Advances in Microbial Physiology, Volume 57.pdf

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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Academic Press is an imprint of Elsevier 

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ISBN: 978-0-12-381045-8 

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Contents 

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

Ammonia-Oxidising Archaea – Physiology, Ecology and Evolution 

Christa Schleper and Graeme W. Nicol 

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 

2. Discovery of Archaea in Moderate Aerobic Habitats . . . . . . . . . . . . . . . . 4 

3. First Insights into the Physiology of Ammonia-oxidising Archaea . . . 6 

4. Model Organisms of Ammonia-Oxidising Archaea . . . . . . . . . . . . . . . . . . 9 

5. Membrane Lipids of Ammonia-Oxidising Archaea . . . . . . . . . . . . . . . . . 13 

6. Genomes and Metagenomes of Ammonia-Oxidising Archaea . . . . . . 15 

7. Diversity, Distribution and Activity of Ammonia-oxidising 

Archaea in the Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 

8. 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 

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 

2. Scope ............................................................... 46 

3. The Concept of Reductive Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 

4. Overview: General Physiological Characteristics of 

Mycobacterium Tuberculosis ........................................ 49

vi CONTENTS 

5. Reductive Sinks in Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 

6. Redox Sinks in Mycobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 

7. 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 

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 

1. Protein Quality Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 

2. CtsR-Regulated Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 

3. Cellular Functions of Genes Regulated by CtsR . . . . . . . . . . . . . . . . . . . 125 

4. Mechanisms for the Inactivation of the CtsR Repressor . . . . . . . . . . . 129 

5. Control of CtsR Degradation by the Regulated Adaptor McsB . . . . 136 

6. 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 


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 and 

Biotechnology, Aruna Asaf Ali Marg, New Delhi, India 

ANUP SRIVASTAVA, Department of Microbiology, University of Alabama at 


ADRIE J.C. STEYN, Department of Microbiology, University of Alabama at 

Birmingham, AL, USA


Ammonia-Oxidising Archaea – Physiology, 

Ecology and Evolution 

Christa Schleper 1 and Graeme W. Nicol 2 

1 Department of Genetics in Ecology, University of Vienna, Vienna, Austria 

2 Institute of Biological & Environmental Sciences, Cruickshank Building, 

University of Aberdeen, Aberdeen, UK 

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 Copyright Ó 2010 by Elsevier Ltd. 

ISSN: 0065-2911 All rights reserved 

DOI:10.1016/B978-0-12-381045-8.00001-1

2 CHRISTA SCHLEPER AND GRAEME W. NICOL 

organisms. The cultivation of a single marineisolateandtwoenrichments 

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. 

Abbreviations . . . ............................................ 3 

1. Introduction ................................................ 3 

2. Discovery of archaea in moderate aerobic habitats. . ................ 4 

3. First insights into the physiology of ammonia-oxidising archaea ........ 6 

4. Model organisms of ammonia-oxidising archaea. ................... 9 

5. Membrane lipids of ammonia-oxidising archaea .................... 13 

6. Genomes and metagenomes of ammonia-oxidising archaea .......... 15 

6.1. Metagenomic Studies of Uncultivated Ammonia Oxidisers. ....... 15 

6.2. Predictions from Complete Genome Sequences of Two Marine 

Archaea . . . ............................................ 

16 

6.3. Ammonia Oxidisers: A Distinct Phylum within the Archaea . . . .... 19 

7. Diversity, distribution and activity of ammonia-oxidising archaea in the 22 

environment ................................................ 

7.1. AOA in the Soil Environment ............................... 23 

7.2. AOA Activity in the Soil Environment. . . ...................... 25 

7.3. AOA in the Marine Environment . ........................... 28 

7.4. AOA Activity in the Marine Environment ...................... 29 

7.5. AOA in Sediments ....................................... 30 

7.6. AOA in Geothermal Environments .......................... 32 

7.7. AOA Associated with Marine Invertebrates. ................... 33 

8. Concluding remarks . . . ....................................... 33 

Acknowledgement ........................................... 34 

References. ................................................ 34 

22

AMMONIA-OXIDISING ARCHAEA 3 

ABBREVIATIONS 

AOA ammonia-oxidising archaea 

AOB ammonia-oxidising bacteria 

GDGT glycerol dialkyl glycerol tetraether 

16S rRNA 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,


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),

[(Figure_1)TD$FIG] 

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). 

AMMONIA-OXIDISING ARCHAEA 5


marine and freshwater sediments (MacGregor et al., 1997;Schleperet al., 

1997; Vetriani et al.,1998;Keoughet 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;Teiraet 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 andamoB 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 andamoB in a BCA gene order and contrasted with the CAB 

arrangement found in ammonia-oxidising bacteria (AOB) (Nicol and


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.) 

AMMONIA-OXIDISING ARCHAEA 7



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



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 

to those found on the fosmid clone 54d9 (K€onneke et al., 2005). These 

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):


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 10 7 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 (


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

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

Characteristics Nitrosopumilus 

maritimus 

Cenarchaeum 

symbiosum 

Nitrosocaldus 

yellowstonii 

Nitrososphaera 

gargensis 

Origin Tropical marine Marine sponge symbiont Terrestrial hot 

Terrestrial warm spring 

aquarium 

spring 

Culture Pure laboratory Inside Axinella mexicana Laboratory 

Laboratory enrichment 

culture 

(aquarium) 

enrichment 

Affiliation Group 1.1a 

Group 1.1a 

HWCGIII Group 1.1b 

Thaumarchaeota Thaumarchaeota 

Thaumarchaeota 

Growth 

temperature 

28 C 10 C(8–18 C) 72 C (65–74 C) 46 C 

Shape Rod Curved rod Spherical Spherical, coccoid 

Genome 1.65 Mb 2.05 Mb n.d. >2.6 Mb, draft 

Reference K€onneke et al. (2005) Preston et al. (1996) de la Torre et al. (2008) Hatzenpichler et al. (2008), 

Spang et al. (2010) 

12 CHRISTA SCHLEPER AND GRAEME W. NICOL


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 

C 40 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;



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


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


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 (Beja 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


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 (c 554 and c 552) 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 (N 2O). 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 O 2 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



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 F0F 1-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 F0F 1-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).


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 acetyl- 

CoA 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


studies. Using a concatenated dataset of 53 ribosomal proteins from C. 

symbiosum which are shared by archaea and eukaryotes, Brochier- 

Armanet 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. 


Euryarchaeota 

Crenarchaeota 

Ribosomal proteins 

r-protein S25e + + + 



r-protein L13e + + 

r-protein L14e + (some) + + 

r-protein L34e + (some) + + 

r-proteins L38e W 

r-protein L29p + + + 

r-protein Lxa + (most) + 

Transcription/RNA polymerase 

rpoG (=rpo8) + + 

rpoA – single ORF + split split + 

rpoB – single ORF + W split + + 

Rpc34 + W + 

MBF 1 W + + + 

EIF 1 + + 

Korarchaeota 

(continued)


Table 2 (continued) 


Euryarchaeota 

Crenarchaeota 

DNA polymerases/replication 

DNA pol D + + + 

RPA (Eury-like) + + + 

RPA/SSB + + (some) +/( ) + 

>one PCNA gene + 

Topoisomerases 

Topo IB + 

Topo IA W + + + 

Reverse gyrase + (HT) + + 

Topo IIA W 

Cell division 

ESCRT-III + + 

Vps4 (CdvC) + + 

CdvA + + 

Fts Z + + + 

Smc + + + 

ScpA + ScpB + + (many) + 

Histones (H3/H4) 

+ + (exc. 2) + 

Repair 

Hef-nuclease + 

ERCC4-type nuclease + + + 

ERRC4-type helicase + 

RadB + 

HSP70s, GrpE, Hsp40 + + 

UvrABC + W 

Korarchaeota 

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).



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 AMMONIA- 

OXIDISING 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.



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


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 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.)


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 13 C–CO 2, 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


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.) 

26 CHRISTA SCHLEPER AND GRAEME W. NICOL



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,


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


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 

10 4 –10 5 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


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



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.)


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).


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


nitrogen into the World’s ecosystems is now estimated to be more than 

double that from natural processes (Rockstr€om et al., 2009). The major 

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 Farhana 1 , Loni Guidry 1 , Anup Srivastava 1 , Amit Singh 2 , 

Mary K. Hondalus 3 and Adrie J.C. Steyn 1 

1 Department of Microbiology, University of Alabama at Birmingham, AL, USA 

2 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 

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 CO 2). This increased reductive 



DOI:10.1016/B978-0-12-381045-8.00002-3

44 AISHA FARHANA ET AL. 

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. 

Abbreviations . . . ............................................ 44 

1. Introduction ................................................ 45 

2. Scope. . . .................................................. 46 

3. The Concept of Reductive Stress ............................... 47 

4. Overview: General Physiological Characteristics of Mycobacterium 49 

tuberculosis ................................................ 

4.1. Historic Overview. ....................................... 49 

4.2. Environmental Factors that Influence Metabolism .............. 50 

5. Reductive Sinks in Microbes . . ................................. 59 

5.1. Fermentation ........................................... 59 

5.2. Polymer Deposition ...................................... 64 

5.3. Nitrate Reductase ....................................... 66 

5.4. Phenazine Production . . . ................................. 67 

5.5. Hydrogenases . . . ....................................... 70 

5.6. The Reverse TCA (rTCA) Cycle . ........................... 72 

5.7. Carbon Monoxide (CO) Dehydrogenase (CODH). .............. 74 

5.8. Other Mechanisms ...................................... 75 

6. Redox Sinks in Mycobacteria. . ................................. 76 

6.1. The Mycobacterial Intracellular Redox Environment. . . .......... 76 

6.2. The Mtb 

Dos Dormancy Regulon ........................... 79 

6.3. Mtb WhiB3 is an Intracellular Redox Sensor 

88 

that Counters Reductive Stress. . ........................... 

7. Concluding Remarks . . ....................................... 95 

Acknowledgements . . . ....................................... 98 

References. ................................................ 98 

ABBREVIATIONS 

GSH glutathione 

GSSG glutathione disulfide (oxidised glutathione) 

49 

88

REDUCTIVE STRESS IN MICROBES 45 

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-Guerin) 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


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


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 (CO 2) 

(Florczyk et al., 2003), oxygen (O 2)(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, 

E 0’ = 315 mV; NADP + /NADPH, E 0’ = 320 mV; FAD/FADH2, 

E 0’ = 219 mV; 2GSH/GSSG, E hc = 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


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’, orreferredtoasthe‘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


O 2 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 O 2. 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


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 FADH 2 

(Vemuri et al., 2006) as depicted below: 

Glucose + 8NAD + + 2NADP + + 2FAD + 4ADP + 4Pi 

! 6CO2 + 8NADH + 2NADPH + 2FADH2 + 4ATP + 10H +


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


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 O 2,isused 

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


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 C 16 FA such as 

palmitate (C 16H 32O 2, a putative in vivo carbon substrate of Mtb) ismore 

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,


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


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 CO 2, 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


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


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 pO 2 

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 aClarkeelectrode 

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,


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 m 2 , 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 (KmO2 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


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. 

CO 2 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 CO 2 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 O 2 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 O 2 

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


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) isaparticularly 

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.


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 glycerol- 

3-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


NADH, based upon the absence of transhydrogenase activity (NADH 

+NADP + $ NAD + +NADPH)inyeasts(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 O 2, energy must be supplied by 

either anaerobic respiration coupled to electron acceptors such as nitrate 

(NO 3 ) 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


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 H 2 and CO 2 (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 CO 2 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 H 2, 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, CO 2 and NH 3. 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 O 2 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 CO 2 are the 

excreted products that function as sinks for accumulating reducing equivalents 

to maintain redox balance (Wolfe, 2005).


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


Dawes, 1973). The central regulator of PHA production is the flux of acetyl- 

CoA, 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


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


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 NO 3 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-


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 (E 0’ = 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 Fe 3+ to the more soluble Fe 2+ 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 +


ratios of


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 (H 2ases) 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). H 2ases 

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


(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 H 2ases 

(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 H 2 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 (Pfenninger- 

Li 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 H 2 during citrate fermentation by 

aH 2: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 (S 0 ) is present, H 2S(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 

H 2ase couples the oxidation of ferredoxin and the reduction of 

2H + to H 2 production. Next, a membrane-bound oxidase oxidises ferredoxin 

and reduces NADP, which is used by a NAD(P)H S 0 oxidoreductase to 

reduce S 0 to H 2S. Thus, P. furiosus disposes of excess reductant using protons 

or S 0 as electron acceptors to yield H 2S(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 H 2 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


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 H 2 or produce H 2, 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 H 2ases 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 H 2ases 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 CO 2 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


(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 FADH 2,FMNH 2 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 CO 2, 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


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 CO 2 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% CO 2 (Florczyk 

et al., 2003). Furthermore, CO 2 preserves microaerophilic growth of 

Mycobacterium bovis BCG and prevents growth inhibition when O2 is 

rapidly removed (Florczyk et al., 2003), suggesting a role of CO 2 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 + H 2O $ CO 2 +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 CO 2, 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 acetyl- 

CoA synthase (ACS) (CODH/ACS), acetyl-CoA is generated from CO2,


CoA and a methyl group. In this scenario, CODH/ACS reduces CO 2 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 13 C-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 CO 2 

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 CO 2 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


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


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).


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


(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


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)andM. 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 O 2 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 O 2, 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


(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 O 2 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,


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 O 2 sensor showed that the pO 2 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


the low O 2 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 (NO 3 , 

NO2 ) across the alveolar capillary membrane followed by aerosolisation of 

airway epithelial lining fluid and confinement of NO2 /NO 3 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 NO 3 

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 O 2 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 HO- 

1 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


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 [KmO2 

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 (Fe 2+ ) whereas NO can react with both 

Fe 2+ and ferric iron (Fe 3+ )(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 Fe 2+ state, thereby preventing 

oxidation to the Fe 3+ 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


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; Berrios- 

Rivera 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.



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 CO 2 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 

NO 3 by NarK2 and its corresponding reduction to NO 2 by NarG consumes cellular 

reducing equivalents. As the concentration of cytosolic NO 2 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.


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 NO 3 /NO 2 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 

NO 3 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 O 2 (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 NO 3 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 NO 3 into and NO 2 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 NO 3 and/or NO 2 in vivo? 

Indeed, the presence of NO3 plus NO 2 in saliva (313 mM), NO 3 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


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 

NO 2 and/or NO 3 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


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


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 O 2 

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


(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


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 NO 3 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,


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 [ 14 C] 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



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, apo- 

WhiB binding was also demonstrated for Mtb WhiB2 and a mycobacteria 

phage TM4 WhiB homologue, WhiBTM4 (Rybniker et al., 2010), suggesting


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, ERG ox/ERG red) 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


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.


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. 13 C-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


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. 

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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. 

Abbreviations . . . ........................................... 120 

1. Protein Quality Control. . ..................................... 120 

2. Ctsr-Regulated Genes . . ..................................... 122 

3. Cellular Functions Of Genes Regulated by CtsR. .................. 125 

4. Mechanisms for the Inactivation of the CtsR Repressor ............. 129 

4.1. Heat Inactivation of CtsR. . ............................... 129 

4.2. CtsR Inactivation During Oxidative Stress ................... 132 



DOI:10.1016/B978-0-12-381045-8.00003-5

120 ALEXANDER K.W. ELSHOLZ ET AL. 

4.3. CtsR Inactivation During Other Stress Conditions ............. 134 

5. Control of Ctsr Degradation by the Regulated Adaptor McsB . ........ 136 

6. Summary And Outlook. . ..................................... 136 

Acknowledgement .......................................... 137 

References. ............................................... 137 

ABBREVIATIONS 

HTH helix-turn-helix 

HHP high-hydrostatic pressure 

LMWPTP 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 Hayer- 

Hartl, 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).


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 s 32 (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 

clpP in B. subtilis (Kr€uger et al., 1996; Gerth et al., 1998) as well as for the 

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


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;


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 ALEXANDER K.W. ELSHOLZ ET AL.


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


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., 

1998; Kr€uger et al., 2001; Pan et al., 2001; Nakano et al., 2002; Zellmeier et al., 

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) andO. oeni (Beltramo et al., 2004). ClpP is also responsible for the


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) andS. 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 

promising target for newly developed antibiotics (Br€otz-Oesterhelt et al., 

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),whichareneededforanefficient 

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) andL. monocytogenes (Rouquette et al., 

1996), but a clpC deletion in L. lactis (Ingmer et al., 1999)andS. 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 (Derre et al., 1999a), whereas a clpE deletion in 

L. lactis (Ingmer et al., 1999)orS. 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


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 (Borezee et al., 2001) and listeriolysin O 

(Rouquette et al., 1998)inL. 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)andL. 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).


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) (Derre 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 

became phosphorylated by McsB during heat stress (Kr€uger et al., 2001). 

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


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 

shockregulatorssuchass 32 (Bukau, 1993) orHrcA(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



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).


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 

(Kr€uger et al., 1994), S. aureus (Fleury et al., 2009) and L. lactis (Elsholz 

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.,


2003), H 2O 2 (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


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


(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.


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+


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 

The authors are grateful to Volker Br€ozel (South Dakota State Univ., USA) 

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) 

(c) 

(d) 

[(Plate_2)TD$FIG] 

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.)

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 

Author Index 

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 AUTHOR INDEX 

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 

Beja, 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 

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 

Bessieres, 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 

Borezee, 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 147 

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 

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


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 

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 

Daffe, M., 93 

Daims, H., 11–13, 33 

Daldal, F., 95 

Damste, 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


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 

Derre, 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 

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


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 

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


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 

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


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 

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


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 

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


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 

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


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-Garcıa, 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 

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


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 

Medard, 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 

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 

Moliere, 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


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 

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


Pan, Q., 128, 130 

Pancost, R.D., 6 

Pane-Farre, 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 

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


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 

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 

Rodrıguez-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


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 

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


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 

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


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 

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


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 

Vazquez-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 

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 

Villen, 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


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 

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


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 

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


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 

TEX 86 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 SUBJECT INDEX 

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 

Azotobacter beijerinckii, 66 

Azotobacter vinelandii, 64, 67 

B 

BAC-derived fosmid vector, 15 

Bacillaceae, 127 

Bacillales, 124, 131 

Bacille Calmette-Guerin (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 169 

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 

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


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 

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


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 

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


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 

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


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 

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

Advances in Microbial Physiology, Volume 57.pdf

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