Life Sciences, University of Amsterdam, Science Park, Amsterdam, The Netherlands 2) Nestlé Research Centre, Vers-chez-les-Blanc, Switzerland 3) NIZO food research, Ede, The Netherlands 4) Kluyver centre for Genomics of Industrial Fermentation, The Netherlands 5) Host Microbe Interactomics Group, Wageningen University, Wageningen, The Netherlands Published in PlosONE (2013);8(2):e57235 Summary The probiotic Lactobacillus johnsonii NCC 533 is relatively sensitive to oxidative stress; the presence of oxygen causes a lower biomass yield due to early growth stagnation. We show however that oxygen can also be beneficial to this organism as it relieves the requirement for acetate and CO2 during growth. Both on agar- and liquid-media, anaerobic growth of L. johnsonii NCC 533 requires CO2 supplementation of the gas phase. Switching off the CO2 supply induces growth arrest and cell death. The presence of molecular oxygen overcomes the CO2 dependency. Analogously, L. johnsonii NCC 533 strictly requires media with acetate to sustain anaerobic growth, although supplementation at a level that is 100-fold lower (120 microM) than the concentration in regular growth medium for lactobacilli already suffices for normal growth. Analogous to the CO2 requirement, oxygen supply relieves this acetate-dependency for growth. The L. johnsonii NCC 533 genome indicates that this organism lacks genes coding for pyruvate formate lyase (PFL) and pyruvate dehydrogenase (PDH), both CO2 and acetylCoA producing systems. Therefore, C1- and C2- compound production is predicted to largely depend on pyruvate oxidase activity (POX). This proposed role of POX in C2/ C1-generation is corroborated by the observation that in a POX deficient mutant of L. johnsonii NCC 533, oxygen is not able to overcome acetate dependency nor does it relieve the CO2 dependency. 92 Introduction The Lactobacillus acidophilus group was recognized early as the most prevalent inhabitant of the vaginal microbiota (242, 243) and also as the pioneer bacteria in the developing intestinal microbiota of neonates (16). Various strains and species of the acidophilus group are marketed as functional ingredients in probiotic products, associated with health benefits for the consumer. Therefore, understanding of the physiology of members of this group of lactic acid bacteria is of importance both from a medical and an economical point of view. One of the probiotics belonging to this group is Lactobacillus johnsonii NCC 533, whose genome sequence was published in 2004 (176). Its probiotic functionalities have been explored in detail, including immuno-modulation (169, 182, 183) and pathogen inhibition (171). Additionally, its ability to adhere to the epithelial cell was explored (170, 195). Analogous to many other members of the acidophilus group, L. johnsonii can be considered as a highly auxotrophic species lacking the operons for a range of biosynthetic pathways. The genome of L. johnsonii NCC 533 lacks genes for the synthesis of vitamins, 4 purines, fatty acids and all amino acids (except for the interconversion of L-asparagine and L-aspartate and the interconversion of L-glutamate to L-glutamine) (176, 185). As a consequence, L. johnsonii has fastidious growth requirements. Noteworthy in the context of applicability, the organism does not grow autonomously on milk (244). In addition to the above-mentioned auxotrophies, and analogous to many other closely related species, L. johnsonii may require a source of acetate for growth. C2-compounds are required in many anabolic reactions and acetate-mediated stimulation of growth has been reported for lactic acid bacteria that exhibit a predominant homolactic metabolism on hexose sugars, such as Lactobacillus sakei (245) and Lactobacillus delbrueckii (246). Acetylation of the muramic and glucosamine residues of the peptidoglycan for instance, involves O-acetylation for which a supply of C2 compounds like acetyl-CoA is essential (247). Heterofermentative lactic acid bacteria have the capacity for acetate production, and are therefore assumed to be independent of exogenous acetate addition. However, growth of a Δldh- Lactococcus lactis mutant was reported to be stimulated by acetate which it uses for the conversion to ethanol as a means to regenerate NAD+ in order to rescue its redox balance (248). 93 Another well-described growth requirement is CO2. L. johnsonii, is a so-called capnophilic organism, i.e. it has a requirement for either gaseous CO2 or bicarbonate supplementation for growth, which is a characteristic that is also observed in many other lactic acid bacteria species (249-251). The C-1 source has been proposed to be required for the synthesis of a common intermediate of the pyrimidine and arginine production pathways, carbamoyl-phosphate. In L. plantarum carbamoyl-phosphate can be synthesized from glutamine, ATP and bicarbonate involving two enzymes: pyrimidine-regulated CPS-P (encoded by carAB) and arginine regulated CPS-A (encoded by pyrAaAb) (252). Two regulators of this pathway, PyrR1 and PyrR2 control expression of the pyr-operon in response to pyrimidine and inorganic carbon levels, respectively (253, 254). The genes of the pyr-operon are conserved amongst many lactobacilli, including L, johnsonii NCC 533. Homologues of the argFGH genes for arginine biosynthesis are absent, rendering this species auxotrophic for arginine. The production and consumption of metabolites, like CO2 and acetate, are known to stabilize microbial communities. For example, in yoghurt fermentation, Streptococcus thermophilus and L. delbrueckii show close metabolic relations with the first species providing the second with CO2, acetate, folate, and formate. In exchange, the streptococcal species profits from the proteolytic activities of L. delbrueckii (255). Analogously, it can be anticipated that specific nutritional requirements of microbes play an important role in the composition of the human microbiota. In view of both its industrial potential and its niche in the complex microbial environments where these lactobacilli are generally found, such as the gut, understanding the mechanisms that underlie these growth requirements are important. Growth requirements may be strongly dependent on the growth conditions. For L. johnsonii NCC 533 we observed major differences in growth and viability between aerobic and anaerobic conditions, including a significantly higher viability in the presence of molecular oxygen. This is surprising in view of the observation that L. johnsonii is known to produce H2O2 under aerobic conditions, a compound that is generally assumed to be toxic (38). The study presented here indicates that the anaerobic dependency of L. johnsonii for carbon dioxide and acetate is related to its limited flexibility in pyruvate dissipation pathways, which can be overcome by pyruvate oxidase activity in the presence of oxygen, placing this enzyme in a pivotal position in the central metabolism of L. johnsonii. 94 Materials & Methods Strains and culture conditions Lactobacillus johnsonii NCC 533 was obtained from the Nestlé Culture Collection. NCC 533 plus NCC 9333, a pox-deletion derivative of NCC 533, were routinely cultured in MRS medium at 37°C under anaerobic conditions. Erythromycin was supplemented at 5 µM/ml as required. Anopore growth and micro-colony analyses The AnoporeTM method (inoculation, incubation, imaging) was carried out as described before by den Besten et al. (256). Multiple AnoporeTM inorganic membranes (AnodiscTM) were placed on MRS-agar plates and four dilutions (102-105) of an overnight MRS culture of L. johnsonii NCC 533 were spotted per slide (to ensure that colonies would be physically separated to allow individual quantification). The plates were incubated at 37°C in Oxoid jars that were filled with a defined gas mixture by vacuuming, followed by replacement with the gas mixture of choice, and repeated 3 times at the start of the experiment as well as after every opening of the jar for sampling. Single AnoporeTM slides were removed at different time points and transferred to a microscopic slide 4 with a pronarose layer that contained the cell permeable SYTO9 stain and the cell impermeable propidium iodide stain (baclight live/dead staining, Molecular Probes, Invitrogen). SYTO9 enters all cells but is replaced by propidium iodide whenever the membrane integrity is compromised. For every time point and condition between 20 and 147 microcolonies were randomly selected from one slide and imaged directly without the use of a cover slip or immersion oil. Photographs were taken through both red and green filters with a cooled chargecoupled device camera (Princeton Instruments, SARL, Utrecht, The Netherlands) mounted on an Olympus BX-60 fluorescence microscope. A threshold was applied to create a binary image of the image intensity plots and these were superimposed using ImageJ. An example of such an image is shown in Supplementary materials, Figure S4.1 in which images of several colonies are combined. The colony size and viability were quantified using the ImageJ. The ImageJ plugin ObjectJ was employed to facilitate colony selection. Growth in liquid media and physiological characterization Cells were inoculated at an OD600 of 0.01-0.05 in fresh medium. Growth was monitored 95 in continuously stirred vessels with 400 ml regular MRS medium (205) (for the CO2 dependency experiments) or a chemically defined medium (in case of the acetate dependency experiments). The latter medium was described for Lactobacillus reuteri (257) and has previously been shown to also support growth of L. johnsonii NCC 533. Batches were sparged with specific gas mixtures varying in CO2 and O2 content. Cultures were grown at 37ºC with constant mixing (ca. 200 rpm) and pH was maintained at pH 6.5 by automatic 4M NaOH titration. Cell density was determined by measuring the optical density at 600 nm. Growth rate was determined by fitting an exponential trendline through the data points with a minimal R2 of 0.99. In cases of very slow or no growth a trendline was fitted through the data points of the first five hours of incubation (for instance where chemically defined medium is inoculated in the absence of acetate and oxygen). Construction of the pox-deletion derivative of L. johnsonii NCC533 The deletion of the gene LJ_1853 encoding a predicted pyruvate oxidase enzyme was achieved as described previously (206) with the exception that the plasmid pDP749 was used. In pDP749 the erythromycin resistance gene in pDP600-Ery has been flanked by direct copies of the yeast 2-micron plasmid FLP recombination target sites to facilitate excision of the erythromycin resistance marker by the FLP recombinase. The 5’ region was amplified from L. johnsonii NCC 533 genomic DNA with the primers A (ATATATGAGCTCAGCAAGAACGGCTTCTGC) and B (ATATATGGATCCAGATGCTGCTTCTGGTGC) introducing SacI and BamHI restriction sites, respectively. The amplicon was SacI-BamHI digested and cloned in similarly digested pDP749, yielding an intermediate plasmid. The 3’ region was amplified using the primers C (GTGAACGGCACCAGGACC) plus D (ATATATGGTACCGAAGCATATATTGGGGTC), the amplicon obtained was PstI-KpnI digested and cloned in similarly digested intermediate plasmid to yield the pox-deletion plasmid pDP887. Plasmid pDP887 isolated from Lactococcus lactis was used to transform NCC 533 (185) and loop-in/loop-out gene replacement was achieved as described previously (206). The deletion was confirmed by PCR analysis. Organic acids measurement by HPLC Extracellular metabolite concentrations were determined as described previously (208) using HPLC (LKB and Pharmacia, Oregon City, OR, USA) fitted with a REZEX organic acid analysis column (Phenomenex, Torrance, CA, USA) at 45°C and a RI 1530 refractive 96 index detector (Jasco, Easton, MD, USA). The mobile phase consisted of a 7.2 mM H2SO4 solution. Chromatograms were analysed using AZUR chromatography software (St. Martin D’Heres, France). Results CO2 dependency of L. johnsonii NCC 533 during aerobic and anaerobic microcolony growth. To study CO2 dependency of L. johnsonii, we used a high-resolution and quantitative technique by using AnoporeTM slides to visualize growth on plates that were placed in jars with a controlled atmosphere. This allowed for the rapid assessment of growth requirements and in combination with time-resolved microscopic inspection at using live/dead staining, enabled the generation of additional data related to the organisms’ physiological state, viability and population heterogeneity (256, 258, 259). This set-up was employed to evaluate growth and viability during CO2 limitation under aerobic and anaerobic conditions. To this end, AnoporeTM slides on MRS-agar plates were inoculated with different dilutions of cells and incubated in jars filled with gasmixtures varying in CO2 and O2 content. At regular intervals the viability and size of the 4 colonies were determined using a live/dead baclight stain as described in Materials & Methods. The sum of the propidium iodide stained pixels and the SYTO9 stained pixels was used to estimate the size of the colony. The fraction of SYTO9 over all stained pixels was used as a relative measure of viability. CO2 supplementation to the gas phase (5%) was found to stimulate growth under both aerobic (air) and anaerobic (N2) conditions. When plates were transferred to a CO2 depleted environment, growth stagnated after 7 hours, both in aerobic and anaerobic conditions. In the presence of supplemented CO2, microcolonies continued growth with an estimated growth rate of 0.79 h-1 in the anaerobic, and 0.74 h-1 in the aerobic environment, which is comparable to growth rate in liquid culture (data not shown). This growth rate was estimated by fitting an exponential trend line through the average colony size (Figure 4.1, panels A and B). Growth stagnation was accompanied by loss of membrane integrity observed in microcolonies that are grown without CO2 supplementation, whereas microcolonies grown in CO2 supplemented environments sustained viability above 90% throughout the experiment (Figure 4.1, panels C and D). 97 Figure 4.1. L. johnsonii NCC 533 is grown on AnoporeTM slides that are transferred from a 2 hour preincubation period in an N2+5% CO2 environment to environments that vary in CO2 and O2 content. Average size of microcolonies grown aerobically (A) and anaerobically (B) and average viability of microcolonies grown aerobically (C) and anaerobically (D). Growth after the pre-incubation was either in the presence (closed symbols) or absence (open symbols) of 5% CO2. Data shown are the mean of all colonies counted for that time point and condition ± standard deviation. Notably, microcolonies grown in aerobic atmosphere displayed reduced loss of viability albeit with a higher degree of heterogeneity, as compared to microcolonies grown in a nitrogen atmosphere (Figure 4.1 C and D). This observation was remarkable since it has been documented that L. johnsonii produces H2O2 in the presence of oxygen (38), which was presumed to reduce growth rate and induce considerable cell death under aerobic conditions. Taken together, these results suggest that CO2 depletion leads to loss of membrane integrity and growth stagnation, while oxygenation appears to support extended viability as compared to anaerobic conditions. CO2 dependency of L. johnsonii NCC 533 during aerobic and anaerobic liquid growth. To consolidate the results obtained with the Anopore system in the more routinely employed liquid culture conditions, L. johnsonii NCC 533 was grown in a pH-controlled stirred batch culture, sparged with predefined gas mixtures at a rate of 750 ml/min. When L. johnsonii was grown in MRS medium in this experimental setup, a clear difference between aerobic and anaerobic growth was observed. Anaerobic and aerobic cultures reached an exponential growth rate of 0.85 h-1 and 0.69 h-1, respectively. After 6 hours 98 of incubation aerobic growth strongly slowed down and eventually the culture entered stationary phase, whereas the anaerobic culture continued growth (Figure 4.2A). The aerobic growth stagnation was related to the accumulation of H2O2 in the extracellular growth medium, as is evidenced by the complete prevention of the growth stagnation by the addition of 0.5 mg/ml catalase to the medium (Chapter 2). Figure 4.2 Growth in stirred pH-controlled batch cultures sparged by N2 + 5% CO2 (closed symbols) or N2 + 20% O2 + 5% CO2 (open symbols) as measured at OD600. In panel B, the gas regime was switched after 3 hours of exponential growth from a CO2-rich gas to a CO2-free gas: N2 (closed symbols curve), N2 + 20% O2 (open symbols). The switch is indicated by the dashed line. Growth curves are representative of at least triplicate experiments. 4 To assess the influence of CO2 on growth in these conditions, cultures were grown until early-logarithmic phase of growth while sparging a defined gas composition, aerobic (75% N2, 20% O2 and 5% CO2) or anaerobic (95% N2 and 5% CO2). Subsequently sparging was switched to a CO2-free gas mixture. Depletion of CO2 in anaerobic cultures resulted in growth stagnation and initiation of cell death within one hour (Figure 4.2B), whereas in aerobic cultures this effect was not observed and growth continued until it stagnated at a final OD of approximately 1.5, due to the accumulation of H2O2. Overall, these data show that oxygen supplementation in the gas phase relieves the CO2 requirement for growth, both on solid, as well as in liquid media. Oxygen overcomes the acetate dependency of L. johnsonii NCC 533 In addition to CO2 dependency, growth of many lactobacilli also depends on the presence of acetate in the growth medium (246). L. johnsonii was unable to grow in chemically defined medium without acetate supplementation. Notably, the addition of as little as 12 µM sodium acetate (1/1000 of the regular sodium acetate concentration in the chemically defined medium) allowed for recovery of growth, albeit at a slower rate and yielding lower final biomass concentrations. Acetate supplementation at a 100- 99 fold lower level as compared to its regular concentration in CDM (120 µM) completely restored normal anaerobic growth (Figure 4.3). These results show that although there is a strict acetate-requirement for growth, this requirement is already fulfilled with concentrations that are substantially below the levels that are normally added to typical Lactobacillus-laboratory media, such as MRS or CDM. \ Figure 4.3. Growth of L. johnsonii NCC 533 in a chemically defined medium with varying concentrations of sodium acetate: 12 mM as in standard CDM (closed square symbols) 120 µM (round symbols), 12 µM (triangular symbols) and without any Na-acetate supplemented (open square symbols) in stirred pH controlled cultures sparged with N2 + 5% CO2 at a rate of 500 ml/min. The growth curves are the average of duplicate experiments ± standard error of the mean. To assess whether the acetate requirement of L. johnsonii NCC 533 depended on the growth conditions, the strain was grown in chemically defined medium with or without acetate supplementation (12 mM), under aerobic or anaerobic conditions. Analogous to what was observed with respect to the CO2 dependency, anaerobic growth of L. johnsonii NCC 533 depended more strictly on acetate supplementation as compared to aerobic growth, which could be sustained without an external acetate source, albeit with a slower growth rate and a lower final biomass yield (Figure 4.4). This implies that the endogenous production of acetate under these conditions may be expected to be in the same range as the 12 µM that allowed similar growth restoration under anaerobic conditions (see above). Both aerobic and anaerobic growth of L. johnsonii in chemically defined medium with 12 mM or 120 µM of acetate were analyzed with respect to acetate metabolism: significant change in extracellular acetate were not detected by HPLC analysis nor by a highly specific and sensitive acetate kinase/pyruvate kinase assay (results not shown). This result is likely caused by analytical limitations that did not allow detection of the minute amounts of acetate that are required to sustain growth under these conditions, 100 (estimated detection limit in spent medium is 200 µM). Figure 4.4 Growth of L. johnsonii NCC 533 in a chemically defined medium with 12 mM Na-acetate (square symbols) and without 12 mM Na-acetate (round symbols) in stirred pH controlled cultures sparged with N2 + 5% CO2 (closed symbols) or N2 + 20% O2 + 5% CO2 (open symbols) at a rate of 500 ml/min. Data are average of independent triplicate experiments ± standard deviation. In most organisms acetyl-CoA functions as the central C2-intermediate in several biosynthetic pathways. This metabolite can be produced from pyruvate by reactions 4 catalyzed by pyruvate dehydrogenase (PDH) or pyruvate formate lyase (PFL). However, apart from a homologue for one subunit of pyruvate dehydrogenase, the corresponding genes appeared to be absent in the L. johnsonii NCC 533 genome (176). This genotype is shared with the other members of the L. acidophilus-group (see Supplementary materials, table S4.1A and S4.1B), indicating that these species lack the capacity for autonomous acetyl-CoA production from their central energy metabolism (Figure 4.5). The L. johnsonii genome does encode an enzyme that could provide the cell with both CO2 and acetate, namely pyruvate oxidase (POX). POX catalyzes a reaction that requires molecular oxygen as a co-substrate, and therefore its activity may directly explain the observed physiological consequences of the presence of oxygen (aerobic growth is independent of an external CO2 and acetate source). Therefore, we hypothesized that oxygen availability relieves the CO2 and acetate dependency by the pyruvate oxidase derived supply of both these metabolites. 101 Figure 4.5: Simplified overview of pyruvate metabolism in L. johnsonii. LDH: Lactate dehydrogenase. POX: pyruvate oxidase. ACK: Acetate kinase. PAT: Phosphate acetyltransferase. Acetate and CO2 dependency of a pox-deletion mutant. To test the proposed hypothesis, a pox deletion derivate that lacks the pyruvate oxidase encoding gene was constructed. Under anaerobic condition in an atmosphere supplemented with 5% CO2, the growth rate of the mutant in MRS was similar to that observed for NCC 533. Moreover, under these conditions the wild-type and its poxdeletion derivative displayed a comparable growth arrest upon CO2 depletion. However, under aerobic conditions, shutting down the 5% CO2 supply elicited rapid growth stagnation of the pox mutant (Figure 4.6), which is in clear contrast to the wild-type that continues to grow under these conditions. Clearly, the deletion of pox resulted in a L. johnsonii mutant that depended on exogenous CO2 supplementation for aerobic growth. This fully supports the proposed pivotal role of the pox-encoded pyruvate oxidase enzyme in the generation of this essential C1-source under these conditions. Analogous to the CO2 supply provided by the POX-pathway under aerobic conditions, it would be expected that this pathway also provides an acetate supply when oxygen is available. Consequently, the pox mutant would be expected to be more hampered aerobically in media that lack exogenous acetate as compared to the wild-type strain. Generally, the pyruvate oxidase deficient mutant displayed slower growth rates than the wild type, independent of the presence of oxygen (Figure 4.7A). However, growth of the pox mutant in the absence of acetate differed considerably, i.e., the typical 102 oxygen relief of the acetate dependency that was observed for the wild type was not observed for the pox mutant (Figure 4.7B), which supports our hypothesized role of pyruvate oxidase in generating C2-compounds. Figure 4.6 Growth of the NCC 533 (closed symbols) and NCC 9333 (open symbols) as measured at OD600 in stirred batch cultures sparged with N2 + 20% O2 + 5% CO2. The gas regime was switched after 3 hours of exponential growth to N2 + 20% O2. Data are the average of quadruple independent experiments ± standard deviation. 4 Figure 4.7 Growth rate of L. johnsonii NCC 533 in the standard chemically defined medium with (panel A) and without 12 mM Na-acetate (panel B) in stirred pH controlled aerobic batch cultures (open bars) or anaerobic batch cultures (closed bars). Growth rates were determined as explained in Materials & Methods. Data are average of triplicate experiments (panel A) and duplicate experiments (panel B) ± standard error of the mean. 103 Discussion Lactobacillus johnsonii is generally described as an anaerobic fastidious lactic acid bacterium. Fastidious because its growth is dependent on supplementation of various nutrients to its growth medium, and anaerobic because oxygen cannot be used for respiration. Moreover, L. johnsonii produces H2O2 when grown under aerobic conditions, which inhibits growth. Here we present an example that auxotrophy can be dependent on external conditions that seemingly are not related to the nutrient requirement: we show that anaerobicity actually exacerbates the fastidious nature of L. johnsonii NCC 533 since the presence of oxygen is shown to relieve at least two of its anaerobic growth requirements, i.e., the requirement for acetate and CO2. Both on plates and in liquid culture, L. johnsonii showed clear CO2 dependent growth. However, the oxygen relief of this dependency was more apparent in liquid culture than on solid medium, as illustrated by the observation that aerobic growth on plates without CO2 still resulted in smaller colonies and reduced viability. In contrast, these CO2 dependent phenotypic differences were completely abolished by oxygen supplementation in liquid culture. One explanation for the observed difference could be found in the ambient pH, which is controlled at 6.5 in liquid culture and is uncontrolled in the Anopore experiment. It should be noted in this context that pH influences the equilibrium between the different dissolved carbonic species; CO2 dissolves in water as H2CO3 (pKa 6.1) and the latter species may be deprotonated in a pH dependent manner to generate HCO3- and CO32-, respectively. Thus, lower pH values shift the equilibrium resulting in release of CO2 from the solution. It is to be expected that on solid media especially the local pH within the direct environment of emerging microcolonies drops substantially below 6.1 due to lactic acid production. These micro-scale differences in environmental conditions experienced by bacteria grown in microcolonies versus liquid cultures may explain the observed CO2 dependency differences observed. Like the other species in the L. acidophilus-group (L. delbrueckii, L. gasseri, L. johnsonii, L. crispatus, L. amylovorus, L. helveticus), the genome of L. johnsonii lacks two major systems for the production of C2- and C1-compounds, namely the pyruvate dehydrogenase complex (PDH) and pyruvate-formate lyase (PFL) producing acetyl –CoA (Supplementary material, table S.1A and S4.1B). Instead, the genomes of these species all encode the pyruvate oxidase gene that can provide a metabolic source of C2-compounds whenever molecular oxygen is available for the POX reaction. The primary habitat of L. johnsonii is considered to be the intestine, which is a predominantly anaerobic environment and 104 would therefore not support POX mediated C2-production. However, in close vicinity to the mucosal tissues, local and a steep oxygen gradient may be encountered (41) that may allow for the POX-mediated contribution to metabolism. Notably, preliminary transcriptome studies of L. johnsonii grown under anaerobic, aerobic and CO2 depleted conditions did not reveal regulation of the pox gene expression, suggesting that the enzyme is constitutively expressed. Based on the physiological observations both on plate and in liquid culture, combined with the absence of these genes, we hypothesized that pyruvate oxidase activity would play a pivotal role in the acetate and CO2 supply for the cell. Indeed, a pox-deletion derivative of L. johnsonii did not display a higher growth rate under aerobic conditions in the absence of acetate, such as observed in the wild type strain. Moreover, whereas the wild type strain continued to grow upon a switch to CO2 depletion, growth of the mutant stagnated at a lower biomass concentration. The observed time lapse between the onset of flushing with CO2 free gas and the actual CO2 depletion of the system is most likely due to the slow removal of all carbonic species at a pH higher than 6.1 (the pKa of carbonic acid). Both results show that, in contrast to the wild type, the pox-mutant has lost the ability to aerobically generate CO2 and acetate. This corroborates the proposed role of pyruvate oxidase in the generation 4 of C1 and C2 metabolic intermediates. It was observed that the pox mutant has a lower growth rate, both aerobically and aerobically. Although it can be argued that under aerobic conditions the pox gene might play a role in protection against its reaction product, H2O2 by allowing for a faster production rate of ATP via the production of acetyl-phosphate and subsequent generation of ATP by acetate kinase (91), this argument does not hold for anaerobic growth conditions. So far, no specific role for POX under these conditions can be brought forward and the cause of the effect of the deletion on growth remains to be elucidated. The major dependency of L. johnsonii on pyruvate oxidase for the supply of these compounds was rather unforeseen since many other pathways are known and present in L. johnsonii that can render CO2 and acetate. Phosphoketolase, for instance, catalyzes the deacetylation of xylulose-5-phosphate which yields acetyl-phosphate. Similarly, CO2 can be produced through decarboxylation of amino acids, oxaloacetic acid and phosphopantotenoyl. However, acetate and CO2 are both required for growth of L. johnsonii in the absence of oxygen, even though very low concentrations of acetate (<120µM) already suffice for growth. This suggests that the flux through these pathways compared to pyruvate oxidase is marginal. 105 It is uncertain, however, that the lactobacilli that do possess PDH and PFL encoding genes (Supplementary materials, Table S4.1A and B), can actually employ these pathways for the synthesis of C1 and C2-compounds under aerobic conditions. Literature suggests that L. plantarum does not possess a functional pyruvate dehydrogenase pathway, since acetate production does not require CoA and is not hampered by PDH-inhibitors like arsenate (260, 261). In addition, pyruvate formate lyase activity has been reported to be highly oxygen sensitive and is only considered active under anaerobic conditions (262). The presence of genes predicted to encode PFL or genes that resemble the PDH-genes of other organisms does not preclude that a species still depends on pyruvate oxidase under aerobic conditions for the production of C2 and C1 components, analogous to what we concluded for L. johnsonii. Clear data to support this hypothesis are lacking, although CO2 dependency of L. plantarum was also reported to cause a characteristic growth stagnation under aerobic conditions (250). In addition, another study showed that a pyruvate-oxidase deficient mutant of L. plantarum is hampered in its acetate production capacity (83, 84), supporting the role of this enzyme in aerobic acetate supply in lactobacilli that have a broader genetic arsenal. The effect of deletion of pox in L. johnsonii confirms the role of POX in the generation of both C1 and C2 sources (CO2 and acetate) required for growth. However, a byproduct of pyruvate oxidation by POX is H2O2, of which the accumulation induces oxidative stress that leads to premature growth arrest under aerobic conditions (38). This brings us to the conclusion that oxygen appears to both benefit and harm L. johnsonii. Under aerobic conditions, clearly, a lower biomass yield is reached (chapter 2, Figure 2.1) on the one hand, presumably as a consequence of H2O2 production. On the other, our data also establish clearly that oxygen can increase the metabolic capacity of the strain, relieving some of its fastidious growth requirements. These opposing consequences of oxygen presence suggest that a micro-aerobic environment may be optimal for growth of L. johnsonii NCC 533. Here we have refined the metabolic requirements of L. johnsonii NCC 533 and pinpointed the pivotal role of the pox gene in the requirement for C1 and C2 sources. These findings can provide novel clues for the optimization of growth conditions of these commercially relevant microbes, and may in more general terms facilitate a more efficient regime for the production of probiotics belonging to this group of lactobacilli. 106 Acknowledgements We would like to thank the Molecular Cytology group at SILS, University of Amsterdam for letting us use the BX fluorescence microscope and in particular Norbert Vischer for the assistance with ObjectJ. More information on ObjectJ can be found at http://simon. bio.uva.nl/objectj 4 107 Supplementary materials Figure S4.1 Superimposed image (Composite picture) of baclight-stained microcolonies in which images of colonies after 7 hours of growth in environments that vary in oxygen and CO2 content are grouped. Images were thresholded, colors were assigned artificially and superimposed as described in Materials & Methods. 108 lar_0608 lvis_1410 reuteri DSM 20016 brevis ATCC 367 lvis_1409 lar_0609 lsei_1306 lrhm_1267 lrhm_1266 lsei_1305 rhamnosus GG lsl_0154 lca_1084 lsl_0153 lca_1085 salivarius UCC118 sakei subsp. Sakei 23K casei ATCC 334 lp_2153 lp_2154 plantarum WCFS1 pdhB pdhA Lactobacillus PDH complex lvis_1408 lar_0610 lsei_1307 lrhm_1268 lca_1083 lsl_0155 lp_2152 pdhC lvis_1407 lar_0611 lsei_1308 lrhm_1269 lca_1082 lsl_0156 lp_2151 pdhD - - - - - lhrm_1365 lsei_1410 lhrm_1366 lsei_1412 - lca_0973 lsl_1873 lca_0974 lsl_1872 lca_0973 lp_2596 lp_3313 lp_3314 pflE pflB pflA PFL - - - - lca_0974 - lp_2598 pflF Table S4.1A: Presence of genes for pyruvate dehydrogenase or pyruvate formate lyase in lactobacilli. Overview of pyruvate dehydrogenase and pyruvate formate lyase encoding gene prevalence in lactobacilli (Table A) and in species belonging to the L. acidophilus group (Table S4.1B). If no gene was found, a BLAST search was performed using the protein sequence of the homologue in L. plantarum WCFS1. 4 109 Table S4.1B - Lactobacillus acidophilus group PDH complex Lactobacillus johnsonii NCC 533 pdhA - PFL pdhB pdhC pdhD lj_1267 - (88% 3e ) - lba_1490 pflB pflE pflF - - - - - - - - (95% 2e ) -17 1 acidophilus NCFM lj_1757 pflA -61 2 - (87% 1e21)1 lba_1220 (96% 1e-65)2 helveticus DPC 4571 - - - lhv_1961, (94%, 3e-66)2 - - - - delbrueckii ATCC 11842 - - - ldb_0759 - - - - - - - - (95% 3e-66)2 gasseri ATCC 33323 - - - lgas_1554 (95% 3e-61) 2 1 BLAST homology to a transketolase 2 BLAST homology to a pyrimidine-dinucleotide oxidoreductase 110 4 111 Chapter 5 Genome-wide transcriptome response to CO2 depletion in Lactobacillus johnsonii Summary Lactic acid bacteria belonging to the Lactobacillus acidophilus-group, including Lactobacillus johnsonii, require a wide variety of nutrients for growth, such as amino acids, vitamins, fatty acids, acetate and CO2. In chapter 4, we showed that CO2 depletion during growth of L. johnsonii not only causes growth stagnation but also results in cell death. To study the molecular events following CO2 exhaustion, we followed the transcriptional changes in anaerobically growing cultures that were exposed to a a CO2 depleted condition. The transcriptome analysis revealed an extensive rearrangement of gene expression profiles, with 343 genes that were more than twofold induced or repressed after 120 minutes of CO2 depletion, including substantial rearrangements of the expression of metabolic functions. These encompassed the strong upregulation of the carbamoyl-phosphate synthesis pathway (up to 17-fold). This pathway utilizes CO2, aspartate and glutamine as substrates in the biosynthesis of pyrimidines, suggesting that intracellular depletion of the pyrimidine pool may be a direct consequence of CO2 depletion. However, supplementation of the MRS growth media with pyrimidines could not alleviate CO2 induced growth stagnation. Moreover, addition of several other metabolites of which the synthesis is associated with CO2 dependency in other lactic acid bacteria (arginine and aspartate) could also not complement the CO2 requiring phenotype, leaving the mechanism behind CO2 dependency to be determined. 114 Introduction Lactic acid bacteria (LAB) are Gram-positive bacteria that produce lactic acid as their main metabolic end-product. Several LAB have been recognized to be capnophilic, which means that their growth is stimulated by supplementation with inorganic carbon in the form of bicarbonate, gaseous CO2, formate or ureic acid (249-251). In L. plantarum, the pyr-encoded uridine monophosphate biosynthesis pathway has been shown to be related to its inorganic carbon requirement. This pathway incorporates ammonia (from glutamine), phosphate and CO2 to synthesize carbamoyl-phosphate, which is the precursor for arginine and pyrimidine nucleotides. The inorganic carbon requiring isolates of L. plantarum depend on exogenous pyrimidine and arginine for growth in environments that are low in inorganic carbon (252-254, 263). Aspartate biosynthesis is another metabolic pathway that has been shown to contribute to CO2 requirements in LAB. The enzyme pyruvate carboxylase uses CO2 to produce the aspartate precursor, oxaloacetate (251). Correspondingly, supplementation with either aspartate or citrate (which LAB can convert to oxaloacetate) could complement the CO2 dependent phenotype (264). Additionally, an older report suggests that oleic acid (Tween 80) could also be used to complement growth of certain lactobacilli in a CO2 deficient environment (265). Along with the metabolic pathways that use inorganic carbon as a substrate, several other biological processes may be affected by the level of environmental CO2. Pathogenic bacteria are known to modulate virulence related genes in response to CO2 5 and bicarbonate (266-268). In L. plantarum protein levels other than those belonging to the pyr-regulon undergo significant modulation by elevated CO2 levels, including for example genes associated with purine synthesis (269). Although arginine and uracil supplementation suffice to complement the CO2 requirement of L. plantarum, the supplementation with inorganic carbon could still further stimulate its growth, even in the presence of these compounds (270). These results indicate that a third unidentified metabolic pathway requires CO2 as a substrate. Lactobacillus johnsonii, a bacterium that is marketed as a probiotic, and is recognized for its immunomodulatory properties (182, 183) and pathogen inhibition (38) was also shown to require CO2 for growth. The removal of gaseous CO2, from an exponential growing culture by flushing with dinitrogen gas resulted not only in rapid growth stagnation but was accompanied by extensive cell death (chapter 4). Interestingly, low level exposure to molecular oxygen could overcome this CO2 dependency, while it also 115 relieved the acetate-requirement for growth. These observations were explained by a central role of the pyruvate oxidase pathway in the production of these compounds, as was supported by the observation that a pox deletion derivative of this species remained CO2 and acetate dependent, irrespective of the presence of molecular oxygen. L. johnsonii has many auxotrophies, and is routinely grown in rich and complex media like MRS (205), which are assumed to contain abundant levels of pyrimidines and amino acids (176). Nevertheless, even under these rich conditions CO2 depletion elicited growth stagnation and cell death, which we study here at the molecular level by analyzing the transcriptome pattern changes over time following CO2 depletion. Despite the rich media employed, CO2 depletion elicited a major adaptation of the expression of the carbamoyl-phosphate pathway in L. johnsonii. However, unlike L. plantarum the CO2 dependency of L. johnsonii could not be complemented by exogenous nucleotide supplies, suggesting that CO2 may have an additional role in L. johnsonii. Materials & Methods Strain & cultivation Lactobacillus johnsonii NCC 533 was obtained from the Nestlé Culture Collection and was routinely cultured in MRS medium at 37°C under anaerobic conditions. Growth in liquid and solid MRS media Cells were inoculated in fresh MRS medium (205). Growth was monitored in continuously stirred vessels with 400 ml working volume. Batches were sparged at 750 ml/min with either a gas mix containing 5% CO2 and 95% nitrogen gas or pure nitrogen gas. Cultures were grown at 37°C with constant mixing (ca. 200 rpm) and pH was maintained at pH 6.5 by automatic 4M NaOH titration. Cell density was determined by measuring the optical density at 600 nm (OD600). Viable counts were determined by placing 2 µl drops of culture dilutions on MRS plates in triplicate and incubating these plates overnight in a closed container in the presence of an AnaerogenTM bag. Cultures of L. johnsonii were grown overnight in MRS medium (>16 hours) and serially diluted in triplicate in fresh MRS medium. Drops of 2 µL from each dilution and each replicate were placed on the agar plates. Plates were incubated at 37°C in air-tight jars in CO2-rich or CO2-depleted conditions. Such conditions were achieved by applying three consecutive cycles of vacuum (0.8 bar) and refilling with either 100% N2 gas (CO2 free) or 95% N2 + 5% CO2 gas (CO2 rich). 116 Transcriptome analysis Cells were harvested at different time points: 0 minutes (prior to closing of the CO2 feed) and 30, 60 and 120 minutes after closing the CO2 feed. The first sample was taken at OD600 0.13 ± 0.017 (inoculation level 0.025 ± 0.008, i.e. more than two duplications after inoculation level), and cells were harvested from 50 ml of culture by cold centrifugation (5’, 2600xg, 4°C). Sample volumes of the samples taken at later time points were adjusted in order to obtain a similar amount of biomass for RNA isolation. Cell pellets were resuspended in 0.5 ml ice-cold Tris-EDTA buffer and transferred to screw-cap tubes with 0.5 gram zirconium beads (0.1 mm), 0.25 ml acidic phenol, 30 µl 10% SDS and 30 µl 3 M sodium acetate. After mixing, the samples were immediately frozen in liquid nitrogen and stored at -80°C until further use. Cells were disrupted by bead-beating in 3 rounds of 40 seconds in a Savant FastPrep FP120, with in between cooling on ice. Cell debris was removed by centrifugation (20817 x g, 10’, 4°C) and residual phenol was extracted by addition of ice-cold chloroform followed by centrifugation. RNA was isolated using a High Pure RNA Isolation Kit (Roche Diagnostics, Mannheim, Germany). RNA purity and yield was determined by comparison of absorption at 260 and 280 nanometer (Ultrospec 3000, Pharmacia Biotech, Roosendaal, The Netherlands). RNA quality control was carried out using the RNA 6000 Nano Assay in an Agilent 2100 Bioanalyzer (Agilent technologies, Palo Alto, Ca, USA). The Cyscribe Post-labeling kit was used to synthesize cDNA using 5 µg of total RNA, which was subsequently labeled according to the manufacturer’s protocol 5 (Amersham Biosciences, Amersham, UK). Samples in which the CyDye labeled cDNA concentration was below 24 ng/µl were concentrated prior to cDNA synthesis using a Hetovac VR-1 (Heto Lab Equipment A/S, Birkerod, Denmark). A hybridization scheme was designed that allowed duplicate comparisons between the transcriptome profiles of aerobic and anaerobic grown cultures. 60 Oligomer microarrays (Agilent technologies) were used with 12 ± 2.5 probes per gene and 21841 probes in total (GEO accession number GPL18009). These arrays were employed as previously described (173). In short, two differentially labeled cDNAs (300 ng) were mixed (final-volume 25µl), incubated at 95° C for 3 minutes and subsequently cooled to 68° C. To these mixed cDNAs 25 µl Slidehyb#1 hybridization buffer (Ambion, Austin, USA) and 2X Hi-RPM hybridization buffer (Agilent Technologies) were added and 40 µl of the resulting solution was applied on a 8 * 15K slide preheated at 68°C. Slides were hybridized at 65°C, rotating at 10 rpm for 16 hours in an Agilent hybridization oven (Agilent technologies). Subsequently, slides were washed with wash buffer 1 117 (Agilent technologies) at room temperature for 1 minute and wash buffer 2 (Agilent technologies) at 37 °C. The slides were dried using nitrogen gas and scanned with a ScanArray Express 4000 scanner (Perkin Elmer, Wellesley, MA). Image analysis and processing were performed using the ImaGene Version 7.5 software (BioDiscovery Inc., Marina Del Rey, CA, USA). The microarrays were scanned at different intensities. For each of the individual microarrays the best scan was selected on the basis of signal distribution (combination of a low number of saturated spots and a low number of low signal spots). The data were normalized using Lowess normalization as available in MicroPrep (230). The data were corrected for inter-slide differences on the basis of total signal intensity per slide using Postprep (230). The median intensity of the different probes per gene was selected as the gene expression intensity. CyberT was used to compare the different transcriptomes, taking into account the duplicates (dye swaps) of each of the conditions (231). This analysis resulted in a gene expression ratio and false discovery rate (FDR) for each gene. Differential gene expression values of expression-ratios with FDR values <0.05 were considered to be statistically significant. All microarray data is MIAME compliant and is available in GEO (accession number GSE52876). Data analysis Time dependent gene expression during CO2 depletion was analyzed using Short Time-series Expression Miner (STEM; http://www.cs.cmu.edu/~jernst/st/; (271)). Log normalized expression data time series (with four data points obtained for every gene) were assigned to model profiles. Model profiles are considered significantly enriched when p<.05. Statistical significance of enriched model profiles was corrected for multiple hypothesis testing using the Bonferroni method. Time-resolved gene expression of all genes with significant changes in expression (minimally twofold induction or repression between any of the four timepoints and p<0.05) can be found in Supplementary material (Table S5.1A and S5.1B). Functional classes/categories were assigned to each gene by using the GeneOntology (272) and KEGG platform (273). Categories were considered statistically significant enriched when p<0.05. Results and discussion Inorganic carbon is one of the many growth requirements of Lactobacillus johnsonii. In chapter 4, we have shown that anaerobic growth of this lactic acid bacterium is dependent on supplementation of gaseous CO2. Upon termination of the CO2 supply 118 to an exponentially growing culture, growth quickly stagnates which is accompanied by a considerable loss in cell-viability. Notably, exposure of this bacterium to low-levels of molecular oxygen relieves this CO2 growth dependency in the wild type L. johnsonii but not in a derivative that lacks a functional pyruvate oxidase encoding gene (pox deletion derivative), indicating that endogenously produced CO2 through the pyruvate oxidase pathway relieves the exogenous CO2 dependency (chapter 4). To further explore the processes that play a role in CO2 dependency, the global transcriptome pattern of logarithmic cultures grown under anaerobic conditions during their gradual depletion for CO2 was analyzed. To this end, L. johnsonii was grown in batches in which pH, temperature, mixing and gas flow were kept constant. The gas supply was interrupted by switching from a 5% CO2/95% N2 gas mixture to 100% N2 gas. One hour after the onset of CO2 depletion growth stagnated, whereas growth continued in the culture in which CO2 was continuously supplemented (Figure 5.1). Along with growth stagnation, a 1-log drop in colony counts indicating cell death was observed. These results were in good agreement with experiments presented in chapter 4. 5 Figure 5.1: Optical density (A) or colony forming units (B) of stirred pH-controlled batch cultures sparged by N2 + 5% CO2 (triangular symbols) or switched from N2 + 5% CO2 to N2 (square symbols, dashed line indicates the switch to pure N2). RNA for transcriptome analysis was isolated before the switch to CO2-depletion (sample 1), and 30 (sample 2), 60 (sample 3) and 120 minutes (sample 4) after the intervention. Data shown are the mean of two independent experiments ± standard error of the mean. The gas-intervention elicits a rapid stripping of the inorganic carbon from the system. This CO2 removal is driven by the concentration difference between dissolved and gaseous form of CO2. Many LAB express a carbonic anhydrase (CA), which is generally assumed to convert CO2 to HCO3- (270), which could skew the equilibrium between CO2 and its protonated bicarbonate towards bicarbonate. Three types of CA are described 119 in LAB: the α-type (such as lp_2736 in L. plantarum WCFS1), the b-type (such as icfA/ bl_0616 in Bifidobacterium longum) and the γ-type (such as ef_2918 in Enterococcus faecalis) (55). However, none of the three types of carbonic anhydrases has a homolog in L. johnsonii, implying that L. johnsonii has no means to influence the HCO3- / CO2 equilibrium. Samples were taken from the culture prior to closing of the CO2 supply (reference sample) and at different time points after closing of the CO2 flow (30, 60, and 120 minutes). Notably, the sample taken two hours after closing the CO2 supply represents a culture in which growth has completely halted and in which viability has decreased approximately 10-fold (chapter 4). Global transcriptome analysis For the interpretation of the transcriptome data, the time-dependent pattern of gene expression was analyzed using Short Time Series Expression Miner (STEM), which is a pattern recognition suite that assigns each gene to a model profile based on its pattern of expression over time. Gene expression data from duplicate independent time series were compared and four model profiles were significantly enriched. A total of 384 genes passed the criterion of a minimal twofold induction or repression between any of the four timepoints. The model profile 40 and 42 that both fall in a cluster of model profiles showing a general trend of induction of expression over time, comprised 259 genes. In addition, 34 genes were assigned to model profile 11 and 26 that both fall in a cluster of model profiles that show a general trend of repression over time (see Figure S5.1 for general trend of each model profile). These profiles were statistically significant enriched when either a Bonferroni correction or an fdr-correction for multiple hypothesis testing was applied (significance level p < 0.05). Table 5.1 provides an overview of the induced and repressed model profiles and overrepresented GeneOntology and KEGG categories in these model profiles (See Table S5.1A and B for expression values of the genes assigned to the profile clusters). Analysis of overrepresentation of genes associated with a specific function or pathway amongst the induced and repressed groups showed that expression of a number of predicted regulators and ABC-type transporters was induced. However, since the predicted target genes of many of the regulators are unknown or have unknown function and many of the transporters have unknown substrates, the regulation of 120 these particular genes does not readily provide biological insights in the adaptation that the cell undergoes under these conditions. Table 5.1: Overrepresented GeneOntology groups (p<0.05) and KEGG categories in the STEM model profiles. Model profile # genes assigned GeneOntology group numbers (p<0.05) KEGG categories (p<0.05) 231 GO:0043565 sequence-specific DNA binding GO:0003700 sequence-specific DNA binding transcription factor activity GO:0006355 regulation of transcription, DNA-template GO:0042626 ATPase activity, coupled to transmembrane movement of substances M0051: Uridine monophosphate biosynthesis: glutamine 28 19 No categories enriched (+ PRPP) ==>UMP No categories enriched 15 Carbamoyl-phosphate pathway To identify any metabolic patterns present in these data, a functional category 5 analysis was performed using a KEGG functional categories annotation. This led to the observation that the genes belonging to the carbamoyl-phosphate pathway were significantly overrepresented in the STEM model profiles with a general pattern of induction (p<.0045). Previously, this pyrimidine and arginine synthesis pathway has extensively been studied in L. plantarum. Two types of carbamoyl-phosphate synthase (CPS) enzymes were identified that are involved in the first carboxylation reaction of the synthesis pathway (252). In L. plantarum, expression of CPS-P (encoded by pyrAB) is regulated by intracellular UMP-levels via a transcription attenuation mechanism (263), whereas expression of CPS-A (encoded by carAB) is controlled by intracellular levels of arginine. Moreover, two regulators control this pathway, PyrR1 acts as a repressor when pyrimidine levels in the environment are high (252), while PyrR2 positively regulates expression of the pyr-operon in response to low inorganic carbon levels (253). The L. johnsonii genome encodes homologues of all the enzymes involved in the pyrimidine synthesis pathway described in L. plantarum, including the two PyrR 121 regulators. Moreover, L. johnsonii also encodes two copies of the CPS encoding genes, which are both present in an operon (lj_1276-1277 and lj_1185-1184). The entire carbamoyl-phosphate and pyrimidine synthesis pathways were induced in L. johnsonii 120 minutes after the onset of CO2 depletion, with the highest fold-induction within the entire datasets assigned to the aspartate carbamoyltransferase encoding pyrB and dihydroorotase encoding pyrC genes that were 17- and 13-fold induced, respectively. The CPS-A encoding genes (carAB) did not display any significant change in expression. Moreover, the uracil permease (lj_0709; pyrP) of L. johnsonii was also strongly induced (10-fold higher expression), indicating that similar to L. plantarum (274) the uracil transport capacity is coregulated with the pyr-operon in L. johnsonii. However, the uracil phosphoribosyltransferase enzyme (encoded by lj_0933; upp), which catalyzes the conversion of imported uracil to UMP, was not significantly affected by CO2 depletion. Notably, the absolute level of expression of upp was found to be among the 200 highest expressed genes within the L. johnsonii genome, which may suggest that this function may not be limiting for pyrimidine import. In a second potential uracil salvage pathway, uridine monophosphate is formed via the ribonucleoside uridine. Genes encoding the enzymes that catalyze the two reactions, uridine kinase (lj_0763; udk) and uridine phosphorylase (lj_1217-lj1218; udp), are both present in the L. johnsonii genome but their expression did not show significant induction or repression upon CO2 depletion. A schematic overview of the carbamoyl-phosphate pathway is presented in Figure 5.2 including changes in gene expression. The observed activation of expression of the carbamoyl-phosphate synthesis operon and pyrimidine import strongly suggests that, analogous to what has been shown for L. plantarum, depletion of the pyrimidine pool in L. johnsonii is the main cause of the observed growth stagnation upon removal of inorganic carbon supplies. In L. plantarum, approximately 30% of the strains displayed a high environmental-C1-carbon demand, which could be complemented by supplementation of the growth media with either arginine or uracil (249). L. johnsonii is auxotrophic for arginine, irrespective of the availability of environmental CO2, suggesting that it is unlikely that arginine exhaustion is the main cause of growth stagnation in this species. Moreover, the transcription of the genes encoding the arginine-depletion responsive CPS-A (carAB) were unaffected during CO2 depletion, supporting that intracellular arginine depletion is not occurring. Analogously, supplementation of solid MRS medium with excessive concentrations of arginine (10 mM), did not relieve the CO2 dependency of L. johnsonii (Figure 5.4). Although the current experiments employed the rich laboratory medium MRS for 122 growth of L. johnsonii, the extracellular pyrimidine concentrations or their rate of import may still be limiting to sustain normal growth under CO2 depleted conditions. Moreover, it is well-established that the exhaustion of thymine, another product of the carbamoyl-phosphate pathway, can cause cell death in auxotrophic bacteria, by creating disbalanced endogenous dNTP-pools, leading to single and double strand DNA breaks (275). This so-called thymineless death was also shown to occur in species of the L. acidophilus group (276). Therefore, we evaluated whether pyrimidine exhaustion could be the primary cause of cell death following CO2-depletion, by supplementing MRS with pyrimidines and pyrimidine precursors at a very high level (100 mg/L). Although the genome of L. johnsonii predicts that this bacterium can interconvert uracil to the other pyrimidines (thymidine and cytosine), also thymidine, uracil and the uracil precursor orotic acid were added to the medium. None of these supplementations was able to overcome the growth stagnation and cell death induced by CO2 depletion. It even appeared that the growth arrest of the pyrimidine supplemented culture was even slightly accelerated as compared to the normal MRS media (Figure 5.3). In addition to the experiment in liquid medium, growth of L. johnsonii on solid MRS medium with and without addition of nucleotides was tested. To this end, overnight cultures were spotted on MRS agarose, with or without 100 mg/l of uracil, thymidine, cytosine and orotic acid and colonies were enumerated after growth with or without 5% gaseous CO2. The difference between colony forming units (cfu) in the presence and absence of CO2 on regular MRS medium was approximately 2 log. Analogous 5 to liquid medium, growth in the absence of CO2 on solid media supplemented with excess pyrimidines and pyrimidine precursors did not improve compared to regular medium. We conclude that extracellular pyrimidine supplementation cannot overcome the inorganic carbon requiring phenotype in L. johnsonii, which is in clear contrast to previous observations made for L. plantarum. However, there is no experimental evidence that the PyrP uracil transporter in L. johnsonii is functional, and ineffective import could explain why supplementation of pyrimidines did not complement this cell death. Alternatively, the upp- or the udk/udp-encoded pathway to convert uracil into UMP may be defective in L. johnsonii, which could also explain why the uracil supplementation is not effective. The transcriptome analyses imply clearly that drainage of the intracellular pyrimidine pools may be the causal mechanism of CO2-related cell death in L. johnsonii, but for as yet unknown reasons, extracellular supplementation with the corresponding components failed to overcome such drainage and could thus not overcome the observed cell death. 123 124 1,1 0,9 0,8 1,0 1,1 1,0 1,0 1,0 1,0 0,8 6,7 9,5 Arg argH argG argF Carba-‐P pyrB Pi α-‐KG Gln ATP CO2 + ADP 1,1 1,2 17,5 Asp 1,1 1,2 2,1 OAA 0,9 1,0 1,0 PEP PPi Carba-‐Asp pyrC pyrD pyrE pyrF 1,0 1,2 1,2 1,1 1,1 1,9 1,7 1,5 13,1 4,4 4,8 6,0 UMP udk uridine 1,4 1,5 1,5 1,2 1,3 1,5 1,0 1,1 1,0 ATP PPi DNA synthesis OUT 0,9 1,9 10,9 D-‐ribose-‐1-‐P pyrP ADP udp pyrimidines 0,9 0,9 0,6 H2O + NADH2 NAD++PRPP PPi + CO2 upp PRPP uracil IN uracil Figure 5.2: Aspartate and pyrimidine biosynthesis pathway in L. johnsonii and differences in their expression level 30 min, 60 min and 120 min after closing of the CO2 supply compared to CO2-rich conditions. Pathways were adjusted from Kegg-pathway (273) and (249, 274). PEP: phosphoenolpyruvate. OAA: oxaloacetate. Asp: aspartate. Gln: glutamine. α-KG: α-ketoglutarate. Carba-P: carbamoyl-phosphate. Arg: arginine. Carba-Asp: carbamoyl-aspartate. PRPP: phosphoribosyl pyrophosphate UMP: uridine monophosphate. carAB and pyrAB: carbamoyl-phosphate synthase. argF-H: ornithine carbamoyltransferase, argininosuccinate synthase, argininosuccinate lyase. pepCK: phosphoenolpyruvate carboxykinase. aspC: aspartate transaminase pyrB: aspartate carbamoyltransferase, pyrC-F dihydroorotase, dihydroorotate dehydrogenase, orotate phosphoribosyltransferase, orotidine 5’-phosphate decarboxylase. pyrP: uracil permease. upp: uracil phosphoribosyltransferase. udp: uridine phosphorylase. udk: uridine kinase. Reactions/enzyme whose gene is absent in the L. johnsonii genome (argFGH) are depicted in grey. Green indicates induced expression (upper limit 5-fold induced), red indicates repressed expression (lower limit at 5-fold repressed). pyrAB carAB HCO3-‐ + NH4+ (Gln) 2 ATP 2 ADP aspC pepCK glycolysis Figure 5.3: Optical density of L. johnsonii cultures grown in regular MRS (closed symbols) or MRS supplemented with 100 mg/l of uracil, thymidine and orotic acid (open symbols). The dashed line indicates the time point where gas regime is switched from sparging N2 + 5% CO2 to pure N2. Data shown are the mean of two independent experiments ± standard error of the mean. Aspartate biosynthesis In other LAB, aspartate biosynthesis has been identified as one of the CO2 requiring pathways (251, 277, 278). Aspartate is required in the carbamoyl-phosphate pathway as a substrate for PyrB, the aspartate carbamoyl transferase. This amino acid is one of the few amino acids that L. johnsonii is predicted to be able to synthesize autonomously from intermediates of the glycolytic pathway. Aspartate transaminase (aspC, lj_1390) (176) uses oxaloacetate and glutamine as substrates to produce aspartate and α-ketoglutarate. The transcription of the aspC gene was 2.1-fold induced after 120 5 minutes of CO2 depletion, relative to CO2 rich conditions. Oxaloacetate is formed by carboxylation of phospoenolpyruvate (PEP), catalyzed by a PEP-carboxylase (PEPC) that uses primarily bicarbonate as a substrate and does not produce an additional ATP, or by PEP carboxykinase (PEPCK) that uses CO2 and produces one ATP (279). The PEPC function in L. johnsonii appears to be disrupted by a single point mutation that introduces a premature stop-codon and thus is classified as a pseudogene (lj_1272 and lj_1273). In contrast the PEPCK function appears intact and is encoded by the lj_0149 gene. Thus, PEPCK and ASPC could synthesize aspartate from central metabolism (PEP), using glutamine and CO2 as co-substrates (Figure 5.2). Although transcription of aspC is induced and transcription of pepCK is not significantly changed, we hypothesize that aspartate may be synthesized through a CO2 requiring pathway. Although aspartate is assumed to be presented in sufficient amounts in the MRS medium, we verified that excess supplementation with this amino acid (30 mM) did not complement the CO2 requirement (Figure 5.4). 125 Figure 5.4: Enumeration of colony forming units of L. johnsonii on solid MRS agar supplemented with aspartate (30 mM), arginine (10 mM) or pyrimidine nucleotides and precursors (100 mg/L) incubated under a N2 + 5% CO2 (open bars) or N2 (grey bars) atmosphere. Concentrations of the supplements were approximately 10-fold higher than those normally present in chemically defined medium (see Materials & Methods section of chapter 4). Data represent average of two independent experiments + standard error. Concluding remarks The experiments in chapter 4 show that L. johnsonii requires CO2 for growth. In other LAB, the CO2 growth-dependency was shown to be related to the biosynthesis pathways of pyrimidines (uracil) and/or amino acids (arginine and aspartate). Our results clearly show that, analogous to L. plantarum, the carbamoyl-phosphate pathway is regulated in response to CO2 levels. However, supplementation of uracil, thymidine and the uracil-precursor orotic acid could not prevent the stagnation of growth related to CO2 depletion, nor could it prevent the induction of cell death. This implies that besides pyrimidine synthesis, an alternative pathway is responsible for the CO2 demand. However, it may also be that the pyrimidine uptake system is compromised in L. johnsonii, which could explain why pyrimidine supplementation of the media did not overcome the growth stagnation. The constitutively high expression of the uracil phosphoribosyltransferase-encoding upp gene, may indicate that the catalytic function of this enzyme is compromised. This could imply that it is part of a futile regulatory circuit, in which the importer is transcriptionally upregulated but fails to increase intracellular levels of its substrate and therefore provide negative feedback on its expression. One of the possibilities to study the uracil transport capacity is to use the toxic uracil analog 5-fluorouracil. If the uracil transport function in L. johnsonii is impaired, the cells should be resistant to this cytotoxic molecule. Furthermore, uridine supplementation should be able to bypass the postulated impairment of uracil import and conversion, and could thus prevent the CO2-related cell death, provided that drainage of the 126 pyrimidine pool is indeed the prime cause of cell death. An alternative pathway that could depend on CO2 is aspartate biosynthesis, where the first step in the biosynthesis pathway involves the carboxylation of phosphoenolpyruvate, leading to the formation of the aspartate-precursor oxaloacetate and ATP. However, aspartate supplementation to the medium did also not complement the CO2 requirement. Nevertheless, we cannot rule out that, besides aspartate, also other metabolites depend on oxaloacetate as an intermediate and their synthesis may thereby be dependent on an adequate CO2 supply. Overall we conclude that L. johnsonii has limited decarboxylases that could generate endogenous CO2. A comparison of the CO2 dependency of a wild type strain of L. johnsonii and its pyruvate oxidase deficient derivative in the presence of oxygen suggested that oxidation of pyruvate could provide an additional CO2 source (chapter 4). L. johnsonii lacks most other genes encoding C1-generating enzymes, such as several known amino acid decarboxylases, pyruvate dehydrogenase and pyruvate formate lyase. Interestingly, L. johnsonii’s natural habitat is the intestinal tract of mammals, where these bacteria may encounter substantial levels of CO2 and bicarbonate in certain areas of the GI-tract, e.g., due to bicarbonate-rich pancreatic secretions (280) or as a result of the metabolic activity of other microbiota members. We speculate that, analogous to the loss of the capacity to synthesize many of the amino acids, or vitamins, evolutionary adjustment to the CO2 rich environment of the GI-tract may include the loss of CO2 5 producing capacities. Finally, it is noteworthy that at present many of the genes that were modulated by the CO2 depletion lack a function prediction and/or do not contain recognizable domains, including several genes of which the expression was quite drastically changed like lj_1790 (10.2-fold induced) and lj_1300 (6.6-fold repressed). These genes may play a significant role in the CO2 depletion response, but such a role remains to be established, which illustrates the importance of continued elucidation of the biological functions of such genes. 127 Acknowlegements We thank Dr Anne de Jong from the Molecular Genetics Group of Groningen University for the compilation of the gene assignments to functional categories used in our analyses. 128 Supplementary material Figure S5.1: General trend of expression of STEM model profiles. 120 min 30 min STEM locus 60 min Table S5.1A: Expression ratio of all genes with minimal twofold induction between any of the four timepoints (p<0.05). NCBI annotation is shown where available. Alternatively Gene Ontology classification of proteins or conserved domains is shown (in grey). Where no Gene Ontology classification could be assigned, the closest BlastP result is shown (displayed in grey, bold and italic). PTS: phosphotransferase system. PEP: phosphoenolpyruvate. LJ_0020 LJ_0021 LJ_0025 LJ_0027 LJ_0030 LJ_0031 LJ_0034 LJ_0036 LJ_0037 LJ_0038 LJ_0039 LJ_0040 LJ_0041 LJ_0042 LJ_0047 LJ_0048 LJ_0050 LJ_0059 LJ_0062 LJ_0063 LJ_0072 LJ_0073 LJ_0074 LJ_0077 LJ_0083 LJ_0084 LJ_0089 LJ_0090 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 18 18 18 18 40 42 40 40 1,3 1,4 1,1 1 1,2 1,4 1,3 1,1 1,3 1,2 1,4 1,1 1 1,2 1,1 1,2 1,3 1,2 1,1 1,2 0,9 1 1 1 1,2 1,3 1,5 1,1 1,4 1,4 1,3 1,1 1,1 1,5 1,4 1,3 1,2 1 1,3 1 0,9 1,1 1,1 1,2 1,1 1,3 1 1,1 1,1 1,2 1,2 0,9 1,4 1,7 1,6 1,2 2,3 2,1 2,4 3,7 2,1 3,2 5,7 2,4 2,4 4,9 6 3,8 3,5 2,1 3,2 2,2 2 2,1 3 4,5 2,7 3,2 4,1 1,8 2,2 3,1 2,5 3 LJ_0091 18 1 1,1 2,9 LJ_0092 40 1 1,3 3,9 LJ_0093 LJ_0098 LJ_0099 LJ_0101 LJ_0107 LJ_0123 LJ_0128 LJ_0143 LJ_0154 LJ_0162 LJ_0163 40 42 40 40 40 40 42 40 40 40 40 1,3 1,4 1,1 1,2 1,1 1,3 1,6 1,1 1,2 1,1 1,2 1,5 1,6 1 1,3 1,1 1,3 2,2 1,7 1,5 1 1,1 2,9 2,3 2,3 2,2 3,2 2,4 2,7 3,7 2,7 2 3,7 NCBI annotation Transcriptional regulator Molybdenum cofactor biosynthesis, MoeB EDD domain protein, DegV family Transcriptional regulator Endopeptidase O Transcriptional regulator Mg2+ transporter protein, CorA-like/Zinc transport protein ZntB Exodeoxyribonuclease Helicase, C-terminal Cell surface hydrolase Cell surface hydrolase Uncharacterized protein Predicted permease FAD-dependent pyridine nucleotide-disulphide oxidoreductase Adhesion exoprotein Homeobox protein, antennapedia type, conserved site LysR, substrate-binding Phospholipid/glycerol acyltransferase Glycosyl transferase, family 8 Phospholipid/glycerol acyltransferase ABC transporter permease component ABC transporter ATPase component ABC transporter permease LemA-like protein HD containing hydrolase-like enzyme HNH endonuclease MarR family transcriptional regulator Membrane protein Methylated DNA-protein cysteine methyltransferase DNA binding domain Methylated DNA-protein cysteine methyltransferase ribonucleaselike domain Sirtuin family Ribonuclease H domain Uncharacterized, extracellular protein Alpha/beta superfamily hydrolase SCP-like protein Uncharacterized membrane protein Universal stress protein LPXTG-motif cell wall anchor domain protein Uncharacterized small conserved protein Putative secreted protein Sensory box protein 5 129 130 LJ_0165 LJ_0172 LJ_0181 LJ_0185 LJ_0186 LJ_0187 LJ_0197 LJ_0201 LJ_0222 LJ_0227 LJ_0228 LJ_0234B LJ_0243 LJ_0253 LJ_0254 LJ_0259 40 40 42 40 40 40 40 40 40 40 40 18 27 42 42 14 LJ_0260 40 1,2 1,2 2,9 LJ_0261 LJ_0262 LJ_0288 LJ_0292 LJ_0293 LJ_0294 LJ_0304 LJ_0305 LJ_0306 LJ_0309 LJ_0310 LJ_0324 LJ_0325 LJ_0328 LJ_0329 LJ_0376 LJ_0377 LJ_0378 LJ_0379 LJ_0383 LJ_0394 LJ_0396 LJ_0418 LJ_0420 LJ_0421 LJ_0422 LJ_0452 LJ_0456 LJ_0457 LJ_0458 LJ_0459 LJ_0459B LJ_0462 LJ_0463 LJ_0464 LJ_0481 LJ_0482 LJ_0483 LJ_0486 LJ_0494 LJ_0495 LJ_0496 LJ_0499 LJ_0502 LJ_0522 LJ_0524 LJ_0525 LJ_0528 LJ_0529 40 40 40 40 18 40 40 40 18 14 40 14 40 40 40 40 40 40 40 14 40 40 18 40 42 40 40 40 40 40 40 40 40 40 40 42 40 40 40 40 40 40 40 18 40 40 18 40 40 1,1 1 1,2 1,1 1 1,2 1,1 1,2 1 0,9 1,2 0,9 1,2 1,1 1,2 1,1 1,2 1,1 1,3 0,9 1,2 1,1 0,8 1 1,3 1,1 1,2 1,3 1,3 1,4 1,3 1,2 1,1 1,2 1,2 1,1 1,1 1,2 1,3 1,1 1,2 1,1 1,1 1 1,1 1,2 1 1,1 1 1,1 1 1,2 1,1 1 1,2 1,3 1,4 1,3 0,7 1,1 0,8 1,3 1,1 1,2 1,1 1,4 1,2 1,4 0,8 1,3 1 1 1,1 1,9 1,2 1,4 1,4 1,5 1,7 1,6 1,6 1,5 1,4 1,2 1,8 1,1 1,2 1,5 1 1,3 1,1 1,2 1,4 1,4 1,3 1,1 1,2 1,3 2,4 2 2,2 2,6 3,1 4,5 3,1 2,4 2,3 2,3 2,2 3,3 4,7 4,4 5,9 2,9 5 2,2 2,1 2,4 3,5 2,5 1,8 2,3 3 3,7 2,1 3,4 3 3,5 3 3,8 3,7 2,3 3,6 2,2 2 2,5 2,2 2,3 3,5 2,2 2 2,6 2,3 3 2,7 2,6 2,1 1,2 1,2 1,2 1,4 1 1,3 1,2 1 1,2 1,3 1,2 0,8 1,1 1 1,2 1 1,6 1,1 1,6 1,5 1,2 1,4 1,3 1,3 1,3 1,5 1,4 1 1,9 1,6 1,8 0,9 3,2 2,1 2,1 2,6 2,3 2,3 3,8 8,2 2,3 2,5 2 1,8 0,8 2,5 2,1 1,9 Toxin-antitoxin system Cell envelope-like function transcriptional regulator Heat shock protein Hsp20 Transcription regulator, GntR Transposase IS204/IS1001/IS1096/IS1165 Uncharacterized Uncharacterized Cation-transporting P-type ATPase Uncharacterized Xanthine phosphoribosyltransferase Beta-carotene 15,15-monooxygenase Uncharacterized Uncharacterized Uncharacterized Uncharacterized Raffinose operon transcriptional regulator Raffinose permease containing PEP-dependent sugar PTS EIIA domain Alpha-galactosidase Sucrose phosphorylase Lj965 prophage integrase Lj965 prophage protein Lj965 prophage repressor Lj965 prophage repressor Lj965 prophage protein Lj965 prophage protein Lj965 prophage protein Lj965 prophage portal protein Lj965 prophage protein Lj965 prophage protein Lj965 prophage protein Lj965 prophage holin Lj965 prophage lysin ArsR-type DNA-binding domain Cation efflux protein Uncharacterized Uncharacterized Amino acid/polyamine transporter I Beta-1,6-galactofuranosyltransferase LytR family transcriptional regulator Uncharacterized Putative flagellar protein FliS Uncharacterized tRNA-specific adenosine deaminase Uncharacterized Lysine decarboxylase PEP-dependent sugar PTS EIIC cellobiose specific LytTR DNA-binding domain Uncharacterized Uncharacterized Transcriptional regulator Putative transcriptional regulator ABC transporter ATPase and permease components uncharacterized hydrocarbon binding protein Glutamate racemase Non-canonical purine NTP pyrophosphatase Small mechanosensitive ion channel protein Glycerophosphoryl diester phosphodiesterase Uncharacterized 5'-Nucleotidase, C-terminal uncharacterized membrane protein General stress protein 13 Glutamine amidotransferase MFS transporter permease Tat pathway signal sequence domain protein / cytochrome C5 Transcriptional regulator protein RpiR Beta-lactamase class A LJ_0536 LJ_0553 LJ_0573 LJ_0574 LJ_0589 LJ_0595 LJ_0599 LJ_0600 LJ_0613 LJ_0614 LJ_0615 LJ_0617 LJ_0618 LJ_0623 LJ_0637 LJ_0640 LJ_0644 LJ_0645 LJ_0646 LJ_0647 LJ_0648 LJ_0649 LJ_0652 LJ_0653 LJ_0669 LJ_0672 LJ_0673 LJ_0674 LJ_0675 LJ_0709 LJ_0712 LJ_0722 LJ_0723 LJ_0724 LJ_0725 LJ_0726 LJ_0730 LJ_0738 LJ_0741 LJ_0748 LJ_0749 LJ_0750 LJ_0751 LJ_0757 LJ_0758 LJ_0759 LJ_0760 LJ_0761 LJ_0762 LJ_0783 LJ_0784 LJ_0785 LJ_0788 LJ_0814 LJ_0815 LJ_0836 LJ_0864 LJ_0900 LJ_0902 LJ_0923 LJ_0965 LJ_1004 LJ_1005 LJ_1039 LJ_1042 LJ_1049 40 40 40 40 42 40 40 40 40 18 40 40 40 40 18 40 42 40 18 18 40 40 40 40 40 40 40 40 40 18 40 18 40 40 40 37 40 37 14 37 40 40 40 42 40 40 40 42 40 18 40 40 42 29 40 25 40 37 40 25 18 40 30 42 40 40 1,1 1,2 1,2 1,1 1,5 1,1 1,4 1,2 1,1 1 1,3 1,1 1,1 1,6 0,9 1,3 1,4 1 1 0,9 1 1 1,2 1 1,1 1,2 1,1 1,1 1,1 0,9 1,3 1 1,1 1,1 1,1 1,3 1,3 1 0,9 1,3 1,3 1 1 1,4 1,1 1,1 1,1 1,4 1,2 1 1 1,2 1,1 1,1 1,3 1 1,1 1,1 1 1 0,9 1,1 1 1,3 1,1 1,1 1,1 1,2 1,1 1,1 1,7 1,3 1,7 1,3 1 1 1,5 1,2 1,4 1,5 1 1,6 1,7 1,2 1,2 1,1 1,2 1,1 1,3 1 1,1 1,2 1,2 1 1,1 1,9 1,4 1 1,3 1,3 1,4 0,9 1,4 0,9 0,7 1 1,5 1,1 1 2,6 1,1 1 1,3 2,3 1,5 1 1,1 1,2 1,5 3,2 1,4 0,9 1,1 0,9 1,1 0,8 1,4 1,1 2 1,6 1,2 0,9 3,5 4,7 2,3 2,4 2,5 2 3,2 2,8 4,3 5,1 13,8 3,6 2,8 2,4 2 3,1 2,4 2,2 2,1 2,2 2,3 2,1 3,4 2,1 2,1 3,7 5,7 2,5 2,6 10,9 2,3 3,6 6,1 6,6 5 3,5 2,9 1,8 1,5 2,5 7,6 3,4 2,7 3,5 2,4 2,2 2 6,6 2,7 2,6 2,5 3,9 2,2 5,1 2,4 1,8 2,1 1,9 2,1 2,2 2,1 2,1 1,2 2,3 2,6 4,6 Alpha/beta hydrolase fold domain Colicin V production, CvpA YSIRK Gram-positive signal peptide Gram-positive signal peptide protein Plasmid maintenance toxin/Cell growth inhibitor Uncharacterized GTPase HflX ABC transporter ATPase component Cation-transporting ATPase Cation-transporting ATPase Metallopeptidase, catalytic domain Cysteine protease YvpB SGNH hydrolase-type esterase domain CAT RNA-binding domain Transcriptional regulator protein RpiR Large-conductance mechanosensitive channel Oxidoreductase Peptidase T Uncharacterized Uncharacterized membrane protein Sir2 family NAD-dependent protein DNA polymerase III DNA/RNA non-specific endonuclease Dipeptidase SNARE-like domain protein Transcriptional regulator Alpha/beta hydrolase fold-3 Uncharacterized membrane protein Acyl-CoA N-acyltransferase Uracil permease Phosphoglycerate mutase Major facilitator superfamily permease Uncharacterized Transcription regulator, MarR-type Uncharacterized Uncharacterized Phophatidylserine decarboxylase Beta-galactosidase PEP-dependent sugar PTS EIID, probable fructose specific Cell surface hydrolase Transcriptional regulator, MarR family ABC transporter ATPase and permease components ABC transporter ATPase and permease components Uncharacterized Trehalose-6-phosphate hydrolase Trehalose operon repressor PEP-dependent sugar PTS EIIABC, probable trehalose specific Uncharacterized Uncharacterized Cation-transporting P-type ATPase Cation-transporting P-type ATPase Acyl transferase/acyl hydrolase/lysophospholipase Acyl-CoA N-acyltransferase Uncharacterized ATP-dependent clp protease ATP-binding subunit clpE Glucose/ribitol dehydrogenase UvrABC system protein A Alpha/beta superfamily hydrolase DNA/pantothenate metabolism flavoprotein Glycopeptide antibiotics resistance protein Uncharacterized DNA polymerase III 30S ribosomal protein S20 Uncharacterized Uncharacterized dTDP-glucose 4,6-dehydratase 5 131 LJ_1050 LJ_1051 LJ_1052 LJ_1059 LJ_1060 LJ_1063 LJ_1064 LJ_1066 LJ_1070 LJ_1088 LJ_1107 LJ_1108 LJ_1118 LJ_1121 LJ_1122 LJ_1128 LJ_1136 LJ_1137 LJ_1146 LJ_1166 LJ_1194 LJ_1195 LJ_1202 LJ_1203 LJ_1204 LJ_1215 LJ_1229 LJ_1230 LJ_1231 LJ_1237 LJ_1238 LJ_1239 LJ_1247 LJ_1248 LJ_1249 LJ_1250 LJ_1252 LJ_1254 LJ_1255 LJ_1260 LJ_1266 LJ_1270 LJ_1271 LJ_1275 LJ_1276 LJ_1277 LJ_1278 LJ_1279 LJ_1280 LJ_1281 LJ_1282 LJ_1283 LJ_1284 LJ_1287 LJ_1325 LJ_1388 LJ_1390 LJ_1417 LJ_1422 LJ_1435 LJ_1436 LJ_1438 LJ_1441 LJ_1454 LJ_1457 LJ_1458 132 40 18 18 40 40 40 40 40 40 18 40 40 40 40 42 37 40 29 40 40 40 40 37 14 40 42 14 18 40 40 40 18 40 40 18 40 18 40 40 40 37 40 40 18 18 18 40 40 18 40 40 40 18 42 20 40 40 18 40 18 40 40 42 29 40 40 1 1 0,9 1,1 1,2 1,2 1,2 1,1 1,1 0,8 1,1 1,1 1,1 1,3 1,7 1,1 1,4 0,8 1,7 1,3 1,5 1,5 1 1 1 1,1 0,9 1 1,2 1,4 1,3 0,7 1,1 1,3 1 1,5 1 1 1,1 1,1 1,1 1,3 1,1 1 0,8 1 1 1,1 1 1,2 1,1 1,2 1 1,5 0,7 1,1 1 0,8 1,2 0,9 1,4 1,1 1,3 1,3 1,2 1 0,9 0,9 0,9 1,2 1,3 1,3 1,1 1,3 1,1 1,1 1 1 1,2 1,5 2,5 0,9 1,5 3,2 1,7 1,3 1,3 1,5 0,9 0,9 1 1,5 0,7 0,9 1,3 1,5 1,3 1,1 1,2 1,4 0,9 1,9 0,9 1 1 1 0,9 1,4 1,2 0,8 1 1 1,1 1,2 1,6 1,9 1,5 1,7 0,9 2,3 1,5 1,4 1,2 0,9 1,2 1,1 1,4 1,1 2 2,2 1,3 1,1 3,5 2,5 2,5 2,6 2,5 2,4 2,1 2,4 4,1 1,9 2,8 2,1 2,1 2,4 3,9 2,2 2,8 3 4,9 2,4 2,1 5,3 1,9 1,9 1,9 2,1 1,8 2 2,1 3,6 3,5 1,7 3 5,4 4,3 3,9 1,9 2,5 2,4 3,1 2,3 5,9 2,3 4,3 6,7 9,4 13,1 17,5 4,3 4,4 6 4,8 4 2,5 0,9 2,3 2,1 1,8 2,5 2,1 3,5 2,6 2,4 2 3,5 2,1 Glucose-1-phosphate thymidylyltransferase dTDP-4-dehydrorhamnose 3,5-epimerase dTDP-4-dehydrorhamnose reductase Uncharacterized NUDIX hydrolase domain Uncharacterized Rib/alpha-like repeat protein Oxidoreductase UvrABC system protein C ECF transporter, Riboflavin transporter RibU SMF protein DNA topoisomerase 1 Transcription regulator, LysR Uncharacterized ABC transporter ATPase component Domain of unknown function DUF1542 Uncharacterized Branched-chain amino acid transport system carrier protein Uncharacterized ABC transporter ATPase component Cell division protein GpsB UPF0398 protein ATP-dependent helicase, C-terminal ATP-dependent helicase/nuclease subunit A ATP-dependent helicase/deoxyribonuclease subunit B Bifunctional protein FolD 1 Uncharacterized Phospholipase D-like domain Uncharacterized Uncharacterized Uncharacterized Acyl-CoA N-acyltransferase ABC transporter ATPase and permease components ABC transporter ATPase component ABC transporter ATPase and permease components SPFH/Band 7/PHB domain protein Sugar-phosphate isomerase Aldolase-type TIM barrel Aldolase-type TIM barrel Lambda repressor-like, DNA-binding domain Transketolase Armadillo-type fold Glycosyl transferase, family 1 Bifunctional protein PyrR Carbamoyl-phosphate synthase large chain Carbamoyl-phosphate synthase small chain Dihydroorotase Aspartate carbamoyltransferase Bifunctional protein PyrR Dihydroorotate dehydrogenase A (fumarate) Orotidine 5'-phosphate decarboxylase Orotate phosphoribosyltransferase Beta-carotene 15,15-monooxygenase UPF0145 protein 30S ribosomal protein S21 SGNH hydrolase-type esterase domain Aspartate aminotransferase Phosphatidic acid phosphatase type 2/halopero Lj928 prophage protein Lj928 prophage major head protein Lj928 prophage minor head protein Lj928 prophage protein Lj928 prophage terminase large subunit Lj928 prophage protein Lj928 prophage repressor protein Lj928 prophage protein LJ_1465 LJ_1466 LJ_1501 LJ_1502 LJ_1504 LJ_1505 LJ_1509 LJ_1510 LJ_1511 LJ_1523 LJ_1544 LJ_1556B LJ_1557 LJ_1565 LJ_1566 LJ_1567 LJ_1568 LJ_1571 LJ_1574 LJ_1575 LJ_1576 LJ_1577 LJ_1588 LJ_1590 LJ_1610 LJ_1615B LJ_1621 LJ_1657 LJ_1658 LJ_1659 LJ_1660 LJ_1666 LJ_1679 LJ_1686 LJ_1687 LJ_1688 LJ_1689 LJ_1690 LJ_1702 LJ_1724 LJ_1725 LJ_1727 LJ_1728 LJ_1729 LJ_1730 LJ_1738 LJ_1740 LJ_1747B LJ_1751 LJ_1752 LJ_1753 LJ_1754 LJ_1765 LJ_1774 LJ_1778 LJ_1779 LJ_1785 LJ_1786 LJ_1787 LJ_1788 LJ_1789 LJ_1790 LJ_1791 LJ_1794 LJ_1796 LJ_1802 40 40 40 40 42 40 40 40 42 42 42 40 40 40 40 40 18 40 42 40 40 42 40 40 40 30 40 40 40 40 40 40 40 18 40 40 40 42 40 18 40 14 14 40 40 42 40 40 40 40 40 40 49 18 42 18 18 40 40 40 40 40 40 40 40 40 1,3 1,3 1,2 1,1 1,1 1,1 1,4 1,5 1,5 1,5 1,3 1,1 1,2 1,1 1,2 1,2 1 1,2 1,3 1,1 1,3 1,5 1,1 1,2 1,1 0,9 1 1,1 1 1,1 1,2 1,4 1,2 1 1,3 1 1 1,4 1,3 1 1,1 0,9 0,9 1,3 1,1 1,3 1,2 1,2 1,2 1,3 1,1 1,5 1,4 1 1,3 0,9 1 1 1 1,2 1,2 1,2 1,1 1,1 1 1,2 1,4 1,3 1,2 1,3 1,4 1,3 1,4 1,5 1,6 1,7 1,7 1,5 1,1 1,1 1,6 1,3 1 1,4 2 1,3 1,6 2,3 1 1,2 1,4 2,2 1,1 1,3 1,1 1,2 1,3 1,4 1,4 1,3 1,3 1,1 1,1 1,7 1,5 1 1 0,7 0,8 1,6 1,2 1,7 1,2 1,4 1,1 1,2 1,1 1,6 2,1 1,3 1,8 1,2 1,2 1,2 1,2 1,5 1,4 1,9 1,1 1,6 1,2 1,4 2,1 2,7 2,1 2,8 2,1 2,2 2,3 2,3 2,2 2,3 2,7 3,5 2,6 2,7 2,8 3,3 3,2 4,1 2,1 2,2 4,1 5 2,1 2 3 1,6 2 4,4 2,4 2,1 2,5 3,2 2,8 2,4 2,8 2,7 4,1 2,4 4,2 2 2 2,3 2,4 3,4 2,1 2,1 2,3 2,1 2,2 2,5 2,6 2,7 1,3 2 3,6 2,2 2,7 4,6 5,6 4,6 4,1 10,2 2,6 3,1 4,4 2,7 Lj928 prophage integrase LysR, substrate-binding DNA methylase, N-6 adenine-specific, conserved protein Phospholipid/glycerol acyltransferase ABC transporter ATPase and permease components ABC transporter ATPase and permease components Uracil-DNA glycosylase NUDIX hydrolase domain Signal peptidase I Oligopeptide ABC transporter solute-binding component Uncharacterized Uncharacterized Uncharacterized membrane protein Uncharacterized membrane protein Alpha/beta hydrolase fold-3 DNA binding domain, AraC-type RmlC-like cupin domain Uncharacterized ABC transporter solute-binding component Transcriptional regulator Major facilitator superfamily permease Uncharacterized RelA/SpoT K homology domain-like, alpha/beta Alpha/beta hydrolase Uncharacterized ABC transporter permease GntR family transcriptional regulator Two-component system histidine kinase Two-component system response regulator Bacterial extracellular solute-binding, family 1 Uncharacterized Pseudouridine synthase Uncharacterized Uncharacterized Aminoglycoside 3-phosphotransferase Beta-lactamase Uncharacterized DNA-directed RNA polymerase subunit delta Zinc-metalloprotease Glycosyl transferase Polysaccharide biosynthesis protein CDP-glycerol glycerophosphotransferase Transcriptional regulator ABC transporter ATPase and permease components Glycerol-3-phosphate cytidylyltransferase Uncharacterized Phosphate-starvation-induced PsiE Replication initiation factor ATP transporter Uncharacterized Uncharacterized Uncharacterized Amino acid transporter Uncharacterized Major facilitator superfamily permease Uncharacterized Quinonprotein alcohol dehydrogenase-like superfamily Uncharacterized Uncharacterized Uncharacterized Uncharacterized Cystathionine beta-synthase, core Uncharacterized Transcriptional regulator Uncharacterized 5 133 LJ_1803 LJ_1804 LJ_1805 LJ_1806 LJ_1807 LJ_1816 LJ_1817 LJ_1818 LJ_1819 LJ_1822 LJ_1823 LJ_1828 LJ_1831 LJ_1833 LJ_1834 LJ_1839 LJ_1841 LJ_1843 LJ_1845 LJ_1848 LJ_1850 134 40 18 40 40 40 40 40 40 40 40 40 40 40 40 40 40 40 18 42 40 40 1,4 0,9 1,1 1,1 1,1 1,1 1,1 1,4 1,2 1,4 1,3 1,4 1,2 1,3 1,3 1 1,7 1 1,6 1 1,3 1,6 1 1,3 1,3 1,2 1,5 1,3 1,3 1,4 1,5 1,7 1,6 1,3 1,4 1,6 1 2,1 0,9 2 1,1 1,4 3,2 2,5 2,3 2,3 3,1 3,5 3,3 2,3 2,2 3,9 3,6 2,5 2,3 2,3 3,6 2 4,5 1,9 2,8 2,1 3,1 Uncharacterized PEP-dependent sugar PTS EIIC, probable galactitol specific Uncharacterized membrane protein CAAX amino terminal protease Uncharacterized membrane protein Basic membrane lipoprotein Transcriptional regulator protein RpiR Aldolase-type TIM barrel N-acetylmuramic acid 6-phosphate etherase Aldo/keto reductase Glucosamine-6-phosphate deaminase DNA binding domain, putative Transcriptional regulator Uncharacterized Beta-carotene 15,15-monooxygenase Protein of unknown function DUF1085 Foldase protein PrsA 2 Ornithine decarboxylase Acyl-CoA N-acyltransferase ABC transporter ATPase component DNA mismatch repair protein MutS, C-terminal 60 min 120 min LJ_0085 LJ_0086 LJ_0087 LJ_0129 LJ_0132 LJ_0240 LJ_0268 LJ_0302 LJ_0312 LJ_0313 LJ_0318 LJ_0319 LJ_0320 LJ_0326 LJ_0416 LJ_0490 LJ_0506 LJ_0512 LJ_0604 LJ_0605 LJ_0627 LJ_0628 LJ_0629 LJ_0630 LJ_0700 LJ_0745 LJ_0803 LJ_1012 LJ_1027 LJ_1028 LJ_1029 LJ_1030 LJ_1031 LJ_1035 LJ_1132 LJ_1156 LJ_1174 LJ_1211 LJ_1262 LJ_1299 LJ_1300 LJ_1301 LJ_1302 LJ_1320 LJ_1392 LJ_1415 LJ_1426 LJ_1429 LJ_1448 LJ_1451 LJ_1473 LJ_1476 LJ_1533 LJ_1625 LJ_1745 LJ_1746 LJ_1763 LJ_1827 30 min locus model profile Table S5.1B: Expression ratio of all genes with minimal twofold repression between any of the four timepoints (p<0.05). NCBI annotation is shown where available. Alternatively Gene Ontology classification of proteins or conserved domains is shown (in grey). Where no Gene Ontology classification could be assigned, the closest BlastP result is shown (displayed in grey, bold and italic). PTS: phosphotransferase system. PEP: phosphoenolpyruvate. NCBI annotation 15 27 11 9 26 23 11 9 34 9 22 9 11 38 34 11 26 22 11 26 15 11 25 11 11 26 26 26 15 11 11 11 11 22 26 9 26 26 9 26 26 11 34 11 11 9 20 11 26 26 11 11 26 34 27 11 26 14 0,8 1 0,7 0,8 0,8 0,8 0,7 0,6 1,2 1 0,8 0,8 0,8 1,3 1,1 0,9 1 0,8 0,8 0,9 0,8 0,7 0,9 0,7 0,8 0,9 0,9 0,8 0,7 0,9 0,8 0,8 0,8 0,8 1 1,1 1 1,1 1 1 1 0,6 1 0,8 0,7 1 0,5 0,8 1,1 1,1 0,8 0,8 0,9 1,1 0,9 0,8 1 0,7 1,1 1,3 0,9 0,5 0,9 0,5 0,4 0,3 1 0,7 0,5 0,5 0,8 1,5 0,9 0,7 0,8 0,4 0,9 1,1 1 0,8 0,4 0,8 0,9 0,9 0,8 0,9 0,9 0,6 0,7 0,8 0,7 0,5 0,9 0,6 1 0,9 0,7 0,8 0,8 0,5 0,7 0,5 0,5 0,7 1,5 0,8 1,2 1,1 0,6 0,8 0,9 0,9 1,3 0,9 0,8 0,5 0,5 0,6 0,4 0,2 0,4 0,5 0,1 0,1 0,6 0,4 0,6 0,5 0,5 0,7 0,6 0,4 0,4 0,5 0,4 0,5 0,5 0,5 1,9 0,5 0,4 0,4 0,4 0,4 0,5 0,4 0,3 0,4 0,5 0,7 0,5 0,3 0,5 0,5 0,5 0,5 0,5 0,2 0,5 0,2 0,2 0,4 1 0,4 0,5 0,5 0,2 0,5 0,5 0,5 0,5 0,4 0,5 1,6 Uncharacterized ABC transporter ATPase component ABC transporter permease component Phosphonate-binding periplasmic protein Phosphate/phosphonate ABC transporter permease component Transcriptional regulator 50S ribosomal protein L31 type B Lj965 prophage antirepressor Lj965 prophage scaffold protein Lj965 prophage protein Lj965 prophage protein Lj965 prophage major tail protein Lj965 prophage protein Lj965 prophage protein 50S ribosomal protein L7/L12 Xaa-Pro dipeptidase Seryl-tRNA synthetase Drug/metabolite transporter ABC transporter ATPase component ABC transporter permease protein BceB-type PEP-dependent sugar PTS EIIAB, probable mannose specific PEP-dependent sugar PTS EIIC, probable N-acetylgalactosamine specific PEP-dependent sugar PTS EIID, probable N-acetylgalactosamine specific Uncharacterized Glycerol uptake facilitator protein Dipeptide/tripeptide permease Probable xylulose-5-phosphate/fructose-6-phosphate phosphoketolase Probable GTP-binding protein EngB Glycosyl transferase, family 1 Glycosyl transferase, family 1 Glycosyl transferase, family 1 Glycosyltransferase Oligosaccharide repeat unit polymerase Acyltransferase 3 Two-component system response regulator Transcription regulator, MarR-type ATP-dependent DNA helicase RecQ Major facilitator superfamily permease CAT RNA-binding domain Uncharacterized ABC transporter permease ABC transporter ATPase component Transcription regulator, GntR Glycine--tRNA ligase alpha subunit Histidine--tRNA ligase SWEET sugar transporter Lj928 prophage protein Lj928 prophage protein Lj928 prophage protein Lj928 prophage recombination protein Amino acid transporter Sortase 50S ribosomal protein L28 Phenylalanine--tRNA ligase alpha subunit Surface protein, aggregation promoting factor Surface protein, aggregation promoting factor Carboxymuconolactone decarboxylase Purine permease PbuG-like 5 135 Chapter 6 General discussion and outlook This thesis addresses the consequences of oxygen and CO2 exposure on the metabolism and transcriptome profile of the lactic acid bacterium Lactobacillus johnsonii. These two gaseous components were found to have a major influence on metabolism, growth, gene expression, yield, and viability. Here we discuss several questions that logically follow from our findings and we suggest future experiments that could bring us closer to answering these questions. Furthermore, we discuss the relevance of the research presented in this thesis for the interactions between L. johnsonii and the mucosal surfaces of host organisms. CO2 growth dependency and cell death The growth requirement of L. johnsonii for CO2 in the absence of oxygen is one of the clear novel findings in this thesis. Previously, other LAB, including S. thermophilus and a subset of L. plantarum and L. lactis strains (251, 252) were shown to depend on an exogenous C-1 source under certain conditions. Given the similarities in LAB metabolism, we expect that CO2 dependency is a characteristic that is conserved among a variety of LAB. One of the transcriptional responses to CO2 depletion was the induction of the carbamoyl-phosphate pathway genes, reported in chapter 5. Regulation of this pathway was previously shown to play a role in governing CO2 and pyrimidine metabolism in L. plantarum (252, 263, 269). Expression of the pyr-operon was also found to be controlled in several other LAB by a variety of conditions. For example, the pyr-operon was repressed during bile and acid stress of L. rhamnosus GG (281, 282), coculturing of L. lactis with S. cerevisiae led to substantial repression of the pyr-operon in L. lactis (283), and growth of L. casei in ‘soy milk’ compared to bovine milk induced the expression of this operon (284). We consider that many of these environmental changes also include a change in the availability of an environmental C-1 supply; bile solutions may contain bicarbonate, and low pH influences the CO2/HCO3- equilibrium, while cocultivation with S. cerevisiae is expected to increase environmental CO2 levels. Taken together, we consider it likely that many of the environmental conditions known to control the expression of the pyr-operon in LAB, may in fact be responses to the availability to an appropriate C-1 source for growth, which thereby plays an important role in metabolic control in these bacteria. There are three distinctive characteristics of CO2 dependency that we report here, that were not described before. Firstly, in contrast to other LAB, the CO2 dependency of L. 138 johnsonii cannot be relieved by addition of pyrimidine nucleotides, arginine or aspartate. Even with addition of excessive amounts of these nucleotides and amino acids, the typical 2-fold difference in viability of L. johnsonii grown under pure nitrogen versus CO2-rich conditions could not be complemented. One of the possible causes may be a dysfunctional pathway for incorporation of exogenous uracil. An indication for this is the high level of expression (amongst the 200 most highly expressed genes in L. johnsonii) of the upp-encoded uracil phosphoribosyltransferase in normal, CO2-rich, conditions. This enzyme catalyzes the conversion of imported uracil to uracil monophosphate. Its high-level transcription would be expected in case L. johnsonii would depend on the continuous import of uracil for regular growth in MRS. However, preliminary studies with chemically defined medium from which uracil was omitted, did not reveal uracil auxotrophy in L. johnsonii in medium in which CO2 is not actively removed. These findings imply that the carbamoyl-phosphate pathway is functional in vivo. The metabolic mechanism underlying CO2 dependency in L. johnsonii deserves further exploration, since no supplementation strategy could be identified to complement this dependency by specific nutrients, which is clearly different from what has been found in various other LAB (see also chapter 5). A second characteristic that was not previously identified in LAB, is the substantial induction of cell death that was associated with CO2 depletion. Both in microcolonies (Anopore studies) and in liquid cultures, we observed that CO2 depletion led to loss of membrane integrity (as demonstrated by propidium iodide staining) and abolished the capacity to grow (as demonstrated by viability plating). This is a remarkable observation, since the absence of components required for growth is normally bacteriostatic rather than bactericidal. Following our hypothesis on the relation between CO2 and pyrimidine biosynthesis, we speculate that a depletion of the intracellular pyrimidine pool is 6 the mechanism underlying the lethality of CO2 depletion. Fatal DNA breaks due to disbalanced nucleotide pools is a well-known consequence of thymidine exhaustion in both prokaryotes and eukaryotes (275, 276), and such a mechanism may explain the CO2-related cell death in our experiments. This cell death is especially of relevance for the application of L. johnsonii in the food industry, in particular in its application as a probiotic. Survival of probiotic bacteria in industrial processes as well as in the final products can be considered as a prerequisite for the health beneficial effect they elicit in the consumer. It is an important notion that not only oxidative stress may lead to viability loss, but depletion of the available C-1 nutrients in the industrial and product environments may also be an important cause 139 for loss of bacterial (probiotic) viability. A third surprising characteristic of CO2 depletion was the relation to environmental oxygen, reported in chapter 4. In contrast to the expectations, bubbling a fermenter with N2 + O2 resulted in higher growth rates and biomass yields than bubbling with pure N2. Similarly, L. johnsonii displayed significantly higher growth rates in the absence of acetate under aerobic conditions as compared to anaerobic conditions. This growth stimulatory effect of oxygen was shown to be dependent on pyruvate oxidase-mediated CO2 and acetate production, since it could be abolished in a pox deletion derivative of L. johnsonii. These findings imply that oxygen, besides its deleterious consequences in terms of hydrogen peroxide production (see below), also has beneficial effects by reducing the fastidious growth requirements of L. johnsonii. The role of NFR and NOX in aerotolerance of Lactobacillus johnsonii. One of the most prominent differences between aerobic and anaerobic growth of L. johnsonii is the production of substantial amounts of H2O2. Accumulation of this H2O2 results in an approximately 10-fold lower biomass yield in the presence of oxygen, due to premature H2O2-induced growth stagnation. Continuous removal of H2O2 through the addition of exogenous catalase prevents aerobic growth stagnation. In chapter 1 and 2 we discuss the role of two enzymes, NOX and NFR, in H2O2 production of L. johnsonii. Based on our findings we propose that NFR is constitutively expressed, whereas NOX is expected to complement NFR after longer-term oxygen exposure. This redundancy for H2O2 production capacity, in combination with the strongly increased oxygen sensitivity observed in a L. johnsonii derivative that lacks both NFR and NOX, implies that NADH oxidation, oxygen consumption and/or hydrogen peroxide production are important to sustain aerotolerance and to allow aerobic growth of L. johnsonii. However, our experimental approaches did not directly enable the identification of the precise role of H2O2 production in the overall aerobic physiology of L. johnsonii, and below we discuss some of the possible mechanistic explanations. Can H2O2:NADH oxidases confer aerotolerance by scavenging oxygen? One of the most striking observations on the role of the H2O2 producing enzymes NOX and NFR, is that they appear to be essential for aerotolerance of L. johnsonii: a deletion of the genetic loci that encode these enzymes resulted in a strain that 140 displayed significantly lower growth rate and final OD under aerobic conditions. This finding provides an interesting paradox; H2O2 production is the main source of growth inhibiting stress under aerobic conditions, but at the same time appears to be essential for growth under these conditions. Preliminary experiments comparing oxygen consumption rates in wild type L. johnsonii and its Δnfr and Δnfr Δnox derivatives, establish that nfr and nox encode the main oxygen scavenging capacity in L. johnsonii, since the deletion of both genes led to a complete elimination of oxygen consumption. Based on these results and the other findings presented in this thesis, we postulate that NFR and NOX may contribute to aerotolerance by scavenging oxygen. In this proposition, we consider the possibility that molecular oxygen and possibly superoxide derivatives resulting from the spontaneous autoxidation of cellular components, may under certain conditions be more damaging than H2O2. The most predominant damaging reaction in bacteria due to H2O2 is the result of Fenton chemistry, demonstrated by the effectivity of iron chelators to reduce H2O2-induced cell death (131, 157, 285). L. johnsonii has a remarkably low number of enzymes that are predicted to contain an iron-sulfur cluster (3 vs 145 in E. coli). If L. johnsonii maintains very low intracellular iron pools, similar to other LAB (151, 286), hydroxyl formation through Fenton chemistry would be rare. Thereby, the cellular make-up of L. johnsonii would render it intrinsically tolerant against low-levels of H2O2. Moreover, we assume that in the natural habitat in which this type of lactobacilli are encountered, they are only transiently exposed to limited amounts of oxygen. Oxygen is thought to freely diffuse over the bacterial membrane, driven by the intra- and extra-cellular concentration difference. Once inside the cell, oxygen could react with cellular components such as flavins, resulting in superoxide production and launching a cascade of damage inducing reactions. However, superoxide formation and its 6 detrimental consequences could be prevented by the effective scavenging of oxygen by NADH oxidases that convert it into H2O2. The small, uncharged H2O2 molecule can freely diffuse out of the cell as long as the extracellular H2O2 levels remain low (287). In a crowded environment such as the intestinal microbiota, where L. johnsonii co-occurs with numerous catalase and peroxidase producing bacteria, one could plausibly assume that the released H2O2 would be quickly scavenged and is unlikely to accumulate to substantial levels. The argumentation raised above could explain how in certain environments H2O2producing NADH oxidation could contribute to aerotolerance, by the prevention of 141 superoxide mediated damage, but at the same time preventing the excessive accumulation of H2O2 by exploiting the ecosystem’s H2O2 defusing capacity. Consequently, the absence of elaborate oxidative stress defense mechanisms in L. johnsonii may be a result of its adaptation to its natural habitat, where it expresses only a minimal defense against the limited oxygen exposure, and exploits its environment for effective detoxification of the H2O2 it produces, which is relatively nontoxic to L. johnsonii anyway. The argumentation raised above could explain how in certain environments H2O2producing NADH oxidation could contribute to aerotolerance, by the prevention of superoxide mediated damage, but at the same time preventing the excessive accumulation of H2O2 by exploiting the ecosystem’s H2O2 defusing capacity. Consequently, the absence of elaborate oxidative stress defense mechanisms in L. johnsonii may be a result of its adaptation to its natural habitat, where it expresses only a minimal defense against the limited oxygen exposure, and exploits its environment for effective detoxification of the H2O2 it produces, which is relatively undamaging to L. johnsonii anyway. The notion that LAB use oxidases to scavenge oxygen from its environment is supported by two studies. Rezaiki et al. showed that L. lactis grown under respiratory-permissive condition in Erlenmeyer-flasks can effectively create an anaerobic environment, indicated by low GFP fluorescence and ceased H2O2 production (53). In addition, Gibson et al. demonstrate that the deletion of a protein identified as a water-forming NOX in S. pyogenes resulted in a H2O2 producing strain that displayed a severe aerobic growth defect. One of the possible explanations of this observation is that the protein that was annotated as NOX is in fact an NADH peroxidase, since it is difficult to distinguish between these enzymes (see above; Table S1.1). However, if the enzyme was correctly identified as an NADH oxidase, it could scavenge oxygen to a level that is sufficient to prevent H2O2 production via the aerobic lactate utilization pathway (288). We propose to explore this hypothesis by the following set of experiments: • a careful assessment of the toxicity of the different reactive oxygen species, for instance by engineering superoxide dismutase and catalase expressing variants of L. johnsonii. • an analysis of the intracellular iron pools, or the total iron content compared to species that do express ROS-scavenging enzymes. This could provide support for the hypothesis that L. johnsonii can withstand low H2O2 levels due to its intrinsic resistance against H2O2 damage. 142 • quantification of superoxide production in cultures and cell free extracts of L. johnsonii that lack both NOX and NFR (nox,nfr deletion derivative) could provide support for the theory that oxygen scavenging by NOX and NFR prevents superoxide production. Interestingly, preliminary experiments with the introduction of plasmid borne expression of a SOD gene from L. paracasei in wildtype L. johnsoni and its Δnfr and Δnfr Δnox derivatives, indicated that SOD expression recovers H2O2 production in the Δnfr Δnox background, indicating that these cells contain superoxide radicals that act as substrate in the SOD catalized reaction leading to H2O2 production. Further experiments should include a comparison of aerotolerance levels in these strains. In case SOD expressions indeed improves aerotolerance in the Δnfr Δnox mutant, this would substantiate the proposed primary role of the H2O2 forming NADH oxydases in oxygen scavenging to prevent the formation of the more detrimental superoxide. Perspectives: the relevance of ROS in host/microbiota interactions The experiments that are presented in this thesis observe L. johnsonii in a relatively unnatural single-species environment which provides useful insights in the metabolism and physiology of L. johnsonii, but neglects all metabolic and physiological effects in complex microbial communities associated with mucosal surfaces that can be considered as its natural habitat. Here, we would like to take a broader perspective on bacterial H2O2 production and discuss how it may affect host/microbe interactions. Role of ROS in host defense and signaling Reactive oxygen species have a central role in the non-specific innate immune response. 6 Pathogenic bacteria are engulfed in phagocytic vesicles of dedicated immune cells such as macrophages and neutrophils. Activity of phagocytic-NADPH oxidases (Nox2) generates high levels of superoxide, referred to as respiratory or oxidative burst. The high levels of superoxide results in the formation of secondary ROS such as H2O2 and hypochlorous acid (HOCL) collectively leading to the death of the phagocytized bacterial cell. Besides this dedicated anti-bacterial use of ROS, H2O2 produced in non-phagocytic tissue by NOX (1,3,5) and DOUX (1,2) plays a role as signaling molecules in a variety of pathways. In many of these processes H2O2 does not transduce the signal directly but oxidizes thiol 143 peroxidases, thioredoxin or glutathione that function as secondary messengers. The cysteine residues of many regulatory enzymes, such as tyrosine phosphatases, protein kinase phosphatases, NF-κB and ubiquitins are sensitive to oxidation by these thiols and have been reported to be modulated by such redox signaling. Additionally, H2O2 produced by epithelial NOX can also interfere with signaling processes in pathogenic bacteria such as Campylobacter jejuni, where the oxidation of a tyrosine kinase prevents capsule formation, which is an important virulence factor (289). NOX-related ROS production serves a variety of roles: it can act directly antibacterial, and simultaneously recruit an immune response by activating inflammasome, cytokines and prostaglandins (290). Dysregulation of this process can result in excessively high levels of ROS production and can induce hyperinflammation, which is a typical characteristic encountered in Inflammatory bowel disease (IBD). Hyperinflammation cascades induce mucosal tissue morphological destruction, including the formation of lesions, ulcerations and fibrosis (291). Interestingly, the oral administration of LAB that overexpress superoxide dismutase or catalase has been shown to attenuate disease symptoms in experimental animal models for IBD, like the chemically induced rodent (mouse or rat) colitis models (138, 292, 293). At the same time, ROS-producing NOX, provides negative feedback on this sytem and plays an important role in suppressing the inflammatory cascades in inflamed tissue to recover its homeostatic, non-inflamed status. A defect in the Nox2 causes chronic granulomatous disease in humans, which is not only characterised by recurrent infections, but also by hyperinflammation. Similarly, Nox2 expression was found to play an essential role in the prevention of insulin resistance and diet-induced obesity in mice. These apparently dualistic roles of ROS indicate that its effect is highly dependent on its temporal, spatial and quantitative production pattern (294). Role of ROS in host/microbiota interactions The epithelial cells that line the digestive tract are continuously exposed to enormous amounts of bacteria: ranging from ~103/g in the upper parts of the small intestine to ~1012/gram in the colon. It is of utmost importance that immune responses are balanced and appropriately eradicate pathogenic and invading bacteria, whereas they should not respond excessively to the bulk of commensal bacteria in the gut. An intriguing question is how H2O2 produced by bacteria may impact gut homeostasis and host signal transduction. Essential to bacterial H2O2 to emerge and affect host 144 immune responses is the presence of sufficient oxygen in the GI-tract. Studies on the oxygen content demonstrate that the majority of the gut volume is anaerobic, due to continuous scavenging of oxygen by the microbiota. Oxygen is continuously leaking in from epithelial tissue, creating a very steep gradient from the mucosal surface to the lumen (40, 41). Another gradient appears to be present in which the proximal regions of the small intestine (duodenum and jejunum) are considered to contain higher levels of oxygen compared to the strictly anaerobic colon (196). Oxygen availability in the micro-environments of the intestinal tract may be one of the dominant drivers of the spatial distribution of microbial specialists, in which microaerophilic microbes are generally found in samples taken close to the mucosal surfaces whereas the obligate anaerobic species are mostly found in the more anaerobic environments of the lumen and the colon (41, 295, 296). There are several studies detailing how intestinal oxygen levels impact bacterial metabolism, including the modulation of pathogenicity factors of Shigella (41), respiration of E. coli (297) and thiol/flavin export by Faecalibacterium prauznitsi (298). We consider that aerotolerance is an important factor for bacteria to reside in close proximity of epithelial surfaces, which can be considered as an important driver for direct modulation of host immune responses. In such environments with fluctuating oxygen levels, ROS produced by LAB, could potentially have an impact on health and disease in the host. In in vitro cell culture systems it was shown that superoxide produced by E. faecalis leads to host-cell lipid oxidation generating the reactive compound 4-hydroxy-2-nonenal which induces DNA damage (62, 299). Also, H2O2-producing streptococci, such as S. pyogenes, S. mutans and S. pneumoniae are associated with disease in humans. S. pneumoniae is regularly found amongst the upper-respiratory tract microbiota of healthy individuals, but can cause bacteremia, meningitis and pneumonia in immunocompromised individuals. 6 S. pyogenes can show a similar transformation from harmless commensal to lethal pathogen, whereas S. mutans and S. gordoni are mostly associated with dental caries. As we described in chapter 1, H2O2 production in these species is mostly catalyzed by the lactate and pyruvate oxidation pathways (LOX, POX, ACK). Importantly, expression of the pyruvate oxidase encoding spxB gene has been identified as an important factor for virulence of S. pneumoniae (88, 90, 300-302). Similarly, SpxB and H2O2 production was also correlated with the ability of oral streptococci to adhere to the tooth surface and release DNA, which is an important factor for biofilm formation and natural competence (85, 303-306), although the exact mechanism behind these correlations remains unresolved. Bacterial H2O2 production could play a role in pathogenesis on basis 145 of its cytotoxic effect on epithelial tissue, but at the same time the bacterial production pathways are of importance for their aerotolerance and oxidative stress resistance. Other studies indicate that modulation of characteristics like aerobic respiration or expression of ROS-scavenging enzymes are also considered as virulence factors, while they also contribute to survival of the bacterial cell (57, 116, 117). Thereby, it becomes difficult to distinguish between a direct role in the virulence cascade and the ability to persist in the host environment. Obviously, the latter capacity is a prerequisite to allow bacteria to exert their detrimental and pathogenic effects, creating a intrinsic dilemma in the design of adequate experiments to discriminate between direct, and persistence-driven effects on pathogenesis. In many of the aforementioned cases, bacterial ROS production is associated with adverse health outcomes. However, there are several reports indicating that bacterial H2O2 may also have a beneficial, anti-inflammatory effect. One study, by Voltan et al., shows that L. crispatus derived H2O2 induces PPAR-g expression in epithelial cell cultures. Such increased PPAR-g expression was previously associated with improved gut homeostasis and reduced severity of colitis (201), through its capacity to decrease NF-κB expression and thereby suppress inflammatory cascades. Administration of H2O2-producing L. crispatus in mice, could substantially reduce severity of (chemically induced) colitis, whereas a low hydrogen-peroxide producing (spontaneous) mutant of L. crispatus did not show such an effect. This study illustrated in multiple ways that H2O2 is essential for PPAR-regulation. However, since the spontaneous low-level-H2O2 producing mutants failed to adhere to the epithelial cells while the wild-type strain adhered effectively, the prerequisite of close proximity of the bacteria and the epithelia may have been a strong confounder in this study. Thereby, secondary mechanisms to explain the differences between the two bacterial strain that are independent of their H2O2 production capacity can not be excluded to play a role in the observed protection against colitis (50). Another study described the effect of L. johnsonii on rats that are genetically predisposed to develop type 1 diabetes (T1D), which to a certain extent could also be linked to the bacterial capacity to produce H2O2. The L. johnsonii strain used in this study, strain N6.2 was previously found to be overrepresented in the microbiota of T1D-susceptible rats that did not develop diabetes (307). Administration of this isolate to the rats was shown to increase a variety of mucosal proteins associated with gut barrier function and decrease the inflammatory cytokine IFNγ. Furthermore, it decreased the oxidative stress response and the overall incidence of T1D in this rat model, while these effects 146 were not seen with another Lactobacillus strain (of the species L. reuteri) that also negatively correlated with type 1 diabetes symptoms (308). Interestingly, L. johnsonii administration was shown to inhibit the expression of indoleamine 2,3-dioxygenase (IDO) in host ileal mucosa. IDO is primarily involved in tryptohan catabolism, but also plays a central role in inflammation cascades, and its activity has been associated with several adverse health outcomes (309). L. johnsonii administration was shown to lead to higher ileal H2O2 levels, and in in vitro cell cultures it could be shown that H2O2 (produced by L. johnsonii) abolished IDO activity. Taken together these results create a tentative connection between the IDO inhibition and the H2O2 producing capacity of this bacterium in vivo, although it can not be excluded that the elevated ileal H2O2 levels were (in part) host-derived, and thus may not have depended (exclusively) on the production by L. johnsonii (203). Nevertheless, this study provides a strong indication that bacterially produced H2O2 could modulate IDO expression in vivo, and it would be very interesting to evaluate the effect of the L. johnsonii mutants described in this thesis that are unable to produce H2O2 in this rat model. Collectively, these studies indicate that H2O2 may act as a bacterial effector molecule that can evoke immune modulations in the host. We hypothesize that analogous to hostrelated H2O2 production, the effect of bacterial H2O2 depends on its temporal, spatial and quantitative release in the GI-tract. Intestinal conditions that stimulate high level production of ROS by bacteria could potentially induce epithelial and mucosal damage, whereas low-level production of ROS by bacteria like L. johnsonii may successfully suppress excessive host immune responses by modulating the expression of central metabolic regulatory nodes, such as PPAR-g and IDO that have established roles in metabolism-immune cross talk. 6 Relevance of bacterial H2O2 in the vaginal microbiota Compared to the oral and intestinal microbiota, where ROS-producing bacteria form only a small subset of the overall community, the vaginal microbiota stands out by its high abundance of such bacteria. The vaginal microbiota of the vast majority of women in the reproductive age is dominated by one (or more) of four different species (L. crispatus, L. gasseri, L. jensenii and L. iners) belonging to the L. acidophilus group, of which at least three (L. gasseri, jensenii and crispatus) can produce substantial amounts of H2O2. Although colonization patterns vary as a function of the menstrual cycle, health and disease and are influenced by sexual intercourse and pregnancy, all major studies that have analyzed the constituents of the vaginal microbiota identify this 147 lactobacilli dominated ecosystem in healthy women (310-313). In addition, the absence of lactobacilli in this ecosystem is often accompanied by overgrowth of a collection of opportunistic species such as Gardnerella vaginalis and Atopobium vaginae (314316). This microbial disbalance is referred to as bacterial vaginosis (BV), which affects approximately 10-20% of European women (317). Women with BV-type microbiota are also at a significant higher risk to acquire sexually transmitted disease, and increased risk of womb infections that can elicit preterm births (199, 200, 318, 319). The lactobacilli-dominated microbiota of the vagina provide an intriguing contrast to the mucosal surfaces of the digestive tract that are colonized by tens to hundreds of different species, none of which is really dominating the ecosystem. Conversely, the vaginal environment is colonized by only a handful of dominant species with very similar features, including their capacity for H2O2 production (94, 187, 320). A longstanding hypothesis states that bacterial H2O2 production in the vaginal niche is the molecular trait by which lactobacilli prevent the colonization of other organisms (319, 321). However, this hypothesis is debated, and as an alternative explanation it has been proposed that the high levels of lactic acid in combination with a low pH may be more effective in creating colonization resistance. In addition, the oxygen availability, as well as the stability of H2O2 in the vaginal environment have been contested (312, 322). The precise role of bacterial H2O2 in vaginal homeostasis remains to be determined. The identification and mutation of the H2O2 producing enzymes in a species closely related to vaginal lactobacilli, that we presented in this thesis, may advance our understanding of this topic. We envision that a comparison between the ability of wildtype and nonH2O2 producing lactobacilli to prevent bacterial vaginosis in animal models could help understand the influence of this physiology on stability of the vaginal microbiota. Relevance of bacterial H2O2 in the neonatal gut The composition of the vaginal microbiota is not only related to health of women, but may also contribute to the health of an infant after vaginal delivery. Although it has been proposed that fetuses already acquire microbes prenatally (323), the predominant colonization of the infant gut takes place after birth. The vaginal bacteria of the birth canal may present an important part of microbial transfer between mother and infant. Comparison of microbial constituents of the neonatal gut following vaginal delivery with those found after caesarian delivery, revealed prominent differences and supported the idea that the vaginal (and/or fecal) microbes of the mother are amongst the pioneer colonizers of the neonatal gut after birth (16, 324, 325). Interestingly, these 148 lactobacilli containing early microbial colonization patterns are inversely correlated with the prevalence of necrotizing enterocolitis (NEC) in preterm infants (326, 327). The administration of probiotic lactobacilli has been shown to be effective in lowering the incidence of NEC (25). Oxygen levels are generally assumed to show a significant drop during subsequent phases of colonization of the infant gut, which is reflected by the aerotolerance levels of the colonizing species. The first microbial groups that are found in the neonatal gut in the first 48h after birth are mostly aerotolerant, facultative or obligate aerobic species (16). Outgrowth of the strictly anaerobic groups, such as clostridiae, faecalibacteria and sulfate-reducing bacteria, is typically found at later stages in the development of the microbiota (328). Although this dynamic colonization is influenced by many parameters, the establishment of the aforementioned oxygen gradients in the developing intestinal environment may play an important role in suppressing outgrowth of the more aerosensitive members of the microbiota. Some authors even suggest that oxygen consumption by the first aerobic colonizers is essential to pave the way for colonization of their anaerobic successors (40). The observations described above raise the question whether H2O2 produced in the neonatal gut by the vaginal lactobacilli that were transferred during passage through the birth canal, may play a role in initiating or shaping the early immune system. This hypothesis is based on three assumptions: (1) that the absence of high numbers of bacteria lead to high oxygen levels in the GI-tract during the first hours after birth, (2) that mother-to-child microbial transfer leads to prominent colonization of the infant gut with H2O2-producing lactobacilli and (3) that bacterial H2O2 can modulate host-immune reactions. The third assumption is supported by observations described above that highlight the potential role of bacterially produced H2O2 in signaling to the mucosal via 6 PPAR and IDO. Moreover, the capacity to scavenge oxygen could help the colonization of less aerotolerant species such as the bifidobacteria that are typically overrepresented in the intestinal microbiota of breastfed infants. However, one can also not rule out that the bacterially produced H2O2 may in certain cases also have adverse effects, since the presence of high intestinal oxygen levels and the absence of catalase expressing bacteria in the infant intestine may also cause ROS induced mucosal damage. Also here, small amounts may go a long way. The balance between ROS-production and ROSscavenging are key in maintaining a healthy intestinal environment. 149 Perspectives: how the findings in this thesis could help unravel the role of bacterial H2O2 in host/microbe interactions An important bottleneck in studying the effect of ROS in immune responses is the difficulty in teasing apart host and bacterially derived H2O2. Adequate negative controls that exclusively eliminate bacterial H2O2 from the interplay are currently not available. Especially the effects of ecosystem-derived catalase and SOD may scavenge both host and bacterially-derived ROS and may influence the colonization by specific microbial groups including the lactobacilli. A promising approach to resolve this issue was followed by Voltan et al., who used a spontaneous non-H2O2 producing isolate of L. crispatus (strain MU5) as a negative control. Unfortunately, this isolate was affected in various phenotypic traits besides its H2O2-producing capacity, including a non-aggregating phenotype and the inability to adhere to epithelial tissue cells (329). These deviations may be of critical importance for the in vivo effect that these bacteria could elicit in the intestinal tract of a host model, which was also supported by the finding that epithelial adherence was essential for the immunomodulatory capacity of L. crispatus. The identification of the H2O2 producing enzymes in L. johnsonii and the availability of deficient mutants, opens novel avenues to further study the relevance of bacterial H2O2 in various host/microbiota interactions. The isogenic strains that are no longer able to produce H2O2, developed and studied in this thesis, NCC 9359 (Δnfr) and NCC 9360 (Δnfr, Δnox) could be instrumental in studies with epithelial tissue cultures and/or animal models. Moreover, similar mutants may more readily be constructed in other members of the L. acidophilus group, now that key-enzymes involved in H2O2 production (NFR and NOX) are identified in this group of bacteria. Inversely, H2O2-overproducing variants of L. johnsonii or other members of the L. acidophilus group could also be of interest. Although transformation with NFR encoding multi-copy plasmids (Chapter 2) did not lead to elevated H2O2 production by L. johnsonii in our experiments, it can not be excluded that alternative expression systems or specific (micro-aerobic) environments may allow the construction or exploitation of H2O2 overproducing strains in the same cell-based or animal models. 150 References 1. de Kruif P. 1926. Microbe hunters. 2. Sebald M, Hauser D. 1995. Pasteur, oxygen and the anaerobes revisited. Anaerobe. 1:11-16. 3. Naqui A, Chance B, Cadenas E. 1986. Reactive oxygen intermediates in biochemistry. Annu. Rev. Biochem. 55:137-166. doi: 10.1146/annurev.bi.55.070186.001033. 4. Imlay JA. 2013. The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium. Nat. Rev. Microbiol. 11:443-454. doi: 10.1038/nrmicro3032. 5. Flint DH, Tuminello JF, Emptage MH. 1993. The inactivation of Fe-S cluster containing hydro-lyases by superoxide. J. Biol. Chem. 268:22369-22376. 6. Imlay JA. 2003. Pathways of oxidative damage. Annu. Rev. Microbiol. 57:395-418. doi: 10.1146/annurev. micro.57.030502.090938. 7. Mishra S, Imlay J. 2012. Why do bacteria use so many enzymes to scavenge hydrogen peroxide? Arch. Biochem. Biophys. 525:145-160. doi: 10.1016/j.abb.2012.04.014. 8. Imlay JA. 2008. Cellular defenses against superoxide and hydrogen peroxide. Annu. Rev. Biochem. 77:755-776. doi: 10.1146/annurev.biochem.77.061606.161055. 9. Imlay JA. 2011. Redox pioneer: professor Irwin Fridovich. Antioxid. Redox Signal. 14:335-340. doi: 10.1089/ars.2010.3264. 10. Schnell DM, St Clair D. 2014. Redox Pioneer: Professor Joe M. McCord. Antioxid. Redox Signal. 20:183188. doi: 10.1089/ars.2013.5291. 11. McCord JM, Keele BB,Jr, Fridovich I. 1971. An enzyme-based theory of obligate anaerobiosis: the physiological function of superoxide dismutase. Proc. Natl. Acad. Sci. U. S. A. 68:1024-1027. 12. Panek HR, O’Brian MR. 2004. KatG is the primary detoxifier of hydrogen peroxide produced by aerobic metabolism in Bradyrhizobium japonicum. J. Bacteriol. 186:7874-7880. doi: 10.1128/JB.186.23.78747880.2004. 13. Cosgrove K, Coutts G, Jonsson IM, Tarkowski A, Kokai-Kun JF, Mond JJ, Foster SJ. 2007. Catalase (KatA) and alkyl hydroperoxide reductase (AhpC) have compensatory roles in peroxide stress resistance and are required for survival, persistence, and nasal colonization in Staphylococcus aureus. J. Bacteriol. 189:10251035. doi: 10.1128/JB.01524-06. 14. Ohara N, Kikuchi Y, Shoji M, Naito M, Nakayama K. 2006. Superoxide dismutase-encoding gene of the obligate anaerobe Porphyromonas gingivalis is regulated by the redox-sensing transcription activator OxyR. Microbiology. 152:955-966. doi: 10.1099/mic.0.28537-0. 15. Kawasaki S, Watamura Y, Ono M, Watanabe T, Takeda K, Niimura Y. 2005. Adaptive responses to oxygen stress in obligatory anaerobes Clostridium acetobutylicum and Clostridium aminovalericum. Appl. Environ. Microbiol. 71:8442-8450. doi: 10.1128/ AEM.71.12.8442-8450.2005. 16. Karlsson CL, Molin G, Cilio CM, Ahrne S. 2011. The pioneer gut microbiota in human neonates vaginally born at term-a pilot study. Pediatr. Res. 70:282-286. doi: 10.1203/PDR.0b013e318225f765. 17. Martin R, Jimenez E, Heilig H, Fernandez L, Marin ML, Zoetendal EG, Rodriguez JM. 2009. Isolation of bifidobacteria from breast milk and assessment of the bifidobacterial population by PCRdenaturing gradient gel electrophoresis and quantitative real-time PCR. Appl. Environ. Microbiol. 75:965-969. doi: 10.1128/AEM.02063-08. 18. Fernandez L, Langa S, Martin V, Maldonado A, Jimenez E, Martin R, Rodriguez JM. 2013. The human milk microbiota: origin and potential roles in health and disease. Pharmacol. Res. 69:1-10. doi: 10.1016/j.phrs.2012.09.001. 19. Krzysciak W, Jurczak A, Koscielniak D, Bystrowska B, Skalniak A. 2013. The virulence of Streptococcus mutans and the ability to form biofilms. Eur. J. Clin. Microbiol. Infect. Dis. . doi: 10.1007/s10096-013-1993-7. 7 151 20. Aas JA, Paster BJ, Stokes LN, Olsen I, Dewhirst FE. 2005. Defining the normal bacterial flora of the oral cavity. J. Clin. Microbiol. 43:5721-5732. doi: 10.1128/JCM.43.11.5721-5732.2005. 21. Johnston BC, Ma SS, Goldenberg JZ, Thorlund K, Vandvik PO, Loeb M, Guyatt GH. 2012. Probiotics for the prevention of Clostridium difficile-associated diarrhea: a systematic review and meta-analysis. Ann. Intern. Med. 157:878-888. 22. Hempel S, Newberry SJ, Maher AR, Wang Z, Miles JN, Shanman R, Johnsen B, Shekelle PG. 2012. Probiotics for the prevention and treatment of antibiotic-associated diarrhea: a systematic review and metaanalysis. JAMA. 307:1959-1969. doi: 10.1001/jama.2012.3507. 23. Allen SJ, Martinez EG, Gregorio GV, Dans LF. 2010. Probiotics for treating acute infectious diarrhoea. Cochrane Database Syst. Rev. (11):. doi: 10.1002/14651858.CD003048.pub3;. 24. Pattani R, Palda VA, Hwang SW, Shah PS. 2013. Probiotics for the prevention of antibiotic-associated diarrhea and Clostridium difficile infection among hospitalized patients: systematic review and meta-analysis. Open Med. 7:e56-67. 25. Bernardo WM, Aires FT, Carneiro RM, Sa FP, Rullo VE, Burns DA. 2013. Effectiveness of probiotics in the prophylaxis of necrotizing enterocolitis in preterm neonates: a systematic review and meta-analysis. J. Pediatr. (Rio J). 89:18-24. doi: 10.1016/j.jped.2013.02.004. 26. Foolad N, Brezinski EA, Chase EP, Armstrong AW. 2013. Effect of nutrient supplementation on atopic dermatitis in children: a systematic review of probiotics, prebiotics, formula, and fatty acids. JAMA Dermatol. 149:350-355. 27. van de Guchte M, Serror P, Chervaux C, Smokvina T, Ehrlich SD, Maguin E. 2002. Stress responses in lactic acid bacteria. Antonie Van Leeuwenhoek. 82:187-216. 28. Van Tassell ML, Miller MJ. 2011. Lactobacillus adhesion to mucus. Nutrients. 3:613-636. doi: 10.3390/ nu3050613. 29. Bron PA, Tomita S, Mercenier A, Kleerebezem M. 2013. Cell surface-associated compounds of probiotic lactobacilli sustain the strain-specificity dogma. Curr. Opin. Microbiol. 16:262-269. doi: 10.1016/j. mib.2013.06.001. 30. Bron PA, van Baarlen P, Kleerebezem M. 2011. Emerging molecular insights into the interaction between probiotics and the host intestinal mucosa. Nat. Rev. Microbiol. 10:66-78. doi: 10.1038/nrmicro2690. 31. Yamada T, Takahashi-Abbe S, Abbe K. 1985. Effects of oxygen on pyruvate formate-lyase in situ and sugar metabolism of Streptococcus mutans and Streptococcus sanguis. Infect. Immun. 47:129-134. 32. Condon S. 1987. Responses of lactic acid bacteria to oxygen. FEMS Microbiology Reviews. 46:269-269280. 33. Melchiorsen CR, Jokumsen KV, Villadsen J, Johnsen MG, Israelsen H, Arnau J. 2000. Synthesis and posttranslational regulation of pyruvate formate-lyase in Lactococcus lactis. J. Bacteriol. 182:4783-4788. 34. Sedewitz B, Schleifer KH, Gotz F. 1984. Physiological role of pyruvate oxidase in the aerobic metabolism of Lactobacillus plantarum. J. Bacteriol. 160:462-465. 35. Tawalkar A, Kailasapathy L. 2004. The role of oxygen in the viability of probiotic bacteria with reference to Lactobacillus acidophilus and Bifidobacterium spp. Curr. Issues Intest. Microbiol. 5:1-8. 36. Villegas E, Gilliland SE. 2006. Hydrogen Peroxide Production by Lactobacillus delbrueckii Subsp. Lactis I at 5°C. Journal of Food Science. 63:1070-1074. 37. Marty-Teysset C, de la Torre F, Garel J. 2000. Increased production of hydrogen peroxide by Lactobacillus delbrueckii subsp. bulgaricus upon aeration: involvement of an NADH oxidase in oxidative stress. Appl. Environ. Microbiol. 66:262-267. 38. Pridmore RD, Pittet AC, Praplan F, Cavadini C. 2008. Hydrogen peroxide production by Lactobacillus johnsonii NCC 533 and its role in anti-Salmonella activity. FEMS Microbiol. Lett. 283:210-215. doi: 10.1111/j.1574-6968.2008.01176.x. 152 39. Lucey CA, Condon S. 1986. Active role of oxygen and NADH oxidase in growth and energy metabolism of Leuconostoc. J. Gen. Microbiol. 132:1789--1796. 40. Espey MG. 2013. Role of oxygen gradients in shaping redox relationships between the human intestine and its microbiota. Free Radic. Biol. Med. 55:130-140. doi: 10.1016/j.freeradbiomed.2012.10.554. 41. Marteyn B, West NP, Browning DF, Cole JA, Shaw JG, Palm F, Mounier J, Prévost M, Sansonetti P, Tang CM. 2010. Modulation of Shigella virulence in response to available oxygen in vivo. Nature. . doi: 10.1038/nature08970. 42. Saldena TA, Saravi FD, Hwang HJ, Cincunegui LM, Carra GE. 2000. Oxygen diffusive barriers of rat distal colon: role of subepithelial tissue, mucosa, and mucus gel layer. Dig. Dis. Sci. 45:2108-2114. 43. Watson ME,Jr, Nielsen HV, Hultgren SJ, Caparon MG. 2013. Murine vaginal colonization model for investigating asymptomatic mucosal carriage of Streptococcus pyogenes. Infect. Immun. 81:1606-1617. doi: 10.1128/IAI.00021-13. 44. Wagner G, Bohr L, Wagner P, Petersen LN. 1984. Tampon-induced changes in vaginal oxygen and carbon dioxide tensions. Am. J. Obstet. Gynecol. 148:147-150. 45. Hill DR, Brunner ME, Schmitz DC, Davis CC, Flood JA, Schlievert PM, Wang-Weigand SZ, Osborn TW. 2005. In vivo assessment of human vaginal oxygen and carbon dioxide levels during and post menses. J. Appl. Physiol. 99:1582-1591. doi: 10.1152/japplphysiol.01422.2004. 46. Miller CW, Nguyen MH, Rooney M, Kailasapathy K. 2002. The influence of packaging materials on the dissolved oxygen content of probiotic yoghurt. Packag. Technol. Sci. 15:133-138. 47. Mills S, Stanton C, Fitzgerald GF, Ross RP. 2011. Enhancing the stress responses of probiotics for a lifestyle from gut to product and back again. Microb. Cell. Fact. 10 Suppl 1:S1--S19. doi: 10.1186/14752859-10-S1-S19. 48. van Niel EW, Hofvendahl K, Hahn-Hagerdal B. 2002. Formation and conversion of oxygen metabolites by Lactococcus lactis subsp. lactis ATCC 19435 under different growth conditions. Appl. Environ. Microbiol. 68:4350-4356. 49. Herve-Jimenez L, Guillouard I, Guedon E, Boudebbouze S, Hols P, Monnet V, Maguin E, Rul F. 2009. Postgenomic analysis of Streptococcus thermophilus cocultivated in milk with Lactobacillus delbrueckii subsp. bulgaricus: involvement of nitrogen, purine, and iron metabolism. Appl. Environ. Microbiol. 75:20622073. doi: 10.1128/AEM.01984-08. 50. Voltan S, Martines D, Elli M, Brun P, Longo S, Porzionato A, Macchi V, D’Inca R, Scarpa M, Palu G, Sturniolo GC, Morelli L, Castagliuolo I. 2008. Lactobacillus crispatus M247-derived H2O2 acts as a signal transducing molecule activating peroxisome proliferator activated receptor-gamma in the intestinal mucosa. Gastroenterology. 135:1216-1227. doi: 10.1053/j.gastro.2008.07.007. 51. Brooijmans RJ, de Vos WM, Hugenholtz J. 2009. Lactobacillus plantarum WCFS1 electron transport chains. Appl. Environ. Microbiol. 75:3580-3585. doi: 10.1128/AEM.00147-09. 52. Brooijmans R, Smit B, Santos F, van Riel J, de Vos WM, Hugenholtz J. 2009. Heme and menaquinone induced electron transport in lactic acid bacteria. Microb. Cell. Fact. 8:28-2859-8-28. doi: 10.1186/14752859-8-28;. 53. Rezaiki L, Lamberet G, Derre A, Gruss A, Gaudu P. 2008. Lactococcus lactis produces short-chain quinones that cross-feed Group B Streptococcus to activate respiration growth. Mol. Microbiol. 67:947-957. doi: 10.1111/j.1365-2958.2007.06083.x. 54. Morishita T, Tamura N, Makino T, Kudo S. 1999. Production of menaquinones by lactic acid bacteria. J. Dairy Sci. 82:1897-1903. doi: 10.3168/jds.S0022-0302(99)75424-X. 55. Gruss A, Borezee-Durant E, Lechardeur D. 2012. Environmental heme utilization by heme-auxotrophic bacteria. Adv. Microb. Physiol. 61:69-124. doi: 10.1016/B978-0-12-394423-8.00003-2. 7 153 56. Pritchard GG, Wimpenny JW. 1978. Cytochrome formation, oxygen-induced proton extrusion and respiratory activity in Streptococcus faecalis var. zymogenes grown in the presence of haematin. J. Gen. Microbiol. 104:15-22. 57. Yamamoto Y, Poyart C, Trieu-Cuot P, Lamberet G, Gruss A, Gaudu P. 2005. Respiration metabolism of Group B Streptococcus is activated by environmental haem and quinone and contributes to virulence. Mol. Microbiol. 56:525-534. doi: 10.1111/j.1365-2958.2005.04555.x. 58. Brooijmans RJ, Poolman B, Schuurman-Wolters GK, de Vos WM, Hugenholtz J. 2007. Generation of a membrane potential by Lactococcus lactis through aerobic electron transport. J. Bacteriol. 189:52035209. doi: 10.1128/JB.00361-07. 59. Duwat P, Sourice S, Cesselin B, Lamberet G, Vido K, Gaudu P, Le Loir Y, Violet F, Loubiere P, Gruss A. 2001. Respiration capacity of the fermenting bacterium Lactococcus lactis and its positive effects on growth and survival. J. Bacteriol. 183:4509-4516. doi: 10.1128/JB.183.15.4509-4516.2001. 60. Rezaiki L, Cesselin B, Yamamoto Y, Vido K, van West E, Gaudu P, Gruss A. 2004. Respiration metabolism reduces oxidative and acid stress to improve long-term survival of Lactococcus lactis. Mol. Microbiol. 53:1331-1342. doi: 10.1111/j.1365-2958.2004.04217.x. 61. Watanabe M, van der Veen S, Nakajima H, Abee T. 2012. Effect of respiration and manganese on oxidative stress resistance of Lactobacillus plantarum WCFS1. Microbiology. 158:293-300. doi: 10.1099/ mic.0.051250-0. 62. Huycke MM, Abrams V, Moore DR. 2002. Enterococcus faecalis produces extracellular superoxide and hydrogen peroxide that damages colonic epithelial cell DNA. Carcinogenesis. 23:529-536. 63. Huycke MM, Joyce W, Wack MF. 1996. Augmented production of extracellular superoxide by blood isolates of Enterococcus faecalis. J. Infect. Dis. 173:743-746. 64. Korshunov S, Imlay JA. 2010. Two sources of endogenous hydrogen peroxide in Escherichia coli. Mol. Microbiol. 75:1389-1401. doi: 10.1111/j.1365-2958.2010.07059.x. 65. Jansch A, Freiding S, Behr J, Vogel RF. 2011. Contribution of the NADH-oxidase (Nox) to the aerobic life of Lactobacillus sanfranciscensis DSM20451T. Food Microbiol. 28:29-37. doi: 10.1016/j.fm.2010.08.001. 66. Lopez de Felipe F, Kleerebezem M, de Vos WM, Hugenholtz J. 1998. Cofactor engineering: a novel approach to metabolic engineering in Lactococcus lactis by controlled expression of NADH oxidase. J. Bacteriol. 180:3804-3808. 67. Neves AR, Ramos A, Costa H, van S,II, Hugenholtz J, Kleerebezem M, de Vos W, Santos H. 2002. Effect of different NADH oxidase levels on glucose metabolism by Lactococcus lactis: kinetics of intracellular metabolite pools determined by in vivo nuclear magnetic resonance. Appl. Environ. Microbiol. 68:6332-6342. 68. Messner KR, Imlay JA. 2002. Mechanism of superoxide and hydrogen peroxide formation by fumarate reductase, succinate dehydrogenase, and aspartate oxidase. J. Biol. Chem. 277:42563-42571. doi: 10.1074/ jbc.M204958200. 69. Kawasaki S, Satoh T, Todoroki M, Niimura Y. 2009. b-type dihydroorotate dehydrogenase is purified as a H2O2-forming NADH oxidase from Bifidobacterium bifidum. Appl. Environ. Microbiol. 75:629-636. doi: 10.1128/AEM.02111-08. 70. Talwalkar A, Kailasapathy K, Hourigan J, Peiris P, Arumugaswamy R. 2003. An improved method for the determination of NADH oxidase in the presence of NADH peroxidase in lactic acid bacteria. J. Microbiol. Methods. 52:333-339. 71. Meitzler JL, Ortiz de Montellano PR. 2009. Caenorhabditis elegans and human dual oxidase 1 (DUOX1) “peroxidase” domains: insights into heme binding and catalytic activity. J. Biol. Chem. 284:18634-18643. doi: 10.1074/jbc.M109.013581. 72. Niimura Y, Nishiyama Y, Saito D, Tsuji H, Hidaka M, Miyaji T, Watanabe T, Massey V. 2000. A hydrogen peroxide-forming NADH oxidase that functions as an alkyl hydroperoxide reductase in Amphibacillus xylanus. J. Bacteriol. 182:5046-5051. 154 73. Storz G, Jacobson FS, Tartaglia LA, Morgan RW, Silveira LA, Ames BN. 1989. An alkyl hydroperoxide reductase induced by oxidative stress in Salmonella typhimurium and Escherichia coli: genetic characterization and cloning of ahp. J. Bacteriol. 171:2049-2055. 74. Poole LB, Higuchi M, Shimada M, Calzi ML, Kamio Y. 2000. Streptococcus mutans H2O2-forming NADH oxidase is an alkyl hydroperoxide reductase protein. Free Radic. Biol. Med. 28:108-120. 75. Higuchi M, Yamamoto Y, Kamio Y. 2000. Molecular biology of oxygen tolerance in lactic acid bacteria: Functions of NADH oxidases and Dpr in oxidative stress. J. Biosci. Bioeng. 90:484-493. 76. Higuchi M, Yamamoto Y, Poole LB, Shimada M, Sato Y, Takahashi N, Kamio Y. 1999. Functions of two types of NADH oxidases in energy metabolism and oxidative stress of Streptococcus mutans. J. Bacteriol. 181:5940-5947. 77. Lopez de Felipe F, Starrenburg M, Hugenholtz J. 1997. The role of NADH-oxidation in acetoin and diacetyl production from glucose in Lactococcus lactis subsp. lactis MG1363. FEMS Microbiol. Lett. 156:15-19. 78. Bongers RS, Hoefnagel MH, Kleerebezem M. 2005. High-level acetaldehyde production in Lactococcus lactis by metabolic engineering. Appl. Environ. Microbiol. 71:1109-1113. doi: 10.1128/AEM.71.2.11091113.2005. 79. Sedewitz B, Schleifer KH, Gotz F. 1984. Purification and biochemical characterization of pyruvate oxidase from Lactobacillus plantarum. J. Bacteriol. 160:273-278. 80. Hager LP, Lipmann F. 1961. Coupling between phosphorylation and flavin adenine dinucleotide reduction with the pyruvate oxidase of Lactobacillus delbrueckii enzyme. Proc. Natl. Acad. Sci. U. S. A. 47:1768-1772. 81. Goffin P, Lorquet F, Kleerebezem M, Hols P. 2004. Major role of NAD-dependent lactate dehydrogenases in aerobic lactate utilization in Lactobacillus plantarum during early stationary phase. J. Bacteriol. 186:66616666. doi: 10.1128/JB.186.19.6661-6666.2004. 82. Zhao R, Zheng S, Duan C, Liu F, Yang L, Huo G. 2013. NAD-dependent lactate dehydrogenase catalyses the first step in respiratory utilization of lactate by Lactococcus lactis. FEBS Open Bio. 3:379-386. doi: 10.1016/j.fob.2013.08.005. 83. Goffin P, Muscariello L, Lorquet F, Stukkens A, Prozzi D, Sacco M, Kleerebezem M, Hols P. 2006. Involvement of pyruvate oxidase activity and acetate production in the survival of Lactobacillus plantarum during the stationary phase of aerobic growth. Appl. Environ. Microbiol. 72:7933-7940. doi: 10.1128/ AEM.00659-06. 84. Lorquet F, Goffin P, Muscariello L, Baudry JB, Ladero V, Sacco M, Kleerebezem M, Hols P. 2004. Characterization and functional analysis of the poxB gene, which encodes pyruvate oxidase in Lactobacillus plantarum. J. Bacteriol. 186:3749-3759. doi: 10.1128/JB.186.12.3749-3759.2004. 85. Zheng L, Itzek A, Chen Z, Kreth J. 2011. Environmental influences on competitive hydrogen peroxide production in Streptococcus gordonii. Appl. Environ. Microbiol. 77:4318-4328. doi: 10.1128/AEM.00309-11. 86. Seki M, Iida K, Saito M, Nakayama H, Yoshida S. 2004. Hydrogen peroxide production in Streptococcus pyogenes: involvement of lactate oxidase and coupling with aerobic utilization of lactate. J. Bacteriol. 186:2046-2051. 87. Kietzman CC, Caparon MG. 2010. CcpA and LacD.1 affect temporal regulation of Streptococcus pyogenes virulence genes. Infect. Immun. 78:241-252. doi: 10.1128/IAI.00746-09;. 88. Carvalho SM, Kloosterman TG, Kuipers OP, Neves AR. 2011. CcpA ensures optimal metabolic fitness of Streptococcus pneumoniae. PLoS One. 6:e26707. doi: 10.1371/journal.pone.0026707; 10.1371/journal. pone.0026707. 89. Taniai H, Iida K, Seki M, Saito M, Shiota S, Nakayama H, Yoshida S. 2008. Concerted action of lactate oxidase and pyruvate oxidase in aerobic growth of Streptococcus pneumoniae: role of lactate as an energy source. J. Bacteriol. 190:3572-3579. doi: 10.1128/JB.01882-07. 7 155 90. Regev-Yochay G, Trzcinski K, Thompson CM, Lipsitch M, Malley R. 2007. SpxB is a suicide gene of Streptococcus pneumoniae and confers a selective advantage in an in vivo competitive colonization model. J. Bacteriol. 189:6532-6539. doi: 10.1128/JB.00813-07. 91. Pericone CD, Park S, Imlay JA, Weiser JN. 2003. Factors contributing to hydrogen peroxide resistance in Streptococcus pneumoniae include pyruvate oxidase (SpxB) and avoidance of the toxic effects of the fenton reaction. J. Bacteriol. 185:6815-6825. 92. Chen L, Ge X, Dou Y, Wang X, Patel JR, Xu P. 2011. Identification of hydrogen peroxide productionrelated genes in Streptococcus sanguinis and their functional relationship with pyruvate oxidase. Microbiology. 157:13-20. doi: 10.1099/mic.0.039669-0. 93. Yi X, Kot E, Bezkorovainy A. 1998. Properties of NADH oxidase from Lactobacillus delbrueckii ssp bulgaricus. J Sci Food Agric. 78:527-527-534. 94. Martin R, Suarez JE. 2010. Biosynthesis and degradation of H2O2 by vaginal lactobacilli. Appl. Environ. Microbiol. 76:400-405. doi: 10.1128/AEM.01631-09. 95. Seaver LC, Imlay JA. 2004. Are respiratory enzymes the primary sources of intracellular hydrogen peroxide? J. Biol. Chem. 279:48742-48750. doi: 10.1074/jbc.M408754200. 96. Frankenberg L, Brugna M, Hederstedt L. 2002. Enterococcus faecalis heme-dependent catalase. J. Bacteriol. 184:6351-6356. 97. Abriouel H, Herrmann A, Starke J, Yousif NM, Wijaya A, Tauscher B, Holzapfel W, Franz CM. 2004. Cloning and heterologous expression of hematin-dependent catalase produced by Lactobacillus plantarum CNRZ 1228. Appl. Environ. Microbiol. 70:603-606. 98. An H, Zhou H, Huang Y, Wang G, Luan C, Mou J, Luo Y, Hao Y. 2010. High-level expression of hemedependent catalase gene katA from Lactobacillus sakei protects Lactobacillus rhamnosus from oxidative stress. Mol. Biotechnol. 45:155-160. doi: 10.1007/s12033-010-9254-9. 99. Wolf G, Hammes WP. 1988. Effect of hematin on the activities of nitrite reductase and catalase in lactobacilli. Arch Microb. 149:220--224. 100. Kono Y, Fridovich I. 1983. Functional significance of manganese catalase in Lactobacillus plantarum. J. Bacteriol. 155:742-746. 101. Johnston MA, Delwiche EA. 1962. Catalase of the Lactobacillaceae. J. Bacteriol. 83:936--938. 102. Johnston MA, Delwiche EA. 1965. Isolation and characterization of the cyanide-resistant and azideresistant catalase of Lactobacillus plantarum. J. Bacteriol. 90:352-356. 103. Kono Y, Fridovich I. 1983. Isolation and characterization of the pseudocatalase of Lactobacillus plantarum. J. Biol. Chem. 258:6015-6019. 104. Flohe L, Toppo S, Cozza G, Ursini F. 2011. A comparison of thiol peroxidase mechanisms. Antioxid. Redox Signal. 15:763-780. doi: 10.1089/ars.2010.3397. 105. Poole LB. 2005. Bacterial defenses against oxidants: mechanistic features of cysteine-based peroxidases and their flavoprotein reductases. Arch. Biochem. Biophys. 433:240-254. doi: 10.1016/j.abb.2004.09.006. 106. Lu J, Holmgren A. 2014. The thioredoxin antioxidant system. Free Radic. Biol. Med. 66:75-87. doi: 10.1016/j.freeradbiomed.2013.07.036. 107. Zhang J, Du GC, Zhang Y, Liao XY, Wang M, Li Y, Chen J. 2010. Glutathione protects Lactobacillus sanfranciscensis against freeze-thawing, freeze-drying, and cold treatment. Appl. Environ. Microbiol. 76:2989-2996. doi: 10.1128/AEM.00026-09. 108. Pophaly SD, Singh R, Pophaly SD, Kaushik JK, Tomar SK. 2012. Current status and emerging role of glutathione in food grade lactic acid bacteria. Microb. Cell. Fact. 11:114-2859-11-114. doi: 10.1186/14752859-11-114. 109. Seaver LC, Imlay JA. 2001. Alkyl hydroperoxide reductase is the primary scavenger of endogenous hydrogen peroxide in Escherichia coli. J. Bacteriol. 183:7173-7181. doi: 10.1128/JB.183.24.7173-7181.2001. 156 110. Arner ES, Holmgren A. 2000. Physiological functions of thioredoxin and thioredoxin reductase. Eur. J. Biochem. 267:6102-6109. 111. Holmgren A. 1985. Thioredoxin. Annu. Rev. Biochem. 54:237-271. doi: 10.1146/annurev. bi.54.070185.001321. 112. Zeller T, Klug G. 2006. Thioredoxins in bacteria: functions in oxidative stress response and regulation of thioredoxin genes. Naturwissenschaften. 93:259-266. doi: 10.1007/s00114-006-0106-1. 113. Serrano LM, Molenaar D, Wels M, Teusink B, Bron PA, de Vos WM, Smid EJ. 2007. Thioredoxin reductase is a key factor in the oxidative stress response of Lactobacillus plantarum WCFS1. Microb. Cell. Fact. 6:29. doi: 10.1186/1475-2859-6-29. 114. Serata M, Iino T, Yasuda E, Sako T. 2012. Roles of thioredoxin and thioredoxin reductase in the resistance to oxidative stress in Lactobacillus casei. Microbiology. 158:953-962. doi: 10.1099/mic.0.053942-0. 115. Vido K, Diemer H, Van Dorsselaer A, Leize E, Juillard V, Gruss A, Gaudu P. 2005. Roles of thioredoxin reductase during the aerobic life of Lactococcus lactis. J. Bacteriol. 187:601-610. doi: 10.1128/ JB.187.2.601-610.2005. 116. Hajaj B, Yesilkaya H, Benisty R, David M, Andrew PW, Porat N. 2012. Thiol peroxidase is an important component of Streptococcus pneumoniae in oxygenated environments. Infect. Immun. 80:43334343. doi: 10.1128/IAI.00126-12. 117. La Carbona S, Sauvageot N, Giard JC, Benachour A, Posteraro B, Auffray Y, Sanguinetti M, Hartke A. 2007. Comparative study of the physiological roles of three peroxidases (NADH peroxidase, Alkyl hydroperoxide reductase and Thiol peroxidase) in oxidative stress response, survival inside macrophages and virulence of Enterococcus faecalis. Mol. Microbiol. 66:1148-1163. doi: 10.1111/j.1365-2958.2007.05987.x. 118. Potter AJ, Trappetti C, Paton JC. 2012. Streptococcus pneumoniae uses glutathione to defend against oxidative stress and metal ion toxicity. J. Bacteriol. 194:6248-6254. doi: 10.1128/JB.01393-12. 119. Li Y, Hugenholtz J, Abee T, Molenaar D. 2003. Glutathione protects Lactococcus lactis against oxidative stress. Appl. Environ. Microbiol. 69:5739-5745. 120. Zhang YW, Tiwari MK, Gao H, Dhiman SS, Jeya M, Lee JK. 2012. Cloning and characterization of a thermostable H2O-forming NADH oxidase from Lactobacillus rhamnosus. Enzyme Microb. Technol. 50:255262. doi: 10.1016/j.enzmictec.2012.01.009. 121. Sherrill C, Fahey RC. 1998. Import and metabolism of glutathione by Streptococcus mutans. J. Bacteriol. 180:1454-1459. 122. Jansch A, Korakli M, Vogel RF, Ganzle MG. 2007. Glutathione reductase from Lactobacillus sanfranciscensis DSM20451T: contribution to oxygen tolerance and thiol exchange reactions in wheat sourdoughs. Appl. Environ. Microbiol. 73:4469-4476. doi: 10.1128/AEM.02322-06. 123. Brenot A, King KY, Janowiak B, Griffith O, Caparon MG. 2004. Contribution of glutathione peroxidase to the virulence of Streptococcus pyogenes. Infect. Immun. 72:408-413. 124. Higuchi M, Shimada M, Yamamoto Y, Hayashi T, Koga T, Kamio Y. 1993. Identification of two distinct NADH oxidases corresponding to H2O2-forming oxidase and H2O-forming oxidase induced in Streptococcus mutans. J. Gen. Microbiol. 139:2343-2351. 125. Fujishima K, Kawada-Matsuo M, Oogai Y, Tokuda M, Torii M, Komatsuzawa H. 2013. dpr and sod in Streptococcus mutans are involved in coexistence with S. sanguinis, and PerR is associated with resistance to H2O2. Appl. Environ. Microbiol. 79:1436-1443. doi: 10.1128/AEM.03306-12; 10.1128/AEM.03306-12. 126. Claiborne A, Ross RP, Parsonage D. 1992. Flavin-linked peroxide reductases: protein-sulfenic acids and the oxidative stress response. Trends Biochem. Sci. 17:183-186. 127. Stehle T, Ahmed SA, Claiborne A, Schulz GE. 1991. Structure of NADH peroxidase from Streptococcus faecalis 10C1 refined at 2.16 A resolution. J. Mol. Biol. 221:1325-1344. 7 157 128. Park JT, Hirano J, Thangavel, V., Riebel, B.R., Bommarius, A. S. 2011. NAD(P)H oxidase V from Lactobacillus plantarum (NoxV) displays enhanced operational stability even in absence of reducing agents. Journal of Mol. Cat. B: Enz. 71:159--165. 129. Massey V. 2002. The reactivity of oxygen with flavoproteins. International Congress Series. 1233:3--11. 130. Massey V. 1994. Activation of molecular oxygen by flavins and flavoproteins. J. Biol. Chem. 269:2245922462. 131. Keyer K, Imlay JA. 1996. Superoxide accelerates DNA damage by elevating free-iron levels. Proc. Natl. Acad. Sci. U. S. A. 93:13635-13640. 132. Sanders JW, Leenhouts KJ, Haandrikman AJ, Venema G, Kok J. 1995. Stress response in Lactococcus lactis: cloning, expression analysis, and mutation of the lactococcal superoxide dismutase gene. J. Bacteriol. 177:5254-5260. 133. De Angelis M, Gobbetti M. 1999. Lactobacillus sanfranciscensis CB1: manganese, oxygen, superoxide dismutase and metabolism. Appl. Microbiol. Biotechnol. 51:358-363. 134. Amanatidou A, Bennik MH, Gorris LG, Smid EJ. 2001. Superoxide dismutase plays an important role in the survival of Lactobacillus sake upon exposure to elevated oxygen. Arch. Microbiol. 176:79-88. 135. Yesilkaya H, Kadioglu A, Gingles N, Alexander JE, Mitchell TJ, Andrew PW. 2000. Role of manganese-containing superoxide dismutase in oxidative stress and virulence of Streptococcus pneumoniae. Infect. Immun. 68:2819-2826. 136. Nakayama K. 1992. Nucleotide sequence of Streptococcus mutans superoxide dismutase gene and isolation of insertion mutants. J. Bacteriol. 174:4928-4934. 137. Gibson CM, Caparon MG. 1996. Insertional inactivation of Streptococcus pyogenes sod suggests that prtF is regulated in response to a superoxide signal. J. Bacteriol. 178:4688-4695. 138. Bruno-Barcena JM, Andrus JM, Libby SL, Klaenhammer TR, Hassan HM. 2004. Expression of a heterologous manganese superoxide dismutase gene in intestinal lactobacilli provides protection against hydrogen peroxide toxicity. Appl. Environ. Microbiol. 70:4702-4710. doi: 10.1128/AEM.70.8.47024710.2004. 139. Gregory EM, Fridovich I. 1974. Oxygen metabolism in Lactobacillus plantarum. J. Bacteriol. 117:166169. 140. Quatravaux S, Remize F, Bryckaert E, Colavizza D, Guzzo J. 2006. Examination of Lactobacillus plantarum lactate metabolism side effects in relation to the modulation of aeration parameters. J. Appl. Microbiol. 101:903-912. doi: 10.1111/j.1365-2672.2006.02955.x. 141. Archibald FS, Fridovich I. 1981. Manganese and defenses against oxygen toxicity in Lactobacillus plantarum. J. Bacteriol. 145:442-451. 142. Jakubovics NS, Jenkinson HF. 2001. Out of the iron age: new insights into the critical role of manganese homeostasis in bacteria. Microbiology. 147:1709-1718. 143. Nierop Groot MN, de Bont JA. 1999. Involvement of manganese in conversion of phenylalanine to benzaldehyde by lactic acid bacteria. Appl. Environ. Microbiol. 65:5590-5593. 144. Stadtman ER, Berlett BS, Chock PB. 1990. Manganese-dependent disproportionation of hydrogen peroxide in bicarbonate buffer. Proc. Natl. Acad. Sci. U. S. A. 87:384-388. 145. Anjem A, Varghese S, Imlay JA. 2009. Manganese import is a key element of the OxyR response to hydrogen peroxide in Escherichia coli. Mol. Microbiol. 72:844-858. doi: 10.1111/j.1365-2958.2009.06699.x. 146. Anjem A, Imlay JA. 2012. Mononuclear iron enzymes are primary targets of hydrogen peroxide stress. J. Biol. Chem. 287:15544-15556. doi: 10.1074/jbc.M111.330365. 147. Wang X, Tong H, Dong X. 2014. PerR-regulated manganese ion uptake contributes to oxidative stress defense in an oral Streptococcus. Appl. Environ. Microbiol. . doi: 10.1128/AEM.00064-14. 148. Hao Z, Reiske HR, Wilson DB. 1999. Characterization of cadmium uptake in Lactobacillus plantarum and isolation of cadmium and manganese uptake mutants. Appl. Environ. Microbiol. 65:4741-4745. 158 149. Hao Z, Chen S, Wilson DB. 1999. Cloning, expression, and characterization of cadmium and manganese uptake genes from Lactobacillus plantarum. Appl. Environ. Microbiol. 65:4746-4752. 150. Groot MN, Klaassens E, de Vos WM, Delcour J, Hols P, Kleerebezem M. 2005. Genome-based in silico detection of putative manganese transport systems in Lactobacillus plantarum and their genetic analysis. Microbiology. 151:1229-1238. doi: 10.1099/mic.0.27375-0. 151. Duhutrel P, Bordat C, Wu TD, Zagorec M, Guerquin-Kern JL, Champomier-Verges MC. 2010. Iron sources used by the nonpathogenic lactic acid bacterium Lactobacillus sakei as revealed by electron energy loss spectroscopy and secondary-ion mass spectrometry. Appl. Environ. Microbiol. 76:560-565. doi: 10.1128/ AEM.02205-09. 152. Archibald FS. 1983. Lactobacillus plantarum, an organism not requiring iron. FEMS Microbiol. Lett. 19:29--32. 153. Imbert M, Blondeau R. 1998. On the iron requirement of lactobacilli grown in chemically defined medium. Curr. Microbiol. 37:64-66. 154. Elli M, Zink R, Rytz A, Reniero R, Morelli L. 2000. Iron requirement of Lactobacillus spp. in completely chemically defined growth media. J. Appl. Microbiol. 88:695-703. 155. Andrews SC, Robinson AK, Rodriguez-Quinones F. 2003. Bacterial iron homeostasis. FEMS Microbiol. Rev. 27:215-237. 156. Ishikawa T, Mizunoe Y, Kawabata S, Takade A, Harada M, Wai SN, Yoshida S. 2003. The iron-binding protein Dps confers hydrogen peroxide stress resistance to Campylobacter jejuni. J. Bacteriol. 185:1010-1017. 157. Tsou CC, Chiang-Ni C, Lin YS, Chuang WJ, Lin MT, Liu CC, Wu JJ. 2008. An iron-binding protein, Dpr, decreases hydrogen peroxide stress and protects Streptococcus pyogenes against multiple stresses. Infect. Immun. 76:4038-4045. doi: 10.1128/IAI.00477-08. 158. Yamamoto Y, Higuchi M, Poole LB, Kamio Y. 2000. Role of the dpr product in oxygen tolerance in Streptococcus mutans. J. Bacteriol. 182:3740-3747. 159. Troxell B, Zhang JJ, Bourret TJ, Zeng MY, Blum J, Gherardini F, Hassan HM, Yang XF. 2014. Pyruvate protects pathogenic spirochetes from H2O2 killing. PLoS One. 9:e84625. doi: 10.1371/journal. pone.0084625. 160. O’Donnell-Tormey J, Nathan CF, Lanks K, DeBoer CJ, de la Harpe J. 1987. Secretion of pyruvate. An antioxidant defense of mammalian cells. J. Exp. Med. 165:500-514. 161. Ramakrishnan N, Chen R, McClain DE, Bunger R. 1998. Pyruvate prevents hydrogen peroxideinduced apoptosis. Free Radic. Res. 29:283-295. 162. Smart JB, Thomas TD. 1987. Effect of oxygen on lactose metabolism in lactic streptococci. Appl. Environ. Microbiol. 53:533-541. 163. Bignucolo A, Appanna VP, Thomas SC, Auger C, Han S, Omri A, Appanna VD. 2013. Hydrogen peroxide stress provokes a metabolic reprogramming in Pseudomonas fluorescens: enhanced production of pyruvate. J. Biotechnol. 167:309-315. doi: 10.1016/j.jbiotec.2013.07.002. 164. Hung J, Cooper D, Turner MS, Walsh T, Giffard PM. 2003. Cystine uptake prevents production of hydrogen peroxide by Lactobacillus fermentum BR11. FEMS Microbiol. Lett. 227:93-99. 165. Lo R, Turner MS, Barry DG, Sreekumar R, Walsh TP, Giffard PM. 2009. Cystathionine gamma-lyase is a component of cystine-mediated oxidative defense in Lactobacillus reuteri BR11. J. Bacteriol. 191:18271837. doi: 10.1128/JB.01553-08. 166. Sreekumar R, Al-Attabi Z, Deeth HC, Turner MS. 2009. Volatile sulfur compounds produced by probiotic bacteria in the presence of cysteine or methionine. Lett. Appl. Microbiol. 48:777-782. doi: 10.1111/j.1472-765X.2009.02610.x. 7 159 167. Hung J, Turner MS, Walsh T, Giffard PM. 2005. BspA (CyuC) in Lactobacillus fermentum BR11 is a highly expressed high-affinity L-cystine-binding protein. Curr. Microbiol. 50:33-37. doi: 10.1007/s00284-0044408-2. 168. Berger B, Pridmore RD, Barretto C, Delmas-Julien F, Schreiber K, Arigoni F, Brussow H. 2007. Similarity and differences in the Lactobacillus acidophilus group identified by polyphasic analysis and comparative genomics. J. Bacteriol. 189:1311-1321. doi: 10.1128/JB.01393-06. 169. Ibnou-Zekri N, Blum S, Schiffrin EJ, von der Weid T. 2003. Divergent patterns of colonization and immune response elicited from two intestinal Lactobacillus strains that display similar properties in vitro. Infect. Immun. 71:428-436. 170. Granato D, Perotti F, Masserey I, Rouvet M, Golliard M, Servin A, Brassart D. 1999. Cell surfaceassociated lipoteichoic acid acts as an adhesion factor for attachment of Lactobacillus johnsonii La1 to human enterocyte-like Caco-2 cells. Appl. Environ. Microbiol. 65:1071-1077. 171. Bernet MF, Brassart D, Neeser JR, Servin AL. 1994. Lactobacillus acidophilus LA 1 binds to cultured human intestinal cell lines and inhibits cell attachment and cell invasion by enterovirulent bacteria. Gut. 35:483-489. 172. Fujisawa T, Benno Y, Yaeshima T, Mitsuoka T. 1992. Taxonomic study of the Lactobacillus acidophilus group, with recognition of Lactobacillus gallinarum sp. nov. and Lactobacillus johnsonii sp. nov. and synonymy of Lactobacillus acidophilus group A3 (Johnson et al. 1980) with the type strain of Lactobacillus amylovorus (Nakamura 1981). Int. J. Syst. Bacteriol. 42:487-491. 173. Sarmiento-Rubiano LA, Berger B, Moine D, Zuniga M, Perez-Martinez G, Yebra MJ. 2010. Characterization of a novel Lactobacillus species closely related to Lactobacillus johnsonii using a combination of molecular and comparative genomics methods. BMC Genomics. 11:504-2164-11-504. doi: 10.1186/14712164-11-504. 174. Kullen MJ, Sanozky-Dawes RB, Crowell DC, Klaenhammer TR. 2000. Use of the DNA sequence of variable regions of the 16S rRNA gene for rapid and accurate identification of bacteria in the Lactobacillus acidophilus complex. J. Appl. Microbiol. 89:511-516. 175. Du Plessis EM, Dicks LM. 1995. Evaluation of random amplified polymorphic DNA (RAPD)-PCR as a method to differentiate Lactobacillus acidophilus, Lactobacillus crispatus, Lactobacillus amylovorus, Lactobacillus gallinarum, Lactobacillus gasseri, and Lactobacillus johnsonii. Curr. Microbiol. 31:114-118. 176. Pridmore RD, Berger B, Desiere F, Vilanova D, Barretto C, Pittet AC, Zwahlen MC, Rouvet M, Altermann E, Barrangou R, Mollet B, Mercenier A, Klaenhammer T, Arigoni F, Schell MA. 2004. The genome sequence of the probiotic intestinal bacterium Lactobacillus johnsonii NCC 533. Proc. Natl. Acad. Sci. U. S. A. 101:2512-2517. 177. Kleerebezem M, Boekhorst J, van Kranenburg R, Molenaar D, Kuipers OP, Leer R, Tarchini R, Peters SA, Sandbrink HM, Fiers MW, Stiekema W, Lankhorst RM, Bron PA, Hoffer SM, Groot MN, Kerkhoven R, de Vries M, Ursing B, de Vos WM, Siezen RJ. 2003. Complete genome sequence of Lactobacillus plantarum WCFS1. Proc. Natl. Acad. Sci. U. S. A. 100:1990-1995. doi: 10.1073/pnas.0337704100. 178. Boekhorst J, Siezen RJ, Zwahlen MC, Vilanova D, Pridmore RD, Mercenier A, Kleerebezem M, de Vos WM, Brussow H, Desiere F. 2004. The complete genomes of Lactobacillus plantarum and Lactobacillus johnsonii reveal extensive differences in chromosome organization and gene content. Microbiology. 150:3601-3611. doi: 10.1099/mic.0.27392-0. 179. Bernet-Camard MF, Lievin V, Brassart D, Neeser JR, Servin AL, Hudault S. 1997. The human Lactobacillus acidophilus strain LA1 secretes a nonbacteriocin antibacterial substance(s) active in vitro and in vivo. Appl. Environ. Microbiol. 63:2747-2753. 180. Michetti P, Dorta G, Wiesel PH, Brassart D, Verdu E, Herranz M, Felley C, Porta N, Rouvet M, Blum AL, Corthesy-Theulaz I. 1999. Effect of whey-based culture supernatant of Lactobacillus acidophilus (johnsonii) La1 on Helicobacter pylori infection in humans. Digestion. 60:203-209. doi: 7660. 160 181. Kaburagi T, Yamano T, Fukushima Y, Yoshino H, Mito N, Sato K. 2007. Effect of Lactobacillus johnsonii La1 on immune function and serum albumin in aged and malnourished aged mice. Nutrition. 23:342-350. doi: 10.1016/j.nut.2007.02.001. 182. Haller D, Serrant P, Granato D, Schiffrin EJ, Blum S. 2002. Activation of human NK cells by staphylococci and lactobacilli requires cell contact-dependent costimulation by autologous monocytes. Clin. Diagn. Lab. Immunol. 9:649-657. 183. Haller D, Bode C, Hammes WP, Pfeifer AM, Schiffrin EJ, Blum S. 2000. Non-pathogenic bacteria elicit a differential cytokine response by intestinal epithelial cell/leucocyte co-cultures. Gut. 47:79-87. 184. Muller JA, Ross RP, Sybesma WF, Fitzgerald GF, Stanton C. 2011. Modification of the technical properties of Lactobacillus johnsonii NCC 533 by supplementing the growth medium with unsaturated fatty acids. Appl. Environ. Microbiol. 77:6889-6898. doi: 10.1128/AEM.05213-11. 185. van der Kaaij H, Desiere F, Mollet B, Germond JE. 2004. L-alanine auxotrophy of Lactobacillus johnsonii as demonstrated by physiological, genomic, and gene complementation approaches. Appl. Environ. Microbiol. 70:1869-1873. 186. Anwar MA, Kralj S, van der Maarel MJ, Dijkhuizen L. 2008. The probiotic Lactobacillus johnsonii NCC 533 produces high-molecular-mass inulin from sucrose by using an inulosucrase enzyme. Appl. Environ. Microbiol. 74:3426-3433. doi: 10.1128/AEM.00377-08. 187. Eschenbach DA, Davick PR, Williams BL, Klebanoff SJ, Young-Smith K, Critchlow CM, Holmes KK. 1989. Prevalence of hydrogen peroxide-producing Lactobacillus species in normal women and women with bacterial vaginosis. J. Clin. Microbiol. 27:251-256. 188. Rochat T, Gratadoux JJ, Gruss A, Corthier G, Maguin E, Langella P, van de Guchte M. 2006. Production of a heterologous nonheme catalase by Lactobacillus casei: an efficient tool for removal of H2O2 and protection of Lactobacillus bulgaricus from oxidative stress in milk. Appl. Environ. Microbiol. 72:51435149. doi: 10.1128/AEM.00482-06. 189. Thomas EL, Pera KA. 1983. Oxygen metabolism of Streptococcus mutans: uptake of oxygen and release of superoxide and hydrogen peroxide. J. Bacteriol. 154:1236-1244. 190. Pericone CD, Overweg K, Hermans PW, Weiser JN. 2000. Inhibitory and bactericidal effects of hydrogen peroxide production by Streptococcus pneumoniae on other inhabitants of the upper respiratory tract. Infect. Immun. 68:3990-3997. 191. Korshunov S, Imlay JA. 2010. Two sources of endogenous hydrogen peroxide in Escherichia coli. Mol. Microbiol. 75:1389-1401. doi: 10.1111/j.1365-2958.2010.07059.x. 192. Seki M, Iida K-, Saito M, Nakayama H, Yoshida S-. 2004. Hydrogen peroxide production in Streptococcus pyogenes: involvement of lactate oxidase and coupling with aerobic utilization of lactate. J. Bacteriol. 186:2046--2051. doi: 10.1128/JB.186.7.2046-2051.2004. 193. Kietzman CC, Caparon MG. 2010. CcpA and LacD1 affect temporal regulation of Streptococcus pyogenes virulence genes. Infect. Immun. 78:241-252. doi: 10.1128/IAI.00746-09. 194. Hecht HJ, Erdmann H, Park HJ, Sprinzl M, Schmid RD. 1995. Crystal structure of NADH oxidase from Thermus thermophilus. Nat. Struct. Biol. 2:1109-1114. 195. Neeser JR, Granato D, Rouvet M, Servin A, Teneberg S, Karlsson KA. 2000. Lactobacillus johnsonii La1 shares carbohydrate-binding specificities with several enteropathogenic bacteria. Glycobiology. 10:11931199. 196. He G, Shankar RA, Chzhan M, Samouilov A, Kuppusamy P, Zweier JL. 1999. Noninvasive measurement of anatomic structure and intraluminal oxygenation in the gastrointestinal tract of living mice with spatial and spectral EPR imaging. Proc. Natl. Acad. Sci. U. S. A. 96:4586-4591. 197. Ding SZ, O’Hara AM, Denning TL, Dirden-Kramer B, Mifflin RC, Reyes VE, Ryan KA, Elliott SN, Izumi T, Boldogh I, Mitra S, Ernst PB, Crowe SE. 2004. Helicobacter pylori and H2O2 increase AP endonuclease-1/redox factor-1 expression in human gastric epithelial cells. Gastroenterology. 127:845-858. 7 161 198. Hawes SE, Hillier SL, Benedetti J, Stevens CE, Koutsky LA, Wolner-Hanssen P, Holmes KK. 1996. Hydrogen peroxide-producing lactobacilli and acquisition of vaginal infections. J. Infect. Dis. 174:1058-1063. 199. Donati L, Di Vico A, Nucci M, Quagliozzi L, Spagnuolo T, Labianca A, Bracaglia M, Ianniello F, Caruso A, Paradisi G. 2010. Vaginal microbial flora and outcome of pregnancy. Arch. Gynecol. Obstet. 281:589-600. doi: 10.1007/s00404-009-1318-3. 200. Schwebke JR. 2005. Abnormal vaginal flora as a biological risk factor for acquisition of HIV infection and sexually transmitted diseases. J. Infect. Dis. 192:1315-1317. doi: 10.1086/462430. 201. Desreumaux P, Dubuquoy L, Nutten S, Peuchmaur M, Englaro W, Schoonjans K, Derijard B, Desvergne B, Wahli W, Chambon P, Leibowitz MD, Colombel JF, Auwerx J. 2001. Attenuation of colon inflammation through activators of the retinoid X receptor (RXR)/peroxisome proliferator-activated receptor gamma (PPARgamma) heterodimer. A basis for new therapeutic strategies. J. Exp. Med. 193:827-838. 202. Su CG, Wen X, Bailey ST, Jiang W, Rangwala SM, Keilbaugh SA, Flanigan A, Murthy S, Lazar MA, Wu GD. 1999. A novel therapy for colitis utilizing PPAR-gamma ligands to inhibit the epithelial inflammatory response. J. Clin. Invest. 104:383-389. doi: 10.1172/JCI7145. 203. Valladares R, Bojilova L, Potts AH, Cameron E, Gardner C, Lorca G, Gonzalez CF. 2013. Lactobacillus johnsonii inhibits indoleamine 2,3-dioxygenase and alters tryptophan metabolite levels in BioBreeding rats. FASEB J. 27:1711-1720. doi: 10.1096/fj.12-223339;. 204. Felis GE, Dellaglio F. 2007. Taxonomy of Lactobacilli and Bifidobacteria. Curr. Issues Intest Microbiol. 8:44-61. 205. de Man JC, Rogosa M, Sharpe MEA. 1960. A medium for the cultivation of lactobacilli. Journal of Applied Bacteriology. 130-130-135. 206. Denou E, Pridmore RD, Berger B, Panoff JM, Arigoni F, Brussow H. 2008. Identification of genes associated with the long-gut-persistence phenotype of the probiotic Lactobacillus johnsonii strain NCC533 using a combination of genomics and transcriptome analysis. J. Bacteriol. 190:3161-3168. doi: 10.1128/ JB.01637-07. 207. Platteeuw C, Simons G, de Vos WM. 1994. Use of the Escherichia coli beta-glucuronidase (gusA) gene as a reporter gene for analyzing promoters in lactic acid bacteria. Appl. Environ. Microbiol. 60:587-593. 208. Bekker M, de Vries S, Ter Beek A, Hellingwerf KJ, de Mattos MJ. 2009. Respiration of Escherichia coli can be fully uncoupled via the nonelectrogenic terminal cytochrome bd-II oxidase. J. Bacteriol. 191:55105517. doi: 10.1128/JB.00562-09. 209. Zdobnov EM, Apweiler R. 2001. InterProScan--an integration platform for the signature-recognition methods in InterPro. Bioinformatics. 17:847-848. 210. Mi H, Muruganujan A, Thomas PD. 2013. PANTHER in 2013: modeling the evolution of gene function, and other gene attributes, in the context of phylogenetic trees. Nucleic Acids Res. 41:D377-86. doi: 10.1093/ nar/gks1118;. 211. Ackerley DF, Gonzalez CF, Park CH, Blake R, Keyhan M, Matin A. 2004. Chromate-reducing properties of soluble flavoproteins from Pseudomonas putida and Escherichia coli. Appl. Environ. Microbiol. 70:873--882. doi: 10.1128/AEM.70.2.873-882.2004. 212. Gonzalez CF, Ackerley DF, Lynch SV, Matin A. 2005. ChrR, a soluble quinone reductase of Pseudomonas putida that defends against H2O2. J. Biol. Chem. 280:22590-22595. doi: 10.1074/jbc.M501654200. 213. Eswaramoorthy S, Poulain S, Hienerwadel R, Bremond N, Sylvester MD, Zhang YB, Berthomieu C, Van Der Lelie D, Matin A. 2012. Crystal structure of ChrR--a quinone reductase with the capacity to reduce chromate. PLoS One. 7:e36017. doi: 10.1371/journal.pone.0036017. 214. Jensen LJ, Kuhn M, Stark M, Chaffron S, Creevey C, Muller J, Doerks T, Julien P, Roth A, Simonovic M, Bork P, von Mering C. 2009. STRING 8--a global view on proteins and their functional interactions in 630 organisms. Nucleic Acids Res. 37:D412-6. doi: 10.1093/nar/gkn760. 162 215. Copeland A, Sikorski J, Lapidus A, Nolan M, Del Rio TG, Lucas S, Chen F, Tice H, Pitluck S, Cheng JF, Pukall R, Chertkov O, Brettin T, Han C, Detter JC, Kuske C, Bruce D, Goodwin L, Ivanova N, Mavromatis K, Mikhailova N, Chen A, Palaniappan K, Chain P, Rohde M, Goker M, Bristow J, Eisen JA, Markowitz V, Hugenholtz P, Kyrpides NC, Klenk HP, Detter JC. 2009. Complete genome sequence of Atopobium parvulum type strain (IPP 1246). Stand. Genomic Sci. 1:166-173. doi: 10.4056/sigs.29547. 216. Pot B, Hertel C, Ludwig W, Descheemaeker P, Kersters K, Schleifer KH. 1993. Identification and classification of Lactobacillus acidophilus, L. gasseri and L. johnsonii strains by SDS-PAGE and rRNA-targeted oligonucleotide probe hybridization. J. Gen. Microbiol. 139:513-517. 217. von Canstein H, Ogawa J, Shimizu S, Lloyd JR. 2008. Secretion of flavins by Shewanella species and their role in extracellular electron transfer. Appl. Environ. Microbiol. 74:615-623. doi: 10.1128/AEM.0138707. 218. Ellis RJ. 2001. Macromolecular crowding: an important but neglected aspect of the intracellular environment. Curr. Opin. Struct. Biol. 11:114-119. 219. Ohnishi K, Niimura Y, Yokoyama K, Hidaka M, Masaki H, Uchimura T, Suzuki H, Uozumi T, Kozaki M, Komagata K. 1994. Purification and analysis of a flavoprotein functional as NADH oxidase from Amphibacillus xylanus overexpressed in Escherichia coli. J. Biol. Chem. 269:31418-31423. 220. Pan N, Imlay JA. 2001. How does oxygen inhibit central metabolism in the obligate anaerobe Bacteroides thetaiotaomicron? Mol. Microbiol. 39:1562-1571. 221. Martins BM, Dobbek H, Cinkaya I, Buckel W, Messerschmidt A. 2004. Crystal structure of 4-hydroxybutyryl-CoA dehydratase: radical catalysis involving a [4Fe-4S] cluster and flavin. Proc. Natl. Acad. Sci. U. S. A. 101:15645-15649. doi: 10.1073/pnas.0403952101. 222. Routray W, Mishra HN. 2011. Scientific and Technical Aspects of Yogurt Aroma and Taste: A Review. Comprehensive Reviews in Food Science and Food Safety. 10(4):208--220. 223. FAO/WHO (Food and Agriculture Organization/World Health Organization). 2001. Expert Consultation on Evaluation of Health and Nutritional Properties of Probiotics in Food Including Powder Milk with Live Lactic Acid Bacteria Córdoba, Argentina. . 224. Ma D, Forsythe P, Bienenstock J. 2004. Live Lactobacillus rhamnosus is essential for the inhibitory effect on tumor necrosis factor alpha-induced interleukin-8 expression. Infect. Immun. 72:5308-5314. doi: 10.1128/IAI.72.9.5308-5314.2004. 225. Gobbetti M, Cagno RD, De Angelis M. 2010. Functional microorganisms for functional food quality. Crit. Rev. Food Sci. Nutr. 50:716-727. doi: 10.1080/10408398.2010.499770. 226. Hertel C, Schmidt G, Fischer M, Oellers K, Hammes WP. 1998. Oxygen-dependent regulation of the expression of the catalase gene katA of Lactobacillus sakei LTH677. Appl. Environ. Microbiol. 64:1359-1365. 227. Storz G, Jacobson FS, Tartaglia LA, Morgan RW, Silveira LA, Ames BN. 1989. An alkyl hydroperoxide reductase induced by oxidative stress in Salmonella typhimurium and Escherichia coli: genetic characterization and cloning of ahp. J. Bacteriol. 171:2049-2055. 228. Hertzberger R, Arents J, Dekker HL, Pridmore RD, Gysler C, Kleerebezem M, Teixeira de Mattos MJ. 2014. H2O2 production in species of the Lactobacillus acidophilus group, a central role for a novel NADH dependent flavin reductase. Appl. Environ. Microbiol. . doi: 10.1128/AEM.04272-13. 229. Pedersen MB, Garrigues C, Tuphile K, Brun C, Vido K, Bennedsen M, Mollgaard H, Gaudu P, Gruss A. 2008. Impact of aeration and heme-activated respiration on Lactococcus lactis gene expression: identification of a heme-responsive operon. J. Bacteriol. 190:4903-4911. doi: 10.1128/JB.00447-08;. 230. van Hijum SA, Garcia de la Nava J, Trelles O, Kok J, Kuipers OP. 2003. MicroPreP: a cDNA microarray data pre-processing framework. Appl. Bioinformatics. 2:241-244. 231. Baldi P, Long AD. 2001. A Bayesian framework for the analysis of microarray expression data: regularized t -test and statistical inferences of gene changes. Bioinformatics. 17:509-519. 7 163 232. Pick E, Keisari Y. 1980. A simple colorimetric method for the measurement of hydrogen peroxide produced by cells in culture. J. Immunol. Methods. 38:161-170. 233. Hartley JL, Donelson JE. 1980. Nucleotide sequence of the yeast plasmid. Nature. 286:860-865. 234. Ventura M, Canchaya C, Pridmore D, Berger B, Brussow H. 2003. Integration and distribution of Lactobacillus johnsonii prophages. J. Bacteriol. 185:4603-4608. 235. Ventura M, Canchaya C, Pridmore RD, Brussow H. 2004. The prophages of Lactobacillus johnsonii NCC 533: comparative genomics and transcription analysis. Virology. 320:229-242. doi: 10.1016/j. virol.2003.11.034. 236. Waksman G, Krishna TS, Williams CH,Jr, Kuriyan J. 1994. Crystal structure of Escherichia coli thioredoxin reductase refined at 2 A resolution. Implications for a large conformational change during catalysis. J. Mol. Biol. 236:800-816. 237. van Kranenburg R, Marugg JD, van S,II, Willem NJ, de Vos WM. 1997. Molecular characterization of the plasmid-encoded eps gene cluster essential for exopolysaccharide biosynthesis in Lactococcus lactis. Mol. Microbiol. 24:387-397. 238. Storz G, Tartaglia LA, Ames BN. 1990. The OxyR regulon. Antonie Van Leeuwenhoek. 58:157-161. 239. Nishiyama Y, Massey V, Takeda K, Kawasaki S, Sato J, Watanabe T, Niimura Y. 2001. Hydrogen peroxide-forming NADH oxidase belonging to the peroxiredoxin oxidoreductase family: existence and physiological role in bacteria. J. Bacteriol. 183:2431-2438. doi: 10.1128/JB.183.8.2431-2438.2001. 240. Looijesteijn PJ, Boels IC, Kleerebezem M, Hugenholtz J. 1999. Regulation of exopolysaccharide production by Lactococcus lactis subsp. cremoris By the sugar source. Appl. Environ. Microbiol. 65:50035008. 241. van Kranenburg R, van S,II, Marugg JD, Kleerebezem M, de Vos WM. 1999. Exopolysaccharide biosynthesis in Lactococcus lactis NIZO B40: functional analysis of the glycosyltransferase genes involved in synthesis of the polysaccharide backbone. J. Bacteriol. 181:338-340. 242. Aagaard K, Riehle K, Ma J, Segata N, Mistretta TA, Coarfa C, Raza S, Rosenbaum S, Van den Veyver I, Milosavljevic A, Gevers D, Huttenhower C, Petrosino J, Versalovic J. 2012. A metagenomic approach to characterization of the vaginal microbiome signature in pregnancy. PLoS One. 7:e36466. doi: 10.1371/journal.pone.0036466. 243. Lamont RF, Sobel JD, Akins RA, Hassan SS, Chaiworapongsa T, Kusanovic JP, Romero R. 2011. The vaginal microbiome: new information about genital tract flora using molecular based techniques. BJOG. 118:533-549. doi: 10.1111/j.1471-0528.2010.02840.x;. 244. Elli M, Zink R, Reniero R, Morelli L. 1999. Growth requirements of Lactobacillus johnsonii in skim an UHT milk. International Dairy Journal. 9:507-507-513. 245. Iino T, Uchimura T, Komagata K. 2002. The effect of sodium acetate on the growth yield, the production of L- and D-lactic acid, and the activity of some enzymes of the glycolytic pathway of Lactobacillus sakei NRIC 1071(T) and Lactobacillus plantarum NRIC 1067(T). J. Gen. Appl. Microbiol. 48:91-102. 246. Chervaux C, Ehrlich SD, Maguin E. 2000. Physiological study of Lactobacillus delbrueckii subsp. bulgaricus strains in a novel chemically defined medium. Appl. Environ. Microbiol. 66:5306-5311. 247. Bernard E, Rolain T, Courtin P, Guillot A, Langella P, Hols P, Chapot-Chartier MP. 2011. Characterization of O-acetylation of N-acetylglucosamine: a novel structural variation of bacterial peptidoglycan. J. Biol. Chem. 286:23950-23958. doi: 10.1074/jbc.M111.241414. 248. Hols P, Ramos A, Hugenholtz J, Delcour J, de Vos WM, Santos H, Kleerebezem M. 1999. Acetate utilization in Lactococcus lactis deficient in lactate dehydrogenase: a rescue pathway for maintaining redox balance. J. Bacteriol. 181:5521-5526. 164 249. Bringel F, Hubert JC. 2003. Extent of genetic lesions of the arginine and pyrimidine biosynthetic pathways in Lactobacillus plantarum, L. paraplantarum, L. pentosus, and L. casei: prevalence of CO(2)dependent auxotrophs and characterization of deficient arg genes in L. plantarum. Appl. Environ. Microbiol. 69:2674-2683. 250. Stevens MJ, Wiersma A, de Vos WM, Kuipers OP, Smid EJ, Molenaar D, Kleerebezem M. 2008. Improvement of Lactobacillus plantarum aerobic growth as directed by comprehensive transcriptome analysis. Appl. Environ. Microbiol. 74:4776-4778. doi: 10.1128/AEM.00136-08. 251. Arioli S, Roncada P, Salzano AM, Deriu F, Corona S, Guglielmetti S, Bonizzi L, Scaloni A, Mora D. 2009. The relevance of carbon dioxide metabolism in Streptococcus thermophilus. Microbiology. 155:19531965. doi: 10.1099/mic.0.024737-0. 252. Nicoloff H, Hubert JC, Bringel F. 2000. In Lactobacillus plantarum, carbamoyl phosphate is synthesized by two carbamoyl-phosphate synthetases (CPS): carbon dioxide differentiates the arginine-repressed from the pyrimidine-regulated CPS. J. Bacteriol. 182:3416-3422. 253. Arsene-Ploetze F, Kugler V, Martinussen J, Bringel F. 2006. Expression of the pyr operon of Lactobacillus plantarum is regulated by inorganic carbon availability through a second regulator, PyrR2, homologous to the pyrimidine-dependent regulator PyrR1. J. Bacteriol. 188:8607-8616. doi: 10.1128/ JB.00985-06. 254. Bringel F, Vuilleumier S, Arsene-Ploetze F. 2008. Low carbamoyl phosphate pools may drive Lactobacillus plantarum CO2-dependent growth phenotype. J. Mol. Microbiol. Biotechnol. 14:22-30. doi: 10.1159/000107966. 255. Sieuwerts S, Molenaar D, van Hijum SA, Beerthuyzen M, Stevens MJ, Janssen PW, Ingham CJ, de Bok FA, de Vos WM, van Hylckama Vlieg JE. 2010. Mixed-culture transcriptome analysis reveals the molecular basis of mixed-culture growth in Streptococcus thermophilus and Lactobacillus bulgaricus. Appl. Environ. Microbiol. 76:7775-7784. doi: 10.1128/AEM.01122-10. 256. den Besten HM, Ingham CJ, van Hylckama Vlieg JE, Beerthuyzen MM, Zwietering MH, Abee T. 2007. Quantitative analysis of population heterogeneity of the adaptive salt stress response and growth capacity of Bacillus cereus ATCC 14579. Appl. Environ. Microbiol. 73:4797-4804. doi: 10.1128/AEM.0040407. 257. Santos F, Teusink B, Molenaar D, van Heck M, Wels M, Sieuwerts S, de Vos WM, Hugenholtz J. 2009. Effect of amino acid availability on vitamin B12 production in Lactobacillus reuteri. Appl. Environ. Microbiol. 75:3930-3936. doi: 10.1128/AEM.02487-08. 258. den Besten HM, Garcia D, Moezelaar R, Zwietering MH, Abee T. 2010. Direct-imaging-based quantification of Bacillus cereus ATCC 14579 population heterogeneity at a low incubation temperature. Appl. Environ. Microbiol. 76:927-930. doi: 10.1128/AEM.01372-09. 259. Ingham CJ, Beerthuyzen M, van Hylckama Vlieg J. 2008. Population heterogeneity of Lactobacillus plantarum WCFS1 microcolonies in response to and recovery from acid stress. Appl. Environ. Microbiol. 74:7750-7758. doi: 10.1128/AEM.00982-08. 260. Dirar H, Collins EB. 1973. Aerobic utilization of low concentrations of galactose by Lactobacillus plantarum. J. Gen. Microbiol. 78:211-215. 261. Murphy MG, Condon S. 1984. Correlation of oxygen utilization and hydrogen peroxide accumulation with oxygen induced enzymes in Lactobacillus plantarum cultures. Arch. Microbiol. 138:44-48. 262. Melchiorsen CR, Jokumsen KV, Villadsen J, Israelsen H, Arnau J. 2002. The level of pyruvateformate lyase controls the shift from homolactic to mixed-acid product formation in Lactococcus lactis. Appl. Microbiol. Biotechnol. 58:338-344. doi: 10.1007/s00253-001-0892-5. 263. Nicoloff H, Elagoz A, Arsene-Ploetze F, Kammerer B, Martinussen J, Bringel F. 2005. Repression of the pyr operon in Lactobacillus plantarum prevents its ability to grow at low carbon dioxide levels. J. Bacteriol. 187:2093-2104. doi: 10.1128/JB.187.6.2093-2104.2005. 7 165 264. Henriksen CM, Curic M, Nilsson D. 2000. Citrate can partially replace carbon dioxide required for growth of Lactococcus lactis subsp. lactis biovar diacetylactis. Lett. Appl. Microbiol. 30:415-418. 265. Deibel RH, Niven CF,Jr. 1955. Reciprocal replacement of oleic acid and CO2 in the nutrition of the minute streptococci and Lactobacillus leichmannii.. J. Bacteriol. 70:134-140. 266. Hammerstrom TG, Roh JH, Nikonowicz EP, Koehler TM. 2011. Bacillus anthracis virulence regulator AtxA: oligomeric state, function and CO(2) -signalling. Mol. Microbiol. 82:634-647. doi: 10.1111/j.13652958.2011.07843.x. 267. Herbert S, Newell SW, Lee C, Wieland KP, Dassy B, Fournier JM, Wolz C, Doring G. 2001. Regulation of Staphylococcus aureus type 5 and type 8 capsular polysaccharides by CO(2). J. Bacteriol. 183:4609-4613. doi: 10.1128/JB.183.15.4609-4613.2001. 268. Hester SE, Lui M, Nicholson T, Nowacki D, Harvill ET. 2012. Identification of a CO2 responsive regulon in Bordetella. PLoS One. 7:e47635. doi: 10.1371/journal.pone.0047635. 269. Bringel F, Hammann P, Kugler V, Arsene-Ploetze F. 2008. Lactobacillus plantarum response to inorganic carbon concentrations: PyrR2-dependent and -independent transcription regulation of genes involved in arginine and nucleotide metabolism. Microbiology. 154:2629-2640. doi: 10.1099/mic.0.2008/018184-0. 270. Arsene-Ploetze,F., Bringel,F. 2004. Role of inorganic carbon in lactic acid bacteria. Lait. 84:49--59. 271. Ernst J, Bar-Joseph Z. 2006. STEM: a tool for the analysis of short time series gene expression data. BMC Bioinformatics. 7:191. doi: 10.1186/1471-2105-7-191. 272. Ashburner M, Ball CA, Blake JA, Botstein D, Butler H, Cherry JM, Davis AP, Dolinski K, Dwight SS, Eppig JT, Harris MA, Hill DP, Issel-Tarver L, Kasarskis A, Lewis S, Matese JC, Richardson JE, Ringwald M, Rubin GM, Sherlock G. 2000. Gene ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat. Genet. 25:25-29. doi: 10.1038/75556. 273. Kanehisa M, Goto S, Sato Y, Kawashima M, Furumichi M, Tanabe M. 2014. Data, information, knowledge and principle: back to metabolism in KEGG. Nucleic Acids Res. 42:D199-205. doi: 10.1093/nar/ gkt1076. 274. Arsene-Ploetze F, Nicoloff H, Kammerer B, Martinussen J, Bringel F. 2006. Uracil salvage pathway in Lactobacillus plantarum: Transcription and genetic studies. J. Bacteriol. 188:4777-4786. doi: 10.1128/ JB.00195-06. 275. Ahmad SI, Kirk SH, Eisenstark A. 1998. Thymine metabolism and thymineless death in prokaryotes and eukaryotes. Annu. Rev. Microbiol. 52:591-625. doi: 10.1146/annurev.micro.52.1.591. 276. Reich J, Soska J. 1970. Thymineless death in Lactobacillus acidophilus R-26. Folia Microbiol. (Praha). 15:40-47. 277. Zelle RM, Trueheart J, Harrison JC, Pronk JT, van Maris AJ. 2010. Phosphoenolpyruvate carboxykinase as the sole anaplerotic enzyme in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 76:5383-5389. doi: 10.1128/AEM.01077-10. 278. Henriksen CM, Curic M, Nilsson D. 2000. Citrate can partially replace carbon dioxide required for growth of Lactococcus lactis subsp. lactis biovar diacetylactis. Lett. Appl. Microbiol. 30:415-418. 279. Cooper TG, Tchen TT, Wood HG, Benedict CR. 1968. The carboxylation of phosphoenolpyruvate and pyruvate. I. The active species of “CO2” utilized by phosphoenolpyruvate carboxykinase, carboxytransphosphorylase, and pyruvate carboxylase. J. Biol. Chem. 243:3857-3863. 280. Novak I, Wang J, Henriksen KL, Haanes KA, Krabbe S, Nitschke R, Hede SE. 2011. Pancreatic bicarbonate secretion involves two proton pumps. J. Biol. Chem. 286:280-289. doi: 10.1074/jbc. M110.136382. 281. Koponen J, Laakso K, Koskenniemi K, Kankainen M, Savijoki K, Nyman TA, de Vos WM, Tynkkynen S, Kalkkinen N, Varmanen P. 2012. Effect of acid stress on protein expression and phosphorylation in Lactobacillus rhamnosus GG. J. Proteomics. 75:1357-1374. doi: 10.1016/j.jprot.2011.11.009. 166 282. Koskenniemi K, Laakso K, Koponen J, Kankainen M, Greco D, Auvinen P, Savijoki K, Nyman TA, Surakka A, Salusjarvi T, de Vos WM, Tynkkynen S, Kalkkinen N, Varmanen P. 2011. Proteomics and transcriptomics characterization of bile stress response in probiotic Lactobacillus rhamnosus GG. Mol. Cell. Proteomics. 10:M110.002741. doi: 10.1074/mcp.M110.002741. 283. Maligoy M, Mercade M, Cocaign-Bousquet M, Loubiere P. 2008. Transcriptome analysis of Lactococcus lactis in coculture with Saccharomyces cerevisiae. Appl. Environ. Microbiol. 74:485-494. doi: 10.1128/AEM.01531-07. 284. Wang J, Wu R, Zhang W, Sun Z, Zhao W, Zhang H. 2013. Proteomic comparison of the probiotic bacterium Lactobacillus casei Zhang cultivated in milk and soy milk. J. Dairy Sci. 96:5603-5624. doi: 10.3168/ jds.2013-6927. 285. Verneuil N, Maze A, Sanguinetti M, Laplace JM, Benachour A, Auffray Y, Giard JC, Hartke A. 2006. Implication of (Mn)superoxide dismutase of Enterococcus faecalis in oxidative stress responses and survival inside macrophages. Microbiology. 152:2579-2589. doi: 10.1099/mic.0.28922-0. 286. Imbert M, Blondeau R. 1998. On the iron requirement of lactobacilli grown in chemically defined medium. Curr. Microbiol. 37:64-66. 287. Bienert GP, Schjoerring JK, Jahn TP. 2006. Membrane transport of hydrogen peroxide. Biochim. Biophys. Acta. 1758:994-1003. doi: 10.1016/j.bbamem.2006.02.015. 288. Gibson CM, Mallett TC, Claiborne A, Caparon MG. 2000. Contribution of NADH oxidase to aerobic metabolism of Streptococcus pyogenes. J. Bacteriol. 182:448-455. 289. Corcionivoschi N, Alvarez LA, Sharp TH, Strengert M, Alemka A, Mantell J, Verkade P, Knaus UG, Bourke B. 2012. Mucosal reactive oxygen species decrease virulence by disrupting Campylobacter jejuni phosphotyrosine signaling. Cell. Host Microbe. 12:47-59. doi: 10.1016/j.chom.2012.05.018;. 290. Niethammer P, Grabher C, Look AT, Mitchison TJ. 2009. A tissue-scale gradient of hydrogen peroxide mediates rapid wound detection in zebrafish. Nature. 459:996-999. doi: 10.1038/nature08119;. 291. Lambeth JD, Neish AS. 2014. Nox enzymes and new thinking on reactive oxygen: a double-edged sword revisited. Annu. Rev. Pathol. 9:119-145. doi: 10.1146/annurev-pathol-012513-104651. 292. Carroll IM, Andrus JM, Bruno-Barcena JM, Klaenhammer TR, Hassan HM, Threadgill DS. 2007. Anti-inflammatory properties of Lactobacillus gasseri expressing manganese superoxide dismutase using the interleukin 10-deficient mouse model of colitis. Am. J. Physiol. Gastrointest. Liver Physiol. 293:G729-38. doi: 10.1152/ajpgi.00132.2007. 293. Han W, Mercenier A, Ait-Belgnaoui A, Pavan S, Lamine F, van S,II, Kleerebezem M, SalvadorCartier C, Hisbergues M, Bueno L, Theodorou V, Fioramonti J. 2006. Improvement of an experimental colitis in rats by lactic acid bacteria producing superoxide dismutase. Inflamm. Bowel Dis. 12:1044-1052. doi: 10.1097/01.mib.0000235101.09231.9e. 294. Teixeira GS, Carvalho FP, Arantes RM, Nunes AC, Moreira JL, Mendonca M, Almeida RB, Farias LM, Carvalho MA, Nicoli JR. 2012. Characteristics of Lactobacillus and Gardnerella vaginalis from women with or without bacterial vaginosis and their relationships in gnotobiotic mice. J. Med. Microbiol. 61:10741081. doi: 10.1099/jmm.0.041962-0;. 295. Morris RL, Schmidt TM. 2013. Shallow breathing: bacterial life at low O(2). Nat. Rev. Microbiol. 11:205-212. doi: 10.1038/nrmicro2970. 296. Ceja-Navarro JA, Nguyen NH, Karaoz U, Gross SR, Herman DJ, Andersen GL, Bruns TD, PettRidge J, Blackwell M, Brodie EL. 2014. Compartmentalized microbial composition, oxygen gradients and nitrogen fixation in the gut of Odontotaenius disjunctus. ISME J. 8:6-18. doi: 10.1038/ismej.2013.134;. 297. Jones SA, Chowdhury FZ, Fabich AJ, Anderson A, Schreiner DM, House AL, Autieri SM, Leatham MP, Lins JJ, Jorgensen M, Cohen PS, Conway T. 2007. Respiration of Escherichia coli in the mouse intestine. Infect. Immun. 75:4891-4899. doi: 10.1128/IAI.00484-07. 7 167 298. Khan MT, Duncan SH, Stams AJ, van Dijl JM, Flint HJ, Harmsen HJ. 2012. The gut anaerobe Faecalibacterium prausnitzii uses an extracellular electron shuttle to grow at oxic-anoxic interphases. ISME J. 6:1578-1585. doi: 10.1038/ismej.2012.5;. 299. Wang X, Allen TD, Yang Y, Moore DR, Huycke MM. 2013. Cyclooxygenase-2 generates the endogenous mutagen trans-4-hydroxy-2-nonenal in Enterococcus faecalis-infected macrophages. Cancer. Prev. Res. (Phila). 6:206-216. doi: 10.1158/1940-6207.CAPR-12-0350. 300. Spellerberg B, Cundell DR, Sandros J, Pearce BJ, Idanpaan-Heikkila I, Rosenow C, Masure HR. 1996. Pyruvate oxidase, as a determinant of virulence in Streptococcus pneumoniae. Mol. Microbiol. 19:803813. 301. Ramos-Montanez S, Tsui HC, Wayne KJ, Morris JL, Peters LE, Zhang F, Kazmierczak KM, Sham LT, Winkler ME. 2008. Polymorphism and regulation of the spxB (pyruvate oxidase) virulence factor gene by a CBS-HotDog domain protein (SpxR) in serotype 2 Streptococcus pneumoniae. Mol. Microbiol. 67:729-746. doi: 10.1111/j.1365-2958.2007.06082.x. 302. Carvalho SM, Farshchi Andisi V, Gradstedt H, Neef J, Kuipers OP, Neves AR, Bijlsma JJ. 2013. Pyruvate oxidase influences the sugar utilization pattern and capsule production in Streptococcus pneumoniae. PLoS One. 8:e68277. doi: 10.1371/journal.pone.0068277. 303. Zhu L, Xu Y, Ferretti JJ, Kreth J. 2014. Probing oral microbial functionality--expression of spxB in plaque samples. PLoS One. 9:e86685. doi: 10.1371/journal.pone.0086685;. 304. Zhu L, Kreth J. 2012. The role of hydrogen peroxide in environmental adaptation of oral microbial communities. Oxid Med. Cell. Longev. 2012:717843. doi: 10.1155/2012/717843;. 305. Itzek A, Zheng L, Chen Z, Merritt J, Kreth J. 2011. Hydrogen peroxide-dependent DNA release and transfer of antibiotic resistance genes in Streptococcus gordonii. J. Bacteriol. 193:6912-6922. doi: 10.1128/ JB.05791-11. 306. Xu Y, Kreth J. 2013. Role of LytF and AtlS in eDNA release by Streptococcus gordonii. PLoS One. 8:e62339. doi: 10.1371/journal.pone.0062339. 307. Lau K, Benitez P, Ardissone A, Wilson TD, Collins EL, Lorca G, Li N, Sankar D, Wasserfall C, Neu J, Atkinson MA, Shatz D, Triplett EW, Larkin J,3rd. 2011. Inhibition of type 1 diabetes correlated to a Lactobacillus johnsonii N6.2-mediated Th17 bias. J. Immunol. 186:3538-3546. doi: 10.4049/ jimmunol.1001864. 308. Valladares R, Sankar D, Li N, Williams E, Lai KK, Abdelgeliel AS, Gonzalez CF, Wasserfall CH, Larkin J, Schatz D, Atkinson MA, Triplett EW, Neu J, Lorca GL. 2010. Lactobacillus johnsonii N6.2 mitigates the development of type 1 diabetes in BB-DP rats. PLoS One. 5:e10507. doi: 10.1371/journal. pone.0010507; 10.1371/journal.pone.0010507. 309. Mellor AL, Munn DH. 2004. IDO expression by dendritic cells: tolerance and tryptophan catabolism. Nat. Rev. Immunol. 4:762-774. doi: 10.1038/nri1457. 310. Gajer P, Brotman RM, Bai G, Sakamoto J, Schutte UM, Zhong X, Koenig SS, Fu L, Ma ZS, Zhou X, Abdo Z, Forney LJ, Ravel J. 2012. Temporal dynamics of the human vaginal microbiota. Sci. Transl. Med. 4:132ra52. doi: 10.1126/scitranslmed.3003605;. 311. Romero R, Hassan SS, Gajer P, Tarca AL, Fadrosh DW, Nikita L, Galuppi M, Lamont RF, Chaemsaithong P, Miranda J, Chaiworapongsa T, Ravel J. 2014. The composition and stability of the vaginal microbiota of normal pregnant women is different from that of non-pregnant women. Microbiome. 2:4-2618-2-4. doi: 10.1186/2049-2618-2-4. 312. Ma B, Forney LJ, Ravel J. 2012. Vaginal microbiome: rethinking health and disease. Annu. Rev. Microbiol. 66:371-389. doi: 10.1146/annurev-micro-092611-150157. 313. Ravel J, Gajer P, Abdo Z, Schneider GM, Koenig SS, McCulle SL, Karlebach S, Gorle R, Russell J, Tacket CO, Brotman RM, Davis CC, Ault K, Peralta L, Forney LJ. 2011. Vaginal microbiome of reproductiveage women. Proc. Natl. Acad. Sci. U. S. A. 108 Suppl 1:4680-4687. doi: 10.1073/pnas.1002611107;. 168 314. Yeoman CJ, Thomas SM, Miller ME, Ulanov AV, Torralba M, Lucas S, Gillis M, Cregger M, Gomez A, Ho M, Leigh SR, Stumpf R, Creedon DJ, Smith MA, Weisbaum JS, Nelson KE, Wilson BA, White BA. 2013. A multi-omic systems-based approach reveals metabolic markers of bacterial vaginosis and insight into the disease. PLoS One. 8:e56111. doi: 10.1371/journal.pone.0056111. 315. Lewis WG, Robinson LS, Gilbert NM, Perry JC, Lewis AL. 2013. Degradation, foraging, and depletion of mucus sialoglycans by the vagina-adapted Actinobacterium Gardnerella vaginalis. J. Biol. Chem. 288:12067-12079. doi: 10.1074/jbc.M113.453654;. 316. Gilbert NM, Lewis WG, Lewis AL. 2013. Clinical features of bacterial vaginosis in a murine model of vaginal infection with Gardnerella vaginalis. PLoS One. 8:e59539. doi: 10.1371/journal.pone.0059539. 317. Kenyon C, Colebunders R, Crucitti T. 2013. The global epidemiology of bacterial vaginosis: a systematic review. Am. J. Obstet. Gynecol. . doi: 10.1016/j.ajog.2013.05.006;. 318. Antonio MA, Rabe LK, Hillier SL. 2005. Colonization of the rectum by Lactobacillus species and decreased risk of bacterial vaginosis. J. Infect. Dis. 192:394-398. doi: 10.1086/430926. 319. Gupta K, Stapleton AE, Hooton TM, Roberts PL, Fennell CL, Stamm WE. 1998. Inverse association of H2O2-producing lactobacilli and vaginal Escherichia coli colonization in women with recurrent urinary tract infections. J. Infect. Dis. 178:446-450. 320. Cruickshank R. 1931. Döderlein’s Vaginal Bacillus: A Contribution to the Study of the Lacto-Bacilli. J Hyg. 3:375--381. 321. Ocana VS, de Ruiz Holgado AA, Nader-Macias ME. 1999. Growth inhibition of Staphylococcus aureus by H2O2-producing Lactobacillus paracasei subsp. paracasei isolated from the human vagina. FEMS Immunol. Med. Microbiol. 23:87-92. 322. O’Hanlon DE, Moench TR, Cone RA. 2011. In vaginal fluid, bacteria associated with bacterial vaginosis can be suppressed with lactic acid but not hydrogen peroxide. BMC Infect. Dis. 11:200-2334-11-200. doi: 10.1186/1471-2334-11-200;. 323. Mshvildadze M, Neu J, Shuster J, Theriaque D, Li N, Mai V. 2010. Intestinal microbial ecology in premature infants assessed with non-culture-based techniques. J. Pediatr. 156:20-25. doi: 10.1016/j. jpeds.2009.06.063;. 324. Dominguez-Bello MG, Costello EK, Contreras M, Magris M, Hidalgo G, Fierer N, Knight R. 2010. Delivery mode shapes the acquisition and structure of the initial microbiota across multiple body habitats in newborns. Proc. Natl. Acad. Sci. U. S. A. 107:11971-11975. doi: 10.1073/pnas.1002601107;. 325. Jakobsson HE, Abrahamsson TR, Jenmalm MC, Harris K, Quince C, Jernberg C, Bjorksten B, Engstrand L, Andersson AF. 2014. Decreased gut microbiota diversity, delayed Bacteroidetes colonisation and reduced Th1 responses in infants delivered by Caesarean section. Gut. 63:559-566. doi: 10.1136/ gutjnl-2012-303249;. 326. Hallstrom M, Eerola E, Vuento R, Janas M, Tammela O. 2004. Effects of mode of delivery and necrotising enterocolitis on the intestinal microflora in preterm infants. Eur. J. Clin. Microbiol. Infect. Dis. 23:463-470. doi: 10.1007/s10096-004-1146-0. 327. Torrazza RM, Ukhanova M, Wang X, Sharma R, Hudak ML, Neu J, Mai V. 2013. Intestinal microbial ecology and environmental factors affecting necrotizing enterocolitis. PLoS One. 8:e83304. doi: 10.1371/ journal.pone.0083304;. 328. El Aidy S, Van den Abbeele P, Van de Wiele T, Louis P, Kleerebezem M. 2013. Intestinal colonization: how key microbial players become established in this dynamic process: microbial metabolic activities and the interplay between the host and microbes. Bioessays. 35:913-923. doi: 10.1002/bies.201300073;. 329. Castagliuolo I, Galeazzi F, Ferrari S, Elli M, Brun P, Cavaggioni A, Tormen D, Sturniolo GC, Morelli L, Palu G. 2005. Beneficial effect of auto-aggregating Lactobacillus crispatus on experimentally induced colitis in mice. FEMS Immunol. Med. Microbiol. 43:197-204. doi: 10.1016/j.femsim.2004.08.011. 7 169 Summary (for a scientific audience) Lactic acid bacteria (LAB) are intimately entwined with human life. They ferment several key food products in our diets and they reside on the mucosal surfaces of the mouth, intestine and vagina. Administration of adequate amounts of certain LAB species has been shown to provide health benefits, such as attenuation of antibiotic-associated diarrhoea. Lactobacillus johnsonii is a LAB that is used in the food industry for such health-benefit, or probiotic effects. The functionality of L. johnsonii and other related LAB in the food industry and in the host-related environment are strongly dependent on environmental factors. Especially oxygen and carbon dioxide were found to have a major influence on metabolism, growth, gene expression, yield, and viability. This thesis addresses the consequences of exposure to these two gaseous components on L. johnsonii. Chapter 1 contains a general overview of research literature on aerobic metabolism and oxidative stress of lactic acid bacteria. LAB are classified as aerotolerant anaerobes; although they commonly grow well in the presence of oxygen they do not show aerobic respiration unless a hemin source is added. Instead, oxygen is used for direct oxidation of various metabolic intermediates. Facultative heterolactic LAB produce acetate and CO2 aerobically through the pyruvate oxidase and lactate oxidase pathway. These products are more oxidized than lactate and therefore additional oxidation of the reducing equivalent NADH is required. When exogenous hemin is added to the medium, NADH can be channelled through the electron transport chain. This respiratory growth is associated with higher yield, robustness and lower oxidative stress. Alternatively, water or H2O2-forming NADH oxidases can regenerate NAD+. NADH oxidase is a central switch in the metabolism of heterolactic LAB. Activity of these oxidases and autoxidation of other cellular components results in the generation of reactive oxygen species, such as hydrogen peroxide and superoxide, which cause oxidative stress. We provide an overview of the anti-oxidative enzymes (ROS-scavengers) and physiological adaptation found in LAB to reduce oxidative stress. A prominent characteristic that L. johnsonii shares with several other lactobacilli and streptococci is the accumulation of substantial amounts (>1 mM) of hydrogen peroxide in its environment. L. johnsonii lacks the key ROS scavenging enzymes, such as catalase, alkyl hydroperoxide reductase and superoxide dismutase. Hydrogen peroxide accumulation is the primary cause of oxidative stress in L. johnsonii. It induces a premature stationary phase and a ~10-fold lower biomass yield in the presence of 170 oxygen. Addition of catalase abolishes this growth stagnation. In chapter 2 and 3 we report on the identification and characterization of two proteins that contribute to hydrogen peroxide production of L. johnsonii. In chapter 2 we showed that H2O2 production is unrelated to expression of pyruvate, lactate and NADH oxidase. In cell extracts of L. johnsonii an enzymatic hydrogen peroxide forming activity could be detected upon addition of flavin and NADH (not NADPH). Partial purification of the unidentified enzyme displaying this activity showed that two small flavoproteins, LJ_0548 and LJ_0549, were overrepresented in the hydrogen peroxide forming fraction. Genetic disruption and overexpression confirmed that these proteins constitute an NADH flavin reductase (NFR). A deletion derivative of these two genes did not produce any hydrogen peroxide when exposed to oxygen, indicating that these enzymes catalyze the reaction that produces hydrogen peroxide in L. johnsonii. However, after prolonged cultivation in the presence of oxygen, the NFR deletion derivative regained partial H2O2 producing capacity. In chapter 3 we report on the identification of a second H2O2 producing enzyme, an NADH oxidase (NOX), encoded by the LJ_1254-1255 locus. Expression of this locus was 2.1-fold induced in the wildtype under aerobic growth conditions and 3.7-fold in the NFR-deletion derivative. Deficiency of this NOX activity did not impact H2O2 production of the wildtype, but completely abolished all H2O2 production in its NFR deficient derivative. Intriguingly, this mutant also showed hampered growth and lower biomass yield in the presence of oxygen, despite its H2O2-negative phenotype. We conclude that the oxygen-induced NADH oxidase produces hydrogen peroxide in the absence of NADH flavin reductase, and may also contribute to hydrogen peroxide production during longer exposure to oxygen. Oxygen is not in all cases detrimental for growth and viability of L. johnsonii. In the second part of this thesis we focus on the growth stagnation observed during growth under a N2 atmosphere, observed both in liquid (sparged batch cultures) as on solid media (AnoporeTM slides). The cause of this growth stagnation was shown to reflect a lack of CO2 which L. johnsonii requires for growth. Two aspects of this observation were unexpected. Firstly, oxygen could fully relieve this CO2 growth dependency, and secondly, lack of CO2 apparently led to cell death of L. johnsonii. Especially the latter 7 effect is unusual, since removal of essential nutrients generally only halts bacterial growth. We further study these two factors in chapter 4 and 5. In chapter 4, we showed that this oxygen relieve of CO2 dependency also accounted for the acetate growth dependency: L. johnsonii cannot grow in an environment without acetate unless the culture is aerated. Both these effects of oxygen could be traced back to one 171 common denominator, which is the pyruvate oxidase reaction leading to both CO2 and acetate production. Pyruvate oxidase could therefore allow L. johnsonii to overcome its acetate and CO2 dependency in conditions that include oxygen exposure. A pyruvate oxidase deletion derivative confirmed this hypothesis as it rendered the organism both aerobically and anaerobically dependent on acetate and CO2 supplementation. Our results demonstrate that certain growth requirements of L. johnsonii are not hardwired but depend on environmental factors. In chapter 5, we attempted to further understand the metabolic requirement for CO2 by analyzing genome-wide transcriptome changes in an anaerobically growing culture upon CO2 depletion. We detected an extensive rearrangement of gene expression, including many transporters and regulators. Additionally, expression of the pyr-operon encoded pyrimidine biosynthesis pathway, also referred to as the carbamoyl-phosphate pathway, was strongly upregulated (up to 17-fold). In other LAB, this pathway has been associated with CO2 dependency, which is required in the first enzymatic step of the pathway and generally growth stagnation due to CO2 depletion could in these LAB be prevented by addition of sufficient levels of pyrimidines. However, in L. johnsonii such a relation between pyrimidine supplementation and CO2 dependency was not observed. Supplementation of other compounds that have previously been associated with CO2 growth dependency in LAB, such as arginine and aspartate also could not prevent CO2 induced growth stagnation. We speculate that L. johnsonii is unable to incorporate exogenous pyrimidines and propose further experiments to test this preposition. In chapter 6, we present a general discussion of the results in this thesis and place them in the environmental context of the intestinal and vaginal microbiota. One of the most prominent questions that remains is why NOX and NFR activity is required for aerotolerance. We propose that H2O2 production of L. johnsonii may be a means to scavenge oxygen in an physiologically attractive way, i.e. without spending much NADH which is ultimately required for lactate production. High levels of intracellular molecular oxygen and possible resulting superoxide formation could be more hazardous than hydrogen peroxide, which quickly diffuses out of the cell and is potentially scavenged by neighboring catalase-expressing bacteria or other environmental factors. We propose several experiments to study this hypothesis. Lastly, we discuss how hydrogen peroxide production and aerotolerance could play a role in the interactions of L. johnsonii with a host organism. Reactive oxygen species play a central role in a wide variety of immune reactions. We discuss how bacterially- 172 derived superoxide and hydrogen peroxide could contribute to gut homeostasis through immune signaling. Especially in the neonatal intestine, which generally contains higher levels of oxygen, H2O2-producing lactobacilli, transferred from the mother to the infant during birth, could produce substantial levels of hydrogen peroxide, which may contribute to shaping the neonatal immune system. We propose how the NOX and/ or NFR deletion derivatives that were constructed in the scope of this thesis, can be employed to testing the role of bacterial H2O2 in host/microbiome interactions. 7 173 Samenvatting (voor breed publiek) Melkzuurbacteriën zijn belangrijke organismen in het leven van de mens. Niet alleen zijn ze essentieel in de bereiding van gefermenteerde producten in ons voedselpatroon, zoals yoghurt, kaas, olijven, zuurdesem en vleesproducten, maar ze zijn ook prominente bewoners van ons lichaam, bijvoorbeeld in de mond, darm en vagina. Verder hebben sommige melkzuurbacteriën zogenaamde probiotische eigenschappen: als je er voldoende van consumeert kan dat een gezondheidseffect teweeg brengen, zoals het verminderen van de ernst van diarree. In dit proefschrift presenteren we onderzoek over één van deze probiotische melkzuurbacteriën, Lactobacillus johnsonii en we concentreren ons specifiek op de invloed van zuurstof (O2) en koolstofdioxide (CO2) op metabolisme en fysiologie van deze bacterie. Met deze gassen komt L. johnsonii veel in aanraking, zowel in de voedingsmiddelenindustrie als in het menselijk lichaam. Het is daarom belangrijk om de nogal ingrijpende rol van deze twee gassen beter te begrijpen. In hoofdstuk 1, geven we een overzicht van de huidige kennis over de invloed van zuurstof op melkzuurbacteriën (LAB). Zuurstof is voor veel levensvormen schadelijk, zelfs voor soorten die geen vijf minuten zonder zuurstof kunnen overleven zoals de mens. Tijdens de ademhaling wordt zuurstof omgezet in water (H2O), maar als bijproduct worden giftige zuurstofradicalen geproduceerd, zoals waterstof peroxide (H2O2) en superoxide (O2-), die continue uit de weg moeten worden geruimd. Melkzuurbacteriën kunnen niet ademhalen, maar ze gebruiken zuurstof wel op andere manieren, bijvoorbeeld in de omzetting van melkzuur naar azijnzuur. In dat proces worden veel zuurstofradicalen geproduceerd, met name H2O2. Ook L. johnsonii produceert veel H2O2 maar heeft geen adequate mogelijkheid om dit op te ruimen. Ophoping van die H2O2 zorgt ervoor dat deze bacterie niet meer verder kan groeien na een paar uren beluchting. Uit ons onderzoek blijkt dat, in tegenstelling tot veel andere melkzuurbacteriën, de azijnzuurproductie geen prominente bron is van H2O2. In hoofdstuk 2 presenteren we een aantal experimenten waarin we op zoek gaan naar de oorsprong van die H2O2 in L. johnsonii. In cellen worden chemische reacties gefaciliteerd (gekatalyseerd) door eiwitten, en eiwitten worden gemaakt door het kopiëren van erfelijk materiaal (het DNA, de genen). We hebben het eiwit dat verantwoordelijk is voor de H2O2 productie kunnen identificeren, door alle eiwitten van L. johnsonii te fractioneren en de H2O2 productie in elk van die fracties te meten. Door de fracties telkens kleiner te maken konden we uiteindelijk de chemische reactie toeschrijven aan een handjevol eiwitten. Na analyse van die eiwitten, met zogeheten massaspectrometrie, konden we twee kleine eiwitten detecteren met een aantal eigenschappen die erop wezen dat ze inderdaad de H2O2 174 productie in L. johnsonii zouden kunnen faciliteren. Die suggestie werd bevestigd: als we de genen die voor die specifieke eiwitten coderen, uit de bacterie verwijderen, maakt de resulterende gemuteerde bacterie ook geen H2O2 meer. Alhoewel, na langdurige blootstelling aan zuurstof, blijkt ook in deze mutant bacterie de H2O2 productie weer terug te komen. Er is blijkbaar dus nog een tweede eiwit dat H2O2 vorming faciliteert. In hoofdstuk 3 beschrijven we de zoektocht naar dat tweede eiwit. Door te meten welke genen L. johnsonii aan en uitzet bij zuurstof blootstelling, kwamen we erachter dat het gen dat het sterkst wordt aangeschakeld door zuurstof, voor een eiwit codeert dat ook eigenschappen heeft die erop wijzen dat het een rol kan spelen in H2O2 productie. Als we het gen van dit eiwit verwijderen, verandert er niets aan de bacterie, maar het verwijderen van beide genen (ook het gen dat we hadden gevonden in hoofdstuk 2) levert een bacterie op die helemaal geen H2O2 meer maakt. Deze bacterie bleek alleen ook niet meer goed te kunnen groeien bij beluchting. Dat was een opvallende karakteristiek, omdat juist die H2O2 vorming zo schadelijk was. Kennelijk spelen deze twee eiwitten L. johnsonii een dubbelrol, waarbij ze zowel het schadelijke H2O2 vormen, maar tegelijkertijd op een tot nu toe onbekende manier de bacterie beschermen tegen andere schadelijke bijproducten van zuurstof. In het tweede deel van dit proefschrift beschrijven we het effect van kooldioxide, CO2. L. johnsonii heeft CO2 nodig om te kunnen groeien en om in leven te blijven. Het is opvallend dat als we zuurstof toevoegen aan de atmosfeer waarin de bacteriën groeien, er ineens wél gegroeid kan worden zonder CO2. In hoofdstuk 4 zijn we op zoek gegaan naar de oorzaak van dit zuurstof-effect. Ten eerste constateerden we dat wat geldt voor CO2 afhankelijkheid, ook blijkt te gelden voor azijnzuur-afhankelijkheid. L. johnsonii heeft azijnzuur nodig om te kunnen groeien, maar in de aanwezigheid van zuurstof is die azijnzuur afhankelijkheid opgeheven. Eén reactie blijkt hierin een centrale rol te spelen. Pyruvaat is een metaboliet dat kan reageren met zuurstof waarbij CO2 en azijnzuur worden gevormd. Deze reactie wordt gefaciliteerd door het eiwit pyruvaat oxidase. De hypothese was dat L. johnsonii zonder azijnzuur en CO2 kan groeien als zuurstof aanwezig is, omdat onder die condities beide stoffen gemaakt kunnen worden door de reactie tussen pyruvaat en zuurstof. Dit bleek te kloppen, 7 omdat na uitschakeling van het gen voor pyruvaat oxidase L. johnsonii ook niet meer zonder azijnzuur en CO2 kon groeien als er zuurstof aanwezig was. We concluderen dat zuurstof niet alleen maar schadelijk is voor deze bacterie, maar dat zuurstof er ook voor kan zorgen dat de bacterie kan groeien zonder azijnzuur en CO2. 175 In hoofdstuk 5 concentreren we ons verder op de vraag waarom L. johnsonii afhankelijk is van CO2. Zulk soort afhankelijkheid wordt in andere melkzuurbacteriën veroorzaakt doordat CO2 nodig is als grondstof voor het maken van pyrimidine, een bouwsteen van het DNA. We analyseren welke genen in L. johnsonii aan en uit worden gezet wanneer CO2 wordt verwijderd uit de omgeving. De expressie (aan/uit) van zeer veel genen blijkt te reageren op de aan of afwezigheid van CO2, voornamelijk genen die betrokken zijn bij membraantransport en genregulatie. Daarnaast zien we dat genen die betrokken zijn bij de eerder genoemde productie van DNA-bouwstenen (uit CO2) zeer sterk worden aangeschakeld bij verwijdering van CO2, wat erop kan duiden dat de productie van DNA bouwstenen inderdaad de belangrijkste oorzaak is van de CO2 behoefte van L. johnsonii. In andere melkzuurbacteriën kon het toevoegen van deze bouwstenen aan de groeimedia de afhankelijkheid voor CO2 opheffen. In onze experimenten bleek dit niet op te gaan voor L. johnsonii. Onze hypothese is dat L. johnsonii de toegevoegde DNA-bouwstenen niet goed kan inbouwen en we stellen een aantal experimenten voor waarmee die hypothese zou kunnen worden getest. In hoofdstuk 6 presenteren we een algemene analyse van al onze resultaten. Een prominente vraag die overblijft is waarom L. johnsonii de H2O2-producerende eiwitten nodig heeft om in de aanwezigheid van zuurstof te kunnen groeien. Onze hypothese is dat deze bacterie zuurstof liever omzet in H2O2 dat snel naar buiten verdwijnt en daar uit de weg wordt geruimd door omliggende bacteriën. Misschien kan L. johnsonii daarmee voorkomen dat er nog schadelijkere zuurstof radicalen zoals superoxide worden gevormd. Verder bespreken we hoe H2O2 productie door bacteriën een rol kan spelen in de interacties met de gastheer. Het is welbekend dat zuurstofradicalen, geproduceerd door de cellen van de darmwand, een regulatie effect uitoefenen op ons immuunsysteem. De vraag is of zuurstofradicalen die door darmbacteriën worden geproduceerd ook kunnen bijdragen aan deze immuunregulatie. Dit zou vooral belangrijk kunnen zijn in de darm van pasgeboren baby’s, waar vermoedelijk veel zuurstof aanwezig is om zuurstofradicalen te produceren. De vaginale bacteriën van de moeder, die tijdens de geboorte worden overgedragen aan het kind, zijn typisch H2O2-producerende lactobacillen, sterk verwant aan L. johnsonii. We stellen voor dat dit een manier is waarop bacteriën een rol kunnen spelen bij de vroege vormgeving en regulatie van het immuunsysteem. De kennis die is ontwikkeld in dit promotieonderzoek, en de mutanten die we hebben geconstrueerd, zouden een bijdrage kunnen leveren aan het bevestigen van de voorgestelde rol die bacteriën zoals L. johnsonii zouden kunnen 176 spelen in de interacties met de gastheer. List of Abbreviations ACK AHP / AHPR ATP / ADP CCP-A COX DLD FAD FMN GSH(R) HemCat LAB LDH LOX MnCat MnSOD NADH / NAD NOX NPR PAT PDH PFL POX ROS SOD spx TRX(R) acetate kinase alkyl hydroperoxide (reductase) adenosine tri/diphosphate catabolite control protein A cytochrome oxidase dihydrolipoamide dehydrogenase flavin adenine dinucleotide flavin mononucleotide Glutathione (reductase) hemin-dependent catalase lactic acid bacteria lactate dehydrogenase lactate oxidase manganese dependent catalase manganese-dependent SOD nicotinamide adenine dinucleotide NADH oxidase NADH peroxidase phosphate acetyltransferase pyruvate dehydrogenase pyruvate formate lyase pyruvate oxidase reactive oxygen species superoxide dismutase pyruvate oxidase encoding gene Thioredoxin (reductase) 7 177 Dankwoord Dit proefschrift is het stoffelijke eindproduct van vijf jaar promotieonderzoek. Vijf jaren van verrassende inzichten en harde lessen, fundamentele en toegepaste vragen, inspirerende ontmoetingen en goede gesprekken, Eureka-momenten en grote teleurstellingen, anoniem zwoegen en publiek koketteren. Het is in vele opzichten een verrijkende tijd geweest. Daarvoor ben ik veel mensen dankbaar. Te beginnen bij mijn promotores: Joost Teixeira de Mattos. Nadat Jeroen Hugenholtz dit project helaas vroegtijdig moesten verlaten, was jij bereid om de supervisie namens de UvA over te nemen. Je was de tweede die ik tegenkwam op de UvA en dat was een hele prettige kennismaking die ervoor zorgde dat ik graag bij SILS kwam werken. Ik bewonder je bevlogenheid, je liefde voor het vak en voor het onderwijs. Je had een onverwoestbaar vertrouwen in mij als wetenschapper en in de goede afloop van dit onderzoek. Dat heb ik als heel waardevol ervaren. Je zorgde regelmatig voor licht en lucht in dit project, met je aanstekelijke positivisme en humor. Michiel Kleerebezem. Nadat Eddy Smit dit project helaas vroegtijdig moest verlaten, was jij bereid om de supervisie namens NIZO over te nemen. Een enorme aanwinst! Ondanks je overvolle schema, nam je met regelmaat uitgebreid de tijd om van gedachten te wisselen. Onvermoeibaar tilde je mijn schrijfsels naar een hoger niveau. Je voortdurende nieuwsgierigheid en je aanmoediging voor nieuwe experimenten, betere teksten, consistentere notaties waren een zegen voor dit proefschrift en voor mijn ontwikkeling. Ik heb ontzettend veel van je geleerd, het was een privilege om met je samen te werken. Christof Gysler and David Pridmore. It was a pleasure to have worked together with you. This project gained a lot from the continuous interest from Nestlé’s side and your input during our monthly updates and the Kluyver meetings. The mutant strains were an essential element in this thesis, which we owe to the excellent cloning skills of David and his perseverance. Ik wil de Leescommissie bedanken voor het kritisch lezen van mijn manuscript. Het was een genoegen om onderdeel uit te maken van de MMP groep, met zijn vele bijzondere persoonlijkheden. Zowel in kantoor C3.258 als in het 12:00 lunchgroepje, werd er veel gediscussieerd, geroddeld, gescholden, gesnoept, gewerkt, gelanterfant, en ook veel en hard gelachen. Dit waren voor mij ook vooral hele gezellige jaren. Dankzij 178 jullie, lieve collega’s! In het bijzonder: Martijn, jij was een baken van enthousiasme, uitbundigheid en onredelijk positivisme in het overheersend cynische academische landschap. Gertien, ik heb veel van je geleerd en ik bewonder je ambitie, je kracht, je toewijding en je onvermoeibare streven naar meer en beter wetenschappelijk onderzoek en onderwijs. Jos, ik heb jouw creatieve projecten en prachtige foto’s zeer gewaardeerd! Jeroen, Andreas, Pascal, Aniek, jullie hebben me geholpen met de meest uiteenlopende zaken zoals boekjes drukken, figuren maken, wetenschapper worden en hoe je moet omgaan met de nieuwste Android-versie op je telefoon. Milou, Poonam dank voor de vriendschap en alle mooie tijden die we samen beleefden. Felipe, you arrived a bit late but in those few months you added a lot of energy, new ideas and a lot of fun. Johan, in lab, kantoor en kantine ben jij de afgelopen jaren een dierbare vriend geworden en ik ben dankbaar dat je mijn paranimf wilt zijn. Sarah, jij bent een representant van de vele intelligente, ambitieuze en lieve vrienden die ik de afgelopen jaren in Amsterdam heb gemaakt. Dank dat je mijn paranimf wilt zijn. All other colleagues and friends, although I will not list all your names, you know who you are, thanks for everything! Annereinou, Iris, Marjo en vele andere NIZO’ers, ondanks dat ik geen échte NIZO’er was, voelde ik me altijd welkom en was het niet alleen nuttig, maar ook ontzettend gezellig om naar Ede te komen. Dank! Dr. Rebecca ten Cate, mama, jij leerde mij dat een promotieonderzoek een voorrecht is. Je bracht me liefde voor de wetenschap bij, doorzettingsvermogen en geduld. Je leerde me hoe je zeven levens kan hebben en elke tegenslag te boven kan komen. Je bent een blijvend voorbeeld voor mij. Papa, een aantal eigenschappen die essentieel waren voor dit project en andere activiteiten, heb ik aan jou te denken. Schaamteloos ambitieus zijn, groot denken, hoffelijkheid en positivisme. Ik ben gezegend met een inspirerende familie. Liefste Arjen, zonder jou was dit boekje er ook gekomen, maar door jou was ik een hele gelukkige, vrolijke en verliefde promotiestudent. Dank je wel, dat je geduldig was, dat je me altijd steunt en dat je me telkens weer aanmoedigt om hoger te mikken. Ik 7 heb ontzettend veel zin in ons volgende hoofdstuk in St. Louis. 179 About the author Rosanne Y. Hertzberger (1984, Rotterdam) completed a BSc and MSc in Life science & Technology from Delft University of Technology and Leiden University, including courses at the École nationale du genie rural, des eaux et des fôrets in Paris. During her master thesis she studied micronutrient accumulation in Saccharomyces cerevisiae at the Nestlé Research Centre in Switzerland. In April 2009 she started her PhD project as a collaboration between the Nestlé Research Centre and NIZO Food Research, carried out as a part of the Molecular Microbial Physiology lab of the Swammerdam Institute for Life Sciences, University of Amsterdam. During her PhD studies she wrote a weekly column for the Dutch newspapers NRC Handelsblad and nrc.next. From July 8th 2014, she will start a postdoctoral research project on the role of vaginal lactobacilli in women’s health at the Centre for Women’s Infectious Disease Research at Washington University in St Louis, Missouri, USA. 180