Advances in Microbial Physiology, Volume 57

Advances in

MICROBIAL PHYSIOLOGY VOLUME 57

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Advances in

MICROBIAL PHYSIOLOGY Edited by

ROBERT K. POOLE West Riding Professor of Microbiology Department of Molecular Biology and Biotechnology The University of Sheffield Firth Court, Western Bank Sheffield, UK

VOLUME 57

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London • New York • Oxford Singapore • Sydney • Tokyo

Academic Press is an imprint of Elsevier

ACADEMIC PRESS

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Contents

CONTRIBUTORS TO VOLUME 57 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

vii

Ammonia-Oxidising Archaea – Physiology, Ecology and Evolution Christa Schleper and Graeme W. Nicol

1. 2. 3. 4. 5. 6. 7. 8.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Discovery of Archaea in Moderate Aerobic Habitats . . . . . . . . . . . . . . . . 4 First Insights into the Physiology of Ammonia-oxidising Archaea . . . 6 Model Organisms of Ammonia-Oxidising Archaea . . . . . . . . . . . . . . . . . . 9 Membrane Lipids of Ammonia-Oxidising Archaea . . . . . . . . . . . . . . . . . 13 Genomes and Metagenomes of Ammonia-Oxidising Archaea . . . . . . 15 Diversity, Distribution and Activity of Ammonia-oxidising Archaea in the Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Acknowledgement. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

Reductive Stress in Microbes: Implications for Understanding Mycobacterium tuberculosis Disease and Persistence Aisha Farhana, Loni Guidry, Anup Srivastava, Amit Singh, Mary K. Hondalus and Adrie J.C. Steyn

1. 2. 3. 4.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Scope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46 The Concept of Reductive Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Overview: General Physiological Characteristics of Mycobacterium Tuberculosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

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CONTENTS

5. 6. 7.

Reductive Sinks in Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Redox Sinks in Mycobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98

Regulation of CtsR Activity in Low GC, Gram+ Bacteria Alexander K.W. Elsholz, Ulf Gerth and Michael Hecker

1. 2. 3. 4. 5. 6.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Protein Quality Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 CtsR-Regulated Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Cellular Functions of Genes Regulated by CtsR . . . . . . . . . . . . . . . . . . . 125 Mechanisms for the Inactivation of the CtsR Repressor . . . . . . . . . . . 129 Control of CtsR Degradation by the Regulated Adaptor McsB . . . . 136 Summary and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137

AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 SUBJECT INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167

Contributors to Volume 57

ALEXANDER K.W. ELSHOLZ, Ernst-Moritz-Arndt-University Greifswald, Institute of Microbiology, Greifswald, Germany AISHA FARHANA, Department of Microbiology, University of Alabama at Birmingham, AL, USA ULF GERTH, Ernst-Moritz-Arndt-University Greifswald, Institute of Microbiology, Greifswald, Germany LONI GUIDRY, Department of Microbiology, University of Alabama at Birmingham, AL, USA MICHAEL HECKER, Ernst-Moritz-Arndt-University Greifswald, Institute of Microbiology, Greifswald, Germany MARY K. HONDALUS, Department of Infectious Diseases, University of Georgia, Athens, GA, USA GRAEME W. NICOL, Institute of Biological & Environmental Sciences, Cruickshank Building, University of Aberdeen, Aberdeen, UK CHRISTA SCHLEPER, Department of Genetics in Ecology, University of Vienna, Vienna, Austria AMIT SINGH, International Center for Genetic Engineering Biotechnology, Aruna Asaf Ali Marg, New Delhi, India

and

ANUP SRIVASTAVA, Department of Microbiology, University of Alabama at Birmingham, AL, USA ADRIE J.C. STEYN, Department of Microbiology, University of Alabama at Birmingham, AL, USA

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Ammonia-Oxidising Archaea – Physiology, Ecology and Evolution Christa Schleper1 and Graeme W. Nicol2 1

Department of Genetics in Ecology, University of Vienna, Vienna, Austria Institute of Biological & Environmental Sciences, Cruickshank Building, University of Aberdeen, Aberdeen, UK

2

ABSTRACT Nitrification is a microbially mediated process that plays a central role in the global cycling of nitrogen and is also of economic importance in agriculture and wastewater treatment. The first step in nitrification is performed by ammonia-oxidising microorganisms, which convert ammonia into nitrite ions. Ammonia-oxidising bacteria (AOB) have been known for more than 100 years. However, metagenomic studies and subsequent cultivation efforts have recently demonstrated that microorganisms of the domain archaea are also capable of performing this process. Astonishingly, members of this group of ammonia-oxidising archaea (AOA), which was overlooked for so long, are present in almost every environment on Earth and typically outnumber the known bacterial ammonia oxidisers by orders of magnitudes in common environments such as the marine plankton, soils, sediments and estuaries. Molecular studies indicate that AOA are amongst the most abundant organisms on this planet, adapted to the most common environments, but are also present in those considered extreme, such as hot springs. The ecological distribution and community dynamics of these archaea are currently the subject of intensive study by many research groups who are attempting to understand the physiological diversity and the ecosystem function of these

ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 57 ISSN: 0065-2911 DOI:10.1016/B978-0-12-381045-8.00001-1

Copyright Ó 2010 by Elsevier Ltd. All rights reserved

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CHRISTA SCHLEPER AND GRAEME W. NICOL

organisms. The cultivation of a single marine isolate and two enrichments from hot terrestrial environments has demonstrated a chemolithoautotrophic mode of growth. Both pure culture-based and environmental studies indicate that at least some AOA have a high substrate affinity for ammonia and are able to grow under extremely oligotrophic conditions. Information from the first available genomes of AOA indicate that their metabolism is fundamentally different from that of their bacterial counterparts, involving a highly copper-dependent system for ammonia oxidation and electron transport, as well as a novel carbon fixation pathway that has recently been discovered in hyperthermophilic archaea. A distinct set of informational processing genes of AOA indicates that they are members of a distinct and novel phylum within the archaea, the ‘Thaumarchaeota’, which may even be a more ancient lineage than the established Cren- and Euryarchaeota lineages, raising questions about the evolutionary origins of archaea and the origins of ammonia-oxidising metabolism.

1. 2. 3. 4. 5. 6.

7.

8.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Discovery of archaea in moderate aerobic habitats. . . . . . . . . . . . . . . . . . First insights into the physiology of ammonia-oxidising archaea . . . . . . . . Model organisms of ammonia-oxidising archaea. . . . . . . . . . . . . . . . . . . . Membrane lipids of ammonia-oxidising archaea . . . . . . . . . . . . . . . . . . . . Genomes and metagenomes of ammonia-oxidising archaea . . . . . . . . . . 6.1. Metagenomic Studies of Uncultivated Ammonia Oxidisers. . . . . . . . 6.2. Predictions from Complete Genome Sequences of Two Marine Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Ammonia Oxidisers: A Distinct Phylum within the Archaea . . . . . . . Diversity, distribution and activity of ammonia-oxidising archaea in the environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. AOA in the Soil Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. AOA Activity in the Soil Environment. . . . . . . . . . . . . . . . . . . . . . . . . 7.3. AOA in the Marine Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. AOA Activity in the Marine Environment . . . . . . . . . . . . . . . . . . . . . . 7.5. AOA in Sediments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6. AOA in Geothermal Environments . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7. AOA Associated with Marine Invertebrates. . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 3 4 6 9 13 15 15 16 19 22 22 23 25 28 29 30 32 33 33 34 34

AMMONIA-OXIDISING ARCHAEA

3

ABBREVIATIONS AOA AOB GDGT 16S rRNA

ammonia-oxidising archaea ammonia-oxidising bacteria glycerol dialkyl glycerol tetraether 16S ribosomal ribonucleic acid

1. INTRODUCTION In many ecosystems and in the biosphere as a whole, microorganisms are considered to constitute the largest component, in terms of both biomass and biological activity (Whitman et al., 1998). They are major players in regulating the biosphere through their participation in global biogeochemical cycles. Until recently, the role of archaea in these global cycles, as well as their phylogenetic and physiological diversity, had been largely underestimated. Only the distribution of methanogenic archaea in many different anaerobic habitats worldwide, as well as their role in the global carbon cycle, has been well described (Garcia et al., 2000). All other archaea were considered extremophiles, with specific adaptations allowing them to inhabit environments considered inhospitable for most other organisms, such as salt-saturated lakes, high-temperature terrestrial springs and deep-sea vents (Woese, 1987). However, with the help of culture-independent molecular techniques, involving the amplification of 16S rRNA genes directly from environmental samples, it has been shown over the past two decades that archaea are not confined to extreme habitats. By contrast, they have a ubiquitous distribution on this planet and occur in significant numbers in common environments such as soils, marine plankton and sediments as well as in the deep subsurface (DeLong, 1998; Schleper et al., 2005). However, since their initial discovery, it took over a decade before any aspect of the physiology or ecological role of moderate, aerobic archaea could be determined. Only very recently, metagenomic and cultivation studies have provided evidence that moderate archaea of terrestrial and marine environments are capable of ammonia oxidation and thus potentially represent important players in the global nitrogen cycle. Nitrification, the biological conversion of ammonia to nitrate via nitrite, is a central component of the natural nitrogen cycle (Prosser, 1989; Kowalchuk and Stephen, 2001). It is a two-step, aerobic, microbially mediated process, with ammonia first oxidised to nitrite by ammonia-oxidising microorganisms,

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CHRISTA SCHLEPER AND GRAEME W. NICOL

and nitrite subsequently oxidised to nitrate by nitrite-oxidising microorganisms. The first step is considered to be rate limiting for this process (Kowalchuk and Stephen, 2001). Nitrification ensures the conversion of ammonia (derived from organic nitrogen during decomposition and mineralisation of biomass) into the oxidised and more soluble form of nitrate, and provides the substrate for denitrification, which returns nitrogen back to the atmosphere. Nitrate is the preferred substrate of plants and aerobic microorganisms. However, the consequences of nitrification for human activities are also considerable. The process is used in wastewater treatment plants to remove urea and ammonia from sewage. In agricultural soils, the oxidation of ammonia to nitrate increases the availability of nitrogen for plants, but it also has negative consequences, because this results in loss of vast amounts of nitrogen fertiliser from agricultural land by leaching of the more soluble nitrate, resulting in groundwater pollution. Ammonia oxidisers have a further substantial environmental impact as contributors to greenhouse gas emissions via both ammonia oxidation directly as well as nitrifier-denitrification mechanisms (Wrage et al., 2001). Since the recognition of ammonia and nitrite-oxidising bacteria by Percy Faraday Frankland and Sergei Winogradsky and others over 100 years ago, only proteobacteria of the beta- and gamma-subdivisions were considered as capable of performing aerobic ammonia oxidation (Purkhold et al., 2000). Here, we summarise the current knowledge of the recently discovered ammonia-oxidising archaea (AOA), their physiology, genomic potential and distribution and activity in various environments.

2. DISCOVERY OF ARCHAEA IN MODERATE AEROBIC HABITATS Archaea in moderate aerobic habitats were first recognised by Fuhrman et al., 1992Fuhrman and colleagues (1992) and DeLong (1992), based on 16S rRNA gene surveys of marine environments. They were initially grouped into three previously undetected lineages that were termed Group I, affiliated to the kingdom of Crenarchaeota, and Groups II and III within the kingdom Euryarchaeota (DeLong, 1992) (Fig. 1a). In particular, organisms within Group I (a lineage distinct from, but specifically associated with, cultured hyperthermophilic organisms) were found in many moderate habitats including soils (Bintrim et al., 1997; Buckley et al., 1998; Sandaa et al., 1999; Jurgens et al., 2000; Simon et al., 2000; Ochsenreiter et al., 2003), the ocean’s plankton (DeLong, 1992; Fuhrman et al., 1992), estuaries (Crump and Baross, 2000),

AMMONIA-OXIDISING ARCHAEA

[(Figure_1)TD$IG]

Figure 1 (a) Phylogenetic relationship between various archaeal 16S rRNA gene-defined lineages, including sequences from cultivated organisms and environmental samples. Groups 1, 2 and 3 represent lineages which were originally discovered in planktonic marine habitats, with Group 1 sequences now recovered in nearly all terrestrial and aquatic habitats. (b) Phylogenetic relationships of Group I archaea. Black triangles represent groups in which amoA genes have been discovered (adapted from Prosser and Nicol, 2008).

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marine and freshwater sediments (MacGregor et al., 1997; Schleper et al., 1997; Vetriani et al., 1998; Keough et al., 2003), but also in the deep subsurface (Takai et al., 2001). Extensive surveys of 16S rRNA gene sequences from marine and soil samples revealed that Group I Crenarchaeota can be separated into a number of distinct clades, with the majority of soil and marine sequences placed within two of these, referred to as Group 1.1a and 1.1b lineages, respectively (Fig. 1b). While aspects of the physiology and energy metabolism of these organisms remained unknown for a long time, some initial indirect insights into the carbon metabolism of marine archaea were obtained using stable isotope, microautoradiography or natural radiocarbon analyses. They indicated that both modes of carbon assimilation occurred within marine archaea, that is autotrophy (using inorganic carbon as a nutrient source) (e.g. Kuypers et al., 2001; Pearson et al., 2001; Wuchter et al., 2003) and heterotrophy (using organic carbon compounds as nutrients) (Ouverney and Fuhrman, 2000; Herndl et al., 2005; Ingalls et al., 2006; Teira et al., 2006).

3. FIRST INSIGHTS INTO THE PHYSIOLOGY OF AMMONIA-OXIDISING ARCHAEA The first insight into a specific energy metabolism of Group I archaea stemmed from a fosmid clone derived from a soil metagenomic library (Treusch and Schleper, 2004; Treusch et al., 2004a,b, 2005). It contained an insert of about 43 kb. Based on 16S and 23S rRNA genes, clone ‘54d9’ was identified as belonging to the Group 1.1b lineage (Fig. 2, deposited in GenBank, February 2004). In addition, it contained homologues to bacterial genes involved in nitrogen cycling. Specifically, it contained two open reading frames (ORFs) coding for putative alpha and beta subunits (AmoA and AmoB, respectively) of an ammonia monooxygenase (AMO) as well as a gene whose product was highly similar to copperdependent nitrite reductases (NirK) (Treusch et al., 2005). An in silico comparison to environmental sequences deposited in public databases showed that the soil-derived archaeal amoA and amoB genes were highly similar to archaea-associated scaffolds from the whole-genome shotgun (WGS) sequencing project of the Sargasso Sea (Venter et al., 2004; Schleper et al., 2005). Additionally, the genomic fragments from marine archaea assembled in the Sargasso Sea project contained genes coding for the C-subunit of an AMO, apparently organised in a cluster together with amoA and amoB in a BCA gene order and contrasted with the CAB arrangement found in ammonia-oxidising bacteria (AOB) (Nicol and

AMMONIA-OXIDISING ARCHAEA

[(Figure_2)TD$IG]

Figure 2 Schematic diagram showing predicted ORFs on the 43 kb soil fosmid 54d9, some of which are potentially involved in nitrogen transformations (amoA and amoB: genes for potential subunits of ammonia monooxygenase; nirK for nitrite reductase gene, ORF38 conserved in amo clusters) (Treusch et al., 2005). Homologues present in scaffolds assembled from the Sargasso Sea sequencing project (Venter et al., 2004) presented underneath. (Adapted from Schleper et al. (2005), with permission.)

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[(Figure_3)TD$IG]

Figure 3 Maximum-likelihood tree of derived amino acid sequences showing the phylogenetic relationship of ammonia monooxygenases and particulate methane monooxygenases (AMO and pMMO respectively) of bacteria and archaea. The phylogenetic tree is based on 156 unambiguously aligned positions.

Schleper, 2006). Sequence comparison of the soil- and marine-derived archaeal AmoA sequences to the alpha subunits of the bacterial AMO and the particulate methane monooxygenase (pMMO) from bacterial methane oxidisers indicated that they were quite distinct (Fig. 3), with only about 40% similarity (25% identity) at the amino acid level. In contrast, the similarity between the two related proteins in AMO and pMMO in bacteria is much higher with up to 74% (50% identity). Furthermore, the putative amo/pmo genes of archaea were considerably shorter than those of their bacterial homologues. A comparison with structural data obtained of the pMMO of Methylococcus capsulatus (Lieberman and Rosenzweig, 2005) brought further evidence that the respective archaeal ORFs indeed coded for subunits of an AMO/pMMO-related protein, as many amino acid

AMMONIA-OXIDISING ARCHAEA

9

[(Figure_4)TD$IG]

Figure 4 Aerobic ammonia oxidation during nitrification. The oxidation of ammonia is performed by ammonia oxidisers (archaea and bacteria), and the nitrite produced is subsequently oxidised by nitrite-oxidising bacteria. In bacteria, ammonia is oxidised to nitrite via the intermediate hydroxylamine and the enzyme hydroxylamine oxidoreductase (HAO). No HAO homologue has yet been identified in archaea, and oxidation of ammonia to nitrite may occur via a different biochemical pathway (see hypotheses Fig. 7).

residues potentially involved in copper binding metal centres were also found to be conserved in the archaeal variants (Treusch et al., 2005). Moreover, microcosm experiments with soil slurries were conducted to study transcription of the archaeal amoA genes. Upon incubation with NH4+ a significant increase in transcriptional activity of the putative amoA gene was observed, suggesting that the amo-like genes indeed coded for a monooxygenase involved in the oxidation of ammonia (Treusch et al., 2005). The ultimate support for this hypothesis came from the cultivation of an autotrophic AOA, Nitrosopumilus maritimus, that contains amoA genes highly related €nneke et al., 2005). These to those found on the fosmid clone 54d9 (Ko findings indicated that archaea could be involved in ammonia oxidation (Fig. 4) in many terrestrial and marine environments because 16S rRNA and amoA gene sequences related to those of N. maritimus and 54d9 were found in many habitats around the planet.

4. MODEL ORGANISMS OF AMMONIA-OXIDISING ARCHAEA Chemolithoautotrophic ammonia oxidisers, bacteria or archaea, are notoriously difficult to grow and maintain in (pure) laboratory cultures. This is also illustrated by the fact that strain collections of AOB are only kept in a few specialised laboratories (J. Prosser, University of Aberdeen, pers. comm.). The isolation of the first AOA, Candidatus N. maritimus [nitrosus (latin):

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CHRISTA SCHLEPER AND GRAEME W. NICOL

nitrous; pumilus (Latin): dwarf; maritimus (Latin): of the sea] (K€ onneke et al., 2005) has therefore represented a breakthrough for the characterisation of AOA, providing the first insights into the general physiology and specific metabolism of these organisms. N. maritimus strain SCM1 was isolated from a tropical marine aquarium and grows with a near-stoichiometric conversion of ammonia into nitrite under aerobic conditions. It grows to a maximum density of about 1.4  107 cells mL 1 at 28  C in defined mineral medium supplemented with bicarbonate and 500 mM ammonium (K€ onneke et al., 2005), and its growth is inhibited when organic compounds are added even in low amounts. N. maritimus is a straight, relatively small rod with a diameter of 0.17–0.22 mm and a length of 0.5–0.9 mm (Fig. 5a). Based on 16S rRNA gene phylogeny, N. maritimus, like the earlier described sponge symbiont Cenarchaeum symbiosum (Preston et al., 1996), belongs to the Group 1.1a lineage of marine archaea that is found in large numbers in the open ocean. Using the sequence information from metagenomic studies, K€ onneke et al. (2005) designed primers against amoA, amoB and amoC sequences and were able to amplify the respective homologous genes from N. maritimus. This analysis provided the direct link between the metagenomic predictions and physiological studies. In contrast to AOB, N. maritimus is apparently adapted to an extremely oligotrophic lifestyle (Martens-Habbena et al., 2009). It possesses a half-saturation constant (Km = 133 nM total ammonium) and a substrate threshold (<10 nM) closely resembling ammonia oxidation kinetics in the open ocean, indicating that marine AOA can thrive under extreme nutrient limitation as typically found in the open ocean.

[(Figure_5)TD$IG]

Figure 5 Electron micrograph images of ammonia-oxidising archaea. (a) Au/Pdsputtered cells of Nitrosopumilus maritimus, length of cells approximately 0.5–0.9 mm (K€ onneke et al., 2005, with permission). (b) Fluorescent in situ hybridisation of C. symbiosum in a cell preparation obtained from tissue of the sponge Axinella mexicana, bar represents 2 mm (kindly provided by Chris Preston). (c) Uranyl acetate contrasted, electron microscopic picture of Nitrososphaera gargensis (kindly provided by Eva Spieck). (d) Scanning electron microscopic picture of strain EN76 cultivated from Vienna soil, diameter of cells between 0.3 and 0.8 mm (M. Tourna, C. Schleper, unpublished).

AMMONIA-OXIDISING ARCHAEA

11

Two enrichments of thermophilic AOA have further demonstrated the capability of archaea to grow autotrophically with ammonia as a sole energy source. In an enrichment of ammonia-oxidising microorganisms from a microbial mat from the Siberian Garga hot spring (Lebedeva et al., 2005), cells of Candidatus Nitrososphaera gargensis [nitrosus (Latin): nitrous; sphaera (Latin): spherical; gargensis: from Garga spring] (Hatzenpichler et al., 2008) were visualised using CARD-FISH and simultaneous incorporation of radioactive labelled bicarbonate indicating ammonia-dependent autotrophic growth at concentrations of 0.14 and 0.79 mM NH4+. However, at higher concentrations, around 3 mM ammonium, or at lower concentrations but in the presence of allylthiourea, a potent inhibitor of ammonia oxidation, this incorporation was considerably reduced. Interestingly, N. gargensis belongs to the lineage of AOA distinct from marine AOA, the Group 1.1b lineage, which is mostly represented by environmental sequences from soils and other terrestrial habitats (Ochsenreiter et al., 2003; Nicol et al., 2006). It is therefore the first representative of this second major lineage which has been obtained in laboratory enrichments. The thermophilic AOA Candidatus Nitrosocaldus yellowstonii [nitrosus (Latin): nitrous; caldus (Latin): hot; yellowstonii: from Yellowstone] (de la Torre et al., 2008) obtained from a hot spring located in the Yellowstone National Park has an even higher optimal growth temperature between 65 and 72  C, again growing with a stoichiometric conversion of ammonia to nitrite. This organism was particularly interesting as it not only extended the temperature limit for cultivated ammonia oxidisers, but broadened the known 16S rRNA and amoA gene diversity associated with ammonia oxidation (Table 1, Fig. 3). C. symbiosum [cen from Greek kainos/koinos: recent/common refers to the recent (derived from thermophiles) and widespread (common) occurrence of this group of archaea; (Preston et al., 1996)] was detected by 16S rRNA-based PCR surveys among the complex microbial communities within the tissues of the marine sponge Axinella mexicana (Preston et al., 1996). Since it can be ‘quasi’-cultivated in the laboratory by maintaining it in stable association with its host under controlled conditions, it was the first described mesophilic crenarchaeon. The Cenarchaeum/Axinella association provided the first tractable system for the study of marine archaea and gave access to relatively large amounts of biomass of this species, allowing the study of lipids, genomic DNA and cell structure by microscopy. As it is maintained within the tissues of the sponge and cannot be cultivated independently, ammonia oxidation by C. symbiosum has not been proven directly. However, the genome sequence of this organism, the presence of archaeal amoA genes and measured ammonia oxidation activity in other

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Table 1 Ammonia-oxidising archaea in laboratory cultures or enrichments. Nitrosopumilus maritimus

Cenarchaeum symbiosum

Nitrosocaldus yellowstonii

Nitrososphaera gargensis

Origin

Tropical marine aquarium Pure laboratory culture Group 1.1a Thaumarchaeota 28  C

Marine sponge symbiont

Terrestrial warm spring

Inside Axinella mexicana (aquarium) Group 1.1a Thaumarchaeota 10  C (8–18  C)

Terrestrial hot spring Laboratory enrichment HWCGIII

Rod 1.65 Mb K€ onneke et al. (2005)

Curved rod 2.05 Mb Preston et al. (1996)

Culture Affiliation Growth temperature Shape Genome Reference

72  C (65–74  C) Spherical n.d. de la Torre et al. (2008)

Laboratory enrichment Group 1.1b Thaumarchaeota 46  C Spherical, coccoid >2.6 Mb, draft Hatzenpichler et al. (2008), Spang et al. (2010)

CHRISTA SCHLEPER AND GRAEME W. NICOL

Characteristics

AMMONIA-OXIDISING ARCHAEA

13

marine sponges (Taylor et al., 2007; Bayer et al., 2008; Steger et al., 2008; Hoffmann et al., 2009) strongly support the hypothesis that C. symbiosum, like its close free-living relatives in marine water, is capable of ammonia oxidation.

5. MEMBRANE LIPIDS OF AMMONIA-OXIDISING ARCHAEA Since their discovery as a separate domain, distinct from the eukaryotes (or Eukarya) and bacteria based on 16S rRNA gene phylogenies (Woese and Fox, 1977), the archaea have been found to possess a number of cellular and molecular features that clearly distinguishes them from the other two domains. The best distinguishing feature is the membrane lipids of archaea, which are fundamentally different from all representatives in the other two domains (Brown and Doolittle, 1997). The phospholipids of archaea are glycerol–ether lipids in which isoprenoid side chains, not fatty acids like in bacteria or Eukarya, are linked via an ether bond (instead of an ester bond) to a glycerol moiety, and have a stereochemistry that is reverse of that in bacteria and Eukarya. Furthermore, the C20-isoprenoid side chains are often linked to each other, which leads to the formation of a lipid monolayer with C40 side chains, instead of the typical membrane bilayer found in other organisms. Thus, the core, apolar component of archaeal cellular membrane lipids (in particular those of Crenarchaeota) are dominated by glycerol dialkyl glycerol tetraethers (GDGTs, see Fig. 6). The side chains may contain multiple cyclopentane moieties (see Fig. 5), whose numbers have been shown to increase in response to an increase in growth temperature of Archaeoglobus species (Lai et al., 2008). Interestingly, a specific structure, a unique GDGT with a cyclohexane moiety in addition to the four cyclopentane rings (Fig. 5, bottom), was found in natural samples from moderate environments as well as in the marine sponge A. mexicana, which harbours C. symbiosum, a member of the marine Group 1.1a (Damste et al., 2002). It was thus termed ‘crenarchaeol’, although this lipid compound had never been found in cultivated hyperthermophilic Crenarchaeota and seems to be present in ‘mesophilic’ archaea only. In support of this, crenarchaeol and the other GDGT compounds have also been isolated from the laboratory cultures of N. maritimus (Schouten et al., 2008). In addition, with the finding that the moderate thermophile N. gargensis (Pitcher et al., 2009) and the extremely thermophilic N. yellowstonii (de la Torre et al., 2008) also contain crenarchaeol, and with the recovery of crenarchaeol from various hot springs around the world (Pearson et al., 2004; Zhang et al., 2006;

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CHRISTA SCHLEPER AND GRAEME W. NICOL

[(Figure_6)TD$IG]

Figure 6 (a) Isoprenoidal glycerol dialkyl glycerol tetraethers (GDGTs) of archaea, including crenarchaeol that is so far exclusively found in cultured Thaumarchaeota and in many natural environments. (b) Correlation between GDGT abundance and AmoA gene copies (eight different soils types). (Adapted from data obtained by Leininger et al. (2006), with permission.)

Reigstad et al., 2008; Pitcher et al., 2009), it is now evident that this compound is not an indicator for mesophilic archaea per se, but rather for certain lineages of archaea to which ammonia oxidisers are affiliated (Pitcher et al., 2009). At least in some environments, crenarchaeol could be a suitable biomarker for AOA (Wuchter et al., 2004; Coolen et al., 2007; Schouten et al., 2007). For example, in 8 out of 12 investigated soils, the abundance of crenarchaeol (as well as the abundance of total GDGTs) correlated significantly with the abundance of amoA genes (Leininger et al., 2006). Only the analysis of more isolates of AOA and of other more remotely related lineages of archaea will help to clarify whether this lipid compound is restricted to ammonia oxidisers. It is interesting to note in this context that the relative abundance of crenarchaeol in the different laboratory cultures of AOA seems to vary quite considerably. The GDGTs of N. gargensis (grown at 46  C) consisted mainly of crenarchaeol, its regioisomer and a novel GDGT, while crenarchaeol was a minor compound in the other organisms. Varying relative amounts of the crenarchaeol or its regioisomer in different lineages of AOA could potentially confuse the TEX86 index that is used for paleothermometry, that is for reconstructing average temperatures experienced by plankton in earlier

AMMONIA-OXIDISING ARCHAEA

15

times by measuring the relative composition of sedimentary archaeal membrane lipids (Wuchter et al., 2004).

6. GENOMES AND METAGENOMES OF AMMONIA-OXIDISING ARCHAEA Inspired by the rapid advances in genomic techniques applied to cultivated microorganisms, Stein et al. (1996) used a BAC-derived fosmid vector to prepare a large-insert library from marine water of the North-Eastern Pacific in order to characterise marine planktonic archaea. A 38.5-kb genomic fragment of an uncultivated mesophilic Crenarchaeota was identified within 3552 clones using archaea-specific 16S rRNA gene probes. This study was the beginning of a novel and now exploding field of microbial environmental genomics or ‘metagenomics’. Large genome fragments are cloned into bacterial artificial chromosomes (BACs) or, more commonly, BAC-derived fosmid vectors (a hybrid of a cosmid and BAC) which are archived in Escherichia coli clone libraries (Handelsman, 2004; Treusch and Schleper, 2005). Alternatively, large-scale sequencing using a whole-genome-shotgun approach allows the generation of small sequence reads from environmental samples that can be assembled into larger fragments in silico (Venter et al., 2004). These techniques are now increasingly used for the characterisation of microbial communities, particularly since novel deep sequencing technologies allow increasingly cheaper and high-throughput analysis. The first complete genome of a potential AOA (C. symbiosum) was also assembled from a metagenomic library, and now with cultivated or enriched organisms available, complete reference genomes of the first model organisms are easily obtained. While the complete genome sequences will be invaluable for the reconstruction of full metabolic pathways, the metagenomic datasets of AOA are of great importance to study the distribution and the genomic potential of the organisms in the various environments.

6.1. Metagenomic Studies of Uncultivated Ammonia Oxidisers Several metagenomic libraries from microorganisms associated with a marine sponge, marine plankton or soil have been produced to characterise the genomic content of Group I archaea (Schleper et al., 1998; Beja et al., 2000, 2002; Quaiser et al., 2002; López-Garcıa et al., 2004; Treusch et al., 2004). Soil fosmid 54d9 was detected in such a library and led to the discovery of amo-related genes in archaea (Treusch et al., 2005). Comparative analyses

16

CHRISTA SCHLEPER AND GRAEME W. NICOL

of metagenomic clones from the marine plankton indicated the conservation of gene order around the 16S rRNA gene, thus confirming the close relationship of the planktonic archaea, even in strains from different oceanic provinces (Bej a et al., 2002; López-Garcıa et al., 2004). Conversely, considerable genomic variation was dissected, including microheterogeneity, in protein-encoding regions and intergenic spacers, when genome fragments with otherwise identical or almost identical 16S rRNA genes were compared from the same DNA library (Schleper et al., 1998; Beja et al., 2002). Given the abundance and ubiquity of marine planktonic archaea, it is plausible that large numbers of archaeal genes would be detected in global random sequencing surveys of DNA obtained from filtered waters (Venter et al., 2004; Nealson and Venter, 2007). The huge databases from BAC and fosmid libraries (Treusch et al., 2004; Martin-Cuadrado et al., 2008; Konstantinidis et al., 2009) as well as large-scale shotgun sequencing efforts are a valuable resource for dissecting the diversity and distribution of AOA, particularly since the first genomes of cultivated isolates are now available that allow us to test hypotheses on gene functions and metabolisms experimentally.

6.2. Predictions from Complete Genome Sequences of Two Marine Archaea Although C. symbiosum has not been cultivated or completely physically separated from the tissues of its host (the marine sponge A. mexicana) and from the co-existing bacteria, cell fractions that were enriched with the archaeon were used for the construction of large-insert genomic libraries (Schleper et al., 1997, 1998), facilitating the isolation of genome fragments and leading to a genome sequencing project that was completed in 2006 (Hallam et al., 2006a, 2006bHallam et al., 2006a,b). The second genome from N. maritimus, the first cultivated isolate of marine AOA, has recently been completed (Walker et al., 2010). The C. symbiosum genome has a considerably higher G + C content (>55%) (Schleper et al., 1998; Hallam et al., 2006a) than its relatives in the marine plankton (approximately 34%), which might reflect adaptation to the lifestyle in the metazoan host, rather than a large evolutionary distance. Despite this difference, however, the two organisms show high overlap in gene content with each other (approx. 1200 genes) and with marine metagenomes, indicating that they can serve as suitable models to study the globally distributed and abundant marine planktonic archaea (Walker et al., 2010). With a few minor exceptions, both genomes share similar gene content with respect to potential energy metabolism and carbon fixation

AMMONIA-OXIDISING ARCHAEA

17

pathways, both of which seem to be clearly different from known bacterial ammonia oxidisers. AOA contain amoA, B and C genes for the three subunits of a potential archaeal AMO, but no homologous genes of the bacterial hydroxylamine oxidoreductase (HAO) complex that catalyses the second step of this process in bacteria, that is the oxidation of hydroxylamine to nitrite (Hallam et al., 2006b; Walker et al., 2010). In AOB, this complex delivers electrons back to the AMO and to an electron transport chain with cytochrome c proteins (c554 and c552) required for electron flow to ubiquinone (Fig. 6). These cytochromes are also not present in AOA, but they do possess numerous copper-containing proteins such as multicopper oxidases, small blue copper-containing proteins (Hallam et al., 2006a; Bartossek et al., 2010; Walker et al., 2010) as well as potential thiol-disulphide oxidoreductases, which may functionally replace cytochromes (Walker et al., 2010). In principle, it appears that the energy metabolism of AOA relies on copperrather than on iron-containing electron transfer systems (Hallam et al., 2006a; Walker et al., 2010). While C. symbiosum (like marine metagenomes) contains genes for urease, indicating a potential broader substrate spectrum, N. maritimus does not. Both organisms also do not seem to contain a homologue of nitric oxide reductase, that is part of the denitrification pathway of AOB, reducing nitric oxide (NO) to nitrous oxide (N2O). N. maritimus (Walker et al., 2010), as well as soil archaea (Treusch et al., 2004a; Bartossek et al., 2010), does, however, contain homologues of nitrite reductase, the first enzyme involved in dentrification, that (in bacteria) reduces nitrite, the product of ammonia oxidation, to nitric oxide. In total, the genetic makeup of archaeal ammonia oxidisers indicates that the biochemistry underlying ammonia oxidation to nitrite could be fundamentally different from that of AOB. Even the key metabolic enzyme, AMO, shows only little sequence similarity to bacterial AMO and pMMOs. It is possible that AOA use the same pathway as AOB, that is oxidising ammonia via hydroxylamine to nitrite, but with different enzymes. Alternatively, a fundamentally different pathway may be operating. Martin Klotz (University of Louisville, Kentucky, USA) has proposed an alternative pathway for AOA that does not involve hydroxylamine as an intermediate, but rather nitroxyl (HNO) (see Walker et al., 2010) (Fig. 7). A nitroxyl oxidoreductase could then operate to oxidise nitroxyl to nitrite (Walker et al., 2010). In his extended model, Klotz proposes that the activation of AMO could be achieved by recycling NO, the product of nitrite reduction via nitrite reductase (see Fig. 7). The activation of O2 by NO would then result in the production of N2. If this or a similar pathway for ammonia oxidation is indeed operating, it would indicate most probably that AOA do not produce the greenhouse gas N2O, as do their bacterial counterparts. It is

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CHRISTA SCHLEPER AND GRAEME W. NICOL

[(Figure_7)TD$IG]

Figure 7 Proposed pathways of nitrogen, oxygen and electron flow in the quinone-oxidising and -reducing branches of the electron transport chains (ETC) in ammonia-oxidising bacteria (AOB) and archaea (AOA). (a) Basic inventory encoded in all AOB. Ammonia monooxygenase (AMO); N-oxide-Ubiquinone Redox Module (NURM) consisting of hydroxylamine oxidoreductase (HAO), cytochrome c554 and quinone reductase cM552; cytochrome c552; cytochromes b and c1 (complex III), with several proton-pumping oxygen-reducing heme-copper oxidases (complex IV) and several NO-reducing heme-copper oxidases (HCO-NOR) in the membrane-associated branch, and copper-containing nitrite reductase (NirK) and cytochrome c NOreductases in the soluble branch of the ETC, plus bacterial F0F1-type ATP synthetase. The question mark indicates that a direct quinol oxidase function of bacterial AMO/ pMMO has not yet been demonstrated. (b) Basic inventory predicted from genome sequences in AOA. AMO; NURM consisting of nitroxyl oxidoreductase (NxOR) and plastocyanin-domain containing membrane-associated putative quinone reductase (Pcy); plastocyanins (pcy); cytochrome b and associated plastocyanin (complex III) with several proton-pumping oxygen-reducing plastocyanin-copper oxidases (complex IV) plus F0F1-type ATP synthetase in the membrane-associated branch. Copper-containing nitrite reductase (NirK), active in the soluble branch of the ETC, has been identified in all AOA genomes with the exception of Cenarchaeum symbiosum (Bartossek et al., 2010). No inventory encoding cytochrome c and hemecopper NO-reductases has been identified. The asterisks indicate the need for activation of oxygen in the monooxygenase reaction facilitated by AMO. Because a quinol oxidase function of archaeal AMO is not part of the model, it is predicted that oxygen activation in AOA is accomplished by utilising the NO radical produced in the soluble branch of the ETC. This would produce one-half molecule of dinitrogen gas per oxidised ammonia and introduce AOA as non-classical aerobic denitrifiers. This chemistry needs to be experimentally tested as indicated by ‘??’ (Figure and legend kindly provided by Prof. Martin G. Klotz, University of Louisville, USA).

AMMONIA-OXIDISING ARCHAEA

19

interesting to note in this context that Bartossek et al. (2010) recently found variants of genes encoding for copper-dependent nitrite reductases in soils and other environments, indicating that this enzyme might indeed be widely distributed in AOA. Furthermore, transcription of the nitrite reductase homologue in soil archaea was observed even under aerobic (and potentially ammonia-oxidising) conditions, rather than under low-oxygen conditions that favour denitrification (Bartossek et al., 2010). Thus, the copperdependent nitrite reductase of archaea might indeed fulfill a different function in AOA metabolism than is assumed for the bacterial counterpart. From the genome sequences of the two marine organisms one can also deduce a possible pathway for carbon fixation. Whereas autotrophic AOB fix carbon with the ribulose bisphosphate carboxylase/oxygenase (RubisCO) in the Calvin–Bassham–Benson cycle, N. maritimus and C. symbiosum seem to contain a different pathway, similar to that recently described for the hyperthermophilic archaeon Metallosphaera sedula (Berg et al., 2007). The pathway involves the transformation of acetyl-CoA with two bicarbonate molecules via 3-hydroxyproprionate to succinyl-CoA. This intermediate is reduced to 4-hydroxybutyrate and subsequently converted into two acetylCoA molecules. Key enzymes for this pathway are found in both AOA organisms, such as the biotinylated acetyl-CoA/propionyl-CoA carboxylase, methylmalonyl-CoA epimerase and mutase and 4-hydroxybutyrate dehydratase, but some proteins of the M. sedula pathway are also missing, indicating the AOA use a variant of this pathway or may possess nonorthologous gene replacements (Hallam et al., 2006b; Walker et al., 2010). While autotrophic growth of N. maritimus and its inhibition by organic compounds has been reported, both AOA contain components of an oxidative TCA cycle that can potentially be utilised for the consumption of organic carbon or for the production of intermediates for amino acid and cofactor biosynthesis. Mixotrophic growth has not been shown yet for any of the cultivated or enriched AOA, but it might well be possible that this growth mode will be found as more organisms are obtained in laboratory cultures.

6.3. Ammonia Oxidisers: A Distinct Phylum within the Archaea Based on 16S rRNA sequence phylogeny, AOA were originally placed as a sister group of the Crenarchaeota (DeLong, 1992; Fuhrman et al., 1992), suggesting that these archaea might have ancestors in hot springs and only later radiated into moderate environments. The AOA have since been referred to as Crenarchaeota in all following 16S rRNA-based diversity

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CHRISTA SCHLEPER AND GRAEME W. NICOL

studies. Using a concatenated dataset of 53 ribosomal proteins from C. symbiosum which are shared by archaea and eukaryotes, BrochierArmanet et al. (2008) calculated that ‘moderate Crenarchaeota’ constitute a separate phylum of the archaea that branches off before the separation of Crenarchaeota and Euryarchaeota. Including the genomic information of N. maritimus and N. gargensis (draft genome) in this analysis confirmed the separation of AOA (Spang et al., 2010). The name Thaumarchaeota (from the Greek word ‘thaumas’ for wonder) was proposed for the new phylum (Brochier-Armanet et al., 2008). Several information processing genes, whose presence or absence is characteristic for Euryarchaeota and/or Crenarchaeota, show a pattern in the three investigated Thaumarchaeota genomes that is distinctive from either of the two described phyla and this points to fundamental differences in cellular processes. Thus, this gene content comparison strongly supports the Thaumarchaeota proposal (Table 2 and Brochier-Armanet et al., 2008; Spang et al., 2010). Most notably, Thaumarchaeota possess unsplit RNA

Table 2 Distribution of core informational processing genes in the four different phyla of archaea. Thaumarchaeota Ribosomal proteins r-protein S25e + r-protein S26e + r-protein S30e + r-protein L13e r-protein L14e r-protein L34e r-proteins L38e r-protein L29p + r-protein Lxa Transcription/RNA polymerase rpoG (=rpo8) rpoA – single ORF + rpoB – single ORF + Rpc34 + MBF 1 W EIF 1

Euryarchaeota

+ (some) + (some) + + (most) split W split W +

Crenarchaeota

Korarchaeota

+ + + + + + W + +

+ + + + + +

+ split + + + +

+ + + + + (continued)

AMMONIA-OXIDISING ARCHAEA Table 2

21

(continued) Thaumarchaeota

DNA polymerases/replication DNA pol D + RPA (Eury-like) + RPA/SSB + >one PCNA gene Topoisomerases Topo IB + Topo IA W Reverse gyrase Topo IIA Cell division ESCRT-III + Vps4 (CdvC) + CdvA + Fts Z + Smc + ScpA + ScpB + Histones (H3/H4) + Repair Hef-nuclease ERCC4-type nuclease + ERRC4-type helicase + RadB HSP70s, GrpE, Hsp40 + UvrABC +

Euryarchaeota

+ + + (some)

+ + (HT) W

Crenarchaeota

+/( ) + + +

Korarchaeota

+ + +

+ +

+ + + + + + (many)

+ + + (exc. 2)

+

+

+ +

+

+ + W

polymerase A and B genes, both topoisomerases IA and B, and histones, but lack the archaea-specific r-protein LXa. Thaumarchaeota, like all other archaea, contain genes for central information processing machinery (replication, transcription, etc.) that are shared with eukaryotes or are more closely related to eukaryotes than to bacteria (Bell and Jackson, 1998; Garrett and Klenk, 2007). In several phylogenetic analyses with information processing factors, such as RNA polymerase subunits (see Fig. 8), Thaumarchaeota branch off early, indicating that a more detailed characterisation of this previously enigmatic group might change our perception of the early evolution within the archaeal and eukaryotic lineage (Spang et al., 2010).

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[(Figure_8)TD$IG]

Figure 8 Phylogenetic tree of RNA polymerase subunit A (rpoA) and schematic overview of gene arrangements in different archaeal lineages and in bacteria and eukaryotes. Like the latter two, Thaumarchaeota have an unsplit rpoA gene. In line with this finding, they form a separate and deeply branching lineage within the archaea in the phylogenetic analysis of the corresponding protein RpoA. (Modified from Spang et al. (2010), with permission.)

7. DIVERSITY, DISTRIBUTION AND ACTIVITY OF AMMONIAOXIDISING ARCHAEA IN THE ENVIRONMENT Within the archaeal domain, the ability to oxidise ammonia appears thus far to be restricted to the Group 1 lineage. After the initial identification of AOA amo genes in soil and marine environments (Venter et al., 2004; Treusch et al., 2005), oligonucleotide primers were designed to target conserved regions of this gene and used in PCR, cloning and sequencing approaches to examine the diversity and abundance of these organisms in DNA extracted from environmental samples. It became clear that these organisms are globally distributed in most, if not all, environments (Fig. 9), similar to the findings for 16S rRNA gene surveys (Fig. 1). Current efforts are to determine the fundamental aspects of their ecophysiology and their ecological importance to nitrification processes globally by linking environmental factors to AOA and AOB population dynamics.

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[(Figure_9)TD$IG]

Figure 9 Phylogenetic tree describing major ammonia-oxidising archaeal amoA gene-defined lineages and environments of origin. Analyses were performed on an alignment of 486 positions from 188 sequences representative of known amoA diversity and the shape of triangular blocks are proportional to the number of sequences and maximum branch lengths within. Multifurcating branches indicate where the relative branching order of major lineages could not be determined in the majority of bootstrap replicates using various treeing methods (distance, parsimony and maximum likelihood analyses). Lineages with cultivated representatives are highlighted together with soil fosmid clone 54d9.

7.1. AOA in the Soil Environment Thaumarchaeota are the dominant archaea in most soil systems where they constitute up to 5% of all prokaryotes (e.g. Ochsenreiter et al., 2003; Lehtovirta et al., 2009). There are a number of thaumarchaeal lineages found in high numbers in soil, and Group 1.1b (Fig. 1) is the dominant lineage in

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most soil systems (Auguet et al., 2010). This lineage includes the archaeon from which soil fosmid 54d9 was derived, therefore indicating that the most abundant archaea in soil may be capable of ammonia oxidation. Using quantitative PCR of amoA genes, Leininger et al. (2006) examined the abundance of AOA and AOB in 12 surface soils sampled across Europe. These soils represented a wide range of physicochemical properties and land-use types. Without exception, AOA represented the dominant group, with the ratio of AOA to AOB amoA genes ranging from 1.5 to over 230 (Fig. 10). This observation was confirmed for most soils examined so far (e.g. He et al., 2007; Nicol et al., 2008; Jia and Conrad, 2009; Schauss et al., 2009; Di et al., 2010), although there are exceptions (Boyle-Yarwood et al., 2008). Another general trend observed is that the relative abundance of AOA to AOB changes with soil depth, with AOA numbers remaining relatively constant but AOB numbers decreasing dramatically (Leininger et al., 2006; Jia and Conrad, 2009; Di et al., 2010), indicating that AOA may be

[(Figure_0)TD$IG]

Figure 10 AOA and AOB amoA gene abundance in 12 soils representing a wide geographical distribution and contrasting physicochemical properties. Gene abundances (copies per dry weight soil gram) were calculated using two specific quantitative PCR assays. (From Leininger et al. (2006), with permission.)

AMMONIA-OXIDISING ARCHAEA

25

particularly well adapted to conditions with low levels of available nutrients and oxygen.

7.2. AOA Activity in the Soil Environment With the co-occurrence of AOA and AOB in the soil environment, a major focus has been to determine the relative activities of both groups and what conditions determine their growth. Schauss and colleagues (2009) were the first to demonstrate growth of AOA in response to (organic) fertiliser additions. They demonstrated differences in the growth characteristics of AOA and AOB, but perhaps also provided some evidence for potential functional redundancy, with AOA and AOB both responding to the same source of ammonia: in microcosms amended with the antibiotic sulphadiazine, growth of AOB was inhibited while nitrification still occurred. Model calculations revealed that in such microcosms, a substantial contribution of ammonia oxidation must be attributed to AOA activity (Schauss et al., 2009). In some agricultural soils receiving significant N inputs, AOA have been shown to make a relatively small contribution to overall ammonia oxidation, with only the growth of AOB correlating with measured nitrification activity. Jia and Conrad (2009) demonstrated that in microcosms of agricultural soil receiving regular amendments of 7 mM inorganic ammonium fertiliser, growth of AOB populations (only) correlated with ammonia oxidation activity, and growth of Group 1.1b AOA populations occurred even when all nitrification activity was inhibited by acetylene. In addition, growing AOA populations did not take up 13C–CO2, indicating that they may possess heterotrophic metabolism. Di et al. (2009, 2010) reported similar findings in a number of experimental soils in New Zealand which were amended with high concentrations of urea–N (in the form of collected urine). Again, in these field soil experiments, only AOB growth showed a positive relationship to nitrification activity. However, AOA growth (and not AOB) was observed in unamended (control) soils with low levels of nitrification fuelled by mineralised organic nitrogen (Fig. 11), indicating AOA growth associated with low levels of ammonia. Conclusive evidence of AOA growth in soil associated with nitrification activity was provided by studies of an agricultural soil, again receiving no fertiliser. Tourna et al. (2008) demonstrated that with different rates of nitrification (controlled as a function of temperature), changes were associated specifically with the transcript profiles of AOA amoA. These transcriptionally active populations grew during nitrification (Offre et al., 2009), and their growth was completely inhibited when

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[(Figure_1)TD$IG] CHRISTA SCHLEPER AND GRAEME W. NICOL

Figure 11 Contrasting response of AOA and AOB communities to nitrogen deposition in a New Zealand agricultural soil. (a) Nitrification kinetics in soils receiving no amendment (control) and a high nitrogen load (collected dairy cow urine and added at an equivalent rate of 1000 kgN/ha). Growth dynamics of AOA (b) and AOB (c) communities in response to the different ammonia concentrations. (Adapted from data obtained by Di et al. (2010), with permission.)

AMMONIA-OXIDISING ARCHAEA

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[(Figure_2)TD$IG]

Figure 12 Growth of acetylene-sensitive ammonia-oxidising archaea in nitrifying soil microcosms. (a) DGGE analysis of amoA-defined AOA communities. Arrow indicates the growth of a specific population for which a specific qPCR assay was developed. (b) Inhibition of ammonia-oxidising activity in microcosms with a 10 Pa acetylene headspace partial pressure. (c) qPCR analysis demonstrating growth of AOA (group 1.1a archaea) only in microcosms with active nitrification. (Adapted from data obtained by Offre et al. (2009), with permission.)

nitrification was inhibited with the addition of low concentrations of acetylene (Fig. 12), thus providing, for the first time, a direct link between soil nitrification and archaeal activity. However, one has to note that the active AOA phylotypes in these experiments were affiliated to Group 1.1a archaea typically found in the marine environment whereas Group 1.1b archaea (typically found in soils) did not exhibit activity in these experiments. Although based on a limited number of studies, published data describing the growth dynamics of AOA and AOB populations do hint at fundamental differences in AOA and AOB physiology. From the relatively large number of cultivated AOB strains, it is known that there is a range of physiological diversity (adaptation to different ranges of ammonia concentrations,

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temperature optima, contrasting ureolytic capabilities, etc.) and it would seem likely that similar physiological diversity could be found within the AOA. Therefore, it is probable that some populations of AOA and AOB do share or compete within a similar distinct ecological niche present in soil. However, current evidence suggests the opposite may be the general rule, with populations of each lineage residing in distinct ecological niches in the soil, and ammonia concentration (also pH) being the major driver for the relative activity of AOA and AOB. In addition, the source of ammonia may be a critical factor in determining relative growth. In all soil-based studies to date, where substantial AOA growth has been demonstrated, ammonium has been supplied to the system in the form of mineralised organic N derived from composted manure (Schauss et al., 2009) or soil organic matter (Offre et al., 2009; Di et al., 2010) and AOB-dominated nitrification activity associated with ammonia from inorganic fertiliser (Jia and Conrad, 2009) or (hydrolysed) urea (Di et al., 2010).

7.3. AOA in the Marine Environment In the marine water column, nearly all AOA are placed within a specific lineage which is distinct from those associated with soil environments (Fig. 3), and is congruent with the phylogenetic partitioning of 16S rRNA genes. Thaumarchaeota (formerly crenarchaeota) are found in very large numbers throughout the water column and they have been estimated to represent approximately 20% of prokaryotic cells in the water column (Karner et al., 2001). Indeed, the relative numbers of archaea decrease much less than bacteria and therefore generally represent a greater proportion of total prokaryotic numbers at depth. However, although thaumarchaeota are distributed throughout the water column, there is a clear phylogenetic separation of distinct AOA groups, with well-defined ‘shallow’ and ‘deep’ water lineages (Francis et al., 2005; Hallam et al., 2006a; Mincer et al., 2007; Beman et al., 2008) and with only a small amount of overlap. Using specific qPCR assays for the ‘deep’ and ‘shallow’ lineages, Beman et al. (2008) demonstrated that the shallow AOA lineage was also found in deeper samples, but the deep lineage demonstrated a more restricted distribution and did not occur in the shallower waters. This observation was also reflected in the detection of amoA mRNA transcripts of both these groups (Santoro et al., 2010). It is unclear whether all these thaumarchaeota are capable of ammonia oxidation and autotrophic growth. A recent study examining the ratio of thaumarchaeal 16S rRNA and AOA amoA genes indicated that all shallow

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archaeal populations may be capable of ammonia oxidation, with 16S rRNA: amoA gene ratios approaching 1:1 (Agogue et al., 2008), and is consistent with the ratio found in the limited number of AOA genomes sequence thus far. However, with decreasing depth this ratio increases, with 16S rRNA: amoA gene ratio greater than 100:1 found at depths greater than 1000 m, thus indicating that archaea in the deep ocean may not all be autotrophic ammonia oxidisers but heterotrophs (Agogue et al., 2008). However, analysis of radiocarbon data in archaeal lipids recovered from samples taken from the North Pacific Gyre have confirmed that the dominant thaumarchaeotal metabolism at depth does appear to be autotrophy (Ingalls et al., 2006), and potential discrepancies between amoA and 16 rRNA gene copy number may be due to a lack of coverage associated with certain AOA amoA primer sets (Konstantinidis et al., 2009). There is strong correlative evidence that Group 1.1a archaea are not the only AOA lineage present in the World’s oceans. Using quantitative PCR, Mincer et al. (2007) observed that there was a discrepancy in the abundance of AOA amoA and Group 1.1a 16S rRNA genes. However, when the abundance of a novel archaeal group related to the pSL12 clade [a lineage originally discovered in terrestrial hot springs (Barns et al., 1996)] was taken into account, a strong correlation was observed. Despite the relatively large sequence divergence between these two lineages, the amoA genes of this lineage appear to be indistinguishable from Group 1.1a.

7.4. AOA Activity in the Marine Environment It is perhaps in the marine environment that the clearest correlations between AOA abundance and nitrification activity are observed, and where AOA do appear to be both numerically dominant and functionally more active (relative to AOB). Abundances of AOA amoA and thaumarchaeal 16S rRNA genes show a high correlation with nitrification rates, with up to 104–105 gene copies mL 1 in zones of high activity, and contrasts with the abundances of their bacterial counterparts which are frequently detected in low numbers or are even undetectable (Ward, 2000; Wuchter et al., 2006; Mincer et al., 2007). AOA amoA abundance correlates with nitrite maxima in both oxygenated shallow waters and deeper waters in the oxygen minimum zone (Coolen et al., 2007; Herfort et al., 2007; Beman et al., 2008). Nitrification activity (and AOA numbers) is greatest in the water column at the bottom of the euphotic zone. This may be due to competition for ammonia between nitrifiers and phytoplankton and/or light inhibition of the AMO enzyme (Ward, 2005). There is an inverse correlation between

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thaumarchaeota and chlorophyll a (Murray et al., 1998), supporting the idea that phytoplankton have a negative effect on nitrifier communities (Ward, 2005; Herfort et al., 2007). Additionally, previous studies have failed to find a correlation between bacterial ammonia-oxidising community structures and nitrification rates in ocean waters (O’Mullan and Ward, 2005). In an impressive time-series experiment over 11 months in the North Sea, Wuchter et al. (2006) quantified the abundance of inorganic nitrogen concentrations together with bacterial and archaeal amoA gene copy numbers (Fig. 13). Ammonia concentrations were greatest in autumn and winter and decreased in the spring months. This decrease in ammonia concentration correlated with not only increases in nitrite and nitrate concentrations (indicative of aerobic ammonia oxidation), but also concomitant increases in archaea amoA and 16S rRNA gene copies and also thaumarchaeotal cell numbers (enumerated by fluorescent in situ hybridisation). Besides looking at the correlation between inorganic nitrogen concentrations and gene copies, Beman et al. (2008) also measured the oxidation of 15 N-labelled ammonium pools in water samples taken from between the surface and 100 m depth in the Gulf of California. This study demonstrated that there was a correlation between AOA cell numbers and actual rates of ammonia oxidation. Correlations with the activity of other organisms involved in the nitrogen cycle are also observed. For example, increases in the abundance of AOA populations together with nitrite peaks in suboxic zones indicate that they may supply the nitrite required for planctomycete bacteria performing the anammox processes (Coolen et al., 2007). In aerobic sub-surface waters at the bottom of the euphotic zone, correlations are observed between the abundance of AOA and nitrite-oxidising Nitrospina, suggesting that the two groups are metabolically linked with AOA providing the nitrite substrate for Nitrospina populations and completing aerobic nitrification (Mincer et al., 2007; Santoro et al., 2010).

7.5. AOA in Sediments AOA amoA sequences have been recovered from both freshwater (Herrmann et al., 2009) and estuarine, coastal and deep-water marine sediments (e.g. Francis et al., 2005; Dang et al., 2009). There is a wide diversity of AOA in sediments, with sequences not only placed within the marine water/ sediment cluster where they are found in specific groups, but also affiliated to the major soil/sediment clade (Fig. 9). However, unlike in the marine column and most soils, the numerical dominance of AOA over AOB is not so

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[(Figure_3)TD$IG]

Figure 13 Correlation of fluxes in inorganic nitrogen concentrations and archaeal/AOA abundances during an 11-month sampling time series. (a) Ammonia, nitrite and nitrate concentrations. (b) Enumeration of crenarchaeol (thaumarchaeal) abundance as determined by qPCR and CARD-FISH with microscopy. (c) Abundance of AOA and AOB populations, as determined by measuring amoA gene copy numbers. (Adapted from data obtained by Wuchter et al. (2006), with permission.)

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prevalent. Both communities show strong patterns of selection with different physicochemical properties (including salinity, ammonia and oxygen concentrations) (Mosier and Francis, 2008; Santoro et al., 2008), with AOA preferring lower salinities and lower ammonia concentrations. Estuaries are particularly important as they probably experience the highest concentrations of anthropogenic N inputs in the marine environment (particularly from agricultural run-off) and therefore represent an important area of nitrifying transformations on a global scale.

7.6. AOA in Geothermal Environments Although often referred to as ‘mesophilic’ or ‘nonthermophilic’ (cren) archaea, organisms associated with this lineage were known to be present in terrestrial hot springs, through the detection of 16S rRNA genes (Kvist et al., 2005, 2007) or archaeal-specific isoprenoid GDGTs lipids such as crenarchaeol (e.g. Pearson et al., 2004; Schouten et al., 2007). These findings therefore raised the possibility that AOA would also be found in such environments. This concept was particularly fascinating as no known AOB had ever been found in such an environment, and it also raised questions about the potential origins of prokaryotic ammonia oxidation. There is now conclusive evidence that thaumarchaeota possessing AMO are found in terrestrial environments of high temperature, with AOA amoA genes detected in a variety of habitats. These include speleothems (mineral deposits), water and biofilms (Weidler et al., 2008) of geothermal caves and mines, as well as terrestrial hot springs. Thermal springs (from where sulphur-dependent Crenarchaeota are typically cultured) which represent a wide range of temperatures and broad pH ranges located on the Russian Kamchatka peninsula and on Iceland (Reigstad et al., 2008) as well as in Yellowstone National Park (de la Torre et al., 2008) and further terrestrial hot springs in the USA, China and Russia (Zhang et al., 2008) all harbour AOA gene markers. Reigstad et al. (2008) measured actual nitrification activity in an acidic hot muddy pool of 80  C under in situ conditions, demonstrating that this process is indeed found at considerable levels in terrestrial hot-springs. A thermophilic AOA (N. yellowstonii) was grown in enrichment culture obtained from a hot spring located in the Yellowstone National Park with an optimal growth temperature between 65 and 72  C, growing with a stoichiometric conversion of ammonia to nitrite. Not only does this organism grow at the highest temperature for any known ammonia oxidiser, but it represents a separate lineage outwith the ‘marine’ and ‘soil’ dominated groups (de la Torre et al., 2008).

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7.7. AOA Associated with Marine Invertebrates Associations between AOA and a variety of marine invertebrates are known, including marine sponges and corals, where they may play an important role in potentially complex nitrogen-cycling interactions within the host ‘ecosystem’. The first Group 1 ‘model’ archaeon was the sponge symbiont C. symbiosum (Preston et al., 1996), which is found in the tissues of the marine sponge A. mexicana, and is closely related to those thaumarchaeota dominating planktonic archaeal communities. The genome of this organism was the first to be sequenced within the AOA lineage (Hallam et al., 2006a,b) and possessed many interesting attributes, including the absence of some genes found in planktonic AOA (probably reflecting the specific association with the sponge), and also had near-complete components of a 3-hydroxypropionate/4-hydroxybutyrate as well as TCA cycles, indicative of both autotrophic and heterotrophic modes of growth, respectively. Recent studies of AOA amoA sequences in marine sponges (e.g. Meyer et al., 2008; Steger et al., 2008; Hoffmann et al., 2009) and corals (Beman et al., 2007; Siboni et al., 2008) have demonstrated that there are specific lineages of AOA adapted to association with marine invertebrates (Fig. 9), an observation previously found in 16S rRNA-based surveys. The association with AOA and marine sponges appears to be a continuous and stable one, with the transmission of AOA from adults to offspring in the larval stage observed in a number of different sponge species (Steger et al., 2008). A range of nitrogen transformative processes have been observed in sponges, with complex communities including anammox planctomycetes, nitrite oxidisers, denitrifiers, as well as AOA and AOB (Hoffmann et al., 2009; Mohamed et al., 2010). These sponge-associated communities therefore may represent a nitrogen cycling ecosystem which is distinct from that in the surrounding water, and one which is essential for sponge health and cycling of waste.

8. CONCLUDING REMARKS Based on the quantification of genes and cell numbers, Thaumarchaeota range among the most abundant microorganisms on this planet. Although their metabolic activity and versatility are still not entirely understood, there is no doubt that many of them are capable of ammonia oxidation and thus contribute significantly to global nitrogen and carbon cycling. Due to the extensive use of fertilisers in agriculture, the anthropogenic input of fixed

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nitrogen into the World’s ecosystems is now estimated to be more than €m et al., 2009). The major double that from natural processes (Rockstro consequence of this shift in the equilibrium of the nitrogen cycle is an acceleration of nitrification, as well as eutrophication of freshwater and estuarine environments. Another major consequence of accelerated rates of global nitrification is the increased release of nitrogen oxides into the atmosphere, which are produced by the denitrification activity of many bacteria, including ammonia oxidisers. However, it remains to be elucidated whether archaea also contribute to this process, with analyses of the first AOA genomes indicating that ammonia oxidation is performed by a fundamentally different metabolic pathway. This new area of microbiology eagerly anticipates the results of current and future research which will compare the fundamental differences (or similarities) between bacterial and archaeal ammonia oxidation in various environments to understand whether these two groups of organisms have competing (or rather complementing) roles in various ecosystem processes.

ACKNOWLEDGEMENT The authors would like to thank Martin G. Klotz for discussions and for permission to include his hypotheses on the ammonia-oxidising metabolism of AOA, and provision of Fig. 7 used in this article. The authors also gratefully acknowledge the formatting work of Nathalia Jandl and support for Table 2 as well as Fig. 8 from Anja Spang.

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Reductive Stress in Microbes: Implications for Understanding Mycobacterium tuberculosis Disease and Persistence Aisha Farhana1, Loni Guidry1, Anup Srivastava1, Amit Singh2, Mary K. Hondalus3 and Adrie J.C. Steyn1 1 Department of Microbiology, University of Alabama at Birmingham, AL, USA International Center for Genetic Engineering and Biotechnology, Aruna Asaf Ali Marg, New Delhi, India 3 Department of Infectious Diseases, University of Georgia, Athens, GA, USA

2

ABSTRACT Mycobacterium tuberculosis (Mtb) is a remarkably successful pathogen that is capable of persisting in host tissues for decades without causing disease. Years after initial infection, the bacilli may resume growth, the outcome of which is active tuberculosis (TB). In order to establish infection, resist host defences and re-emerge, Mtb must coordinate its metabolism with the in vivo environmental conditions and nutrient availability within the primary site of infection, the lung. Maintaining metabolic homeostasis for an intracellular pathogen such as Mtb requires a carefully orchestrated series of oxidation–reduction reactions, which, if unbalanced, generate oxidative or reductive stress. The importance of oxidative stress in microbial pathogenesis has been appreciated and well studied over the past several decades. However, the role of its counterpart, reductive stress, has been largely ignored. Reductive stress is defined as an aberrant increase in reducing equivalents, the magnitude and identity of which is determined by host carbon source utilisation and influenced by the presence of host-generated gases (e.g. NO, CO, O2 and CO2). This increased reductive

ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 57 ISSN: 0065-2911 DOI:10.1016/B978-0-12-381045-8.00002-3

Copyright Ó 2010 by Elsevier Ltd. All rights reserved

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power must be dissipated for bacterial survival. To recycle reducing equivalents, microbes have evolved unique electron ‘sinks’ that are distinct for their particular environmental niche. In this review, we describe the specific mechanisms that some microbes have evolved to dispel reductive stress. The intention of this review is to introduce the concept of reductive stress, in tuberculosis research in particular, in the hope of stimulating new avenues of investigation.

1. 2. 3. 4.

5.

6.

7.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Scope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Concept of Reductive Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Overview: General Physiological Characteristics of Mycobacterium tuberculosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Historic Overview. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Environmental Factors that Influence Metabolism . . . . . . . . . . . . . . Reductive Sinks in Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Polymer Deposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Nitrate Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4. Phenazine Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5. Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6. The Reverse TCA (rTCA) Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.7. Carbon Monoxide (CO) Dehydrogenase (CODH) . . . . . . . . . . . . . . . 5.8. Other Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Redox Sinks in Mycobacteria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. The Mycobacterial Intracellular Redox Environment. . . . . . . . . . . . . 6.2. The Mtb Dos Dormancy Regulon . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Mtb WhiB3 is an Intracellular Redox Sensor that Counters Reductive Stress. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

ABBREVIATIONS GSH GSSG

glutathione glutathione disulfide (oxidised glutathione)

44 45 46 47 49 49 49 50 59 59 64 66 67 70 72 74 75 76 76 79 88 88 95 98 98

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Sometimes scientific progress is not based on a discovery or the generation of new data but on a change of viewpoint that allows one to see a set of already existing data in a new light’ (Michael Reth)

1. INTRODUCTION Tuberculosis (TB), caused by Mycobacterium tuberculosis (Mtb), is a disease of great international concern and the leading cause of death worldwide from a curable infectious disease. Across the globe, one human life is lost to TB every 15 s (WHO Factsheet, 2009). The situation is further exacerbated by the coexistent HIV epidemic, and the emergence of multidrug resistant (MDR), extensively drug resistant (XDR) and Super-XDR (S-XDR) Mtb strains (Gandhi et al., 2006; Pillay and Sturm, 2007; Velayati et al., 2009). Despite the availability of ample genomic, proteomic and bioinformatic information on Mtb, it is estimated that in 2009 more TB-related deaths occurred than at any time in history (Fauci, 2008). The variably efficacious BCG (Bacille Calmette-Gu
erin) vaccine remains the only available TB vaccine and no new anti-mycobacterial drug has been deployed since the discovery of rifampicin in 1963 (Duncan, 2004; Young et al., 2008; Kaufmann et al., 2010). The slow pace in the development of TB intervention strategies compared to an overwhelming increase in global TB incidence compromises the achievements made in TB control. An important hurdle to the development of successful TB treatment regimes is the lack of knowledge concerning the mechanisms by which Mtb is able to persist in a dormant state, unresponsive to anti-mycobacterial drugs (Gomez and McKinney, 2004; Sacchettini et al., 2008; Ma et al., 2010). We have yet to understand the physiological status of the persisting mycobacterial organisms or the environmental cues which lead to reactivation of disease. Detailed knowledge of this persistent state of Mtb is crucial for the establishment of efficacious TB eradication schemes. Mtb displays a remarkable capacity to persist in latent form and switch between replicative and non-replicative (dormant) states in response to environmental signals generated by the host immune responses (Cosma et al., 2003; Warner and Mizrahi, 2007; Rustad et al., 2009). Mtb harbours the machinery necessary to synthesise almost all essential vitamins, amino acids and enzyme cofactors, providing the organism with the ability to alter its metabolic state enabling an aerobic (e.g. oxidative phosphorylation) and possibly an anaerobic mode of respiration (Wheeler and Ratledge, 1994; Muttucumaru et al., 2004). Importantly, this metabolic flexibility ensures

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bacilli survival in the varied environments within the human host ranging from that of high oxygen tension in the lung alveolus to microaerophilic conditions within the tuberculous granuloma (Ulrichs and Kaufmann, 2006). Most studies of the physiology and biochemistry of mycobacteria were carried out in the early 1910–1980s, and it was discovered that fundamental differences exist in the metabolism of Mtb cultured in vitro and that of bacilli growing in vivo. An important difference includes demonstrating that Mtb harvested from lungs showed inactive respiratory responses to various carbohydrates and glycolytic intermediates, whereas positive responses were obtained to these same substrates by the same strain when cultured in vitro (Dubos, 1953; Artman and Bekierkunst, 1961; Segal, 1965, 1984; Brezina et al., 1967). Unfortunately, much of the data generated from these classical studies remain hidden in the historical ‘archives’ not accessible through PubMed or similar literature database searches. In part, the goal of this review is to excavate some of this ‘buried’ information on Mtb and integrate it with the current understanding of metabolic paradigms of prokaryotic and lower eukaryotic organisms.

2. SCOPE In this review, we aim to introduce the idea of ‘reductive stress’ in TB research. A strong emphasis is placed on the historical knowledge of Mtb physiology obtained by in vivo studies performed in the earlier half of the last century, because in some respects, these analyses are a lost art in the modern era of molecular techniques. This is then followed by a discussion of the in vivo factors that affect Mtb growth and metabolic mechanisms, such as redox sinks, which microorganisms have evolved to maintain redox homeostasis in response to oxido-reductive stress. Parallels between oxidoreductive pathways in mycobacteria versus other bacteria and yeast are highlighted. Metabolic engineering approaches that modulate reductive stress are also described. Next, the intracellular redox environment of Mtb is discussed followed by a description of the best-known paradigm for signal transduction in Mtb: the Dos dormancy regulon, and its role in maintaining redox balance. Lastly, the role of the intracellular redox sensor, Mtb WhiB3, in maintaining redox homeostasis is discussed. This review does not cover oxidative stress per se, but it is considered when appropriate to the theme. Regarding virulence and persistence, and general mycobacterial metabolism, we refer the reader to several articles that discuss these issues in detail (Ramakrishnan et al., 1972; Cosma et al., 2003; Boshoff and Barry, 2005; Hett

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and Rubin, 2008; Barry et al., 2009; Rustad et al., 2009; Meena and Rajni, 2010; Paige and Bishai, 2010). In sum, we aim to present a clear analysis of the current knowledge of reductive stress in microorganisms in order to provide a better foundation for future interpretation of the physiological events associated with Mtb infection.

3. THE CONCEPT OF REDUCTIVE STRESS The physiology and metabolism of Mtb are unique, allowing it to survive under a wide range of in vitro and in vivo environmental conditions. This flexibility is evident from the fact that Mtb is exposed to a plethora of products including carbohydrates, organic acids, lipids, amino acids, ions, etc., as well as gases such as nitric oxide (NO) (Voskuil et al., 2003, 2009) carbon monoxide (CO) (Kumar et al., 2007, 2008), carbon dioxide (CO2) (Florczyk et al., 2003), oxygen (O2) (Voskuil et al., 2003) and its corresponding free radicals in vivo. These molecules subsequently inflict either oxidative or reductive stress within the bacteria. Over the years, oxidative stress and the critical role it plays in a wide range of diseases has been well studied; however, the role of its counterpart, namely reductive stress, has largely been underappreciated. The likely reasons for this, primarily, include a lack of understanding of the concept of reductive stress and the dearth of experimental techniques for examining it (Ghyczy and Boros, 2007). A crucial element in reductive stress is redox coupling, which entails electron transfer. More specifically, redox reactions involve the transfer of electrons and hydrogen atoms from an electron donor (reductant or reducing agent) to an electron acceptor (oxidant or oxidising agent), which together function as a redox couple. These redox couples (e.g. NAD+/NADH, E0’ =
315 mV; NADP+/NADPH, E0’ =
320 mV; FAD/FADH2, 0’ E =
219 mV; 2GSH/GSSG, Ehc =
250 mV [10 mM]) are vital to both anabolic and catabolic reactions. NADH functions as an energy-rich electron transfer coenzyme, which generates almost three ATPs for every NADH to NAD+ oxidation event, whereas NAD+ functions as a sink for electrons. In contrast, NADPH is the primary source of electrons for reductive synthesis or anabolism of fatty acids (FAs) and reduction of the glutathione system, which is the key cellular antioxidant defence system. Thus, the NAD+/ NADH coenzyme system required for catabolism contrasts with the NADPH/NADP+ system that is required for anabolism (Voet et al., 2008). The above redox couples are often thermodynamically linked because elevated levels of either reductant or pro-oxidant are deleterious to the

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microbial cell (Schafer and Buettner, 2001). Balanced rates of oxidation and reduction of these molecules are necessary for optimal metabolic function as redox imbalance in cells can lead to either oxidative or reductive stress, of which the latter has mostly escaped the attention of the scientific investigator. Reductive stress can be defined as an abnormal increase in reductive equivalents (e.g. NADH, NADPH, GSH, etc.) or reducing power (Dimmeler and Zeiher, 2007; Ghyczy and Boros, 2007; Zhang et al., 2010). The central focus of this review is to examine the mechanisms used by microorganisms and Mtb in particular to recycle reducing equivalents in order to maintain redox balance. The formal concept of reductive stress emerged little over a decade ago. Using animal models for diabetes, several studies reported that the metabolic imbalance linked to an increased blood flow to the retina, kidney and peripheral nerve is cytosolic reductive stress. This increased NADH/NAD+ ratio or hypoxia-like state is linked to the increased oxidation of substrates such as sorbitol, glucoronic acid and non-esterified FAs, and to the reduction of NAD+ to yield NADH (Ido et al., 1997; Tilton, 2002; Ido, 2007). Since an increase in NADH was observed under hypoxic conditions, the observation was termed ‘pseudohypoxia’, or referred to as the ‘reductive stress hypothesis’ (Ido, 2007). In a seminal study, Rajasekaran et al. (2007) reported reductive stress in mice expressing the mutant human ab-crystallin gene. In this study, because of increased activity of glucose-6-phosphate dehydrogenase (G6PD), enhanced levels of NADPH and GSH caused protein aggregation, cardiomyopathy and increased expression of heat shock proteins (Hsp) including Hsp27. Since NADPH is a cofactor of NADPH oxidases and NO synthases, these findings established a link between reductive stress and oxidative or nitrosative stress signaling pathways. Subsequently, it was shown that overexpression of Hsp27 induces reductive stress in the heart (Zhang et al., 2010), which was evident by an increase in 2GSH/GSSG, myocardial glutathione peroxidase activity and decreased levels of reactive oxygen species (ROS). Interestingly, 2GSH levels rose but GSSG levels remained unaltered. In another study, G6PD was overexpressed in Drosophila melanogaster, which resulted in increased levels of NADH and NADPH, and increased GSH/GSSG ratio. The presence of high amounts of these reducing equivalents enhanced resistance to oxidative stress and were associated with an extension of life span in the transgenic flies (Legan et al., 2008). On the other hand, increased reductive stress may also lead to an increased oxidative stress, as was demonstrated in hypoxic injury studies (Gores et al., 1989; Khan and O’Brien, 1995). In those studies, it was proposed that reduction of electron carriers that are normally oxidised under aerobic conditions (reductive stress) promotes formation of toxic ROS upon

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O2 availability. In an attempt to address the possible mode of action, it was shown that reducing equivalents can release redox-active iron leading to oxidative stress and cell injury (Staubli and Boelsterli, 1998). In conclusion, it is clear that normal cellular functions essentially depend on maintaining redox homeostasis. A redox imbalance can lead to either oxidative or reductive stress. Lastly, it is evident that reductive stress might be a common mechanism in many eukaryotic diseases but, unfortunately, its implications in microbial pathogenesis are poorly understood. The concept of reductive stress in bacteria, particularly as it applies to Mtb, is an example of a shift in perspective.

4. OVERVIEW: GENERAL PHYSIOLOGICAL CHARACTERISTICS OF MYCOBACTERIUM TUBERCULOSIS 4.1. Historic Overview Mtb is a prototrophic, obligate aerobe that is able to survive periods of extended anaerobiosis, although conditions are yet to be identified wherein the bacilli are capable of replication in the absence of O2. In fact, studies reported in 1933 established that Mtb could survive for up to 12 years in sealed tubes and remain fully virulent (Corper and Cohn, 1933). Mycobacteria can utilise a wide range of carbon compounds for growth in vitro including carbohydrates, lipids and proteins, which suggests that the bacilli are able to assimilate a wide range of host substrates for growth in vivo. For example, micromolar quantities of organic acids (e.g. lactate, pyruvate, citrate, succinate, malate, acetoacetate, etc.), micro to millimolar quantities of carbohydrates (glucose, glycogen, fructose, etc.), micromolar quantities of virtually all amino acids, nucleic acid precursors, nucleotides, and milligram to gram/litre quantities of lipids (including total and free FAs, triacylglycerol [TAG] and total cholesterol) are available as sources of metabolic energy (Wheeler and Ratledge, 1994). The degradative pathways of the above substrates converge on common intermediates, including in many cases acetyl-coenzyme A (acetyl-CoA) that eventually produce ATP. Decades ago, many of the scientific studies of Mtb metabolism and physiology were performed on in vivo-derived bacilli (Segal and Bloch, 1956; Artman and Bekierkunst, 1961; Segal, 1962, 1965, 1984). The complex and laborious approaches (e.g. isolating, purifying and characterising bacilli from infected mouse lungs) yielded a wealth of information regarding the

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differences between the in vivo and in vitro physiology of Mtb. In addition, differences between virulent and avirulent Mtb strains were noted. In recent years, a number of eloquent molecular and metabolic studies of Mtb have been performed. Although valuable information was obtained, an inherent weakness of much of that work was the use of in vitro cultured bacilli. The chemical makeup of artificial growth media critically influences the biochemical activity of the organism, and the applicability of data thus generated is limited to the conditions under which it was derived. Therefore, keeping the complexity of the host environment in mind, certain aspects of Mtb physiology will undoubtedly have to be revisited when examining Mtb in vivo. Nonetheless, ample data exist to indicate that Mtb adjusts its metabolism in response to the availability of nutrients and environmental gases during different stages of infection (Wheeler and Ratledge, 1994; McKinney et al., 2000; Boshoff and Barry, 2005; Tian et al., 2005; Munoz-Elias et al., 2006; Jain et al., 2007; Barry et al., 2009). It is therefore important to understand how this metabolic response permits the bacterium to persist long term in the human host.

4.2. Environmental Factors that Influence Metabolism 4.2.1. The TCA Cycle The global metabolic pathway of a microbial cell is an interlinked network of chemical reactions through which the cell breaks down substrate compounds into smaller organic molecules, which then serve as precursors for the biosynthesis of diverse macromolecules. Microorganisms employ different metabolic strategies of which the ultimate goal is to generate a proton motive force and cellular energy, ATP. The TCA cycle is present in all aerobic organisms and serves as a means to oxidise carbohydrates such as glucose to CO2 and H2O and the energy released is efficiently harvested by the electron transport chain (ETC). The TCA cycle is amphibolic because it can be used for both anabolic and catabolic processes, and yields much more energy per mole of glucose (38 moles of ATP) when completely oxidised than the 1–4 moles of ATP generated via anaerobic fermentation. The complete oxidation of 1 mole of glucose via glycolysis and the TCA cycle of Escherichia coli yields 10 moles of NAD(P)H and 2 moles of FADH2 (Vemuri et al., 2006) as depicted below: Glucose + 8NAD+ + 2NADP+ + 2FAD + 4ADP + 4Pi ! 6CO2 + 8NADH + 2NADPH + 2FADH2 + 4ATP + 10H+

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Most of the TCA cycle enzymes are repressed by glucose and further repressed by anaerobiosis. Under aerobic conditions, succinate is formed through the oxidation of a-ketoglutarate (KG), whereas under anaerobic conditions bacteria form succinate through reduction of fumarate. Under anaerobic growth, the 2-oxoglutarate dehydrogenase complex (ODHC) and succinate dehydrogenase (SDH) are also repressed, causing the activity of the TCA cycle to virtually cease. The cycle is thus transformed into its branched or non-cyclic form in which the carbon flows into independent oxidative and reductive pathways leading to the formation of 2-oxoglutarate (glutamate), and succinate and succinyl-CoA respectively (Amarasingham and Davis, 1965; Spencer and Guest, 1987; Guest and Russell, 1992). In the branched or reductive pathway, SDH is replaced by fumarate reductase (FRD) which enables fumarate to be used as an electron acceptor in anaerobic respiration. Alternative anaerobic routes to succinate production occur either via aspartate involving aspartate oxaloacetate aminotransferase, or via isocitrate catalysed by isocitrate lyase (Spencer and Guest, 1987; Guest and Russell, 1992). Environmental factors such as the availability of O2 and the nature (e.g. carbon oxidation state, COS; see Section 4.2.2) and quantity of the carbon source profoundly affect the status of the TCA cycle (Spencer and Guest, 1987; Clark, 1989; Guest and Russell, 1992). O2 is a poisonous lethal gas, which allows aerobic microbes to survive as they have developed appropriate antioxidant defence mechanisms (Halliwell, 2008). TCA cycle enzymes known to be inhibited by the O2 radical, superoxide anion (O2
), and under high pO2 include aconitase, isocitrate dehydrogenase, a-ketoglutarate dehydrogenase (KDH) and fumarase (Halliwell, 2008). Several of these enzymes contain 4Fe–4S clusters, which fall apart when targeted by O2 or O2
. The inactivation of these enzymes leads to the release of iron, which can then promote the production of OH via the Fenton reaction. In addition, NO, a host-generated gas can effectively and irreversibly react with the Fe–S clusters of TCA cycle enzymes leading to the formation of a proteinbound DNIC complex, which affects specific enzymatic activity and overall metabolic activity of the cell (Imlay, 2006, 2008; Duan et al., 2009). In accordance with the critical role of the TCA enzymes in the production or consumption of reducing equivalents, it is logical to believe that O2, O2
and NO also affect redox balance. Nonetheless, the impact of these diatomic gases and oxygen radicals on the components of the Mtb TCA cycle is an understudied area. The first evidence of TCA cycle activity during in vivo growth of Mtb was the demonstration of SDH activity in in vivo-derived bacilli (Segal, 1962). Subsequently, the activity of all the TCA cycle enzymes with the exception of *

*

*

*

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KDH was established through the analysis of Mycobacterium lepraemurium harvested from murine lepromas (Mori et al., 1971; Tepper and Varma, 1972). Another important study which characterised the enzymes of the Mtb TCA cycle noted that all the dehydrogenases, unlike those present in other organisms, are NADP+-dependent with the exception of malate dehydrogenase which is NAD+-dependent (Murthy et al., 1962). Likely explanations for this finding are: (i) the NADP+ dependence of the Mtb dehydrogenases ‘guarantees’ the presence of substantial quantities of NADPH, the reducing agent necessary for metabolic biosynthesis of essential lipids, and (ii) the presence of NADH oxidase ensures that adequate NAD+ is continuously available as an oxidising agent during these processes (Murthy et al., 1962). Present-day genomic analysis seems to support the above interpretations in that a considerable portion of the Mtb genome is dedicated to lipid anabolism, which requires NADPH. The work of Wayne and others (Segal, 1984) identified a switch from aerobic to anaerobic metabolism during in vivo growth and found this to be an important factor in virulence. Notably, the identification of hypoxia as an in vivo signal for metabolic transformation profoundly affected future scientific studies and led many years later to the widely used Wayne model of in vitro dormancy (Wayne and Hayes, 1996). This in turn facilitated the identification of the 48-member Mtb Dos dormancy regulon, a genetic response induced by hypoxia, NO and CO (Sherman et al., 2001; Ohno et al., 2003; Voskuil et al., 2003; Kumar et al., 2008; Shiloh et al., 2008), which has become a paradigm for Mtb signal transduction in response to host cues (see Section 6.2 for a complete discussion). The implication is that the metabolic adaptation or response of Mtb to the lack of O2 or the presence of NO and CO in vivo induces the Dos regulon and allows establishment of a latent infection. This raises several important questions such as which terminal electron acceptor, besides O2, is used in vivo, and how are reducing equivalents re-oxidised to maintain redox balance? 4.2.2. The Carbon Oxidation State (COS) Experimental evidence in support of FAs as potential in vivo carbon sources for Mtb was provided several decades ago (Segal and Bloch, 1956; Segal, 1984), and is supported by many recent studies (McKinney et al., 2000; Munoz-Elias and McKinney, 2005). In vivo grown Mtb and M. lepraemurium were shown to robustly oxidise long-chain FA such as n-heptanoic, octanoic, oleic, palmitic, steric, linoleic, linolenic and luric

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acids, but failed to utilise carbohydrates (Segal and Bloch, 1956; Segal, 1984). This disagrees with the impression that bacilli in infected tissue exist in a reduced state of metabolic activity. These findings are further supported by recent Mtb genome data revealing the presence of 36 homologs of fadE and fadD genes catalysing the first step of b-oxidation. Other bacteria such as E. coli and Salmonella enterica serovar Typhimurium, have only a single fadE gene (Campbell and Cronan, 2002). Several studies strongly suggest that isocitrate lyase (icl), an enzyme of the glyoxylate cycle, enabling the recycling of acetyl-CoA (formed via b-oxidation), plays an important role in FA carbon source utilisation in vivo. Studies of an Mtb Dicl mutant in mice (McKinney et al., 2000) showed that this mutant strain is attenuated at the onset of adaptive immunity (3 weeks post-infection) in immunocompetent animals, but remains virulent in g-IFN-deficient mice. Further research on icl (Munoz-Elias and McKinney, 2005; Munoz-Elias et al., 2006) substantiates the importance of lipids as an important in vivo carbon source for Mtb. The effect of a carbon source (e.g. glucose vs. FA) on the ‘spontaneity’ of a process, as defined by the Gibbs free energy (G) (Voet et al., 2008) is illustrated by the fact that the complete oxidation of glucose yields DG ’ =
2850 kJ/mol, whereas oxidation of a C16 FA such as palmitate (C 16H32O2, a putative in vivo carbon substrate of Mtb) is more exergonic and yields DG ’ =
9781 kJ/mol. Palmitate and oleate have highly reduced carbon oxidation states (COSs) of
28 and
30 respectively, compared to other FA precursors such as propionate (COS =
1), valerate (COS =
6) and carbohydrates such as glucose (COS = 0) and sorbitol (COS =
1). Subsequent b-oxidation of palmitate generates 106 ATP, whereas oxidation of glucose produces 38 ATP. Importantly, b-oxidation of FA yields one NADH and one FADH2 molecule for every acetyl-CoA generated, a condition which has the potential to cause cellular redox imbalance leading to reductive stress if the consequent buildup of reducing equivalents is not dissipated. Thus, the oxidation state of the carbon source determines the amount of reducing equivalents [e.g. NAD(P)H] to be recycled and consequently also the excreted products (see Section 4.2.3). In E. coli, it has been shown that the COS, extracellular oxido-reduction potential and environmental pH (Kleman and Strohl, 1994) influence the composition of excreted fermentation products. For example, in E. coli, oxidation of glucose and sorbitol generates two and three reducing equivalents respectively, whereas utilisation of the highly oxidised sugar glucuronic acid (COS = +2) results in no NADH production. Thus, in order to recycle the NADH produced during growth on the more reduced substrate, sorbitol,

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E. coli excretes reduced ethanol (COS =
2). In contrast, cells grown on glucuronic acid are redox balanced and do not need to produce ethanol; rather, glucuronic acid is converted to acetate (COS = 0) (Wolfe, 2005). Other studies have shown that as the pH drops, E. coli produces lactate rather than acetate or formate (Bunch et al., 1997), and that the rate of glycolysis is dramatically reduced (Ogino et al., 1980). Clearly, the physicochemical properties of a particular carbon source (e.g. FA or glucose and therefore the COS), environmental factors and the metabolites produced during substrate utilisation profoundly affect redox balance and thus overall microbial physiology. 4.2.3. Excretion of Metabolites and Redox Balance E. coli regenerates NAD+ under anaerobic conditions via the production and excretion of partially oxidised metabolic intermediates such as D-lactate, succinate, formate and ethanol. Similarly, acetate excretion by E. coli occurs anaerobically during mixed acid fermentation in order to regenerate the NAD+ consumed by glycolysis and to recycle Coenzyme A (CoASH) utilised during the conversion of pyruvate to acetyl-CoA (Wolfe, 2005). Acetate can also be excreted during aerobic growth on high concentrations of glucose (Crabtree effect), which inhibits respiration (Ko et al., 1993; Wolfe, 2005). Because NAD+ is required by the glycolytic enzyme glyceraldehyde-3phosphate dehydrogenase (GAPDH), E. coli must re-oxidise NADH to maintain a working glycolytic pathway. In the absence of a functional TCA cycle during anaerobic growth, the reducing equivalents are recycled by the production of metabolic intermediates such as D-lactate, ethanol, succinate and formate, which are secreted along with acetate into the culture medium. However, acetate excretion produces ATP, whereas the other metabolites are not used as energy harvesting molecules, but rather consume reducing equivalents (Wolfe, 2005). Thus, under anaerobic conditions, bacteria excrete a range of products in order to regenerate NAD+ and to maintain redox balance. Other studies have yielded a few clues as to the intermediary metabolic changes mycobacteria undergo during aerobic respiration and oxidation of diverse substrates. An interesting observation made in 1930 (Merrill, 1930) was that mycobacteria utilise carbohydrates without the production of acids, suggesting that carbohydrates are completely oxidised, leaving insignificant amounts of partially oxidised products (e.g. acids) in the medium (Merrill, 1930; Edson, 1951). Initially, precise manometer measurements of respiratory changes (the respiratory quotient) were determined by measuring O2 consumption and CO2 production of bacilli growing on carbon sources such as

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glucose or glycerol. However, it quickly became clear that in order to accurately interpret the respiratory quotients, lipid and protein content of the bacilli had to be determined. Subsequently, ‘starved’ bacilli (achieved by floating bacilli on phosphate buffered saline for several days) rather than ‘washed’ cell suspensions were used in manometer techniques. Notably, autorespiration was barely impaired after 1–4 days of starvation. It subsequently became clear that glycerol, acetate and FA enhanced respiration whereas glucose stimulation was weak, and arabinose, fructose, mannose and inositol showed no effect (Edson, 1951). However, an intriguing and important observation was that there did not seem to be a direct correlation between growth and the respiring capacity of a substrate. In fact, a substrate that promotes respiration could inhibit or induce bacterial growth, or may have no influence at all (Bloch et al., 1947). Carbon balance experiments have shown that when glucose was used as a carbon source, 34% of its carbon was recovered as CO2, 63% was found in the bacilli and 2–6% remained in the media (Edson, 1951). Several studies demonstrated that human and bovine tubercle bacilli growing in glycerol medium generated alkaline culture supernatants (reviewed in Merrill, 1930). This contrasts with the vast majority of bacteria, which produce organic acids as cleavage products upon the utilisation of carbohydrates. Some researchers also argued that minute quantities of acids were formed from the oxidation of glycerol, whereas others believed that glycerol was completely utilised without the production of intermediates. However, unconfirmed studies (Fowler et al., 1960) claimed that virulent and avirulent mycobacterial strains accumulate acetic acid, succinic acid, malic acid, citric acid, oxalic acid and pyruvic acid in the culture filtrate. Further, Mycobacterium butyricum became a model organism for studying acid formation, since it was noted that M. butyricum acidifies its culture medium. Using this organism, several studies demonstrated the excretion of a-ketoglutaric acid (2-oxoglutaric acid), succinic acid, acetic acid and pyruvic acid into the culture filtrate (Hunter, 1953; Wright, 1959). Succinic, acetic and fumaric acids and DL-5-carboxymethylhydantoin were also isolated as crystalline products from Mtb and Mycobacterium ranae cultured in a defined medium containing asparagine, glycerol and trace quantities of citrate (Fowler et al., 1960). Acetyl L-isoleucine and acetyl L-leucine were also identified in culture filtrates of M. ranae (Fowler et al., 1961). Although a recent mass spectrometry-based study identified small quantities of pyruvate (18 mM), succinate (15 mM) and lactate (15 mM) (Goodwin et al., 2006) in the culture supernatants of Mtb, the experimental conditions were limited, necessitating a more comprehensive investigations to examine excreted metabolic intermediates of Mtb under a range of environmental conditions. Thus, unlike E. coli, mycobacterial species in general appears not to excrete

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large amounts of intermediary metabolites and therefore has a distinct metabolic mechanism to maintain intracellular redox balance. 4.2.4. The Balancing Act In Vitro and In Vivo In agreement with prior findings (Segal and Bloch, 1956), many studies have confirmed that there are metabolic distinctions between in vivo and in vitro grown mycobacteria. For example, differences exist between phtiocol, tuberculostearic acid, phtioic acid, specific polysaccharides (Anderson et al., 1943) and the lipid content of in vitro cultured Mtb and that of bacilli in tuberculous lung tissue (Sheehan and Whitwell, 1949). The subsequent development of differential centrifugation techniques to purify Mtb from animal lung tissue (Segal and Bloch, 1956), and biochemical comparison with in vitro grown Mtb led to a profoundly new understanding of the phenotypic and metabolic states of Mtb grown in vitro and in vivo. In these studies, separation of tubercle bacilli from infected lungs involved the use of isotonic sucrose and differential centrifugation at low temperatures to yield considerable quantities of highly purified bacilli. Using Warburg manometry (which measures O2 consumption) and testing the hydrogen transfer capacity of bacilli in the presence of a range of substrates and the electron acceptor 2,3,5-triphenyl tetrazolium chloride, it was shown that the metabolic activity of in vivo grown Mtb was very low compared to that of in vitro cultured Mtb, which was maintained for at least 20 h post-purification. In addition, the respiratory response of in vivo grown Mtb to glucose, glycerol, sodium lactate, sodium acetate and sodium pyruvate was shown to be virtually absent. On the other hand, salicylate and the FA n-heptanoic acid, octanoic acid and oleic acid, stimulated respiration to the same degree in in vivo grown bacilli as that observed in in vitro cultured Mtb. The robust respiratory responses of the bacilli isolated from the lungs towards FAs suggest that in vivo bacilli do not exist in a reduced state of metabolic activity. Intriguingly, in vitro culturing of the lung-derived Mtb in standard culture medium rapidly reversed the in vivo phenotype to that of the in vitro cultured bacilli (Segal and Bloch, 1956). An additional study (Segal, 1962) raised concerns regarding the validity of in vitro based experiments by demonstrating that untreated whole cells of in vivo grown Mtb exhibit active SDH activity, whereas in vitro cultured Mtb cells are negative for SDH activity (cell-free extracts of the latter were shown to be positive for SDH activity). In another study, the lack of respiratory responses of in vivo grown Mtb for succinate, fumarate, a-oxoglutarate, malate and glyoxylate was in fact due to the impermeability of the bacilli because cell free extracts, but not intact cells, oxidised these substrates in the presence of an electron acceptor (Murthy et al., 1962). This metabolic

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disparity between in vitro and in vivo grown Mtb was hypothesised to be (i) initial repression of the TCA cycle and/or repression of the ETC during in vivo growth, (ii) host-induced inhibition of Mtb oxidative enzymes during in vivo growth or (iii) substrate impermeability during in vivo growth (Segal, 1984). Collectively, the metabolic dissimilarity between in vivo and in vitro grown bacilli pointed to a metabolic shift away from the respiratory pathway towards anaerobic glycolysis (Segal, 1984). The gross morphological and biochemical differences between in vivo and in vitro cultured Mtb, as indicated by the above-mentioned studies, are in agreement with modern expression studies demonstrating differential expression of genes encoding proteins needed for cell wall synthesis, virulence lipid anabolism and energy production in macrophages, animal models and humans (Triccas et al., 1999; Talaat et al., 2004, 2007; Shi et al., 2005; Rachman et al., 2006; Srivastava et al., 2007, 2008; Fontan et al., 2008). In a seminal study, transcriptional analysis of Mtb derived from infected lung samples (Rachman et al., 2006) found dramatic changes in genes involved in cell envelope, lipid biosynthesis, FA and mycolic acid biosynthesis, and anaerobic respiration. In addition, using the mouse model for TB, in vivo lipidomics studies have suggested a link between host lipid catabolism and increased production of Mtb virulence lipid (PDIM, SL-1) (Jain et al., 2007). In an elegant study examining the transcriptional profile of Mtb in sputum, genes involved in anaerobic respiration and tgs1, which encodes for the enzyme responsible for producing TAG (Garton et al., 2008), were found to be overexpressed. Tgs1 is under the strict control of the Dos dormancy regulon that is induced by hypoxia, NO and CO (Sherman et al., 2001; Ohno et al., 2003; Voskuil et al., 2004). TAG production in sputum contradicts the assumption that sputum contains aerobically replicating bacilli. It has been argued that hypoxic conditions do not exist in sputum and thus the Dos dormancy regulon would not be induced (Barry et al., 2009). While the pO2 concentration of tuberculous sputum has not been established, Worlitzsch et al. (2002) reported an in situ pO2 concentration of 2.5 mm Hg in the mucus of cystic fibrosis (CF) patients, a measurement made via a Clarke electrode attached to a fibre-optic bronchoscope. The latter finding along with the severely restricted diffusion of O2 through mucopurulent luminal material (Worlitzsch et al., 2002) provides good evidence that a hypoxic environment can indeed be generated in sputum. Using fluorescent dyes and cell surface antibodies to examine the ultrastructure of Mtb in mice and guinea pigs, it was shown that Mtb exists in different subpopulations (Ryan et al., 2010). This suggests that Mtb in vivo exist as varying stochastic phenotypes, which may allow the bacilli to adapt to a changing host environment. Collectively, modern-day gene expression,

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cellular and morphological data are in support of the classical biochemical studies (Anderson et al., 1943; Segal and Bloch, 1956; Segal, 1965, 1984), which reported profound differences in cell wall architecture, virulence, lipid production and energy metabolism between in vivo grown Mtb and in vitro cultured bacilli. 4.2.5. The Gaseous Environment of the Lung An irrefutable finding, based upon 100 years of study, ascertained that Mtb cannot replicate in the absence of O2. The rate of Mtb multiplication decreases rapidly as the partial O2 pressure (pO2) falls below that of room air (Dubos, 1953; Wayne and Hayes, 1996). The pO2 of atmospheric O2 is 150–160 mm Hg and drops substantially in the lung (60–150 mm Hg) and blood (104 mm Hg) (Aly et al., 2006; Brahimi-Horn and Pouyssegur, 2007), the rat spleen (16 mm Hg) and thymus (10 mm Hg) (Braun et al., 2001) and the TB granuloma (1.59 mm Hg) (Via et al., 2008). The total alveolar surface area encountered by O2 in inhaled air is 130 m2, which allows for optimum O2 exchange (Murray, 2010). Besides O2, another gas, NO, is encountered by Mtb during infection. NO is a small, highly diffusible free radical. Inducible nitric oxide synthase (iNOS) and therefore NO production are crucial for protection of mice against Mtb (MacMicking et al., 1997; Chan et al., 2001). Importantly, human macrophages present in Mtb-infected tissues have been demonstrated to express iNOS (Nicholson et al., 1996). Similarly, increased exhaled NO and NO3
levels in patients with active pulmonary TB were shown to be due to increased iNOS production (Wang et al., 1998). In macrophages, iNOS uses NADPH and O2 as cofactors and produces NO and its oxidative products NO2
and NO3
. The diffusion distance of NO is 175 mm (Leone et al., 1996) and the concentrations in skin, a single endothelial cell, and rat lung ranges from 0.14 to 0.95 mM (Clough et al., 1998; Brovkovych et al., 1999). Although it is intuitively assumed that NO is present in TB lesions, the fact that iNOS requires O2 as cofactor for its enzymatic function (K mO2 ¼ 135 mM) (Dweik, 2005) suggests that NO production would be severely inhibited within these hypoxic granulomas [1.59 mm Hg (Via et al., 2008)]. CO is a diatomic gas that is endogenously produced by heme oxygenase-1 (HO-1) in the human lungs in response to oxidative stress. HO-1 enzymatic activity requires three moles of molecular O2 per heme molecule oxidised and NADPH or NADH (albeit only in vitro) as reducing equivalents (Chung et al., 2009; Ryter and Choi, 2009). A credible role for CO in Mtb persistence was first discovered during the biochemical and biophysical characterisations of DosS and DosT (Kumar et al., 2007; Sousa et al., 2007). CO was

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subsequently shown to induce the complete Mtb Dos dormancy regulon and was demonstrated to be produced in the lungs of Mtb-infected mice (Kumar et al., 2008; Shiloh et al., 2008). It is known that hypoxia, NO (Voskuil et al., 2003) and CO (Davidge et al., 2009) each individually inhibits respiration and that combinations of these gases may even act synergistically to do the same. CO2 has also been shown to be extremely unfavourable for in vitro survival (Dubos, 1953) and survival of Mtb in the cavities of TB patients (Haapanen et al., 1959). On the other hand, low concentrations of CO2 have been shown to enhance survival of BCG under hypoxic conditions (Florczyk et al., 2003). Differences exist in ventilation and perfusion of various areas of the lung, as do the degree of blood oxygenation and pO2. In the upper lung regions, higher O2 tension is present (Rich and Follis, 1942; Rasmussen, 1957; Riley, 1957; West, 1977). As proposed earlier (Kumar et al., 2008), Mtb likely encounters gradients of gases (e.g. NO, CO, O2 or CO2) during the course of infection, which contribute towards producing varying microenvironments and distinct granulomatous populations. These microenvironments may include caseous, fibrotic and non-necrotising granulomas all occurring within the same lung (Barry et al., 2009). Thus, it is reasonable to conclude that if anaerobic granulomas exist, Mtb will not be able to survive in them. However, hypoxic (as opposed to anaerobic) granulomas provide bacilli with the capacity to respire, albeit at a low metabolic state, allowing survival and promoting antimicrobial tolerance (see Section 6.2.4.1 on the role of NO3
/NO2
in maintaining Mtb viability under anaerobic conditions). Mtb has an extraordinary capacity to persist for decades in the human lung despite conditions that should be detrimental to its survival. Low levels of O2 in TB granulomas and the presence of diatomic host gases inhibit respiration, effects which profoundly influence the metabolic state of the tubercle bacillus. Detailed knowledge of the metabolic state of Mtb within the granulomatous host environment is lacking and severely hampers our understanding of the mechanism(s) of Mtb persistence.

5. REDUCTIVE SINKS IN MICROBES 5.1. Fermentation 5.1.1. Saccharomyces cerevisiae A vast literature shows that mycobacterial species are incapable of fermentation. Nonetheless, since Mtb is exposed to a hypoxic environment in vivo

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and encounters host defence molecules such as NO and CO, which inhibit respiration, it is important to identify the metabolic pathways involved in the reoxidation of reducing equivalents in order to maintain redox balance. It therefore becomes imperative to look at examples provided by other model organisms to identify parallels of such metabolic events. The lower eukaryote Saccharomyces cerevisiae (S. cerevisiae) is a particularly attractive model organism since, as is the case for eukaryotes, its cytoplasm is highly reduced [2GSH/GSSG = 70–190:1 (Grant et al., 1998; Garrido and Grant, 2002)], whereas the endoplasmic reticulum (ER), where oxidative protein folding occurs, is oxidised [2GSH/GSSG = 1–3:1 (Hwang et al., 1992)]. Compartmentalisation of these oxido-reductive events is essential for proper cellular functioning of the organism, and more specifically, to protect intracellular components from non-specific oxidation or reduction. In order to dissect the oxido-reductive events associated with these functionally distinct cellular compartments, the strong reducing agent dithiothreitol (DTT) was used as a molecular tool to promote reductive stress. Comprehensive microarray analysis showed that S. cerevisiae treated with DTT generates a transcriptional response that is distinct from other stresses such as hyperosmotic stress, starvation and heat shock (Gasch et al., 2000). DTT exposure also induces the upregulation of protein disulfide isomerase, protein folding chaperones localised to the ER and other genes that respond to changes in the cellular redox potential. Furthermore, the upregulation of genes involved in cell wall synthesis and signaling pathways responsive to cell wall damage was noted and led the investigators to conclude that cell wall defects eventually initiate the environmental stress response (ESR). Further investigations using DTT exposure showed that the loss of S. cerevisiae genes encoding two thioredoxins (trx1, trx2) causes sensitivity to DTT (Trotter and Grant, 2002). Since thioredoxins are small oxidoreductases that typically protect cells against oxidative stress, the observed sensitivity to DTT was intriguing. Given that thioredoxin loss was previously shown to cause an imbalance in the 2GSH:GSSG ratio (Muller, 1996; Garrido and Grant, 2002), the findings suggest that the yeast cellular redox machinery requires precise regulation to protect against oxidative and reductive stress, and that thioredoxins help maintain redox homeostasis in response to both oxidative and reductive stress. The mode of action of the yeast redox machinery is functionally distinct from that of bacteria, since loss of bacterial thioredoxin results in sensitivity to the thiol-oxidising agent diamide (Ritz et al., 2000), whereas loss of trx1 and trx2 in S. cerevisiae produced diamide resistance (Trotter and Grant, 2002). Thioredoxins, glutaredoxins and GSH are thermodynamically linked, since oxidised thioredoxins are reduced by NADPH and thioredoxin reductase, whereas oxidised glutaredoxins are reduced by GSH and NADPH.

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In continuation of the above studies (Rand and Grant, 2006), S. cerevisiae was used in a screen to identify mutants sensitive to DTT. A large number of mutations were found that affected gene expression, metabolism and components of the secretory pathway, including a mutation that resulted in the loss of TSA1, encoding a peroxiredoxin. It was observed that TSA1 mutants accumulate aggregated ribosomal proteins, thus impairing translation initiation (Rand and Grant, 2006). A complementary and physiologically relevant approach to studying DTT exposure included analysis of a glycerol3-phosphate dehydrogenase (GPD2) mutant shown to have altered intracellular NADH levels. Excessive NADH levels are thought to be the underlying reason for the attenuated anaerobic growth of S. cerevisiae lacking GPD2 (Ansell et al., 1997). During S. cerevisiae fermentation, NADH/NAD+ redox balance is maintained when acetaldehyde is reduced to ethanol, which is redox neutral. During anaerobic growth, glycerol is following ethanol and CO2, the most abundant byproduct, which is produced by the NADH-mediated reduction of dihydroxyacetone phosphate (DHAP) to glycerol-3-phosphate, followed by dephosphorylation (van Dijken and Scheffers, 1986). Nonetheless, the reduction of NAD+ to NADH occurs via metabolite and biomass synthesis and NADH in turn must be re-oxidised to NAD+ to maintain redox balance. Thus, intrinsically, glycerol functions as a redox sink for anaerobic growth of S. cerevisiae, and as such, glycerol must be continuously produced to maintain redox balance (van Dijken and Scheffers, 1986). S. cerevisiae Dgpd1Dgpd2 cells are unable to synthesise glycerol under anaerobic conditions (Ansell et al., 1997) and consequently cannot re-oxidise NADH, which leads to NADH accumulation and growth arrest. However, provision of exogenous acetoin or acetaldehyde to the media (Ansell et al., 1997) as electron acceptors restored redox balance and growth. 2D-PAGE analyses of anaerobically grown S. cerevisiae Dgpd2 showed increased expression of Tdh1p, the minor isoform of G3PD, an effect which could be reversed by the addition of acetoin. Since deletion of TDH1 improved anaerobic growth of S. cerevisiae Dgpd2, it was speculated that TDH1 functions as a reporter for intracellular NADH reductive stress (Valadi et al., 2004). The requirement of glycerol formation as a redox sink for NADH in anaerobically cultured S. cerevisiae was abrogated by the NADHdependent reduction of acetic acid to ethanol (Medina et al., 2010). In this study, the E. coli mhpF gene, encoding the acetylating NAD+-dependent acetaldehyde dehydrogenase, was expressed in S. cerevisiae Dgpd1Dgpd2 and was able to restore growth of the mutant under anaerobic conditions when the medium was supplemented with acetate as an electron acceptor (Medina et al., 2010). An alternative approach for the reoxidation of

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NADH, based upon the absence of transhydrogenase activity (NADH + NADP+ $ NAD+ + NADPH) in yeasts (van Dijken and Scheffers, 1986) was attempted, wherein the ultimate goal was to heterologously introduce a different pathway for reoxidation of NADH in yeast when glycerol synthesis was impaired. This would lead to an increased ethanol production under aerobic and anaerobic conditions. Unfortunately, the system appeared to be more complex than anticipated, and attempts to introduce an alternative NADH oxidation pathway by expressing the transhydrogenase of Azotobacter vinelandii in S. cerevisiae Dgpd1Dgpd2 was unsuccessful (Nissen et al., 2000). Formate is a particularly important NADH-generating substrate utilised by S. cerevisiae under aerobic conditions, since CO2 [product of the formate dehydrogenase (FDH) reaction] does not accumulate in solution. In a metabolic engineering approach, formate, which cannot act as a carbon source for biomass formation, was used to increase glycerol production under anaerobic conditions (Geertman et al., 2006). However, formate oxidation was shown to be incomplete. Since low intracellular NAD+ concentrations negatively affect the in vivo Km of FDH for formate, GPD2 was overexpressed to re-oxidise NADH. The concurrent overexpression of FDH1 with GPD2 demonstrated a synergistic effect that resulted in consumption of 70% of the supplied formate (Geertman et al., 2006). In sum, overproduction of NAD(P)H has to be balanced by NAD(P)Hconsuming pathways. In order to maintain redox balance, yeast may secrete ethanol, polyalcohols, monocarboxylic acids and di- and tricarboxylic acids. 5.1.2. Escherichia coli Bacteria have evolved a variety of mechanisms to balance the rate of oxidation and reduction. Reoxidation of NAD(P)H requires electron acceptors acquired from the environment (external acceptors) or they may be generated intracellularly. When electron transfer occurs in a membrane-bound process, NADH oxidation may be linked to respiration (either aerobic or anaerobic) (de Graef et al., 1999), whereas electron transfer that occurs in the cytosol, aka fermentation, also re-oxidises NADH to generate NAD+ (Wolfe, 2005). Pyruvate catabolism is the major switch point between the respiratory and fermentative responses. In the absence of O2, energy must be supplied by either anaerobic respiration coupled to electron acceptors such as nitrate (NO3
) and fumarate, or by fermentation (Gray et al., 1966; Clark, 1989). For example, facultative anaerobes such as E. coli can use terminal electron acceptors such as fumarate, NO3
or DMSO in the process of anaerobic

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respiration. Alternatively, if no terminal electron acceptor is available, E. coli switches to fermentation, which makes use of endogenous organic compounds as electron acceptors to generate soluble products including acetate, ethanol, lactate, formate and succinate and gaseous products such as H2 and CO2 (Clark, 1989). Notably, because redox balance has to be maintained, the ratio of these products is influenced by the number of reducing equivalents generated during breakdown of the substrate. In the ethanolic fermentation pathway, ethanol, propionic acid and CO2 are the end products whose formation is coupled with the conversion of NADH to NAD+. In the case of mixed acid fermentation carried out by E. coli and members of the genera Salmonella and Shigella, pyruvate is converted into ethanol, acetate, succinate, formate, molecular H2, lactate and CO2 (Gray et al., 1966; Wolfe, 2005). During synthesis of these products, NADH is re-oxidised to NAD+, and acetate production is accompanied by ATP formation via substrate-level phosphorylation. In addition to carbohydrates, amino acids such as arginine can be fermented by Clostridium, Streptococcus and Mycoplasma spp. to ornithine, CO2 and NH3. Clostridia can ferment multiple amino acids through the Stickland reaction in which one amino acid functions as an electron donor and the other as an electron acceptor, allowing regeneration of reducing equivalents (Atlas, 1996). The intracellular redox state of E. coli as indicated by the NADH/NAD+ ratio is strongly influenced by the availability and nature of the external electron acceptors present in the extracellular environment (de Graef et al., 1999). Both fumarate and nitrate were shown to be electron acceptors capable of functioning as effective reductive (NADH) sinks. The highest NADH/NAD+ ratios occurred during fermentation followed by fumarate respiration and nitrite respiration. Furthermore, in examining the relationship between dissolved O2 tension (DOT) and the intracellular levels of reducing equivalents, the NADH/NAD+ redox ratio was found to inversely correlate with DOT; the lower the DOT, the higher the ratio, such that the most anaerobic condition examined (DOT of 0.1%) was associated with the largest ratio (de Graef et al., 1999). As described earlier, acetogenesis, or the excretion of acetic acid into the culture medium, can either occur as a result of growth involving a high rate of glucose consumption in the presence of ample O2 (Crabtree effect), or during growth under anaerobic conditions when the TCA cycle is not operating. Although acetic acid production can be viewed as a mechanism to reduce NAD(P)H accumulation, it subsequently leads to the production of ATP, whereas D-lactate, succinate, ethanol, formate and CO2 are the excreted products that function as sinks for accumulating reducing equivalents to maintain redox balance (Wolfe, 2005).

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To examine the role of the NADH/NAD+ ratio in acetic acid overflow metabolism, the redox ratio in E. coli was modulated by overexpressing the Streptococcus pneumoniae nox gene (encoding water-forming NADH oxidase), which decouples NADH oxidation from respiration (Vemuri et al., 2006). The data demonstrated that an increase in oxidation of excess NADH led to decreased acetate formation and biomass yield, and increased the glucose consumption rate by 50%. As expected, the redox ratio was always greater for nox
bacteria than for the nox+ strain. However, acetate formation for both strains occurred at an identical NADH/NAD+ ratio of 0.06, thereby establishing a relationship between the redox state of the cell and overflow metabolism (Vemuri et al., 2006). Conversely, the effect of increasing intracellular NADH was studied by substituting the native cofactor-independent FDH with an NAD+-dependent FDH from Candida boidinii (Berrios-Rivera et al., 2002a,b,c). Overexpression of the yeast FDH in E. coli under anaerobic conditions caused an increase in NADH and favoured the production of more reduced metabolites such as ethanol, which also generated a three- to fourfold increase in the ethanol/acetate ratio (Berrios-Rivera et al., 2002b). In fact, the increased availability of NADH induced a shift towards fermentation in the presence of O2 evident by the production of lactate, ethanol and succinate, all metabolites typically produced during anaerobic fermentation (Berrios-Rivera et al., 2002a,b).

5.2. Polymer Deposition 5.2.1. Polyhydroxyalkonate (PHA), Poly-b-Hydroxybutyrate (PHB) and Triacylglycerol (TAG) Biosynthesis of polyketide or lipid-like molecules in response to a change in intracellular redox balance may be a general compensatory mechanism found in many bacteria. In fact, polyhydroxyalkonate (PHA) and polyb-hydroxybutyrate (PHB) are accumulated by diverse bacteria as carbon and reductive-power storage molecules (Encarnacion et al., 1995; Cevallos et al., 1996). As in the case of the TCA cycle, carbon flow through the pathways necessary for PHB or PHA accumulation is greatly influenced by growth and environmental conditions (e.g. O2 concentration) (Senior and Dawes, 1973). Biosynthesis of PHA, a storage lipid, in Azotobacter beijerinckii was shown to be regulated by O2 concentration and carbon source (Senior and

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Dawes, 1973). The central regulator of PHA production is the flux of acetylCoA, which may be oxidised via the TCA cycle or can serve as a precursor for PHA synthesis depending upon oxygen concentration. Under oxygen limitation, when the NADH/NAD+ ratio increases, the activities of the TCA cycle enzymes, citrate synthase and isocitrate dehydrogenase are inhibited by NADH and as a consequence, acetyl-CoA could no longer enters the TCA cycle. Instead, acetyl-CoA is converted to acetoacetyl-CoA by 3ketothiolase, the first enzyme in the PHA biosynthesis pathway. Based on these findings, it was proposed that PHA serves not only as a reserve carbon and energy source, but also as a reductant sink, similar to what was suggested for TAG (Senior and Dawes, 1973). Direct evidence in support of PHA in regulating intracellular redox balance was provided by Page and Knosp (1989), using an NADH oxidase-deficient strain of A. vinelandii. This strain is unable to re-oxidise NADH via oxygen-dependent respiration and instead accumulates large amounts of PHA, as a mechanism for the disposal of excess reductants (Page and Knosp, 1989). Species belonging to the genera Rhizobium, Bradyrhizobium and Azorhizobium synthesise PHB during symbiosis and in free-living state (Encarnacion et al., 1995; Cevallos et al., 1996). Interestingly, even though Rhizobium spp. are strict aerobes that are well adapted to survive microaerophilically, a fermentative response was also described. Besides serving as a sink of reductive power, PHB is also a fermentative product, which is secreted like other organic acids and amino acids (Encarnacion et al., 1995). Studies in Rhizobium etli have shown that a PHB mutant excreted significantly more pyruvate, fumarate, lactate, acetate and b-hydroxybutyrate compared to the wild-type (wt) strain (Cevallos et al., 1996). This data, together with the observation that the NAD+/NADH ratio is much reduced, indicates that the oxidative capacity of the organism is significantly decreased because of the absence of a sink for reductive power (Cevallos et al., 1996). TAG is a water-insoluble triester of glycerol with FA and an excellent reserve substrate because of the reduced COS relative to carbohydrates or proteins. Because of these properties, it yields significantly more energy when oxidised (Alvarez and Steinbuchel, 2002; Waltermann et al., 2007). b-Oxidation of the FA chains of TAG generates large quantities of reducing equivalents, which require subsequent oxidation. This requirement might be the reason why TAG-producing bacteria are all aerobes. Consistent with this notion, it has been suggested that, in actinomycetes, TAG could serve as a sink for excess reductants accumulated in the absence of terminal electron acceptors (Alvarez and Steinbuchel, 2002). Rhodococcus ruber is capable of accumulating both TAG and PHA and

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disruption of PHA biosynthesis leads to increased accumulation of TAG, suggesting a metabolic link between these two triesters (Alvarez et al., 2000; Alvarez and Steinbuchel, 2002). Studies in Rhodococcus opacus PD630 suggested that TAG production can be promoted by culturing bacteria under limited aeration (Alvarez and Steinbuchel, 2002). Similarly, increased expression of Mtb tgs1 leading to TAG accumulation occurs under hypoxic conditions and during exposure to NO and CO (Sherman et al., 2001; Ohno et al., 2003; Voskuil et al., 2003; Kumar et al., 2008). Consistent with the biological function of TAG and PHA, it appears that inhibition of respiration by NO or lack of O2 as terminal electron acceptor leads to an increased amount of reducing equivalents, which can be dissipated via TAG, PHA or PHB anabolism. Recent reports provide important mechanistic links between reductive stress, polyketide and TAG anabolism (see Section 6.2.4.2). It has been shown that during persistence Mtb accumulates NAD(P)H, confirming the physiological presence of reductive stress in Mtb pathogenesis (Boshoff et al., 2008). Furthermore, in the mouse model for TB (Jain et al., 2007) and in the in vitro model for dormancy (Daniel et al., 2004), Mtb induces production of complex polyketides, such as PDIM and SL-1, and the storage lipid TAG respectively. Since accumulation of NADH can eventually lead to oxidative stress by auto-oxidation and reduction of O2 to generate O2
, it has been proposed that polyketide and TAG anabolism could serve as efficient reductant disposal mechanisms utilised by Mtb to alleviate reductive stress for long-term persistence (Singh et al., 2009). TAG is metabolised by Mtb upon reactivation from the Wayne model of in vitro dormancy (Deb et al., 2006), indicating a possible role for TAG in emergence from a persistent state. *

5.3. Nitrate Reductase Under normal growing conditions, aerobic respiration in bacteria is primarily required to generate a proton motive force for ATP synthesis. Several studies suggest that inhibition of aerobic respiration, due to the lack of oxygen or exposure to NO, results in accumulation of reducing equivalents and depletion of ATP (de Graef et al., 1999; San et al., 2002; Berrios-Rivera et al., 2004; Sanchez et al., 2005; Vemuri et al., 2006; Husain et al., 2008). It has been shown that in addition to hypoxia and NO, oxidative metabolism of highly reduced carbon substrates (e.g. palmitate, caproate, butyrate, oleate) also results in intracellular reductive stress even in the presence of oxygen (Alam and Clark, 1989; Clark, 1989; Sears et al., 2000; Berrios-Rivera et al., 2004; Lin et al., 2005; Sanchez et al., 2005). These studies suggest that under

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conditions of reductive stress, respiration is not coupled to proton translocation; rather, disposal of excess reductants from cells without ATP generation may be the dominant function of respiration (Sears et al., 2000). For example, in Paracoccus pantotrophus, electrons flow from ubiquinol to periplasmic nitrate reductase (Nap) without proton translocation during growth on reduced carbon substrates such as acetate and butyrate (Ellington et al., 2002). P. pantotrophus is capable of both aerobic and anaerobic respiration on NO3
in the presence of a variety of carbon sources with broad oxidation states. It has been shown that Nap is required to maintain cellular redox homeostasis by providing an alternate route for the oxidation of excess reductants generated from the oxidative metabolism of highly reduced carbon substrates (Richardson, 2000). Furthermore, demonstrating that highly reduced carbon substrates (e.g. caproate and butyrate) increase Nap activity whereas oxidised carbon substrates (e.g. succinate and malate) decrease Nap activity points to a clear correlation between Nap activity and the COS of a carbon substrate (Sears et al., 2000). More importantly, transcription of the nap operon is shown to be dependent on carbon source, implying that nap transcription responds to a change in the cellular redox state due to the metabolism of the reduced carbon substrates (Sears et al., 2000). A strict hierarchical preference for carbon sources was exhibited by P. pantotrophus. The less reduced carbon source, succinate, is preferred over acetate, which in turn is preferred over butyrate for growth. This hierarchical preference for carbon substrates is directly correlated with the expression of nap, such that nap is maximally induced during growth on butyrate followed by acetate and then succinate (Ellington et al., 2002). As was demonstrated by the low growth rate, P. pantotrophus cells grown on butyrate were ‘strained’ as compared to succinate. This suggests a growth-restricting effect of reduced carbon source. Consistent with cellular energetics, butyrate as a carbon substrate is more reduced than most of the sugars and thus generates an excess of toxic reducing equivalents, which must be dissipated. The excess reductants can be disposed only if there is a mechanism for uncoupling the respiratory electron flow from ATP synthesis. In P. pantotrophus, the ubiquinol-Nap nitrate reductase pathway is one such mechanism (Richardson, 2000; Sears et al., 2000). See Section 6.2.4.1 for a description of the role of Mtb nitrate reductase in reductive stress.

5.4. Phenazine Production Phenazines are redox-active heterocyclic compounds produced naturally and modified at different positions on their rings by various phenazine-

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generating bacterial species. To date, there are approximately 100 natural phenazine products that are almost exclusively produced in high levels (mg to g/L) by eubacteria (Mavrodi et al., 2006). Many members of the genus Streptomyces, which are high G + C content bacteria, also produce simple and complex phenazines (Mavrodi et al., 2006; Mentel et al., 2009; Winstanley and Fothergill, 2009). Pyocyanin (PYO; 5-N-methyl-1-hydroxyphenazine) was the first described phenazine and is produced by Pseudomonas aeruginosa. PYO naturally occurs as a zwitterion that has hydrophobic and hydrophilic regions, which can easily penetrate cytoplasmic membranes, and can undergo cellular redox cycling in the presence of NADPH, NADH and O2 to generate O2
and H2O2 (Dietrich et al., 2008; Mentel et al., 2009). PYO and phenazine-1-carboxylic acid (another major phenazine produced by P. aeruginosa) have redox potentials of
34 mV (Friedheim and Michaelis, 1931) and
116 mV (Price-Whelan et al., 2007) respectively, and therefore can be reduced by NADH (E0’ =
320 mV). Not surprisingly, establishing that NADH can react with PYO in vitro led to the conclusion that bacteria may use PYO as a mechanism to maintain intracellular redox homoeostasis (Friedheim, 1931; Price-Whelan et al., 2006). Consistent with this, recent studies have shown that PYO can directly activate SoxR, a Fe–S cluster and regulatory protein that is typically upregulated in response to oxidative stress (Dietrich et al., 2006). It was also suggested that excreted phenazines reduce Fe3+ to the more soluble Fe2+ form that can be taken up by siderophores (Hernandez et al., 2004). Thus, it is clear that phenazines have the potential to generate substantial oxidative stress on cells. Because phenazines have low mid-point redox potentials, it has been suggested that they can be directly reduced by NADH or GSH (Hernandez and Newman, 2001). Several groups observed that both synthetic and natural phenazines are reduced by prokaryotes, but in most of these studies the physiological effect of this reduction was not examined. However, Methanobacterium mazei Go1, an archeon producing methanophenazine (a phenazine derivative), has been shown to utilise phenazines instead of quinones in the ETC in order to generate ATP (Deppenmeier, 2002). Thus, in M. mazei, phenazine production is not only crucial to energy metabolism, but also for re-oxidising NADH (Deppenmeier, 2004). This raises the possibility that phenazines may be similarly involved in dissipating reductive stress in pseudomonads. In an interesting report examining the NAD+ and NADH concentrations in Clostridium welchii, Klebsiella aerogenes, E. coli, Staphylococcus albus and P. aeruginosa (Wimpenny and Firth, 1972), it was demonstrated that all the species, with the exception of P. aeruginosa, have NADH/NAD+ *

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ratios of <1. In stark contrast, P. aeruginosa has a NADH/NAD+ ratio of 1.33 (Wimpenny and Firth, 1972), which suggests that this organism has evolved mechanisms to maintain viability despite experiencing significant reductive stress. In a subsequent genetic study examining NADH/NAD+ in a P. aeruginosa mutant defective in PYO generation, it was shown that the mutant accumulated more NADH in stationary phase than the wt strain. It was proposed that O2 limitation increases NADH levels that leads to reduction of PYO, which upon normoxic conditions is oxidised via electron transfer to O2 (Price-Whelan et al., 2007). Lastly, in order to maintain redox balance, it was suggested that O2
generated by PYO or a PYO radical itself may target the lipoamide cofactor of pyruvate dehydrogenase, leading to the excretion of pyruvate (Price-Whelan et al., 2007). Because of the recognised involvement of phenazines in quorum sensing and biofilm formation in P. aeroginosa (Hernandez and Newman, 2001; Alvarez-Ortega and Harwood, 2007; Dietrich et al., 2008; Winstanley and Fothergill, 2009), the latter has become a popular model organism for studying phenazines. Phenazines have been shown to regulate many aspects of pulmonary infection, ranging from controlling the establishment of acute and chronic P. aeroginosa lung infections, influencing neutrophil influx into lung tissue, causing oxidant formation in human airway epithelial cells and disturbing the host cell’s redox homeostasis by generating ROS (Linnane et al., 1998; Worlitzsch et al., 2002; Alvarez-Ortega and Harwood, 2007; Winstanley and Fothergill, 2009). It has also been shown that P. aeruginosa is located in hypoxic mucopurulent debris in airway lumens of CF patients (Worlitzsch et al., 2002). In order to determine the pO2 in vivo, a Clark type O2 probe was fixed to the tip of the bronchoscope and inserted into the upper lobar bronchi of CF patients. The pO2 in the bronchial lumen was 180 mm Hg whereas it was 2.5 mm Hg in the mucopurulent material, thereby providing conclusive evidence of a hypoxic environment occupied by P. aeruginosa. Since the diffusion rate of O2 through mucus is slow and because live P. aeruginosa cells consume available O2 within the mucus, cells at the base of the mucopurulent material become hypoxic (Worlitzsch et al., 2002). In the absence of a O2 as a terminal electron acceptor, P. aeruginosa cells within this hypoxic environment experience reductive stress [accumulation of NAD(P)H] and depletion of ATP. Under these conditions, hypoxic cells could utilise excreted redox active phenazines as electron acceptors in lieu of O2 to re-oxidise accumulated NADH and generate ATP, thereby dissipating toxic reductants (Worlitzsch et al., 2002; Alvarez-Ortega and Harwood, 2007). Despite the advances in understanding the redox biology of phenazines, the precise mechanism of phenazine reduction remains a matter of debate. *

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Phenazines have the potential to directly associate with the respiratory chain to either couple their reduction to ATP generation or dissipate reductive stress (Hernandez and Newman, 2001; Mavrodi et al., 2006; Price-Whelan et al., 2006, 2007).

5.5. Hydrogenases Hydrogenases (H2ases) are redox metalloenzymes found throughout the eukaryotic and prokaryotic genera, the majority of which contain an iron– sulphur (Fe–S) cluster or alternatively two metal atoms (two Fe atoms or an Ni–Fe combination) (Vignais et al., 2001; Jenney and Adams, 2008). H2ases catalyse the reversible conversion/oxidation of hydrogen gas (Nandi and Sengupta, 1998; Meyer, 2007; Jenney and Adams, 2008): H2 $ 2H+ + 2e
Membrane-bound hydrogenases split the protons from H2, thus creating a proton gradient across the cytoplasmic membrane. The electrons (e
) produced in the H2 oxidation process are transferred to electron carriers in the bacterial membrane and ultimately to terminal electron acceptors such as O2 (aerobic respiration) or fumarate, NO3
, SO4
2 and CO2 (anaerobic respiration) (Vignais et al., 2001; Vignais and Colbeau, 2004). In addition, some bacteria can use the protons as oxidants to dispose of excess reducing power, thereby reoxidising their coenzymes in the process (Cammack et al., 2001; Vignais et al., 2001). Since the oxidation of H2 is reversible, the H2ase will produce H2 in the presence of an electron donor and will act as a H2 uptake enzyme in the presence of an electron acceptor. H2ase activity is modulated by numerous regulatory pathways and responds to changes in the valance state of the metal cofactors, O2 levels and pH (Vignais and Colbeau, 2004). H2 is a high-energy reductant that is present in the mucus layer of the mouse stomach at 43 mM, in the liver at 50 mM (Olson and Maier, 2002; Maier et al., 2003) and spreads throughout host tissues by diffusion, or alternatively is carried through the bloodstream to organs such as the lungs (Levitt, 1969). H2 has a likely role to play in microbial pathogenesis as it was recently demonstrated that H2 is highly effective in reducing OH and alleviating OH -induced cytotoxicity without affecting other ROS (Ohsawa et al., 2007). H2ases can be classified into three categories based on the composition of their metal centres: (i) [NiFe]-hydrogenases, (ii) [FeFe]-hydrogenases and (iii) the Fe–S cluster-free hydrogenases (Nandi and Sengupta, 1998; Vignais and Colbeau, 2004; Burgdorf et al., 2005; Jenney and Adams, 2008). Since the enzymatic activity of uptake H2ases can increase under anaerobic conditions *

*

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(reductive activation) (Maier et al., 2003), it has implications for a wide range of pathogens. For example, in a seminal study, a role for H2 in Helicobacter pathogenesis was reported (Olson and Maier, 2002). It was shown that H2 produced by the gastric flora in mice functions as a respiratory substrate for Helicobacter pylori and substantially increases its ability to colonise the stomach (Olson and Maier, 2002). On the other hand, Salmonella H2ases (e.g. Hyd) may oxidise H2 with O2 as the terminal electron acceptor with the intention of conserving energy during different stages of host infection (Zbell et al., 2008). Genetic studies in Salmonella have shown that H2 can be generated and oxidised by the same organism (Zbell and Maier, 2009). Using H2 as an energy source in vivo by bacterial pathogens is not unique and has been reported previously (Olson and Maier, 2002; Zbell et al., 2007; da Silva et al., 2008). Since the TCA cycle is repressed under anaerobic conditions, fermentative growing bacteria must oxidise reducing equivalents such as NADH to ensure continuous conversion of the substrate. Klebsiella pneumoniae has evolved a clever mechanism by which the bacteria gain limited quantities of the reducing equivalent NADPH from the oxidation of citrate (PfenningerLi and Dimroth, 1992), but because of the downregulation of the TCA cycle, sufficient quantities of NADH is lacking. K. pneumoniae solves this problem by generating NADPH via the oxidation of H2 during citrate fermentation by a H2:NAD(P)+ oxidoreductase (hydrogenase). Thus, reducing equivalents are provided for the production of biomass (Steuber et al., 1999). Pyrococcus furiosus employs a different strategy. This organism can use a range of carbohydrates, converting them to acetate, CO2, H2 and, if elemental sulphur (S0) is present, H2S (Ma et al., 1993). An important characteristic is that ferredoxin serves as electron acceptor during glycolysis and that no NAD(P)+ is generated. Subsequently, it was found that a H2-evolving membrane-bound H2ase couples the oxidation of ferredoxin and the reduction of 2H+ to H2 production. Next, a membrane-bound oxidase oxidises ferredoxin and reduces NADP, which is used by a NAD(P)H S0 oxidoreductase to reduce S0 to H2S. Thus, P. furiosus disposes of excess reductant using protons or S0 as electron acceptors to yield H2S (Ma et al., 1993; Schut et al., 2007; Jenney and Adams, 2008). The production of H2 via hydrogenases is a specific mechanism to dispose of surplus reducing equivalents (Nandi and Sengupta, 1998). A recent study suggested a feasible and physiologically relevant role for mycobacterial hydrogenases in scavenging H2 as an energy source or as a redox sink (Berney and Cook, 2010). Microarray analysis of hypoxic, carbonlimited continuous cultures of Mycobacterium smegmatis identified a number of hydrogenases that are differentially regulated under energy- or

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oxygen-limiting conditions. Since it remains unclear how mycobacteria recycle reducing equivalents under hypoxic conditions lacking exogenous electron acceptors, it was hypothesised that (i) mycobacteria switch to NAD+/ NADH-independent enzymes and use ferredoxins as electron carriers, which is typical for anaerobic bacteria, and that (ii) hydrogenases either oxidise H2 or produce H2, or perform both. The investigators cited the significant upregulation of many ferredoxin-reducing and oxidising enzymes identified in their study, as well as the results of another study identifying an anaerobic type a-ketoglutarate:ferredoxin oxidoreductase (KOR), which upon disruption requires CO2 for growth (Baughn et al., 2009), to provide credence to their hypotheses (Berney and Cook, 2010). In order to investigate the role of H2ases under conditions of energy limitation, a highly upregulated putative hydrogen:quinone oxidoreductase of Msm was disrupted. The Msm mutant showed a 20% reduction in biomass, thus demonstrating the essential role of this particular H2ase in adapting to energy limitation. Finally, based on the Msm studies, it was argued that acidic, hypoxic and lipid-rich environments (which contribute to a highly reductive intracellular milieu) may require H2ases to function as reductive sinks, or alternatively, that H2 could serve as an energy source for Mtb (Berney and Cook, 2010). The role of H2ases in Mtb pathogenesis is a highly relevant, albeit unexplored area of investigation.

5.6. The Reverse TCA (rTCA) Cycle The TCA cycle is widely distributed in aerobic microorganisms and is an energy acquisition pathway that is exergonic (spontaneous). Although this cycle only operates under aerobic conditions, it is also present in anaerobes (Srinivasan and Morowitz, 2006). The reverse TCA (rTCA) cycle is generally regarded as the phylogenetic origin of the TCA cycle and uses CO2 as the key source for carbon fixation. The rTCA cycle is, in fact, a reversal of the oxidative TCA cycle and is an endergonic (unfavourable, non-spontaneous) anabolic pathway that requires reducing equivalents such as NADH, NADPH and FADH2 to complete the cycle (Hugler et al., 2005; Srinivasan and Morowitz, 2006; Aoshima, 2007; Ikeda et al., 2010). The end result is the fixation of two molecules of CO2 and the production of one molecule of acetyl-CoA, which is reductively carboxylated to pyruvate from which many central metabolites can be formed. Three enzymes allow the TCA cycle to operate in the reverse direction and their activity is indicative of a functioning rTCA cycle (Hugler et al., 2005; Srinivasan and Morowitz, 2006). These enzymes are ATP citrate lyase, 2-oxoglutarate:ferredoxin oxidoreductase

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(OGFO) and FRD, which catalyse the ATP-dependent cleavage of citrate to acetyl-CoA and oxaloacetate, the carboxylation of succinyl-CoA to 2oxoglutarate and the reduction of fumarate to form succinate, respectively. Having these three enzymes to drive the rTCA cycle provides pathogens with the flexibility to effectively respond to diverse environments in order to survive in their physiological niche (Hugler et al., 2005; Srinivasan and Morowitz, 2006; Aoshima, 2007). S. cerevisiae contains two FRDs that use FADH2, FMNH2 or reduced riboflavin as electron donors to irreversibly reduce fumarate to succinate. It has been suggested that these FRDs are involved in maintaining redox homeostasis during anaerobiosis (Enomoto et al., 2002; Camarasa et al., 2007). The oxidative and reductive branches of the TCA cycle can also operate under anaerobic fermentation. For example, during yeast fermentation, if aspartate is the nitrogen source, succinate can be generated via the reductive branch. On the other hand, if glutamate is the nitrogen source, succinate can be made via the oxidative branch (Srinivasan and Morowitz, 2006). Thus, it is clear that depending on the environmental conditions such as O2 concentration and energy source availability, the rTCA cycle can operate in either a reductive or oxidative direction. Since Mtb resides in a hypoxic, presumably nutrient-starved environment, it was speculated that these environmental conditions may severely restrict the ability of the organism to re-oxidise reducing equivalents to maintain redox homeostasis (Boshoff and Barry, 2005; Srinivasan and Morowitz, 2006; Leistikow et al., 2010), which would eventually lead to a redox imbalance and death. Since tubercle lesions generate substantial quantities of CO2 (Haapanen et al., 1959), the rTCA cycle, with its capacity to fix CO2, may be an effective mechanism to dissipate reducing equivalents generated by hypoxic or microaerophilic conditions and host gases (NO or CO) that inhibit Mtb respiration or by the catabolism of highly reduced in vivo carbon sources such as FA. Intriguingly, Mtb appears to lack a CoA-dependent KDH because the corresponding enzymatic activity was absent in crude cellular extracts (Tian et al., 2005). This finding suggests that Mtb might operate separate oxidative and reductive TCA half-cycles. Therefore, the study proposed that the oxidative branch (that produces a-ketoglutarate [KG] and glutamate) and reductive branch (that produces succinate) may be linked by a-ketoglutarate decarboxylase (KGD) and succinic semialdehyde dehydrogenase (SSADH) to generate succinate from KG via succinic semialdehyde (SSA) (Tian et al., 2005). In conflict with the above interpretation are the results of an independent study wherein an anaerobic type CoA-dependent KG dehydrogenase activity was assayed in Mtb (Baughn et al., 2009). An interesting

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finding was that this KOR is highly stable under aerobic conditions. KOR is expendable for growth when the glyoxylate shunt is non-functional whereas KGD is critical for this bypass. Collectively, these findings point to a pathway that operates concomitantly with b-oxidation (KOR-dependent) and another pathway that functions in the absence of b-oxidation (KGDdependent). Intriguingly, an Mtb korAB mutant strain required substantial amounts of CO2 for growth, suggesting that the KOR-dependent decarboxylation of KG is a valuable source of CO2 in Mtb metabolism (Baughn et al., 2009). This finding may have important physiological implications in vivo as alveolar air, blood and interstitial fluids contain 5–6% CO2 (Florczyk et al., 2003). Furthermore, CO2 preserves microaerophilic growth of Mycobacterium bovis BCG and prevents growth inhibition when O2 is rapidly removed (Florczyk et al., 2003), suggesting a role of CO2 in mycobacterial persistence. Despite differences in mechanisms proposed (Tian et al., 2005; Baughn et al., 2009), the production of succinate under lowoxygen conditions via the rTCA cycle is a feasible survival mechanism under this physiological condition. In agreement with this, FRD was recently speculated to be involved in survival and ATP production during hypoxia (Rao et al., 2008).

5.7. Carbon Monoxide (CO) Dehydrogenase (CODH) Carboxydotrophic microbes are aerobic organisms that use CO as the sole source of carbon and energy during chemolithoautotrophic growth. These CO-utilising microorganisms use the enzyme CODH to reversibly oxidise CO according to the following reaction (Ferry, 1995; Ragsdale, 2004; Seravalli and Ragsdale, 2008): CO + H2O $ CO2 + 2H+ + 2e
CODH enzymes are divided into two classes according to metal or cofactor content or metabolic role and catalytic activity. Aerobic carboxydotrophic bacteria uses a Cu-, Fe- and Mo-containing flavoenzyme, whereas anaerobic and archaebacteria use an oxygen-sensitive Ni- and Fe-containing CODH (Ragsdale, 2004; Jeoung and Dobbek, 2007; Oelgeschlager and Rother, 2008). Genes encoding aerobic CODH are usually denoted as carbon monoxide oxidases (cox). CODH oxidises CO to generate CO2, which is then fixed into biomass via the rTCA cycle, the Calvin–Benson–Bassham cycle, the 3-hydroxypropionate cycle or the Wood–Ljungdahl pathway (Ragsdale, 2004; Seravalli and Ragsdale, 2008). When CODH is coupled with acetylCoA synthase (ACS) (CODH/ACS), acetyl-CoA is generated from CO2,

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CoA and a methyl group. In this scenario, CODH/ACS reduces CO2 to CO, which supplies the carbonyl group of acetyl-CoA in the Wood–Ljungdahl pathway. Acetyl-CoA is then used in ATP production or for biomass synthesis (Oelgeschlager and Rother, 2008). Diverse respiratory events including oxygen respiration, hydrogenesis, desulphurication, acetogenesis and methanogenesis can all be coupled to CO oxidation (Oelgeschlager and Rother, 2008). These findings serve to demonstrate the range of microbes capable of using CO as an energy source. In a recent 13C-metabolic flux analysis study, (McKinlay and Harwood, 2010) it was reported that the Calvin–Benson–Bassham (Calvin) cycle plays an important role in reoxidising approximately half of the reduced cofactors generated by conversion of acetate into biomass. Thus, the Calvin cycle is an electron accepting process and helps maintain redox balance by fixing CO2 and oxidising the reduced cofactor NADH to NAD+. Phylogenetic and growth analyses (King, 2003; Song et al., 2010), and examination of crude extracts suggest the presence of CODH (Park et al., 2003) with NO dehydrogenase (NODH) activity (Park et al., 2007) in Mtb. The presence of CODH subunit homologues (Rv0373c, Rv0374c, Rv0375c) in the Mtb genome are consistent with the above observation. Furthermore, CODH can join the Calvin cycle and the rTCA cycle (see above) as a means to fix CO2 into biomass (Meyer and Schlegel, 1983; Ferry, 1995). However, since the Calvin cycle appears to be absent in Mtb (Park et al., 2003), the rTCA is a plausible mechanism for conversion of exogenous CO2 to biomass. A sub-lethal source of CO2 may be the cavities of lesions found in TB patients (Haapanen et al., 1959), which are essential for Mtb growth (Baughn et al., 2009). Thus, the rTCA cycle in Mtb may operate when encountering reductive stress generated by host FA catabolism, hypoxia, NO and CO (and other signals), in order to fix CO2 and to consume reducing equivalents. The presence of CODH and NODH activities in Mtb suggest that the tubercle bacillus may detoxify host-generated NO and/or CO in order to survive and persist in the host. The roles of CODH and NODH in Mtb pathogenesis represent an important area of research.

5.8. Other Mechanisms Under aerobic culture conditions, Staphylococcus aureus primarily excretes acetate whereas under fermenting growth conditions the organism produces L-lactate, ethanol and formate. In an elegant study, it was shown that upon NO exposure the cells exclusively produce L-lactate from both aerobically

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respiring and fermenting cells (Richardson et al., 2008). Subsequently, it was demonstrated that S. aureus exposure to NO caused an increase in reducing equivalents, which was metabolically balanced via L-lactate dehydrogenase activity to generate L-lactate. Thus, the NO-inducible L-lactate dehydrogenase dissipates reductive stress by reoxidising NADH to generate NAD+ and L-lactate (Richardson et al., 2008). In the case of Enterococcus faecalis, the regeneration of NAD+ from excess NADH was accomplished by generating extracellular a-ketoisocaproic acid, 1,1-dihydroxy-4-methyl-2-pentanone (Ward et al., 2000). Other mechanisms the microorganisms have evolved to dissipate excess reducing power include the use of the Calvin cycle as a reductive sink (Xanthobacter flavus) (Van Keulen et al., 2000), the synthesis of branched chain amino acids (Aspergillus nidulans) (Shimizu et al., 2010) and the overproduction of L-alanine dehydrogenase by non-pathogenic mycobacteria (Hutter and Dick, 1998; Feng et al., 2002) and Arthrobacter oxydans (Hashimoto and Katsumata, 1999).

6. REDOX SINKS IN MYCOBACTERIA 6.1. The Mycobacterial Intracellular Redox Environment In aerobic microbes, the production of ROS is not perfectly balanced by antioxidants. The latter dampens the effects of the former rather than eliminating them (Halliwell, 2008). If the balance is severely disturbed, a state of either oxidative or reductive stress ensues. The presence of substantial quantities of redox buffers such as glutathione or mycothiol in the microbial cytoplasm generates a reducing environment. Nonetheless, during aerobic respiration, redox couples such as NAD+/NADH exist predominantly in the oxidised, NAD+ state because of rapid electron flow to the ETC (Green and Paget, 2004). To study the redox environment of a microbial cell, it is impractical and impossible to measure the concentration of all linked redox couples. Rather, quantification of a representative redox couple can be used to infer the overall redox environment of the cell. For example, in eukaryotes and many bacteria, the glutathione disulfide–glutathione couple (GSSG/2GSH) is the major thiol-disulfide redox buffer and the redox state of this couple can be used to infer the status of the redox environment. As elegantly discussed (Schafer and Buettner, 2001), the reduction potential of GSSG/2GSH is dependent on the absolute concentration of GSH, as well as the GSSG/ 2GSH ratio. This circumstance differs from that of NAD+/NADH and

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NADP+/NADPH couples in which assessment of the absolute concentration of the individual components is not necessary and measurement of the ratio of the oxidised and reduced species of the redox couples is sufficient. Therefore, the redox state of a redox couple is defined by the half-cell reduction potential and the reducing capacity of that couple (Schafer and Buettner, 2001). As is the case for GSH, thioredoxin acts as an antioxidant by facilitating the reduction of cystine residues in proteins by cysteine thiol-disulphide exchange. GSH is typically present in millimolar concentration in bacterial and mammalian cells, whereas the concentration of thioredoxin [TrxSS/Trx (SH)2] is in the micromolar range in bacteria (Halliwell, 2008). Notably, since the Trx and GSH systems use NADPH as a reducing factor, the NADP+/ NADPH, GSSG/2GSH and TrxSS/Trx(SH)2 redox systems are not isolated systems and are thermodynamically linked to one another (Schafer and Buettner, 2001). Intriguingly, mycobacterial species do not produce glutathione; instead, they produce millimolar quantities of mycothiol (MSH) as a redox buffer. 6.1.1. Mycothiol: The Mycobacterial Redox Buffer In addition to acting as the major redox buffer in mycobacteria, the lowmolecular-weight (LMW) thiol, mycothiol, is involved in the removal of toxic compounds from the cell. Besides, another antioxidant, ergothioneine, (ERG; ERGox/ERGred), is also synthesised by mycobacteria (Genghof and Van Damme, 1964, 1968), although little is known so far about its role in these bacteria. MSH is produced in a five-step process and all but one of the enzymes responsible for its synthesis have been identified. The first step involves linking 1L-myo-inositol-1-phosphate (derived from glucose-6-phosphate) to UDP-N-acetylglucosamine to produce N-acetylglucosaminylinositol phosphate, catalysed by the glycosyltransferase encoded by mshA (Newton et al., 2003). MshA2, the MSH phosphatase whose gene is not yet identified, then dephosphorylates N-acetylglucosaminylinositol phosphate (Newton et al., 2006) followed by deacetylation by MshB, the MSH deacetylase, to produce glucosaminylinositol [1-O-(2-amino-1-deoxy-a-D-glucopyranosyl)D-myo-inositol] (Newton et al., 2000a). Cysteine is ligated to this compound via its carboxyl group in an ATP-dependent reaction catalysed by the MSH ligase, MshC (Sareen et al., 2002). The final step is the acetylation of the amino group of cysteine by MshD, the MSH acetyltransferase (Koledin et al., 2002), a reaction which serves to make mycothiol more resistant to autoxidation than free cysteine (Newton et al., 1995, 2008).

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Mutants in the mycothiol biosynthetic pathway have been isolated in M. smegmatis (Msm) and Mtb. In Msm and the Mtb strains H37Rv, Erdman and CDC1551, the loss of MshA activity results in undetectable levels of MSH and its intermediates (Newton et al., 1999, 2003; Vilcheze et al., 2008). mshB mutants of Msm and Mtb display accumulation of pathway intermediate, N-acetylglucosaminylinositol, but are still able to produce low levels of MSH, which may be attributed to an uncharacterised enzyme with a complementary deacetylase function (Buchmeier et al., 2003; Rawat et al., 2003). An mshC mutant in Msm does not produce detectable levels of MSH but has increased glucosaminylinositol levels (Rawat et al., 2002). Efforts to produce an mshC mutant of Mtb Erdman were unsuccessful and the researchers attributed this inability to the essentiality of mycothiol for Mtb survival (Sareen et al., 2003). mshD mutants of Msm and Mtb have been shown to produce decreased MSH, increased levels of its immediate precursor and two novel thiols (Newton et al., 2005; Buchmeier et al., 2006). Based on the phenotypes of various MSH mutants in response to redox stress, it is clear that MSH contributes greatly to the maintenance of redox homeostasis in mycobacteria. Increased sensitivity to oxidative stress is a common theme, and in agreement to this, the Msm mycothiol mutants have decreased survival compared to wild type after treatment with hydrogen peroxide and plumbagin, a superoxide generator (Newton et al., 1999; Rawat et al., 2002, 2007). In Mtb, the mshB and mshD mutants have increased sensitivity to cumene hydroperoxide and hydrogen peroxide respectively (Buchmeier et al., 2003, 2006). Additionally, it has been shown that the MSH mutants of Msm are more sensitive than wild type to reducing stress based on disk assays in the presence of 10 mM DTT, a reductant (Rawat et al., 2007). Antibiotic resistance is also altered in the mutants; increased resistance to ethionamide (ETH) has been reported for all Msm mutants except mshC (Rawat et al., 2003, 2007). However, the mshA and mshD Msm mutants are more resistant to isoniazid (INH) (Koledin et al., 2002; Newton et al., 2003). In Mtb, the mshA mutant has increased resistance to INH and ETH (Vilcheze et al., 2008), and INH resistance has also been reported for the mshB mutant (Buchmeier et al., 2003). The regulation of MSH biosynthesis has not yet been fully elucidated. Transcriptional regulators are located upstream of mshA and mshD in Mtb, but not a single study has been carried out to determine their role in MSH regulation (Newton et al., 2008). MSH levels are influenced by growth phase in Mtb and are increased greater than threefold in stationary phase as compared to early exponential phase (Buchmeier et al., 2006). However, it is not known what causes this effect. No direct transcriptional analysis of the genes involved in MSH biosynthesis at these different phases has been performed

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(Newton et al., 2008), but microarray analysis to identify genes whose expression levels differ in exponential and stationary phases did not identify the MSH-encoding genes (Voskuil et al., 2004). MshB activity is inhibited in the presence of MSH; thus, MSH biosynthesis may be regulated in part by feedback inhibition (Newton and Fahey, 2002). Many MSH-dependent enzymes are present in mycobacteria such as MSH disulfide reductase, or Mtr. In the course of maintaining the reducing environment of the cell, MSH becomes oxidised (MSSM) and must be re-reduced to ensure proper functioning of cellular processes. This crucial task is performed by Mtr, which uses FAD as a cofactor in a reaction that consumes NADPH (Rietveld et al., 1994; Patel and Blanchard, 1998, 1999). MSH has a role in the removal of toxic compounds such as antibiotics, via the action of mycothiol S-conjugate amidase (Mca). MSH fuses with the toxic compound and then Mca breaks down the complex into glucosaminylinositol that is used to synthesise more MSH and a mercapturic acid, which is subsequently excreted from the cell (Newton et al., 2000b; Newton and Fahey, 2002). Similarly, the detoxification of formaldehyde is accomplished due to the action of formaldehyde dehydrogenase (MscR), an enzyme that also acts as a nitrosothiol reductase. MSH can react with formaldehyde, and in the presence of NAD+, the formaldehyde is oxidised by MscR to the less toxic formate (Vogt et al., 2003). The nitrosothiol reductase activity of MscR is greater than its formaldehyde dehydrogenase activity (Vogt et al., 2003). This activity protects the cell from NO, and in fact, it was shown that mycothiol plays a major part in protection of mycobacteria from NO (Miller et al., 2007). MSNO, an adduct of MSH and NO, is formed, which is subsequently broken down into nitrate and MSH by the action of MscR in the presence of NADH (Vogt et al., 2003). It has been established that MSH is the most abundant LMW thiol in mycobacteria, which maintains redox homeostasis and is thermodynamically linked with a variety of enzymes to eliminate toxic compounds. Unfortunately, little is known thus far of the mechanisms utilised by mycobacteria to regulate the production of MSH.

6.2. The Mtb Dos Dormancy Regulon 6.2.1. Biological Role and Function The Dos two-component system was first identified in a virulent strain of Mtb (Kinger and Tyagi, 1993) wherein DosR (Rv3133c) was demonstrated to possess homology to response regulator proteins of the NarL/UhpA/ FixJ subfamily and DosS (Rv3132c) to sensor histidine kinases. The

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two-component pair genes, dosR and dosS, are cotranscribed and conserved in Mtb and M. bovis BCG (Dasgupta et al., 2000) and their expression is hypoxia-responsive in pathogenic and non-pathogenic mycobacteria including Mtb (Sherman et al., 2001), M. bovis BCG (Boon et al., 2001) and M. smegmatis (Mayuri et al., 2002). A series of studies revealed that significant overlap exists between the expression profiles of Mtb cells cultured under hypoxic conditions and those exposed to NO and that the transcriptional response is under direct control of DosR (Sherman et al., 2001; Ohno et al., 2003; Voskuil et al., 2003). Subsequently, DosT (Rv2027c), a homologue of DosS, was discovered and was also shown to phosphorylate DosR (Roberts et al., 2004; Saini et al., 2004), indicating a crosstalk between DosS, DosT and DosR. The role of DosT remains enigmatic as it is an orphan sensor kinase, has no cognate response regulator and is not upregulated under any known condition. Thus, DosS/T/R can be regarded as a ‘three’-component system believed to facilitate transition from active to the latent form of infection. Detailed studies have shown that DosT is active early in the hypoxia response. As O2 becomes more limiting, DosT loses its activity and DosS continues to induce the regulon thereafter. Thus, it has been proposed that Mtb responds to hypoxia in a stepwise manner (Honaker et al., 2009). More recently, significant studies have demonstrated that in addition to hypoxia and NO, the complete Dos regulon is also induced by CO (Kumar et al., 2007, 2008; Shiloh et al., 2008). Because of the acknowledged role of hypoxia and NO (and likely also CO) in Mtb persistence, the Dos regulon is a paradigm for Mtb signal transduction, illustrating that sensing and relaying of physiologically relevant host information induces an adaptive bacterial transcriptional response. The response of the Dos regulon to O2, NO and CO consists of the altered expression of 100 genes (Sherman et al., 2001; Voskuil et al., 2003, 2009; Kumar et al., 2007, 2008; Shiloh et al., 2008) that include the repression of well-characterised genes of protein synthesis, DNA synthesis/cell division, lipid or amino acid synthesis, aerobic metabolism and the induction of 48 genes which are mostly uncharacterised. Nearly all of the 48 genes initially induced by hypoxia, NO and CO require DosR for their induction (Honaker et al., 2008, 2009). Among these 48 genes, with known or predicted function, many are speculated to play a role in adaptation to hypoxic stress, such as acr (rv2031c; chaperone function), narX (rv1736c; unknown function), nark2 (rv1737c; nitrate/nitrite transport), fdxA (rv2007c; ferredoxin), nrdZ (rv0570; ribonuclease reductase), tgs1 (rv3130; triglyceride synthase) and Mtb orthologues of the universal stress protein family (rv1996, rv2005c, rv2028c, rv2623, rv2624c, rv3134c) (Voskuil et al., 2009). A particularly interesting finding is that isolates of the W-Beijing lineage of Mtb constitutively overexpress the Dos regulon under in vitro conditions

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(Reed et al., 2007), suggesting that the Dos regulon may confer an adaptive advantage for growth under conditions that inhibit aerobic respiration. Studies of a Mtb Ddos rv3134c/rv3133c/rv3132c mutant deficient in expression of the Dos regulon (Leistikow et al., 2010) demonstrate its essential role in Mtb survival in the Wayne model for in vitro dormancy wherein O2 is limiting. Notably, the Dos regulon is important for resumption of growth once Mtb exits this anaerobic state (Leistikow et al., 2010). More recently, an additional transcriptional response, defined as the enduring hypoxia response, was identified and comprised a set of 230 genes induced subsequent to the Dos dormancy response and expressed long term (Rustad et al., 2008). Several lines of evidence suggest that the diatomic gases O2, NO and CO likely play roles in mycobacterial persistence. First, it was demonstrated that tubercle bacilli require oxygen for growth, and since reactivation disease occurs primarily in the oxygen-rich upper lung lobes (Medlar, 1948), the role of oxygen tension in TB has received wide attention (Dubos, 1953; Wayne and Hayes, 1996; Boon et al., 2001; Voskuil et al., 2009; Gengenbacher et al., 2010). Rapid withdrawal of oxygen is lethal to Mtb; however, gradual depletion allows time for adaptation and bacterial survival (Wayne and Hayes, 1996). Furthermore, TB is associated with the most O2-rich site (upper region of the lung) within the body (Rich and Follis, 1942; Rasmussen, 1957; Riley, 1957), and significantly more bacilli are present in TB lung lesions connected to open airways versus lesions closed to airways (Medlar, 1948; Haapanen et al., 1959). Secondly, iNOS and therefore NO production is crucial for protection of mice against Mtb (MacMicking et al., 1997), and human macrophages in Mtb-infected tissues are also shown to express iNOS (Nicholson et al., 1996; Nathan and Shiloh, 2000). Convincing studies have demonstrated iNOS RNA and protein in bronchoalveolar lavage specimens from active pulmonary TB patients (Nicholson et al., 1996). Studies have also shown that increased exhaled NO and nitrite in patients with active pulmonary TB is due to an increased iNOS production (Wang et al., 1998) and that a polymorphism in iNOS influences susceptibility to TB (Gomez et al., 2007). Thirdly, significant overlap exists between the gene expression profiles of Mtb cells treated in vitro with NO or CO and that of bacilli cultured under hypoxic conditions (Voskuil et al., 2003, 2009). Fourthly, in recent studies it was shown that NO, O2 and CO (Ioanoviciu et al., 2007; Kumar et al., 2007, 2008; Sousa et al., 2007; Yukl et al., 2007) are modulatory ligands of the heme sensor kinases DosT and DosS and that they either bind directly to the heme-irons or oxidise the heme irons, suggesting that the bacilli have the machinery to continuously monitor O2, NO and CO levels during the course of infection. Regardless of the aforementioned data,

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the evidence linking hypoxia, NO and CO to latent TB in humans remains circumstantial. Is there a role for CO in human TB? Intriguingly, a role for environmental CO in TB was first described as early as 1923 (Hazleton, 1923) when individuals exposed to coal gas (which contains large quantities of CO) contracted TB. In observing a large number of similar cases, the investigator concluded that the inhalation of small quantities of coal gas for a considerable period of time might have been a predisposing cause of TB. Also, more recently, in examining the historical statistics on coal consumption and TB, an interesting hypothesis that links TB and coal-generated CO (and thus air pollution) was developed (Tremblay, 2007). The newly discovered role of binding of CO to DosS and DosT (Kumar et al., 2007; Sousa et al., 2007) and evidence suggesting that mycobacteria can oxidise CO (King, 2003; Park et al., 2003, 2007) have implications for understanding Mtb disease and persistence. Lastly, a seminal study demonstrated a role for HO-1 generated CO in protection against experimental cerebral malaria (Pamplona et al., 2007). Thus, hostgenerated CO production may be part of a generalised host response to a variety of pathogens. 6.2.2. The Mtb Dos Dormancy Regulon and Virulence For many decades, it has been postulated that pO2 influences the progression of human TB (Rich and Follis, 1942; Medlar, 1948; Rasmussen, 1957). Present-day studies using the hypoxia marker pimonidazole hydrochloride yielded the surprising discovery that granulomatous tissue in the Mtbinfected mouse model is relatively aerobic (37 mm Hg) (Aly et al., 2006; Tsai et al., 2006), but that tuberculous pulmonary lesions in macaques, rabbits and guinea pigs are hypoxic (Via et al., 2008). In fact, a detailed analysis using a fibre-optic O2 sensor showed that the pO2 in rabbit pulmonary lesions was 1.59 mm Hg (Via et al., 2008), demonstrating that the TB granuloma is indeed hypoxic. In vivo virulence studies have shown that an Mtb dosR mutant is attenuated in the guinea pig TB infection model (Malhotra et al., 2004; Converse et al., 2009). However, studies in mice (Parish et al., 2003; Rustad et al., 2008) have produced conflicting results. The discrepancies in data obtained in the various in vivo studies have been thoroughly discussed elsewhere (Converse et al., 2009). The pronounced expression of the Dos regulon and the presence of TAG in bacilli procured from the sputum of Mtb-infected individuals (Garton et al., 2008) suggest that sputum contains non-replicating bacilli and has significant implications for understanding the physiological makeup of the host environment that allows Mtb to persist. For example, because of

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the low O2 diffusion capability in mucus (Worlitzsch et al., 2002), it is plausible that the sputum is depleted for O2, which leads to upregulation of the Dos regulon. Furthermore, the direct induction of the Dos regulon by NO, CO or a combination thereof cannot be excluded, since TB patients (including healthy individuals) exhale NO (Gustafsson et al., 1991; Borland et al., 1993; Wang et al., 1998). Lastly, the diffusion of NO metabolites (NO3
, NO2
) across the alveolar capillary membrane followed by aerosolisation of airway epithelial lining fluid and confinement of NO2
/NO3
to the oropharyngeal tract and trachea (Marteus et al., 2005; Brindicci et al., 2009; Fitzpatrick et al., 2009), point to an environment in which access to NO3
may enable Mtb to oxidise excess NADH via NarG (see Section 6.2.4.1). 6.2.3. The Dos Regulon and CO The biochemical and biophysical basis of the Mtb response to O2, NO and CO involving the heme-irons of the sensor kinases DosS and DosT has been previously reviewed (Voskuil et al., 2009). However, since the role of CO in induction of the Dos regulon has only recently been discovered, a short overview will be given here on the biological relevance of this host gas. CO is a LMW gas that is endogenously produced by HO-1 in humans during normal metabolism and it is increased in response to oxidative stress (Ferry, 1995; Chung et al., 2009). CO binding to the heme-irons of DosS and DosT modulates their autokinase activity leading to the upregulation of key members of the Dos regulon (e.g. dosR, hspX and fdxA) (Kumar et al., 2007). This finding led the investigators to hypothesise that CO is capable of inducing the Dos regulon. Indeed, subsequent microarray studies performed by two independent groups have shown that exposure to CO induces the 48-member Dos regulon (Kumar et al., 2008; Shiloh et al., 2008). HO-1 requires O2 and NADPH as cofactors and utilises heme as a substrate, ultimately generating biliverdin, iron and CO. HO-1 confers protection against oxidative stress via the anti-inflammatory and anti-apoptotic activities of its byproducts (Chung et al., 2009; Ryter and Choi, 2009). Infection of macrophages by Mtb induces the expression of HO-1 mRNA, protein levels and enzymatic activity (Kumar et al., 2008). The ability of HO1 generated CO to induce the Dos regulon was demonstrated by infection of HO-1+/+ and HO-1
/
bone marrow-derived macrophages with Mtb, followed by expression analysis of the Dos regulon members (Kumar et al., 2008; Shiloh et al., 2008). Importantly, upregulation of HO-1 was found to be independent of the NO pathway (Kumar et al., 2008). In addition, HO-1 was found to be produced in the lungs of Mtb-infected mice (Kumar et al., 2008; Shiloh et al., 2008). Thus, the combined presence of NO and CO at the site of

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Mtb infection, which might also be hypoxic, provides new insight into how gaseous gradients may influence disease progression (Kumar et al., 2008). However, as mentioned earlier (Section 6.2.1), since O2 is a cofactor for iNOS [K mO2 ¼ 135 mM; (Dweik et al., 1998; Dweik, 2005)], the hypoxic nature of the granuloma (Via et al., 2008) might prevent production of significant quantities of NO. Although CO typically inhibits respiration in other bacteria (Davidge et al., 2009), Mtb is unusual in the sense that it is capable of tolerating relatively high concentrations of CO [80 mM (Shiloh et al., 2008)]. An important functional difference between NO and CO is that CO can only react with ferrous iron (Fe2+) whereas NO can react with both Fe2+ and ferric iron (Fe3+) (Kumar et al., 2007). Recently, a protective role for CO in experimental cerebral malaria was demonstrated and it was proposed that CO locks cell free haemoglobin in the Fe2+ state, thereby preventing oxidation to the Fe3+ state, which upon reaction with ROS would lead to disruption of the blood–brain barrier (Pamplona et al., 2007). A similar model termed the ‘sense-and-lock’ model was proposed for the mechanisms of DosS and DosT in sensing O2, NO and CO (Kumar et al., 2007). However, a definite role for CO in Mtb pathogenesis has yet to be demonstrated, and represents an important but underexplored area of investigation. 6.2.4. The Mtb Dos Regulon and Reductive Stress In physiological terms, it makes bioenergetic sense for an aerobe such as Mtb to express energy-dissipating systems under hypoxic conditions in order to maintain redox balance. For example, when Mtb encounters highly reduced carbon sources in vivo, the carbon source must be oxidised to the point of assimilation. If oxidation releases more reducing equivalents, for example via b-oxidation, than is needed for ATP generation (reductive stress), the bacillus must initiate mechanisms to dispose of excess reductants, otherwise the growth rate will be slowed to the rate at which NADH can be oxidised or ADP be phosphorylated. Using an in vitro dormancy model, studies have shown that in an Mtb Ddos rv3134c/rv3133c/rv3132c mutant, the ATP level decreased to 10% of that measured under aerobic growth. In contrast, the ATP level in the wt strain stabilised at 25% of the aerobic level (Leistikow et al., 2010). These findings are in agreement with an independent study demonstrating that, during hypoxia, Mtb has a much reduced but continuous pool of ATP (Rao et al., 2008). In fact, it was shown that further reduction of ATP (three to fourfold) under hypoxic conditions increases cell death to more than 90% whereas a

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depletion of more than 90% of ATP was required to see a killing effect under aerobic growth conditions. Consistent with these findings, it was demonstrated that the membrane of Mtb cultured under hypoxia is fully energised and is required for ATP production (Rao et al., 2008). NAD+ and NADH concentrations decrease by 50% when wt Mtb is grown under hypoxic conditions, yet the overall redox couple ratio is maintained. This is an intriguing observation as the NADH/NAD+ ratio typically increases in bacteria as O2 levels decrease (de Graef et al., 1999; BerriosRivera et al., 2002a). One explanation might be that increased NADH generated under hypoxia is consumed during TAG synthesis, thereby maintaining an ‘optimal’ NADH/NAD+ ratio. Regardless, in the Mtb Ddos mutant, the NAD+ levels dropped to 10% of the wt level, and the NAD+/NADH ratio was sixfold less under hypoxia. Thus, the data clearly suggest that the Mtb Dos regulon plays a role in maintaining redox homeostasis (Leistikow et al., 2010). Furthermore, it appears that ndh-2 rather than ndh-1 contributes to the generation of the proton motive force for ATP production under hypoxic conditions and that Ndh-2 is responsible for replenishing NAD+ in Mtb grown under hypoxic conditions (Rao et al., 2008). Two reports provide direct evidence for the presence of reductive stress in Mtb. First, the accumulation of extraordinarily high levels of NAD(P)H relative to NAD(P)+ was found in bacilli derived from the lungs of infected mice (Boshoff et al., 2008). Secondly, polyketide and TAG anabolism have been demonstrated to occur in vivo and the synthesis of such may serve as efficient reductant disposal mechanisms to alleviate reductive stress for longterm persistence (Singh et al., 2009) (Fig. 1) (see also Section 6.2.4.2). In conclusion, a role for the Dos regulon in reductive stress has largely been underappreciated. Nonetheless, the DosR-controlled NarGHJI/NarK2 and Tgs1 proteins are likely candidates for disposing of excess reducing equivalents and thereby maintaining redox balance. 6.2.4.1. Nitrate reductase Several functions can be attributed to nitrate reduction in bacteria (Cole, 1996; Richardson et al., 2001; Morozkina and Zvyagilskaya, 2007) and include (i) NO3
assimilation (using NO3
as a nitrogen source), (ii) NO3
dissimilation (disposal of excess reducing equivalents to maintain redox balance) and (iii) NO3
respiration (using NO3
as the terminal electron acceptor for the production of energy). Mtb contains two possible nitrate reductases, the NarGHJI (RV1161–1164) complex, which catalyses the reduction of NO3
to NO2
and NarX that has no known function to date.

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[(Figure_1)TD$IG]

Figure 1 Model depicting the dissipation of excess reductants through the Dos regulon to maintain redox homeostasis. Hypoxia, NO and likely CO, which inhibit respiration, are sensed by DosS/T leading to activation of the Dos regulon via DosR. Included in the Dos regulon are genes encoding for components of the rTCA cycle, and narGHJI-narK. These pathways serve as alternate electron sinks for reductive stress dissipation. The excess NADH, NADPH and FADH2 that are produced upon inhibition of respiration or b-oxidation of host FA are utilised for the generation of metabolic intermediates. (a) The rTCA cycle incorporates one molecule of CO2 in an energy-consuming process, wherein the energy is provided by the reducing equivalents. This allows the recycling of excess reductants for the production of essential metabolites, thereby lowering their intracellular concentrations. (b) The uptake of NO3
by NarK2 and its corresponding reduction to NO2
by NarG consumes cellular reducing equivalents. As the concentration of cytosolic NO2
increases to reach toxic levels, it is pumped out via NarK. As extracellular levels of NO3
decrease, NO2
is transported back into the cell and reduced via NirBD to NH3. (c) NADPH is a cofactor in the FASI pathway and is consumed during anabolism of the storage lipid, TAG.

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Several studies have shown that nitrate reductase activity increases dramatically during culturing of Mtb under microaerophilic conditions (Sohaskey and Wayne, 2003; Sohaskey, 2005, 2008). The narGHJI operon is constitutively expressed in Mtb whereas narK2, encoding for the NO3
/NO2
transporter, is under control of DosR and therefore a component of the Dos regulon that is highly induced during hypoxia and upon exposure to NO and CO (Sherman et al., 2001; Voskuil et al., 2003; Kumar et al., 2008). Depending on the organism, nitrate reductases require NADH or NADPH as cofactors (Richardson et al., 2001; Morozkina and Zvyagilskaya, 2007). Genetic experiments suggest that nitrate reductase activity in Mtb is not coupled to proton translocation and ATP synthesis (Sohaskey and Wayne, 2003). Rather, it has been concluded that nitrate reductase may be involved in maintaining redox balance during microaerophilic growth by dissipating excess reducing equivalents (Sohaskey, 2008). The redox potential of NO3
/NO2
is +420 mV and the change in Gibbs free energy approximates
82 kJ/mol (Seifritz et al., 1993), which suggests that NO3
is an effective electron sink. Although Mtb can utilise NO3
as a terminal electron acceptor under anaerobic conditions to maintain some degree of viability or to prolong survival, active replication does not occur in the absence of O2 (Sohaskey and Wayne, 2003; Aly et al., 2006; Sohaskey, 2008; Gengenbacher et al., 2010). Since Mtb does not actively respire anaerobically, it is tempting to theorise that the release of Gibbs free energy from the reduction of NO3
to NO2
contributes towards maintaining viability of the bacilli when encountering hypoxia in vivo. Complementation of an E. coli nark narU mutant with Mtb narK2 strongly suggests that NarK2 transports NO3
into and NO2
out of Mtb. Thus, under low-O2 conditions, NO3
is transported into the cell, reduced to NO2
and, when toxic levels of NO2
are reached, exported out. An interesting parallel exists between Mtb and Staphylococcus carnosus. In case of the latter, the exported NO2
is transported back into the cell and reduced to ammonia, which again accumulates in the medium (Neubauer and Gotz, 1996). This event is reminiscent of the increased alkalinity (as opposed to acidity) detected in the culture supernatant of Mtb, which was shown to be due to buildup of ammonia (Merrill, 1930). It is tempting to speculate that NirB (Rv0252) and NirD (Rv0253) are Mtb nitrite reductase subunits that reduce NO2
to ammonia without liberating intermediate products and may explain the observed alkalinity by Merrill (1930). An obvious question is: does Mtb encounter NO3
and/or NO2
in vivo? Indeed, the presence of NO3
plus NO2
in saliva (313 mM), NO3
in trachael secretions (144–421 mM) (Grasemann et al., 1998; Linnane et al., 1998; Worlitzsch et al., 2002), NO3
(0.5–37 mM) and NO2
(1.4–15 mM) in

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bronchoalveolar lavage fluid (Dweik et al., 2001; Fitzpatrick et al., 2009), and NO2
/NO3
in exhaled breath condensate (17 mmol) and sputum (449 mmol) (Brindicci et al., 2009) provides evidence that Mtb is likely to encounter NO2
and/or NO3
during the full spectrum of disease. 6.2.4.2. TAG production An underappreciated consequence of Mtb utilisation of host lipids for energy is the production of large quantities of reducing equivalents that are essential cofactors for the production of storage lipids (TAG) and virulence lipids such as SL-1, PDIM and PAT/DAT. At least two widely known biological functions of TAG in prokaryotes include its use as an effective reserve carbon source because of its reduced COS, and its service as an electron sink for excess reducing equivalents (Alvarez and Steinbuchel, 2002). Thus, it appears that TAG anabolism is a plausible mechanism for dissipation of reductive stress in Mtb (Singh et al., 2009). TAG production is under the control of the DosR regulon and its generation is highly increased when Mtb cells are exposed to hypoxia, NO or CO (Voskuil et al., 2009). Interestingly, TAG is found in the sputum of TB patients (Garton et al., 2008). Studies using the in vitro Wayne model for dormancy have shown that TAG is catabolised by Mtb upon reactivation (Deb et al., 2006) and thus may implicate a possible role for TAG in emergence of Mtb from a persistent state in vivo.

6.3. Mtb WhiB3 is an Intracellular Redox Sensor that Counters Reductive Stress 6.3.1. The WhiB Family of Proteins The first WhiB-like protein was characterised in Streptomyces in 1992 (Davis and Chater, 1992) and was predicted to have a putative helix-turn-helix (HTH) motif at its C-terminus. WhiB-like proteins are LMW (81–122 amino acids) and are restricted to actinomycetes (den Hengst and Buttner, 2008). WhiB members contain four conserved Cys residues, with one exception (Cole et al., 1998). Mtb contains seven homologues of WhiB (WhiB1, Rv3219; WhiB2, Rv3260c; WhiB3, Rv3416; WhiB4, Rv3681c; WhiB5, Rv0022c; WhiB6, Rv3862c and WhiB7, Rv3197A) that show strong homology to regulatory proteins of Streptomyces spp. critical for sporulation (Chater, 1972; Flardh et al., 1999). In one of the initial studies on mycobacterial WhiB proteins, it was shown that Msm WhmD is essential and is involved in septum formation and fragmentation (Gomez and Bishai, 2000). Subsequently, other members

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of the WhiB family have been shown to play a role in combating oxidative stress in Corynebacterium glutamicum (Kim et al., 2005). An important finding was the establishment of Mtb WhiB3 as a virulence factor (Steyn et al., 2002) (see Section 6.3.2 below for a complete description). Moreover, the observations that Mtb WhiB7 plays a role in antibiotic resistance and that it is highly upregulated in the presence of palmitic acid, a putative in vivo carbon source, are particularly interesting (Morris et al., 2005) and suggest that, during infection, the specific in vivo environment might allow Mtb to effectively resist chemotherapeutic intervention strategies. Examination of the differential expression of all seven Mtb whiB genes during growth and upon exposure to a wide range of antibiotics or to a variety of in vitro stress conditions indicates that the Mtb WhiB family is strongly reactive to a wide range of environmental stress conditions (Geiman et al., 2006). Consistent with the presence of conserved Cys residues in the WhiB family, it was demonstrated that WhiD, a Streptomyces member of the WhiB family, contains a 4Fe–4S cluster (Jakimowicz et al., 2005). Subsequently, it was demonstrated that Mtb WhiB3 also possesses a 4Fe–4S cluster (Singh et al., 2007), suggesting that this feature might be common to all seven Mtb homologues. Studies on Mtb apo-WhiB1 (Garg et al., 2007) and apo-WhiB4 (Alam et al., 2007) suggest that they function as disulphide reductases. Using the yeast two-hybrid system, the same group has shown that WhiB1 interacts with GlgB (an a-1,4-glucan branching enzyme) and reduces the intra-molecular disulfide bond of GlgB (Garg et al., 2009). Additionally, a biochemical study provides some evidence that all the Mtb WhiB’s contain Fe–S clusters (Alam et al., 2009). However, since the study predominantly used UV-visible spectroscopy and not electron paramagnetic resonance spectroscopy to characterise the Fe–S clusters, the type of Fe–S cluster (4Fe–4S, 3Fe–4S or 2Fe–2S) of each homologue was not conclusively demonstrated. Lastly, an interesting study has shown that the WhiB-like protein of mycobacteriophage TM4 leads to the downregulation of Mtb whiB2 with subsequent filamentation and growth inhibition (Rybniker et al., 2010). The likely redox-mediated regulation of a host mycobacterial protein by a viral factor is an exciting finding and warrants further investigations. 6.3.2. Mtb WhiB3 and Virulence In 1995, a single point mutation (Arg515–His) in the 4.2 region of rpoV (encoding the principal s-factor SigA) was shown to be responsible for the loss of virulence of wt M. bovis (Steyn et al., 2002). rpoV was the first gene identified in the Mtb complex required for virulence. The biological

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mechanism of attenuation caused by this mutation remained unknown until a yeast two-hybrid screen, which employed the wt rpoV 4.2 region allele as bait, identified a small regulatory protein, WhiB3, as an interacting partner. Significantly, WhiB3 interaction with the attenuated RpoV allele containing the single point mutation described above was severely impaired (Steyn et al., 2002). Therefore, loss of interaction with WhiB3 was the likely reason for the attenuating nature of the RpoV R-H mutation. Although whiB3 (Rv3416) deletion mutants of both Mtb H37Rv and M. bovis displayed no observable phenotypic changes during in vitro growth, both were shown to be attenuated in vivo. Infection of guinea pigs with the M. bovis DwhiB3 mutant demonstrated that this strain was unable to colonise the spleen. Perhaps a more striking finding was that the Mtb DwhiB3 deletion mutant showed wt-like growth in all organs, yet the mean survival time of C57BL/6 mice infected with the Mtb DwhiB3 deletion strain (280 days) was virtually twice to that of mice similarly infected with wt Mtb H37Rv (150 days). Furthermore, despite identical organ burdens, a dramatic difference in lung pathology induced by mutant and wt was observed. The lungs of the Mtb DwhiB3-infected mice showed less pathology and cellular infiltration compared to those of mice infected with wt Mtb, an observation that suggested that infection with the Mtb DwhiB3 mutant was not associated with a harmful inflammatory response (Steyn et al., 2002). Owing to the reduced immunopathology caused by this mutant in mice and because a Streptomyces whiD mutant has repressed expression of whiE that encodes a polyketide synthase (Kelemen et al., 1998), a connection between whiB3, the production of Mtb polyketides and detrimental immunopathology was proposed (Steyn et al., 2002). Consistent with the idea that WhiB3 is needed for the synthesis of cellsurface polyketides is the observation of differences in colony morphology displayed by the wt and the DwhiB3 mutant strains (Singh et al., 2007). Finally, whiB3 expression was found to inversely correlate with bacterial density in vivo in the mouse system, suggesting a possible role for WhiB3 in quorum sensing (Banaiee et al., 2006). An important finding was the demonstration that Mtb contains a cysteine desulphurase (IscS; Rv3025c) capable of assembling the WhiB3 4Fe–4S cluster (Singh et al., 2007). The four WhiB3 Cys residues were subsequently shown to be essential in coordinating a Fe–S cluster. Exposure of purified WhiB3 to air or NO led to cluster degradation or formation of a dinitrosyl iron–dithiol complex respectively. Therefore, WhiB3 reacts directly with O2 and NO and likely functions as an endogenous sensor of these two dormancy gases (Singh et al., 2007). Aside from the identification of a functional IscS, not much is known about Fe–S cluster assembly systems in mycobacteria. A putative Mtb SUF

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(mobilisation of sulphur) system (SufBCD) was identified through protein– protein interaction studies (Huet et al., 2005), and formation of this protein complex depends on the protein splicing (via an intein) of SufB (Huet et al., 2006). Mtb contains more than one IscS homolog and the SufBCD system, which are virtually uncharacterised in terms of general Fe–S cluster assembly. It has now been demonstrated that WhiB3 is a regulator of cellular metabolism, wherein it is important for growth in the presence of glucose, pyruvate, succinate and fumarate as sole carbon sources (Singh et al., 2007). The defective growth exhibited by the Mtb DwhiB3 mutant on glucose or succinate as the sole carbon source can be rescued by complementation with wt WhiB3, but not with WhiB3 that has all four Cys residues mutated. Thus, the [4Fe–4S]+ cluster is necessary for WhiB3 to function as a metabolic regulator. Intriguingly, the Mtb DwhiB3 mutant grows much better in media containing the short-chain FA acetate, suggesting that WhiB3 is a potential regulator of FA metabolism in Mtb. It has been proposed that WhiB3 senses or responds to cellular substrates via its [4Fe–4S]+ cluster in order to affect a metabolic switchover to the use of FA as the preferred carbon source in vivo (Singh et al., 2007). The role of WhiB3 in core intermediary metabolism is consistent with studies on FNR and ArcA, which regulate key enzymes in the glycolytic and TCA cycles in response to carbon source starvation (Levanon et al., 2005; Shalel-Levanon et al., 2005). In conclusion, the pathological defect induced by Mtb DwhiB3 in the mouse model (Steyn et al., 2002) as well as the altered colony morphology and growth properties of Mtb DwhiB3 on carbon-limiting media, in particular FAs (Singh et al., 2007), suggests that WhiB3 is involved in maintaining redox homeostasis by regulating catabolism and polyketide biosynthesis in Mtb. The precise mechanism by which this occurs remains to be investigated. 6.3.3. WhiB3 and Reductive Stress Mtb WhiB3 is essential for maintaining bacterial cell shape and size, and modulates the biosynthesis of complex virulence lipids including PAT, DAT, SL-1 and PDIM (Singh et al., 2009). An interesting finding was that Mtb DwhiB3 displayed defective production of the methyl-branched polar lipids PAT, DAT and SL-1 but showed increased synthesis of PDIM and, to a lesser extent, TAG. The finding that Mtb DwhiB3 accumulates PDIM and TAG is unique and has not yet been reported for any Mtb mutant to date. Examination of the mycolic acid profiles demonstrated a decrease in a,a’-trehalose dimycolate (TDM) and a,a’-trehalose monomycolate (TMM) in Mtb DwhiB3 as compared to wt Mtb. Importantly, further studies

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revealed that WhiB3 regulates the production of PAT, PDIM and TAG in a redox-dependent manner in vitro (Singh et al., 2009). The discovery of a redox-switching mechanism of Mtb virulence lipid production is the first of its kind and provides important insight into how oxido-reductive host factors could modulate the synthesis of lipids involved in virulence. In these experiments, diamide and DTT were used as a thiol-specific oxidant and reductant respectively. An interesting observation was that under reductive stress conditions (treatment with DTT), a significant increase in TAG production was observed in Mtb DwhiB3 compared to wt Mtb (Singh et al., 2009). These findings have several important biological implications. First, they establish a link between the extracellular heme-based Dos dormancy signaling pathway (TAG is under DosR control), the intracellular 4Fe–4S cluster WhiB3 signaling pathway and reductive stress. Secondly, they suggest that intracellular oxido-reductive stress modulates diverse polyketides, which are implicated in virulence (Cox et al., 1999; Trivedi et al., 2005), and TAG production that might be essential for emergence from a persistent state (Deb et al., 2006). Thirdly, since it is well known that TAG can function as an effective electron sink in bacteria (Alvarez and Steinbuchel, 2002), the data implicate WhiB3 in maintaining intracellular redox homeostasis. As host FAs are degraded via b-oxidation, potentially toxic levels of propionate, a byproduct of odd chain FA assimilation, are generated (Jain et al., 2007; Upton and McKinney, 2007). The fact that Mtb DwhiB3 was able to grow on much higher concentrations of propionate compared to wt Mtb, and that it accumulated PDIM and TAG during intra-macrophage growth, suggests that the increased resistance to propionate toxicity is because propionate is channelled into PDIM via the methyl-malonyl CoA (MMCoA) pathway, and into TAG (Singh et al., 2009). Thus, WhiB3-mediated regulation of virulence lipid anabolism is a plausible mechanism for the detoxification of excess levels of propionate in vivo. In fact, it was shown that increased flux of MMCoA augments the size and abundance of PDIM and SL-1. The increased synthesis and mass of PDIM that occurred during infection of mice provides credence to the notion that propionate toxicity in vivo is relieved via PDIM anabolism (Jain et al., 2007). Large quantities of NADH and NADPH are generated from b-oxidation of host FA (e.g. palmitate) (see Section 4.2.2), which, if not properly balanced, can lead to reductive stress. A central question is: How does Mtb WhiB3 modulate the disposal of, or dissipate excess reducing equivalents to maintain redox balance? In light of the fact that Mtb does not use NO3
as a terminal electron acceptor under anaerobic conditions to promote replication, does not ferment and, to the best of our knowledge, does not secrete redox active molecules or use electron bifurcation mechanisms (Thauer,

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1988; Darrouzet and Daldal, 2003; Xia et al., 2007; Thauer et al., 2008; Costa et al., 2010) for energy production; this raises the question as to what the primary electron sink in Mtb is? Although this is an emerging area of investigation and not much is known at this point of time, several recent findings suggest that Mtb lipid anabolism, which requires NAD(P)H as cofactors, functions as an effective electron sink and that WhiB3 plays an important regulatory role in this process. First, in vivo studies have shown that Mtb isolated from infected mouse lungs have extraordinarily high levels of NAD (P)H, providing evidence that Mtb experiences severe reductive stress (Boshoff et al., 2008). Second, studies have shown that PDIM levels and bacterial mass increase significantly in infected mice (Jain et al., 2007). Third, WhiB3 was shown to modulate the differential anabolism of mycobacterial lipids including methyl-branched FAs (PDIM, SL-1, PAT and DAT) and TAG during growth in vitro and in macrophages (Singh et al., 2009). Fourth, Mtb DwhiB3 treated with DTT increased synthesis of TAG dramatically (Singh et al., 2009). Fifth, using [14C] nicotinamide incorporation and enzymatic cycling assays, it was shown that Mtb DwhiB3 accumulated large quantities of NADPH in macrophages and therefore experienced reductive stress (Singh et al., 2009). Collectively, the data suggest that Mtb lipid anabolism in vitro and in vivo demand substantial quantities of NAD(P) H and therefore functions as an effective electron sink to alleviate excess reducing equivalents and reductive stress. In order to explain these events, a model (Fig. 2) was proposed to illustrate the catabolism of highly reduced host FAs, as well as exposure of the bacilli to hypoxic conditions or NO, thereby generating reductive stress. In order to dispose or dissipate the excess reducing equivalents [NAD(P)H], Mtb anabolises PAT/DAT, SL-1, PDIM and predominantly TAG, which function as electron sinks. In conclusion, the mode of action of Mtb WhiB3 represents an elegant avenue for examining the mechanistic basis of reductive stress dissipation in bacterial pathogens and should open new areas of investigation. 6.3.4. WhiB3 and DNA Binding For more than 15 years, WhiB homologues in actinomycetes have been speculated to be putative DNA-binding transcription factors (den Hengst and Buttner, 2008). However, a formal proof demonstrating their DNA binding activity was lacking until recently. Singh et al. (2009) provided the first evidence that a member of the WhiB family can bind to DNA. Gel retardation studies have shown that the redox state of the Mtb WhiB3 4Fe– 4S cluster does not influence DNA binding and that oxidised apo-WhiB3 binds DNA tightly compared to holo-WhiB3. This is unusual because either

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[(Figure_2)TD$IG]

Figure 2 Model depicting the role of Mtb WhiB3 in sensing and dissipating reductive stress to maintain redox homeostasis. (a) Generation of reductants. b-oxidation of host FA and the inhibition of respiration by hypoxia, NO and possibly CO lead to the generation of NADH and NADPH that are major contributors to reductive stress in Mtb. (b) Regulation. WhiB3 maintains intracellular redox homeostasis through its Fe–S cluster via an alteration in its apo and holo redox states. This modulates DNA binding and regulates the expression of genes necessary for lipid anabolism. (c) Dissipating reductive stress. The induction of lipid anabolism by WhiB3 leads to the dissipation of excess reductants. Anabolism of a wide range of lipids implicated in virulence (PAT, DAT, PDIM and SL-1) via MMCoA, and the synthesis of storage lipid (TAG) by the FASI pathway functions as electron sinks. Note that the Dos dormancy regulon and the WhiB3 signaling pathway are linked via TAG anabolism (Singh et al., 2009).

classic bacterial Fe–S cluster proteins such as FNR and SufR require an Fe–S cluster for DNA binding (Green and Paget, 2004), or binding is dictated by the redox state of the Fe–S cluster (Shen et al., 2007). In a recent study, apoWhiB binding was also demonstrated for Mtb WhiB2 and a mycobacteria phage TM4 WhiB homologue, WhiBTM4 (Rybniker et al., 2010), suggesting

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that apo-WhiB binding of DNA might be a common feature of the WhiB family. Although DNA binding does not provide categorical proof for transcriptional regulation, evidence from a previous study demonstrating that WhiB3 selectively interacts with the 4.2 domain of the Mtb principal sigma factor, RpoV (SigA) (Steyn et al., 2002), suggests a plausible role for WhiB3 in transcriptional regulation. Consistent with this observation, WhiB3 directly regulates the transcription of various lipid/polyketide biosynthetic genes including pks2 (for SL-1 production), pks3 (PAT/DAT production), fbpA (TDM biosynthesis), mas, ppsA, fadD26 and fadD28 (PDIM production) by directly binding to their promoter regions (Singh et al., 2009). However, the role of the WhiB3 4Fe–4S cluster and the oxidation state thereof in transcriptional regulation of those genes remains to be determined.

7. CONCLUDING REMARKS Development of new intervention strategies against latent TB hinges on gaining a substantially better understanding of the in vivo physiology of Mtb, including the critical anabolic and catabolic pathways which allow the bacillus to persist in spite of host immune responses and/or chemotherapy. The crucial role of redox couples (e.g. NADH/NAD+, NADPH/NADP+, FADH2/FAD, MSH/MSSM, ERGox/ERGred) in metabolic homeostasis clearly suggests that oxidation–reduction reactions play an indispensible role in the maintenance of latency. Even though the balance of such reactions can swing either way, research has focused primarily on oxidative stress and its counterpart, reductive stress, has largely been ignored. An emerging hypothesis regarding the triggering events resulting in the Mtb latency response might be that of ‘reductive stress’. As stated earlier, reductive stress is defined as increased ‘reducing power’, usually caused by an excess of high-energy reductant including NADH or NADPH, or a failure of the mechanism(s) to counter those excesses, leading to a reductive cytosolic environment (Ido et al., 1997). Although very little is known about reductive stress in bacterial biology, it is possibly more prevalent than oxidative stress and may also be the main source of ROS (Ghyczy and Boros, 2007). The phenomenon of reductive stress is well established in eukaryotic biology and its role has been demonstrated in various human diseases including diabetes and cardiomyopathy (Boucher, 2007; Zhang et al., 2010). Although the role of reductive stress in microbial-induced diseases is inviting

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an increasing research interest, very few such studies have been reported. Nonetheless, the concept of reductive stress has provided researchers with a plethora of new research avenues that include the potential for development of new diagnostic and therapeutic intervention strategies. Numerous studies have demonstrated increased sensitivity to both oxidative and reductive stress in MSH mutants, emphasising the importance of MSH as the major redox buffer in mycobacteria. However, ERG in mycobacteria has not been studied for the past 50 years and warrants further investigation. Also, the link between redox homeostasis and drug efficacy in mycobacteria is important, since many anti-mycobacterial drugs are prodrugs that are activated upon reduction in the mycobacterial cytoplasm. This leads to the logical conclusion that mechanisms involved in maintaining the intracellular redox homeostasis of Mtb are important factors in drug resistance studies. However, the genetically intractable nature of this pathogen and the lack of novel tools allowing measurement of the intracellular redox environment of Mtb hamper novel drug discovery approaches in this regard. The wealth of anabolic and catabolic information gathered from a diverse spectrum of non-pathogenic and pathogenic bacteria, lower eukaryotes and other model microbes has revealed numerous electron sinks (e.g. fermentation, intracellular lipid accumulation, Calvin cycle, rTCA cycle, hydrogenase activity, lipid anabolism, etc.) for the recycling of reducing equivalents. These recycling sinks are unique for the particular environmental niche the microbe occupies. Despite some recent progress (Singh et al., 2009), their place in Mtb physiology is poorly defined at present. As more is learned about the role of electron sinks in model microbes, Mtb researchers should be compelled to revisit the existing paradigms for bacilli persistence, and to seek new testable hypotheses. As is clear from many historical studies, Mtb utilises carbohydrates during in vitro growth without the production of acids; rather, the culture media becomes alkaline. The latter finding begs the questions of how carbohydrate metabolism by Mtb is distinct from other bacteria that produce acids, and what are the generated end products, if any? Data indicating that lipids are the preferred in vivo carbon sources are compelling, but are based upon indirect gene knockout inferences and genome sequence information. Detailed in vivo studies using radiolabeled FA precursors should be undertaken. The effect of exogenous host lipid catabolism on endogenous Mtb lipid anabolism must be explored. Insights into how these metabolic events are coordinated to recycle reducing equivalents are needed. Furthermore, direct measurement of the intracellular redox environment of Mtb during in vitro growth and during infection will prove invaluable.

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The majority of investigations addressing the fundamental aspects of in vivo Mtb physiology (meaning the study of Mtb isolated from infected lung tissue) are decades old. Although they remain remarkably perceptive, there is an urgent need to revisit those studies using modern molecular biological tools. Mtb is an excellent candidate for in vivo study because answers to key questions regarding the mechanisms of persistence have not yet been obtained using in vitro model systems. In principle, the need to study in vivo-derived organisms was recognised long ago, particularly when it was elegantly revealed that the respiratory responses to a spectrum of carbohydrates and FAs were vastly different between in vitro-cultured and in vivo-isolated bacilli (Segal and Bloch, 1956). The marked response of in vivo-derived bacilli to FAs and lack of utilisation of carbohydrates suggest physiological and biochemical adaptation to widely diverse environments, changes which should be analysed by modern-day molecular tools such as DNA microarray, mass spectrometry and metabolic profiling. Using gene knock-out strategies and enzyme activity assays, a systematic in vitro and in vivo analysis of the Krebs cycle components and constituents of other pathways (e.g. glycolysis) should be undertaken. 13C-metabolic flux analysis of in vivo-isolated Mtb versus in vitro-cultured bacilli should reveal important insights into pathways that are specific for infection, and perhaps persistence. As discussed, the in vivo carbon source(s) used by Mtb is important from the perspective of energy generation and the choice almost certainly influences metabolic homeostasis. The combined effect of in vivo carbon substrate utilisation (most likely lipids and cholesterol) and the presence of environmental gases (e.g. NO, CO2) that inhibit respiration results in the accumulation of reducing equivalents that require recycling to maintain redox balance. The admittedly limited biochemical and physiological data on reductive stress in Mtb points to lipid anabolism as an electron sink for recycling reducing equivalents (Singh et al., 2009). The latter finding raises hopes for the future unveiling of novel features of the in vivo Mtb metabolome, lipodome and redoxome. Furthermore, the potential roles of hydrogenases, CODH and the rTCA cycle in dissipating reductive stress in Mtb are unexplored and therefore represent new research areas in need of investigations. Insight into the role of WhiB3 in the maintenance of redox homeostasis by diverting reductive stress in macrophages (Singh et al., 2009) and the observation that an extraordinary amount of reducing equivalents are produced in the lungs of Mtb-infected mice (Boshoff et al., 2008) represent starting points for other innovative avenues of research. Studies on the microenvironment of the granuloma wherein Mtb resides in vivo remain at an early stage of investigation, and few reports are

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available. The current paradigm, which suggests that Mtb resides in distinct and dynamic microenvironments within the lung (Barry et al., 2009), represents a biological predicament for the mycobacteriologist, since the metabolic state of the bacilli within each independent granuloma may be dissimilar from that of another. For example, the microaerophilic or anaerobic state of the particular granuloma, presence of gases, and exposure of the bacilli to a vast range of host factors that vary throughout the course of infection, illustrate the challenges of examining Mtb in vivo. Creatively exploiting genome-wide technologies that examine the transcriptional state and metabolic profile of Mtb within distinct granulomas is essential towards identifying new pathways and metabolites, and establishing whether existing pathways are ‘active’ during infection. As more is learned about the physiology and biochemistry of Mtb, a better understanding of TB is obtained which is key to the development of effective anti-mycobacterial strategies.

ACKNOWLEDGEMENTS We wish to thank members of the Steyn laboratory for critical reading of this manuscript. Research in our laboratories is supported in whole or in part, by National Institutes of Health Grants AI058131, AI076389 (to A.J.C.S.) and AI060469 (to M.K.H.) This work was also supported by the University of Alabama at Birmingham (UAB) Center for AIDS Research, UAB Center for Free Radical Biology and UAB Center for Emerging Infections and Emergency Preparedness (A.J.C.S.). A.J.C.S. is a Burroughs Wellcome Investigator in the Pathogenesis of Infectious Diseases.

REFERENCES Alam, K.Y. and Clark, D.P. (1989) Anaerobic fermentation balance of Escherichia coli as observed by in vivo nuclear magnetic resonance spectroscopy. J. Bacteriol. 171, 6213–6217. Alam, M.S., Garg, S.K. and Agrawal, P. (2007) Molecular function of WhiB4/ Rv3681c of Mycobacterium tuberculosis H37Rv: a [4Fe–4S] cluster co-ordinating protein disulphide reductase. Mol. Microbiol. 63, 1414–1431. Alam, M.S., Garg, S.K. and Agrawal, P. (2009) Studies on structural and functional divergence among seven WhiB proteins of Mycobacterium tuberculosis H37Rv. FEBS J. 276, 76–93. Alvarez-Ortega, C. and Harwood, C.S. (2007) Responses of Pseudomonas aeruginosa to low oxygen indicate that growth in the cystic fibrosis lung is by aerobic respiration. Mol. Microbiol. 65, 153–165.

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Regulation of CtsR Activity in Low GC, Gram+ Bacteria Alexander K.W. Elsholz, Ulf Gerth and Michael Hecker Ernst-Moritz-Arndt-University Greifswald, Institute of Microbiology, Greifswald, Germany

ABSTRACT CtsR is the global transcriptional regulator of the core protein quality networks in low GC, Gram+ bacteria. Balancing these networks during environmental stress is of considerable importance for moderate survival of the bacteria, and also for virulence of pathogenic species. Therefore, inactivation of the CtsR repressor is one of the major cellular responses for fast and efficient adaptation to different protein stress conditions. Historically, CtsR inactivation was mainly studied for the heat stress response, and recently it has been shown that CtsR is an intrinsic thermosensor. Moreover, it has been demonstrated that CtsR degradation is regulated by a two-step mechanism during heat stress, dependent on the arginine kinase activity of McsB. Interestingly, CtsR is also inactivated during oxidative stress, but by a thiol-dependent regulatory pathway. These observations suggest that dual activity control of CtsR activity has developed during the course of evolution. 1. 2. 3. 4.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Protein Quality Control. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ctsr-Regulated Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cellular Functions Of Genes Regulated by CtsR. . . . . . . . . . . . . . . . . . . Mechanisms for the Inactivation of the CtsR Repressor . . . . . . . . . . . . . 4.1. Heat Inactivation of CtsR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. CtsR Inactivation During Oxidative Stress . . . . . . . . . . . . . . . . . . .

ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 57 ISSN: 0065-2911 DOI:10.1016/B978-0-12-381045-8.00003-5

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ALEXANDER K.W. ELSHOLZ ET AL. 4.3. CtsR Inactivation During Other Stress Conditions . . . . . . . . . . . . . Control of Ctsr Degradation by the Regulated Adaptor McsB . . . . . . . . . Summary And Outlook. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABBREVIATIONS HTH HHP LMWPTP

helix-turn-helix high-hydrostatic pressure low-molecular-weight protein tyrosine phosphatase

1. PROTEIN QUALITY CONTROL The maintenance of proper protein homeostasis is important for viability and growth of all living organisms, and cells have evolved two major strategies. Protein quality networks ensure the correct protein function as molecular chaperones promote protein folding and mediate refolding while ATPdependent proteases degrade misfolded or aggregated proteins to prevent cell injury when refolding by molecular chaperones has failed. Stress conditions such as heat, oxidative stress and extreme pH values result in damage of the protein structure that may lead to aggregation of proteins which fail to fulfill their physiological function and are often lethal for the cell. Hence, enhanced expression of these critical protein classes is needed for moderate survival of the cell during severe protein stress conditions (Gething and Sambrook, 1992; Wickner et al., 1999; Sauer et al., 2004; Hartl and HayerHartl, 2009). The cellular chaperone machinery assists in non-covalent folding or unfolding of macromolecular complexes, but molecular chaperones are not involved in the normal enzymatic reactions of these complexes. In all three domains of life, a plethora of structurally unrelated chaperones have evolved (Young et al., 2004), but general modes of operation are conserved for all molecular chaperones. In general, all chaperones recognise hydrophobic residues and/or unstructured backbone regions in proteins, that is structural features normally buried upon completion of folding but typically exposed by non-native proteins. This ensures that

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 121 only incorrectly folded proteins become targets for molecular chaperones (Hartl and Hayer-Hartl, 2009). Energy-dependent protein degradation is achieved by large cylindrical assemblies with a common ring-stacking architecture of diverse complexity. This architecture is highly conserved in all living organisms and operates with similar principles. It is composed of a chaperone ring comprising ATPase domains of the AAA+-type (ATPase associated with a variety of cellular activities) that caps both ends of self-compartmentalised proteases whereby the active sites are located in an internal chamber and are thereby separated from the cytosol (Wickner et al., 1999; Sauer et al., 2004). Hydrolysis of ATP by the AAA+ superfamily of proteins is translated into force that unfolds substrates and translocates them into the proteolytic chamber of the protease subunit where the peptide bonds are hydrolysed (Neuwald et al., 1999; Baker and Sauer, 2006). Protein degradation in eukaryotes is performed by the 26S proteasome comprised of a 19S cap particle with six different AAA+ proteins at its base, and a 20S core particle which contains the proteolytic site (Pickart and Cohen, 2004). A 26S proteasome system is not present in eubacteria, except for some actinobacteria (Darwin, 2009). However, at least four ATPdependent proteases (Clp, HslUV, Lon and FtsH) operating on a similar principle have evolved in eubacteria (Gottesman, 2003). In low GC, Gram+ bacteria the Clp machinery seems to be the major system for general protein turnover (Frees and Ingmer, 1999; Kr€ uger et al., 2000; Kock et al., 2004b; Gerth et al., 2008). The Clp protease is constituted of Hsp 100/Clp proteins of the AAA+ superfamily and an associated barrel-like protease complex formed by ClpP (Gottesman, 2003). The multimeric barrellike structure of ClpP is formed by two stacked heptameric rings of ClpP homomers which create a catalytic cavity wherein the 14 proteolytic active serine residues are enclosed (Wang et al., 1997). ClpP, on its own, is only able to degrade small peptides and with very restricted efficiency (Jennings et al., 2008). Therefore, a regulatory AAA+ protein of the HSP 100/Clp family flanks both ends of the ClpP complex, displaying a gateway to the protease. These regulatory proteins form hexameric rings with a narrow pore in the middle through which the substrate is translocated into the internal chamber of the peptidase (Zolkiewski, 2006). Substrate unfolding and threading through the narrow pore into the proteolytic chamber by the Hsp 100/Clp proteins is ATP dependent (Weber-Ban et al., 1999). Once the unfolded polypeptide enters the ClpP proteolytic chamber, it is rapidly degraded, without further utilisation of ATP, to short peptides of 7–10 amino acids length (Thompson et al., 1994).

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2. CtsR-REGULATED GENES In the Gram-negative model organism Escherichia coli the regulation of classical heat shock genes depends mainly on the level of the alternative transcription factor s32 (Yura, 1996). However, in the low GC, Gram+ model organism Bacillus subtilis regulation of protein quality control systems differs significantly. To date, at least six different classes have been distinguished for induced expression of heat shock proteins (Schumann, 2004). Class one was defined as genes regulated by the global repressor HrcA, class two encompasses genes under the control of the alternative sigma factor SigB, class three genes belong to the CtsR regulon, class four contains only the htpG gene, class five is controlled by the CssRS two-component system and class six is reserved for genes whose expression is induced during heat stress but with no known regulatory mechanism (Schumann, 2004). The central core of the protein quality control in all low GC, Gram+ bacteria is under the control of CtsR (class three stress gene repressor), as long as a ctsR gene is present in the corresponding genome. To date, the only bacilli lacking a ctsR gene belong to the Lactobacillus acidophilus group (van de Guchte et al., 2006). Although these six different classes of heat shock proteins are not present in all Gram+ bacteria, regulation of heat shock induction remains complex and is also divided into several groups in other low GC, Gram+ bacteria. Nevertheless, these regulons are not very committed in their specific contributions and thus partial or full overlap between different classes can be found in some species, whereas these classes are very distinct in others. The alternative sigma factor SigB is only conserved in the order Bacillales (Hecker et al., 2007) and is defined as class two in the heat shock response of B. subtilis. SigB overlaps with CtsR in the regulation of the clpC operon and €ger et al., 1996; Gerth et al., 1998) as well as for the clpP in B. subtilis (Kru clpC, clpP and clpB operons in Listeria monocytogenes (Hu et al., 2007). However, in Staphylococcus aureus no functional overlap between SigB and CtsR was found. The clpC, clpP and clpB operons are solely under CtsR control, whereas SigB alone is responsible for heat induction of clpL (Gertz et al., 2000; Chastanet et al., 2003). Dual regulation of heat shock gene regulation by SigB and CtsR does not mean that a SigB stimulus is sufficient for an effective induction of transcription while CtsR is still active as a transcriptional repressor. Such a scenario leads to only a slight induction of the dually regulated clpC operon in B. subtilis, for example during different SigB activating stress conditions such as glucose limitation or ethanol stress (Kr€ uger et al., 1994). SigB activity alone does not lead to an appropriate induction of the target genes because the impact of CtsR repression on transcription is super-ordinated, and thus CtsR inactivation is stringent for

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 123 adequate expression. The major task of SigB for dually regulated genes maybe is to enhance the induction of transcription when CtsR is inactivated. Heat shock class one in B. subtilis is regulated by the global repressor HrcA (heat shock regulation by CIRCE) (Zuber and Schumann, 1994; Yuan and Wong, 1995) and regulates only two operons coding for the classical molecular chaperones dnaK and groE (Homuth et al., 1997). In B. subtilis and close relatives, the HrcA and the CtsR regulon are distinct from each other. However, in other groups of low GC, Gram+ bacteria, CtsR is also able to regulate expression of HrcA-dependent genes (Chastanet et al., 2003). In most cases, HrcA-regulated genes are under dual control of both transcriptional regulators, and only in Streptococcus salivarius and Streptococcus thermophilus clpP is in addition to CtsR under HrcA control (Chastanet and Msadek, 2003). In S. aureus both HrcA-dependent operons also belong to the CtsR regulon, whereas in the order of Lactobacillales only the groE operon stands under dual regulation (Chastanet et al., 2003). Moreover, in species where one of the two global heat shock repressors is absent, the other has taken over the regulation of both regulons. For example, CtsR is the global stress repressor in Oenococcus oeni (Grandvalet et al., 2005), a lactic acid bacterium missing hrcA in its genome. In contrast, HrcA is the corresponding regulator of clp expression in the L. acidophilus group, which lack ctsR in their genome (van de Guchte et al., 2006) (Fig. 1). In contrast to dual regulation by a transcriptional ‘activator’ such as SigB and a repressor such as CtsR the interplay between two transcriptional repressors seems odd. The major effect of this crosstalk is that basal transcription under repressor active conditions is lower compared to expression when only one repressor is responsible for regulation (Chastanet et al., 2003). HrcA activity is directly dependent on protein stress (Mogk et al., 1997), whereas the CtsR repressor senses different stress inputs (Elsholz et al., 2010, manuscript submitted) and is also inactivated during virulence-related stress conditions. This gives space for speculation that dual regulation of the HrcA regulon ensures that the molecular chaperones, which are also crucial for virulence (see Section 3), are strongly induced during both specific stress and virulence conditions (Chastanet et al., 2003). The ctsR gene itself stands under its own control and is auto-regulated (Derre et al., 1999b). The ctsR gene is co-transcribed with clpC in an operon for all low GC, Gram+ bacteria and the AAA+ protein ClpC belongs to the CtsR regulon in all cases when ctsR gene is present in a genome. In the order Bacillales of low GC, Gram+ bacteria the clpC operon is tetra-cistronic and, in addition to ctsR and clpC, also encodes the two modulators of CtsR activity, mcsA and mcsB. However, genes for these two modulators are absent in the order of Lactobacillales (Varmanen et al., 2000;

ALEXANDER K.W. ELSHOLZ ET AL.

Figure 1 Graphical presentation of the distribution of CtsR-regulated proteins for different low GC, Gram+ species and known overlaps with other regulators such as SigB or HrcA. An arrow indicates specific influence of transcriptional regulator for the expression of the corresponding gene. Proteins whose expression is solely dependent on CtsR activity are depicted in red, whereas proteins that are dually regulated by CtsR and SigB (green) are shown in blue. Proteins controlled by CtsR as well as HrcA are presented in grey, and proteins that are regulated only by HrcA are shown in purple (See Colour Plate Section).

124

[(Figure_1)TD$IG]

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 125 Chastanet et al., 2001). In these species, ctsR is co-transcribed only with clpC in a bi-cistronic operon. The proteolytic component of the ATP-dependent protease ClpP is also a permanent member of the CtsR regulon in all low GC, Gram+ bacteria, as long as the ctsR gene is present in the corresponding genome. With the exception of S. salivarius and S. thermophilus, where dual regulation with CtsR and HrcA is present (Chastanet and Msadek, 2003), expression of the clpP operon stands solely under control of CtsR. In low GC Gram+ bacteria, clpP is mostly encoded by a mono-cistronic gene (Gerth et al., 1996), but in O. oeni clpP can be also co-transcribed with clpL (Beltramo et al., 2004). The molecular chaperones ClpB and ClpL lack an IGF loop and thus are probably not able to interact directly with the proteolytic component ClpP (Kim et al., 2001; Frees et al., 2004; Weibezahn et al., 2004). ClpB and ClpL are not present in the families Bacillaceae and Listeriaceae, but can be found in Staphylococcaceae and more often in Lactobacillaceae. Both genes stand under CtsR control with the exception of clpL in staphylococci (Gertz et al., 2000). ClpE, another Hsp 100/Clp protein, always stands under CtsR control as far as both clpE and ctsR genes are present in the genome. Interestingly, new studies have revealed so far unknown and not typical members of the CtsR regulon. In Lactobacillus plantarum the ATP-dependent membrane-bound AAA+ protease FtsH and the small heat shock protein Hsp1 are under direct CtsR control (Fiocco et al., 2008, 2010). In addition, a second small heat shock protein, Hsp18, was found regulated by CtsR in O. oeni (Grandvalet et al., 2005). In O. oeni, a CtsR consensus sequence was surprisingly found in front of the clpX operon, but no influence of CtsR was detected for clpX transcription (Grandvalet et al., 2005). It was speculated that this binding site was an evolutionary remnant. Nevertheless, regulation of clpX expression remains unclear for all low GC, Gram+ bacteria, and to date, no CtsR-dependent regulation has been shown for ClpX.

3. CELLULAR FUNCTIONS OF GENES REGULATED BY CtsR The physiological function of the CtsR regulon members lies not only in their important role in protein quality control and the resulting effect in stress adaptation and general cellular processes, but also in their specific role in regulated degradation of key regulators for essential cellular and physiological programmes in response to temporal, spatial or environmental stimuli. Not only the specific regulated proteolysis of transcription factors is

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important, but also the re-arrangement of the complete proteome after a specific stress input to attain a better adaptation of the cell (Sauer et al., 2004; Neher et al., 2006). However, specific adaptor proteins are needed for Clpspecific degradation. To date, only a few adaptor proteins are known for the low GC, Gram+ model organism B. subtilis, but additional adaptor proteins must exist (Kirstein et al., 2009), suggesting that induction of the Clp proteolytic machinery by inactivation of CtsR alone is not sufficient for regulated degradation, with the exception of McsB-dependent proteolysis. Thus, the CtsR regulon seems to be embedded into stress-specific modulons, where additional regulatory mechanisms induce expression of stress-specific adaptor proteins allowing a concatenated stress response. Consequently, regulated protein degradation seems to be under multiple control. The peptidase ClpP is the proteolytic component of the Clp degradation machinery and is mainly involved in (1) protein quality control (degradation of irreversibly damaged or ssrA-tagged proteins) (Kr€ uger et al., 2000; Wiegert and Schumann, 2001); (2) specific degradation of regulatory proteins such as CtsR, ComK, SpoIIAB, Spx, DegU-P or RsiW (Turgay et al., €ger et al., 2001; Pan et al., 2001; Nakano et al., 2002; Zellmeier et al., 1998; Kru 2006; Ogura and Tsukahara, 2010); and (3) unemployed proteins such as biosynthetic enzymes, which are no longer needed in starving cells such as MurAA, IlvB, PurF or PyrB (Kock et al., 2004a; Gerth et al., 2008) and also for virulence of pathogenic low GC, Gram+ bacteria (Frees et al., 2007). Synthesis of clpP is induced during several environmental stress conditions that causes damage to the protein structure (Gerth et al., 1998, 2004; Frees and Ingmer, 1999; Gaillot et al., 2000; Chastanet et al., 2001; Frees et al., 2003a). Consequently, clpP inactivation in low GC, Gram+ bacteria leads to a pleiotropic mutant phenotype indicating the extraordinary role of the Clp machinery for protein quality control in low GC, Gram+ bacteria (Gerth et al., 1998; Msadek et al., 1998). A B. subtilis clpP mutant failed to activate specific developmental programmes due to failure in regulated degradation of key regulators, resulting in a deficiency in competence development and sporulation (Msadek et al., 1998). Both deficiencies depend on ClpCP-dependent degradation of either the master regulator of competence development ComK (Turgay et al., 1998) or the anti-sigma factor SpoIIAB, which regulates the sporulation-specific sigma factor s F in the forespore (Pan et al., 2001). A clpP mutant is more sensitive against heat, oxidative or low pH stress in Lactococcus lactis (Frees and Ingmer, 1999), L. monocytogenes (Gaillot et al., 2000), Streptococcus pneumoniae (Chastanet et al., 2001), Streptococcus mutans (Lemos and Burne, 2002), S. aureus (Frees et al., 2003a) and O. oeni (Beltramo et al., 2004). ClpP is also responsible for the

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 127 controlled degradation of the MazE antitoxins and probably all other antitoxins in S. aureus (Donegan et al., 2010). In L. lactis, activity of the transcriptional stress repressor HdiR (heat and DNA damage-inducible regulator) is controlled by ClpP-dependent degradation (Savijoki et al., 2003). Finally, ClpP is also involved in the virulence of pathogenic low GC, Gram + bacteria. ClpP is essential in a mouse infection model and for intracellular survival of L. monocytogenes (Gaillot et al., 2000), in a murine skin abscess model for S. aureus (Frees et al., 2003a), for survival of S. pneumoniae in mice (Robertson et al., 2002) and for expression of virulence genes in S. mutans (Kajfasz et al., 2009). In the last few years, different roles for ClpP in the expression of virulence factors were described. In L. monocytogenes, ClpP affects expression of listeriolysin O (Gaillot et al., 2000) and SvpA (Borezee et al., 2001). In S. aureus, ClpP influences expression of extracellular virulence factors such as hla (Frees et al., 2003a). In S. pneumoniae (Robertson et al., 2002; Kwon et al., 2003; Ibrahim et al., 2005) and S. mutans (Kajfasz et al., 2009), it was demonstrated that ClpP influences virulence gene expression. Based on the important cellular and physiological function of ClpP for stress adaptation and virulence of pathogenic bacteria, ClpP seems to be a €tz-Oesterhelt et al., promising target for newly developed antibiotics (Bro 2005; B€ ottcher and Sieber, 2008). ClpP needs a Clp ATPase to perform efficient protein degradation. In low GC, Gram+ bacteria only ClpX, ClpC and ClpE are able to perform this function, but only clpC and clpE are under CtsR control. Interestingly, B. subtilis ClpC needs specific adaptor proteins for oligomerisation and ATPase activity (Kirstein et al., 2006), which are needed for an efficient ClpCP activity. Contrarily, B. subtilis ClpE possesses an intrinsic ATPase activity, which depends on a putative zinc finger (Miethke et al., 2006). A clpC mutant exhibits thermosensitivity in B. subtilis (Msadek et al., 1994), S. aureus (Frees et al., 2003a) and L. monocytogenes (Rouquette et al., 1996), but a clpC deletion in L. lactis (Ingmer et al., 1999) and S. pneumoniae (Chastanet et al., 2001) did not affect growth at higher temperatures. Opposite observations were made for ClpE. A clpE mutant has no obvious phenotype in B. subtilis (Derr e et al., 1999a), whereas a clpE deletion in L. lactis (Ingmer et al., 1999) or S. pneumoniae (Chastanet et al., 2001) has a severe thermosensitive phenotype. One could speculate that this observation is linked to the occurrence of the specific heat-activated ClpC adaptor McsB, which is present in the order of Bacillales but is absent in Lactobacillales. ClpC is the major ATPase for protein turnover and regulated degradation in B. subtilis (Kr€ uger et al., 2000; Gerth et al., 2008) and a corresponding

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mutant has a severe phenotype (Kr€ uger et al., 1994; Msadek et al., 1994). Therefore, degradation of key regulators or ‘unemployed’ proteins such as SpoIIAB, ComK, MurAA and GlmS are mainly performed by the ClpCP system (Turgay et al., 1998; Pan et al., 2001; Kock et al., 2004a; Gerth et al., 2008). ClpC is also important in other low GC, Gram+ bacteria and a clpC mutant thus has also a severe phenotype in L. monocytogenes (Rouquette et al., 1998), S. pneumoniae (Charpentier et al., 2000) and S. aureus (Frees et al., 2004; Chatterjee et al., 2005). ClpC is involved in the virulence of pathogenic bacteria such as L. monocytogenes (Rouquette et al., 1996, 1998; Nair et al., 2000b), S. aureus (Frees et al., 2004) and S. pneumoniae (Charpentier et al., 2000). Similar to ClpP, ClpC also affects expression of virulence factors such as SvpA (Borez ee et al., 2001) and listeriolysin O (Rouquette et al., 1998) in L. monocytogenes or general virulence expression in S. pneumoniae (Ibrahim et al., 2005). In B. subtilis, clpE does not show a severe phenotype (Derre et al., 1999a) and a specific function for ClpE in B. subtilis could only be linked to CtsR (Miethke et al., 2006). Deletion of clpE affects the re-repression of CtsRdependent transcription, suggesting a role for ClpE in the re-activation of CtsR in B. subtilis (Miethke et al., 2006) and L. lactis (Varmanen et al., 2003). ClpE also participates in the degradation of CtsR, partially in B. subtilis (Miethke et al., 2006) and fully in L. lactis (Elsholz et al., manuscript submitted). In other low GC, Gram+ bacteria a clpE mutant is deficient in stress survival in L. monocytogenes (Nair et al., 1999), L. lactis (Ingmer et al., 1999) and S. pneumoniae (Chastanet et al., 2001). In addition, ClpE is also needed for virulence of L. monocytogenes (Nair et al., 1999) and S. pneumoniae (Zhang et al., 2009). Effects of clpL or clpB knock-outs in S. aureus can only depend on chaperone activity because ClpL and ClpB lack the IGF loop which is crucial for interaction with ClpP (Kim et al., 2001). A clpL mutant displays a heatsensitive phenotype and decreased virulence for S. pneumoniae (Kwon et al., 2003) and S. mutans. ClpL is involved in stress tolerance and viability (Kajfasz et al., 2009). ClpB in L. monocytogenes is needed for virulence and induction of thermotolerance, but not for other stress conditions (Chastanet et al., 2004). In S. aureus a clpB mutant showed decreased virulence and thermotolerance (Frees et al., 2004). The molecular chaperones DnaK and GroEL are highly conserved among pro- and eukaryotes (Craig, 1985), and are important for stress resistance. In S. aureus, both proteins are important for virulence (Qoronfleh et al., 1993, 1998). Deletion of either dnaK or groEL leads to reduced virulence and stress sensitivity (Singh et al., 2007). S. mutans dnaK and groEL are needed for general stress tolerance (Lemos et al., 2007).

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 129 The two modulators McsA and McsB in Bacillales were shown to act as an adaptor complex for ClpC (Kirstein et al., 2007). Recently, it has also been demonstrated that the McsAB adaptor complex mediates the delocalisation of competence proteins from cell poles in B. subtilis (Hahn et al., 2009). Inactivation of CtsR itself leads to an enhanced stress tolerance according to the increased expression of the protein quality control proteins in L. monocytogenes (Nair et al., 2000a; Karatzas et al., 2003), L. lactis (Varmanen et al., 2000), S. thermophilus (Zotta et al., 2009) and Lactobacillus sakeii (H€ ufner et al., 2007; H€ ufner and Hertel, 2008), where the utilisation of a ctsR mutant represents an advantage for industrial production in a fermenter.

4. MECHANISMS FOR THE INACTIVATION OF THE CtsR REPRESSOR 4.1. Heat Inactivation of CtsR CtsR, the first gene of the clpC operon, was described to contain a putative helix-turn-helix (HTH) DNA binding motif in low GC, Gram+ bacteria (Kr€ uger et al., 1997). Later, CtsR was identified as the corresponding repressor for clpC and clpP expression in B. subtilis. CtsR directly binds to the promoters and recognises the direct heptanucleotide repeat sequence (A/ GGTCA A ANANA/GGTCA A A) (Derr e et al., 1999b). This consensus sequence is highly conserved among low GC, Gram+ bacteria. Moreover, the ctsR gene itself is also highly conserved. However, CtsR is very specific for low GC, Gram+ bacteria because no homologues were found in either Gram-negative bacteria or the actinobacteria branch. An initial characterisation in B. subtilis revealed that CtsR is only active as a dimer (Derre et al., 2000). The 2nd and 3rd genes of the tetra-cistronic clpCoperon in B. subtilis were renamed McsA and McsB (modulators of CtsR activity) according to their influence on CtsR activity. McsB shows homology to ATP:guanidino phosphotransferases such as arginine kinases and inhibited CtsR DNA binding in vitro. 2D Western analysis suggested that CtsR €ger et al., 2001). became phosphorylated by McsB during heat stress (Kru McsA, a putative zinc finger protein, was shown to be essential for CtsR activity in vivo because CtsR is no longer active in a mcsA mutant. Furthermore, it became clear that CtsR is degraded in ClpCP-dependent manner during heat stress (Kr€ uger et al., 2001). ClpE is needed for an efficient and timely re-repression of CtsRdependent transcription. In a L. lactis clpE mutant, re-repression was

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delayed, but still occurred with a time lag. Therefore, ClpE does not seem solely responsible for the reconstitution of CtsR activity. In addition, the zinc finger of ClpE was identified to be crucial for re-repression (Varmanen et al., 2003). This observation was later also confirmed for B. subtilis. Moreover, the zinc finger of ClpE was linked to the autonomic ATPase function of ClpE (Miethke et al., 2006), as was also shown for ClpX in E. coli (Banecki et al., 2001). McsB was characterised as a tyrosine kinase in vitro that specifically phosphorylates McsA, CtsR, ClpC as well as itself (Kirstein et al., 2005). However, McsB kinase activity needs McsA as an activator for efficient phosphorylation. Specific phosphorylation sites for McsB, tyrosine residue 155 and 210, as well as tyrosine residues for McsA were described. Inter(McsB!McsA) and intramolecular (McsB!McsB) phosphate transfer was observed. The low-molecular-weight protein tyrosine phosphatase (LMWPTP) YwlE was identified as the cognate McsB phosphatase that dephosphorylates CtsR, McsA, McsB as well as ClpC in vitro. Furthermore, McsB is the only direct interaction partner of CtsR and McsBdependent release of CtsR from the DNA in vitro is strongly enhanced when McsB is active as a kinase. Additionally, ClpCP-dependent CtsR degradation during heat stress is also dependent on McsA and McsB. Based on the observation that ClpC inhibits McsB activity and this repression is restricted when protein aggregates are present, a titration model was postulated (Kirstein et al., 2005). Such a titration model is also known for other heat shock regulators such as s32 (Bukau, 1993) or HrcA (Mogk et al., 1997), and seems to be an elegant model for regulation of CtsR activity as well (Fig. 2). A recent in vitro study with CtsR and McsB from Bacillus stearothermophilus revealed that McsB is an arginine kinase instead of a tyrosine kinase (Fuhrmann et al., 2009). Furthermore, the structure of CtsR binding to its DNA operator was solved. It was shown that CtsR is a winged HTH protein where the HTH domain binds in the major groove of the DNA whereas the b-hairpin wing grabs into the minor groove. It was also demonstrated that McsB phosphorylates CtsR on specific arginine residues within the winged HTH region in vitro. Because the determined phospho-sites are also involved in DNA binding of CtsR, it was thought that McsB is only able to bind and phosphorylate free CtsR (Fuhrmann et al., 2009). Such a model supports only a transient induction of gene expression, but not rapid and strong induction within 1–2 min, as was observed for CtsRdependent transcription during heat stress (Kr€ uger et al., 1994). Exclusive phosphorylation of free CtsR would result in slow enhancement of transcription peaking long after the stress input because inactivation of CtsR would depend on prior dissociation of DNA-bound CtsR. In addition, it has long

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 131

[(Figure_2)TD$IG]

Figure 2 Graphical presentation of regulation and degradation of CtsR in the different low GC, Gram+ orders under specific stress conditions. (a) CtsR inactivation and degradation during heat stress in the order Bacillales. Under control conditions CtsR binds to its DNA operator and McsB kinase is repressed by binding to ClpC. Elevated temperatures lead to a loss of CtsR DNA binding. In addition, McsB is released from ClpC and becomes activated as a kinase by McsA. This activation results in targeting of free CtsR for a ClpCP-mediated proteolysis. (b) CtsR inactivation and degradation during heat stress in the order Lactobacillales that misses the two modulators McsA and McsB. Under control conditions CtsR binds to its DNA operator. Elevated temperatures lead to a loss of CtsR DNA binding and ClpEPmediated degradation. (c) CtsR inactivation during oxidative stress conditions in the order Bacillales. Under control conditions CtsR binds to its DNA operator and McsB kinase is repressed by binding to ClpC. During oxidative stress critical thiols are oxidised within the oxidative stress sensor protein McsA. This oxidation disturbs interaction of McsA/McsB and activates a specific McsB function. Consequently, McsB is now able to bind and inactivate DNA-bound CtsR, but CtsR cannot be degraded due to the inactive McsB kinase. (d) CtsR inactivation during oxidative stress in the order Lactobacillales. Under control conditions CtsR binds to its DNA operator. Oxidative stress leads to oxidation of critical thiols within ClpE. This oxidation causes an activation of ClpE which is now able to target DNA-bound CtsR (See Colour Plate Section).

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been known that the two modulators McsA and McsB are absent in the Lactobacillales order (Varmanen et al., 2000; Lemos and Burne, 2002). Recently, Elsholz and co-workers revealed that McsB is not involved in the regulation of CtsR activity during heat stress in vivo, supporting a hypothesis that CtsR activity is identically regulated in all low GC, Gram+ bacteria. It was suggested that CtsR is able to sense and respond to elevated temperatures by acting as a protein thermometer in all low GC, Gram+ bacteria, which adjusts its activity intrinsically to the surrounding temperatures. CtsR is able to bind to its cognate DNA operator sequence with high affinity under control conditions, but DNA binding under heat shock conditions is dramatically reduced. Furthermore, the CtsR protein is adapted to the ecological niche of the specific low GC, Gram+ bacteria, and thus can respond to the very species-specific heat shock temperatures (Elsholz et al., 2010). The auto-inactivation of CtsR is not an on/off decision, where CtsR activity is switched off above a specific temperature. CtsR is only partially inactivated at modest heat temperatures and becomes fully inactivated only at maximal heat stress temperatures, as observed for CtsR from B. subtilis €ger et al., 1994), S. aureus (Fleury et al., 2009) and L. lactis (Elsholz (Kru et al., 2010). This mechanism secures that only the needed amounts of heat shock proteins become expressed under specific circumstances and excludes that an excess of protein quality systems are expressed under a modest protein stress which would waste important cellular resources. The b hairpin wing with its tetra-glycine loop was identified as the region that is responsible for thermosensing of CtsR in all low GC, Gram+ bacteria (Elsholz et al., 2010). This loop occupies an outstanding position as it is directly located at the tip of the wing implicating a potential regulatory role (Fuhrmann et al., 2009). It has long been known that glycine residues exhibit great conformational freedom and can adopt a conformation to a much wider range than the other amino acid residues due to their free phi and psi angles (Matthews et al., 1987). As a result, glycine residues have more backbone conformational flexibility, which increases the configurational entropy of the denatured state. Therefore, more free energy is needed to maintain the structural integrity of the native state. This effect results in decreased thermostability of the protein (Elsholz et al., 2010).

4.2. CtsR Inactivation During Oxidative Stress It has long been known that, in addition to heat stress, CtsR is also inactivated during other stress conditions such as disulfide stress (Leichert et al.,

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 133 2003), H2O2 (Mostertz et al., 2004), carbonyl electrophiles (Nguyen et al., 2009) and low pH (Frees et al., 2003b). As long as a titration model regarding McsB was applicable, the induction mechanism seemed to be clear and unique because all these stress conditions should result in damage of protein structure which may lead to protein aggregation (Hartl and Hayer-Hartl, 2009). However, since the observation that CtsR is an intrinsic heat sensor (Elsholz et al., 2010), the assumption that a common mechanism for CtsR inactivation exists for all stress conditions turned out to be obsolete. Thus, dual activity control of CtsR activity must have evolved for CtsR inactivation during different stress conditions. This model is also underlined by the fact that CtsR-dependent transcription differs significantly during oxidative stress and heat stress (Elsholz et al., manuscript submitted). A new mechanism for CtsR inactivation during oxidative stress was uncovered that depends solely on McsB, but not on its kinase activity. McsB activity during oxidative stress is regulated by McsA, which acts as a redox-sensing protein to adjust McsB function. Not only CtsR activity is dually regulated, but also McsA acts as a dual regulator of McsB. Under control and heat stress conditions, McsA and McsB are associated, and McsA can activate McsB kinase as long as McsB is not inhibited by ClpC. On the other hand, McsA can also act as an inhibitor of McsB, and McsB ceases to inactivate DNA-bound CtsR, as long as both proteins are associated. Under oxidative stress conditions, the critical thiols of the second zinc finger of McsA become oxidised. As a result, these thiols act as a molecular redox switch to regulate the activity of McsB by decreasing the interaction of McsA with McsB. When McsB is free of McsA, it is able to bind and inactivate DNA-bound CtsR, resulting in induced transcription of target genes. But due to the inactive McsB kinase, CtsR is not degraded. McsA is processed in a ClpCP-dependent manner, with the essential help of the adaptor protein McsB, resulting in a cleaved McsA protein, which may prevent binding to McsB. Interestingly, reversible oxidation of McsA seems to be required for correct cleavage of McsA (Elsholz et al., manuscript submitted). McsA, with its crucial cysteine residues, is a redox-sensing protein which specifically regulates the activity of McsB and thus is responsible for a modest response to oxidative stress. Yet the oxidative stress sensing complex McsA/B is not present in the order of Lactobacillales. The observation that CtsR-dependent transcription in L. lactis is strongly induced during oxidative stress suggests that another mechanism for CtsR inactivation exists. It was found that ClpE is essential for CtsR inactivation during oxidative stress in L. lactis, revealing a new ClpE-mediated inactivation mechanism for CtsR. Remarkably, L. lactis clpE expression is

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relatively higher under control conditions when compared with B. subtilis to secure that ClpE is always present and can act as an redox-sensor protein (Elsholz et al., manuscript submitted). This is due to the fact that the number of L. lactis CtsR binding sites in front of clpE is lower compared with B. subtilis (Varmanen et al., 2000). It was demonstrated that the previously described thiol-dependent cleavage of ClpE by ClpP (Varmanen et al., 2003) depends on the reversible oxidation of the critical thiols. Interestingly, this cleavage was not observed for B. subtilis ClpE, suggesting that L. lactis ClpE specifically gained this feature in the course of evolution (Elsholz et al., manuscript submitted). Signal transduction during oxidative stress resulting in CtsR inactivation is mediated by different signalling pathways in low GC, Gram+ bacteria, dependent on the presence or evolutionary development of the redoxsensing proteins. Nevertheless, both aforementioned mechanisms depend on critical thiols, which act as nano-switches to adjust protein activity in order to counteract oxidative stress (Elsholz et al., manuscript submitted). Both pathways are also completely different in comparison to heat inactivation, demonstrating dual activity control of CtsR, which may be important for appropriate adaptation to different stress conditions.

4.3. CtsR Inactivation During Other Stress Conditions Two distinct mechanisms are known for the inactivation of CtsR, a temperature-dependent one for the response to elevated temperatures and a thioldependent one for sensing of oxidative stress. Nevertheless, a few stress conditions which result in CtsR inactivation cannot be explained sufficiently by these two mechanisms. Inactivation of CtsR by protein stress caused by specific antibiotics such as puromycin (Kr€ uger et al., 1996; Frees and Ingmer, 1999) or low pH (Frees et al., 2003b) is not explainable by the two identified mechanisms. Neither a heat-sensing domain nor critical thiols are known to respond to protein aggregates. CtsR inactivation during these conditions may depend on activation of McsB kinase by protein aggregates. However, one could also speculate that an additional mechanism has become established for these conditions. Interestingly, induction of CtsR-dependent protein quality control systems in L. monocytogenes under hydrostatic pressure was shown not to depend on a transcriptional signalling cascade, but is regulated at the genomic level by genetic variation. Most of the isolated L. monocytogenes spontaneous high-hydrostatic pressure (HHP)-tolerant mutants bear a mutation in a specific region of CtsR which results in an inactive CtsR protein

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 135 (Karatzas et al., 2003). The mutated region codes for the b hairpin wing and the mutations are located in the tetra-glycine loop. This loop is encoded in L. monocytogenes by a short sequence repeat of GGT and all mutations were located within this DNA repeat (Karatzas et al., 2005). Such hot spots of genetic variation, where Rec-independent mutations accumulate at high frequency, have been known for a long time. DNA repeats are very common in this group and most likely cause strand slippage of the DNA polymerase, thus promoting genetic variability (Treangen et al., 2009). It was demonstrated for E. coli that such tandem repeats are very common in stress response genes to ensure a moderate stress survival (Rocha et al., 2002). In addition, such specific DNA repeats in virulence genes are of tremendous relevance for the adaptation of pathogenic bacteria to their specific host (Moxon et al., 2006). As noted above, a ctsR mutant displays an increased stress tolerance due to the permanent induction of the protein quality control genes. Thus, transient gene silencing of ctsR would enhance fitness and stress tolerance and give the bacteria an advantage in the adaptation to a new ecological niche and their specific conditions. In fact, inactivation of L. monocytogenes ctsR by a single codon deletion enhances the stress tolerance and decreases the virulence (Karatzas et al., 2003), most probably to escape the immune system and persist within the host. It seems likely that evolution has allowed an emergency exit for low GC, Gram+ bacteria to cope with conformational stress conditions when activation of protein quality control genes through regular transcriptional regulation of CtsR is not applicable. As a consequence, a regulatory layer at the genomic level was introduced to give the bacteria an opportunity to escape stress conditions by increasing the amount of protein quality systems. Hence, specific mutations accumulate in a DNA repeat within the ctsR gene during evolutionary pressure to inactivate CtsR and to give the bacteria a growth advantage, ensuring that a subset of cells survive the stress. However, such a stress-specific genetic mutation is only shown for piezo-tolerant SCV of L. monocytogenes (Karatzas et al., 2003) and not for other low GC, Gram+ bacteria. In addition, in piezo-tolerant SCV of S. aureus no mutations for CtsR were detected (Karatzas et al., 2007), and the DNA repeats of the tetra-glycine domain in ctsR are not so conserved in low GC, Gram+ bacteria as at the protein level (Karatzas et al., 2005). To date, only these two studies have been reported, but this could be a widely used mechanism. Therefore, additional genetic studies must be performed to investigate more protein stress conditions in different low GC, Gram+ bacteria.

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5. CONTROL OF CTSR DEGRADATION BY THE REGULATED ADAPTOR McsB Controlled degradation of key transcriptional regulators plays a critical role in many bacterial regulatory circuits (Gottesman, 2003) However, CtsR degradation is not needed for induction of ctsR-dependent genes. In vivo and in vitro experiments demonstrated that CtsR is a substrate for the ClpCP protease (Kr€ uger et al., 2001), but CtsR is very stable under control conditions and only becomes degraded under stress conditions such as heat or puromycin treatment (Kr€ uger et al., 2001). It was revealed that the two modulators of CtsR activity, McsA and McsB, in B. subtilis are necessary for CtsR degradation during heat stress (Kirstein et al., 2005). It was shown by in vitro experiments that specifically McsB kinase is essential for CtsR degradation, but CtsR phosphorylation is not sufficient for CtsR degradation (Kirstein et al., 2007). Additionally, YwlE was identified as the cognate phosphatase of the McsB kinase in vitro (Kirstein et al., 2005). Recently, it has been demonstrated that McsB kinase activity is needed for an efficient CtsR degradation in vivo, underlining the role of McsB as an adaptor for ClpC. CtsR degradation depends on the activation of McsB as an adaptor protein by auto-phosphorylation and not on a direct CtsR phosphorylation. Only phosphorylated McsB is able to bind CtsR. Consequently, a sophisticated regulatory two-step mechanism for CtsR degradation was uncovered as CtsR is only accessible for McsB when it is not bound to its DNA operator, and McsB can only target free CtsR when heat activated as an adaptor protein (Elsholz et al., 2010). YwlE was shown to counteract McsB adaptor activity in vivo. When, dephosphorylation of McsB by YwlE failed, McsB adaptor activity is shutdown rapidly by degradation when phosphorylated. This regulatory switch prevents degradation of re-activated CtsR (Elsholz et al., 2010). CtsR was also shown to be rapidly degraded during heat stress in the order of Lactobacillales, where mcsA and mcsB are not present. CtsR degradation in these bacteria was shown to depend on ClpE (Elsholz et al., 2010), but the precise mechanism of CtsR targeting and degradation, that is whether ClpE is heat activated or not, remains to be uncovered.

6. SUMMARY AND OUTLOOK The regulation of CtsR activity and its controlled degradation has become one of the best-studied regulatory heat stress mechanisms in low GC, Gram+

REGULATION OF CtsR ACTIVITY IN LOW GC, GRAM+ BACTERIA 137 bacteria. In recent years, CtsR was established as a model system par excellence to study precise and fine-tuned regulatory mechanisms in molecular detail. Generally, elucidation of CtsR activity provides deeper insights into a fundamental, highly conserved and global bacterial stress response system. Nevertheless, ‘solved’ problems tend to raise more questions. Thus, a further characterisation of the structural details of how CtsR gets inactivated during heat stress and how CtsR is re-activated as a DNA binding repressor, as well as a molecular characterisation of CtsR degradation, remains an interesting subject for future studies. The identification of other protein stress conditions that lead to CtsR inactivation, as well as the underlying molecular mechanisms, will provide a deeper understanding of the function and role of CtsR in low GC, Gram+ bacteria. In addition, new members of the CtsR regulon will probably also contribute to the physiological important function of CtsR. One of the major challenges is to connect the different mechanisms pertaining to the regulation of CtsR activity and stability, to investigate why different mechanisms have become established for the same physiological function in the different low GC, Gram+ orders and to place all the information into a systems biology context.

ACKNOWLEDGEMENT €zel (South Dakota State Univ., USA) The authors are grateful to Volker Bro for critical reading of the manuscript and helpful comments.

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[(Plate_1)TD$FIG] Plate 1 Graphical presentation of the distribution of CtsR-regulated proteins for different low GC, Gram+ species and known overlaps with other regulators such as SigB or HrcA. An arrow indicates specific influence of transcriptional regulator for the expression of the corresponding gene. Proteins whose expression is solely dependent on CtsR activity are depicted in red, whereas proteins that are dually regulated by CtsR and SigB (green) are shown in blue. Proteins controlled by CtsR as well as HrcA are presented in grey, and proteins that are regulated only by HrcA are shown in purple. (For b/w version, see page 126 in the volume.)

(a)

(b)

[(Plate_2)TD$FIG]

(c)

(d)

Plate 2 Graphical presentation of regulation and degradation of CtsR in the different low GC, Gram+ orders under specific stress conditions. (a) CtsR inactivation and degradation during heat stress in the order Bacillales. Under control conditions CtsR binds to its DNA operator and McsB kinase is repressed by binding to ClpC. Elevated temperatures lead to a loss of CtsR DNA binding. In addition, McsB is released from ClpC and becomes activated as a kinase by McsA. This activation results in targeting of free CtsR for a ClpCP-mediated proteolysis. (b) CtsR inactivation and degradation during heat stress in the order Lactobacillales that misses the two modulators McsA and McsB. Under control conditions CtsR binds to its DNA operator. Elevated temperatures lead to a loss of CtsR DNA binding and ClpEPmediated degradation. (c) CtsR inactivation during oxidative stress conditions in the order Bacillales. Under control conditions CtsR binds to its DNA operator and McsB kinase is repressed by binding to ClpC. During oxidative stress critical thiols are oxidised within the oxidative stress sensor protein McsA. This oxidation disturbs interaction of McsA/McsB and activates a specific McsB function. Consequently, McsB is now able to bind and inactivate DNA-bound CtsR, but CtsR cannot be degraded due to the inactive McsB kinase. (d) CtsR inactivation during oxidative stress in the order Lactobacillales. Under control conditions CtsR binds to its DNA operator. Oxidative stress leads to oxidation of critical thiols within ClpE. This oxidation causes an activation of ClpE which is now able to target DNA-bound CtsR. (For b/w version, see page 133 in the volume.)

Author Index Abbas, B., 14, 30, 32 Abee, T., 131, 137 Abomoelak, B., 68, 90, 94 Abraham, A.-L., 137 Abranches, J., 129, 130 Abu-Soud, H.M., 86, 90 Adams, M.W., 73 Adams, M.W.W., 72, 73 Adegbola, R.A., 59, 84, 90 Adler, L., 63 Agarwal, A., 49, 54, 61, 68, 85, 86, 89 Agarwal, N., 91 Agashe, V.R., 122 Agogue, H., 29 Agrawal, P., 91 Aguilera, J.A., 80 Aharonowitz, Y., 79 Ahring, B.K., 32 Ahuja, E.G., 70 Alam, K.Y., 68 Alam, M.S., 91 Alawi, M., 11 Albracht, S.P.J., 72 Alexeeva, S., 64, 65, 68, 87 Alland, D., 80 Allen, S.S., 60, 84, 86 Alm, E.W., 6 Almering, M.J.H., 63 Alonso, S., 76, 86, 87 Altman, E., 52, 66, 68 Aluwihare, L.I., 6, 29 Alvarez, H.M., 67, 68, 90, 94 Alvarez-Ortega, C., 71 Alving, K., 85 Aly, S., 60, 84, 89 Amann, R., 4 Amarasingham, C.R., 53 Amelung, W., 23, 25, 28 Anaya, J.M., 83

Anderberg, S.J., 80 Anderson, R.J., 58, 60 Andrews, J., 47 Andrews-Pfannkoch, C., 34 Ansari, M.Z., 94 Ansell, R., 63 Antelmann, H., 135 Aoshima, M., 74, 75 Arai, H., 74 Arata, Y., 56 Aravind, L., 15, 16, 123 Arikawa, Y., 75 Aris, V., 59 Arora, P., 94 Arp, D.J., 16, 17 Arscott, L.D., 81 Artman, M., 48, 51 Aslund, F., 62 Asoh, S., 72 Asterinou, K., 131 Atlas, R.L., 65 Attarian, R., 80 Av-Gay, Y., 79–81 Baas, M., 13 Badcock, K., 90 Baden-Tillson, H., 6, 7, 10, 15, 16, 22, 34 Bagwell, C., 33 Bajpai, R., 91 Bakalarski, C.E., 128 Baker, T.A., 122, 123, 127, 128, 130 Balasubramanian, R., 96 Balla, G., 84, 86 Balla, J., 84, 86 Banaiee, N., 92 Bancroft, G.J., 84 Bandow, J.E., 129 Banecki, B., 132 Bange, F.C., 60, 84, 89

146 Bannantine, J.P., 59 Barbe, V., 124, 125 Barletta, R.G., 78 Barnes, P.J., 85, 90 Barrell, B.G., 90 Barry 3rd, C.E., 47–49, 52, 59, 61, 68, 75, 83, 87, 90, 95, 99, 100 Barry, C.E., 57 Barry, W.H., 50 Bartek, I.L., 75, 82, 83, 86, 87 Bartossek, R., 17, 18 Basaraba, R.J., 59 Basham, D., 90 Batto, J.-M., 124, 125 Baughn, A.D., 74–77 Bayer, K., 13 Bayliss, C., 137 Becker, P., 130 Becktel, W.J., 134 Beckwith, J., 62 Beeson, K., 34 B
ej a, O., 15, 16 Bekierkunst, A., 48, 51 Bellon, G., 59, 71, 85, 89 Beltramo, C., 125, 127, 128 Beman, J.M., 29, 30, 32, 33 Ben-Dov, E., 33 Benitez-Nelson, B.C., 6 Benjamin, I.J., 50, 97 Bennett, A.R., 60 Bennett, G.N., 66, 68, 87, 93 Bennik, M.H.J., 131, 137 Benoit, S.L., 73 Bensen, D.C., 15, 16 Bentley, W.E., 56 Berche, P., 128–131 Beretti, J.L., 129, 130 Berg, I.A., 19 Berger, J., 59, 71, 85, 89 Berks, B.C., 68, 69, 87, 89 Bermingham, E., 34 Berney, M., 73, 74 Bernhard, A.E., 9, 10, 12 Berrios-Rivera, S.J., 66, 68, 87

AUTHOR INDEX Berry, A., 81 Berthet, F.X., 59 Bertozzi, C.R., 52, 59, 68, 94, 95 Berube, P.M., 10 Besra, G.S., 59, 84, 90 Bessi eres, P., 124, 125 Bewley, C.A., 79 Beyer, D., 129 Bhakoo, K.K., 69 Billman-Jacobe, H., 80 Billoud, B., 72 Bintrim, S.B., 4 Birrer, P., 59, 71, 85, 89 Bischoff, M., 130 Bishai, W.R., 49, 90, 91 Blackwood, K., 68 Blakis, A., 30 Blanchard, J.S., 81 Blankenfeldt, W., 70, 72 Bloch, H., 51, 54, 57, 58, 60, 99 Blokker, P., 6 Blomberg, A., 63 Bloom, B.R., 91–93, 97 Bock, E., 11 Boechat, N., 60, 83 Boehm, A.B., 32 Boelsterli, U.A., 51 Bolisetty, S., 49, 54, 61, 68, 85, 86, 89 Bolla, J.M., 129, 130 Bolon, D.N., 122, 123, 128 Bolotin, A., 124, 125 Bonch-Osmolovskaya, L., 4, 11, 23 Bonecini-Almeida Mda, G., 60, 83 Bonilla-Rosso, G., 34 Boon, C., 82, 83 Boon, J.P., 30 Boor, K.J., 124 Borez
ee, E., 129, 130 Borland, C., 85 Boros, M., 49, 50, 97 Bortner, C.A., 130 Boschker, H.T., 6 Boshoff, H.I., 48, 49, 52, 57, 59, 61, 68, 75, 87, 95, 99, 100

AUTHOR INDEX Bossy, R., 124, 125 Botstein, D., 62 Bott, M., 73 B€ ottcher, T., 129 Bottomley, P.J., 24 Botzenhart, K., 59, 71, 85, 89 Boucher, B.J., 97 Boucher, R.C., 59, 71, 85, 89 Boumann, H., 13 Bourret, T.J., 68 Boussau, B., 20 Bowatte, S., 24, 26–28 Boyle-Yarwood, S.A., 24 Braff, J., 29 Brahimi-Horn, M.C., 60 Brandau, S., 60, 84, 89 Braun, R.D., 60 Bregenholt, S., 128, 129 Breinbauer, R., 70 Brezina, O., 48 Brian, P., 92 Bridger, S.L., 73 Briley, K., 131 Brindicci, C., 85, 90 Brink, M., 29 Brochier, C., 15, 16 Brochier-Armanet, C., 16, 17, 20, 22, 23 Brosch, R., 90 Br€ otz-Oesterhelt, H., 129 Brovkovych, V., 60 Brown, D., 90 Brown, L.A., 85, 90 Brown, P.O., 62 Bryant, D.A., 96 Bryk, R., 52, 75, 76 Bryson, K., 124, 125 Buchmeier, N., 80, 81 Buchmeier, N.A., 80 Buckel, W., 19, 94, 95 Buckley, D.H., 4 Buettner, G.R., 50, 78, 79 Buhrke, T., 72 Bukau, B., 127, 129, 132 Bunch, P.K., 56

147 Burgdorf, T., 72 Burghoorn, J., 128, 130 Burn, J.A., 95 Burne, R.A., 128, 130, 134 Burton, B.M., 122, 123, 128 Burton, R.E., 122, 123, 128 Buttner, M.J., 90–92, 95 Bzymek, K.P., 79, 80 Caceres, N.E., 78 Cadena, J., 83 Cadiz, V., 80 Calaycay, J., 60, 83 Camacho, L.R., 68, 87, 95, 99 Camarasa, C., 75 Cameron, K., 24, 26–28 Cameron, K.C., 26 Camien, M.N., 57 Cammack, R., 72 Campbell, J.W., 55 Capozzi, V., 127 Carmel-Harel, O., 62 Carrillo, J., 60, 84, 86 Carter, N., 127 Casali, N., 84 Casciotti, K.L., 29, 32 Castaing, J.P., 93 Cekici, A., 59, 71, 85, 89 Cellek, S., 60 Cevallos, M.A., 66, 67 Chaillou, S., 131 Chain, P.S.G., 16, 17 Chakravarty, S., 84 Chakravorty, S., 84 Chamberlain, N.R., 130 Chamberlin, L., 92 Chan, E.D., 60 Chan, J., 54, 59, 60, 68, 82, 84 Chan, P.P., 16, 17 Chan, W.T., 52, 54, 55 Charpentier, E., 130, 132, 134 Chastanet, A., 124, 125, 127–130 Chater, K.F., 90–92 Chatterjee, I., 130

148 Cheesman, M.R., 91 Chen, B., 52, 54, 55, 80, 94 Chen, J.Q., 33 Cheng, Y., 50, 97 Cherian, J., 52, 55 Cheung, A.L., 129 Chillingworth, T., 90 Cho, C.M., 34 Choi, A.M., 60, 85 Choi, M.-H., 129, 130 Chora, A., 84, 86 Christians, E.S., 50 Chu, D., 54, 59, 68, 82 Chung, K.F., 60, 83, 85 Chung, S.W., 60, 85 Church, M.J., 29, 30, 32 Church, M.K., 60 Churcher, C., 90 Claiborne, A., 78 Clark, D.P., 53, 56, 64, 65, 68 Clausen, T., 132, 134 Claverys, J.P., 127–130 Clough, G.F., 60 Cohen, G., 79 Cohen, R.E., 123 Cohn, M.L., 51 Colangeli, R., 80 Colbeau, A., 72 Cole, J., 87 Cole, S.T., 90 Coleman, D.C., 3 Collins, D.M., 91–93, 97 Collins, M., 127 Comhair, S.A., 90 Connell, P., 50 Connor, R., 90 Conrad, R., 23, 24, 26, 28 Converse, P.J., 84 Cook, G.M., 73, 74 Coolen, M.J., 14, 30, 32 Coolen, M.J.L., 14, 15, 30 Corper, H.J., 51 Cosma, C.L., 47, 48 Costa, K.C., 95 Costello, C.M., 71, 89

AUTHOR INDEX Coucheney, F., 125, 127 Couloux, A., 124, 125 Cox, A.G., 61, 86 Cox, J.S., 52, 54, 59, 61, 68, 82, 85, 86, 94, 95 Cox, Y., 85 Craig, E.A., 130 Creighton, M.M., 58, 60 Crick, D.C., 59 Cronan Jr., J.E., 55 Crossman, D.K., 49, 54, 61, 68, 85–87, 89, 90, 93–99 Crutz-Le Coq, A.-M., 131 Cunha-Rodrigues, M., 84, 86 Currenti, E., 49, 61, 76 da Silva, S.M., 73 Daff
e, M., 93 Daims, H., 11–13, 33 Daldal, F., 95 Damst
e, J.S.S., 6, 13–15, 30, 32 Dang, H., 13 Daniel, J., 68, 90, 94 Daniel, S.L., 89 Darrouzet, E., 95 Dartois, V., 49, 52, 59, 61, 100, 128 Darwin, K.H., 123 Das, T.K., 82, 84 Dasgupta, N., 82 Davidge, K.S., 61, 86 Davies, J., 79 Davies, R., 90 Davis, A.A., 4, 19 Davis, B.D., 53 Davis, N.K., 90 Dawes, E.A., 66, 67 Dawes, I.W., 62 Dawes, S., 59 Dawson, T.L., 50 de Chastellier, C., 130 de Graef, M.R., 64, 65, 68, 87 De Groot, H., 89 de la Torre, J., 16, 17, 29, 33 de la Torre, J.R., 9–13, 33 de Montellano, P.R., 83

AUTHOR INDEX de Nys, R., 13, 33, 34 de Sieyes, N.R., 32 Deb, C., 68, 90, 94 DeFlaun, M., 6 delCardayre, S.B., 80 DeLong, E.F., 4, 10–12, 15, 16, 19, 29, 30, 32, 33 den Hengst, C.D., 90, 95 Denizot, F., 128 Deonarain, M.P., 81 Deppenmeier, U., 70 Dequin, S., 75 DeRiemer, K., 83 Derr
e, I., 124, 125, 128–131 Dervyn, R., 124, 125 Deshane, J.S., 49, 54, 61, 68, 85, 86, 89 Devine, K., 129, 130 Devlin, K., 90 Dewhirst, M.W., 60 Di, H., 23, 24, 26–28 Di, H.J., 26 Dick, T., 49, 52, 59, 61, 76, 78, 82, 83, 86, 87, 89, 100 Dietrich, L.E., 70–72 Dijkhuizen, L., 78 Dimmeler, S., 50 Dimroth, P., 73 Dinasquet, J., 29 Ding, H., 53 Ding, Z., 50, 97 Doan, B., 62 Dobbek, H., 76 Dodsworth, J.A., 4, 95 Dolganov, G.M., 49, 54, 61, 68, 82, 83, 89 Donegan, N.P., 129 Donohue, T.J., 4 Doring, G., 59, 71, 85, 89 Dougan, D.A., 127–129, 131, 138 Dowd, C.S., 68, 87, 95, 99 Drake, H.L., 89 Driver, E.R., 59 Drobnica, L., 48 Drobnicova, I., 48 Druffel, E.R., 6, 29 Duan, X., 53

149 Dubey, V.S., 68, 90, 94 Dubnau, D., 128, 130, 131 Dubos, R.J., 48, 60, 61, 83 Duncan, K., 47 Dunn, M., 66, 67 Dunn, M.S., 57 Dweik, R.A., 60, 86, 90 Dwyer, T.J., 79 Eck, J., 15 Edson, N.L., 56, 57 Eglinton, T.I., 6 Eguiarte, L.E., 34 Ehlers, S., 60, 84, 89 Ehrlich, S.D., 124, 125 Ehrt, S., 49, 52, 59, 61, 68, 87, 91, 95, 99, 100 Eiamphungporn, W., 135 Eiglmeier, K., 90 Eisen, J.A., 6, 7, 10, 15, 16, 22, 34 Eisen, M.B., 62 Eisenberg, D., 59 Eiteman, M.A., 52, 66, 68 Ellington, M.J., 69 Emmerson, P.T., 127 Encarnacion, S., 66, 67 Endermann, R., 129 Engelmann, S., 124, 127 Enger, O., 4 Enomoto, K., 75 Eom, C.Y., 77, 84 Epiphanio, S., 84, 86 Erbacher, J., 6 Ernst, J.D., 92 Erzurum, S.C., 86, 90 Escuyer, V., 130 Esser, L., 95 Ettinger-Epstein, P., 13, 33, 34 Eum, S.Y., 60, 84, 86 Fahey, R.C., 79–81 Falcón, L.I., 34 Fang, F.C., 78 Farnia, P., 47 Farver, C., 90

150 Faucet, V., 75 Fauci, A.S., 47 Faulkner, D.J., 79 Feldman, R.A., 15, 16 Feltwell, T., 90 Feng, Z., 78 Ferguson, S.J., 68, 69 Fernandes, C.L.V., 73 Ferrari, M.R., 34 Ferreira, A., 84, 86 Ferry, J.G., 76, 77, 87 Fert, J., 124, 125, 127 Fiencke, C., 11 Findlay, K.C., 90 Fiocco, D., 127 Firth, A., 70, 71 Fitzgerald, M.X., 71, 89 Fitzpatrick, A.M., 85, 90 Flanagan, J.M., 123 Flardh, K., 90, 92 Flarsheim, C.E., 50 Fleury, B., 134 Florczyk, M.A., 49, 61, 76 Flynn, J., 49, 52, 59, 61, 100 Flynn, J.A., 84 Flynn, J.M., 122, 123, 128 Focks, A., 23, 25, 28 Foley, J., 129 Follis Jr., R.H., 61, 83, 84 Fontan, P., 59 Forterre, P., 20 Fothergill, J.L., 70, 71 Fournier, D., 93 Fouts, D.E., 6, 7, 10, 15, 16, 22 Fowler, A.V., 57 Fox, G.E., 13 Foxwell, N.A., 60 Fraczkowska, K., 127, 130 Francis, C.A., 29, 30, 32, 33 Frazier, M., 34 Fredrickson, J.K., 6 Freeman, J., 34 Freeman, K.H., 13, 32 Frees, D., 123, 127–130, 135, 136

AUTHOR INDEX Frehel, C., 130 Freitag, T.E., 26 Freundlich, J.S., 47 Frey, M., 72 Friedheim, E., 70 Friedland, G., 47 Friedman, R., 34 Friedrich, B., 72 Frimpong, I., 75, 83, 86, 87 Fu, Z., 129 Fuchs, G., 19, 74, 75 Fuhrman, J.A., 4, 6, 19 Fuhrmann, J., 132, 134 Fujii, T., 78 Fujiwara, S., 56 Furst, V.W., 60 Gagneux, S., 83 Gahan, C.G.M., 131, 137 Gaillot, O., 128, 129, 131 Gallardo, V., 34 Gallone, A., 127 Gandhi, N.R., 47 Gao, L., 33 Gao, X., 50, 97 Garcia, J.-L., 3 Garforth, S.J., 74–77 Garg, S., 91 Garg, S.K., 91 Garnier, T., 90 Garrett, R.A., 22 Garrido, E.O., 62 Garrigues, C., 15 Garsin, D.A., 128, 130 Garton, N.J., 59, 84, 90 Gas, S., 90 Gasch, A.P., 62 Gaston, B., 90 Gatfield, J., 91 Geenevasen, J.A., 13 Geertman, J.M., 64 Geiman, D.E., 91 Geng, J., 60, 83 Gengenbacher, M., 83, 89

AUTHOR INDEX Genghof, D.S., 79 Gennaro, M.L., 59 Gensini, G., 61, 75, 77, 83 Gentles, S., 90 Georgopoulos, C., 132 Gerbl, F.W., 33 Gerth, U., 123, 124, 127–130, 132, 138 Gertz, S., 124, 127 Gething, M.J., 122 Ghanavi, J., 47 Ghanny, S., 59 Ghyczy, M., 49, 50, 97 Gibrat, J.-F., 124, 125 Gicquel, B., 59 Giles, G.I., 91–93 Gilles-Gonzalez, M.A., 60, 83, 84 Gilmour, R., 129 Glockner, F., 4 Gokhale, R.S., 94 Golbeck, J.H., 96 Gollabgir, A., 16, 17 Gomez, J.E., 47, 90 Gomez, L.M., 83 Gonzales, J., 60, 84, 86 Gonzalez, G., 60, 83, 84 Gonzalez-Juarrero, M., 59 Goodman, R.M., 4 Goodwin, M.B., 57 Gopinathan, K.P., 48 Gordon, S.V., 90 Gores, G.J., 50 Gorovitz, B., 79, 80 Gossner, A., 89 Gottesman, S., 122, 123, 138 G€ otz, F., 89, 134 Govender, T., 47 Graber, J.R., 4 Granath, K., 63 Grandvalet, C., 125, 127, 128 Grant, C.M., 62, 63 Grant, R.A., 122, 123, 128 Grasemann, H., 89 Gray, C.T., 64, 65 Green, J., 78, 96

151 Gregoire, I.P., 84, 86 Gribaldo, S., 20 Grimaldi, C., 124, 125 Grode, L., 59 Grosset, J.H., 84 Grundmeier, M., 130 Guest, J.R., 53 Guidry, L., 68, 87, 90–99 Gustafsson, L., 63 Gustafsson, L.E., 85 Gutterridge, J.M.C., 53, 78, 79 Guzzo, J., 125, 127, 128 Gygi, S.P., 128 Haapanen, J.H., 61, 75, 77, 83 Hacker, J., 124, 127 Hackett, M., 95 Hadd, A., 15 Hahn, J., 128, 130, 131 Hall, S.R., 60, 85 Hallam, S.J., 16, 29, 33 Halliwell, B., 53, 78, 79 Halpern, A.L., 6, 7, 10, 15, 16, 22, 34 Hamann, C.W., 64 Hamlin, N., 90 Hammel, J., 90 Hammer, K., 129, 130, 132, 136 Handelsman, J., 4, 34 Hanschke, R., 123, 128, 129 Hansman, R.L., 6, 29 Harraghy, N., 130 Harrell, M.I., 49, 54, 59, 61, 68, 82–84, 89 Harris, D., 90 Hartl, F.U., 122, 123, 135 Hartling, J.A., 123 Hartmann, P., 91, 96 Harwood, C.R., 127 Harwood, C.S., 71, 77 Hashimoto, S., 78 Hatzenpichler, R., 11, 12, 20, 22, 23 Hayer-Hartl, M., 122, 123, 135 Hayes, J.M., 6 Hayes, L.G., 54, 60, 83 Hazbon, M.H., 80

152 Hazleton, E.B., 84 He, J., 23, 24, 26–28 He, J.Z., 26 Hecker, M., 123, 124, 127–132, 134–136, 138 Hedderich, R., 94, 95 Hedlund, B.P., 33 Heidelberg, J.F., 6, 7, 10, 15, 16, 22, 34 Heidelberg, K.B., 34 Heifets, L., 91 Helmann, J.D., 135 Hemp, J., 16, 17 Henninger, K., 129 Hentschel, U., 13, 33, 34 Heo, J., 77, 84 Herbaud, M.L., 128 Herfort, L., 30 Herman, B., 50 Hernandez, M.E., 70–72 Herndl, G.J., 6, 29, 30 Herrmann, M., 32, 130 Hersch, G.L., 122, 123, 128 Hertel, C., 131 Hett, E.C., 48 Heuer, H., 23, 25, 28 Hicks, R.E., 6 Higenbottam, T., 85 Hill, C., 131, 137 Hill, P., 127, 130 Hill, P.J., 128, 129 Hill, R.T., 34 Hinds, J., 47, 59, 84, 90 Hinojosa, R., 83 Hinzen, B., 129 Ho, J.L., 60, 83 Hoff, D.R., 59 Hoffart, L.M., 96 Hoffman, J., 6, 7, 10, 15, 16, 22 Hoffman, J.M., 34 Hoffmann, F., 13, 33, 34 Hoffner, S.E., 47 Hofreiter, M., 33 Hohmann, S., 63 Hol, W.G., 82

AUTHOR INDEX Holben, W., 6, 16 Holguin, F., 85, 90 Holroyd, S., 90 Hols, P., 127 Holtappels, M., 13, 33, 34 Homuth, G., 125, 132, 135 Honaker, R.W., 49, 82, 83, 85, 90 Hondalus, M.K., 91–93, 97 Honer zu Bentrup, K., 52, 54, 55 Hood, D., 137 Hopmans, E.C., 13, 14, 30, 32 Horn, R., 79 Hornsby, T., 90 Horton, E., 68 Horwich, A.L., 123 Howard, S.T., 59 Hu, Y., 124 Huang, Z., 13, 32 Huang, Z.Y., 33 Huet, G., 93 H€ ufner, E., 131 H€ ugler, M., 16, 17, 74, 75 Humphrey, T.J., 137 Hunter, G.J.E., 57 Husain, M., 68 Hussey, G., 47 Hutte, R., 86 Hutter, B., 78 Hwang, C., 62 Hwang, E.H., 77, 84 Ibrahim, Y.M., 129, 130 Ido, Y., 50 Igarashi, Y., 74 Ikeda, T., 74 Imlay, J.A., 53 Ingalls, A.E., 6, 11–13, 29, 32, 33 Ingmer, H., 123, 125, 127–131, 134–136 Inskeep, W.P., 33 Ioannidis, I., 89 Ioanoviciu, A., 83 Ishii, M., 74 Ishikawa, M., 72

AUTHOR INDEX Ito, K., 85, 90 Itoh, M., 52, 75, 76 Izzo, A., 82 Jackson, L.S., 130 Jacobs Jr., W.R., 52, 54, 55, 59, 68, 74–77, 80, 82, 91–94, 97 Jagels, K., 90 Jain, A., 59 Jain, M., 52, 59, 68, 94, 95 Jain, S.K., 84 Jakimowicz, P., 91 Jayaswal, R.K., 130 Jeney, V., 84, 86 Jenney Jr., F.E., 72, 73 Jennings, L.D., 123 Jeoung, J.H., 76 Jia, Z., 23, 24, 26, 28 Jin, W., 13 Johnson, C., 80 Johnson, T., 81 Johnston, S.A., 59 Jones, A.K., 72 Jones-Carson, J., 68 Jonuscheit, M., 3, 6, 7 Joshi, S.A., 122, 123, 128 Jovanovich, S.B., 15 Juretschko, S., 4 Jurgens, G., 3, 4, 6, 7 Jyothisri, K., 82 Kajfasz, J.K., 129, 130 Kalscheuer, R., 68 Kana, B.D., 59 Kaneko, F., 86 Kao, C.M., 62 Kappler, A., 70 Kapur, V., 82 Karakousis, P.C., 84 Karatzas, K.A.G., 131, 137 Karl, D.M., 29, 30, 32, 34 Karner, M.B., 29 Karr, E.A., 16, 17 Kass, I., 61, 75, 77, 83

153 Kaster, A.K., 94, 95 Katayama, Y., 72 Katsumata, R., 78 Katsura, K., 72 Kaufmann, S.H., 47, 48 Kaufmann, S.H.E., 59 Kaupenjohann, M., 23, 25, 28 Kavuru, M., 90 Kawakami, R.P., 91–93, 97 Keatings, V.M., 71, 89 Kelemen, G.H., 92 Keller, C., 60, 84, 89 Kelley, W.L., 134 Kelly, R.M., 73 Kendall, S., 84 Kenniston, J.A., 122, 123, 128 Keough, B.P., 6 Kerr, A.R., 129, 130 Kesavan, A.K., 84 Khan, S., 50 Kharitonov, S.A., 85, 90 Khodursky, A.B., 52, 66, 68 Kielland-Brandt, M.C., 64 Kilo, C., 50 Kilstrup, M., 129, 130 Kim, E., 77, 84 Kim, H.J., 91 Kim, J.A., 77, 84 Kim, J.H., 77 Kim, L., 125, 132 Kim, P., 91 Kim, S.W., 77, 84, 129, 130 Kim, S.Y., 77, 84 Kim, T.H., 91 Kim, Y., 91 Kim, Y.I., 127, 130 Kim, Y.M., 77, 84 King, G.M., 77, 84 Kinger, A.K., 81 Kinkel, H., 6 Kirstein, J., 128, 129, 131, 132, 138 Kishan, K.V., 91 Kleerebezem, M., 127 Klein, E., 60, 84, 86

154 Klein, M.R., 82 Kleman, G.L., 55 Klenk, H.-P., 6, 7, 9, 10, 15–18, 22 Kletzin, A., 6, 7, 9, 10, 15, 17, 22 Klichko, V.I., 50 Klinkenberg, L.G., 84 Klotz, M.G., 16, 17 Knap, A.H., 6, 7, 10, 15, 16, 22 Knosp, O., 67 Ko, M., 80 Ko, Y.F., 56 Kobayashi, Y., 125 Koch, C., 84 Kock, H., 123, 128–130 Kockelkorn, D., 19 Kohno, S., 54, 59, 68, 82 Kolattukudy, P.E., 68, 90, 94 Koledin, T., 79, 80 K€ onneke, M., 9–13, 16, 17, 33 Konstantinidis, K.T., 16, 29, 33 Koo, H., 129, 130 Koonin, E.V., 15, 16, 123 Koops, H.P., 4 Kosaka, K., 54 Kosmiadi, G.A., 59 Kotzerke, A., 23, 25, 28 Kovacevic, S., 80 Kowalchuk, G.A., 3, 4 Kramer, N., 131 Kramnik, I., 49, 54, 61, 68, 85, 86, 89 Kravitz, S., 34 Krebs, C., 96 Krebs, W., 73 Krogh, A., 90 Kroll, H.-P., 129 Kr€ uger, E., 123, 124, 128–132, 134, 136, 138 Kumar, A., 49, 54, 60, 61, 68, 82–86, 89 Kunst, F., 128–130 Kuo, H.P., 60, 83, 85 Kushmaro, A., 33 Kusters, I., 123, 128–130 Kuypers, M.M., 14, 30, 32 Kuypers, M.M.M., 6, 13, 33, 34

AUTHOR INDEX Kvist, T., 32 Kwon, H.-Y., 129, 130 Labischinski, H., 129 LaCourse, R., 60, 83 Ladel, C., 129 Lai, D., 13 Lalk, M., 135 Lalloo, U., 47 Lambert, P.H., 47 Lamichhane, G., 84 Lancaster Jr., J.R., 49, 60, 82–86 Lancaster, J.R., 91–93 Lang, D., 16, 17 Lanzen, A., 17, 18 Lanzen, J.L., 60 Lapa e Silva, J.R., 60, 83 Laskowski, D., 86 Laughlin, J., 68 Lavik, G., 13, 33, 34 Lawton, T.J., 16, 17 Leary, J.A., 52, 59, 68, 94, 95 Leavell, M.D., 52, 59, 68, 94, 95 Lebedeva, E., 11 Lebedeva, E.V., 11, 12 Lee, H.S., 91 Lee, K.H., 77, 84 Lee, N., 56 Lee, S., 127 Lee, S.M., 59, 84, 90 Legan, S.K., 50 Lehner, A., 132, 134 Lehtovirta, L.E., 23 Leichert, L.I.O., 134 Leigh, J.A., 95 Leija, A., 66, 67 Leininger, S., 6, 7, 9, 10, 14, 15, 22–25, 28 Leistikow, R.L., 75, 82, 83, 86, 87 Lemasters, J.J., 50 Lemos, J.A., 129, 130 Lemos, J.A.C., 128, 134 Lenaerts, A.J., 59 Lenz, O., 72 Leone, A.M., 60, 85

AUTHOR INDEX Leopold, J.A., 50 Levanon, S.S., 93 Levchenko, I., 122, 123, 127, 128, 130 Levitt, M.D., 72 Levy, S., 6, 7, 10, 15, 16, 22 Lew, D., 134 Li, C., 50, 97 Li, J., 13 Li, K., 34 Li, Q., 84 Li, R., 82, 83 Li, T., 13 Li, W.J., 33 Liao, R., 54, 59, 68, 82–84, 89 Liao, R.P., 82 Libby, S.J., 78 Licht, S., 123 Lie, T.J., 95 Liebeke, M., 135 Lieberman, R.L., 8 Lienhardt, C., 47 Likolammi, M., 4 Lilie, H., 129 Lin, H., 68 Lin, H.C., 60, 83, 85 Lin, P.L., 60, 84, 86 Linhares, C., 60, 83 Linnane, S.J., 71, 89 Liu, C.Y., 60, 83, 85 Liu, L., 50, 97, 130 Liu, W., 80 Liu, X., 34 Liu, Y., 91 Liu, Z., 80 Locht, C., 59 Lodish, H.F., 62 Lomas, M.W., 6, 7, 10, 15, 16, 22 López-Garc
ıa, P., 15, 16 Lopez-Nevot, M.A., 83 Loscalzo, J., 50 Losick, R., 128, 130 Loss, C., 124 Lothrop, W.C., 58, 60 Loux, V., 124, 125

155 Lowe, T., 16, 17 Ludwig, H., 128, 131, 138 Lun, D.S., 123 Luzardo, Y., 130 Ly, L.H., 84 Lyons, R., 59 Ma, K., 73 Ma, Y., 50, 97 Ma, Z., 47 MacGregor, B.J., 6 MacMicking, J.D., 60, 83 M€ ader, U., 135 Maglica, Z., 127 Maguin, E., 124, 125 Mai, D., 68, 87, 90–99 Maier, R.J., 72, 73 Maier, S.E., 73 Malhotra, V., 82, 84 Malinski, T., 60 Mallidis, C.G., 137 Malm, S., 60, 84, 89 Malzan, A., 60, 84, 89 Manabe, Y.C., 84 Mangenot, S., 124, 125 Manjunatha, U., 60, 84, 86 Mann, B.E., 61, 86 Manning, G., 16, 17 Manzanillo, P., 54, 61, 82, 85, 86 Manzei, S., 32 Markieton, T., 131 Marsh, T.L., 15 Martens-Habbena, W., 10, 16, 17 Marteus, H., 85 Martin, J., 83 Martinez, A.R., 129, 130 Martinez, G.J., 80 Masjedi, M.R., 47 Massana, R., 30 Masuda, S., 125 Masuo, S., 78 Matic, I., 137 Mat-Jan, F., 56 Matter, E., 57

156 Matthews, B.W., 134 Matthies, M., 23, 25, 28 Maurizi, M.R., 122, 123 Mavrodi, D.V., 70, 72 Mayuri, Bagchi, G., 82 McCallum, K., 4, 19 McCollister, B.D., 68 McCue, L.A., 49, 61, 76 McDonough, K.A., 49, 61, 76 McIlleron, H., 47 McKinlay, J.B., 77 McKinney, J.D., 47, 52, 54, 55, 94 McLean, J., 90 McLeod, C.W., 61, 86 McLoughlin, P., 71, 89 McNeil, M., 94 McNichol, A.P., 6 Mechtler, K., 132, 134 M
edard, M., 123 Medina, V.G., 63 Medlar, E.M., 83, 84 Meena, L.S., 49 Meijer, W.G., 78 Mengewein, A., 32 Mentel, M., 70 Merrill, M.H., 56, 57, 89 Meurer, G., 6, 15, 17 Meurer, G.b., 15 Meyer, J., 72 Meyer, K.C., 59, 71, 85, 89 Meyer, M., 33 Meyer, O., 77 Michaelis, L., 70 Michalik, S., 123, 128–130 Miczak, A., 52, 54, 55 Middelburg, J.J., 30 Middlebrook, G., 61, 75, 77, 83 Miethke, M., 128–130, 132 Miller, C.C., 81 Mills, G., 13 Milohanic, E., 130 Min, X., 50, 97 Mincer, T.J., 29, 30, 32, 33 Minch, K.J., 47, 49

AUTHOR INDEX Miranker, A.D., 123 Mitchell, T.J., 129, 130 Mizrahi, V., 47, 59 Mockett, R.J., 50 Moenne-Loccoz, P., 83 Mogk, A., 125, 127, 129, 132 Mohamed, N.M., 34 Mohan, V.P., 54, 59, 68, 82 Mohanty, D., 94 Moli ere, N., 128 Molinski, T.F., 10–12, 33 Moll, A., 47 Mollenkopf, H., 59 Monbouquette, H.G., 13 Moncada, S., 60, 85 Monk, C.E., 61, 86 Montonen, L., 4 Mora, J., 66, 67 Mora, Y., 66, 67 Morbidoni, H.R., 68 Moreira, D., 15, 16 Mori, T., 54 Morowitz, H.J., 74, 75 Morozkina, E.V., 87, 89 Morris, R.P., 91 Morton, R.A., 75, 83, 86, 87 Moser, D.P., 6 Mosier, A.C., 32 Mossman, M.R., 64, 65 Mostertz, J., 128, 135 Mota, M.M., 84, 86 Motterlini, R., 61, 86 Mougous, J.D., 52, 59, 68, 94, 95 Moule, S., 90 Moxon, R., 137 Moynihan, J.B., 71, 89 Msadek, T., 124, 125, 127–131, 136 Mudgett, J.S., 60, 83 Muller, E.G., 62 Mumford, R., 60, 83 Munch, J.C., 23, 25, 28 Munoz-Elias, E.J., 52, 54, 55 Munster, U., 4 Muratsubaki, H., 75

AUTHOR INDEX Murphy, L., 90 Murray, A.E., 30 Murray, J.F., 60 Murthy, P.S., 48, 54, 58 Muscariello, L., 127 Muttucumaru, D.G., 47 Myrold, D.D., 24 Nair, S., 128–131 Nakano, M.M., 128 Nakano, S., 128 Namsaraev, B., 11 Nandi, R., 72, 73 Narasimhulu, K.V., 91–93 Narayanan, P.R., 84 Nathan, C., 52, 60, 75, 76, 83 Nathan, C.F., 60, 83 Nealson, K., 6, 7, 10, 15, 16, 22, 34 Nealson, K.H., 6, 16 Neher, S.B., 122, 123, 128 Nelson, K.E., 6, 7, 10, 15, 16, 22 Nelson, W., 6, 7, 10, 15, 16, 22 Neubauer, H., 89 Neuwald, A.F., 123 Newman, D.K., 70–72 Newton, G.L., 79–81 Ng, W.-L., 129 Nguyen, K., 91 Nguyen, L., 91, 130 Nguyen, L.P., 15 Nguyen, T.T.H., 135 Nicholson, H., 134 Nicholson, S., 60, 83 Nicol, G., 17, 18, 26, 28 Nicol, G.W., 6, 11, 14, 23–26 Nicolas, P., 124, 125 Nielsen, J., 64 Nieminen, A.L., 50 Nishimaki, K., 72 Nissen, N., 91, 96 Nissen, T.L., 64 Norbeck, J., 63 North, R.J., 59, 60, 83 Novak, R., 130

157 Nowag, A., 91, 96 Nunn, A.J., 47 O’Brien, P.J., 50 O’Callaghan, M., 24, 26–28 O’Connor, C.M., 71, 89 O’Mullan, G.D., 30 Oakes, E.C., 128 Oakes, E.S.C., 122, 123, 128 Oakley, B.B., 29, 32 Ochsenreiter, T., 4, 6, 11, 15, 17, 23 Odelberg, S.J., 50 Oelgeschlager, E., 76, 77 Oenema, O., 4 Offre, P., 26, 28 Ogino, T., 56 Ogunniyi, A.D., 129, 130 Ogura, M., 128 Oh, J.I., 77, 84 Oh, N.N., 80 Ohlmeier, S., 123, 128, 129, 131 Ohmori, D., 74 Ohno, H., 54, 59, 68, 82 Ohsawa, I., 72 Ohta, S., 72 Olczak, A., 72, 73 Oliver, K., 90 Ollivier, B., 3 Olson, J., 72, 73 Olson, J.W., 72, 73 Oman, J., 60 Onstott, T.C., 6 Orosz, A., 50 Orr, W.C., 50 Orsi, R.H., 124 Osborne, J., 90 Ouverney, C.C., 6 Oztas, S., 124, 125 Page, W.J., 67 Paget, M.S., 78, 96 Paige, C., 49 Palva, A., 129, 130, 132, 136 Pamplona, A., 84, 86

158 Pan, Q., 128, 130 Pancost, R.D., 6 Pan
e-Farr
e, J., 124 Parente, E., 131 Parish, T., 47, 84 Park, H., 77, 84 Park, J.S., 91 Park, S.-H., 129, 130 Park, S.J., 77 Park, S.W., 77, 84 Park, T.H., 68, 87, 95, 99 Parkhill, J., 90 Parsons, R., 6, 7, 10, 15, 16, 22 Patel, B.K.C., 3 Patel, H., 62 Patel, M.P., 81 Patel, R.P., 49, 60, 82–86 Paton, J.C., 129, 130 Paulsen, H., 129 Paulsen, I., 6, 7, 10, 15, 16, 22 Pawinski, R., 47 Pearson, A., 6, 13, 29, 32 Pelicic, V., 59 Pellegrini, E., 128–130 Penaud, S., 124, 125 Pereira, I.A.C., 73 Perham, R.N., 81 Perkins, M.D., 47 Pernthaler, A., 6 Pernthaler, J., 6 Perrella, M.A., 60, 85 Perrone, G., 62 Persson, M.G., 85 Peshock, R.M., 50 Peters, G., 130 Petersen, A., 70 Peterson, J., 6, 7, 10, 15, 16, 22 Pethe, K., 68, 76, 83, 86, 87, 89, 95, 99 Petzold, C.J., 52, 59, 68, 94, 95 Pfannkoch, C., 6, 7, 10, 15, 16, 22 Pfenninger-Li, X.D., 73 Pickart, C.M., 123 Pierre, F., 127, 128 Pieters, J., 91

AUTHOR INDEX Pillay, M., 47 Pinel, N., 16, 17 Pitcher, A., 13, 14 Platt, T., 34 Plum, G., 91, 96 Pommerening-R€ oser, A., 4 Poole, R.K., 61, 86 Popp, B.N., 29, 30 Portugal, S., 84, 86 Pouyssegur, J., 60 Pratt, C.W., 49, 55 Preston, C., 16, 29, 30, 32, 33 Preston, C.M., 10–12, 15, 16, 30, 33 Price-Whelan, A., 70–72 Proctor, R.A., 130, 134 Pronk, J.T., 63, 64 Prosser, J., 3 Prosser, J.I., 14, 23–28 Prudhomme, M., 127–130 Purkayastha, A., 49, 61, 76 Purkhold, U., 4 Putnam, N., 16, 29, 33 Puzewicz, J., 132 Pyo, S.-N., 129, 130 Qazi, S., 127, 130 Qazi, S.N.A., 128, 129 Qi, J., 14, 23–25 Qi, R., 82, 83 Qian, B., 50, 97 Qoronfleh, M.W., 130 Quail, M.A., 90 Quaiser, A., 4, 6, 11, 15, 17, 23 Quivey, R.G., 129, 130 Rachman, H., 59 Radax, R., 13, 33, 34 Raddatz, G., 6, 15, 17 Raddatz, S., 129 Radha Kishan, K.V., 91 Radyuk, S.N., 50 Raengpradub, S., 124 Raghunand, T.R., 91 Ragsdale, S.W., 76

AUTHOR INDEX Rajakumar, K., 59, 84, 90 Rajandream, M.A., 90 Rajasekaran, N.S., 50 Rajni, 49 Ramakrishnan, L., 47, 48 Ramakrishnan, T., 48, 54, 58 Ramalingam, B., 59 Ramanathan, V.D., 84 Rand, J.D., 63 Rand, L., 76, 86, 87 Randell, S., 59, 71, 85, 89 Rao, S.P., 76, 83, 86, 87, 89 Rapoport, G., 125, 128–131 Rapp, H.T., 13, 33, 34 Rasmussen, K.N., 61, 83, 84 Ratjen, F., 89 Ratledge, C., 47, 51, 52 Rattei, T., 20, 22, 23 Rawat, M., 79–81 Ray, S.M., 60, 84, 86 Rebrin, I., 50 Redding, K.E., 91–93 Reed, M.B., 83 Reeves, R.E., 58, 60 Reid, B.G., 123 Reigstad, L.J., 13, 33 Reinthaler, T., 6 Remington, K., 6, 7, 10, 15, 16, 22, 34 Ren, B., 53 Renfrow, M.B., 68, 87, 90, 93–99 Reysenbach, A.L., 14, 32 Rhee, D.-K., 129, 130 Ricciardi, A., 131 Rich, A.R., 61, 83, 84 Richardson, A.R., 78 Richardson, D.J., 68, 69, 87, 89 Richardson, P.M., 16, 29, 33 Richter, A., 11–13, 33 Rietveld, P., 81 Rieu, A., 127 Rijpstra, W.I., 14, 32 Riley, R.L., 61, 83 Ripio, M.T., 129, 130 Ritz, D., 62

159 Rivera-Ramos, I., 129, 130 Ro, Y.T., 77, 84 Robert, C., 124, 125 Roberts, D.M., 82 Roberts, G., 47 Roberts, G.P., 4 Roberts, K., 33 Roberts, K.J., 29, 32 Robertson, G.T., 129 Robinson, N., 91, 96 Robson, R., 72 Rocha, E.P.C., 137 Rodrigues, C.D., 84, 86 Rodr
ıguez-Valera, F., 15, 16 Rogers, J., 90 Rogers, Y.H., 6, 7, 10, 15, 16, 22, 34 Rohwer, F., 33 Rom, W., 60, 83 Romanek, C.S., 13, 32, 33 Rondon, E., 59 Rose, M., 129, 130 Rosenzweig, A.C., 8, 16, 17 Ross, C., 33 Rossano, R., 131 Rother, M., 76, 77 Rouanet, C., 59 Rouquette, C., 129, 130 Rubin, B.K., 89 Rubin, E.J., 47, 48 Rudolph, F.B., 68 Rusch, D., 6, 7, 10, 15, 16, 22 Rusch, D.B., 34 Russell, D.A., 87, 89 Russell, D.G., 52, 54, 55 Russell, G.C., 53 Rustad, T.R., 47, 49, 83, 84 Rutter, S., 90 Ryan, G.J., 59 Rybniker, J., 91, 96 Ryter, S.W., 60, 85 Saano, A., 4 Sacchettini, J.C., 47, 52, 54, 55, 80 Sachdeva, S., 82

160 Sahl, H.-G., 129 Saini, D.K., 82, 84 Saito, K., 34 Sambrook, J., 122 San, K.Y., 66, 68, 87, 93 Sanchez, A.M., 68 Sandaa, R.A., 4 Sanguinetti, G., 61, 86 Sangurdekar, D.P., 52, 66, 68 Santoro, A.E., 29, 32 Santos, G.M., 6, 29 Sarath, G., 78 Sareen, D., 79, 80 Sariyar, B., 68 Sathyendranath, S., 34 Sauer, R.T., 122, 123, 127, 128, 130 Saunders, A.M., 32 Saves, I., 93 Savijoki, K., 128, 129 Sawers, G., 68, 69 Sayavedra-Soto, L.A., 16, 17 Schafer, F.Q., 78, 79 Scharf, C., 124, 127, 134, 135 Schauss, K., 23, 25, 28 Schedin, U., 85 Scheffers, W.A., 63, 64 Schelle, M.W., 52, 59, 68, 94, 95 Schicho, R.N., 73 Schittone, S., 82 Schl€ appy, M.-L., 13, 33, 34 Schlegel, H.G., 77 Schleper, C., 3, 4, 6, 7, 9–11, 13–18, 20, 22–25, 28, 29, 33, 34 Schlieker, C., 127 Schloter, M., 14, 23–25, 28 Schlothauer, T., 129 Schluger, N.W., 60 Schmid, F.X., 125, 132 Schmid, M.C., 4 Schmid, R., 124, 127 Schmidt, A., 132, 134 Schmidt, T.M., 4, 6 Schmitt, S., 13

AUTHOR INDEX Schnappinger, D., 49, 52, 54, 59, 61, 68, 82, 83, 87, 89, 91, 95, 99, 100 Scholz, C., 125, 132 Schoolnik, G.K., 49, 54, 59, 61, 68, 81–83, 89 Schouten, S., 6, 13–15, 30, 32 Schramm, A., 32 Schroeder, W., 129 Schuchhardt, J., 59 Schumann, W., 124, 125, 128, 132 Schuster, S.C., 6, 7, 9, 10, 14, 15, 17, 22–25 Schut, G.J., 73 Schwab, U., 59, 71, 85, 89, 124 Schwark, L., 13, 14, 23–25, 33 Schwartzberg, P., 130 Scrutton, N.S., 81 Sears, H.J., 68, 69 Seedorf, H., 94, 95 Seeger, K., 90 Segal, W., 48, 51, 53, 54, 58–60, 99 Seifritz, C., 89 Seitz, H., 15, 16 Selezi, D., 4, 11, 23 Sengupta, S., 72, 73 Senior, P.J., 66, 67 Senner, C., 59, 84, 90 Sensen, C.W., 15 Seravalli, J., 76 Shah, S.K., 60, 83 Shah, S.R., 6, 29 Shakila, H., 84 Shalel-Levanon, S., 93 Sharma, D., 84 Sharma, S., 23, 25, 28 Sheehan, H.L., 58 Shen, G., 96 Shen, J., 23, 24, 26–28 Shen, J.P., 26 Sherman, D.R., 47–49, 54, 59, 61, 68, 82–84, 89 Sherratt, A.L., 59, 84, 90 Sherrid, A.M., 47, 49 Shi, L., 59

AUTHOR INDEX Shiloh, M.U., 54, 61, 82, 83, 85, 86 Shimizu, M., 78 Shin, M., 16, 17 Shizuya, H., 15 Shock, E.L., 33 Siboni, N., 33 Siddiqui, S.M., 122, 123, 128 Sieber, S.A., 129 Siegers, K., 122 Sievert, S.M., 16, 17, 74, 75 Silva, N.A., 129, 130 Silver, R.F., 84 Simon, H.M., 4 Singh, A., 68, 87, 90–99 Singh, K.K., 82 Singh, S.K., 123 Singh, V.K., 130 Sinha, B., 130 Sinskey, A.J., 62 Sirakova, T.D., 68, 90, 94 Sirsi, M., 54, 58 Sivan, A., 33 Skelton, J., 90 Small, P.M., 83 Smalla, K., 23, 25, 28 Smith, D.A., 84 Smith, H., 34 Smith, H.O., 6, 7, 10, 15, 16, 22 Smith, I., 59 Smith, R.J., 59, 84, 90 Smittenberg, R.H., 13, 32 Snoep, J.L., 64, 65, 68, 78, 87 Snyder, S.A., 60 Soares, M.P., 84, 86 Sohal, R.S., 50 Sohaskey, C.D., 59, 89 Somerville, G.A., 130 Song, T., 77, 84 Song, Z.Q., 33 Soni, V., 91 Sørensen, K., 127, 130 Soteropoulos, P., 59 Sousa, E.H., 60, 83, 84 Souza, V., 34

161 Spang, A., 20, 22, 23 Spano, G., 127 Spellman, P.T., 62 Spencer, J.S., 59 Spencer, M.E., 53 Spieck, E., 11, 12, 20, 22, 23 Spiess, S., 132, 134 Spiro, S., 87, 89 Spouge, J.L., 123 Springstead, J.R., 13 Squares, R., 90 Squares, S., 90 Sridharan, V., 94 Srinivasan, V., 74, 75 Srivastava, B.S., 59 Srivastava, R., 59 Srivastava, V., 59 Stabler, R.A., 47 Stahl, D.A., 6, 9–13, 16, 17, 20, 22, 23, 33 Standfest, S., 13 Stan-Lotter, H., 33 Staubli, A., 51 Steenkamp, D.J., 81 Steffek, M., 79 Steger, D., 13, 33, 34 Stein, J.L., 15, 16 Stein, L., 34 Steinbuchel, A., 67, 68, 90, 94 Stenzel, U., 33 Stephen, J.R., 3, 4 Steuber, J., 73 Stevenson, T.J., 50 Stewart, A., 82 Stewart, C., 34 Steyn, A.J., 49, 54, 60, 61, 68, 82–87, 89–99 Steyn, A.J.C., 49, 82, 83, 85, 90–93 Stoecker, K., 11, 12 Stoker, N.G., 84 Stolarczyk, E., 60 Storz, G., 62 Stoveken, T., 67 Strausberg, R.L., 34 Streit, W., 20, 22, 23

162 Strohl, W.R., 55 Strong, M., 30, 59 Stuehr, D.J., 86 Sturm, A.W., 47 Suematsu, M., 52, 75, 76 Sugahara, J., 16, 29, 33 Suhail Alam, M., 91 Sulston, J.E., 90 Suter, E., 57 Sutton, G., 34 Suzuki, M.T., 15 Swanson, I., 95 Swanson, R.V., 15, 16 Swenson, D., 52, 54, 55 Switzer, R.L., 123, 128–130 Ta, P., 79, 80 Tabarsi, P., 47 Taddei, F., 137 Tahlan, K., 68, 87, 95, 99 Takahashi, K., 72 Takai, K., 6 Takaya, N., 78 Tal, Y., 34 Talaat, A.M., 59 Tamayo-Castillo, G., 34 Tan, G., 53 Tanaka, K., 84 Tanaka, Y., 54 Tarran, R., 59, 71, 85, 89 Tascon, R.I., 129, 130 Tassou, C.C., 137 Tavano, C., 59 Taylor, C.D., 74, 75 Taylor, C.J., 87, 89 Taylor, K., 60, 84, 86, 90 Taylor, L.T., 15, 16, 29, 30, 32 Taylor, M.W., 13, 33, 34 Taylor, R.P., 50 Teague, W.G., 85, 90 Teal, T.K., 70, 71 Teira, E., 6, 30 Teixeira de Mattos, M.J., 64, 65, 68, 87 Tekaia, F., 90

AUTHOR INDEX Tepper, R., 54 Tessarz, P., 127 Thauer, R.K., 94, 95 Thevelein, J.M., 63 Thiele-Bruhn, S., 23, 25, 28 Thomashow, L.S., 70, 72 Thomassen, M.J., 90 Thompson, E.T., 129 Thompson, M.W., 123 Thomson, A.J., 91 Thorpe, J., 34 Thunnissen, F.B., 90 Tian, F., 13 Tian, J., 52, 75, 76 Tickoo, R., 94 Tilton, R.G., 50 Timmers, P., 30 Tischendorf, G., 129 Tischer, K., 124, 127 Tischler, P., 20, 22, 23 Toledo, J.C., 49, 60, 82–86 Tomboulian, P., 60 Tomkiewicz, R.P., 89 T€ ornberg, D.C., 85 Torre, O., 85, 90 Torsvik, V., 4 Touchon, M., 137 Tourna, M., 26 Tran, B., 34 Treangen, T.J., 137 Tremblay, G.A., 84 Treusch, A.H., 6, 7, 9, 10, 15, 17, 22 Triccas, J.A., 59 Trivedi, O.A., 94 Trombley, J., 91–93 Trotter, E.W., 62 Tsai, F.T.F., 127 Tsai, M.C., 84 Tsukahara, K., 128 Tuckerman, J.R., 60, 83, 84 Tufariello, J.A., 84 Tuomanen, E., 130 Turgay, K., 128–132, 134, 138 Tyagi, J.S., 81, 82, 84

AUTHOR INDEX Ulrich, M., 59, 71, 85, 89 Ulrichs, T., 48, 59 Unson, M.D., 80 Upton, A.M., 52, 55, 94 Urakawa, H., 10, 16, 17 Urich, T., 13, 14, 23–25, 33 Utaida, S., 130 Utterback, T., 34 Vadali, R.V., 68 Valadi, A., 63 Valadi, H., 63 Valdramidis, V.P., 137 Valente, F.M.A., 73 Van Aken, H., 6 Van Beusichem, M.L., 4 van Bleijswijk, J., 14, 30, 32 Van Damme, O., 79 van de Guchte, M., 124, 125 Van der Linden, E., 72 van der Meer, M.T., 14, 32 Van Der Merwe, M.J., 78 Van Der Weijden, C.C., 78 van Dijken, J.P., 63, 64 van Duin, A.C., 13 van Gumpel, E., 91, 96 Van Keulen, G., 78 van Maris, A.J.A., 63 Varcamonti, M., 131 Varma, K.G., 54 Varmanen, P., 125, 128–132, 134, 136 Vats, A., 94 Vaudaux, P., 134 V
azquez-Boland, J.A., 129, 130 Vazquez-Torres, A., 68 Velayati, A.A., 47 Veldhuis, M.J.W., 30 Velez, L., 83 Velthof, G.L., 4 Vemuri, G.N., 52, 66, 68 Venceslau, S.S., 73 Venter, J.C., 6, 7, 10, 15, 16, 22, 34 Venter, J.E., 34

163 Veth, C., 6 Via, L.E., 60, 84, 86 Vignais, P.M., 72 Vilchez, J.R., 83 Vilcheze, C., 74–77, 80 Villacorta, R., 15 Villadsen, J., 64 Vill
en, J., 128 Visconti, K., 91 Visconti, K.C., 49, 54, 59, 61, 68, 81–83, 89 Voet, D., 49, 55 Voet, J.G., 49, 55 Vogensen, F.K., 125, 129–132, 134–136 Vogt, R.N., 81 V€ olker, U., 124, 130, 132, 134 Voskuil, M., 54, 59, 68, 82, 89 Voskuil, M.I., 49, 54, 59, 61, 68, 75, 81–83, 85–87, 89, 90, 93–99 Waddell, S.J., 59, 84, 90 Wagner, K., 60, 75, 83, 84, 86, 87, 89 Wagner, M., 4, 11–13, 20, 22, 23, 33, 34 Wah, D.A., 122, 123, 128 Wakeham, S.G., 14, 30, 32 Waldminghaus, T., 128 Walker, C.B., 9–13, 16, 17, 33 Waltermann, M., 67 Walunas, T., 124, 125 Wang, C.H., 60, 83, 85 Wang, H., 130 Wang, J., 123 Wang, T., 95, 96 Wang, X., 47 Ward, B.B., 30 Ward, D.E., 78 Ward, D.M., 14, 32 Ward, S.K., 59 Warner, D.F., 47 Watanabe, M., 72 Watanabe, Y., 16, 29, 33 Waterbury, J.B., 9, 10, 12 Wawrzynow, A., 132 Wayne, L.G., 54, 60, 83, 89

164 Weber-Ban, E.U., 123, 127 Wegley, L., 33 Weibezahn, J., 127 Weidler, G.W., 33 Weidmann, S., 127 Weidner, J.R., 60, 83 Weigand, W.A., 56 Weiss, T., 59, 71, 85, 89 Weissenbach, J., 124, 125 Weitzberg, E., 85 Wells-Bennik, M.H.J., 137 Weraarchakul, W., 130 West, J.B., 61 Westerhoff, H.V., 78 Westermann, P., 32 Whalan, S., 13, 33, 34 Wheeler, P.R., 47, 51, 52 White, O., 6, 7, 10, 15, 16, 22 Whitehead, S., 90 Whiteley, M., 70 Whitman, W.B., 3 Whitwell, F., 58 Wickner, S., 122, 123 Wiebe, W.J., 3 Wiedmann, M., 124 Wiegel, J., 13, 32, 33 Wiegert, T., 128 Wiklund, N.P., 85 Wilke, B.M., 23, 25, 28 Wilkinson, B.J., 130 Wilkinson, R.J., 49, 52, 59, 61, 100 Williams Jr., C.H., 81 Williams, K.J., 68, 87, 95, 99 Williamson, J.R., 50 Williamson, S., 34 Willms, K., 66, 67 Wimpenny, J.W., 64, 65 Wimpenny, J.W.T., 70, 71 Winefield, C., 24, 26–28 Winefield, C.S., 26 Winkler, M.E., 129 Winstanley, C., 70, 71 Wipat, A., 127 Wirsen, C.O., 74, 75

AUTHOR INDEX Wisedchaisri, G., 82 Witt, E., 123, 128, 129, 131, 138 Woebken, D., 13, 33, 34 Woese, C.R., 13 Wolfe, A.J., 56, 64, 65 Wolin, M.J., 49, 61, 76 Wong, P.M., 95 Wong, S.L., 125 Woodruff, R.V., 127, 130 Worlitzsch, D., 59, 71, 85, 89 Wouters, J.A., 131, 137 Wrage, N., 4 Wright, D.E., 57 Wu, C.W., 59 Wu, D., 6, 7, 10, 15, 16, 22, 34 Wu, K., 30 Wu, K.-F., 130 Wu, K.Y., 10–12, 15, 16, 33 Wu, L., 34 Wu, Y., 96 Wuchter, C., 6, 14, 15, 30 Xia, D., 95 Xie, Q.W., 60, 83 Xu, J., 84 Xu, M., 23 Xu, S.-X., 130 Xu, W.-C., 130 Xu, X., 68, 87, 95, 99 Yamagata, K., 72 Yamamoto, M., 74 Yan, B.S., 49, 54, 61, 68, 85, 86, 89 Yan, L.J., 50 Yan, T., 34 Yang, J., 53 Yang, Y.T., 68 Yankaskas, J.R., 59, 71, 85, 89 Yao, Y., 50, 97 Ye, Q., 33 Yee, C.H., 61, 86 Yi-Lian, L., 13 Yin, Y.-B., 130 Yooseph, S., 34

AUTHOR INDEX Young, D., 49, 52, 59, 61, 100 Young, D.B., 47 Young, J.C., 122 Yu, C.A., 95 Yu, C.T., 60, 83, 85 Yu, J.Y., 77 Yu, L., 95 Yuan, G., 125 Yukl, E.T., 83 Yun, C.S., 68, 87, 95, 99 Yura, T., 124 Zagorec, M., 131 Zahn, R., 127 Zbell, A.L., 73 Zeiher, A.M., 50 Zeller, K., 47 Zellmeier, S., 128 Zentgraf, H., 127 Zervos, A., 137 Zhang, C.L., 13, 32, 33

165 Zhang, L., 23 Zhang, Q., 130 Zhang, X., 13, 50, 97 Zhang, X.-M., 130 Zhang, X.Q., 50 Zhao, W.D., 33 Zheng, G., 128 Zheng, M., 62 Zheng, R., 81 Zheng, Y., 23 Zhou, J., 34 Zhu, G., 54, 59, 68, 82, 84 Zhu, Y., 23 Ziazarifi, A.H., 47 Ziebandt, A.K., 124, 127 Zolkiewski, M., 123 Zotta, T., 131 Zuber, P., 128 Zuber, U., 125 Z€ uhlke, D., 128, 131, 132, 138 Zvyagilskaya, R.A., 87, 89 Zylicz, M., 132

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Subject Index Note: The page numbers taken from figures and tables are given in italics.

A Acetyl-CoA synthase (ACS), 76 Acetyl-coenzyme A (acetyl-CoA), 51 N-Acetylglucosaminylinositol, 79, 80 Aerobic ammonia oxidation, 4 Aerobic archaea, 3 Aerobic carboxydotrophic bacteria, 76 Aerobic microorganisms, 4 Allylthiourea, 11 Ammonia, 3, 4 competition for, 30 conversion to nitrite, 33 oxidation, 3, 4, 13, 17, 25, 30 source of, 25 Ammonia monooxygenase (AMO), 6 pMMO-related protein, 8 PMO, phylogenetic relationship, 8 Ammonia oxidation, 3, 4 Ammonia oxidisers as distinct phylum within archaea, 4, 19–22 phylogenetic tree, 23 processing genes, distribution, 20–22 uncultivated, metagenomic studies, 15–16 amo-related genes, 15 confirming close relationship of, 16 microheterogeneity, 16 Ammonia-oxidising archaea (AOA), 4, 6–13 ammonia oxidation kinetics, 10 amo/pmo genes, 8

associated with marine invertebrates, 33–34 sponge-associated communities, 34 Cenarchaeum/Axinella association, 11 contrasting response to nitrogen deposition, 27 cultivated, 12 diversity, distribution and activity, 22–23 phylogenetic tree, 24 enrichments, 11 genomes and metagenomes, 15 BAC-derived fosmid vectors, 15 WGS approach, 15 in geothermal environments, 32–33 isolation, 9 in marine environment, 28–29 activity, 30–32 fluxes in inorganic nitrogen concentrations, 31 membrane lipids of, 13–15 GDGT, 13, 14 phospholipids, 13 recovery of crenarchaeol, 13 TEX86 index, 14 oligotrophic lifestyle, 10 open reading frames (ORFs) coding for, 6 phylogenetic relationship, of AMO and PMO, 8 predicted ORFs, on 43 kb soil fosmid 54d9, 7 in sediments, 32

168 in soil environment, 23–24 activity, 25–28 amoA gene abundance in, 25 growth of acetylene-sensitive ammonia-oxidising archaea in, 28 16S rRNA-based PCR surveys, 11 Ammonia-oxidising bacteria (AOB), 4, 6 amoA gene abundance in soils, 25 autotrophic, fixation of carbon, 19 contrasting response to nitrogen deposition, 27 environmental factors, 22 growth characteristics, 25 growth dynamics, 26 nitrification activity, 28 pathways of nitrogen, oxygen and electron flow, 18 ratio of AOA to AOB amoA genes, 23, 25 amoA genes, 9, 14, 29 Anti-mycobacterial drug, 47 Anti-sigma factor, SpoIIAB, 128 AOA. See Ammonia-oxidising archaea (AOA) AOB. See Ammonia-oxidising bacteria (AOB) Archaea carbon metabolism, 6 in moderate aerobic habitats, 4 phylogenetic relationship, 5 role in global cycles, 3 16S rRNA gene-defined lineages, 5 Arthrobacter oxydans, 78 Aspergillus nidulans, 78 ATPase, 123, 129 ATP citrate lyase, 74 ATP-dependent proteases, 123 ATP hydrolysis, 123 ATP synthesis, 69, 89 Auto-phosphorylation, 138 Axinella mexicana, 11, 13, 16, 33 Azorhizobium, 67

SUBJECT INDEX Azotobacter beijerinckii, 66 Azotobacter vinelandii, 64, 67

B BAC-derived fosmid vector, 15 Bacillaceae, 127 Bacillales, 124, 131 Bacille Calmette-Gu erin (BCG) vaccine, 47 survival, under hypoxic conditions, 61 Bacillus stearothermophilus, 132 Bacillus subtilis, 124, 125, 128 Bacterial artificial chromosomes (BACs), 15 Bacterial regulatory circuits, 138 Blood–brain barrier, 86 Bone marrow-derived macrophages, 85 Bradyrhizobium, 67

C Calvin–Bassham–Benson cycle, 19 Calvin cycle, 76, 77 Candida boidinii, 66 Carbon cycle, 3 Carbon dioxide (CO2), 49 Carbon monoxide (CO), 49 binding of CO to DosS and DosT, 84 Dos regulon and, 85–86 as energy source, 77 in Mtb persistence, 60 oxidation, 77 utilising microorganisms use enzyme CODH to, 76 Carbon monoxide (CO) dehydrogenase (CODH), 76–77 Carbon oxidation state (COS), 54–56 Carboxydotrophic microbes, 76 Cellular chaperone machinery, 122 Cenarchaeum symbiosum, 10, 11 ammonia oxidation by, 11

SUBJECT INDEX from concatenated dataset of ribosomal proteins, 20 fluorescent in situ hybridisation, 10 genes for urease, 17 genome of, 15, 16 G + C content, 16 pathway for carbon fixation, 19 Chemolithoautotrophic ammonia oxidisers, 9 Clostridium welchii, 70 clpC, clpP and clpB operons, 124 clpL in staphylococci, 127 Clp machinery, 123 ClpP complex, 123 Clp protease, 123 Clp-specific degradation, 128 clpX expression, regulation of, 127 13 C-metabolic flux analysis, 99 CoA-dependent KDH, 75 Corynebacterium glutamicum, 91 Crabtree effect, 65 Crenarchaeol, 13, 14, 31, 32 Crenarchaeota, 4, 6, 20 hyperthermophilic, 13 CtsR degradation McsB kinase activity, 138 role of McsB as adaptor, 138 YwlE, to counteract, 138 ctsR gene, 124, 125, 127, 137 CtsR inactivation, 124–125, 136 during oxidative stress, 135 CtsR protein, 134 CtsR-regulated genes, 124–127 cellular functions of, 127–131 CtsR-regulated proteins, distribution, 126 CtsR regulon, physiological function, 127 CtsR repressor, 124, 131 mechanisms for inactivation of, 131 heat inactivation of CtsR, 131–134 during other stress conditions, 136–137

169 during oxidative stress, 134–136 Cysteine, 79 Cystic fibrosis (CF), 59 Cytochromes, 17

D Denitrification, 4, 17, 19, 34 Dihydroxyacetone phosphate (DHAP), 63 Dissolved O2 tension (DOT), 65 Dithiothreitol (DTT), 62 DNA damage-inducible regulator, 129 Dos two-component system, 81 Drosophila melanogaster, 50

E Ecosystems, 3 Energy-dependent protein degradation, 123 Enterococcus faecalis, 78 Environmental stress response (ESR), 62 Escherichia coli, 15, 56, 64–66, 70, 124 acetate excretion, 56 fermentation, 64–66 (see also Reductive sinks) intracellular redox state, 65 nark narU mutant, 89 TCA cycle, 52 Estuaries, 4 Ethionamide (ETH), 80 Euryarchaeota, 4, 20 Extensively drug resistant (XDR), 47 Extremophiles, 3

F Fatty acids (FAs), 49 Ferredoxins, 74 Fe–S cluster proteins, 96

170 Formate dehydrogenase (FDH) reaction, 64 Fumarate reductase (FRD), 53, 75

G Genetic mutation, 137 Genomic techniques, advancement, 15 Gibbs free energy, 89 Glucuronic acid, 50 Glucose-6-phosphate dehydrogenase (G6PD), 50 Glutathione disulfide–glutathione couple (GSSG/2GSH), 78 Glycerol dialkyl glycerol tetraethers (GDGTs), 13, 14 Glycerol formation, 63 Glycerol-phosphate dehydrogenase (GPD2) mutant, 63 G3PD isoform, 63 Greenhouse gas, 4 Groundwater pollution, 4 GSH/GSSG ratio, 50

H Heat shock proteins (Hsp), 50, 124, 127, 134 Hsp18, 127 Hsp27, 50 Hsp 100/Clp, 123, 127 Helicobacter pylori, 73 Helix-turn-helix (HTH), 90, 131 Heme oxygenase-1 (HO-1), 60 High-hydrostatic pressure (HHP)tolerant mutants, 136 HrcA-dependent genes, 125 HrcA repressor, 125 Hsp. See Heat shock proteins (Hsp) Hydrogenases (H2ases), 54, 72–74 Hydroxylamine oxidoreductase (HAO), 9, 17

SUBJECT INDEX Hyperthermophilic organisms, 4 Hypoxia, 50, 76, 84

I Inducible nitric oxide synthase (iNOS), 60, 83 Isoniazid (INH), 80

K a-Ketoglutarate (KG), 53 a-Ketoglutarate decarboxylase (KGD), 75 a-Ketoglutarate:ferredoxin oxidoreductase (KOR), 74 Klebsiella aerogenes, 70 Klebsiella pneumoniae, 73

L Lactobacillaceae, 127 Lactobacillales, 125, 138 Lactobacillus acidophilus, 124 Lactobacillus plantarum, 127 Lactobacillus sakeii, 131 Listeriaceae, 127 Listeria monocytogenes, 124, 130, 137 Lactate, 77, 78 Low-molecular-weight (LMW), 79 Low-molecular-weight protein tyrosine phosphatase (LMWPTP), 132

M Macrophages, 59, 60, 85, 95, 99 Marine archaea carbon metabolism, 6 complete genome sequences, predictions from, 16–19 genomic fragments from, 6 tractable system for study of, 11 McsB adaptor, 138

SUBJECT INDEX McsB kinase, 136, 138 Metallosphaera sedula, 19 Methanobacterium mazei Go1, 70 Methanogenic archaea, 3 5-N-Methyl-1-hydroxyphenazine, 70 Methyl-malonyl CoA (MMCoA), 94 Methylococcus capsulatus, 8 Microarray analysis, 62, 73, 81 Microautoradiography, 6 Molecular chaperones dnaK and groE, 125 Mono-cistronic gene, 127 MSH acetyltransferase, 79 MSH-dependent enzymes, 81 MSH disulfide reductase, 81 MSH mutants, 80, 98 Msm mutant, 74, 80 Mtb. See Mycobacterium tuberculosis (Mtb) Mtb dos dormancy regulon, 81 biological role, and function, 81–84 Dos regulon and CO, 85–86 and reductive stress, 86–87 nitrate reductase, 87–90 TAG production, 90 and virulence, 84–85 Mtb DwhiB3 mutant, 92, 93 Mtb tgs1, expression of, 68 Mtb virulence lipid, 59, 94 Mtb WhiB7 depicting, sensing and dissipating reductive stress to, 96 as intracellular redox sensor, 90 Mtb whiB genes, 91 Multidrug resistant (MDR), 47 Mycobacteriophage TM4, 91 Mycobacterium bovis, 76 Mycobacterium smegmatis, 73 Mycobacterium tuberculosis (Mtb), 45, 47 culture in vitro, 48

171 Dos dormancy regulon, and role in, 48 DwhiB3 deletion strain, 92 environmental factors, influencing metabolism, 52 balancing act in vitro and in vivo, 58–60 carbon oxidation state (COS), 54–56 gaseous environment, of lung, 60–61 metabolites, excretion of, 56–58 redox balance, 56–58 TCA cycle, 52–54 historical knowledge, 48, 51–52 physiological characteristics, 51–61 W-Beijing lineage, 82 WhiB3 and virulence, 91–93 Mycolic acid, 93 Mycothiol (MSH), 79 biosynthetic pathway, 80 Mycothiol S-conjugate amidase (Mca), 81

N NADH/NAD+ ratio, 50, 66, 87 NAD+/NADH-independent enzymes, 74 NAD+/NADH ratio, 67, 71 NADPH/NADP+ system, 49 Nitrate reductase (Nap), 69, 87–90 Nitric oxide (NO), 49, 92 Nitrification, 3, 4 aerobic ammonia oxidation during, 9 Nitrifier-denitrification mechanisms, 4 Nitrite, 3 Nitrite-oxidising bacteria, 4, 9 Nitrogen cycle, 3 Nitrosocaldus yellowstonii, 11 Nitrosopumilus maritimus, 9, 10, 17, 19, 20 strain SCM1, 10

172 Nitrososphaera gargensis, 11 belongs to lineage of AOA distinct from, 11 contain crenarchaeol, 13 GDGTs of, 14 genomic information, 20 Nitrosothiol reductase, 81 Nitroxyl (HNO), 17 NO dehydrogenase (NODH), 77

O Oenococcus oeni, 125 Open reading frames (ORFs), 6 b-Oxidation, 67, 76, 86, 88, 94 Oxidation–reduction reactions, 97 metabolic homeostasis, 45 Oxidative phosphorylation, 47 Oxidative stress, 45, 91, 122, 135 importance, 45 role, 49 2-Oxoglutarate dehydrogenase complex (ODHC), 53 2-Oxoglutarate:ferredoxin oxidoreductase (OGFO), 74–75 Oxygen (O2), 49 Oxygen radicals, 53

P Paleothermometry, 14 Paracoccus pantotrophus, 69 Particulate methane monooxygenase (pMMO), 8 PHB mutant, 67 Phenazines, 69–72 Poly b-hydroxybutyrate (PHB), 66, 67 Polyhydroxyalkonate (PHA) biosynthesis, 66 central regulator, 67 support, direct evidence, 67 Polyketide, 66, 68, 87, 93 Polymer deposition

SUBJECT INDEX poly-b-hydroxybutyrate (PHB), 66–68 polyhydroxyalkonate (PHA), 66–68 triacylglycerol (TAG), 51, 59, 66–68, 84, 90, 93, 94, 96 Polymorphism, in iNOS, 83 Proteases, 123 26S proteasome system, 123 Protein degradation in eukaryotes, 123 Protein homeostasis, 122 Protein turnover, 123 Proteobacteria, 4 Proton motive force, 87 Pseudomonas aeruginosa, 70, 71 Pyocyanin, 70 Pyrococcus furiosus, 73 Pyruvate catabolism, 64, 65

R Reactive oxygen species (ROS), 50, 72 Rec-independent mutations, 137 Redox couples, 49 Redox homeostasis, 88, 98 Redox reactions, 49 Reductive sinks in microbes carbon monoxide (CO) dehydrogenase (CODH), 76–77 fermentation, 61–66 hydrogenases, 72–74 nitrate reductase, 68–69 phenazine production, 69–72 polymer deposition, 66–68 reverse TCA (rTCA) cycle, 74–76 in mycobacteria, 78 Dos dormancy regulon, 81–90 intracellular redox environment, 78–81 WhiB proteins, as intracellular redox sensor, 90–97 Reductive stress, 46, 49, 97 concept of, 49–51

SUBJECT INDEX Regulatory AAA+ protein, 123 Regulatory protein, 70 Reverse TCA (rTCA) cycle, 74–76 enzymes for, 74–75 Rhizobium etli, 67 Rhodococcus opacus, 68 Rhodococcus ruber, 67 Ribulose bisphosphate carboxylase/ oxygenase (RubisCO), 19 16S rRNA gene sequences, 6

S Saccharomyces cerevisiae fermentation, 61–64 FRDs, 75 Salmonella, 55, 65, 73 Sense-and-lock model, 86 Shigella, 65 Sigma factor, 97, 124 Single point mutation, 91 Sorbitol, 50 ssrA-tagged proteins, 128 Staphylococcaceae, 127 Staphylococcus albus, 70 Staphylococcus aureus, 77, 78, 134 Staphylococcus carnosus, 89 Streptococcus mutans, 128 Streptococcus pneumoniae, 128 in mice, 129 nox gene, 66 Streptococcus salivarius, 125, 127 Streptococcus thermophilus, 125, 127 Streptomyces spp., 70, 90 Stress protein, 82 Succinate dehydrogenase (SDH), 53 Succinic semialdehyde (SSA), 75 Succinic semialdehyde dehydrogenase (SSADH), 75 SufBCD system, 93 Super-XDR (S-XDR), 47

173

T TAG anabolism, 68 TAG-producing bacteria, 67 TAG production, 90, 93 TCA cycle, 19, 33, 52–54, 67 Thaumarchaeota, 20, 23, 29 Thiol-oxidising agent diamide, 62 Thioredoxin, 79 Transcription factors, 95, 127 Transhydrogenase activity, 64 a,a’-Trehalose dimycolate (TDM), 93 a,a’-Trehalose monomycolate (TMM), 93 Tuberculosis (TB), 45 infection model, 84 treatment regimes, 47 Tuberculous granuloma, 48

V Virulence factors, 129

W Warburg manometry, 58 Wastewater, treatment plants, 4 Wayne model of in vitro dormancy, 54, 68, 90 role in Mtb survival, 83 WhiB family of proteins, 90–91 and DNA binding, 95–97 and reductive stress, 93–95 and virulence, 91–93 Whole-genome shotgun (WGS), 6 Wood–Ljungdahl pathway, 76

X Xanthobacter flavus, 78

Z Zwitterion, 70

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