Glutathione, Altruistic Metabolite in Fungi The Role of the Flavodiiron Proteins in Microbial Nitric...

Contents

CONTRIBUTORS TO VOLUME 49 vii

Glutathione, Altruistic Metabolite in FungiIstvan Pocsi, Rolf A. Prade and Michel J. Penninckx

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31. Introduction – Why study glutathione in fungi? . . . . . . . . . . . . 42. Glutathione and other low molecular weight

non-protein thiols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63. GSH metabolism in fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94. Glutathione in stress responses . . . . . . . . . . . . . . . . . . . . . . . . 235. GSH-dependent detoxification processes . . . . . . . . . . . . . . . . . 406. Aging and autolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 487. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

The Role of the Flavodiiron Proteins in Microbial NitricOxide Detoxification

Lıgia M. Saraiva, Joao B. Vicente and Miguel Teixeira

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 782. The family of flavodiiron proteins . . . . . . . . . . . . . . . . . . . . . . 883. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119

Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

Stress Responsive Bacteria: Biosensors as Environmental MonitorsAmy Cheng Vollmer and Tina K. Van Dyk

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1332. Reporters of gene expression. . . . . . . . . . . . . . . . . . . . . . . . . . 1363. Macromolecular damage. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1424. Nutrient limitation/imbalance . . . . . . . . . . . . . . . . . . . . . . . . . 1505. Panels and arrays. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1556. Future trends. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162


Bacterial Naþ- or H

þ-coupled ATP Synthases Operating at

Low Electrochemical PotentialPeter Dimroth and Gregory M. Cook

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1762. ATP synthesis in anaerobic bacteria at low electrochemical

potential . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1793. Alkaliphilic bacteria growing at low �mHþ . . . . . . . . . . . . . . . 200


Dissimilatory Fe(III) and Mn(IV) ReductionDerek R. Lovley, Dawn E. Holmes and Kelly P. Nevin

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2212. Environmental considerations . . . . . . . . . . . . . . . . . . . . . . . . . 2223. Major groups of Fe(III)- and Mn(IV)-reducing

microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2374. Physiological diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2435. Mechanisms for Fe(III) and Mn(IV) reduction . . . . . . . . . . . . . 2546. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

Author index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 000Subject index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 000

vi CONTENTS

Contributors to Volume 49

GREGORY M. COOK, Department of Microbiology, Otago School of MedicalSciences, University of Otago, P.O. Box 56, Dunedin, New Zealand

PETER DIMROTH, Institut fur Mikrobiologie, Eidgenossische TechnischeHochschule, ETH-Zentrum, Schmelzbergstrasse 7, CH-8092 Zurich,Switzerland

DAWN E. HOLMES, Department of Microbiology, University ofMassachusetts-Amherst, Amherst, MA 01003, USA

DEREK R. LOVLEY, Department of Microbiology, University ofMassachusetts-Amherst, Amherst, MA 01003, USA

KELLY P. NEVIN, Department of Microbiology, University ofMassachusetts-Amherst, Amherst, MA 01003, USA

MICHEL J. PENNINCKX, Laboratoire de Physiologie et Ecologie Microbienne,Universite Libre de Bruxelles, c/o Institut Pasteur, 642 Rue Engeland,B-1180 Brussels, Belgium

ISTVAN POCSI, Department of Microbiology and Biotechnology, Facultyof Sciences, University of Debrecen, P.O. Box 63, H-4010, Debrecen,Hungary

ROLF A. PRADE, Department of Microbiology and Molecular Genetics,Oklahoma State University, Stillwater, OK 74078, USA

LIGIA M. SARAIVA, Instituto de Tecnologia Quımica e Biologica, Universi-dade Nova de Lisboa, Apartado 127 Avenida da Republica (EAN),2781-901 Oeiras, Portugal

MIGUEL TEIXEIRA, Instituto de Tecnologia Quımica e Biologica,Universidade Nova de Lisboa, Apartado 127 Avenida da Republica(EAN), 2781-901 Oeiras, Portugal

TINA K. VAN DYK, DuPont Central Research and Development, Exper-imental Station E173/216, P.O. Box 80173, Wilmington, DE 19880-0173,USA

JOAO B. VICENTE, Instituto de Tecnologia Quımica e Biologica, UniversidadeNova de Lisboa, Apartado 127 Avenida da Republica (EAN), 2781-901Oeiras, Portugal

AMY CHENG VOLLMER, Department of Biology, Swarthmore College,500 College Avenue, Swarthmore, PA 19081, USA

viii CONTRIBUTORS TO VOLUME 47

Glutathione, Altruistic Metabolite in Fungi

Istvan Pocsi1, Rolf A. Prade2 and Michel J. Penninckx3,*

1Department of Microbiology and Biotechnology, Faculty of Sciences, University of

Debrecen, P.O. Box 63, H-4010 Debrecen, Hungary2Department of Microbiology and Molecular Genetics, Oklahoma State University,

Stillwater, OK 74078, USA3Laboratoire de Physiologie et Ecologie Microbienne, Universite Libre de Bruxelles,

c/o Institut Pasteur, 642 Rue Engeland, B-1180 Brussels, Belgium

Though the mills of God grind slowly,yet they grind exceedingly small;Though with patience He stands waiting,with exactness grinds He all.

Friedrich von Logau (1614–1655)

ABSTRACT

Glutathione (GSH; g-L-glutamyl-L-cysteinyl-glycine), a non-protein thiolwith a very low redox potential (E0

0 ¼ �240 mV for thiol-disulfideexchange), is present in high concentration up to 10 mM in yeasts andfilamentous fungi. GSH is concerned with basic cellular functions as wellas the maintenance of mitochondrial structure, membrane integrity, andin cell differentiation and development. GSH plays key roles in theresponse to several stress situations in fungi. For example, GSH is animportant antioxidant molecule, which reacts non-enzymatically witha series of reactive oxygen species. In addition, the response to

*Corresponding author. Tel.: 32 2 3733303; Fax: 32 2 3733309;

E-mail: [email protected]

ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 49 Copyright � 2004, Elsevier Ltd.

ISBN 0-12-027749-2 All rights reserved.

DOI 10.1016/S0065-2911(04)49001-8

oxidative stress also involves GSH biosynthesis enzymes, NADPH-dependent GSH-regenerating reductase, glutathione S-transferase alongwith peroxide-eliminating glutathione peroxidase and glutaredoxins.Some components of the GSH-dependent antioxidative defence systemconfer resistance against heat shock and osmotic stress. Formation ofprotein–SSG mixed disulfides results in protection against desiccation-induced oxidative injuries in lichens. Intracellular GSH and GSH-derivedphytochelatins hinder the progression of heavy metal-initiated cellinjuries by chelating and sequestering the metal ions themselves and/or byeliminating reactive oxygen species. In fungi, GSH is mobilized to ensurecellular maintenance under sulfur or nitrogen starvation. Moreover,adaptation to carbon deprivation stress results in an increased toleranceto oxidative stress, which involves the induction of GSH-dependentelements of the antioxidant defence system. GSH-dependent detoxifica-tion processes concern the elimination of toxic endogenous metabolites,such as excess formaldehyde produced during the growth of themethylotrophic yeasts, by formaldehyde dehydrogenase and methyl-glyoxal, a by-product of glycolysis, by the glyoxalase pathway.Detoxification of xenobiotics, such as halogenated aromatic andalkylating agents, relies on glutathione S-transferases. In yeast, theseenzymes may participate in the elimination of toxic intermediates thataccumulate in stationary phase and/or act in a similar fashion as heatshock proteins. GSH S-conjugates may also form in a glutathione S-transferases-independent way, e.g. through chemical reaction betweenGSH and the antifugal agent Thiram. GSH-dependent detoxification ofpenicillin side-chain precursors was shown in Penicillium sp. GSHcontrols aging and autolysis in several fungal species, and possesses ananti-apoptotic feature.

Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31. Introduction – Why study glutathione in fungi? . . . . . . . . . . . . . . . . . . . 42. Glutathione and other low molecular weight non-protein thiols . . . . . . . . . . 63. GSH metabolism in fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

3.1. The g-glutamyl cycle, biosynthesis and degradation of GSH. . . . . . . . 93.2. GSH metabolism under unstressed conditions . . . . . . . . . . . . . . . 113.3. Uptake and storage of GSH and GSH-conjugates . . . . . . . . . . . . . 133.4. Degradation and recycling of GSH . . . . . . . . . . . . . . . . . . . . . . 153.5. Stabilization of physiological GSH/GSSG redox

balance – glutathione reductases . . . . . . . . . . . . . . . . . . . . . . . 173.6. GSH – extracellular functions . . . . . . . . . . . . . . . . . . . . . . . . . 173.7. GSH in cell differentiation and development . . . . . . . . . . . . . . . . . 18

2 ISTVAN POCSI, ROLF A. PRADE AND MICHEL J. PENNINCKX

3.8. Is GSH essential in unstressed cells? . . . . . . . . . . . . . . . . . . . . . 194. Glutathione in stress responses . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

4.1. Oxidative stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234.2. Heat and osmotic shock . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334.3. Desiccation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 344.4. High cell density cultures. . . . . . . . . . . . . . . . . . . . . . . . . . . . 344.5. Heavy metal stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354.6. Nutrient deprivation stress . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

5. GSH-dependent detoxification processes. . . . . . . . . . . . . . . . . . . . . . 405.1. Elimination of toxic metabolites . . . . . . . . . . . . . . . . . . . . . . . . 405.2. Detoxification of xenobiotics . . . . . . . . . . . . . . . . . . . . . . . . . . 425.3. Glutathione, regulator of b-lactam antibiotic synthesis . . . . . . . . . . . 44

6. Aging and autolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 487. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

ABBREVIATIONS

ACV d-(L-a-aminoadipyl)-L-cysteinyl-D-valineCG L-cysteinyl-glycine dipeptidaseCPC cephalosporin CDMDT dimethyldithiocarbamic acidFaDH formaldehyde dehydrogenasegGCS g-L-glutamyl-L-cysteinyl synthetaseGLO glyoxalaseGPx glutathione peroxidaseGR glutathione reductaseGrx glutaredoxinGS glutathione synthetaseGSH glutathioneGSSG glutathione disulfideGST glutathione S-transferaseGS-X glutathione S-conjugategGT g-glutamyltranspeptidaseNES nuclear export sequenceNNG N0-nitro-N-nitrosoguanidinesNPT non-protein thiolPC phytochelatinROS reactive oxygen speciesSOD superoxide dismutaseTrx thioredoxin

GLUTATHIONE METABOLISM IN FUNGI 3

1. INTRODUCTION – WHY STUDY GLUTATHIONE IN FUNGI?

Fungi, in particular the baker’s yeast Saccharomyces cerevisiae, have servedas biological model systems that assisted the unraveling of the role andfunction of glutathione in cellular processes. This molecule was initiallydescribed as ‘‘philothion’’ by Rey-Pahlade over 120 years ago, as a sub-stance having the property to reduce elemental sulfur releasing hydrogensulfide (Meister, 1988). This ‘‘sulfur-loving’’ compound was isolated by theEnglish biochemist Sir Frederic Gowland Hopkins and renamedglutathione. In terms of chemistry, glutathione (GSH) was found to be athiol tripeptide with an unusual g-glutamyl linkage (g-L-glutamyl-L-cysteinyl-glycine). Research on the role and function of GSH in animaltissues has been active during the last 40 years, and enriched withcontributions of Alton Meister and his colleagues (Meister and Anderson,1983). Most research studies focusing on animal GSH are multidisciplinary,comprising biochemical, physiological, toxicological and clinical aspectsof its biological role. In contrast, the biology of GSH in microbialsystems has received less attention, even though it is widely recognized thatGSH is a physiologically relevant non-protein thiol (NPT) present inmost microorganisms as reviewed by Penninckx and Elskens (1993) andHell (1997).The biology of GSH has been approached in at least two different

ways. First, in biological systems where GSH molecules are abundant(in some types of cells GSH can make up to 1% of the dry weight(Penninckx et al., 1980)) studies commonly focus on ‘‘in vivo’’ biochemicalstudies, and second, GSH metabolism has been examined indirectly as acomponent of other cellular processes (Fig. 1).For example, GSH metabolism is involved in stress response,

specifically the detoxification of oxidative stressed cells (Grant andDawes, 1996; Grant et al., 1996a; Stephen and Jamieson, 1996; Emri et al.,1997a, 1999a; Gasch et al., 2000; Grant, 2001). Because GSH metabolismis tightly associated with the response to oxidative and other kindsof environmental stress and balancing redox potentials in different sub-cellular compartments, its understanding has also been of interest tothe food and pharmaceutical industries, which frequently use yeastsand filamentous fungi under stressful conditions in bioprocesses (Walker,1998).The last review considering GSH metabolism in microorganisms

appeared in 1993 (Penninckx and Elskens, 1993) and recent reviews des-cribe specialized aspects in yeast (Penninckx, 2000, 2002). The purpose of


this review is to offer a current overview of GSH metabolism focused onyeast and filamentous fungi, but relating to bacterial, animal and plantsystems where needed. We hope that this contribution will inspire newresearch in the future.

De novo GSH

synthesis

Sulphate reduction

Asexual and sexual

sporulation

Proper assembly of microtubuli

NADPH production in pentose phosphate

shunt

Amino acid and

ion transport Reduction

of gluta-redoxins

Synthesis of deoxy-

ribose

Cys, Glu and Gly synthesis

Detoxifi-cation of drugs and

xenobiotics

Elimination

of ROS

Anti-ageing and anti-apoptotic

effects

Stabilisation of cellular membranes and proteins

Sulphur storage

GSSG

GSH

Reduc-tion of GSSG

Protection of mito-

chondrial DNA

Detoxifi-cation of

heavy metals

GSH and GSSG

GSH production

GSH consumption - dispensible

GSH consumption - indispensible

Figure 1 GSH production and consumption machines in the metabolic networkof fungi.


2. GLUTATHIONE AND OTHER LOW MOLECULAR WEIGHTNON-PROTEIN THIOLS

GSH and other NPTs are present in all living cells from bacteria (Fahey,2001), fungi, plants (Hell, 1997) and mammals (Meister and Anderson,1983). The biological significance of GSH and NPTs in general is directlyassociated with the strong reducing potential derived from the free sulfydrylgroup found in these molecules.The reduced sulfydryl group in GSH, when oxidized, produces a disulfide

bond between two GSH molecules forming glutathione disulfide, or‘‘oxidized glutathione’’ (GSSG). The reduced form of GSH (E0

0 ¼�0.24 Vfor thiol-disulfide exchange) is maintained by NADPH-dependentglutathione reductase (GR) (Table 1), and the GSH/GSSG ratio found incells is typically greater than 20 (Emri et al., 1997a). The equilibriumbetween the formation and dissolution of disulfide bonds among GSHmolecules driven by the redox potential of a cellular state creates a highcapacity oxidoreductive buffer within the cell (Meister and Anderson, 1983),as originally suggested by the French discoverer of GSH, Rey-Pahlade(Meister, 1988).Owing to its thiol group, GSH is also a strong nucleophile undergoing

conjugation with a range of electrophilic compounds including coenzyme A,cysteine, proteins (Meister and Anderson, 1983; Penninckx and Elskens,1993; Hell, 1997; Fahey, 2001) and numerous xenobiotics (Vuilleumier,1997). Another advantageous chemical characteristic of GSH is that itsunusual g-glutamyl peptide bond results in an increased resistance towardsproteolytic degradation (Meister and Anderson, 1983).In most eukaryotes, GSH appears to be the most abundant low molecular

weight thiol (Fahey et al., 1984). An alternative, trypanothione, the oxidizedform of N1,N6-bis(glutathionyl) spermidine, is found in trypanosomatidssuch as Trypanosoma cruzi and the insect-parasitic Crithidia fasciculata(Fairlamb et al., 1985). On the other hand, in prokaryotes, GSH ispredominant only in aerobic, gram-negative bacteria, and is less frequent inanaerobic, gram-positive bacteria. However, bacteria with no detectableamounts of GSH accumulate other non-proteinous, low molecular weightthiols, many of which remain structurally unknown (reviewed in Fahey,2001). Thus, the wide occurrence of free thiols in cells may indicate acrucial role for thiol-based redox buffer systems in the maintenance ofcellular metabolism. Some of the GSH alternate thiol forms described inbacteria are mercaptoethanesulfonic acid (coenzyme M) and g-L-glutamyl-L-cysteine in Archaebacteria (Newton and Javor, 1985, Ferry, 1994)


Table 1 Principal enzyme and transport systems associated with different functions ofGSH in fungi.

1) Biosynthesis of GSH

g-L-glutamyl-L-cysteine synthetase (gGCS: EC 6.3.2.2): þ L-glutamateþ L-cysteineþATP

! g-L-glutamyl-L-cysteineþADP phosphate

Genes: S. cerevisiae: GSH1 (Ohtake and Yabuchi, 1991); S. pombe: gcs1 (Mutoh et al.,1995; Wood et al., 2002); Candida albicans: GCS1/GSH1 (Baek et al., 1999); Neurosporacrassa: GSH1 (Mannhaupt et al., 2003); Hansenula polymorpha: GSH2 (Ubiyvovk et al.,2002).

Comments: gGCS has not been purified in fungi; subunit size is from 65 to 82 kDa asdeduced from gene sequences. Gene disruption was lethal in S. cerevisiae in the absenceof exogenous GSH.

Glutathione synthetase (GS: EC 6.3.2.3): g-L-glutamyl-L-cysteineþ glycineþATP !

GSHþADP

Genes: S. cerevisiae: GSH2 (Mooz and Meister, 1967; Goffeau et al., 1996); S. pombe(Nakagawa et al., 1993; Wood et al., 2002; Phlippen et al., 2003); C. albicans (Baek et al.,1999); Pichia angusta (Berardi et al., 2001); Nectria lugdunensis (Braha et al., 2003);Pneumocystis carinii (Smulian et al., 2001); Aspergillus niger (Murata et al., 1989).

Comments: GSs were purified in S. cerevisiae and S. pombe. In S. cerevisiae, the enzyme isa homodimer of two identical subunits of 56 kDa. In S. pombe, GS is either a homodimerof two identical 56 kDa subunits or a heterotetramer of two 32 kDa and two 24 kDasubfragments. Gene disruption was not lethal in S. cerevisiae.

2) Degradation

g-L-glutamyl transpeptidase (gGT: EC 2.3.2.2): GSHþ amino acid (H2O) ! g-L-glutamyl-

amino acid (L-glutamate)þ L-cysteinyl-glycine

Genes: S. cerevisiae: CIS2 (ECM38) (Mehdi et al., 2001; Kumar et al., 2003b); S. pombe(Wood et al., 2002); C. albicans (ca4913 at http://mips.gsf.de/proj/yeast/CYGD).

Comments: In S. cerevisiae, the non-glycosylated precursor form of gGT is 73 kDa.The native glycosylated enzyme is about 90 kDa. The enzyme is a heterodimer with one64 kDa and one 29 kDa subunit. Gene disruption was not lethal in S. cerevisiae.

3) Redox balance

Glutathione reductase (GR: EC 1.6.4.2): GS-SGþNADPHþHþ! 2 GSHþNADPþ

Genes: S. cerevisiae: GLR1 (Muller, 1996); S. pombe: pgr1þ (Lee et al., 1997); N. crassa(CAD70360) (Schulte et al., 2003); C. albicans (Kim et al., 1999).

Comments: GR is a homodimer of two identical subunits in S. cerevisiae. Gene disruptionwas not lethal in S. cerevisiae but was lethal in S. pombe.

(Continued )


Table 1 Continued.

Glutaredoxin (Grx): 2 GSHþ oxidized glutaredoxin ! GSSGþ reduced glutaredoxin

Genes: S. cerevisiae: GRX1, GRX2/TTR2 (Goffeau et al., 1996); GRX3-5 (Luikenhuiset al., 1998; Rodrıguez-Manzaneque et al., 1999); S. pombe: GLR1 and GLR2 (Wood et al.,2002); Encephalitozoon cuniculi (Katinka et al., 2001); N. crassa (Schulte et al., 2001a).

Comments: Grxs are monomeric proteins of 12.4 kDa in S. cerevisiae; grx1 grx2 nullmutants are viable but are sensitive to oxidative stress (Luikenhuis et al., 1998; Collinsonand Grant, 2003).

Glutathione peroxidase: (GPx: EC 1.11.1.9.) 2 GSHþROOH ! GSSGþH2OþROH

Genes: S. cerevisiae: GPX1 and GPX2 (Avery and Avery, 2001); S. pombe: GPX1 (Woodet al., 2002); Candida boidinii (Horiguchi et al., 2001); Blumeria graminis (Zhang andGurr, 2000).

Comments: GPx is a monomer of 19.5 kDa in S. cerevisiae; the null mutant is viable.

4) Detoxification

Glyoxalase I (GLO1: EC 4.4.1.5): GSHþmethylglyoxal ! (R)-S-lactoyl GSH

Genes: S. cerevisiae GLO1 (Inoue and Kimura, 1996); S. pombe (Wood et al., 2002);N. crassa (Schulte et al., 2001b); Paracoccidioides brasiliensis (Castro et al., 2003).

Comments: Glyoxalase I is a monomer with a molecular mass of 37.2 kDa in S. cerevisiae;a GLO1 null mutant is viable but has increased sensitivity to methylglyoxal.

Glyoxalase II (GLO2: EC 3.1.2.6): (R)-S-lactoyl GSHþH2O ! GSHþD (�) Lactic

acid

Gene: S. cerevisiae GLO2 (Bito et al., 1997).

Comments: Glyoxalase II is also monomeric with a molecular mass of 31.3 kDa in S.cerevisiae; a GLO2 null mutant is viable but shows increased sensitivity to methylglyoxal.

Formaldehyde dehydrogenase (FaDH: EC 1.1.1.1): FormaldehydeþGSHþNADþ!

S-formylglutathioneþNADHþHþ

Genes: S. cerevisiae (Wehner et al., 1993; Fernandez et al., 1999); S. pombe (Wood et al.,2002); H. polymorpha (Baerends et al., 2002); Pichia pastoris (Shen et al., 1998); C. boidinii(Lee et al., 2002); Candida maltosa (Sasnauskas et al., 1992).

Comments: FaDH is monomeric (molecular mass 41 kDa) in S. cerevisiae; a null mutant isviable but sensitive to formaldehyde. Hansenula polymorpha null mutant accumulatestoxic formaldehyde when growing on methanol.

Glutathione S-transferase (GST: EC 2.5.1.18): GSHþRX ! GS � XþRH

Genes: S. cerevisiae GTT1 and GTT2 (Choi et al., 1998; Grant, 2001); URE2 (Coschiganoand Magasanik, 1991; Rai et al., 2003); S. pombe gst1þ , gst2þ and gst3þ (Veal et al.,2002); Aspergillus nidulans gstA (Fraser et al., 2002); N. crassa (Galagan et al., 2003);Cunninghamella elegans (Cha et al., 2001, 2002); Issatchenkia orientalis (Tamaki et al.,1990); Gibberella fujikuroi (Yoshida et al., 2001); Pichia augusta (Siverio, 2000);Botryotinia fuckeliana (Prins et al., 1998); Polymyxa betae (Mutasa-Gottgens et al., 1999);E. cuniculi (Katinka et al., 2001).

(Continued )


as well as 2-N-acetylcysteinyl)amido-2-deoxy-a-D-glucopyranosyl-(1! 1)myo-inositol (mycothiol, MSH) in Actinomycetes (Newton et al., 1996).Finally, polymeric forms of GSH (phytochelatins) (Rauser, 1995; Cobbettand Goldsbrough, 2002) and GSH transpeptidation products have also beenobserved in plants and fungi (Kean and Hare, 1980; Kasai et al., 1982;Jaspers et al., 1985).

3. GSH METABOLISM IN FUNGI

3.1. The g-glutamyl Cycle, Biosynthesis and Degradationof GSH

GSH metabolism comprises a ribosome-independent, tripeptide bio-synthetic and catabolic pathway also known as the g-glutamyl cycle

Table 1 Continued.

Comments: GSTs have also been characterized in Aspergillus ochraceus, Mucor javanicus,Phanerochaete chrysosporium, Yarrowia lipolytica, Penicillium crysogenum and inCandida, Hansenula, Penicillium, Pichia and Rhodotorula spp. (reviewed in Sheehanet al., 2001).

GSTs are homodimers of 22–26 kDa subunits in S. cerevisiae. In yeast, GTT1 andGTT2 null mutants are viable, but are heat shock sensitive at stationary phase.S. cerevisiae �URE2 mutants possess the same phenotypes as S. cerevisiae and S. pombeGST mutants. S. pombe gst1, gst2 and gst3 null mutants are sensitive to fluconazole.

5) Transport

Cell surface GSH transporters

Genes: S. cerevisiae OPT1/HGT1/GSH11 (Bourbouloux et al., 2000; Miyake et al., 2002;reviewed in De Hertogh et al., 2002); GSH P-2 (Miyake et al., 1998); S. pombe: isp4 (Satoet al., 1994); C. albicans (Lubkowitz et al., 1997); Y. lipolytica (Gonzalez-Lopez et al.,2002); E. cuniculi (Katinka et al., 2001).

Comments: Hgt1p possesses a molecular mass of 91.6 kDa in S. cerevisiae; null mutant isviable but exhibits loss of plasma membrane GSH transport.

Vacuolar GSH and GS-X transporters

Genes: S. cerevisiae YCF1 (Li et al., 1997; reviewed in Rosen, 2002); BPT1 (Goffeau et al.,1996; Sharma et al., 2002); C. albicans (Theiss et al., 2002).

Comments: Ycf1p is 171.1 kDa and its null mutant is viable but sensitive to Cd2þ . Bpt1pis 176.8 kDa and its null mutant is viable too but lacks approximately 40% of thetransport activity of unconjugated bilirubin into the yeast vacuole.


(Meister and Anderson, 1983). The complete cycle comprises six enzymaticreactions involving two ATP-dependent GSH biosynthetic steps (Reactions1 and 2 in Fig. 2) and four catabolic reactions (Reactions 3–6), with only onebeing ATP-dependent (Reaction 6).As in plant and animal cells (Meister and Anderson, 1983; Rennenberg

et al., 1980), a complete g-glutamyl cycle has been reported in S.cerevisiae (Mooz and Wigglesworth, 1976). However, Reactions 5 and6 (Fig. 2) could not be determined by other groups (Jaspers et al., 1985).Moreover, direct labelling experiments with (U-14C)-GSH and (14C)-5-oxoproline suggest a truncated g-glutamyl cycle, catalyzed by thebiosynthetic enzymes, g-glutamylcysteine synthetase (gGCS) (Reaction 1)and GSH synthetase (GS) (Reaction 2) and degraded by the catabolicenzymes g-glutamyltranspeptidase (gGT) (Reaction 3) and L-cysteinylgly-cine dipeptidase (Reaction 4) (Table 1) (Jaspers et al., 1985). On the otherhand, a gene homologous to rat 5-oxoprolinase has been found both in S.cerevisiae and Schizosaccharomyces pombe with 48.4 and 44% amino acididentity, respectively (Guo-jie et al., 1996). A g-glutamyl cycle may functionin the b-lactam-producing filamentous fungus Acremonium chrysogenumsupplying the cephalosporin C biosynthetic machinery with L-cysteine(Nagy et al., 2003).

Figure 2 Synthesis and degradation of GSH via the g-glutamyl cycle. (1) g-Gluta-mylcysteine synthetase; (2) GSH synthetase; (3) g-glutamyltranspeptidase; (4) cysteinyl-glycine dipeptidase; (5) g-glutamylcyclotransferase; (6) 5-oxoprolinase.


3.2. GSH Metabolism Under Unstressed Conditions

3.2.1. GSH Biosynthesis

The synthesis reaction catalyzed by gGCS involves the formation of an g-L-glutamylphosphate intermediate, followed by addition of L-cysteine andrelease of g-L-glutamyl-L-cysteine. gGCS is feedback-inhibited by GSH,preventing excessive accumulation (Meister and Anderson, 1983).Mammalian gGCS is a heterodimer one chain being the catalytic subunit

(72.8 kDa) GCLC, encoded by GSH1 and the other a modifier subunit (30.8kDa) GCLM, encoded by GSH0 (Huang et al., 1993). Furthermore, Gclm(�/�) knockout mice suggest that GCLM is not essential for viability,however; loss of the modifier subunit results in reduced GSH levels andcells susceptibility to oxidative stress (Yang et al., 2002). The reducedGSH levels in the absence of the GCLM regulatory subunit are the resultof a two-fold increase in the Km for glutamate and more effective feedbackinhibition of GSH on GCLC (Yang et al., 2002).Putative gGCS sequences are available now for different fungal species

(Table 1) including S. cerevisiae (Ohtake and Yabuchi, 1991) and S. pombe(Mutoh et al., 1991; Coblenz and Wolf, 1994). Annotation of all availablegGCS sequences indicated that eukaryotic genes are diverse but likely to beevolved from a bacterial ancestor (May and Leaver, 1994). Moreover,mammalian GCLM showed significant homology to Escherichia coli gGCS(Watanabe et al., 1986) and, therefore, the modifier subunit is also likelyto originate from a prokaryotic prototype of gGCS (Huang et al., 1993).The existence of gGCS modifier subunits in eukaryotes other than mammalsis yet to be demonstrated (Hussein and Walter, 1995; Lueder and Phillips,1996).The catalytic function of GS initiates with ATP-dependent phosphoryla-

tion of the substrate, g-L-glutamy-L-cysteine, yielding an acyl-phosphory-lated intermediate, followed by a ligation reaction with glycine bythe loss of an inorganic phosphate and releasing GSH. GSH syntheta-ses have been purified from budding yeast (Mooz and Meister, 1967),Aspergillus niger (Murata et al., 1989) and S. pombe (Nakagawa et al., 1993).The human enzyme shows cooperative binding for g-glutamyl substrate(Njalsson et al., 2001); however GSs from microorganisms followclassical Michaelis-Menten kinetics (Mooz and Meister, 1967;Meierjohann et al., 2002).Eukaryote GSs apparently form homodimers and in E. coli the same

catalytic activity is found as a homotetrameric protein (Meister, 1974;Murata et al., 1989; Njalsson et al., 2001; Meierjohann et al., 2002).


In S. pombe, a unique heterotetrameric GS association was foundgenerating controversy about the subunit association of GSs in general(Mutoh et al., 1991; Nakagawa et al., 1993; Wang and Oliver, 1997). In arecent study, S. pombe GS was purified as a His-tagged protein andrecovered as a homodimer (Mr,subunit¼ 56 kDa) and a heterotetramer of two32 kDa and two 24 kDa subunits, all peptides being encoded by a singlegene (GSH2) (Phlippen et al., 2003). In addition, it was demonstrated thatsubfragments were generated from the larger peptide by a metalloproteaseactive in cell-free extracts. The homodimer shows full in vivo and in vitroactivity and the physiological relevance of the cleavage reaction remainsunclear (Phlippen et al., 2003).In terms of cell physiology, GS is dispensable for growth under both

normal and oxidative stress conditions because the product of gGCS, thedipeptide g-L-glutamyl-L-cysteine, can substitute, at least in part, for GSH(Grant et al., 1997).

3.2.2. Regulation of GSH Biosynthesis

In S. cerevisiae, gGCS is present in normal conditions (Inoue et al., 1998a;Sugiyama et al., 2000a) and is transcriptionally regulated by the activatorprotein Yap1p (Lee et al., 1999; Carmel-Harel and Storz, 2000; Moye-Rowley, 2002). Yap1p, a b-ZIP containing transcription factor, respondsprimarily to oxidative stress by activating the expression of genes whosepromoters possess a functional Yap1p-response element (YRE) (Fernandeset al., 1997; Cyert, 2001; Toone et al., 2001). Yap1p function is not restric-ted to activation of stress genes, being also active under ‘‘normal’’ vegetativeconditions facilitating mitosis (Dumond et al., 2000).GSH regulates the expression of gGCS (GSH1) via Met4p, a

transcriptional activator of alternate sulfur source metabolism genes(Thomas and Surdin-Kerjan, 1997; Wheeler et al., 2002), and regulates GSHbiosynthesis in ‘‘unstressed’’ cells. The GSH1 promoter contains elementsthat are specifically recognized by Met4p, and the GSH1 promoteractivity was induced in Met4p-dependent manner in a GSH1 mutant, whichis devoid of GSH. The addition of exogenous GSH repressed gGCSexpression (Wheeler et al., 2002). Analysis of a �CIS2 mutant, which doesnot express gGT and hence cannot break down GSH, confirmed that GSHitself and not a GSH metabolic product is the regulatory molecule.However, this is not a general mechanism affecting all Met4p-regulatedgenes, as, for example, MET16 expression was unaffected in a GSH1mutant, and GSH was a poor repressor of MET16 in comparison to


methionine. GSH biosynthesis is thus apparently regulated in parallel withsulfate assimilation by the Met4p protein, but GSH1-specific mechanismsexist that respond solely to GSH availability (Wheeler et al., 2002).In addition, there is evidence that GSH levels are sensitive to the

availability and nature of nitrogen and carbon sources. For example,nitrogen-response elements are present in the promoters of both GSH1 andGSH2 (Springael and Penninckx, 2003), which are likely to be responsiblefor the GSH overproduction under nitrogen starvation (Mehdi andPenninckx, 1997). Moreover, the GSH concentrations observable in lowglucose fed-batch or chemostat cultures were 4 to 6 times higher than thoserecorded under ethanol-producing conditions in S. cerevisiae (Shimizu et al.,1991; Berthe-Corti et al., 1992).

3.3. Uptake and Storage of GSH and GSH-conjugates

GSH is taken up from the environment by transporters in microorganisms(Sherrill and Fahey, 1998), plants (Hell, 1997) and animals (Iantomasi et al.,1997). Evidence that GSH can be assimilated from the medium by yeast cellscame from a metabolic study using radioactively labeled forms of GSH(Jaspers et al., 1985), and the discovery of plasma membrane GSHtransporters (Miyake et al., 1998; Bourbouloux et al., 2000).The Hgt1p high affinity plasma membrane GSH transporter (GSH-P1;

Fig. 3) (Km¼ 54 mM) is a 799-amino acid polypeptide with a predictedmolecular mass of 91.6 kDa and with 12–14 transmembrane domains,which is encoded by the GSH11 gene in S. cerevisiae (Bourbouloux et al.,2000; Miyake et al., 2002). Homologues of Hgt1p are apparently restrictedto fungi, e.g. C. albicans (Lubkowitz et al., 1998; Hauser et al., 2000) andS. pombe (Sato et al., 1994), and plants (Bourbouloux et al., 2000), andconstitute a large family of transporters including also the phytosider-ophores involved in Fe(III) uptake in plants (Curie et al., 2001). Yeaststrains deleted in HGT1 do not transport GSH, and �HGT1�GSH1double mutants are not viable (Bourbouloux et al., 2000). HGT1/GSH11 isregulated by the availability and type of sulfur source (Miyake et al., 2002),and may also be responsive to nitrogen sources (Springael and Penninckx,2003).A second GSH transporter has not yet been identified even though

biochemical evidence has suggested the presence of another low affinitytransport system in both fungi and plants (GSH-P2; Fig. 3) (Miyakeet al., 1998; Foyer et al., 2001).


The ABC-type vacuolar transporter Ycf1p of S. cerevisiae is homologousto the human multidrug resistance protein MRP1 (Borst et al., 2000) butnot to Hgt1p, and has been shown to mediate low-affinity transport ofGSH and GSH-conjugates from the cytoplasm into the vacuoles (Li et al.,1996; Tommasini et al., 1996; Rebbeor et al., 1998; Mehdi et al., 2001)(Fig. 4). Btp1p, a close homologue of Ycf1p, can also participate in thevacuolar transport of GSH and GSH-conjugates but its contribution issubstantially less than that of Ycf1p (Petrovic et al., 2000; Sharma et al.,

Figure 3 Transport and metabolism of sulfur in S. cerevisiae. GSH can be taken upby the yeast cell through two transport systems, GSH-P1 (high affinity) and GSH-P2

(low affinity). Sulfur flows from GSH to other metabolites along the sulfur metabolicpathway. SO2�

4 is also taken up and metabolized as indicated. In the case of totalsulfur deprivation, GSH stored in the cell is used as an endogenous sulfur source.(1) Serine acetyltransferase; (2) cysteine synthase; (3) homoserine acetyltransferase;(4) homocysteine synthase; (5) g-cystathionine synthase; (6) g-cystathionase; (7) b-cystathionase; (8) b-cystathionine synthase; (9) homocysteine methyltransferase;(10) S-adenosylmethionine synthase; (11) S-adenosylmethionine demethylase; (12) adeno-sylhomocysteinase; (13) sulfate reducing pathway; (14) gGCS; (15) GSH synthetase;(16) gGT; (17) CGase. (Reproduced with permission from Penninckx (2002). � ElsevierScience B.V.)


2002). While the expression of YCF1 is clearly under Yap1p control, this isnot the case for BTP1 (Sharma et al., 2002).

3.4. Degradation and Recycling of GSH

Among the GSH degradative enzymes gGT is the only activity that hasbeen detected and characterised in prokaryotes and lower eukaryotes(Penninckx and Elskens, 1993). gGT has been found in many fungiincluding S. cerevisiae (Penninckx et al., 1980; Penninckx and Jaspers, 1985;Mehdi et al., 2001), S. pombe (Wood et al., 2002), C. albicans and N. crassa(Galagan et al., 2003) (Table 1). Interestingly, the ECM38 gene locus codingfor gGT in S. cerevisiae revealed extensive polymorphisms in strains widelyused by yeast researchers (Kumar et al., 2003a).

Figure 4 A model for GSH and GS-X transport and metabolism in S. cerevisiae.GSH and GS-X are transported into the central vacuole by Ycf1p (1) and a V-ATPase-coupled anion uniport system (2 and 3). GSH is further degraded by gGT (4) and CGase(5). Ycf1p is activated by gGT (thick arrow). GSH accumulated in the vacuole exerts afeedback effect on its transport by Ycf1p (dotted arrow). GS–Xs are possibly transportedout of the cell by an ATP-linked system (6). (Reproduced with permission fromPenninckx (2002). � Elsevier Science B.V.)


The yeast gGT enzyme is found as a heterodimer, with 64 kDa and 29kDa subunits (Penninckx and Jaspers, 1985) and the active site on thesmaller subunit (Tate and Meister, 1981). Yeast and human gGT containan N-terminal hydrophobic transmembrane domain, which could accountfor the membrane-bound localization and extracellular exposure of theremaining portion of the protein (Meister and Anderson, 1983). In yeast,gGT is mainly bound to vacuolar membranes (Jaspers and Penninckx, 1984;Mehdi et al., 2001), although the enzyme has been reported in theplasmalemma fraction by other authors (Payne and Payne, 1984).Furthermore, the yeast gGT has a typical type-II topology of integralvacuolar membrane protein, with its short hydrophilic tail extending intothe cytoplasm and with its C terminus bearing the active site inside thevacuolar lumen (Klionsky and Emr, 1989).gGTs catalyze the transfer of the g-glutamyl component of GSH and

other g-glutamyl compounds to amino acids and also the hydrolyticrelease of L-glutamate from GSH, various g-glutamyl compounds, andS-substituted derivatives (Tate and Meister, 1981). In yeast and mammals,L-methionine, L-cysteine and L-glutamine are the most active acceptors intranspeptidation reactions and autotranspeptidation, where g-glutamyl istransferred to GSH, has also been observed (Meister and Anderson, 1983;Penninckx and Jaspers, 1985). Evidence of ‘‘in vivo’’ gGT transpeptidationhas been shown in S. cerevisiae (Jaspers et al., 1985) even though the mainactivity is the hydrolysis of GSH (Mehdi et al., 2001).The physiological role of gGT remains an open-ended question. Initially,

Meister proposed that g-glutamyl cycle and gGT are directly related to renalre-absorption of amino acids (Meister, 1981). In accordance with this, aknockout mutant of mouse deficient in gGT showed the symptoms ofglutathionuria, due to the failure to recover GSH from the renal glomerularfiltrate and hence preventing operation of the g-glutamyl cycle in theepithelial membrane of the renal tubules (Hardings et al., 1997). However,in yeast, deletion of the gGT (CIS2) gene resulted in viable cells (Mehdiet al., 2001; Giaever et al., 2002) characterized by a slower growth rate(Mehdi and Penninckx, 1997; Mehdi et al., 2001) and altered sensitivity tocalcofluor white (Lussier et al., 1997). The participation of gGT in the bulktransport of amino acids has been ruled out in yeast (Jaspers et al., 1985)because it possibly mobilizes GSH reserves as an alternative sulfur (Elskenset al., 1991) and nitrogen source under starvation (Mehdi and Penninckx,1997). CIS2 is a multicopy suppressor of CIK1 and KAR3 nullmutants, genes involved in microtubule assembly (Manning et al., 1997).Therefore, gGT may also play a role in the post-translational processing ofmicrotubules-associated proteins (Mayer and Jurgens, 2002).


In S. cerevisiae, the cellular level of gGT is controlled by the nitrogensource (Penninckx et al., 1980). Biosynthesis of gGT is repressed byammonium, expressed at intermediate levels on glutamine or arginine andat the highest levels with proline, urea, glutamate, or nitrogen starvation.More recently, eight nitrogen and one stress response elements were foundin the promoter region of CIS2 (Springael and Penninckx, 2003). In yeastcells grown on poor nitrogen sources, the GATA zinc-finger transcriptionfactors Nil1p and Gln3p are required for CIS2 expression whileGzf3, another GATA zinc-finger protein and the Gln3p-binding proteinUre2p/GdhCR are negative regulators of the gene induction (Springael andPenninckx, 2003). Interestingly, GSH itself can be a regulator of CIS2expression by binding tightly to and influencing the oligomerisation ofUre2p (Bousset et al., 2001). The degradation of GSH as an exogenoussulfur source was shown to be independent of gGT, which indicates that analternative GSH degradation pathway must exist in S. cerevisiae (Kumaret al., 2003b).

L-Cysteinylglycine dipeptidase (CG) catalyzing the hydrolysis of L-cysteinyl-glycine has been detected in the vacuolar membrane of yeast cells(Jaspers and Penninckx, 1984) and was also found in other microorganismsand mammals (Rankin et al., 1980; Meister and Anderson, 1983; Penninckxand Elskens, 1993; van den Hazel et al., 1996).

3.5. Stabilization of Physiological GSH/GSSG RedoxBalance – Glutathione Reductases

GRs (Table 1) maintain and restore physiological GSH/GSSG balancesunder both stressed and unstressed conditions by reducing GSSG in aNADPH-dependent reaction. It is remarkable that, unlike in S. cerevisiaeand E. coli, GR is absolutely required for the growth of S. pombe (Lee et al.,1997). Stationary phase yeast cells are more resistant to various stressfulconditions than exponentially growing cells (Jamieson, 1992; Lee et al.,1995, 1997; Grant et al., 1996b; Cyrne et al., 2003), which can be attributed,at least in part, to the up-regulated expression of the appropriate GR genes(Grant et al., 1996b).

3.6. GSH – Extracellular Functions

Thiols, including GSH, are often regarded as antioxidant agents but theycan themselves generate free radicals (Halliwell and Gutteridge, 1999).


The generation of GS�thiyl radicals may be harmful for biological

membranes (Riedl et al., 1996) but may facilitate the extracellular degra-dation of polycyclic aromatic hydrocarbons by manganese peroxidasein wood-decaying fungi like Nematoloma frowardii (Sack et al., 1997)and 2-amino-4,6-dinitrotoluene by the white-rot fungus P. chrysosporium(van Aken et al., 2000).Another extracellular enzyme that uses GSH as substrate is the GSH-

dependent ferric reductase, which represents one of the iron acquisitionmechanisms of Histoplasma capsulatum (Timmerman and Woods, 2001).

3.7. GSH in Cell Differentiation and Development

In fungi, transient hyperoxidant states, characterized by redox imbalances,intracellular accumulation of reactive oxygen species (ROS) and activa-tion of antioxidant enzymes, are hypothesized to initiate versatile cell dif-ferentiation processes including germination, conidiation, yeast$myceliumdimorphic conversions and even autolysis (Hansberg and Aguirre, 1990).For example, each morphogenetic step of N. crassa conidiation waspreceded by NAD(P)(H)/NAD(P) and GSH/GSSG redox imbalances(Toledo et al., 1995) and the generation of singlet oxygen was observedduring germination of N. crassa conidia (Lledıas et al., 1999).On the other hand, morphological changes observable in stationary

and autolytic phase submerged Penicillium chrysogenum (Sami et al., 2001a,2003; Pocsi et al., 2003) and A. chrysogenum (Nagy et al., 2003) cultureswere not preceded by GSH/GSSG redox imbalances. In general, GSH/GSSG redox signaling seems to be only one of the factors that may initiatethe genomic expression programs governing morphological transitions infungi.Studies in C. albicans suggest involvement of GSH in the yeast to

mycelium dimorphic switch (Thomas et al., 1991; Manavathu et al., 1996a,1996b). Intracellular GSH levels decreased significantly during thedimorphic switch (Thomas et al., 1991), probably the result of an increasedgGT activity in germ tubes as compared with yeast cells (Manavathu et al.,1996a). These findings led to the hypothesis that GSH levels may signalinitiation of a dimorphic switch (Manavathu et al., 1996a, 1996b). Similarly,GSH levels in Aureobasidium pullulans yeast cells were higher than inmycelia (Fig. 5) even though no detectable difference in the GSH/GSSGredox status was found (Jurgensen et al., 2001).More recently, Lee et al. (2001) used an isogenic homozygous deletant

GSH1 (diploid) S. cerevisiae strain to demonstrate that GSH is essential for


the sporulation of yeast. This finding corroborates previous observationson sake yeast where sporulation was induced by GSH (Kawado et al., 1992;Suizu et al., 1994; Suizu, 1996).

3.8. Is GSH Essential in Unstressed Cells?

A logical approach to determine if GSH is essential in cell physiology andmetabolism is to measure the consequences arising from its deficiency.Partial or total GSH shortage could be obtained by genetically engineered

Figure 5 Similar GSH/GSSG redox balances are found in A. pullulans cells witheither pure yeast (Pure Y) or pure mycelial (Pure M) morphology. cGSH and cGSSG standfor intracellular GSH and GSSG concentrations, respectively. GR activity was alwayshigher in M cells separated from either pure or mixed morphology cultures. Glucose-6-phosphate dehydrogense and gGT activities were similar in cells separated from mixedmorphology cultures independently of the observed cell morphology. Between pure mor-phology cultures, these two enzyme activities were significantly different. Cell morphologywas adjusted simply by varying the inoculum size (Jurgensen et al., 2001).


defects affecting the production of GSH enzymes, or by utilization of GSHbiosynthesis enzyme inhibitors or GSH-depleting drugs. In animal andhuman cells, GSH deficiency has been associated with severe pathologicaldisorders. Deficiency in gGCe or GS activities result in low GSH levels,which are followed by haemolytic anaemia in humans (Larsson andAnderson, 2001).Total block in GSH biosynthesis may be lethal in animals. Disruption

of the mouse Gclc gene coding for gGCS resulted in embryonic lethalityprior to embryonic day 13 (E13) in homozygous mutants (Dalton et al.,2000). A partial GSH deficiency of genetic origin, or consequence of drugintake (e.g. alcohol or acetaminophen), generally leads to impaired liverand kidney functions (Bondy, 1992; Thomas, 1993) and a state of reducedresistance to various infectious agents or stress (De Rosa et al., 2000).HIV-infected subjects with GSH deficiency were found to have a shorter lifespan, by 2 to 3 years, when compared with subjects without GSH deficiency(Herzenberg et al., 1997).GSH dispensability in microorganisms was also addressed in bacteria

and fungi. gGCS (gshA� or gsh1�) and GS (gshB�) deficient E. coli mutantsshow normal growth but are 3 to 10 times more sensitive to chemicalssuch as sulfydryl oxidizing agents diazenes, oxo-aldehydes, heavy metals,pesticides and certain food additives (Apontoweil and Berends, 1975; Fuchsand Warner, 1975). A double mutant gshA� gshB� had the same phenotypeas the gshB� parent, which confirmed that GSH is not a NPT required byE. coli under laboratory growth conditions (Fuchs et al., 1983).Yeast GSH-deficient mutants were first reported by Eckardt

and colleagues (Kistler et al., 1986) as N0-nitro-N-nitrosoguanidine(NNG)-resistant clones. The mutant selection was based on the fact thatNNG needs to be activated by GSH to exert a mutagenic effect (Mohn et al.,1983). Thus, all isolates were GSH1 mutants with a residual gGCS activityof about 5–10% of the parental activity and had a residual GSH contentvarying from 2 to 8% of the wild type. GSH deficiency in these strains wascorrelated with an extension of the lag phase of growth and a decreasein growth rate when cultivated on laboratory media (Kistler et al., 1986).Prototrophic GSH1 derivatives obtained by Elskens et al. (1991) displayedsimilar characteristics in addition to being hypersensitive to heavy metals,oxo-aldehydes, diazenes and dithiocarbamates (Elskens and Penninckx,1995a, 1997). GSH1 mutants are also hypersensitive to ROS includingH2O2, superoxide anion and lipid hydroperoxides (Stephen and Jamieson,1996).�GSH1 ‘‘knockout’’ mutants that lack a functional GSH1 were unable

to synthesize glutathione and required the addition of GSH to grow on


minimal medium (Grant et al., 1996a). However, ‘‘delayed growth stasis’’was reported for �GSH1 mutants (Sharma et al., 2000), where uponwithdrawal of GSH from the medium, cells grew for seven or eightgenerations at wild-type growth rates before entering growth stasis.A similar phenomenon was observed for the anaerobic protozoonEntamoeba histolytica, where, in the absence of exogenous GSH, theparasite was able to grow for at least five to seven generations (Ondarzaet al., 1999). Furthermore, growth of a �gsh1 mutant can be restored withdithiothreitol (DTT), indicating that GSH is required as a reductant duringboth aerobic or anaerobic conditions (Spector et al., 2001).This alternative reductant function is apparently (i) not linked to

ribonucleotide reduction because the �GSH1 strain arrests in G1 phasewith predominantly unbudded cells after GSH withdrawal and (ii) notlinked to a defect in sulfate assimilation because neither methionine norcysteine can rescue the growth defect of �GSH1. Thioredoxin (TRX1and TRX2) may provide reducing equivalents for ribonucleotides andsulfate reduction in �GSH cells growing in the presence of DTT (Spectoret al., 2001).However, it remains to be determined if, in GSH deficient cells,

thioredoxins (Trxs) are overproduced to compensate for GSH shortage. Inyeast, GSH is not essential for bioreduction thus suggesting overlappingfunction with thioredoxins. �TRX1 and �TRX2 mutants showed wild-typegrowth rates and cell morphology, whereas the double mutant showeddecreased rates of DNA replication with a corresponding increasein generation time, as well as being auxotrophic for methionine (Muller,1991). The triple mutant �TRX1 �TRX2 �GLR1, which in addition toTRX genes also lacked GR, was non-viable under aerobic conditions andgrew poorly anaerobically (Muller, 1996). Finally, yeast contains two genes,GRX1 and GRX2, encoding cytoplasmic glutaredoxins (Grxs) (Table 1).The quadruple mutant �TRX1 �TRX2 �GRX1 �GRX2 was non-viablewhereas one single Grx or Trx was necessary and sufficient for growth inS. cerevisiae (Draculic et al., 2000; Trotter and Grant, 2003). This findingclearly demonstrates that there is a functional link between the GSH/Grxand Trx systems in yeast.Contrasting with GR, which is not essential in budding yeast, thioredoxin

reductase (Trr1p) is absolutely required for normal growth in S. cerevisiae.Examination of the redox state of Trxs and Grxs in GLR1 and TRR1mutants shows that Trxs are maintained independently of the GSH/Grxsystem (Trotter and Grant, 2003).GSH via its reducing power might be involved in other basic functions of

the yeast cell, for example detoxification of cryptic harmful intermediates


generated during unstressed cellular metabolism (Penninckx et al., 1983;Grant et al., 1997) or maintenance of the mitotic apparatus and/or othermembrane systems including perhaps mitochondria (Penninckx andElskens, 1993). Although GSH/GSSG does not mediate disulfidebond formation in the endoplasmic reticulum (Tu et al., 2000), recentdata have emphasized the essential role that GSH could have in thematuration of cytosolic iron–sulfur proteins (Sipos et al., 2002), theregulation of the 20 S proteasome (Demas et al., 2003) at least in the yeast ofS. cerevisiae, and the stabilization of yeast vacuolar ATPase (Oluwatosinand Kane, 1997).Although the molecular background of the absolute need of yeast cells

for GSH is not known, the essential function of GSH in the maintenanceof cellular growth requires very low amounts of the tripeptide, abouttwo orders of magnitude less than that observable in unstressed wild-type cells (Lee et al., 2001). This finding was highly supported bysuppressors of the GSH auxotrophy of �GSH1 mutants where specificmutations in PRO2, the gene coding for g-glutamyl phosphate reductase,the second enzyme in the biosynthesis of proline, made the biosynthesis oftrace amounts of GSH possible (Spector et al., 2001). These data indicatethat the indispensable physiological function of GSH cannot be relatedto GSH/GSSG buffering that assumes the presence of the reduced thiol inhigh concentrations.Information concerning an essential role for GSH in other fungi and

eukaryotes is poor. The first gsh1(gcs1þ ) and gsh2(gsh2þ ) mutants wereobtained for the fission yeast S. pombe on the basis of a phenotype ofhypersensitivity to cadmium salts (Mutoh and Hayashi, 1988). Thesemutants were deficient in cadystin (phytochelatins), cadmium-bindingpeptides involved in detoxification of Cd2þ . Other gcs1þ and gsh2þ mutantstrains were obtained by the NNG activation procedure described above(Glaeser et al., 1991). In general, GSH deficient mutants have a normalgrowth phenotype but have lost the ability to neutralize Cd2þ and aremore sensitive to Cu2þ , Zn2þ and Pb2þ ions (Mutoh and Hayashi, 1988;Coblenz and Wolf, 1994). Disruption of gcs1þ and/or gsh2þ led to GSHauxotrophy (Chaudhuri et al., 1997; Kaur et al., 1997).Physiological phenomena related to changes in GSH/GSSG redox

ratios may be connected to increasing GSSG concentrations and not todeclining GSH levels. For example, when the pgr1þ gene, coding forGR (Table 1) in S. pombe was disrupted, the haploid spores were notviable (Lee et al., 1997). This situation is unlikely in S. cerevisiae andE. coli where disruptants for the gene coding for GR showed wild-typegrowth rates on laboratory media (Muller, 1996). The growth defect of the


S. pombe �pgr1þ strain could not be complemented by the addition ofGSH to the medium suggesting that lowered amounts of GSH are notthe cause of the growth arrest. Possibly, the accumulation of oxidizedGSSG in the disruptant (Lee et al., 1997) followed by uncontrolled thiol-disulfide exchange reactions, was the cause of growth inhibition. Howeverthis was not observed in S. cerevisiae where despite an exceptionallyhigh GSSG/GSH ratio, the �glr1 mutant grows with a normal cell cycle(Muller, 1996).What could we conclude at the moment for the essentiality of GSH

in unstressed fungi? Obviously the essential role that GSH could playin yeasts and filamentous fungi in unstressed situation is still notclear. However, some reply to this question, in particular the role ofGSH in the integrity of membranes and mitochondria (Fig. 1), couldpossibly be found in studies on the evolution of the metabolism of thispeptide. For example, it is noteworthy that E. histolytica, the mitochondriaprotist where GSH is absent, derives from an ancestor that once hadmitochondria that were lost (Clark and Roger, 1995). If so, such lossincluded GSH metabolism, as well as other metabolic processes associatedwith mitochondria (Fahey, 2001), and would identify GSH as a metaboliteessential for mitochondrial function. It is also interesting to remember thatE. histolytica never undergo meiosis and do not produce mitotic spindlesduring cell division. This suggests that GSH might participate incytoskeletal functions (Fig. 1).

4. GLUTATHIONE IN STRESS RESPONSES

4.1. Oxidative Stress

‘‘Oxidative stress’’ and related terms like ‘‘adaptation’’ and ‘‘cell injuries’’are often used in fungal physiology but they are quite loosely defined in manycases (Kreiner et al., 2000). By definition, oxidative stress is a ‘‘disturbancein the pro-oxidant-antioxidant balance in favor of the former, leading topotential damage’’ (Sies, 1991) and, in principle, can result from thediminution of antioxidant reserves or the increased production of ROS(Halliwell and Gutteridge, 1999). Among the consequences of oxidativestress, cells may up-regulate antioxidant defence systems to restore theiroxidant/antioxidant balance (adaptation), may suffer a significant loss oftheir functional biomolecules (cell injuries) and eventually die by eithernecrosis or apoptosis (Halliwell and Gutteridge, 1999).


ROS are reduced forms of atmospheric oxygen (O2) that areproduced and accumulated within living cells as the result of the transferof one, two or three electrons to form superoxide (O2

.�), hydrogenperoxide (H2O2) and hydroxyl radicals (.OH), respectively. Theseside products of numerous essential cellular reactions are capable ofthe oxidation of cellular components that, if uncontrolled, lead to theoxidative destruction of the cell (Mittler, 2002). There are manysources of ROS in fungal cells, including different molecular engines,cellular factories and machines (Gasch et al., 2000, 2001; Causton et al.,2001).Antioxidant defence systems and mechanism of adaptation to oxidative

stress are usually induced with ROS-generating chemicals, e.g. H2O2,paraquat, menadione, tert-butyl hydroperoxides, and diamide (azodicar-boxylic acid bis [dimethylamide]). These compounds interfere with a rangeof cellular components and cause accumulation of ROS and/or alter theGSH/GSSG ratio (Halliwell and Gutteridge, 1999).Artificial induction of oxidative stress with chemicals has several

disadvantages: first, these agents affect the concentrations of multipletypes of ROS resulting in a severe redox imbalance (Fortuniak et al., 1996)unless the quantities of the reactants have been carefully optimized(Emri et al., 1997a, 1999a) and, second, the physiological informationgained from artificially induced oxidative stress cannot be applied directlyto predict or explain the stress responses when fungi are exposed to natu-rally occurring oxidative stress conditions, e.g. to increased O2 gassingin fermenters (Kreiner et al., 2000; Kreiner et al., 2002, 2003; Bai et al.,2003).Glutathione is an important antioxidant molecule, which reacts non-

enzymatically with a series of ROS including .OH, HOCl, RO. , RO2. , 1O2,

as well as with many nitrogen and carbon containing radicals through theformation of thiyl (GS. ) radicals (Halliwell and Gutteridge, 1999). GS.

species may generate O2.� which can be neutralized by superoxide dismutase

(SOD)/catalase enzymes or react with other cellular thiols giving rise tomixed disulfides (Halliwell and Gutteridge, 1999). This process mayinactivate enzymes but, on the other hand, may prevent the irreversible,random formation of protein-S–S-protein disulfides. Hence, proteinS-thiolation is regarded as an important antioxidative process, which canbe tightly regulated (Grant et al., 1999; Shenton et al., 2002). It isworth noting that GSH also reacts with the lipid peroxidation metabo-lite 4-hydroxy-2-nonenal (Wonisch et al., 1997), and plays a role in theinitial resistance against malondialdehyde, another highly toxic lipidperoxidation product (Gupta et al., 1996; Turton et al., 1997).


GSH synthetases (gGCS and GS), NADPH-dependent GSH-regeneratingreductase (GR), glutathione S-transferase (GST) along with peroxide-eliminating glutathione peroxidase (GPx) and glutaredoxins (Grxs) aredirectly involved in the elimination of oxidative compounds in yeast andother fungi (Jamieson, 1998; Moradas-Ferreira and Costa, 2000; Costa andMoradas-Ferreira, 2001; Collinson et al., 2002).Under normal growth conditions the accumulation of ROS in cells is low

and does not cause major problems, except in aging cultures where theaccumulation of ROS is presumed (Section 6). Whereas accumulation ofreactive oxygen radicals is low in normal aerobic conditions, environmentalchanges that cause osmotic- or heat stress, UV mediated DNA damageor nutrient deprivation strongly enhance the accumulation of ROS(Godon et al., 1998; Gasch et al., 2000; Causton et al., 2001; Goldmanet al., 2002). Excessive accumulation of these oxidative free radicals posesa real threat to living cells (Osiewacz, 2002). Cell damage results fromoxidative processes such as membrane lipid peroxidations, proteinoxidation, enzyme inhibition and DNA/RNA damage. Eukaryotic cellsposses metabolic systems that remove cell-damaging activated oxygen formsby induction of enzymes such as SOD, catalase and GSH/Grx/Trxmetabolism that require ROS as substrates to produce water, oxygen orcouple electron transfer to NADP/NADPH (Mittler and Tel-Or, 1991;Navarro et al., 1996; Kawasaki et al., 1997; Navarro and Aguirre, 1998;Noventa-Jordao et al., 1999; Kawasaki and Aguirre, 2001; Mittler andBerkowitz, 2001; Mittler, 2002).Figure 6 describes GSH (Fig. 6A), Grx/Trx (Fig. 6B) and SOD/catalase

(Fig. 6C) ROS elimination mechanisms found in fungi. Indicated are theyeast gene names whose amino acid sequence was used to identify andcharacterize homologs in three filamentous fungi with fully sequencedgenomes, A. nidulans, N. crassa and Magnaporthe grisea. Table 2 showsdetails of the homology-based matches.Genes associated with the production of GSH and its oxidation/reduction

reactions are highly conserved amongst the three fungal species (Table 2).In yeast, two GPxs are reported (GPX1 and GPX2), but in the filamentousfungi we could only identify one, GPX2-like gene.In terms of physiology, fungal GPxs are involved in the detoxification of

H2O2 (Yamada et al., 1999) as well as lipid and phospholipidhydroperoxides (Inoue et al., 1995; Emri et al., 1997a; Evans et al., 1998;Inoue and Kimura, 1998; Inoue et al., 1999; Gil-ad et al., 2000; Avery andAvery, 2001). The S. pombe GPx has been shown to be a selenium-freeenzyme (Yamada et al., 1999) while the S. cerevisiae GPxs have beenidentified as functional phospholipid hydroperoxide glutathione peroxidases


Figure 6 Elimination of reactive oxygen species (ROS) by the glutathione–glutaredoxin–thioredoxin system in fungi. Part A. The glutathione cycle in fungi.


(Avery and Avery, 2001). In accordance with their primary role inthe detoxification of lipid hydroperoxides, GPx of Hansenula mrakii isbound to the membrane of mitochondria and cytoplasm (Inoue et al., 1995;Inoue and Kimura, 1998) and the Candida boidinii CbPmp20 proteinpossessing GPx activity is associated with the inner side of the peroxisomalmembrane (Horiguchi et al., 2001).Amongst the most important factors that regulate cellular redox

homeostasis are the GSH/Grx and Trx systems (Grant, 2001) (Section3.8). These small oxidoreductases participate in physiological processes suchas repair of oxidized proteins (Draculic et al., 2000; Trotter and Grant,2003). Along with GR, Trxs have been shown to contribute to themaintenance of high intracellular GSH/GSSG ratios (Muller, 1996; Garridoand Grant, 2002), even though redox regulation of Trxs was independent ofthe GSH/GSSG system (Trotter and Grant, 2003). Although functionalredundancy between Grx and Trx isoforms exists, both systems may befunctional during normal growth and under different stress conditions(Grant, 2001).Saccharomyces cerevisiae possesses two GRX genes (GRX1 and GRX2)

(Section 3.8) and three members of a family of Grx-related proteins encodedby GRX3-5 (Luikenhuis et al., 1998; Rodrıguez-Manzaneque et al., 1999).While Grx1p and Grx2p function through a common dithiol catalytic

Glutathione (GSH) is synthesized from amino acids through two ATP-dependentreactions, g-glutamylcysteine synthetase (GSH1) and glutathione synthetase (GSH2),which catalyze synthesis of the dipeptide g-L-glutamyl-L-cysteine from glutamic acid andcysteine and the tripeptide, g-L-glutamyl-L-cysteinyl-glycine (GSH), respectively. GSH isoxidized to GSSG by reacting with ROS or in combination with reactions catalyzed byglutathione peroxidases (GPX1) or glutathione S-transferase (GTT1) and is regeneratedby glutathione reductase (GLR1) in NADPH-dependent reaction. Degradation of GSHremains unknown in yeast and fungi. Part B. The glutathione–glutaredoxin–thioredoxinsystem in fungi. The oxidized disulfide form of thioredoxin (TRX2) is reduced directly byNADPH and thioredoxin reductase (TRR1). Oxidized glutaredoxin, glutathione-dependent oxidoreductase (GRX1 and GRX2) is reduced by GSH and oxidized GSSGis reduced by NADP-dependent glutathione reductase (GLR1). Thioredoxins act ashydrogen donors for PAPS reductase (MET16) and thioredoxin peroxidases (TSA1,AHP1 and YDR453/YBL064c) and both thioredoxins and glutaredoxins act ashydrogen donors for ribonucleotide reductase in yeast and fungi. Part C. Eliminationof ROS in fungi. O2

.� radicals are converted into H2O2 by superoxide dismutase (SOD1and SOD2), which is converted further into water and oxygen by catalase (CAT1 andCTT1). Reactive oxygen and hydrogen peroxide is also eliminated in reacting directlywith GSH producing GSSG. All the shown protein names are based on yeast gene names,and the presence or absence of homologues and paralogues in M. grisea (Mg), A. nidulans(An) and N. crassa (Nc) are indicated by the bits score of amino acid sequencecomparisons.


Table 2 Glutathione, glutaredoxin thioredoxin oxidation systems and direct elimination of reactive oxygen intermediate in fungi.

Function Yeast gene M. grisea A. nidulans N. crassa

locus e-value bits locus e-value bits locus e-value bits

GSH-GSSG PATHWAY

GSH transporter HGT1 MG10200 E�149 525 1.129 s10 E�158 556 NCU03171 E�148 520g-L-glutamyl-L-cysteinesynthase

GSH1 MG07317 E�120 427 1.51 s3 E�107 375 NCU01157 E�115 411

Glutathione synthetase GSH2 MG06454 4E�67 251 1.241 s82 1E�39 161NCU06191 1E�72 2691.37 s2 2E�25 114

Glutathione reductase GLR1 MG03571 3E�34 141 1.14 s1 E�116 230 NCU03339 E�142 5011.238 s79 3E�18 90 NCU02407 1E�35 146

Glutathione peroxidase GPX1MG07460

4E�30 125 1.51 s3 3E�30 127NCU09534

8E�34 138GPX2 4E�50 192 1.51.s3 3E�50 194 4E�52 198

Glutathione S-transferase GTT1 MG05677 4E�31 130 1.8 s1 5E�30 92 NCU05706 6E�28 119GTT2 no hit found no hit found no hit found

TRX/GRX PATHWAY

Thioredoxin TRX2 MG04236 4E�23 100 1.5 s1 2E�25 109 NCU06556 5E�19 871.157 s12 7E�20 92 NCU05731 4E�17 81

Glutaredoxin, glutathione- GRX1MG05447

8E�17 80 1.68 s4 5E�11 54NCU01219

5E�15 74dependent oxidoreductase GRX2 7E�19 88 1.192 s34 4E�10 59 2E�19 90

Thioredoxin reductase TRR1 MG01284 E�105 375 1.61 s4 E�121 432 NCU08352 E�124 439PAPS reductase MET16 MG03662 4E�76 280 1.80 s5 5E�83 304 NCU02005 1E�77 285Thioredoxin peroxidases TSA1 MG08256 9E�15 75 1.65 s4 3E�12 58 NCU06031 3E�15 77

AHP1 MG02710 3E�20 94 1.160 s13 1E�15 60 NCU03151 3E�19 901.139 s11 5E�12 52 NCU06880 2E�16 80

YDR453 3E�15 77 1.65 s4 4E�11 57 1E�15 78MG08256 1.65 s4 3E�65 171 NCU06031

YBL064c 1E�65 245 1.26 s2 5E�59 124 9E�66 245

28

ISTVAN

POCSI,ROLFA.PRADEAND

MIC

HELJ.PENNIN

CKX

ELIMINATION OF REACTIVE OXYGEN IN INTERMEDIATES (ROI)

Cu, Zn superoxidedismutase

SOD1 MG02625 7E�50 191 1.5 s1 5E�56 213 NCU02133 2E�61 229

Mn-containing SOD2 MG00212 1E�63 238 1.96 s6 7E�56 154 NCU09560 8E�66 245superoxide dismutase MG07697 1E�41 165 1.13 s1 8E�46 180 NCU01213 2E�45 177

Catalase A MG06442 4E�75 277 1.101 s7 E�160 563 NCU05169 E�122 434MG10061 2E�71 265 1.158 s13 1E�73 175 NCU08791 4E�77 284

CAT1 1.157 s12 3E�59 139 NCU00355 6E�76 2801.172 s16 9E�57 187

Catalase T MG10061 4E�69 258 1.101 s7 E�124 442 NCU05169 1E�94 342MG06442 1E�63 239 1.158 s13 1E�75 180 NCU08791 5E�75 277

CTT1 1.172 s16 5E�44 153 NCU00355 4E�63 2381.157 s12 2E�39 112

OTHERS

GS-X pump YCF1 MG01674 0Eþ 00 1268 1.131 s10 0Eþ 00 800 NCU09012 0Eþ 00 1407MDR FLR1 MG01511 2E�79 292 1.161 s13 8E�63 160 NCU05580 6E�74 273HSP70 SSA1 MG06958 0Eþ 00 893 1.88 s6 0Eþ 00 894 NCU09602 0Eþ 00 907

e- and bits-values are from BLASTp comparisons of the yeast gene (query) with fungal peptides refering to probability and homology, respectively.

GLUTATHIO

NEMETABOLISM

INFUNGI

29

mechanism, Grx3p, Grx4p and Grx5p contain only one cysteine residue attheir active sites (Rodrıguez-Manzaneque et al., 1999). Grx1p appears to beinvolved in the elimination of superoxides, while Grx2p accounts for themajority of Grx activity under unstressed conditions and hydrogen peroxidestress. Grx5p appears to be involved in protection against oxidative proteindamage triggered by superoxide or peroxide (Luikenhuis et al., 1998;Rodrıguez-Manzaneque et al., 1999).S. cerevisiae GRX1 and GRX2 genes had significant matches in

filamentous fungi (Fig. 6B) but p- and bits-scores were much lowerthan the ones reported for any of the components of the GSHpathway (Fig. 6A). In addition, it appears that filamentous fungi consi-stently possess only one GRX gene. All other components of the Grx/Trxsystem appear to be highly conserved in yeast and filamentous fungi(Table 2).A more recent paper by Collinson et al. (2002) reported on the peroxidase

activity of S. cerevisiae Grx1 and Grx2, which might result in theformation of alcohols in cells exposed to hydroperoxides. The alcohols weretransported into the vacuoles after GSH S-conjugation via the ABC proteinYcf1p GS-X pumps (Collinson et al., 2002). This finding is consistent withthe presence of oxidative stress-inducible GSTs in S. pombe (Cho et al.,2002; Veal et al., 2002). GSTs are able to eliminate many toxic secondaryproducts of membrane oxidation, e.g. 4-hydroxy-2-nonenal, cholesterola-oxide, but some of them may play a more direct role, as peroxidases, inthe elimination of primary organic hydroperoxides (Veal et al., 2002).Nevertheless, the oxidative stress response of GSTs observed in yeast is notuniversal among fungi. In P. chrysogenum, the specific GST activity wasonly elevated in the presence of the superoxide-generating agent menadione(Emri et al., 1997a, 1999a), the activation and detoxification of which relyon GSH S-conjugation ( _ZZa�dzinski et al., 1998).In yeast, the elimination of ROS is driven by modifying enzymes; a

Mn- and a Cu,Zn-containing superoxide dismutase, SOD1 and SOD2.Both of these enzymes have homologues in the filamentous fungi.Moreover, filamentous fungi appear to encode an additional Mn-containingsuperoxide dismutase (Table 2). Similarly, yeast encode two similarcatalases, CAT1, CTT1 and the filamentous fungi appear to have several:two, three and four in M. grisea, N. crassa and A. nidulans, respectively(Table 2).The response to remove ROS and protect cells against oxidative

damage is regulated through one or more signal transduction cascades thatactivate the expression of a series of genes involved in general stressresponse and a specific response such as GSH metabolism, superoxide


dismutase and catalase production (Aguirre, 1993; Carter et al., 1994;Krems et al., 1995; Blomberg, 1997; Cuppers et al., 1997; Emri et al., 1997a;Noventa-Jordao et al., 1999; Dumond et al., 2000; Stochaj et al.,2000; Jurgensen et al., 2001; Kawasaki and Aguirre, 2001; Cho et al.,2002; Collinson et al., 2002; Garrido and Grant, 2002; Herrero andRos, 2002; Longo and Fabrizio, 2002; Pekker et al., 2002; Westwater et al.,2002; Yoshimoto et al., 2002). Some of the components that have beenidentified in the regulation of oxidative response are SKN7 and HSF1 thatform a two-component signaling system (Krems et al., 1995). Loss of SKN7function exacerbates H2O2 sensitivity and heat shock proteins requireSKN7 for their induction by oxidative stress but not heat shock (Raittet al., 2000).Transcription factors known to participate in the oxidative stress

response are Msn2p/4p and Yap1p (similar to the human AP-1). Yap1pactivates genes required for the response to oxidative stress (Moye-Rowley,2002). Under normal conditions, Yap1p is cytoplasmic and inactive, butcan be activated by nuclear translocation if cells are in an oxidativeenvironment. Yap1p is targeted by Crm1p (beta-karyopherin-likenuclear exporter) because it is constitutively nuclear in a CRM1 mutant(Wemmie et al., 1997; Kuge et al., 1998). Moreover, Crm1p binds toa nuclear export sequence (NES) in Yap1p in the presence of RanGTP(Yan et al., 1998). Finally, Yap1p interaction with Crm1p is inhibited byoxidation, indicating that localization is controlled by nuclear export, andthe state of oxidation interferes with access of Yap1p NES to Crm1pdirectly (Yan et al., 1998).Under oxidative stress, GRX and TRX genes were up-regulated in a Yap1p

(GSH/Grx system) or Yap1pþ Skn7p/Pos9p (Trx system) dependentmanner (Lee et al., 1999; Dumond et al., 2000; Grant, 2001).Interestingly, inactivation of Yap1p appears to be Trx-dependent andnot influenced by Grxs (Izawa et al., 1999; Kuge et al., 2001; Moye-Rowley, 2002).Atf1p is a transcription factor, similar to Pap1p (Yap1p), and

the last member of the SAPK phosphorylation-signaling cascadethat regulates oxidative stress in S. pombe (Moye-Rowley, 2002).Phosphorylated Spc1p kinase migrates into the nucleus where it isneeded to activate Atf1p. Atf1p is homologous to the mammalianfactor ATF-2, which is regulated via SAPK/JNK and p38. In S. pombe,Atf1p regulates catalase (Ctt1) whose induction is observed afterUV irradiation, osmotic stress or heat shock. The expression of thegpd1 (glycerol 3-phosphate dehydrogenase) gene is also up-regulatedby Atf1p under high salinity stress (Shiozaki and Russell, 1996).


Moreover, Ish1p, a nuclear envelope protein regulated by Atf1p-mediatedoxidative stress is also involved in carbon and nitrogen starvation(Taricani et al., 2002). Thus, Atf1p is a clear example suggesting thateven though different kinds of stress are physiologically dissimilar, theymay invariably create a set of overlapping gene products due to thecommon regulatory elements of different stress responses. Finally, Atf1phas also been shown to affect the meiosis-regulating gene ste11 (Takedaet al., 1995).It is worth noting that the transcriptional regulation of GSH-dependent

antioxidant enzymes is often multifactorial and is highly dependent onthe composition of the culture media. For example, the up-regulation ofGSH1 observable under H2O2 stress in S. cerevisiae was amplified in thepresence of L-Glu, L-Gln and L-Lys (Stephen and Jamieson, 1997).Moreover, the increased GSH content of S. cerevisiae cells grownaerobically (in comparison to anaerobic conditions) was attributed to anincreased cystathionine b-synthase activity and, as a consequence, to anelevated intracellular L-Cys concentration instead of any transcriptionalregulation of gGCS (Ohmori et al., 1999).Another interesting aspect of the transcriptional regulation of antioxidant

enzymes came from the analysis of the genomic expression programsof S. cerevisiae initiated under different kinds of environmental stress(Gasch et al., 2000). Interestingly, isoenzymes of GPx, GST, Grx and Trxwere differentially regulated under environmental stress conditionsindicating that they might fulfil very different physiological functions,and only some of them were optimized to contribute to the generalenvironmental stress response. Other isoenzymes were responsive to morespecialized, e.g. oxidative, stress signals or were produced constitutively(Gasch et al., 2000).Fungi may face oxidative stress much more often than we might

think. For example, plant and human pathogenic fungi have to adapt tohigh ROS levels produced by the host organisms in their infection stressresponses (Jamieson et al., 1996; Mayer et al., 2001). Xenobiotics includingantifungal agents (Elskens and Penninckx, 1997; Machida et al., 1999;Nakayama et al., 2002), anticancer anthraquinones (Buschini et al., 2003)and penicillin side chain precursors (Section 5.3) can also trigger oxidativestress by increasing ROS production and/or depleting the intracellular GSHpool. The development of oxidative stress can be a consequence of, orinherently coupled to, other kinds of stress, e.g. heat shock (Section 4.2),desiccation (Section 4.3), ionising radiation (Jaruga et al., 1995), as well asfreeze-thaw (Park et al., 1998) and metabolic (Aminova and Trotsenko,1998; Koerkamp et al., 2002) stress.


Industrial fungi also have to cope with an increased ROS productionwhen the technology itself requires an elevated dissolved oxygen tension(Moresi et al., 1991; Rosenberg et al., 1992; Henriksen et al., 1997).Interestingly, increased intracellular ROS levels might be beneficial for theproduction of secondary metabolites (Jayashree and Subramanyam, 2000)and hydrolytic enzymes (Sahoo et al., 2003). It is worth noting that white-rot fungi themselves generate substantial quantities of peroxide to reach asatisfactory degradation of lignin (Leonowicz et al., 2001). Last but notleast, ROS concentrations may elevate endogenously in fungal cellsdepending on the age of the culture.

4.2. Heat and Osmotic Shock

Aerobic heat shock increases the frequency of mutations and interchromo-somal DNA recombination (Davidson and Schiestl, 2001), and damagesmitochondrial DNA (Sugiyama et al., 2000a) and membranes (Davidsonand Schiestl, 2001). Cytotoxic and genotoxic effects of heat shock canbe attributed to the increased intracellular oxidation level, triggered byintensified respiration (Sugiyama et al., 2000b). The toxicity of heat istherefore in part the result of oxidative stress (Davidson and Schiestl, 2001)and, as a consequence, heat shock produces cross-adaptation to otherforms of oxidative stress including lipid hydroperoxide treatment (Evanset al., 1998) and ionising radiation (Jaruga et al., 1995). Not surprisingly,components of the antioxidative defence, including GSH (Sugiyamaet al., 2000a), the GSH synthesising enzymes gGCS and GS (Sugiyamaet al., 2000a, 2000b), GR (Lee et al., 1997), GST (Choi et al., 1998), GR(Grant et al., 2000) and Trx peroxidase (Lee and Park, 1998) havebeen reported to confer resistance against heat shock. Similar to theregulation of oxidative stress response, the up-regulation of GSH bio-synthesis under heat shock is Yap1p-dependent in S. cerevisiae (Sugiyamaet al., 2000b).Much less is known about the involvement of GSH-dependent systems

in the protection against osmotic shock but both Grx1 and Grx2(Grant et al., 2000) as well as glyoxalase I (Inoue et al., 1998b) wereinduced in S. cerevisiae cells exposed to high concentrations of sodiumchloride. In both cases, the induction was controlled by the HOG1 MAP-kinase pathway (Inoue et al., 1998b, Grant et al., 2000). Moreover,expression of the S. pombe pgr1 (GR) was also up regulated by osmoticshock (Lee et al., 1997).


4.3. Desiccation

The well-known tolerance of lichens to drought, heat and cold is due toa combination of cellular protective and repair mechanisms (Honegger,1998). For example, both the formation of GSSG during desiccation andits reduction upon hydration are important elements of the adaptationmechanism of lichens to drought (Kranner and Grill, 1996, 1997). GSSGreacts with thiol groups of proteins forming protein–SSG mixed disulfides,resulting in protection against desiccation-induced oxidative injuries such asthe irreversible random formation of intramolecular disulfide bridges or theuncontrolled oxidation of thiols to sulfonic acids (Kranner and Grill, 1996,1997). During hydration, the reduction of the oxidized GSSG pool isnecessary because GSSG is needed to regenerate protein thiols. Kranner(2002) found a correlation between the ability of lichens to regeneratetheir GSH pools during hydration and their long-term desiccationtolerance. GSH and GSH-dependent enzymes have also been shown tobe involved in antioxidative defence in mosses (Dhindsa, 1991; Takacset al., 2001), seaweeds (Collen and Davison, 1999; Burritt et al., 2002)and plants (Navari-Izzo et al., 1997; Kranner et al., 2002) under desiccationstress.

4.4. High Cell Density Cultures

Increasing cell density may result in increasing oxidative stress inmicroorganisms as a consequence of metabolic stress triggered by the fastdepletion of nutrients in the culture media (Pinto et al., 2003). In bacteria,the oxidative stress enzymes catalase and SOD have been shown to beregulated by cell density (Crockford et al., 1995; Wood and Sørensen, 2001).Similar effects can be predicted for fungal cultures especially when highdissolved oxygen tension is applied to increase the biomass and improveheterologous protein expression in yeast (Jahic et al., 2002; Lee et al., 2003).In addition, the application of methylotrophic yeasts like Pichia pastorisand Hansenula polymorpha is spreading widely for the expression ofrecombinant proteins (Cox et al., 2000; Hellwig et al., 2001; Shiloach et al.,2003) because these fungi have the ability to grow to a very high celldensity and the expression of heterologous proteins can be controlledrelatively easily using the alcohol oxidase gene promoter. Methanol isused as the inducer and also as the carbon and energy source inthese fermentations (Zhang et al., 2002). Elimination of formaldehydeand H2O2, the products of alcohol oxidase, is a prerequisite for use of


methanol as a carbon source. The oxidation of formaldehyde requiresGSH and is catalyzed by GSH-dependent formaldehyde dehydrogenases(Shen et al., 1998; Baerends et al., 2002) (Section 5.1). Further studiesare needed to characterize the molecular background of the adaptationto high cell density stress in fungal cultures.

4.5. Heavy Metal Stress

Metal pollutants generated by a wide range of industrial activities representa threat to natural populations of different kinds of fungi includingyeasts (Avery, 2001) and ectomycorrhiza-building basidiomycetes(Hartley et al., 1997). Metal ions assimilated by cells generate reactiveoxygen species (ROS) directly (via redox-active metals, like Cu, Fe, Cr, V)or indirectly by displacing redox-active metals from cellular binding sites(Cd, Hg, Ni, Pb) (Avery, 2001).Lipid peroxidation, protein and DNA oxidation contribute to the

observed symptoms of metal toxicity, e.g. reduced growth rates(Avery, 2001). Intracellular GSH and other NPTs (phytochelatins) andmetallothioneins hinder progression of heavy metal-initiated cell injuries bychelating and sequestering the metal ions themselves and/or by eliminatingROS (Perego and Howell, 1997; Avery, 2001; Cobbett and Goldsbrough,2002).Regarding the central role that GSH plays in conferring protection

on yeast cells against both heavy metal and oxidative stress, it is notsurprising that the regulations of sulfur assimilation and GSH productionare inherently coupled and up-regulated in S. cerevisiae on Cd2þ -exposure(Dormer et al., 2000; Vido et al., 2001; Momose and Iwahashi, 2001;Jamieson, 2002). In addition to the oxidative stress-responsive transcrip-tion factor Yap1p (Wu and Moye-Rowley, 1994), expression of GSH1 isalso positively controlled by transcription factors Met4p, Met31p andMet32p, regulators of sulfate assimilation/methionine biosynthesis genes(Dormer et al., 2000).More recent proteome analysis by Fauchon et al. (2002) indicated that

Met4p plays an essential role in the global sulfur-sparing response ofS. cerevisiae to Cd2þ -stress when the production of abundant sulfur-richproteins is reduced and the expression of sulfur-depleted isoenzymes isfavoured to facilitate the recycling of protein–sulfur into the GSHbiosynthetic pathway. This stress response may have evolved in geneticallydistant fungal species living on plants that tend to accumulate highconcentrations of heavy metal ions, including Cd2þ , to gain defence against


herbivores and pathogenic micro-organisms (Fauchon et al., 2002). Furtherwork is needed to determine how wide-spread this stress response isamong fungi (Jamieson, 2002). It is worth noting that the expression ofGSH1 was also up-regulated by Hg2þ but this regulation was Met4p-independent in S. cerevisiae (Westwater et al., 2002).The reason for the induction of GSH synthesis in the presence of Cd2þ

(Fauchon et al., 2002) is the continuous sequestration of bis(glutathio-nato)cadmium complexes into the vacuoles via the ABC transporter Ycf1p(Li et al., 1997). Ycf1p is also suitable to convey As(GS)3 (Ghosh et al.,1999; Rosen, 2002) and Hg(GS)2 (Gueldry et al., 2003) conjugates into thevacuoles. The involvement of the Ycf1p transporter in heavy metaldetoxification processes is not always beneficial. For example, the YCF1mutant of S. cerevisiae was more resistant to selenite than was the wildtype (Pinson et al., 2000). In this case, sequestration of GSH-conjugatedselenite into the vacuoles decreases intracellular GSH levels andtherefore cytoplasmic reduction of selenite to elementary selenium (Pinsonet al., 2000).In S. pombe (Fig. 7), Cd2þ -ions are complexed with phytochelatins

{PC; (g-GluCys)n-Gly, where n is generally in the range of 2 to 5} (Cobbettand Goldsbrough, 2002). These are enzymatically synthesizedoligopeptides and are closely related structurally to GSH (g-GluCysGly).GSH is the physiological substrate for PC synthetases (Clemens et al.,1999; Ha et al., 1999) and are primarily involved in the detoxification ofCd2þ ions although they are reported to interact with other metal ions aswell, including Cu2þ and Agþ (Cobbett and Goldsbrough, 2002). Lowmolecular weight PC-Cd (LMW PC-Cd) complexes are sequestered to thevacuoles through HMT1, an ABC transporter (Ortiz et al., 1995). In thevacuoles, LMW PC-Cd complexes are further processed, namely acid-labilesulfide is built into the molecules, which facilitates the incorporationof further Cd2þ -ions into the aggregates that are called high molecularweight PC-Cd (HMW PC-Cd) complexes. At last, HMW PC-Cd particlesare formed, which consist of a CdS crystallite core coated with PCs(Dameron et al., 1989). Sulfide originates from cysteine sulfinate, a sulfur-containing analogue of aspartate, in a process that follows the steps in theadenine biosynthetic pathway (Speiser et al., 1992; Juang et al., 1993).Two other enzymes related to cadmium stress response in fungi areHEM2 porphobilinogen synthase (Candida glabrata), which is involved inthe biosynthesis of siroheme, a co-factor of sulfite reductase (Hunterand Mehra, 1998) and HMT2 mitochondrial sulfide : quinone oxidoreduc-tase (S. pombe), which detoxifies excess sulfide generated during theformation of HMW PC-Cd (Vande Weghe and Ow, 1999). In S. cerevisiae,


H2S-overproducing MET2 (homoserine O-acetyltransferase) and MET17/MET15 (O-acetylserine and O-acetylhomoserine sulfhydrylase) mutantswere resistant to methylmercury (Ono et al., 1991).Among the antioxidant GSH metabolic enzymes, GR, GPx and GST

have been reported to contribute to the defence against heavy metalstress in different fungi including S. cerevisiae (Pinson et al., 2000; Bronzettiet al., 2001), S. pombe (Kim et al., 2001; Cho et al., 2002; Sa et al., 2002;Shin et al., 2002), A. nidulans (Fraser et al., 2002) and the ectomycorrhiza-building Paxillus involutus (Ott et al., 2002). Paradoxically, chromatesensitivity was found to be reciprocally related to the specific activity ofGR in S. pombe (Pesti et al., 2002, Gazdag et al., 2003). In fission yeast,the NADPH/GR system was the major one-electron Cr(VI) reductant

Figure 7 Proteins and genes contributing to the detoxification of Cd2þ inS. pombe. Abbreviations: GSH, glutathione; PC, phytochelatins; PCS, phytochelatinsynthetase; LMW PC-Cd complexes, low molecular weight phytochelatin-cadmiumcomplexes; HMW PC-Cd complexes, high molecular weight phytochelatin-cadmium complexes; Cyt, cytoplasm, Mt, mitochondrion; V, vacuole. Gene loci indi-cated in the text are hmt1, hmt2, ade2, ade6, ade7 and ade8. (The electron microscopicpicture of S. pombe is presented by courtesy of Dr. Matthias Sipiczki (University ofDebrecen)).


in vivo and, therefore, there was a straight correlation between GR activityand the formation of harmful Cr(V) species (Pesti et al., 2002, Gazdaget al., 2003).

4.6. Nutrient Deprivation Stress

4.6.1. Nitrogen Starvation

In S. cerevisiae cells exposed to nitrogen deprivation, about 90% of totalGSH accumulated in the central vacuole and a transitory stimulation ofGSH biosynthesis was also observed (Mehdi and Penninckx, 1997).The transient overproduction of GSH was inhibited by buthionine-(S,R)-sulfoximine, a specific transition-state-analogue inhibitor of gGCS andwas absent in a GSH-deficient strain (Mehdi and Penninckx, 1997).The transport of GSH molecules into the vacuoles was mainly (about70%) Ycf1p-dependent and a vATPase-coupled system also contributedto this process (about 30%). During nitrogen starvation, gGT activitywas induced and translocated from the Golgi toward the vacuolarmembrane, and the insertion of gGT in the vacuolar membrane facilitatedthe vacuolar GSH transport by increasing the maximal apparent uptakerate (Vapp) three-fold. Further experiments are needed to demonstrateif there is any direct protein–protein interaction between Ycf1p and gGTsimilar to that observed between the human GGTL3B isoform of gGT andthe plasma membrane-associated product of gene CT120 (He et al., 2002)or between yeast gGT and the vacuolar membrane protein of unknownfunction encoded by YOL137w. The gGT-mediated activation of Ycf1pmight be indirect or mediated by an effector-controlled protein–proteininteraction.During nitrogen starvation in S. cerevisiae, about 90% of the total cellular

GSH could be degraded to provide the cells with L-Glu, L-Cys andGly-amino acids that are all easily convertible to other nitrogen-containingorganic molecules through the major metabolic pathways (Mehdi andPenninckx, 1997).In P. chrysogenum, the intracellular GSH concentrations were not

influenced by changes in the nitrogen sources and did not respondto nitrogen deprivation either (Emri et al., 1997b, 1999b). Nevertheless,the specific gGT activity increased under nitrogen starvation enablingthe cells to degrade GSH as does yeast. Both the de novo GSH synthesisand the gGT activity were repressed by NHþ

4 ions in P. chrysogenum (Emriet al., 1997b).


4.6.2. Sulfur Starvation

Saccharomyces cerevisiae can utilize methionine, homocysteine, cysteineas well as GSH as sole sources of sulfur. Exogenous GSH is taken upvia high and low affinity transporters and degraded by gGT and CGase(Fig. 3; Section 3) but alternative gGT-independent GSH-degradationpathways may also exist (Kumar et al., 2003b). A possible pathway for thedegradation of exogenous GSH could involve a membrane-boundcarboxypeptidase releasing glycine and a cyclotransferase-type enzymereleasing cysteine in the cytosol. The existence of carboxypeptidase actingon GSH and its derivatives has been demonstrated in plants (Wolf et al.,1996) but not in fungi.Saccharomyces cerevisiae cells respond to sulfur deprivation by increasing

the turnover rate of GSH and channelling the sulfur content of GSH intocysteine and methionine to maintain protein synthesis at an acceptablelevel. Analogous to nitrogen starvation, yeast cells can mobilize 90% oftheir GSH reserves under sulfur starvation via the gGT-CG pathway(Section 3.4) (Elskens et al., 1991).The mobilization of intracellular GSH reserves under sulfur starvation

was also demonstrated in P. chrysogenum (Emri et al., 1998). Interestingly,neither the GSH producing activity of the cells nor the specific activity ofgGT was influenced by sulfur deprivation in this fungus.

4.6.3. Carbon Starvation

It is well known that adaptation to carbon starvation in yeasts results inan increased tolerance to oxidative stress similar to that observed instationary phase (Jamieson, 1992). Many GSH-dependent and -independentelements of the antioxidant defence system normally functioning duringthe exponential phase of growth are induced under both conditions(Westerbeek-Marres et al., 1988; Lee et al., 1995; Grant et al., 1996a;Flattery-O‘Brien et al., 1997; Jamieson, 1998).The transcriptional factor Yap1p is of primary importance in

the coordination of the antioxidative defence system in S. cerevisiae(Section 4.1) and also exerts a dual role on cell proliferation via the growth-phase dependent regulation of the RPI1 gene, a repressor of the RAS-cAMPpathway (Dumond et al., 2000). In addition, Yap1p has been shown tointeract with the N-myristoylprotein Sip2p, an alternate b subunit of theSnf1p kinase complex, and accumulates in the nucleus during carbon


starvation, which can be inhibited by exogenous GSH (Wiatrowski andCarlson, 2003).In the filamentous fungus P. chrysogenum, both the de novo synthesis

of GSH as well as the intracellular GSH levels increased considerablyunder carbon deprivation and when glucose was replaced with lactose inthe culture medium (Emri et al., 1998). Although the regulation of GSHbiosynthesis is poorly understood in this microorganism, it seems to beindependent of cAMP levels and glucose catabolism (Emri et al., 1998).Unexpectedly, the specific activities of GR and GPx decreased under carbondeprivation and in carbon-depleted stationary phase cultures (Emri et al.,1998; Sami et al., 2001a).

5. GSH-DEPENDENT DETOXIFICATION PROCESSES

5.1. Elimination of Toxic Metabolites

Hansenula polymorpha, C. boidinii, as well as some Kloeckera and Pichia sp.,produce cytoplasmic peroxisome organelles when growing on methanol as acarbon and energy source (Fig. 8). These peroxisomes contain at leasttwo matrix enzymes, a H2O2-generating methanol oxidase and a H2O2-decomposing catalase (Fig. 8) (Sahm, 1977). Formaldehyde produced bythe oxidase is exported towards the cytosol and incorporated in partby central metabolism in yeast. Excess formaldehyde is metabolizedthrough the formaldehyde dehydrogenase (FaDH)-S-formyl glutathione

Figure 8 The metabolism of methanol in methylotrophic yeasts. (1) Methanoloxidase; (2) catalase; (3) formaldehyde dehydrogenase; (4) S-formyl GSH hydrolase;(5) formate dehydrogenase; (6) formaldehyde assimilation pathway. (Reproduced withpermission from Penninckx (2002). � Elsevier Science B.V.)


system, which uses GSH as a cofactor. Methanol induces elevation ofintracellular GSH levels probably via the regulatory action of a YAP1-related, positive transcription factor acting on biosynthesis of gGCS(Ubiyvovk et al., 1999). GSH-deficient mutants of H. polymorpha failedto grow on methanol due to toxic accumulation of formaldehyde. Thiswas also observed with mutants deficient in FaDH, suggesting thatmethanol induces metabolic stress (Sibirniy et al., 1990; Ubiyvovk et al.,1999). The FaDH-pathway is also present in non-methylotrophic yeast likeS. cerevisiae, where it could serve to detoxify formaldehyde formedduring amino acid catabolism (Rose and Racker, 1962). Furthermore,S. cerevisiae deficient in FaDH was viable but sensitive to exogenousformaldehyde (Fernandez et al., 1999).Methylglyoxal (2-oxopropanal) is a 2-oxoaldehyde synthesized both

enzymatically and non-enzymatically in cells (Cooper, 1984; Inoue andKimura, 1995). 2-Oxoaldehydes are harmful to cells since they react withguanidine residues in nucleic acids as well as with arginine, lysineand cysteine moieties in proteins. In situ analysis of methylglyoxalmetabolism in S. cerevisiae showed that its formation was quantitativelyrelated to glycolysis (Martins et al., 2001). Methylglyoxal is generatedas a by-product of glycolysis from triose phosphates and also by thespontaneous decomposition of glyceraldehyde-3-phosphate during growthon glycerol.Glyoxalases (GLO) I and II are two elements of the GSH-dependent

glyoxalase system, which is considered to eliminate methylglyoxal fromcells (Cooper, 1984; Penninckx and Elskens, 1993; Inoue and Kimura,1995). The GLO pathway is highly inducible by glycerol and is subject toglucose repression (Penninckx et al., 1983; Inoue and Kimura, 1996; Bitoet al., 1997). The physiological characterisation of mutants defective inGLOI (encoded by GLO1 in S. cerevisiae) and/or GLOII (the products ofgenes GLO2 and GLO4) did not give a clear-cut answer to the questionas to whether GLOs represent the major route of methylglyoxal disposalin yeast (Penninckx et al., 1983; Inoue and Kimura, 1996; Bito et al., 1997).Nevertheless, S. cerevisiae P27 strain, a GLO1 mutant, showed a suicidephenotype when it was transferred from glucose to glycerol medium(Penninckx et al., 1983), and any impairment of the GLO system resultedin an enhanced sensitivity towards exogenous methylglyoxal (Penninckxet al., 1983; Inoue and Kimura, 1996; Bito et al., 1997). Oxoaldehydes areformed during the catabolism of threonine, valine and isoleucine aswell, and these toxic metabolites may also be detoxified by GLOs (Murataet al., 1986). Moreover, induction of GLO1 expression by osmoticstress in S. cerevisiae is thought to scavenge methylglyoxal, which increased


during glycerol production for stress adaptation (Section 4.2) (Inoue et al.,1998b).

5.2. Detoxification of Xenobiotics

Xenobiotics are man-made chemical substances that are foreign to abiological system. They include environmental agents, fungicides, insecti-cides, drugs, carcinogens, and mutagenic agents. Some of the xenobiotics(e.g. alkaloids) are of natural origin and, in some instances, nucleophilicand redox properties derived from GSH are involved in detoxification.Many xenobiotics have the potential to react either spontaneously with

the SH group of GSH or to form GSH S-conjugates (GS-X) by the aidof glutathione S-transferases (GSTs). Two distinct superfamilies encodeproteins with GST activity: (i) at least 16 genes of cytosolic forms,distributed in 8 families (alpha, mu, theta, pi, zeta, sigma, kappa andomega) and (ii) at least six genes expressed and localized in membranes(Strange et al., 2001). These enzymes were detected in numerousmicroorganisms, including bacteria, protozoa, alga and fungi (Lau et al.,1980; Penninckx and Elskens, 1993; Amstrong, 1994; Vuilleumier, 1997;Sheehan et al., 2001). Relatively little is known about GSTs in fungialthough the activity was detected in more than two dozen strainsdistributed among several yeast genera, e.g. Candida, Hansenula, Pichia,Schizosaccharomyces, Issatchenkia, Rhodotorula and filamentous fungi, e.g.Aspergillus, Neurospora, Penicillium, Mucor, Phanerochaete, Yarrowia(Casalone et al., 1988; Sheehan and Casey, 1993; Dowd et al., 1997) andseveral genes have been sequenced more recently (Table 1).Multiple forms of GSTs are induced by different xenobiotics.

For example, in Issatchenkia orientalis, expression of two GSTs is inducedby o-dinitrobenzene (Tamaki et al., 1999) and microsomal GSTs inAspergillus ochraceus by 3-methylcholanthrene, benzo (a) pyrene andpolychlorinated biphenyls (Datta et al., 1994).Cytosolic GSTs have been purified from filamentous fungi, including

Fusarium oxysporum, Mucor javanicus, I. orientalis, P. chrysosporium andY. lipolytica (Sheehan et al., 2001). Fungal GSTs form 22–25 kDa subunithomodimers (Cohen et al., 1986; Ando et al., 1988; Dowd et al., 1997; Vealet al., 2002), showing limited similarities with the theta-class of animalsand plants (Dowd et al., 1997; Dowd and Sheenan, 1999; Tamaki et al.,1999).Evidence showing that yeast GSTs produce GS-Xs was found when

the cells were exposed to the fungicide chlorothalonil (Tillman et al., 1973;


Lau et al., 1980; Jaspers and Penninckx, 1982; Sheehan and Casey, 1993).Gtt1p and Gtt2p form homodimers and exhibit enzymatic activity with1-chloro-2, 4-dinitrobenzene (Choi et al., 1998). GTT1 is up-regulated byosmotic stress and xenobiotics, and after diauxic shift remain highthroughout the stationary phase. In addition, �GTT1, �GTT2 and �GTT1�GTT2 strains are sensitive to heat shock in the stationary phase(Choi et al., 1998).The physiological functions of Gtt1p and Gtt2p are presently not

well understood. They may participate in elimination of toxic intermediatesthat accumulate in stationary phase, and/or act in a similar fashion asheat shock proteins. Recently, it was shown that the detoxification ofpenicillin side-chain precursors might depend on microsomal GST inP. chrysogenum (Emri et al., 2003)The gstA gene from A. nidulans was found to be homologous to URE2, a

GST-like gene in budding yeast (Coschigano and Magasanik, 1991) and S.pombe (Rai et al., 2003). The mutant �gstA was hypersensitive to severalxenobiotics (Fraser et al., 2002). Three genes, gst1þ , gst2þ and gst3þ

encode theta GSTs in S. pombe (Veal et al., 2002), all show GST activitywith 1-chloro-2,4-dinitrobenzene, and �gst1, �gst2 and �gst3 mutants aresensitive to fluconazole (Sa et al., 2002).A study of the global gene expression response by S. cerevisiae

exposed to methyl methanesulfonate showed induction (>4 fold) of 325transcripts (Jelinsky and Samson, 1999) with 28.8, 8.8 and 7.7 fold inductionof GTT2, CIS2 and PDR10 (an ABC transporter), respectively (Boylandand Chasseaud, 1969; Wolfger et al., 1997; Mehdi et al., 2001).Several experiments have shown that GS-Xs may be transported into

the yeast vacuole (Li et al., 1996; Sharma et al., 2003). As mentioned above,Ycf1p appears as the major GS-X vacuolar transporter in S. cerevisiae.Yeasts might also transport GS-X outside the cells by not yet identifiedsystems ( _ZZa�dzinski et al., 1996).GSH S-conjugates may also form in a GST-independent way, e.g. the

chemical reaction with the antifugal agent Thiram, (CH3)2NC(¼ S)S–S(S¼ )CN(CH3)2, [bis(dimethylthiocarbamoyl)disulfide], will give riseto GS-X GS-S(S¼ )CN(CH3)2 and dimethyldithiocarbamic acid, (CH3)2NC(¼ S)SH (DMDT) (Elskens et al., 1988). The former product istransported into the vacuoles by Ycf1p and is metabolized further todimethylamine, CS2 and H2S, which are all released into the culture medium(Elskens and Penninckx, 1997). Alternatively, the GS-X GS-S(S¼ )CN(CH3)2 can react with GSH, which further increase theintracellular DMDT concentration with the concomitant depletion of theGSH pool (Elskens and Penninckx, 1995a, 1997). The re-oxidation of


DMTD to Thiram by cytochrome c initiates a deleterious redox re-cyclingprocess, which, together with the inactivation of GR by both Thiramand DMTD, prevents the re-establishment of a physiologically relevantGSH/GSSG balance (Elskens and Penninckx, 1995b).

5.3. Glutathione, Regulator of b-lactam Antibiotic Synthesis

GSH shares structural similarities with the b-lactam biosyntheticintermediate d-(L-a-aminoadipyl)-L-cysteinyl-D-valine (ACV) – a tripeptide(Fig. 9), and it has been suggested that penicillin biosynthesis and GSHmetabolism are interconnected in P. chrysogenum (van de Kamp et al.,1999). Furthermore several studies have shown that GSH inhibits bothACV synthetase and isopenicillin N synthetase (Ramos et al., 1985;Nielsen and Jørgensen, 1995; Theilgaard and Nielsen, 1999; van de Kampet al., 1999; Emri et al., 2000; Pocsi et al., 2001). Thus, the pharmaceuticalindustry has had an interest to decrease intracellular GSH concentrationswithout influencing negatively the physiological status of the vegetativeantibiotic-producing tissue. Unfortunately, intracellular GSH levelscould not be affected by selectively feeding mycelia carbon, nitrogen orsulfur sources or by adding GSH-depleting or oxidative stress-generatingcompounds (Emri et al., 1998; Pocsi et al., 2001). To keep the GSH pooleffectively low during the penicillin production phase Pocsi andcollaborators took advantage of the GSH-dependent detoxification abilityof the penicillin side-chain precursors phenylacetic and phenoxyaceticacids, which was attributed to the subsequent action of micro-somal monooxygenase (causing release of toxic epoxide intermediates),microsomal GST and gGT (Figs. 10 and 11) (Emri et al., 1997b, 2000, 2001,2003).Detoxification-based decreases in the GSH pool were provoked by a

well-controlled transient (5 h) lowering of pH to 5.0 at the beginning ofthe production phase in a fed-batch penicillin V fermentation system.

Figure 9 Resemblance of glutathione (GSH) and d-(L-a-aminoadapyl)-L-cysteinyl-D-valine (ACV) structures and metabolism. Enzymes: (1) g-glutamylcysteine synthetase (toppanel) or ACV synthetase (bottom panel); (2) GSH synthetase; (3) gGT; (4) GSH thioltransferase; (5) g-glutamylcyclotransferase (gGCT); (6) 5-oxoprolinase (5OP); (7)GST; (8) glutathione reductase (GR); (9) (di)peptidase(s); (10) NADPH-dependentthioredoxin-based broad-range disulfide reductase sytem (TrxAB); (11) isopenicillin Nsynthetase. Transport steps: A and G, GSX transporters, B and H, GSH transporter; C,


g-glutamyl-amino-acid uptake system; D and E, amino-acid uptake sytsems; F, OPCtransport system. Putative enzymatic steps are indicated by dashed arrows. PM, plasmamembrane; CW, cell wall. (Reproduced with permission from van de Kamp et al. (1999)by the courtesy of the authors. � Kluwer Academic Publishers.)


The increased influx of the protonophoric phenoxyacetic acid into thecells increased the specific GST and gGT activities but the intracellular GSHconcentrations remained unaltered unless the pH of the feed wastransiently lowered below 5.0. At pH 4.6, the GSH pool was depletedrapidly but no antibiotic production was observed, which was explainedby progressing cell death and autolysis. Therefore, the industrialexploitation of the GSH-dependent detoxification of penicillin side-chainprecursors to reduce intracellular GSH-levels in order to avoid theGSH inhibition of the b-lactam biosynthetic enzymes seems unlikely(Emri et al., 2003).Besides acting as a low-molecular-mass regulator of gene expression

in cephalosporin C (CPC) synthesis in Acremonium chrysogenum, thesulfur of methionine (Met) was channelled effectively into cysteine, oneof the amino acid precursors of CPC, through the reverse transsulfurationpathway (Lewandowska and Paszewski, 1988; Kosalkova et al., 2001).A. chrysogenum cells accumulated GSH effectively in the trophophasewhen culture media were supplemented with Met (Lewandowska andPaszewski, 1988; Nagy et al., 2003). The high GSH concentration might bebeneficial for the stabilization of isopenicillin N synthetase (Perry et al.,1988). Met also increased the specific activity of gGT and, hence, theturnover rate of GSH, which facilitated the uptake of Met and made aMet!GSH!CPC sulfur transfer reasonable (Nagy et al., 2003).

Figure 10 Summary of the GSH metabolism of P. chrysogenum NCAIM 0023.(Reproduced with permission from Pocsi et al. (2001). � Akademiai Kiado, Budapest.)


Figure 11 The toxicity of the protonophore side-chain precursors PA and POA islikely to be connected to the formation of toxic epoxide intermediates of hydroxylationreactions on the aromatic rings in P. chrysogenum (Isono, 1954; Eriksen et al., 1994,1998; Emri et al., 2000, 2001, 2003).


6. AGING AND AUTOLYSIS

In most cases, aging and cell death of fungi are connected tointracellular accumulation of ROS (Fig. 12) (Madeo et al., 1997, 1999,2002; Jakubowski et al., 2000; Nestelbacher et al., 2000; Grzelak et al.,2001; Laun et al., 2001; Sami et al., 2001a; Emri et al., 2002). However,it remains unclear if ROS and/or ROS-induced oxidative damages(Costa and Moradas-Ferreira, 2001) are determinants of aging (Wickens,2001).In any case, GSH, as an important antioxidant, is expected to interfere with

these processes, and GSH/GSSG redox imbalances, together with the accu-mulation of ROS, may play an important role in the activation of celldeath programmes. Decreased antioxidant defence, including reducedGSH levels, were observed in yeast during replicative aging (Grzelaket al., 2001) and aging of stationary cultures (Jakubowski et al., 2000).

Figure 12 Intracellular accumulation of ROS may trigger apoptosis in S. cerevisiaeunder many different stressed and unstressed conditions. In yeast, the phenotypic markersof apoptosis include chromatin condensation and margination, DNA breakage as well asthe exposition of phosphatidylserine (Madeo et al., 1999; Laun et al., 2001). ROSalso accumulated in A. nidulans cells exposed to sphingoid long-chain bases prior toapoptosis but there is no causal connection between these phenomena (Cheng et al.,2003).


In catalase-deficient S. cerevisiae mutants, intracellular GSH concentrationswere not affected when the strains were grown on glucose but weresignificantly higher in catalase T and acatalasaemic mutants in theabsence of ethanol as a carbon source (Van Zandycke et al., 2002),indicating that GSH may compensate for the loss of catalase activity(Grant et al., 1998).In S. cerevisiae, mitochondrial accumulation of ROS was observed in

old mother cells prior to apoptosis (Laun et al., 2001), during respiratoryoscillations (Lloyd et al., 2003) and GSH substantially increased the lifespan of yeast cells under oxidative stress (Nestelbacher et al., 2000).The likely involvement of mitochondria in programmed cell death wasalso demonstrated in acetic acid-induced apoptosis of S. cerevisiae(Ludovico et al., 2002) and Zygosaccharomyces bailii (Ludovico et al.,2003). A GSH1 mutant of S. cerevisiae is prone to apoptosis (Madeo et al.,1999) and the apoptotic effect of the mammalian pro-apoptotic proteinBax expressed in yeast was inhibited by the co-expression of tomatoGST/peroxidase, which restored both the intracellular GSH level and themitochondrial membrane potential disturbed by Bax (Kampranis et al.,2000).Although S. cerevisiae is becoming a frequently used model organism

for aging and apoptosis research, those choosing this model should beaware of the substantial differences between the aging and death ofunicellular micro-organisms on one hand and the differentiated, highlyspecialized cells of multicellular organisms on the other hand (Gershon andGershon, 2000). For example, in the absence of homologues to the greatmajority of metazoan apoptosis genes, fungal apoptosis must rely on verydifferent, ‘‘fungus-specific’’ biochemical pathways, which may give specificcharacteristics to this kind of cell death. In fact, deletion of the yeast histonechaperone gene ASF1/CIA1, which resulted in a cell death phenotype, waslargely characterized by classic features of apoptosis but also had somefeatures of necrosis (Yamaki et al., 2001).In filamentous fungi, the situation might be quite different. For example,

ROS did not accumulate and the GSH/GSSG ratio did not decreasein carbon-limited growth and stationary phase of P. chrysogenumcultures prior to autolysis and were therefore unlikely to play a role inthe initiation events of cell death (Fig. 13) (Sami et al., 2001a; Pocsi et al.,2003).Nevertheless, an inherent physiological analogy between autolysis

and apoptosis (McIntyre et al., 1999) cannot be excluded because theaccumulation of ROS is not an obligatory prerequisite of the induction ofapoptosis in more complex eukaryotes (Coppola and Ghibelli, 2000).


In autolytic and post-autolytic phase cells of P. chrysogenum, ROSconcentration increased and cell vitality decreased continuously, althoughsuperoxide dismutase activity and cyanide-resistant alternative respiration –markers of antioxidative defence – were elevated (Sami et al., 2001a, 2001b,2003). Thus, stationary phase hyphae surviving the typical ‘‘yeast-like’’fragmentation of the mycelium (Pusztahelyi et al., 1997a; Pusztahelyi et al.,1997b; 2001b) may undergo oxidative-stress induced apoptosis (Pocsiet al., 2003) and is in agreement with the Free-radical Theory of Aging(Harman, 1993; Jakubowski et al., 2000; Nestelbacher et al., 2000; Launet al., 2001). The term ‘‘aging’’ has been suggested to describe the changescharacteristic of post-autolytic cultures in P. chrysogenum (Sami et al.,2001a; Pocsi et al., 2003).In A. nidulans, GSH was degraded extensively during carbon

starvation due to the action of g-glutamyltranspeptidase, which resultedin a severe GSH/GSSG redox imbalance before the onset of autolysis(Fig. 13) (Emri et al., 2002). Interestingly, other physiological changes,e.g. intracellular accumulation of ROS, were quite similar to those observedin carbon-depleted P. chrysogenum cultures where the GSH/GSSGratios did not decrease prior to, and even increased during, autolysis(Fig. 13) (Sami et al., 2001a; Emri et al., 2002; Pocsi et al., 2003). Therefore,the initiation and/or signal transduction pathways of the autolysis of

Figure 13 Changes in the GSH/GSSG ratios in A. nidulans (j) and P. chrysogenum(u) cultures. Reproduced from Emri et al. (2002) with permission. � Akademiai Kiado,Budapest. Both cultures were carbon-limited and autolysis started at 34 and 50 h ofincubation, respectively (Pusztahelyi et al., 1997a; Emri et al., 2002). Despite temporaldifferences in the GSH/GSSG ratios, the morphological and physiological markers ofautolysis were similar in both cases.


filamentous fungi are unlikely to proceed through the modulation ofintracellular GSH/GSSG levels.

7. CONCLUSIONS

GSH is a crucial multifaceted cellular metabolite in Fungi. Its clearparticipation in the response of suffering cells subjected to situations ofstress places GSH in the category of altruistic compounds. The mostparticular physiological functions of GSH relate indeed to the state ofstress. For microorganisms, stress signifies the response to a chronic orsudden experience of various harmful circumstances like starvation, heat,cold, osmotic shock, alterations in the pH or water potential, exposure toreactive oxygen species or radiation. These situations refer however tocultures grown in ‘‘optimal’’ laboratory media, which rarely represent themost favourable ecological condition or even may hide unexpecteddevelopment of stress, for example the appearance of harmful metabolicby-products normally generated by general carbon and/or nitrogen cellularmetabolism. In this case, GSH plays a quiet role but is far from beingnegligible. In fact, the role of GSH is not limited to extremely stressfulconditions at all. A cell fully deprived of GSH is simply unable to surviveeven under conditions free of stress. One of the main roles of GSH isrelated to the maintenance of cellular architecture, in particular to theintegrity of membrane structures, as well as to cell differentiation anddevelopment. To fulfil this fundamental function, the fungal cell does notneed necessarily to use its full potential for the sythesis of GSH; oftenonly minute amounts of the thiol are sufficient. There are severalhighly important enzyme systems associated with GSH. However, thechemical reactivity of the tripeptide, which modulates its implication inthe cellular redox cycle, should not be forgotten! Undoubtedly, this is oneof the most important features that explains the versatility of thiscompound.Once again, we must emphasize the deep influence exerted by investi-

gators of mammalian physiology, like the late Alton Meister, on thedevelopment of GSH research. Research on GSH in microorganisms,particularly in fungi, has exploded in the last 10 years. This can be attributedin part to the enthusiasm of investigators working in very different fields,e.g. plant and animal physiology, to use microbial model organisms,particularly yeasts. Nevertheless, one of the underlying objectives of thisreview was to show how GSH-oriented research impacts upon microbial


physiology. Currently, the activities of the GSH community extend in allthese directions and we hope that this will continue.

ACKNOWLEDGMENTS

These projects were supported financially by the Walloon Region and theBelgian National Fund for Scientific Research (FRNS) for M.J.P., and theHungarian Office for Higher Education Programmes (grant referencenumber 0092/2001) and by the OTKA (grant reference numbers T034315and T037473) for I.P. The authors’ visits were facilitated by the EuropeanCommunity ERASMUS Exchange Programme. The Hungarian Ministry ofEducation awarded a ‘‘Szechenyi Scholarship for Professors’’ grant to I.P.Many thanks are addressed to numerous colleagues of the GSH communitywho communicated to us valuable pieces of information as well as paperswhich were in press at the time of submission. Last but not least, specialthanks are addressed to our spouses and children for their immeasurablepatience.

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The Role of the Flavodiiron Proteins inMicrobial Nitric Oxide Detoxification

Lıgia M. Saraiva, Joao B. Vicente and Miguel Teixeira*

Instituto de Tecnologia Quımica e Biologica, Universidade Nova de Lisboa,

Apartado 127 Avenida da Republica (EAN), 2781-901 Oeiras, Portugal

ABSTRACT

The flavodiiron proteins (first named as A-type flavoproteins)constitute a large superfamily of enzymes, widespread amonganaerobic and facultative anaerobic prokaryotes, from both theArchaea and Bacteria domains. Noticeably, genes encoding forhomologous enzymes are also present in the gD:\dataset\Poole-Vol-49\PDFenomes of some pathogenic and anaerobic amitochondriateprotozoa. The fingerprint of this enzyme family is the conservation of atwo-domain structural core, built by a metallo-b-lactamase-like domain,at the N-terminal region, harbouring a non-heme diiron site, and aflavodoxin-like domain, containing one FMN moiety. These enzymeshave a significant nitric oxide reductase activity, and there is increasingevidence that they are involved in microbial resistance to nitric oxide.In this review, we will discuss available data for this novel family ofenzymes, including their physicochemical properties, structural andphylogenetic analyses, enzymatic properties and the molecular geneticapproaches so far used to tackle their function.

*Corresponding author. Fax: 351-214411277; E-mail: [email protected]



DOI 10.1016/S0065-2911(04)49002-X

1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 781.1. Chemistry of NO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 781.2. Biological chemistry of NO. . . . . . . . . . . . . . . . . . . . . . . . . . . 801.3. NO and microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

2. The family of flavodiiron proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . 882.1. Structure of the D. gigas enzyme . . . . . . . . . . . . . . . . . . . . . . . 882.2. Modular organisation – classification of the flavodiiron proteins . . . . . . 922.3. Unity and diversity of electron transfer chains . . . . . . . . . . . . . . . . 932.4. Amino acid sequence analysis. . . . . . . . . . . . . . . . . . . . . . . . . 962.5. Physicochemical properties . . . . . . . . . . . . . . . . . . . . . . . . . . 1012.6. Function of flavodiiron proteins . . . . . . . . . . . . . . . . . . . . . . . . 114

3. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

1. INTRODUCTION

Widely present in biological systems, the nitric oxide (NO) molecule hasboth beneficial and deleterious effects. These effects are intimately associatedwith iron and oxygen (and reactive oxygen species, ROS) metabolism. Nitricoxide utilisation will depend on multiple factors, from the environmentalmedium to the biochemical role to be performed, which can range fromintracellular signalling to defence mechanisms against pathogens. Thebiological chemistry of nitric oxide has been extensively reviewed recently(Cooper, 1999; Ignarro, 2000). Therefore, only a short introduction to NOchemistry and biology will be presented in this section.

1.1. Chemistry of NO

Nitric oxide1 is a small diatomic molecule, with a low electric dipolemoment of 0.159 Debye (Lide, 1997), and which has a solubility in waterof � 2 mM at room temperature and a partial pressure of 1 atm. Due to itslow dipole moment, it is more soluble in hydrophobic media than in water,being thus capable of rapid diffusion across biological membranes. NO isa free radical, with one unpaired electron in a �* antibonding molecularorbital. Since the nitrogen atom has a higher orbital coefficient in this �*orbital, the radical chemistry occurs mainly through the nitrogen atom.

1Although the more correct name is nitrogen monoxide, the traditional nomenclature will be

used in this chapter.

78 LIGIA M. SARAIVA, JOAO B. VICENTE AND MIGUEL TEIXEIRA

NO has a short lifetime in a cellular environment, due to its high reactivitywith a wide range of molecules. In fact, the combination of its shortlifetime with its high reactivity explains why it is so widely used in biologicalprocesses, in spite of its toxicity.The solution and redox chemistry of NO is very complex

(Fukuto et al., 2000). The nitrogen atom in NO has a formal oxidationstate of 2þ , intermediate between the þ 5 state, in nitrate (NO�

3 ), andthe –3 state, in ammonia (NH3) (Fig. 1) (Philips and Williams, 1965);hence, it may be either further oxidised or reduced. NO has anintermediate reduction potential, for its immediate reduction products,which is dependent on the spin ground state of the product (E0 NO/3NO� ofþ 0.39 V, E0 NO/1NO�

�0.350 V)2; however, reduction to the more stableform, N2O, has a quite high reduction potential of þ 1.59 V (standard acidicconditions; E0¼ 1.18 V at pH 7).Nitric oxide may react with nucleophilic molecules, namely thiol groups,

forming S-nitrosothiols (RS-NO) (Gaston, 1999; Fukuto et al., 2000).

Figure 1 Oxidation state diagram for nitrogen species. Volt equivalents are plotted as afunction of the oxidation state of the nitrogen species in acid solution (aH¼ 1.0; solid line)and basic solution (aOH¼ 1.0; broken line). (Adapted from Philips and Williams (1965).)

2The 3 and 1 superscripts refer to the triplet and doublet electronic states of NO�.

FLAVODIIRON PROTEINS IN NO DETOXIFICATION 79

In particular, S-nitroglutathione (GSNO), formed by the reaction of thetripeptide glutathione with NO, is considered one of the most importantNO releasers. NO also reacts with metal centres, such as iron, either inthe ferrous or ferric states, yielding iron nitrosyl complexes. These com-plexes can catalyse nitrosylation reactions, enhancing the deleterious effectsof NO. NO can also be reduced to NO�, or HNO; this species can formN2O or react further with NO, forming N2O

�2 , which decomposes into

the hydroxyl radical, .OH, and N2O or, by reaction with NO, intoN3O

�3 (another unstable species, that decomposes into N2O and NO�

2 ).Thus, NO may generate the most dreadful radical, .OH, under anaerobicconditions.In the presence of oxygen, its chemistry is far more complex. Direct

reaction with dioxygen forms nitrogen dioxide, .NO2, a very strongoxidant (E0

.NO2/NO�2 ¼ 1.04 V), which reacts efficiently with thiolates

or hydroxyl anions, forming the respective radical species, or nitrosatedspecies, such as S-nitrosothiols. Dimerisation of .NO2 yields dinitrogentetroxide, N2O4, an unstable species in water, that decomposes intonitrite and nitrate; further reaction of .NO2 with NO generates dinitrogentrioxide, N2O3, also unstable, decomposing into nitrite. The most impor-tant reaction of NO is probably that with the superoxide anion, O�

2. , which

leads to the formation of the very powerful oxidant peroxynitrite, witha rate constant close to the diffusion limit (� 109 M�1 s�1). The reactionof peroxynitrite with NO or O�

2. generates nitrite or .NO2.

Nitric oxide can be produced, abiotically, by mild acidification of nitritesolutions; this process has been considered quite relevant in medicalterms, since intracellular acidiosis occurs after ischemia or shock, togetherwith hypoxia, leading to NO concentrations that may be much higherthan those produced enzymatically (Zweier et al., 1999).The actual reactivity of nitric oxide will depend on both the rate

constants for each reaction as well as on the respective concentrations ofeach nitrogen oxide and oxygen reactive species. All those species, namedReactive Nitrogen Species, RNS, are at the heart of the biological effectsof NO, and at the same time make it difficult to identify the actualreacting nitrogen species. It is also clear that the solution chemistry of NOis intimately related with that of ROS.

1.2. Biological Chemistry of NO

The multiple roles of NO in biological systems are closely associated with itscomplex chemistry (Cooper, 1999; Fukuto et al., 2000; Miranda et al., 2000).


The main biological targets are: (i) iron centres, leading to the formationof iron nitrosyl complexes (heme, iron-sulfur and other iron proteins,liberation of iron from ferritin, the main iron storage protein), (ii) thiolates,with the formation of S-nitrosothiols and (iii) radical species. These reac-tions have multiple and profound biological effects, as they lead to acti-vation or inhibition of enzymes, ion channels, and transcription regulators.In eukaryotes, NO is generated enzymatically by NO synthases (NOS)

(Stuehr, 1997), which use oxygen and NADPH to produce NO by oxidationof L-arginine. At low concentrations (down to the nanomolar level)NO has a major role as a signalling molecule, in processes of neuronalcommunication, vasodilation, smooth muscle relaxation and inhibition ofplatelet aggregation (Ignarro, 2000; Grange et al., 2001; Miranda et al.,2003; Fubini and Hubbard, 2003). At higher concentrations (micromolarto millimolar levels), NO in association with oxygen reactive speciesis responsible for multiple disorders, including tissue inflammation,chronic infection, malignant transformations and degenerative diseases(Fang, 1997; Bogdan et al., 2000). Cells respond to these aggressions(generally named nitrosative stress) by activation of redox/NO responsivegenes, encoding for enzymes involved in detoxification, export, repair andother homeostatic functions. NO constitutes also one of the weaponsof the mammals immune system to fight pathogens (Fang, 1997;MacMicking et al., 1997). It is observed that the NO released frominterferon g or bacterial lipopolysacharides-activated macrophages con-tribute to their citotoxicity (Kim and Ponka, 2000). The microbicidal effectof NO plays also an important role in the different stages of defenceresponses of infected plants (Delledonne et al., 1998; Klessig et al., 2000;Wendehenne et al., 2001). NO appears also to regulate mitochondrialrespiration through its reaction with the heme-copper oxygen reductases(Brunori et al., 1999; Brown, 2001; Brunori, 2001; Cooper, 2002; Sartiet al., 2003).

1.3. NO and Microbes

In prokaryotes, NO is an intermediate in the stepwise process ofdenitrification (reduction of nitrate or nitrite to nitrogen) (Zumft, 1997;de Vries and Schroder, 2002; Wasser et al., 2002). NO is formed by themulticopper or the cd1 nitrite reductases, being subsequently reduced toN2O by the transmembrane enzyme NO reductase (cytochrome orquinol : NO oxidoreductase), which contains a high-spin heme-irondinuclear site as the catalytic centre. It has been recently proposed that


NO may also be formed during ammonification (reduction of nitrate ornitrite to ammonia), by action of the pentaheme nitrite reductases (Corkerand Poole, 2003).Prokaryotes not involved in the nitrogen cycle may nevertheless

encounter NO, produced abiotically (e.g. by decomposition of nitrite), orbiotically, by denitrifiers/ammonifiers and by the action of the immunesystem of their hosts. Additionally, bacteria are also able to synthesiseNO, as observed for example for Lactobacillus fermentum (Morita et al.,1997) and Escherichia coli (Corker and Poole, 2003). In fact, the genomesequences of some prokaryotes contain genes encoding for NO synthase-like proteins. Bacterial NO synthases were already isolated fromStaphylococcus aureus (Hong et al., 2003), Bacillus subtilis (Adak et al.,2002) and Nocardia species (Chen and Rosazza, 1995), and shown to beable to produce NO. Very recently, sulfite reductase flavoproteins wereproposed to be the bacterial NOS reductase domain (Zemojtel et al., 2003).The occurrence of NO synthases in bacteria suggests a physiological rolefor nitric oxide in prokaryotes, which, however, remains to be clarified.The bacteriostatic NO concentrations (in the micromolar to millimolar

range) have deleterious effects in the pathogen, such as protein orlipid nitrosation/nitrosylation, nitrosylation of iron centres (e.g. of theaconitase tetranuclear iron-sulfur centre), and inactivation of ribonucleotidereductase (Lepoivre et al., 1991; De Groote et al., 1995; Roy et al.,1995; Hausladen and Fridovich, 1996; Demple, 1999, 2002). Besides itsdirect action, NO toxicity occurs associated with reactive oxygen species,and some bacteria are much more sensitive to S-nitrosothiols, which areformed in high concentrations in infectious and inflammatory states, thanto NO. NO is also an indirect DNA-damaging agent: while auto-oxidationof NO, yielding N2O3, causes deamination of the DNA bases, peroxy-nitrite induces nitration of the bases (Burney et al., 1999). Thus, bacteriahad to develop several responses to toxic levels of nitric oxide or itsderivatives.The prokaryotic responses, which are not yet completely established,

include at least five distinct types of enzymes/proteins: (i) enzymesthat directly detoxify NO or S-nitrosothiols; (ii) enzymes that detoxifyreactive oxygen species, thus avoiding the formation of reactivenitrogen species that are formed as a result of reaction of NO withROS; (iii) enzymes that allow regeneration of reduced pyridinenucleotides, thus counteracting the effects of oxidative and nitrosativestress; (iv) DNA-repairing enzymes; (v) regulators of iron homeostasis,decreasing the formation of iron-nitrosyl species, that may act as cata-lysts of nitrosylation reactions. Nitrosative stress also leads to a quite


complex signalling network, associated with the responses to oxidativestress agents. The E. coli system will be presented in some detail, as it is themost studied one.

1.3.1. Genetic Responses to Nitrosative Stress

The two main regulons of E. coli related to oxidative stress aggressions, theSoxRS and OxyR systems, are also sensitive to nitrosative stress, throughthe attachment of NO to a metal or thiol centre, a common theme inregulation of cellular functions (Demple, 2002). SoxRS and OxyR mutantsinduce a higher sensitivity of E. coli to NO produced by macrophages, or toS-nitrosothiols (Nunoshiba et al., 1993; Hausladen et al., 1996). Underoxidative stress conditions, the SoxRS system works in two steps: theSoxR protein contains a dinuclear FeS centre ([2Fe–2S]2þ /1þ ), which inthe reduced (1þ ) state is transcriptionally inactive; upon oxidation to the2þ state it triggers the transcription of soxS, which in its turn stimu-lates the transcription of about 65 other genes encoding for key defenceenzymes such as manganese superoxide dismutase (SOD) and endonucle-ase IV. This regulon is activated by nitric oxide (e.g. produced by macro-phages), thus providing E. coli with some resistance to NO: NO bindsto the binuclear centre of SoxR, forming a dinitrosyl-iron dithiolcomplex, which activates soxS (Ding and Demple, 1997). The inductionof enzymes such as SOD is quite important, since it helps to avoidthe formation of peroxynitrite, by the reaction of the superoxide anionwith NO.OxyR, which contains cysteine redox centres, is activated by S-

nitrosylation, S-hydroxylation and S-glutathionylation, and induces theexpression of multiple genes that protect from oxidative and nitrosativestress, such as catalases, glutathione reductase, and alkylhydroperoxidereductase (Hausladen et al., 1996; Kim et al., 2002).The relation between iron homeostasis and NO stress is well

exemplified by the effect of NO on the Fur (ferric uptake regulator)protein. Fur regulates not only the transcription of genes encoding forproteins of iron metabolism, but also for oxidative and acid stress, includingmore than 90 genes (Hantke, 2001). At low intracellular iron contents, Furloses its iron ion, thus releasing the transcriptional repression of multiplegenes. Fur is under the control of the OxyR and SoxRS systems (Zhenget al., 1999). In the presence of NO, an iron-nitrosyl species is formed,which inactivates its repressor activity (D’Autreaux et al., 2002), causinga general derepression of Fur regulated genes.


The general oxygen regulator FNR (Fumarate Nitrate Regulator)contains a [4Fe–4S]2þ /1 centre that is oxygen and NO labile. Since thecluster controls the protein dimerisation and its DNA binding capability,nitrosylation of the centre leads to an inactivated form of FNR (Cruz-Ramos et al., 2002). Although inactivation of FNR by NO was, so far, onlyshown to affect E. coli hmp regulation, it is to be expected that all FNR-regulated genes will respond to NO. As will be discussed later, more systemsare certainly involved in bacterial responses to nitrosative stress.

1.3.2. Prokaryotic Defence Systems Against Nitrosative Stress

Besides the activation of the OxyR and SoxRS systems, which limit theeffects of RNS associated with reactive oxygen species, in prokaryotes atleast two other families of enzymes were shown to be involved in nitricoxide metabolism, besides the newly identified family of flavodiiron NOreductases,3 which will be discussed at length in the second part of thischapter. Those already known enzymes are the membrane-bound heme-iron NO reductases of denitrifiers, and the globins, a family of cytoplasmicproteins present in a wide range of organisms. Additionally, the pentahemenitrite reductase (Nrf) of E. coli seems to be able to perform NOdetoxification, since the nrf minus strain cultured anaerobically with 20 mMof sodium nitrate and exposed to 150 mM of NO suffered a significantgrowth inhibition (Poock et al., 2002). It remains to be clarified if thatbehaviour is indeed a response mechanism, or just a reflection of the factthat these nitrite reductases fully reduce nitrite to ammonia, being alsocapable of reducing intermediates of this six electron process, such ashydroxylamine (Einsle et al., 2002; Rudolf et al., 2002) or nitric oxide (Costaet al., 1990). Cytochromes c0 may also play a role in NO metabolism, asjudged by the example of Rhodobacter capsulatus. In this organism,cytochrome c0 was shown to confer resistance to NO (Cross et al., 2000) andto behave as a NO reductase (Cross et al., 2001).

1.3.2.1. The globin family of proteins. The function of prokaryoticglobins (reviewed in Poole and Hughes (2000), Frey et al. (2002), Frey andKallio (2003)) remained elusive for a long time, but there is growingevidence that they are involved in NO detoxification. The globins can bedivided into two classes: those that are built by a single B-type heme

3We propose to rename this enzyme family as the flavodiiron proteins, FDP, which allows us

to include in its designation the common redox sites, the flavin and the di-iron site, as first

suggested by D. Kurtz and coworkers (Silaghi-Dumitrescu et al., 2003).


containing globin domain (hemoglobins), and those which have an extraNADPþ : ferredoxin oxidoreductase-like domain, containing an FADmoiety and an NAD(P)H binding motif (know as flavohemoglobins). Whenexpressed in E. coli, the single domain hemoglobins of Vitreoscilla andMycobacterium tuberculosis protected against nitrosative stress (Frey et al.,2002; Pathania et al., 2002a, 2002b). Furthermore, fusion of Vitreoscillahemoglobin with a heterologous reductase domain not only increasedsignificantly the rates of NO consumption but also avoided the forma-tion of thionitrosyls or peroxynitrite derivatives at the heme domain(Hausladen et al., 1998; Kaur et al., 2002). However, the Vitreoscillahemoglobin gene expression was not enhanced by RNS or ROS (Freyet al., 2003), contrary to what is generally observed for flavohemoglobins(see below).Flavohemoglobins (Hmp) are widespread among bacteria, as well as in

yeasts and other fungi, and in protozoa. In the presence of oxygen,Hmp oxidises NO to nitrate, probably through a denitrosylase mechanism,with a turnover of � 94 s�1 at 200 mM oxygen and 1 mM NO, for theE. coli enzyme. The range of activities reported varies from 7.4 to 128 s�1,at 20�C (Hausladen et al., 1998; Gardner et al., 1998b; Hausladenet al., 2001; Frey and Kallio, 2003). Anaerobically, HMP reduced NO toN2O, with a much lower activity of ca. 0.14–0.5 s�1 ( Kim et al., 1999; Freyand Kallio, 2003). Nevertheless, a Salmonella typhimurium strain witha deletion on the hmp gene was more sensitive to NO and S-nitrothiols,even anaerobically, than the wild type strain (Crawford and Goldberg,1998). Thus, it has been proposed that Hmps are active mainly underaerobic conditions, although a function under anaerobiosis cannot becompletely excluded.Several experimental findings indicate a role for Hmp in the resistance of

pathogens to their hosts. For example, deletion of hmp in Erwiniachrysamthemi, a plant pathogen, decreased significantly its pathogenicity(Favey et al., 1995). In S. typhimurium, Hmp was reported to contributeto the microbial protection from killing mediated by NO-released fromhuman macrophages (Stevanin et al., 2002).Besides its function as NO scavenger, bacterial globins, either single-

or two-domain, have since their discovery been suggested to play animportant role in aerobic metabolism, due to their capability of reversiblybinding the oxygen molecule, as the canonical hemoglobins. For example,Vitreoscilla globin, when heterologously expressed, increased the cell yield ofthe host bacterium grown aerobically, concomitant with a higher level ofprotein synthesis, ATP production and a higher proton flux per moleculeof oxygen reduced (Frey and Kallio, 2003). A direct interaction between


the Vitresocilla globin and the heme-copper oxygen reductases was alsoproposed (Frey and Kallio, 2003). Other studies showed that stimuli such asoxygen deprivation or oxidative stress increased the expression of the hmpgene: hmp was induced by paraquat, protecting against superoxideeffects (Membrillo-Hernandez et al., 1996, 1997b). In S. typhimurium hmpmutant strain was shown to be more sensitive to oxygen stress than the wildtype strain (Stevanin et al., 2002). It was also observed that the presence ofHmp was required for full induction of oxidative stress response enzymes,such as NADPH : ferredoxin oxidoreductase, a member of the SoxRSregulon. More recently, E. coli Hmp was proposed to act as an alkyl-hydroperoxide reductase (Bonamore et al., 2003b), which is in accordancewith previous results that suggest that Hmp has a peroxidase-like activesite structure (Mukai et al., 2001). Furthermore, Hmp from Ralstoniaeutrophus and E. coli were reported to interact with membranes lipids(Ollesch et al., 1999; Bonamore et al., 2003a), through conserved regions(Ermler et al., 1995; Ilari et al., 2002), therefore suggesting participationin lipid membrane regeneration. In particular, E. coli Hmp was shown tohave specific recognition sites for unsaturated or cyclopropanated fattyacids, their binding spectroscopic alterations in the visible spectrum of theferric heme. Ligation of the lipid was proposed to involve the heme iron andthe region above the heme pocket. Furthermore, the lipid binding was foundto induce changes in the kinetics of CO-binding and oxygen release,suggesting an interplay between both functions (Membrillo-Hernandez et al.,1997a; Bonamore et al., 2003a).Thus, in summary, at present, the role of Hmp in oxidative stress

response and/or as an oxygen delivery system appears to be important.It still remains to be clarified whether these enzymes have diverse functionsin different organisms, or are indeed multifunctional.The mechanisms of hmp regulation are quite complex and not yet

fully understood. In E. coli, hmp was found to be independent of theOxyR and SoxRS regulators (Poole et al., 1996; Membrillo-Hernandezet al., 1997b; Hausladen et al., 1998), although Hmp expression is neededfor full activation of the SoxRS system and for resistance to paraquat, aswell as for full expression of SodA in response to paraquat (Membrillo-Hernandez et al., 1996; Poole et al., 1996). E. coli hmp was found to beinduced by nitrate and, more effectively, by nitrite, apparently independ-ent of the NarLP regulatory systems (Poole et al., 1996). Several studiesshowed that E. coli hmp was also up-regulated by NO through at leastthree factors: FNR, Fur and MetR, the global regulator of the methion-ine biosynthesis pathway. In fact, it was shown that the repression causedby FNR and Fur on E. coli hmp could be relieved by addition of NO,


which directly inhibits FNR and Fur through nitrosylation of therespective iron centres (Cruz-Ramos et al., 2002; D’Autreaux et al., 2002).Also, it was observed that SNP and GSNO cause nitrosation ofhomocysteine, the cofactor of MetR, and that the binding of MetR tothe intergenic glyA-hmp region that occurs under these conditionsresults in an up-regulation of hmp expression (Membrillo-Hernandez et al.,1998).

1.3.2.2. Other enzymatic systems. Microbes may use other strategies torespond to the presence of nitrosative stress, although their generalrelevance is less well known. A process for detoxification of S-nitrosothiolswas recently identified in Arabidopsis (Sakamoto et al., 2002), E. coli,Saccharomyces cerevisiae and mouse macrophages (Liu et al., 2001): S-nitrosoglutathione and S-nitrosated proteins are metabolised by theglutathione-dependent formaldehyde dehydrogenase, with the formationof GSNH2 and, ultimately, formation of ammonia with the concomitantoxidation of NADH. The catalytic rate constant for GSNO is close to thediffusion limit. The glutathione-dependent formaldehyde dehydrogenasesare widely present in living organisms and highly conserved, frombacteria to higher eukaryotes. These observations strongly suggest thatbesides the role of glutathione as a redox buffer, it plays also a major rolein controlling the intracellular levels of nitrosylated species. Homocysteinewas also proposed to act as an endogenous NO antagonist. InS. typhimurium, a mutation in metL, whose gene product catalysesmetabolic steps required for homocysteine biosynthesis, conferred hyper-susceptibility to S-nitrosothiol and lowered the virulence of this pathogenin mice (De Groote et al., 1996).Bacterial defences against peroxynitrite-mediated damage of methionine

residues in proteins were reported as involving the peptide methioninesulfoxide reductase. The deletion in E. coli of this enzyme, which catalysesthe reduction of methionine sulfoxide in proteins to methionine, madethis organism hypersensitive to GSNO and nitrite in aerobic conditions(St John et al., 2001).Another strategy used by bacteria to escape NO was revealed in

Helicobacter (H.) pylori (Gobert et al., 2001). In this system, bacterialresistance occurs indirectly by preventing the NO production bymacrophages through scavenging of arginine, the common substrate ofarginase and NOS. It was shown that while wild type H. pylori inhibitsthe NO produced by activated mice macrophages, inactivation of thearginase gene restored NO production.


2. THE FAMILY OF FLAVODIIRON PROTEINS

The flavodiiron proteins are widespread among Bacteria and Archaea,either in strict or facultative anaerobes; the genomic data on someamitochondriate and anaerobic protozoa show also the presence of genesencoding for putative flavodiiron proteins (Wasserfallen et al., 1998;Frazao et al., 2000; Andersson et al., 2003). These enzymes were firstlynamed A-type flavoproteins (Wasserfallen et al., 1998), and appear in thedatabases with quite diverse designations. The distinctive feature ofthis family is the common core unit, built by two structural modules: theN-terminal domain, with a metallo b-lactamase like fold, harbouringa diiron catalytic site, and the second domain, having a short chainflavodoxin-like fold with a FMN moiety (Frazao et al., 2000). Only a fewmembers of this family were purified from their natural hosts, namely theenzymes from Desulfovibrio (D.) gigas (Chen et al., 1993b), R. capsulatus(Jouanneau et al., 2000), and Methanothermobacter thermoautotrophicus(Nolling et al., 1995; Wasserfallen et al., 1995). The first function assignedto these enzymes was that of the D. gigas enzyme, which was shown toreduce oxygen to water, receiving electrons from a rubredoxin (Chenet al., 1993b; Gomes et al., 1997); therefore, it was named rubredoxin :oxygen oxidoreductase (ROO). However, later on it was shown that theorthologous enzyme from E. coli had a quite considerable NO reductaseactivity (Gomes et al., 2002b). The enzymatic studies so far performed willbe discussed in Section 2.6. In the following sections, the structural andphysicochemical properties of these enzymes will be presented, using asprototypes the two best studied enzymes, from D. gigas and E. coli.

2.1. Structure of the D. gigas Enzyme

The structure of D. gigas ROO is the only one available for this familyof enzymes [PDB entry 1e5d (Frazao et al., 2000)]. The enzyme was iso-lated as a functional homodimer. The X-ray crystal structure, solved at aresolution of 2.5 A, confirmed this quaternary structure: the enzymecrystallises as a dimer, in which the two monomers have a head-to-tailarrangement (Fig. 2). Two structurally distinct domains build up eachmonomer. The N-terminal domain (up to Gln249)4 has an �b/b�arrangement, with the two inner b-sheets surrounded by solvent exposed

4Unless otherwise stated, the aminoacid numbering of the D. gigas enzyme will be used.


helices towards the external faces (Fig. 2). The fold is very similar to thatof class B Zn-b-lactamases (r.m.s. values of C� atoms of 1.4 to 1.7 A),although the amino-acid sequence identities are lower than 15%. But, incontrast with the lactamases, which contain a mono- or a di-zinc centre,ROO has a diiron centre, which nevertheless occupies essentially the samespatial position as the zinc centre in the lactamases. Furthermore, thesubstrate binding residues and cavity required for the lactamase activityare not conserved in ROO: this space is occupied by an additional two-stranded b-sheet, the first helix of the C-half of the sandwich and itspreceding extra loop, and contacts with the other monomer (Fig. 2B), whichaltogether cover the metal site. The second domain (which starts at Lys250)has a typical �b� flavodoxin-fold (Fig. 2A) and contains one flavinmononucleotide (FMN).

Figure 2 Structural features of D. gigas Rubredoxin : oxygen oxidoreductase, a ClassA FDP which is the structural prototype of the FDP family. Panel A: 3D crystallographicstructure of D. gigas ROO (PDB entry: 1e5d) illustrating its quaternary structure as afunctional head-to-tail homodimer, which places the FMN cofactor of one monomer incontact with the diiron centre from the other monomer. Different colours representdifferent monomers, whereas different tones of the same colour represent the differentmodules; Panel B: metallo b-lactamase domain of D. gigas ROO bearing the non-hemediiron centre; Panel C: first coordination sphere of the non-heme diiron centre in D. gigasROO, with the ligands H79, E81, D83, H146, D165, H226. (See colour plate section.)


2.1.1. The Diiron Centre

The iron ions are coordinated by histidines, glutamates and aspartates(H79-X-E81-X-D83-X62-H146-X18-D165-X60-H226, Fig. 2C), and bridgedby a m-oxo (or hydroxo) species and by the carboxylate of Asp165. Fe1,with a square pyramidal geometry, is further bound to His79, Glu81,and His146, while Fe2 assumes a quadrangular planar geometry, beingbound to Asp83 and His226. Close to Fe2 there is a water molecule, whichis at a non-bonding distance. The carboxylate oxygen OD1 of Asp83 iswithin hydrogen bonding distance to the m-oxo (hydroxo) species.Further electron density close to Fe2 was interpreted as an oxygenmolecule at H-bonding distance to the carboxylate oxygen OE2 ofGlu81. The iron-to-iron distance (3.4 A) is compatible with thosedetermined for di-ferric centres. This set of ligands places the ROO ironcentre in the family of the carboxylate/histidine diiron centres, presentin a variety of enzymes, such as methane monooxygenase (MMOH),Type I ribonucleotide reductases (RNR), (hemo)ferritins (Macedoet al., 2003) and hemerythrin (Wallar and Lipscomb, 1996; Kurtz,1997; Solomon et al., 2000). In these enzymes, the diiron centre isinvolved in oxygen binding (hemerythrin, the oxygen carrier in nematodes),or in oxygen activation, as a first step to perform their function(e.g. oxidation of methane to methanol by MMOH, oxidation of ferrousto ferric iron in ferritins, radical generation in RNR). However, theseenzymes have a four-helix bundle structural motif, quite distinct from thelactamase fold of ROO.The dimeric head-to-tail arrangement has a strong functional

implication. In fact, within each monomer, the two redox centres (thediiron site and the FMN) are quite far apart, at ca. 35 A, which limitsconsiderably an efficient electron transfer between them. However, in thedimer, the diiron centre from one monomer is very close to the FMN ofthe other monomer (Fig. 2). Indeed, the FMN methyl group C8M is invan der Waals contact with the carboxylate from Glu81, one of the ligandsto Fe1, assuring a quite efficient electron transfer pathway between thetwo redox centres (Frazao et al., 2000).

2.1.2. The Metallo �-lactamase Domain

The metallo b-lactamase structural domain is found in a large numberof very diverse enzymes, from the three life domains: Eukarya, Bacteriaand Archaea (Daiyasu et al., 2001; Gomes et al., 2002a): glyoxalases II,


which catalyse the hydrolysis of the thioester of S–D lactoglutathione toglutathione and lactic acid, arylsulfatases, DNA and RNA processingenzymes, among others. All these enzymes contain extra domains, besidesthe b-lactamase one. A total of 16 subfamilies have been proposed basedon extensive sequence and structural comparisons (Daiyasu et al., 2001),made for already classified enzymes. However, Daiyasu and co-workersindicated that a large number of still unassigned open reading frames inthe sequenced genomes are also probably members of this large proteinfamily. With the exception of the flavodiiron proteins, all other enzymesappear to contain a mono or di-zinc site, or, in the case of glyoxalases, amixed iron-zinc centre (Zang et al., 2001). The metal site is always located atone edge of the internal �b� sandwich; therefore, the larger aminoacidconservation occurs in aminoacid stretches that include the metal ligands(see Section 2.4). An analysis of the nature of the metal ligands suggestedthat very minor substitutions led from a di-zinc to a diiron site, passing bya mixed-metal centre, keeping the same structural fold. This changecorrelates with the activity of the enzymes, on going from a mainly Lewis-acid catalysis by the zinc site (in b-lactamases) to a redox active diiron site,capable of reducing dioxygen or nitric oxide (in the flavodiiron proteins).Altogether, these analyses point to a possible evolutionary link betweenthese proteins, being a paradigmatic example not only of divergentevolution but also of how nature can assemble diverse structural motifsand cofactors to achieve different functions. These enzymes are also a clearcase where simple extrapolation of fold to function leads to wrongassignments of enzymatic functions.

2.1.3. The Flavodoxin Domain

The flavodoxin domain – starting from residue Lys250 – is composed byan internal b-sheet flanked by �-helices on each side, displaying a typical�b� flavodoxin fold. The FMN cofactor is located on one edge of theb-sheet. Although in each monomer the xylene ring from the FMNisoalloxazine ring is pointing to the surface of the flavodoxin module,the dimeric conformation of the as-isolated functional protein ends upcovering the flavin moiety from bulk solvent exposure. The FMNisoalloxazine ring is parallel to the aromatic ring of Trp347. In comparisonto flavodoxins, the FMN pocket displays a greater preponderance ofbasic over acidic residues, which will be further discussed in the RedoxProperties section.


2.2. Modular Organization – Classification of the FlavodiironProteins

The flavodiiron proteins have in common the two-domain structural core.However, several members of this family have extra domains, fused at theC-terminal. Thus, according to the domain composition, these enzymescan be divided into three classes, A to C (Fig. 3):

– Class A, having only the two domain core, which so far includes thelargest number of members.

– Class B, which have a rubredoxin-like domain, containing an FeCys4binding motif, similar to those of Type I rubredoxins (two sets ofcysteines, with the spacing –CysXXCys-). The recombinant enzymefrom E. coli–K12 was indeed shown to contain an extra rubredoxin-type centre, and was named flavorubredoxin (FlRd) (Wasserfallenet al., 1998; Gomes et al., 2000). Genes encoding for Class B enzymesare so far found only in the genomes of enterobacteria (which containonly this type of flavodiiron proteins) and in Erwinia chrysanthemi(Okinaka et al., 2002).

– Class C, which have an additional module (of ca. 170 residues) withsignificant similarities to NAD(P)H : flavin oxidoreductases, andonly detected in cyanobacteria. The recombinant enzyme fromSynechocystis sp. PCC6803 was purified and contains two flavinmoieties, in agreement with this domain organisation (Vicente et al.,2002). Within this class, two subgroups can be distinguished, accord-ing to the conservation of the iron binding residues (see below).

Figure 3 Modular arrangement of the different classes of flavodiiron proteins. PanelA: Class A FDPs consist of the structural core of the family, being composed of anN-terminal metallo b-lactamase module and a C-terminal flavodoxin module. Panel B:Class B FDPs have an extra rubredoxin-like domain fused at the C-terminal of theflavodiiron core. Panel C: Class C FDPs bear a C-terminal NAD(P)H : flavinoxidoreductase module fused to the flavodiiron core.


A possible fourth class may exist, since an open reading frame in thegenome of Clostridium perfringens (Shimizu et al., 2002) appears to encodefor a flavodiiron protein, fused to a rubredoxin domain, followed by anextra NADH : oxidoreductase domain. To clarify this possible new classof enzymes, biochemical data on it is needed. With the growing availabilityof genome sequences, and with the mosaic nature of this protein family,it is probable that even more classes of these enzymes may come to ourknowledge.

2.3. Unity and Diversity of Electron Transfer Chains

The domain composition of these enzymes and, in part, the genomicorganisation of the respective genes, reflect the different types of electrontransfer chains operative in each organism. The studied chains have alwaysin common the oxidation of NAD(P)H coupled to the reduction of theterminal electron acceptor (NO or O2) (Fig. 4). Its variability is also reflectedin the nature of the physiological partners, either already established orhypothesised on the basis of the genomic data.

2.3.1. Class A Enzymes

Desulfovibrio gigas ROO has been shown to accept electrons directly fromthe one-electron reduced rubredoxin (Rd) partner (Chen et al., 1993b;Gomes et al., 1997), an electron transfer process which appears to begoverned mainly but not exclusively by electrostatics (Victor et al., 2003).The evidence for ROO and Rd to be redox partners was reinforced by thefact that the respective genes form a dicistronic transcriptional unit (Gomeset al., 1997; Frazao et al., 2000; Silva et al., 2001), a situation also observedin several other anaerobic bacteria. The rubredoxin, a small redox proteinof approximately 6 kDa with a Fe–S(Cys)4 centre, is in turn reduced byan NADH : rubredoxin oxidoreductase (NRO) (Chen et al., 1993a). Thiselectron transfer chain provided the first clear example of a function fora rubredoxin in anaerobes.In the genomes of Clostridia species, genes encoding for flavodiiron

proteins are present. For example, the genome of C. perfringens (Shimizuet al., 2002) has three loci coding for different flavodiiron proteins, oneof which is contiguous to a rubredoxin-encoding gene and transcribed inthe same direction. Interestingly, studies on Clostridia species revealedthe presence of rubredoxin reductases in these organisms (Petitdemange


et al., 1981). In fact, the gene encoding the NADH : rubredoxin oxido-reductase in Clostridium acetobutylicum was identified (Guedon andPetitdemange, 2001), and it is contiguous to one of the genes coding for aflavodiiron protein in this organism, and transcribed in the same direction

Figure 4 Schematic representation of the several electron transfer chains involving thedifferent classes of flavodiiron proteins (FDPs). NRO – D. gigas NADH : rubredoxinoxidoreductase; Rd – D. gigas rubredoxin; ROO – D. gigas FDP; Hrb – Moorellathermoacetica high molecular weight rubredoxin; Fe–S|Flv – putative Fe–S flavoprotein;FlRd – flavorubredoxin; FlRd-Red – flavorubredoxin reductase.


(Nolling et al., 2001). Altogether these observations suggest that inClostridia species the same type of electron transfer chain, as the oneobserved in D. gigas, is operative (Fig. 4, Scheme 1).In Moorella thermoacetica, the C-terminal modules from Class B

and Class C flavodiiron proteins, i.e. the Rd and NAD(P)H : flavinoxidoreductase modules, are fused together in a single polypeptide chain,which was named by the authors as high molecular weight rubredoxin(Hrb) (Das et al., 2001). The genes encoding for both Hrb and theflavodiiron protein are part of the same operon, which includes alsoa putative oxidative stress response protein, rubrerythrin. In the samelocus, but divergently transcribed, there is a dicistronic unit, encoding fora two-iron superoxide reductase (desulfoferrodoxin) and its putativeelectron donor, a rubredoxin (Das et al., 2001). Moreover, it was shownthat Hrb efficiently reduces the flavodiiron protein at the expense ofNADH oxidation (Silaghi-Dumitrescu et al., 2003). The Hrb NADH : flavinoxidoreductase module, which also displays a significant degree ofsimilarity towards Archaeoglobus fulgidus ferric reductase (Vicente et al.,2002), was proposed to accept electrons from NADH and to internallytransfer them to the Rd module, which acts as the donor site for theflavodiiron protein (Silaghi-Dumitrescu et al., 2003) (Fig. 4, Scheme 2).While in many genomes, the flavodiiron genes are in loci containing

genes that apparently do not encode for their physiological partners,analyses of other genomes suggest other possibilities in terms ofelectron transfer chains involving flavodiiron proteins. Such is the case ofA. fulgidus: this hyperthermophilic archaeon expresses two rubredoxins andhas several NADH oxidases as candidates to act as NADH : rubredoxinoxidoreductases (Abreu et al., 2000, 2002); nevertheless, the flavodiironprotein is encoded contiguously and in the same direction as the gene codingfor an Fe–S flavoprotein, exactly as observed in the Methanocaldococcusjanaschii genome (Bult et al., 1996). Moreover, in Methanosarcinaacetovorans (Galagan et al., 2002), this gene is present two genesdownstream of the gene coding for the flavodiiron protein. Theseobservations suggest the possible involvement of these uncharacterisedFe–S flavoproteins in electron transfer chains having flavodiiron proteinsas the terminal acceptors (Fig. 4, Scheme 3).

2.3.2. Class B Enzymes

In E. coli, the flavodiiron protein, flavorubredoxin, was shown to interactdirectly with an NADH-dependent (flavo)rubredoxin reductase (FlRd-Red)


(Gomes et al., 2000) (Fig. 4, Scheme 4). Recently, both were shown to forman operon (da Costa et al., 2003), under a complex regulation (see Section2.6.2). The same type of gene organisation was observed in allenterobacterial genomes so far sequenced (Blattner et al., 1997;McClelland et al., 2001; Parkhill et al., 2001; Perna et al., 2001). The directreduction of the terminal enzyme component FlRd by an NADH-dependentpartner was a result of an interesting evolutionary event, which fused theRd component to the flavodiiron structural core. The resulting effect on theelectron transfer efficiency remains a subject of on-going research.Preliminary studies determined that the flavorubredoxin reductase is a 43kDa monomeric NADH oxidase, containing one FAD, which efficientlyreduces FlRd (Gomes et al., 2000).

2.3.3. Class C Enzymes

An even more extreme module fusion is observed in the cyanobacterialflavodiiron proteins, both in the studied member of Synechocystis sp.PCC 6803 (Vicente et al., 2002) and the remainder multiple copies foundin all the cyanobacterial sequenced genomes (Kaneko et al., 1996, 2001;Nakamura et al., 2002). In this case, the multi-component electron transferchains are completely abolished, as the fusion of the NAD(P)H : flavinoxidoreductase module to the flavodiiron core allows the protein to acceptelectrons directly from NAD(P)H and perform several intra-molecularelectron transfer steps onto the diiron centre, which ultimately reduces thediatomic substrate (Fig. 4, Scheme 5).

2.4. Amino Acid Sequence Analysis

The amino acid sequences of all flavodiiron proteins were retrievedfrom the public databases, using either their actual assignment in eachgenome or searching the genomes with sequences from already charac-terised enzymes as queries. A total of 46 sequences were retrieved. Thesesequences were aligned using Clustal X (Thompson et al., 1997), followedby slight manual adjustments on GeneDoc; distinct alignments wereperformed – using the core lactamase and flavodoxin domains (excludingthe additional C-terminal domains from Class B and C enzymes), eachof these domains separately, and the additional domains, which werecompared among themselves and with related proteins from otherorganisms.


2.4.1. Core Domains

Orthologues of the flavodiiron proteins were found in all genomes ofanaerobic prokaryotes, as well as in several facultative anaerobes.Moreover, they are also present in the genomes of some anaerobicprotozoa, such as Giardia lamblia and Entamoeba histolytica (Anderssonet al., 2003). Interestingly, an extensive comparison of protozoan genesequences with prokaryotic ones, showed that among the 15 genes thatmost probably resulted from lateral gene transfers from anaerobicprokaryotes to the protozoa, are the flavodiiron protein ones, as well asthe flavohemoglobins and the Hybrid Cluster proteins (Andersson et al.,2003), enzymes that have been associated with defence mechanisms(although the function of the last one is yet not established). As alreadymentioned, several organisms contain genes encoding for more than oneputative flavodiiron protein, namely the cyanobacteria, which containmultiple distinct homologous genes. The overall aminoacid identities andsimilarities range from � 20 to � 70%, and � 40 to � 90%, respectively.Remarkably, the dendrograms obtained using the two-domain core(Fig. 5), or each domain per se are essentially identical (data not shown),strongly suggesting that the fusion of the lactamase-like domain with theflavodoxin one occurred only once, in an unidentified common ancestor,i.e. these enzymes appear to have a monophyletic origin. Furthermore, theflavodoxin domains are all more similar to each other than to singleflavodoxins. There is no clear distinction associated with the organismalphylogenies, since the sequences from quite diverse organisms are spreadall over the dendrogram. Nevertheless, the sequences from the proteinsbelonging to the classes B and C form subclusters, what suggests again amonophyletic origin within each of these subclasses, i.e. the fusion of therubredoxin or the flavin reductase domains seems also to have occurredonly once (it should be noted again that these comparisons wereperformed excluding these two extra domains). Also, it is worth notingthat so far no single b-lactamase domain homologous to those of theflavodiiron proteins, containing the ligands to the diiron site, could befound in the genomes.Among the ca. 400 residues of the core domains, only a few residues

are strictly, or almost strictly, conserved (Fig. 6). The first obvious onesare those binding the iron centre. According to the nature of these ligands,there are two types of flavodiiron proteins: those containing theligands found in the structure of the D. gigas enzyme (His79, Glu81,Asp83, His146, Asp165, His226), which include almost all members ofClass A (with the exception of one enzyme from M. jannaschii) and B


Figure 5 Sequence alignment of the 3 classes of Flavodiiron Proteins. The proteins aligned represent the different classes of FDPs:D_gigas – Desulfovibrio gigas ROO (Class A); Ec_FlRd – Escherichia coli FlRd (Class B); SynATF573 – Synechocystis sp. PCC6803FDP1 (Class C1); and Syn3 – Synechocystis sp. PCC6803 FDP3 (Class C2). Conservation is highlighted by the shading patterns,resulting from the alignment of a total of 46 FDP sequences, using Clustal X. The diiron ligands (based on the D. gigas ROO structure)are marked with (*); the Trp residue which is placed as an aromatic sandwich for the flavin moiety is marked with a (#). The secondarystructure from D. gigas ROO is illustrated with boxes and block arrows for �-helices and b-sheets, respectively (dark gray shadingrepresents the b-lactamase module and light gray represents the flavodoxin module). The highlighted conserved motifs are discussed inthe text.

98

LIG

IAM.SARAIVA,JOAO

B.VIC

ENTEAND

MIG

UELTEIXEIR

A

Figure 6 Dendrogram of the Flavodiiron Proteins family. The dendrogram wasgenerated with Clustal X and manipulated in TreeView V1.5. A total of 46 sequences fromFDP were aligned and the dendrogram was bootstrapped by excluding gap positions.Class B and C are grouped in two separate branches, which are inserted into boxes with adarker shade of gray. The FDPs which have already been characterised are displayed inwhite-shaded boxes. Abbreviations for the organisms: A_ful – Archaeoglobus fulgidus,C_tep – Chlorobium tepidum, C_act – Clostridium acetobutylicum, C_ per – Clostridiumperfringens, D_des – Desulfovibrio desulfuricans, D_gigas – Desulfovibrio gigas, D_vul –Desulfovibrio vulgaris, Entamoeba – Entamoeba histolytica, Ec_FlRd – Escherichia coliK12, uEc_FlRd – Escherichia coli O157 :H7, F_nuc – Fusobacterium nucleatum, Giardia –Giardia lamblia, M_thm – Methanothermobacter (M.) thermoautotrophicus, M_ jns –Methanocaldococcus janaschii, M_act – Methanosarcina acetivorans, M_maz –Methanosarcina mazei, Moorella – Moorella thermoacetica, Nostoc – Nostoc sp.PCC6120, P_aby – Pyrococcus abyssii, P_ fur – Pyrococcus furiosus, P_hrk –Pyrococcus horikoshii, R. capsulatus – Rhodobacter capsulatus, S_ent – Salmonellaenterobacter, S_typ – Salmonella typhimurium, Synechocystis (Syn) – Synechocystis sp.PCC6803, T_tgc – Thermoanaerobacter tengcongensis, T_elg – Thermosynechococcuselongatus, T_mar – Thermotoga maritima, T_pal – Treponema palidum. Numbers adja-cent to each abbreviation refers to multiple different FDPs in each organism.


enzymes, and some of the Class C enzymes (Class C1). Within the Class Cproteins, three from Nostoc sp. PCC7120, two from Synechoscystis and onefrom Thermosynechococcus elongatus, have substitutions of Glu81 by anasparagine, His146 by an arginine, and Asp165 by a lysine (but an aspartateis present immediately before), which we classify as Class C2 enzymes; onlywhen their structures become available, will it be clear if these aminoacidsare indeed ligands to the iron ions. The other almost strictly conservedresidues occur in several aminoacid segments (Fig. 6): motifs II, III, IV andV contain the iron ligands and, remarkably, are those that have beenshown by Daiyasu et al. (2001) to be the most conserved aminoacid stretchesamong the 16 enzyme families which share the metallo b-lactamase-like fold.Motif I, located close to the N-terminus and forming a b-sheet locatedabove the diiron centre, is highly conserved only in the flavodiiron proteins;however, analyses of the D. gigas structure did not give yet any clue of itspossible role. Finally, motif VI, the only one in the flavodoxin-like domain,is involved in the binding pocket of the FMN moiety. Motifs I, II and VIhad been previously proposed as typical of the flavodiiron proteins (Gomeset al., 1999).

2.4.2. Class B Rubredoxin Domain

The sequence of the Rd-domain of Class B enzymes (the flavorubredoxins)was aligned with the sequences of type I rubredoxins from different sources,including the eukaryotic examples from Guillardia theta and Plasmodiumyoelii, and also other Rd-domains from larger proteins (not shown).Aminoacid identities range between 31 and 62%, and similarities between50 and 83%; the Pseudomonads Rds display the highest degree of identitywith the Rd-domains. Among the Class B domains, the identities are veryhigh (85–100%), as well as the similarities (96–100%). The residues knownto stabilise the overall structure (Tyr/Phe/Trp4, Tyr11, Tyr13, Phe30, Ile/Leu/Val33, Trp37 and Phe49, in Clostridium pasteurianum Rd numbering)are also highly conserved (Meyer and Moulis, 2001). Relevant residueswhich modulate the redox potential of the Rd-centres have been the subjectof many studies and will be further discussed in the Redox Propertiessection.

2.4.3. Class C NAD(P)H : flavin Oxidoreductase Domain

The presence of the NAD(P)H : flavin oxidoreductase (NFOR) domain isuniquely found among the cyanobacterial FDPs (Class C). This domain


shares similarity with flavin reductases and ferric reductases, as well aswith small components of larger enzymes, such as several oxidoreductasesand monooxygenases. Structurally, they are part of a large family(Pfam entry PF01613) which includes a significant number of the abovementioned enzymes. Within themselves, the NFOR domains of Class Cenzymes share identities ranging from 34 to 78% and similarities between55 and 89%. A sequence alignment between the NFOR domains and someof the protein sequences of members of the above mentioned family (notshown) shows that the NFOR domains are much more similar to oneanother than to other enzymes of the family. This is illustrated by thelow identities (15–20%) and low similarities (31–40%) shared betweenthe Class C flavodiiron NFOR domains, and other members of thePfam entry PF01613 family. However, a slightly higher degree of similar-ity is observed between those domains and the NFOR module of theM. thermoacetica Hrb (identity up to 27% and similarity up to 44%).The NFOR domain of the Synechocystis SsATF573 (Vicente et al., 2002)was structurally aligned with the A. fulgidus ferric reductase (24% identitytowards NFOR, PDB entry: 1ios). In this protein, most of the interactionswith the FMN cofactor are established with the main chain nitrogen andoxygen atoms. Interestingly, the corresponding regions identified in theNFOR domain of SsATF573 as being within a 6 A radius of the FMNcofactor are highly conserved within the NFOR domains of the Class Cflavodiiron proteins.

2.5. Physicochemical Properties

Only a few flavodiiron proteins have so far been isolated (either fromwild-type organisms or heterologously expressed) and characterised: theenzymes from D. gigas, R. capsulatus, M. thermoacetica, E. coli,M. thermoautotrophicus and Synechocystis sp. PCC6803. Some of thebasic physicochemical properties reported for these proteins are summa-rised in Table 1, such as the aminoacid length, molecular mass andquaternary structure, and the content on redox centres5 (including the

5D. gigas rubredoxin : oxygen oxidoreductase was isolated from wild type cells containing

two distinct hemes (Chen et al., 1993b): Fe-uroporphyrin I and a modified C-type heme,

together with an as yet unidentified chromophore, with an absorption maximum at 580 nm.

However, it was later realised that the heme content was always substoichiometric and the

enzyme crystal structure did not reveal the presence of the hemes. Thus, its physiological

relevance remains an open question, as all other flavodiiron proteins that were isolated from

their natural hosts lack any heme centre.


Table 1 Biochemical properties of flavodiiron proteins.

Protein Microorganism a.alength

Monomermolecularmassa

Quaternarystructure

Cofactor contentb Reference

Rubredoxin:oxygenoxidoreductase(ROO)

D. gigas 402 43 kDa (44.8) Homodimer 2 Fe/monomer (XRC)1 FMN/monomer (XRC)

Chen et al. (1993b),Frazao et al. (2000)

Flavorubredoxin(FlRd)

E. coli 479 54 kDa (54.2) Homodimer 2.9� 0.5 Fe/monomer(C)

Gomes et al. (2000)

1 FMN/monomer(AE-UVS)

SsATF573 Synechocystis 573 70 kDa (63.5) Homodimer 1.9 Fe/monomer (C) Vicente et al. (2002)0.8 FMN/monomer(AE-HPLC)

Flavodiiron protein M. thermoacetica 399 45 kDa (44.3) Homodimer 3.8� 0.5 Fe/dimer(PAEA)

Silaghi-Dumitrescuet al. (2003)

1.7� 0.1 FMN/dimerFlavoprotein A(FprA)

R. capsulatus 420 48 kDa (46.2) Homodimer 0.9 FMN/monomer(AE-HPLC)

Jouanneau et al. (2000),Wasserfallen et al. (1998)

Flavoprotein A(FprA)

M. thermoautotrophicumstrain �H

409 45 kDa (46.0) Homodimer 1.3 FMN/monomer(AE-HPLC)

Nolling et al. (1995)

1 mol Fe/mol FMN (C)Flavoprotein A(FprA)

M. thermoautotrophicumMarburg

404 43 kDa (45.7) Homotetramer 0.7 FMN/monomer(AE-HPLC)

Wasserfallen et al. (1995)

aBetween brackets, molecular mass estimated from the aminoacid sequence.bBetween brackets, experimental methodology by which the cofactor was identified and quantified: XRC, X-ray crystallography; PAEA, plasma atomic emission analysis;

AE-UVS, acid extraction plus visible spectroscopy; AE-HPLC, acid extraction followed by HPLC analysis; C, colorimetric method.

methods by which they were identified/quantified): as mentioned above, theflavodiiron proteins have in the core domain a diiron centre and an FMNmoiety; the class B and C enzymes contain extra domains, harbouring arubredoxin-like centre or another flavin, respectively. The Class A FDPs,which correspond to the flavodiiron structural core, have a length ofapproximately 400 aminoacids, and thus a molecular mass around 45 kDa;Class B and Class C FDPs have higher molecular masses – around 54 kDaand 65 kDa, respectively – due to the extra C-terminal modules. With theexception of the M. thermoautotrophicus Marburg protein, the quaternarystructure of FDPs is that of a homodimer. As already described, thecrystallographic structure of D. gigas ROO suggests the need for a head-to-tail homodimeric quaternary structure to place the FMN and diiron centresfrom the opposing monomers within efficient electron transfer distance.

2.5.1. Spectroscopic Studies

2.5.1.1. Absorption spectroscopy. The electronic spectra in the Visibleand near UV region of the majority of the flavodiiron proteinsare dominated by the flavin moiety, namely the FMN bound to theflavodoxin module (Fig. 7). As observed for non-heme diiron proteins

Figure 7 Structural modelling of the FMN pocket. The structures of various FDPthat have already been characterised were modelled in SwissModel, with D. gigas ROOstructure (PDB entry 1e5d) as the template. Here, the FMN pocket is highlighted for theD. gigas ROO, stressing the superposition of the Trp347 aromatic ring over thepyrimidine ring of the FMN isoalloxazine core. The FMN pockets of the other modelledFDPs show that with the exception of the two different M. thermoautotrophicus enzymes,the conserved Trp residue is in the same position as for the ROO structure. (See colourplate section.)


(Solomon et al., 2000), and recently proposed for the M. thermoaceticaflavodiiron (Silaghi-Dumitrescu et al., 2003), the absorptivity of the diironcentre is much lower than that of the flavin cofactors (Ghisla and Edmonson,2001). In the case of this protein, the authors propose a putative diironspectrum, based on the subtraction of the Zn-loaded protein spectrum fromthat of the Fe-loaded one (Silaghi-Dumitrescu et al., 2003). Nevertheless,the authors stress that the resulting differences could be due to a flavinperturbation by the bound zinc.The predominance of the flavin cofactors in the spectra of Class A

flavodiiron proteins, is well documented by the spectrum of a truncatedversion of E. coli FlRd lacking the C-terminal rubredoxin module, thusconsisting only of the flavodiiron structural core (Gomes et al., 2000)(Fig. 7). The shape of the spectrum and its absorption maxima (at 455 nmand 368 nm) are almost identical to the spectra recorded for otherflavodiiron proteins such as the M. thermoacetica (Silaghi-Dumitrescuet al., 2003) and R. capsulatus ones (Jouanneau et al., 2000). The broadnessof the typical flavin bands (Ghisla and Edmonson, 2001) contrasts with thespectra of the flavodiiron proteins from M. thermoautotrophicus (Nollinget al., 1995; Wasserfallen et al., 1995) and Synechocystis sp. PCC6803(Vicente et al., 2002) (Fig. 7), which have a broad band around 370 nm anda shouldered band at 455 nm. In both types of spectra, the band atapproximately 450 nm is commonly assigned to charge transfer transitionsfrom the xylene ring to the pyrimidine ring, in the isoalloxazine coreof the flavin cofactor. In the case of the spectra observed for theM. thermoautotrophicus and Synechocystis flavodiiron proteins, this higherresolution around the 450-nm band was proposed to be related with ahydrophobic environment for the flavin moiety (Ghisla and Edmonson,2001).Structural models based on the D. gigas enzyme structure were

obtained for E. coli, M. thermoacetica, R. capsulatus and M. thermoauto-trophicus flavodiiron proteins (Fig. 8). Although the calculated electro-static surface of the FMN pocket is homogeneously hydrophobic, thereis a clear difference between the M. thermoautotrophicus flavodiironproteins and the remainder: the absence of the Trp residue, which isco-planar with the isoaloxazine ring of the FMN. We thus propose thatthe Trp sandwich to the isoaloxazine ring could create a number of micro-states, ultimately contributing to the broadness in the 450 nm CT transi-tion band observed in the E. coli, M. thermoacetica and R. capsulatusflavodiiron proteins.The spectra of Class B flavodiiron proteins display extra features,

resulting from the combination of the flavin cofactor features with the ones


Figure 8 UV–visible spectra of flavodiiron proteins. Panel A, line a – spectrum of atruncated form of E. coli FlRd lacking the C-terminal domain (FlRd�Rd), thus resultingin a Class A-like FDP; line b – spectrum of the red semiquinone radical from FlRd�Rd,upon one electron reduction of the flavin moiety. Panel B – spectrum of E. coliflavorubredoxin, a Class B flavodiiron protein. Panel C – spectrum of Synechocystis sp.PCC6803 FDP1, a Class C enzyme. All spectra recorded at room temperature, typicallyin TrisHCl 50 mM buffer, pH 7.6.


from the Fe-Cys4 centre in the C-terminal rubredoxin module. Rubredoxinshave unique spectral features: two major bands at approximately 380 nmand 490 nm, and a low absorption broad band around 570 nm (Meyer andMoulis, 2001). Thus, the 455 nm flavin band displayed for the E. coliflavorubredoxin flavodiiron core (Fig. 7) is shifted to 474 nm, the flat bandat 368 nm is split in two shoulders at 353 nm and 375 nm, and an extrabroad band appears at around 570 nm.Class C proteins bear a second flavin cofactor, at the C-terminal

NAD(P)H:flavin oxidoreductase (NFO) module. The spectrum ofSynechocystis sp. flavodiiron protein, which was used to illustrate theheterogeneity in the flavin-dominated spectra of the flavodiiron proteins,is in fact a combination of the two flavin moieties (Vicente et al., 2002).Interestingly, the spectrum of a truncated version of Synechocystis sp.flavodiiron protein, consisting only of the NFO module, has exactly thesame features as the intact protein (Vicente et al., 2002). On the otherhand, the studied flavodiiron protein from Synechocystis lacks the key Trpresidue, which is proposed to sandwich the FMN cofactor in the flavodoxinmodule, and thus to influence its spectral features.

2.5.1.2. EPR spectroscopy. In the flavodiiron proteins family, the firstreported EPR studies were on the D. gigas enzyme (Gomes et al., 1997). Thespectra of the oxidised enzyme were dominated by the heme centres, andsuggested a coordination of the heme iron to thiolate ligands. Underreductive conditions, a radical signal at g� 2.0 was attributed to theone-electron reduced semi-quinone form of the flavin, which was deter-mined to be a red anionic radical, from the 1.6 mT line width. An iden-tical signal was observed for the R. capsulatus flavodiiron protein(Jouanneau et al., 2000), where the radical was trapped in an attempt toreductively titrate the flavin cofactor.Escherichia coli flavorubredoxin is the most thoroughly studied fla-

vodiiron protein, for which all the three cofactors were probed by EPRspectroscopy (Fig. 9). The spectrum of native FlRd was consistent with theFe–Cys4 centre present in the rubredoxin module (Gomes et al., 2000).Two different sets of resonances were observed, attributed to two slightlydistinct conformations of the Rd centre: resonances at g� 9.3 and at g¼ 4.8and 4.3, corresponding to a high-spin (S¼ 5/2) ferric site with E/D� 0.3(gmax¼ 9.6, |� 1/2i doublet, gmed¼ 4.30, |� 3/2i doublet) and withE/D¼ 0.24 (gmax¼ 9.35, |� 1/2i doublet, gmed¼ 4.8, |� 3/2i doublet). Thesame radical signal at g� 2.0, present in the EPR spectra of the D. gigasand R. capsulatus flavodiiron proteins (Gomes et al., 1997; Jouanneauet al., 2000), was observed in the FlRd EPR spectrum and assigned to


the FMN cofactor. In the first reductive experiments, using dithionite asthe reducing agent, the only observation was the disappearance of theabove mentioned features, as the Rd centre has an integer spin whenreduced. In these preliminary experiments, no signature for a diironcentre was detected in the EPR spectrum of native (oxidised) or dithio-nite reduced FlRd, as expected for an antiferromagnetically spin-coupledsystem. In recent experiments, the signal from the mixed-valence FeIII–FeII species was detected, with g values at 1.95, 1.80 and 1.78 (Fig. 9)(our own unpublished data), similar to the spectra of the diiron centre

Figure 9 EPR spectra of E. coli flavorubredoxin. Panel A – EPR spectrum of theas-isolated (oxidised) flavorubredoxin from E. coli at 10 K. Microwave power, 2.4 mW;microwave frequency, 9.64 GHz; modulation amplitude, 1 mT; protein concentration� 170 mM, buffer: 50 mM Tris–HCl pH 7.6. Panel B – EPR spectrum of the mixed-valencenon-heme diiron centre of E. coli flavorubredoxin at 7 K. Microwave power, 2.4 mW;microwave frequency, 9.64 GHz; modulation amplitude, 1 mT; protein concentration� 560 mM; buffer: 10 mM Tris–HCl pH 7.6.


from the hydroxylase component of methane monooxygenase (Fox andLipscomb, 1988). Upon full reduction to the FeII–FeII species, the rhombicsignal disappears; a resonance at g� 12 (not shown) is detected usingparallel-mode EPR, indicating an integer spin state for the fully reducedcentre.

2.5.1.3. Mossbauer spectroscopy. Silaghi-Dumitrescu et al. (2003) haveused Mossbauer spectroscopy to study the properties of the diiron centrein M. thermoacetica FDP. The analysis of the data for the 57Fe-enrichedrecombinant protein showed that in the oxidised diferric state the iron ionswere anti-ferromagnetically coupled, yielding a diamagnetic ground state.The different quadrupole splittings and asymmetry parameters for thetwo quadrupole doublets used to fit the data, are assigned to two differentiron environments of the centre, consistent with the coordination sphere ofD. gigas ROO diiron centre (Frazao et al., 2000).

2.5.2. Redox Properties

A summary of the redox properties for the flavodiiron protein family ispresented in Table 2. Most studies on the redox cofactors have focusedon the FMN moiety, due to its prevalent spectroscopic features (discussedabove).

2.5.2.1. Redox properties of the FMN cofactor in the flavodoxinmodule. With the exception of the M. thermoautotrophicus protein, thefirst redox transition of the FMN – from the oxidised state to thesemiquinone one – occurs in a relatively narrow range of potentials(from �140 mV to þ 20 mV). This range is within that observed both forshort-chain and long-chain flavodoxins, (�229 mV to þ 121 mV) (Paulsenet al., 1990). However, one major difference between canonical flavo-doxins and FDPs is the type of semiquinone radical formed uponone-electron reduction of the flavin cofactor. In the vast majority offlavodoxins, if not all, it is a neutral blue radical that is formed, whereasin the flavodoxin module of the FDP family it is a negative red radicalwhich appears upon one-electron reduction. Both types of radicals havequite distinct features in the UV–visible spectrum, allowing its identifi-cation solely by spectral deconvolution. An increased absorbance atapproximately 380 nm and a shift of the 450 nm band to approximately500 nm, with respect to the oxidised form, are characteristic of redsemiquinones, in which the unpaired electron is located at N5 (whichis deprotonated) and the negative charge lies within N1 (Ghisla and


Table 2 Redox properties of flavodiiron proteins.

Protein Organism Redoxcofactors

Redox transitions E� (mV)(vs. SHE)

Method References

FlRd E. coli Rd [Fe–Cys4] [FeIII–Cys4]/[FeII–Cys4] �140 EPR/UV–vis Gomes et al. (2000)FMN FMNox/FMNsq (red sq.) �140 UV–vis Gomes et al. (2000)

FMNsq/FMNred �180Fe–Fe FeIII–FeIII/FeIII–FeII �25 EPR Our own unpublished

dataFeIII–FeII/FeII–FeII �105

FlRd�Rd E. coli FMN FMNox/FMNsq (red sq.) �60 UV–vis Gomes et al. (2000)FMNsq/FMNred �190 Gomes et al. (2000)

Fe–Fe FeIII–FeIII/FeIII–FeII 0 EPR Our own unpublisheddata

FeIII–FeII/FeII–FeII �50FlRd�FD E. coli Rd [Fe–Cys4] [FeIII–Cys4]/[FeII–Cys4] �95 UV–vis Our own unpublished

dataROO D. gigas FMN FMNox/FMNsq (red sq.) 0 UV–vis Gomes et al. (1997)

FMNsq/FMNred �130FDP Moorella FMN FMNox/FMNsq (red sq.) �117 UV–vis Silaghi-Dumitrescu

et al. (2003)FMNsq/FMNred �220

FprA R. capsulatus FMN FMNox/FMNsq (red sq.) þ 20 UV–vis Jouanneau et al. (2000)FprA M. thermoauto-

trophicumFMN FMNox/FMNred þ 230 UV–vis Nolling et al. (1995)

FLAVODIIR

ON

PROTEIN

SIN

NO

DETOXIFIC

ATIO

N109

Edmonson, 2001). The formation and disappearance of the semiquinoneradical can be followed by monitoring the changes of the visible absorbanceat the appropriate wavelength as a function of the redox potential, whichshould yield a bell-shape curve. This is illustrated in Fig. 10, whichrepresents the redox behaviour of the different redox cofactors in E. coliFlavorubredoxin.The redox properties of the FMN cofactor in flavodoxins has been

the subject of intensive studies, mainly on the proteins fromClostridium beijerinckii, Desulfovibrio vulgaris and Nostoc sp. PCC7120(Paulsen et al., 1990; O’Farrell et al., 1998; Hoover et al., 1999; Kasim andSwenson, 2000). For flavodoxins, it is proposed that upon one electronreduction of the FMN, a conformational rearrangement occurs, bringinga backbone carbonyl close to the NH(5) position, thus stabilising theneutral radical by the formation of a hydrogen bond. It is also suggestedthat a glycine is the most suitable residue for this position, since it hasno side chain and thus does not offer any steric hindrance for the

Figure 10 Redox properties of E. coli flavorubredoxin. The solid lines were calcu-lated using Nernst equations with the reduction potentials presented in Table 2. Linea – rubredoxin centre; Line b – diiron centre (mixed valence state); Line c – FMN centre(semiquinone sate).


conformational change to take place. In fact, studies on the Anacystisnidulans flavodoxin, which has an Asn residue in this position, showed ahigher reduction potential for the first redox transition than that offlavodoxins having a Gly residue in the same position (Hoover et al., 1999).This suggests that indeed the semiquinone radical is more stable whena Gly residue is present. The structure of D. gigas ROO reveals a numberof hydrogen bonds between aminoacid side chains and the FMNcofactor (Frazao et al., 2000), namely in key redox atoms of theisoalloxazine ring. In this protein, the residue Asn315 donates a hydrogenbond to N(5) in the oxidised state. If this bond should hold after theone electron reduction to the semiquinone state, it is difficult to considera conformational rearrangement such as it is observed in canonicalflavodoxins. Furthermore, as a red semiquinone is formed in the FDPs,N(5) is deprotonated and thus unable to form a hydrogen bond with abackbone carbonyl as described for the flavodoxins neutral semiquinoneradical. The fact that a red radical is formed in FDPs may be related toa higher prevalence of basic over acidic aminoacid residues in the FMNpocket, as observed in D. gigas ROO structure. The pKa of 8.3 for theequilibrium between the red and blue semiquinone forms of free FMN(Ghisla and Edmonson, 2001) could be lowered in the FDPs due to thepresence of the excess basic residues, and thus result in the formation ofa red semiquinone, in the pH range at which redox studies were performedfor FDPs (pH 7.0 to 8.0). Altogether, these observations may justify theformation of a red instead of a blue semiquinone radical, and account forthe lower stability of the FDPs radical in comparison to the flavodoxins.This lower stability is accounted for by the much higher reduction poten-tial for the second transition, from the semiquinone to the hydroquinoneform (Table 2). In flavodoxins, this transition occurs within the range of�372 mV down to �522 mV (Paulsen et al., 1990), whereas in the FDPs therange goes from �130 mV to �220 mV. The transient character of thesemiquinone radical in FDPs may also arise from an interaction betweenthe FMN cofactor and the diiron centre from the other monomer(in each functional homodimer, see Section 2.1). In fact, the diiron ligandGlu81 is in contact with the methyl group in the C8 position ofthe isoalloxazine ring, a chemical group in which the protons are muchmore labile for exchange than what is expected for a methyl group(Ghisla and Edmonson, 2001). In E. coli flavorubredoxin, this type ofinteraction can also occur with the rubredoxin domain, since engineeredtruncated domains have different redox properties from the ones observedin the whole protein (Table 2).


2.5.2.2. The diiron centre in the �-lactamase module of E. coliflavorubredoxin. As described in the Spectroscopy section, the diironcentre is practically silent in UV–visible spectra, in all its oxidationstates, and for EPR spectroscopy in its oxidised state. Recently we haveperformed EPR studies on the diiron centre of E. coli FlRd. The distinctiverhombic signal corresponding to the mixed-valence FeIII–FeII centre (seeSpectroscopy section) appears upon one-electron reduction of the diferriccentre and disappears upon the full two-electron reduction. We were thuscapable of determining the reduction potentials of the centre (Table 2),both for the whole protein and for a truncated version consisting only ofthe flavodiiron core. The redox potentials are well within what was observedfor the hydroxylase component of methane monooxygenase, a non-hemediiron containing protein, whose redox properties have been thoroughlystudied (Liu and Lippard, 1991; Paulsen et al., 1994). The difference inpotential between the intact and truncated proteins may result from a highersolvent accessibility of the centre, or from a plausible interaction betweenthe different redox cofactors.

2.5.2.3. Redox properties of the C-terminal Rd-domain in Class B FD-NOR. The reduction potential for the one-electron reduction of the Rdcentre in the intact FlRd is �140 mV (Table 2). This is the lowest redoxpotential described for this type of centre, which may be explained bydifferences in key residues close to the ligating cysteines, that result in acumulative decrease of the reduction potential with respect to other Rdcentres (Gomes et al., 2000). For instance, in C. pasteurianum Rd, a Gly-to-Glu mutation in the widely conserved residue next to the second ligatingCys (Cys9–Gly10, in C. pasteurianum Rd) results in a –35 mV decrease inthe redox potential (Kummerle et al., 1997). This residue is replaced by aGln in FlRd’s Rd-domain, which is also expected to yield a decrease inpotential. The highly conserved Gly next to the fourth ligating Cys and theconsecutive Val have also been studied in terms of redox potentialmodulation (Meyer and Moulis, 2001); interestingly, these residues arereplaced in the Rd-domain by a serine and a lysine respectively bothproven to once again lead to a decrease in the redox potential. Altogether,these aminoacid changes contribute to the low redox potential of theRd centre in FlRd. Nevertheless, by studying a truncated version of FlRdconsisting of the Rd domain alone, an apparent reductionpotential of �95 mV is observed (our own unpublished data), whichindicates that other factors are involved in the modulation of this reduc-tion potential, i.e. the overall structure of FlRd plays a role in controllingthe Rd centre reduction potential, possibly through the interaction of


the Rd centre with the flavin moiety in the flavodoxin module. Concerningthe recently proposed topological arrangement for FlRd (Victor et al.,2003), which places the Rd centre near the flavin moiety, we may againsuggest that a possible interaction between both cofactors could yieldchanges in their redox properties. This proposal is consistent withthe different redox properties of the flavin moiety, observed in intactFlRd and a truncated version lacking the Rd-domain (discussedabove in this section). On the other hand, we cannot exclude the fact thatthese shifts could simply be a matter of solvent accessibility. In fact, theRd centre should be stabilised by solvent exposure more in its reducedstate (global charge of the centre is �2) than in the oxidised state (globalcharge �1).The overall redox properties of the E. coli flavorubredoxin and of

its partner are summarised in Fig. 11. As for the D. gigas system (Gomeset al., 1997), the redox groups have very close apparent reductionpotentials, which are arranged so that the electrons flow into the catalyticcentre. Due to the spatial proximity of the several centres, electrostaticinteractions between them certainly exist, which affect the intrinsicreduction potentials of each one. Furthermore, it should be stressed thatthese potentials were determined by equilibrium titrations; during turn-over, and particularly upon binding of the substrate, the actual potentialswill be certainly different. Also, so far there are no data regarding thepossible pH dependence of the reduction potentials; since reduction ofeither NO or O2 involve protons, such a dependence is to be expected.

Figure 11 Redox potentials associated with the different electron transfer processeswithin the NADH to NO electron transfer chain in E. coli involving flavorubredoxin.


2.6. Function of Flavodiiron Proteins

2.6.1. Enzymatic Studies

The first function assigned to a flavodiiron protein was that of oxygenreduction to water, for the enzyme from D. gigas. The same function waslater reported for the enzymes from E. coli and Synechoscystis. However,exact values for the turnover of this reaction remain to be determined.Already in 1998 it was proposed that E. coli possessed a nitric oxidereductase activity, when grown anaerobically, which was insensitive tocyanide, thus excluding the flavohemoglobin as the enzyme responsiblefor that activity (Gardner et al., 1998a). Later on, Gardner et al. (2002)showed, based on several molecular genetics studies (reviewed furtherbelow), and on the fact that the diiron site of E. coli flavorubredoxin wasable to bind NO (Gomes et al., 2000), that flavorubredoxin was anNO reductase. Subsequently, using the recombinant enzyme from E. coli,which is stable under aerobic conditions, we showed that this enzyme hasindeed a significant NO reductase activity, with a turnover number of15–20 s�1 (Gomes et al., 2002b). This value is similar to those of themembrane-bound heme-iron NORs (in the range of 10–50 s�1 (Zumft, 1997;Hendriks et al., 1998)), and considerably higher than those reported forthe flavohemoglobins (ca. two orders of magnitude). The NO reductaseactivity of E. coli FlRd was studied by measuring amperometrically NOconsumption in the presence of different concentrations of the physio-logical partner, FlRd-reductase, and at saturating NADH concentrations(>200 mM). A representative trace is presented in Fig. 12. The activity waslinearly dependent on the flavorubredoxin concentration, and is essentiallyindependent of NO in the concentration range from � 1 mM to � 10 mM.This shows that the enzyme has a high affinity for NO (KM<1 mM).Furthermore, it was shown that the truncated rubredoxin domain hasno activity, and that the presence of this domain is essential for electrondonation from the NADH : oxidoreductase to the flavorubredoxin. Also,and in agreement with the initial findings of Gardner and co-workers(Gardner et al., 1998a, 2002), the NO reductase activity is not inhibitedby cyanide, at least up to 3 mM cyanide (Gomes et al., 2002b).More recently, the recombinant enzyme from M. thermoacetica, was

also shown to reduce NO, with a turnover of � 50 s�1, again in thepresence of NADH and its physiological partner, the high molecularweight rubredoxin, Hrb (Silaghi-Dumitrescu et al., 2003). When expressedin an E. coli flrd minus strain, this protein eliminated the NO growthsensitivity of that E. coli strain (Silaghi-Dumitrescu et al., 2003).


The product of NO reduction by FDPs has not been demonstrated yet.Nevertheless, the NADH/NO stoichiometry and the reaction kineticsstrongly suggest that the product will be N2O (Silaghi-Dumitrescu et al.,2003). Although this gas is essentially harmless, and may be eliminatedthrough diffusion, it is also possible that an unknown N2O reductase ispresent in these organisms.A major question regarding these enzymes is whether the initially

proposed activity as oxygen reductases is physiologically relevant. Theaffinity for oxygen is much lower than that for NO (Gomes et al., 2002b;Silaghi-Dumitrescu et al., 2003), indicating that nitric oxide is the pre-ferential substrate. But, as discussed above for the (flavo)hemoglobins,it may also be that these enzymes are bifunctional, being active both inoxidative or nitrosative stress conditions. In fact, binuclear iron centres inmultiple enzymes are known to reduce oxygen to water, when the enzymaticreaction is decoupled from the substrate, and bind nitric oxide, which is

Figure 12 NO reductase activity of E. coli flavorubredoxin. In anaerobic conditions,four aliquots of NO (yielding a final concentration of 5 mM) were sequentially addedto 3 mM FlRd-red, in 50 mM Tris–HCl, 20% glycerol, pH 7.6. Following the additionof 1 mM NADH, 22 nM of FlRd were added, yielding a fast consumption of NO,whose time course followed zero-order kinetics. To ensure anaerobicity, glucose oxidase(4 units/ml), glucose (3 mM) and catalase (130 uints/ml) were added to the reactionmixture. The assay was performed by following NO consumption with a specific NOelectrode (from World Precision Instruments, Inc.), consisting of a 2mm NO probeconnected to an amplifier (Duo-18).


usually used as a probe for diiron centres. This situation finds also a parallelin the heme-iron NO reductases and the heme-copper oxygen reductases,which although having a much higher affinity for their natural substrate,nitric oxide or oxygen respectively, are nevertheless capable of reactingwith oxygen or nitric oxide, albeit with a much lower turnover (Brunoriet al., 1999; Brunori, 2001; Forte et al., 2001; Sarti et al., 2003).Furthermore, for E. coli, there is no indication that they respond tooxidative stress conditions, e.g. they are not under the regulation of theSoxRS or OxyR regulons. On the contrary, as discussed below, the levelof expression of E. coli FlRd under aerobic conditions is very low. On theother hand, the enzymes from Synechocystis sp. PCC6803 were recentlyproposed to be essential for oxygen photoreduction (Helman et al., 2003).This subject must wait further detailed studies.

2.6.2. Molecular Genetics Studies

The location of the E. coli flrd gene near a gene product (ygaA, alsodesignated as norR) that shares a significant identity to the NO regulator ofRalstonia eutropha (NorR) (Ramseier et al., 1994), and the fact that therecombinant E. coli flavorubredoxin could bind NO (Gomes et al., 2000),firstly suggested its involvement in NO metabolism. Confirmation camefrom the work of Gardner et al. (2002) who showed that an anaerobi-cally grown strain of E. coli deficient in flrd, and exposed to � 2 mM NO,had almost no NO consumption activity. Furthermore, anaerobic growthof the E. coli minus flrd strain, in metabolic conditions under which it isdependent on the function of NO-sensitive enzymes, was significantlyimpaired upon exposure to NO (� 0.5 mM). In particular, E. coli flavo-rubredoxin was shown to be able to protect the NO-sensitive aconitase(Gardner et al., 2002). As described in the previous section, it was laterproven that the recombinant E. coli FlRd is indeed an NO reductase.Furthermore, FlRd is reduced by the FAD-containing NADH oxido-reductase; it was also proven that the two genes encoding for theflavorubredoxin and its reductase form a dicistronic transcriptional unit(da Costa et al., 2003). In agreement with these results, Spiro and co-workers reported that anaerobically grown strains of E. coli defective inflrd/flrd-red were inhibited by 75 mM nitroprusside, while the parentstrain could support growth up to 5 mM nitroprusside (Hutchings et al.,2002).The presence of NO seems to be required for flrd expression, since in

its absence no significant anaerobic NO consumption was observed in


E. coli (Gardner et al., 2002). Also, NO (via nitroprusside) was found toinduce a �( flrd-lacZ) fusion, but activation was completely dependent onthe products of norR or flrd/flrd-red (Hutchings et al., 2002). On the otherhand, it was reported that the E. coli flrd minus strain, carrying a flrd-lacZfusion, only showed b-galactosidase activity when exposed to NO (Gardneret al., 2003). However, the induction only occurred in a limited rangeof NO concentration (between 0.25 mM and 1 mM NO) since NOconcentrations higher than 2 mM led to a decrease in the expression ofthe flrd-lacZ fusion, an observation that was attributed to NO toxicity.However, in D. gigas and M. thermoautotrophicus a significant amountof flavodiiron protein was isolated from these organisms cultured undertheir usual growth conditions, i.e. in the absence of NO. (Chen et al.,1993b; Nolling et al., 1995; Wasserfallen et al., 1995; Gomes et al., 1997;Silva et al., 2001).The E. coli flavorubredoxin mRNA level was measured under different

growth conditions, and the transcription level also increased in anaero-bically grown cells of E. coli submitted to high NO concentrations(>50 mM) (da Costa et al., 2003). In addition, oxygen, nitrate and nitritecontrol the expression of E. coli FlRd. Upon exposure to O2 the �( flrd-lacZ) fusion was repressed (Gardner et al., 2002, 2003). We have shownthat oxygen regulation occurs through the global transcription regulatorFNR since deletion of fnr causes an increase in the transcriptional andprotein levels, meaning that FNR acts as a repressor (da Costa et al.,2003). In E. coli, the flrd mRNA level was strongly induced by nitriteand slightly repressed by nitrate (da Costa et al., 2003). In contrast, theinduction of the flrd-lacZ reporter fusion increased slightly in the pres-ence of nitrate, and this effect was strengthened by the absence of thegene products of the flrd operon (Hutchings et al., 2002). In spite ofthese observations, the E. coli regulatory factors NarL and NarP werefound to be non-essential for the control of the flrd operon (da Costaet al., 2003). Nevertheless, E. coli strains deleted in narP and/or narLand grown in the presence of nitrite showed a higher transcriptionallevel of the flrd operon (da Costa et al., 2003), which however is muchsmaller when compared with the induction caused by the presence ofnitrite in the wild type strain. Concomitantly, in the absence of nitrateor nitrite, the level of flrd/flrd-red mRNA was significantly increasedin anaerobically grown cells of E. coli deficient in narL and/or narP.Altogether, the data suggests that E. coli NarL and NarP may actas negative regulators of the flrd operon in competition with otherregulator(s) of flrd, which may be activated by nitrite (or by a derivativeproduct).


Mechanisms of post-transcriptional control for flrd were also postulateddue to the immunoblotting studies that showed an inverse correlationbetween flavorubredoxin mRNA and protein levels (da Costa et al., 2003).In particular, and contrary to what was observed by transcription analysis,the expression level of FlRd in nitrite growth conditions was similar to theone measured in fermentatively grown E. coli but much lower than innitrate-grown E. coli.Regulation of flrd seems to also involve the E. coli norR gene, which is

divergently transcribed from flrd. Based on sequence similarity with thefamily of two-component response regulators, NorR is organised into threefunctional domains: the N-terminal sensor domain, a central domain thatinteracts with s54-containing polymerase and a C-terminal DNA bindingdomain. Gardner et al. (2003) showed that upon deletion of the NorRsignalling region, NO consumption activity could be detected in theabsence of NO. The involvement of s54 was also confirmed because the�( flrd-lacZ) fusion was significantly impaired in a s54-deficient strain(rpoN deletion).The phenotypes of the norR and flrd minus strains were found to be

similar, showing the norR-deficient strain a lack of NO reductase activitythat could be rescued through the expression of NorR from a multicopyplasmid (Gardner et al., 2003). A possible negative autoregulation wassuggested because the norR promoter activity was highly active in thenorR mutant (Hutchings et al., 2002). However, the lack of promoteractivity in a flrd-red mutant could not be rationalised. While nitroprusside(and nitrite) promotes a slight repression of the norR promoter, the presenceof nitrate induced a small stimulation, which is fully dependent on the flrdgene product (Hutchings et al., 2002).Although not in full agreement with previously reported results, the level

of norR mRNA was also shown to respond to the growth conditions(da Costa et al., 2003): a significant increase in the norR mRNA levelwas measured when the E. coli metabolism was shifted from aerobic toanaerobic conditions, and in the presence of NO or nitrate, while nitriteexerted a repressor effect. Furthermore, the transcription of norR isrepressed by FNR, but independent of narL and narP (da Costa et al.,2003).The action of flavorubredoxin as NO reductase is most probably

maximised in anaerobic conditions, but a role during microaerobicgrowth has also to be considered. In fact, under microaerobic conditionsE. coli FlRd alone (as well as Hmp) was able to protect an E. coli hmpmutant from the severe growth arrest observed in a E. coli hmp/flrddouble mutant (Gardner et al., 2002). Under aerobic growth conditions


the flrd promoter activity behaved differently: activation by nitroprussideof the flrd-lacZ fusion was only observed in rich medium, and in mediumsupplemented with nitrite (Hutchings et al., 2002). Although aerobic flrdtranscription occurred in a low level, no FlRd could be detected byimmunoblotting of aerobically grown cells of E. coli (da Costa et al., 2003)indicating that, if present, the amount of protein is very low.The participation of flavodiiron proteins in bacterial infection mechan-

isms was not yet shown. So far, the only indication available is theobservation that a gene sharing high similarity with these enzymes wasamong the set of genes that were up-regulated during the process ofplant infection by E. chrysanthemi (Okinaka et al., 2002).

3. CONCLUDING REMARKS

In this chapter, we reviewed the recent experimental evidence that led to theestablishment of a novel family of nitric oxide reductases. First isolatedfrom a limited number of bacteria, its general relevance is well demon-strated by the analyses of the large number of sequenced genomes, ofanaerobic or facultative anaerobes, from Archaea, Bacteria and Eukarya.The flavodiiron proteins are a beautiful example of modular evolution ofenzymes, showing how, by putting together distinct structural domainsand by subtle aminoacid changes, new enzymatic functions have beenachieved throughout evolution. Many questions remain to be answered, ina highly exciting new field: from the molecular mechanism of NO reduc-tion by these enzymes, their apparent bifunctionality, to the regulationof their expression and possible impact on pathogenicity. Altogether, themultiple strategies that micro-organisms use to cope with nitric oxidestress, reviewed in this chapter, reveal a common leitmotif in biology: citingH. V. Westerhoff (Koefoed et al., 2002), complexity and sophisticationof biological mechanisms, as well as the apparent redundancy of the mostimportant mechanisms, lead to life robustness.

ACKNOWLEDGEMENTS

Our work described in this chapter was funded by grants from the Fundacaopara a Ciencia e Tecnologia, Portugal. J. B. Vicente is a recipient of aPOCTI SFRH/BD9136/02 grant. We would like to thank Marta C. Justinofor carefully reading the manuscript, and all our co-workers, whose names


appear in the references. We would like to dedicate this chapter to thememory of Jean LeGall, both for his longstanding collaboration and forhis invaluable contribution to the fields of microbial biochemistry andbioinorganic chemistry.

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Stress Responsive Bacteria: Biosensors asEnvironmental Monitors

Amy Cheng Vollmer1 and Tina K. Van Dyk2

1Department of Biology, Swarthmore College, 500 College Avenue,

Swarthmore, PA 19081, USA2DuPont Central Research and Development, Experimental Station E173/216,

P. O. Box 80173, Wilmington, DE 19880-0173, USA

ABSTRACT

The delicate and dynamic balance of the physiological steady state and itsmaintenance is well characterized by studies of bacterial stress response.Through the use of genetic analysis, numerous stress regulons, theirphysiological regulators and their biochemical processes have beendelineated. In particular, transcriptionally activated stress regulons aresubjects of study and application. These regulons include those thatrespond to macromolecular damage and toxicity as well as to nutrientstarvation. The convenience of reporter gene fusions has allowed thecreation of biosensor strains, resulting from the fusion of stress-responsive promoters with a variety of reporter genes. Such cellularbiosensors are being used for monitoring dynamic systems and canreport the presence of environmental stressors in real time. Theyprovide a greater range of sensitivity, e.g. to sub-lethal concentrationsof toxicants, than the simple assessment of cell viability. The underlyingphysiological context of the reporter strains results in the detection ofbioavailable concentrations of both toxicants and nutrients. Cultureconditions and host strain genotypes can be customized so as to maximizethe sensitivity of the strain for a particular application. Collections of

Correspondence: E-mail: [email protected] or [email protected]



DOI 10.1016/S0065-2911(04)49003-1

specific strains that are grouped in panels are used to diagnose targetsor mode of action for unknown toxicants. Further application inmassive by parallel DNA and gene fusion arrays greatly extends theinformation available for diagnosis of modes of action and may lead todevelopment of novel high-throughput screens. Future studies will includemore panels, arrays, as well as single reporter cell detection for a betterunderstanding of the population heterogeneity during stress response.New knowledge of physiology gained from further studies of novelsystems, or using innovative methods of analysis, will undoubtedly yieldstill more useful and informative environmental biosensors.

1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1331.1. Stress response: definition and scope . . . . . . . . . . . . . . . . . . . . 1331.2. Stress response: specificity and sensitivity. . . . . . . . . . . . . . . . . . 1341.3. Cellular biosensors and environmental monitoring. . . . . . . . . . . . . . 135

2. Reporters of gene expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1362.1. lacZ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.2. inaZ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.3. gfp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.4. dsRed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1392.5. cobA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1402.6. luc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1402.7. ruc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1412.8. luxCDABE and luxAB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141

3. Macromolecular damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423.1. Genotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423.2. Heat shock. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1453.3. Oxidation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1463.4. Membrane damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1473.5. Multi-stress responses to toxic metals . . . . . . . . . . . . . . . . . . . . 147

4. Nutrient limitation/imbalance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1504.1. Carbon-starvation response . . . . . . . . . . . . . . . . . . . . . . . . . . 1514.2. Nitrogen-starvation response . . . . . . . . . . . . . . . . . . . . . . . . . 1524.3. Phosphate-starvation response . . . . . . . . . . . . . . . . . . . . . . . . 1524.4. Sulfur-starvation response . . . . . . . . . . . . . . . . . . . . . . . . . . . 1534.5. Multi-nutrient starvation response . . . . . . . . . . . . . . . . . . . . . . . 154

5. Panels and arrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1555.1. Panels of stress responsive gene fusions . . . . . . . . . . . . . . . . . . 1565.2. DNA arrays. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1585.3. Gene fusion arrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1605.4. From arrays to specialized panels. . . . . . . . . . . . . . . . . . . . . . . 161

6. Future trends . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

132 AMY CHENG VOLLMER AND TINA K. VAN DYK

1. INTRODUCTION

Bacterial gene circuits that have arisen during the course of evolutiondisplay a variety of regulatory schemes. Yet they have in common theability to sense and respond, often discriminating and distinguishingbetween a long list of environmental stresses. While the regulation of geneexpression can occur at many levels, from the structure of the DNAtemplate to the post-translational modification and stability of a proteinproduct, the initiation of transcription is a frequent and common step atwhich control is exerted. The upstream DNA sequences that precede thegene coding region serve as sites at which integration of cellular signals,indicative of internal and external conditions, occurs.Such is the case for genes that are members of stress regulons, that is

suites of unlinked genes that become coordinately expressed in response toconditions that we can manipulate in our laboratories. These conditionsmay reflect naturally selective forces that have driven the evolution andmaintenance of these genes in prokaryotic and eukaryotic genomes.Many investigators have been motivated to characterize promoters andregulatory elements (i.e. cis and trans acting factors), the combined actionof which governs transcription initiation.

1.1. Stress Response: Definition and Scope

By ‘‘stress’’ we choose an inclusive working definition of any perturbation ofthe steady state condition. Generally, measurements of bacterial stressresponse are, in fact, ensemble averages of the behavior of a populationof bacteria. The reporters chosen are introduced into clonally relatedorganisms. Even though there is heterogeneity among the haploidprogeny of any single founding cell from a colony, the data collected froma population of these progeny are treated as a whole, with little attentiondrawn to individual variation. Although this summarizes many recentstudies, currently, optical tweezers, more advanced microscopy and sophisti-cated image analysis may allow the description of the distribution ofbehaviors and variance of single cells within a given population.Response to stress ultimately results in repair, restoration or degradation

of the damaged or dysfunctional elements. The result is the re-establishmentof a new steady state and balance of resource influx and energeticoutput (See Fig. 1). Probably more often than not in nature, stresses occurcoincidentally rather than in isolation or in sequence, as our laboratory

STRESS RESPONSIVE BACTERIA 133

experiments are convenient contrivances for ease of study, manipulation,data collection and analysis. Response can be initiated at many levels ofgene expression and by varying macromolecular stability or degradation.Small effector molecules can also play important roles as covalent or non-covalent adducts.

1.2. Stress Response: Specificity and Sensitivity

It is convenient, but not necessarily accurate, to study stress responses inisolation. Many of the regulatory mechanisms and related circuits havebeen defined by studies that focused on only one or a few members of aparticular stress regulon. These strategies reflect the hierarchy of certaincircuits as well as the convenience in measurement of output. Morerealistically, stress responses are probably connected by elements that serveto coordinate the overall cellular physiology. Conservation of resources,integration of many different inputs, recycling of components and changesin growth rate, growth phase and locomotion are but a few pathwaysaffected when an organism adjusts to an upset. Transcriptome analyses,described in a later section, have shed new light on the global response tostress and the subtle connections that have evaded the more narrowstudies of the past. On the other hand, massive genomic analyses have been

Figure 1 A generic outline of transcriptionally regulated microbial stress responses.As the cell encounters an adverse environmental condition, cellular damage will result.This damage leads to generation of a signal for activated transcription of specific genesencoding proteins that combat the toxic agent or repair the damage caused. Thus, analysisof upregulated gene expression can reveal the nature of the biological damage caused bysamples containing unknown agents.


informed by specific knowledge from previous work. At the same time, theselarge microarray experiments suggest targeted, clarifying studies inparticular realms of the genome. When global experiments insinuate thefunction of a ‘‘gene of unknown function’’, there is a rationale for morespecific genetic and physiology experiments centered on that gene.In many cases, the responses are fine-tuned for specific damage or

insult. That is what one might expect, for example, of the hydrogenperoxide-induced activation of katG (encoding ‘‘catalase’’: hydroperoxidaseI). The enzyme catalyzes a specific reaction: the hydrolysis of hydrogenperoxide to water and oxygen. While the specificity of the enzyme’sactive site is fairly strict, the ability of Escherichia coli to activate katG isresponsive to a wide range of organic peroxides as well (Belkin et al.,1996a). This is because of the regulatory protein OxyR and its abilityto assume a different conformation after oxidation (Zheng et al., 1998),and not due to the katG gene product’s specificity. While it is anoxidation event in the OxyR protein that results in the conformationalchange, oxidation itself does not discriminate between organic and inorganicsources of peroxides. In other cases, the same gene (e.g. uspA) appears tobe activated in response to any of a number of seemingly unrelatedstresses (Nystrom and Neidhardt, 1992). This may have to do withthe putative, undefined function of the gene as a global regulator (Taoet al., 1999).In this chapter, we have selected representative studies of well

characterized stress responses and reporters. It is not the intent of thisreview to be exhaustive; rather we have chosen to be broad and illustrativein our descriptions and examples. It should be noted that comprehensivevolumes on stress responses in bacteria have been compiled recently(Storz and Hengge-Aronis, 2000); physiological, genetic and regulatorynetworks continue to be well synopsized in compendia such as Escherichiacoli and Salmonella typhimurium: cellular and molecular biology(Neidhardt et al., 1996).

1.3. Cellular Biosensors and Environmental Monitoring

In the broadest sense, biosensors are devices utilizing a biologicalcomponent to detect a physiological or biochemical change. Accordingly,cellular biosensors are those in which the biological component is awhole cell. Such cellular biosensors have found applications in numerousareas, including monitoring of pollution and toxic chemicals in theenvironment. Pioneering work in this area has used cellular biosensors


based on catabolic regulatory genes to detect organic pollutants, such asnaphthalene (Burlage et al., 1990). This is an excellent approach for thedetection of specific classes of molecules. As a complement to these morespecific sensors, monitoring with stress responses allows detection ofenvironmental toxicants without a requirement for prior knowledge ofpossible chemical contaminants. Furthermore, use of stress responses formonitoring provides more information on the nature of the toxicity andgreater sensitivity than does monitoring approaches based on cellularviability. Use of cellular biosensors for environmental applications hasa potential advantage of reporting on the bioavailability of pollutantsbecause whole cells will respond only to the bioavailable fraction.Additionally, the capability to adjust medium composition, cell density,growth phase and other growth parameters allows for the customization ofa given population of cellular biosensors for specialized conditions.Furthermore, storage of cells in lyophilized form allows for the biosensorsto be used as reagents in standardized assays (Corbisier et al., 1996;Tauriainen et al., 1997; Wagner and Van Dyk, 1998). Several recent reviewshave discussed various aspects of the use of cellular biosensors forenvironmental applications (D’Souza, 2001; Hansen and Sorensen, 2001;Keane et al., 2002; Belkin, 2003). Here we focus on the measurement ofmicrobial stress responses using reporter genes and utility for environmentalmonitoring.

2. REPORTERS OF GENE EXPRESSION

The use of reporter genes to indirectly measure promoter activity is pre-valent for fundamental studies (Silhavy, 2000) and numerous practicalapplications (Daunert et al., 2000; LaRossa and Van Dyk, 2000). Table 1lists several reporter gene systems with easily assayed products. Thesegenes or operons lacking native transcriptional control regions havebeen fused to promoters and upstream regulatory sites by in vitro recom-binant DNA methods or by in vivo transposon techniques. Typically, thegene fusions are ectopic and do not replace the normal stress responsivegene in the chromosome. Upon stress to cells carrying a reporter genefusion to a stress responsive promoter, the product of the reportergene serves as a convenient measure of the transcription initiated fromthe fused upstream region. The choice of reporter systems should becarefully made as the advantages of each should be weighed againstlimitations.


Table 1 Selected reporter gene systems.

Gene Protein(s) Product Detection

cobA Uroporphyrinogen IIImethyltransferase

Red fluorescent molecules,sirohydrochlorin andtrimethylpyrrocorphin

Fluorometry, fluorescence microscopy,or visually

dsRed Red fluorescent protein Red fluorescence followingexcitation

Fluorometry, fluorescence microscopy,or visually

gfp Green fluorescent protein Green fluorescence followingexcitation

Fluorometry, with a fluorescence-activatedcell sorter, fluorescence microscopy, orvisually

inaZ Ice nucleation protein Ice Freezing assays to detect ice formationlacZ b-Galactosidase Chemiluminescent, fluorescent, or

colored molecules with additionof appropriate enzymaticsubstrate

Luminometry, fluorometry,spectrophotometry, visually, orelectrochemically

luc Insect luciferase Light with addition of luciferin Luminometry, Scintillation counting, CCDimaging, with photographic film, or visually

luxAB Bacterial luciferase Light with addition of long chainaldehyde

Luminometry, Scintillation counting, CCDimaging, with photographic film, or visually

luxCDABE Bacterialbioluminescence

Light Luminometry, Scintillation counting, CCDimaging, with photographic film, or visually

ruc Renilla luciferase Light with addition ofcoelenterazine

Luminometry, Scintillation counting, CCDimaging, with photographic film, or visually

STRESSRESPONSIVEBACTERIA

137

2.1. lacZ

The best-known and most widely used reporter gene continues to be lacZ,encoding b-galactosidase from E. coli. Historical development of the lacgenetic system led to many lac based selections and screens (Beckwith, 1996)and numerous methods to quantitate b-galactosidase activity are incurrent use. However, these methods require cell disruption, substrateaddition, incubation, and product measurement. While these steps can beautomated (Menzel, 1989; Griffith and Wolf, 2002), the complexity ofmeasuring b-galactosidase activity has precluded many applications such asenvironmental monitoring and parallel measurements of gene expression.However, electrochemical determination of b-galactosidase used as areporter of gene expression has been recently demonstrated using thesubstrate p-aminophenyl-b-galactopyranoside, the product of which canbe oxidized by an electrode (Scott et al., 1997; Biran et al., 1999).Such electrochemical detection, which can be done on-line, allows thedevelopment of biosensors for in situ use (Biran et al., 2000).

2.2. inaZ

The ice nucleation proteins, from some bacterial species such asPseudomonas syringae, are outer membrane proteins that catalyze iceformation (Lindow, 1983). Thus, the activity of promoters fused to the inaZreporter can be detected by ice nucleation activity in supercooled aqueoussolutions (Loper and Lindow, 2002). The use of inaZ as a reporter ofpathogenicity gene expression in P. syringae pv. phaseolicola proved to bemuch more sensitive than lacZ in the same system (Lindgren et al., 1989).Such sensitivity and the broad range of ice nucleation activity that can bedetected are advantages of the inaZ reporter. Some disadvantages tothe inaZ reporter are the variable response time of the ice nucleationphenotype among bacterial species, the high degree of dependence on thetemperature and osmolarity at which the cultures are grown, the need toestablish the relationship between InaZ protein content and ice nucleationactivity for each microbial species in which inaZ is used, and the laborintensive assay for ice nucleation (Loper and Lindow, 2002).

2.3. gfp

Since the first publication in 1994 demonstrating the use of greenfluorescent protein from Aequorea victoria as a reporter of gene expression


(Chalfie et al., 1994), the use of gfp has attracted much attention (Southwardand Surette, 2002). A notable feature is the ease of measuring fluorescenceactivity after simple irradiation at the excitation wavelength without arequirement to lyse cells or add substrate molecules. This non-invasivedetection of GFP has proven very useful for determination of proteinlocalization in microbial cells (Phillips, 2001) and as a marker of individualmicrobial species in mixed populations (Tombolini and Jansson, 1998). Thegfp reporter has also been used to study kinetics and levels of geneexpression in microbial systems. For example, gfp fusions to E. colipromoters regulated by the heat shock sigma factor, s32, reported onvarious stresses to resting cells with increased fluorescence (Cha et al., 1999).In another study, gfp fusions to promoters of genes encoding flagellarproteins were used to order expression patterns (Kalir et al., 2001).These and other studies with gfp as a reporter of gene expression havefaced limitations. Importantly, the slow formation of the activefluorophore results in a substantial lag time between GFP synthesis andfull activity. Such lags have been reduced in variants of gfp encoding fasterfolding proteins (Cormack et al., 1996; Scholz et al., 2000), yet lag timesof an hour or more are still common. An additional limitation of GFPfor use as a reporter of gene expression is its extreme stability. Thus,analytical methods, such as calculating the rate of change of GFPfluorescence, are needed to accurately define changes in gene expression(Lu et al., 2002). An additional limitation of gfp is the intrinsic fluorescenceof bacterial cells that gives a background signal resulting in poor detectionat low levels of gfp expression. It should also be noted that use of gfp islimited to aerobic conditions because oxygen is required for fluorophoreformation and that the fluorescence of gfp is sensitive to internal pH.

2.4. dsRed

In 1999, Matz and coworkers described the cloning of genes encoding sixnaturally fluorescent proteins from coral (Matz et al., 1999). One of theseproteins from Discosoma sp., now known as DsRed, has excitation andemission maxima at 558 and 583 nm, respectively. At these wavelengths,cellular auto fluorescence is expected to be far less than at wavelengthsused for GFP. Thus, DsRed has been suggested to be useful for variousapplications. However, to date, it has not proven useful as a reporter ofgene expression in bacteria. For example, a comparison of luc, luxCDABE,gfp and dsRed as reporters of gene expression in E. coli was recentlyreported (Hakkila et al., 2002). Of these four reporters, dsRed yielded the


slowest response and lowest sensitivity. Slow kinetics of fluorophoreformation (Gross et al., 2000) are probably related to the slow response.Nonetheless, as protein engineering has resulted in improvements in GFP,so protein engineering may yield a more useful DsRed. As an indicationthat this approach may prove fruitful, an evolved version of DsRed thatfunctions as a monomer, rather than the obligate tetramer, has recently beenreported (Campbell et al., 2002).

2.5. cobA

Following observations of the utility of cobA or cysG, both of whichencode uroporphyrinogen III methyltransferase, in producing red fluores-cence as a cellular marker (Roessner and Scott, 1995), the abilityof Propionibacterium freudenreichii cobA to serve as a reporter of geneexpression was shown (Wildt and Deuschle, 1999). Red fluorescenceobserved upon expression of cobA results from the conversion ofuroporphyrinogen III to two fluorescent molecules. As most cells normallymake the substrate of the enzyme, no exogenous substrate additionis required to obtain fluorescence. However, the intensity of fluorescencemay become substrate-limited (Feliciano and Daunert, 2002). To date, veryfew applications of cobA as a reporter of gene expression have beendeveloped.

2.6. luc

Insect luciferases catalyze the oxidation of benzothiazolyl-thiazoleluciferin in the presence of ATP, oxygen and magnesium with resultinglight production. A gene encoding luciferase was first isolated from theAmerican firefly Photinus pyralis (de Wet et al., 1985), but has also beencloned from other firefly species (Tatsumi et al., 1992) and from click beetlePyrophorus plagiophthalamus (Wood et al., 1989). The light emissionvaries from 550 to 575 nm dependent on the source of the luc gene(Bronstein et al., 1994). Mutant luc genes with single amino acidsubstitutions have light emissions ranging from red to green (Wilson andHastings, 1998). Thus, an advantage of the luc reporter is that related genescan be used to generate distinct signals. The luc reporter is more commonlyused in mammalian cells than in microbes. Nonetheless, luc has been usedsuccessfully to monitor gene expression in several gram negative bacteria(Palomares et al., 1989) and for applications such as detection of responsesto arsenite and antimonite (Tauriainen et al., 1997). A disadvantage of the


luc reporter is that the luciferin substrate of the insect luciferase is arelatively costly reagent and passes through bacterial membranes only atlow pH (Cebolla et al., 1995); thus, continuous reporting of gene expressionis not possible.

2.7. ruc

The ruc gene for luciferase from Renilla reniformis, an anthozoancoelenterate, has been cloned and expressed in E. coli (Lorenz et al.,1991). The Renilla luciferase is a 36 kDa monomeric protein that catalyzesthe oxidative decarboxylation of coelenterazine to oxyluciferin andcarbon dioxide with emission of blue light (Matthews et al., 1977). Theruc gene has been successfully applied as a reporter of gene expression inCandida albicans (Srikantha et al., 1996) and as a secreted reporter inmammalian cells (Liu et al., 1997). Recently, Ruc-GFP fusion proteins havebeen constructed and shown to report on gene expression in cell cultures andlive animals (Wang et al., 2002; Yu and Szalay, 2002); Ruc activity wasdetected in the presence of coelenterazine, while the GFP fluorescencewas observed after excitation with UV light. Despite these successesin eukaroytes and results confirming ruc function in bacterial systems(Jubin and Murray, 1998), applications in bacteria have not been developed.Improvements, such as those in codon usage, may be necessary to optimizeexpression in bacteria.

2.8. luxCDABE and luxAB

The five-gene bacterial bioluminescence system from the marine micro-organism Vibrio fischeri was first used as a reporter of gene expressionin 1985 (Engebrecht et al., 1985). Since then, this reporter gene systemand lux genes from other bioluminescent bacteria have been widely usedbecause of the ease, sensitivity and large dynamic range of lightmeasurements. In addition, the luxCDABE reporter produces a cellularsignal thereby eliminating the need for cell disruption and enzymaticassay, which then allows for continuous monitoring of gene expression.The five lux genes encode proteins that form the heterodimeric luciferase(luxAB) and that result in synthesis of a long chain aldehyde (luxCDE).This is oxidized in association with the reduced flavin mononucleotide(FMNH2), which is the luciferin molecule. Expression of the five lux genesin cells in the presence of oxygen, ATP and reducing power resultsin continuous light production at a maximum emission of 490 nm


(Meighen, 1991). Several luminescent bacteria provide sources ofluxCDABE operons. The gene products of some of these operons, includingthose derived from V. fischeri, Vibrio harveyi and Photobacterimphosphoreum, are unstable at temperatures in the growth range ofmany bacterial species (Chatterjee and Meighen, 1995). However, theluxCDABE-encoded proteins of Photorhabdus luminescens are significantlymore robust and are active when expressed in E. coli at temperatures up to42�C (Szittner and Meighen, 1990). A recent improvement that allowsefficient expression of the lux genes in gram positive bacteria has includedmodifications to the translation initiation regions of each gene anda concomitant reordering of the operon genes to luxABCDE (Francis et al.,2000, 2001). As an alternative to using a five gene operon, two genes, luxABthat encode the heterodimeric luciferase enzyme, are also used as a reporterof gene expression. In this case, light production occurs when a longstraight-chain aldehyde substrate, such as n-nonanal or n-decanal, is addedto the cells. These lipophilic aldehydes readily diffuse across bacterialmembranes, making cell lysis unnecessary. Advantages of the lux reportergenes in addition to the ease and sensitivity of light measurements are therapid response of the bioluminescent signal following induction ofexpression (Belkin et al., 1996a; Van Dyk et al., 1994b), the numerousoptions for measuring light production (Vollmer et al., 1998) that includesseveral commercial instruments, and the development of field portabledevices, such as single use patches of cells immobilized in latex (Lyngberget al., 1999) or bioluminescent bioreporter integrated circuits that directlyintegrate the cellular reporters with a measuring device (Simpson et al.,1998; Bolton et al., 2002). The principle limitation of the lux reporters is therequirement for active cellular metabolism to generate the bioluminescentsignal. Another consideration when using the five-gene lux operon is that,under some circumstances, an unexpected source of long chain aldehyde canresult in increased light production in the absence of increased transcriptionof the reporter genes (Heitzer et al., 1998). Additionally, the oxygenrequirement for the luciferase reaction means that the lux reporter activitymust be analyzed in aerobic conditions.

3. MACROMOLECULAR DAMAGE

3.1. Genotoxicity

Numerous biosensors of genotoxicity have been constructed. Bacteriaare particularly well suited for this type of measurement, as was ably


demonstrated by Ames et al. (1973). The Ames test relies on growth ofrevertants to quantitate induction of error-prone repair systems in responseto DNA damage. Cellular biosensors have now been developed usingreporters of gene expression that give a more rapid and easily assayed signalin response to conditions that result in DNA damage. Two induciblesystems of bacterial DNA damage repair are the recA-independent,ada-controlled adaptive response and the recA-dependent, lexA-controlledSOS response. The former responds specifically to the presence ofmethylated phosphotriesters generated by DNA alkylation. This signalactivates the ada gene product, which, in turn, triggers the transcriptionof genes such as ada, alkA, alkB and aid (Rupp, 1996). In the SOS system,induction occurs via a single-stranded DNA/RecA nucleoprotein filament.This form of RecA stimulates the autodigestion of the LexA repressorresulting in the transcriptional derepression of several genes. These SOSgenes include uvrA, uvrB, recA, and recN, the products of which areinvolved in DNA repair, and others, such as sulA, that couple DNA damageto cell division (Walker et al., 2000; see Fig. 2).Various promoters from genes in the SOS and the ada regulons have

been fused to several reporters. In general, responses of reporter geneactivity to mutagenic effects of chemicals and radiation increase in a dose-dependent manner at sub-lethal doses. In addition, lethal concentrationsof genotoxic agents are detectable if reporters, such as lux, that rely oncellular viability for function, are used. Such non-specific loss of reportersignal can be detected because many of the promoters have a lowconstitutive expression in the absence of the manipulated genotoxicant.In the case of a recA-luxCDABE reporter (Vollmer et al., 1997) the utilityof ‘‘lights on’’ at sub-lethal doses and ‘‘lights off’’ at lethal doses wasfirst demonstrated. In this study, the recA-luxCDABE induction wasdemonstrated to be under the control of the LexA repressor. Further, theresponse was similar, but less sensitive, when a lacZ reporter gene wasused. Fusions of the lacZ reporter to recA (Nunoshiba and Nishioka, 1991),dinD (Orser et al., 1995), or sfi (Quillardet and Hofnung, 1993) and of theumuDC promoter to lacZ (Oda et al., 1985), luc (Schmid et al., 1997), lux(Justus and Thomas, 1998), or gfp (Justus and Thomas, 1999), have showndifferent levels of sensitivity. A brief review on the use of stress-responsiveluminous bacteria for genotoxicity testing has been compiled by Belkin(1998b).As in the case of the Ames assay, metabolic activation of pro-mutagens

by barbiturate-induced P450 enzymes increases the relevance for humanhealth risk. The availability of such P450 enzymes as reagents allows thesystematic detection of potential mutagens. One application of this


enhancement has been in the development and utility of stress-responsivebiosensors panels (see Section 5.1) Cellular response to genotoxic agentscan be heightened by the presence of certain mutations in DNA repair. Inorder to cover a particular concentration range of toxicants, one couldplace, for example, the recA-luxCDABE reporter in different backgrounds,e.g. mut, din, umu.Use of genotoxicity cellular biosensors has been effective in detection

of potential genotoxicants in several environmental settings, such aswastewater (Belkin et al., 1996b, 1997), soil (Ehrlichmann et al., 2000),and air (Hamers et al., 2000). However, results of a recent study sug-gested that the sensitivity of the umu-lacZ based sensor may not be

Figure 2 Construction of a bioluminescent cellular biosensor for DNA damage.When E. coli DNA is damaged, the RecA protein becomes bound to single strandedDNA and in this form stimulates the autocatalytic cleavage of the LexA transcriptionalrepressor. Subsequently, the transcription of dozens of genes with LexA-bindingsites is upregulated. By genetic manipulation, a gene fusion formed with a LexA-regulated promoter driving expression of the luxCDABE reporter is added to the cell.As DNA damage is encountered, transcription of genes for DNA repair, such as recA,recN and uvrA, is upregulated. At the same time, in response to the same signal, thetranscription of the luxCDABE reporter is increased. Thus, the increased lightproduction, which is readily measured, corresponds with the presence of a conditionleading to DNA damage.


adequate for surface water monitoring and thus should be used incombination with other methods to assess genotoxic potential (Dizer et al.,2002).

3.2. Heat Shock

The heat shock response is so called because it was originally characterizedas a stress response to elevated temperatures. However, this response,which is found in essentially all prokaryotes and eukaryotes, is nowunderstood to be a cellular response to damaged proteins. The heatshock response is induced when cells encounter a rapid increase intemperature or many other stress conditions that lead to proteindenaturation, such as changes from anaerobic to aerobic conditions, viralor phage infection, or exposure to many chemicals (Welch, 1990). Chemicalsthat induce the heat shock response in E. coli are wide-ranging, includingtoxic metals, organic solvents, herbicides, fungicides, weak acids, anddetergents (Van Dyk et al., 1995a, 1995b). Thus, monitoring the heat shockresponse is useful as a general indicator of adverse environmentalconditions.Many of the genes upregulated in the heat shock response encode

molecular chaperones and proteases so that the induction of this responseallows the cell to restore or degrade damaged proteins (Lund, 2001; Douganet al., 2002). In E. coli, induction of the heat shock response is primarilymediated by a transient increase in s32. This sigma factor directsRNA polymerase to transcribe several dozen genes (Yura et al., 2000).While the proteins encoded by heat shock genes are in general conservedin bacterial species, the regulatory mechanisms can be quite distinct.Accordingly, in Bacillus subtilis, expression of some heat shock proteins iscontrolled by specific repressors while others are part of the sB general stressregulon (Yura and Nakahigashi, 1999).Biosensors for environmental applications to monitor protein damage

responses were constructed by fusing the luxCDABE reporter tothe promoter regions of heat shock genes in E. coli (Van Dyk et al.,1994b). The strains containing fusions to the dnaK or grpE promotersrespond with increased light production to many pollutant molecules.The bioluminescent response from these fusion strains is induced atsublethal concentrations thereby providing an advantage in detectionlimit over systems that rely on cell death. The sensitivity of these biosensorstrains to hydrophobic chemicals is further enhanced by introduction of


a tolC mutation that prevents efflux pump function and thus enhancesintracellular toxicant accumulation.The utility of heat shock reporter gene fusions for detection of

environmental pollutants has also been demonstrated using other bacterialspecies, such as Pseudomonas sp. (Park et al., 2002), and with reportergenes other than luxCDABE (Cha et al., 1999; Kwak et al., 2000). Theapplicability of monitoring heat shock expression as general indicators oftoxicity in wastewater treatment is suggested by studies demonstrating theinduction of heat shock proteins in microorganisms of activated sludge(Bott et al., 2001; Love and Bott, 2002). However, further development isnecessary before heat shock biosensors, as an alternative to other generaltoxicity tests, are used in field applications (Bierkens, 2000).

3.3. Oxidation

The utility of oxidative-stress responsive reporters has been demonstratedin the ability of an E. coli katG-luxCDABE fusion to report bothhydrogen peroxide and organic peroxides (Belkin et al., 1996a). Theresponse of this reporter relies on the fact that the sensor for oxidationis the OxyR protein (Storz et al., 1990), containing a well-poised pairof sulfhydryls that can become oxidized to form a disulfide. This thiol-disulfide switch immediately conveys a conformational change throughthe protein (Zheng et al., 1998). The oxidized protein becomes atranscriptional activator that binds to numerous promoters, including thekatG promoter. Many of the proteins whose expression is activated byOxyR function as antioxidants (Storz and Zheng, 2000), such ashydroperoxidase I, or catalase, encoded by katG, which inactivateshydrogen peroxide. The katG-luxCDABE reporter was used to show thatoptical tweezers with a laser (�¼ 1064 nm, at 900 mW) induced oxidativestress in tweezed E. coli cells (Gaskell et al., 2003.) Another novelapplication used the katG-luxCDABE reporter to measure the level ofantioxidant activity in epicatechin, a component of green tea (Nobile andVollmer, 2000.)Another oxidative stress response regulon comprises genes regulated

by SoxRS. In this case, the Fe–S cluster of SoxR senses the presence ofsuperoxide and, in the oxidized, active form, enhances soxS transcription.Increased levels of SoxS then up-regulate expression of the regulon,which includes genes encoding proteins that combat oxidative stress(reviewed in Storz and Zheng, 2000). In E. coli, most of the genes of thesoxRS regulon can also be activated by MarA or Rob in response to stresses


other than oxidative. Thus, further analysis is often necessary to uncoverwhich regulatory circuit controls upregulation of genes in this regulon. Forexample, the response of E. coli carrying micF-luxCDABE to cationicantimicrobial peptides was shown to be mediated by Rob using a set ofmutant strains each lacking one of the regulators (Oh et al., 2000).

3.4. Membrane Damage

Membrane damage is not easily measured directly. Certainly, additionof toluene or chloroform to a turbid cell suspension lyse cells andbrief, low-speed centrifugation would yield a cleared supernatant thatcould be visualized spectrophotometrically. But this is neither rapid norquantitative. In E. coli, genes which are activated in response to membraneperturbations include the phage shock protein genes, which are activatedin response to filamentous phage infection (Model et al., 1997), and thoseinvolved in fatty acid biosynthesis. FadR regulates both fatty aciddegradation ( fad genes) and biosynthesis ( fab genes) (Rock and Cronan,1996) and was shown to activate specifically the transcription of fabgenes in response to membrane damage (Cronan and Rock, 1996). In E. coli,the mechanical damage by acoustic cavitation, as the result of highfrequency ultrasound waves, resulted in induction of a pspA-luxCDABEreporter constructed by Halpern et al. (1998) as well as in a strain bearinga fabA-luxCDABE plasmid reporter (Vollmer et al., 1998). The latter fusionhad been previously shown by Belkin et al. (1997) to be transcriptionallyactivated, in a FadR-dependent manner, when the E. coli strain carryingit was treated with solvents and detergents.

3.5. Multi-Stress Responses to Toxic Metals

Exposure of bacterial cells to toxic metal ions induces expression ofseveral stress regulons due to the multiplicity of toxic effects on cellularphysiology. Consider, for example, the response of E. coli cells to thepresence of cadmium, a metal with no known biological role that canbe a hazardous environmental pollutant. The heat shock responseis induced (VanBogelen et al., 1987; Blom et al., 1992; Van Dyk et al.,2000), indicative of cytoplasmic protein damage and consistent withgeneral responsiveness of the heat shock regulon to many stresses.In addition, other genes that respond to numerous adverse environmentalconditions, such as uspA encoding the universal stress protein, are


upregulated upon cadmium treatment (Van Dyk et al., 1995b; Ferianc et al.,1998). Oxidative stress responses (VanBogelen et al., 1987; Van Dyk et al.,2000; Puskarova et al., 2002) are also induced as expected becausecadmium chloride is a potent oxidizing agent. Most proteins of the DNAdamage responsive SOS regulon are not induced by cadmium; however,increased levels of the RecA protein are observed (VanBogelen et al., 1987;Ferianc et al., 1998). This regulation of recA may not be at thelevel of transcription initiation because cadmium treatment did notinduce expression of a luxCDABE gene fusion to the recA promoter region(Van Dyk et al., 2000). A similar negative result for upregulation of anotherSOS responsive lacZ gene fusion supports the overall conclusionthat cadmium is not an inducer of the SOS response and correlates withthe negative result for mutagenesis in the Ames test (Quillardet andHofnung, 1993).As well as induction of the heat shock and oxidative stress responses,

cadmium treatment of E. coli induces stress responses that are morespecific to cadmium treatment. Notable is the upregulation of zntAencoding a P-type ATPase that translocates zinc, cadmium and lead ions(Beard et al., 1997; Rensing et al., 1997, 1998). Transcription of this geneis positively regulated by ZntR (Brocklehurst et al., 1999), a member ofthe MerR family, via a DNA distortion mechanism (Outten et al., 1999).Upregulation of zntA expression is induced by the substrates of the effluxsystem, zinc, cadmium and lead, and also to a limited extent by mercury(Babai and Ron, 1998; Brocklehurst et al., 1999; Binet and Poole, 2000;Noll and Lutsenko, 2000). Consistent with the relative toxicities,cadmium induces expression of zntA at much lower concentrations thandoes zinc. Biosensors that exploit zntA regulation have been developed. Afusion of the zntA to lacZ has been used in electrochemical detection ofcadmium in water, seawater, and soil samples (Biran et al., 2000). Inagreement with other studies, this sensor also responds to mercury ions,but with a lower signal, and to zinc ions, but at a much higher con-centration. Likewise a zntA-luxCDABE biosensor responded to saltsof cadmium, lead, mercury and zinc (Riether et al., 2001). The zntA-lacZbiosensor has recently been used to fabricate an optical imaging fiber-based biosensor in which lacZ expression in single cells was measured(Biran et al., 2003).As cadmium induces both broad and more specific stress responses in

E. coli, so do other toxic metals induce both common and specificstress responses in E. coli and other bacteria. These induced responses reflectthe toxic biological effects of the metal ions. Thus, for instance, aluminumions, like cadmium ions, induce the heat shock response but, unlike


cadmium ions, do not induce oxidative stress responses (Van Dyket al., 2000). In contrast, aluminum, but not cadmium, ions stimulateexpression of a yciG-luxCDABE gene fusion (Van Dyk et al., 2000), theexpression of which is dependent on the general stress response sigmafactor RpoS (Van Dyk et al., 1998). Thus, differential induction of stressresponses distinguishes the modes of action of these two toxic metals.Much work on biosensor development has focused on specific metal

ion defense responses. The first of these exploited the mercury resistancegenes found on Tn21, which were used to construct an E. coli strain carryinga mer-luxAB gene fusion (Condee and Summers, 1992). In this strain,binding of Hg(II) to heterologously expressed MerR activates transcriptionof the merTPCAD promoter thus driving expression of the fused luxABreporter genes. Hg(II) concentrations as low as 30 nM are detected byincreased light production. Other biosensors based on gene fusions tomercury resistance genes have been developed (Selifonova et al., 1993;Tescione and Belfort, 1993; Holmes et al., 1994; Hansen and Sorensen,2000), with sensitivities as low as 0.1 fM reported (Virta et al., 1995).Similarly, development of biosensors based on inducible resistances to othertoxic metals has been an active area of research. In addition to thosealready mentioned, bacterial strains with reporter gene fusions responsive toions of arsenic and antimony (Corbisier et al., 1993; Cai and DuBow, 1997;Ramanathan et al., 1997, 1998; Scott et al., 1997; Tauriainen et al., 1997),cadmium (Corbisier et al., 1993; Tauriainen et al., 1998), chromium(Peitzsch et al., 1998; Corbisier et al., 1999), copper (Holmes et al., 1994;Corbisier et al., 1996, 1998), lead (Tauriainen et al., 1998; Corbisier et al.,1999), or nickel and cobalt (Tibazarwa et al., 2001) have all beenconstructed. The specificity of these sensors varies. However, consistent withthe specific defense role of the regulated genes, responses are typicallyobserved to one or a small set of metal ions.A chief advantage of whole cell toxic metal responsive biosensors

has been asserted to be that the bioavailability of toxic metals is measured.The underlying assumption is that metal available to bacteria fromenvironmental samples will correlate with the metal available to higherorganisms. For toxic metals, as well as other environmental pollutants,the bioavailable concentrations can be dramatically different from thetotal concentrations. This is a currently active area of research as sensorsdeveloped in laboratories are being tested for application in manyenvironments. In some cases, biosensors provide useful indications ofbioavailable metals. For instance, measurement of bioavailable arsenicwith an E. coli strain carrying a firefly luciferase gene fusion regulated byArsR showed that aging and sequestration of arsenic in contaminated


soils around wood treatment plants results in progressively less bioavail-ability (Turpeinen et al., 2003). In other situations, biosensors werefound not to be suitable for particular applications. For example, abacterial biosensor for antimony gave a false positive signal with soilcontaminated from mining and smelting operations (Flynn et al., 2003).The antimony in the soil was not bioavailable, but arsenic and copperthat were found as co-contaminants were bioavailable and the levels ofthese metals were found to correlate with the bioluminescent responsesof the biosensor. In this case, a more specific biosensor may providethe solution. Alternatively, a panel of biosensors could be developedthat would distinguish responses due to antimony from those due to copperand arsenic. Such a panel may contain sensors for specific metals in com-bination with sensors for broader stress responses and thus wouldcharacterize the bioavailable metals and the toxicity profile of theenvironmental sample.

4. NUTRIENT LIMITATION/IMBALANCE

Limitation or starvation for nutrients is a common condition that mostbacteria experience in nature. Instead of sensing, repairing or degradingdamaged macromolecules, the physiology of the bacterial cell is redirectedto a search for alternatives (as in the case of glucose limitation),switching on the expression of higher affinity transporters or catalysts(e.g. for phosphate and ammonia), setting a different redox potential(e.g. during denitrification). The responses to these types of nutritionalstresses are being employed as biosensors with great sensitivity andversatility. In some cases, the environmental applications have been quitesuccessful. Frequently, the cells to be used for the purpose of sensingnutrient limitation must undergo a more prolonged preparation. Whilemacromolecular damage-responsive cells are often the most sensitiveand responsive during log phase growth, starvation responsive cells mustbe grown so that they are still viable, but contain very little of themolecular reserve that can be harvested intracellularly when externalconditions become limiting. For example, phosphate-starvation responseis not induced until the cells are grown into late stationary phase, suchthat the inorganic phosphate and related organic phosphate reserveshave been reduced (Cardemil and Vollmer, 2001). Escherichia coli,Pseudomonas fluorescens and P. putida have been the workhorse strains


used in environmental detection. In both genera, reporters fused to carbon-,nitrogen- and phosphate-starvation responsive promoters have beenreported (see LaRossa and Van Dyk, 2000, for a brief review). Further-more, the E. coli biosensors were shown to be regulated by their cognatephysiological regulators (Smulski et al., 1994; Cardemil and Vollmer,2001). Use of these strains in situ must be accompanied by the additionof the other nutrients so that cell viability (and therefore the reliability ofthe reporter) does not compromise or complicate the detection (Jensen et al.,1998). In addition, failure to prepare the cells accordingly results in alag time of 6–10 h before the biosensor (Cardemil and Vollmer, 2001)is induced.

4.1. Carbon-Starvation Response

The use of lac-luxCDABE fusions enables the detection of generalizedcarbon starvation, under the control of catabolite activation genes cyaand crp (Van Dyk et al., 1994a, 2001b). Another approach cleverlyexploits coupled bacterial respiration of negatively charged, low molecularweight compounds with reduction of NO�

3 to N2O (Meyer et al., 2002). Inthis way a microscale biosensor for the presence of acetate, propionate,isobutyrate and lactate was developed that was insensitive to changes in pHbetween 5.5 and 9 and stable to changes in salinity ranging from 0.2 to3.2%. Furthermore, the sensitivity of the linear response was increasedfive times by raising the temperature from 7 to 19.5�C. The inclusion ofan internal response correction is another mechanism to minimize theinfluence of non-specific effects from environmental changes on the bio-sensor response. Mirasoli et al. (2002) normalize the arabinose-responsiveGFPuv reporter with an IPTG-responsive EYFP reporter, by always addinga constant amount of the non-natural compound IPTG while varying theconcentration of L-arabinose. Only in this way was a clear dose–responsecurve reliably obtained. Furthermore, when tested in ‘‘non-optimalconditions’’ (in the presence of solvents or detergents), the inclusion of theinternal correction system was especially strategic. Exploiting theobservation that pseudomonads are affected by carbon limitation in bulksoil, Koch et al. (2001) chose the ss-dependent fic promoter fused to lacZas a reporter of the shift to stationary phase, due to lack of carbon inP. fluorescens. Their studies in pure culture and in soil showed that thisreporter was specific in its response to carbon, and not nitrogen orphosphorus, starvation.


4.2. Nitrogen-Starvation Response

Intracellular pools of ammonia are maintained through the interconversionof ammonia carriers glutamine, glutamate and a-ketoglutarate, alongwith a myriad of transamination reactions involving additional a-ketoand a-amino acids. Depletion of these ammonia reserves results in theactivation of genes such as glnA, which encodes glutamine synthetase.Elevated levels of this enzyme result in the efficient incorporation of freeammonia into the buffered intracellular pools. Since high concentrationsof free intracellular ammonia are detrimental, this response is under tightcontrol by glnFG (Reitzer and Magasanik, 1985). The second promoter(proximal) of glnA is controlled by glnFG and not by NTRII, a more globalregulator of nitrogen starvation (Magasanik and Neidhardt, 1987). As aconsequence, the sensitivity of the biosensor, E. coli glnAP2-luxCDABE (seeFig. 3), to different concentrations of added ammonia has a shorter linearrange than that of phosphate (Cardemil and Vollmer, 2001), since theaccumulation of free intracellular phosphate does not affect the pH asdrastically. Gillor et al. (2003) report a biosensor strain that fuses theSynechococcus glnA promoter to luxAB and is able to detect threenitrogen species, namely ammonium, nitrate and nitrite. Another nitrogen-responsive promoter fused to luxAB in P. fluorescens was shown to beexpressed when ammonia levels were below 10–90 mM in culture (Jensenet al., 1998). Amino acid starvation is detected with a his-luxCDABEfusion (using the his promoter region without the attenuator) in a stringentresponse-dependent manner (Van Dyk et al., 1994a).

4.3. Phosphate-Starvation Response

Depletion of intracellular phosphate levels in E. coli results in thestimulation of the pho regulon, which is under the control of phoB(Wanner, 1987). One member of this regulon, phoA, encodes a periplasmicalkaline phosphatase that hydrolyzes phosphate groups from a variety oforganic substrates. In E. coli, a phoA-luxCDABE strain, prepared andstored as a frozen reagent, was more sensitive than a commercial kit inmeasuring the levels of bioavailable phosphate in waters from the MiddleAtlantic region of the USA (Cardemil and Vollmer, 2001). The responseof the reporter, ranging from 1.8 to 15 mM, was shown to be under thecontrol of the physiological regulator, phoB. A cyanobacterial reportersystem, using phoA-luxAB has been developed and tested on freshwatersamples (Gillor et al., 2002). In both the E. coli and Synechococcus


systems, a long lag phase characterized the response of the reporter,indicating that intracellular pools sustained the population for several hoursin starvation medium before pho regulon activation occurred. Additionalenvironmental applications of phosphate biosensors include those devel-oped in P. putida and P. fluorescens that have been used to measurephosphate concentrations in bulk soil, sand and in the rhizosphere(de Weger et al., 1994; Kragelund et al., 1997).

4.4. Sulfur-Starvation Response

The cysteine regulon consists of genes involved in both the biosynthesisand transport of L-cysteine. The cysJ gene product is involved in thesulfate assimilation pathway, where ultimately sulfite is reduced to sulfide.Sulfur limitation causes derepression of the cysteine regulon. A hierarchy of

Figure 3 Kinetic data from a glnA-luxCDABE reporter strain that has been starvedof ammonia in stationary phase. Cells were resuspended in defined medium containingthe indicated concentrations of ammonia at time zero. The greater the amount ofammonia present, the less light is produced, since the glnA promoter is transcriptionallyactive when intracellular ammonia levels are low. The rise of each curve as well asthe maximum value is inversely proportional to the concentration of ammonia in themedium. Bioluminescence is reported in Relative Light Units, which is the number ofphotons counted, relative to the luminometer’s internal standard. Commercialluminometers measure light output with a dynamic sensitivity range of over 7 orders ofmagnitude.


derepression exists, where maximal derepression occurs during growthon poor sulfur sources and maximum repression is achieved by growth onL-cysteine (Loudon and Loughlin, 1992). The order of sulfur sourcecorrelates directly with intracellular L-cysteine levels. To test this model,Smulski et al. (1994) constructed a cysJ-luxCDABE fusion in E. coli. Theyfound that sulfur concentrations of 6 to 26 mM induced a dose-dependentresponse and that the reporter is controlled by cysB, the physiologicalregulator. The induction time (time elapsed before the increase in lightoutput) and the peak height (the amount of light observed) correlated withthe degree of starvation. In addition, cells challenged with growth on thepoor sulfur source L-djenkolic acid, ranging from 420 to 6.6 mM, resultedin a dose dependent response yielding inductions that were thousands offold above those in high cysteine growth conditions. However, the responseoccurred many minutes later and was less in terms of light produced,as compared with growth on limiting sulfate leading to the conclusionthat high L-djenkolic acid levels mimic, but do not duplicate, sulfurstarvation.

4.5. Multi-Nutrient Starvation Response

A tripartate microbial reporter system has been described (Standing et al.,2003) that utilizes three lux-marked P. fluorescens strains. This system isthe first to simultaneously report levels of C, N and P in the rhizosphere.The reporters were tested in a soil solution spiked with glucose, ammoniumnitrate and sodium phosphate. They report that the light output in responseto glucose concentrations between between 3 and 120 �g ml�1 was directlyproportional, but non-linear. The response of the N-starvation reportershowed differential sensitivity to ammonia versus nitrate. Phosphatedetection showed a robust linear inverse relationship between light outputand phosphate concentration in the range of 0.001–0.61 mM.It has been shown by Nystrom and Neidhardt (1992) that the uspA

gene is induced under numerous stress conditions, among them carbon,nitrogen, phosphate and sulfate starvation, as well as damage by toxicants.In addition to uspA, the E. coli genome carries several paralogues:uspC, uspD, uspE, uspF which appear to be regulated, in part, by thestringent response. Some experiments support the hypothesis that the geneproducts of these paralogues participate in protecting the cell fromgenotoxic damage (Kvint et al., 2003), while other data show a broaderregulatory role in many different stress responses (Alderete et al., 2001).Interestingly, bacterial species that carry homologues to uspA often have


several different versions. They are found in all of the enterobacteriacaeaas well as in archaea and some gram positive genomes. It remains to bedemonstrated that in these species, the usp-like gene products have somerole in stress response. Thus, the E. coli uspA-luxCDABE fusion wouldappear to be a useful candidate to report starvation in general. The responseof this construct to toxicants was demonstrated by Van Dyk et al. (1995b)and responses to nutritional limitations have also been reported (Aldereteand Vollmer, 2000; Alderete et al., 2001). While the response of uspA is notas dramatic as more specific members of stress regulons in terms of totalfold-induction (Nystrom and Neidhardt, 1992), its response to manydifferent types of stressors make it a ‘‘utility player’’ in the assay of mixturesof potential toxicants.Some bacteria have evolved complex, alternative life cycles (in lieu of

exponential growth) to adjust to the stress of nutrient limitation. Forexample, sporulating genes of bacteria such as B. subtilis are activated inresponse to carbon and nitrogen starvation (Sonenshein, 2000). While earlyspoO genes have been studied extensively, there has not been an effort touse these as stress-response biosensors. Similarly, Myxococcus xanthusrequires a starvation for carbon in order to initiate its developmentalfruiting body formation program. It is sensitive to the accumulation ofa critical combination of amino acids in the medium, possibly a celldensity signal (called A signal – Shimkets, 2002) in order to commence thedevelopmental transition. While numerous genetic analyses have yieldedseveral important signal transduction pathways in the early stages offruiting body formation, there has been no effort to apply this knowledgeto construct relevant biosensors. Such sensors may be of environmentalimportance, since both Gram positive bacilli and gram negativemyxobacteria occupy important niches in the soil.

5. PANELS AND ARRAYS

As highlighted in previous sections, microorganisms have numerousregulatory systems that respond by altering transcription when cellsencounter stress and starvation conditions. Monitoring of these individualstress responses is used to detect specific types of environmental stresses.This concept has been extended by monitoring numerous transcriptionalresponses in parallel, using multiple stress responsive biosensors. Thus,the toxicity of a chemical or environmental sample is characterized by


the stress responses that are induced or not induced. Such patterns of geneexpression yield a fingerprint characteristic of biological modes of action.

5.1. Panels of Stress Responsive Gene Fusions

Stress responsive gene reporter gene fusions in E. coli and Salmonellatyphimurium were developed in two early applications for characterizingchemical and environmental modes of action by stress fingerprinting.In 1994, a panel of 15 genetic fusions to the V. fischeri luxCDABE reporterthat respond to 12 different stress conditions was described (Van Dyk et al.,1994a). Similarly, in 1995, a group of 16 lacZ fusion strains called the‘‘ProTox’’ test was described (Orser et al., 1995). As expected, the biosensorcells containing these gene fusions, whether using the luxCDABE or lacZreporter, responded differentially to damage agents. For example,irradiation with ultraviolet light highly induced expression of arecA-luxCDABE (SOS response) and moderately induced expression ofgrpE-luxCDABE (heat shock response), but did not induce elevatedexpression of a luxCDABE fusion to katG (OxyR regulated peroxideresponse) (Van Dyk et al., 1994a). Furthermore, the pattern, or fingerprint,of expression responses was indicative of the stresses sustained by thecell from chemical treatment. Accordingly, methyl methanesulfone, aDNA alkylating agent, induced expression of ada-lacZ, known torespond to DNA alkylation, and dinD-lacZ (SOS response), but not ofany other stress responses represented in the ‘‘ProTox’’ panel, consistentwith the specificity of this agent for DNA (Orser et al., 1995). In yetanother demonstration of the utility of the approach, several EPA prioritypollutants were tested for induced responses using a panel of fivestress-responsive luxCDABE gene fusions (Belkin et al., 1997). Relatedmolecules, such as 2-nitrophenol and 4-nitrophenol, resulted in very similarpatterns of induction among the gene fusions, while compounds withunrelated structures, such as methylene chloride, induced a distinct pattern.Thus, the concept of distinguishing between biological modes of action bypatterns of induced stress responses was demonstrated.Later work incorporated numerous improvements to fine-tune the

panels of stress responsive fusions used to characterize chemically inducedstress responses. For example, careful selection of stress responses wasused to optimize a panel that distinguished the modes of action of variousclasses of antibiotics (Bianchi and Baneyx, 1999). Improvements in thereporter gene came from the use of thermostable luxCDABE from P.luminescens that improved the temperature range at which a panel of


luxCDABE gene fusions could be utilized (Van Dyk et al., 2000). Otherreporters have also been used in panels of stress responsive gene fusions.For example, several oxidative stress responsive gfp fusions in E. coliresponded specifically to various types of reactive oxygen species and wereused to characterize oxidative effects of antitumor drugs (Albano et al.,2001). Improvement in sensitivity to low chemical concentrations wasaccomplished by treatment with polymyxin B to permeabilize the outermembrane or use of an E. coli host strain carrying a tolC mutation thatresults in loss of function of several efflux pumps (Shapiro and Baneyx,2002; Van Dyk et al., 2000). Furthermore, modulating plasmid copy numberor placing gene fusions at single copy in the chromosome was found tobe useful in tuning basal levels of gene expression to the same range forall gene fusions in a panel (Van Dyk et al., 2000).If panels of stress responsive gene fusions are applied where profiles

of compounds likely to be present in mammals are required, it is importantto consider protocol modifications that model xenobiotic metabolism inmammalian systems. Thus, sample pretreatment with the S9 fraction ofrat liver homogenate, as incorporated in the Ames test for mutagenicity(Ames et al., 1975; Maron and Ames, 1983), has been used. As expected,changed response profiles due to metabolism of the compounds to otheractive forms were observed (Belkin et al., 1997; Van Dyk et al., 2000).For example, aflatoxin B1 prior to S9 treatment modestly inducedexpression of a reporter of the SOS response and also induced expressionof an inaA-luxCDABE reporter regulated by SoxS/MarR/Rob (Van Dyket al., 2000). Following S9 treatment, the SOS response was greatlyenhanced consistent with the increased genotoxicity of aflatoxin B1 epoxidethat forms following metabolic activation (McLean and Dutton, 1995).Interestingly, the inaA-luxCDABE response was greatly decreased afterS9 treatment suggesting that this response is only induced by the non-activated form of aflatoxin B1. It is important to note that addition ofS9 liver homogenate, in some cases, interferes with detection of reportersignals and thus further protocol modifications may be required to allowefficient detection of responses (Dreier et al., 2002).A noteworthy environmental application of a panel of stress

responsive gene fusions is monitoring wastewater treatment (Belkin,1998a). Municipal and industrial wastewater treatments are often primarilybiological processes. Thus, the health of the microorganisms responsiblefor breakdown of undesirable compounds in the wastewater is of concern.In one example of applying panels of stress responsive fusions to moni-toring wastewater treatment, the influent and effluent of an industrialwastewater treatment facility were tested with a panel of four stress


responsive gene fusions (Belkin et al., 1996b). The water flowing into thetreatment plant induced expression of recA-luxCDABE (SOS regulon) andmicF-luxCDABE (SoxR regulon), suggesting that chemicals causing DNAdamage and oxidative damage were present in the complex mixture. Thewater flowing out of this wastewater treatment plant did not induce anystress responses, consistent with the efficacy of the treatment in remov-ing these harmful agents. Thus, the potential for monitoring the resultsof wastewater treatment using stress responsive reporter strains wasestablished. Another possible application of stress panels in wastewatertreatment is in characterization of the influent to detect and predictwhen water coming into the system has high toxic potential that couldlead to upset due to inhibition or death of the microorganisms in thetreatment plant.The stress panel approach, as implemented to date, relies on the

abundance of fundamental data on stress responses that has beendeveloped in well-studied organisms, such as E. coli. Thus, rationalinterpretations of the stress response patterns, the regulatory networksresponsible for the observed patterns, and the biological implications of theinduced responses are often possible. Nevertheless, a strictly empiricalanalysis that relies only on the pattern of induced responses withoutconsidering the microbial physiology underlying the data from stresspanels has also been shown to be useful. For example, clusteringalgorithms used to analyze the induction patterns from a gene fusionpanel successfully grouped chemicals into common modes of action (Ben-Israel et al., 1998).

5.2. DNA Arrays

The opportunity to obtain abundant data with which to employboth rational and empirical analyses is provided by highly parallelhybridization assays using DNA arrays (Rhodius et al., 2002). Since thepublications in 1999 demonstrating genome-wide transcription profiling inE. coli (Richmond et al., 1999; Tao et al., 1999) and Mycobacteriumtuberculosis (Wilson et al., 1999), use of this methodology has rapidlyexpanded to numerous bacterial species. DNA microarrays can beconstructed for any organisms for which there is full or partial genomicsequence available, or random libraries of chromosomal DNA segments.DNA array technology is unlikely to be applied directly to routineenvironmental monitoring because of the many manipulations needed toproduce experimental results. Nonetheless, this technology will have a


dramatic impact on the applications of stress response monitoring as it isused for discovery of novel responses and to define regulons.Instructive examples are provided by recent work with DNA arrays

in defining genome-wide transcriptional alterations upon oxidativestress in E. coli and B. subtilis. Responses of E. coli to treatment withthe redox cycling agent paraquat (methyl viologen), which resultsin intracellular generation of superoxide, were studied using DNAmacroarrays on nylon membranes (Pomposiello et al., 2001). Expressionof 66 genes was found to be significantly activated. As expected, theseincluded genes in the SoxRS regulon; 9 of 16 previously known SoxS-activated genes were upregulated. An additional experiment involvingoverexpression of SoxS from a plasmid led to identification of 37 activatedgenes, of which 14 were in common with the paraquat experiment. Thus,genes that are candidate new members of the SoxRS regulon were found,helping to further define the regulon. Furthermore, genes activated byparaquat treatment that are not part of the SoxS regulon are also ofinterest because they may define novel responses to oxidative stress.Although the regulatory circuitry involved in the non-SoxS activation havenot been defined, it is reasonable to postulate that another mechanismcontrols these transcriptional responses to paraquat. If so, monitoringexpression of genes in such a regulatory circuit may be useful to distinguishparaquat mode of action from that of other compounds that induce theSoxRS regulon but do not induce this putative novel regulatory circuit.It should be noted that some limitations of DNA array experiments areevident from this study. Clearly the upregulated genes must be consideredan underestimate, as several known members of the SoxS regulon werenot found. Additionally, it is important to consider that the statisticalanalyses needed to determine upregulated and downregulated genes willresult in false positives as well as false negatives. Thus, any results from aDNA array study should be confirmed by other methods.Responses of B. subtilis to treatment with the oxidizing agents

hydrogen peroxide and tert-butyl peroxide analyzed with DNA microarrayson glass slides (Helmann et al., 2003) yielded interesting results thatcould be applied to sensor development. Genes in the PerR regulon werefound to be induced by low (8 mM) and higher (58 mM) levels of hydrogenperoxide and to a lesser extent by treatment with tert-butyl peroxide.In contrast, genes in the general stress response, sB regulon, wereupregulated by tert-butyl peroxide and by the high, but not low, level ofhydrogen peroxide. In an important control experiment, treatment with tert-butanol, the corresponding alcohol of tert-butyl peroxide, upregulatedthe sB regulon, but not the PerR regulon. Thus, the sB response is unlikely


to be responding to the oxidative damage caused by tert-butyl peroxide.Fascinatingly, a single gene in B. sublitis, ohrA, stood out by its very highlevel of upregulation in response by tert-butyl peroxide treatment thatcontrasted with a lack of response to tert-butanol. The expression of thisgene, which is known to be regulated by OhrR, was also unresponsive tohydrogen peroxide treatments. Thus, monitoring the expression of thisgene would be an excellent candidate for developing a sensor specificallyresponsive to organic peroxides. As microarray data accumulate, other suchcandidate genes useful for sensor development will become evident.

5.3. Gene Fusion Arrays

As an alternative to DNA arrays for genome-wide gene expressionanalysis in bacteria, gene fusion arrays also have the advantage ofpotential direct application to environmental monitoring. The LuxArray inE. coli consists of a set of luxCDABE reporter gene fusions to 689 ofthe 2584 predicted transcriptional units in E. coli (Van Dyk et al., 2001a).Thus, 27% of the transcription units of E. coli are represented. The genefusions in the LuxArray were obtained beginning with a collection madeby joining random segments of the E. coli chromosome to a bioluminescentreporter (Van Dyk and Rosson, 1998). Each of 8000 random genefusions was sequenced to define the ends and orientation of thechromosomal segment upstream of the reporter genes yielding 5000mapped gene fusions (Van Dyk et al., 2001b). The final step was cul-ling non-functional fusions and redundancies inherent in a randomapproach. The random nature of the set of gene fusions in the LuxArraymakes it likely that most global regulatory circuits will be represented byone or more members, thus making the Lux Array useful for stress responsemonitoring. Additionally, novel expression events may be discoveredbecause these arrays contain fusions to many previously uncharacterizedgenes and operons. Recently, an array of randomly generated biolu-minescent gene fusions in S. typhimurium was described (Goh et al., 2002),thus showing that this technology can be successfully appliedin other bacteria.Reporter gene arrays and hybridization experiments represent indepen-

dent methods of gene expression analysis. To the extent that gene expressionis controlled at the level of transcriptional initiation, these methodsshould yield equivalent results. A major advantage of using arrays of bio-luminescent reporter genes is the non-invasive monitoring of lightproduction that allows detailed kinetic characterizations. Additionally, as


stress responsive fusions are identified, the strains carrying these fusionscan be developed directly into whole cell biosensors. Furthermore,innovative methods for implementation, such as immobilization in latexcopolymers (Lyngberg et al., 1999) or in conjunction with microlumi-nometers (Bolton et al., 2002), make possible the direct use of biolumi-nescent arrays for environmental applications.Currently, two methods of perturbing and collecting data in a parallel

fashion have been implemented for the LuxArray. One approachinvolves ‘‘printing’’ the set of strains containing the gene fusions in ahigh density ordered array to solidified agar (Van Dyk et al., 2001a).Subsequently, images of the signal generated from reporter constructs arecollected with a CCD camera and quantitated by pixel density. Thisapproach obviously relies on the use of reporter genes that generate avisible signal. The other method for working with the LuxArray entailsgrowth and assay in microplates and is aided by laboratory automation(Rozen et al., 2001; Van Dyk, 2002). Such a method is also applicable toreporters that require manipulation, such as cell lysis and substrate addi-tion, to generate a detectable signal.Among other things, the LuxArray has been applied to analyze

transcriptional responses that occur when E. coli is put into seawater(Rozen et al., 2001). The majority of the 22 upregulated gene fusions werein the RpoS regulon. This response was shown to be due primarily tonutrient limitation in seawater. Other responses were to the alkaline andosmotic shocks encountered. Interestingly, a luxCDABE fusion to thepromoter region of a gene lacking known function, yjbG, with over a 400-fold response ratio, was the most highly induced gene fusion.The upregulation of the gene fusion, which was not dependent on RpoS,was shown to be due in large part to the osmotic shock. The regulatorycircuitry controlling this response is not known. Nonetheless, the straincarrying the yjbG-luxCDABE gene fusion can serve as a biosensor ofosmotic shock.

5.4. From Arrays to Specialized Panels

As discussed above, DNA and reporter gene arrays have been used todiscover novel stress responses that may be useful for environmentalmonitoring. As these technologies are widely applied to characterizetranscriptional alterations to chemicals and other adverse environmentalconditions, discoveries of novel and useful stress responses will continueto be made. It is envisioned that, as expression response profiles of


toxicants are placed in databases and analyzed, key signature transcriptionalresponses to environmental stresses of interest will be identified. Thus,selection of a much smaller subset of genes to monitor will provideequivalent information to full genome arrays. These can then be developedinto selected panels of gene fusions optimized for detection of specificchemical classes or specific stress conditions. Use of such smaller subsets ofbiosensors will then allow rapid throughput of sample analysis.

6. FUTURE TRENDS

The past decade has witnessed the development of cellular biosensors thatrespond with sensitivity and specificity to environmental factors. Questionsabout toxicants in the environment can be answered in a manner thatprovides early alerts, and physiologically relevant data. Reporters suchas lux allow for non-invasive, continual real-time reporting, whichis useful for assessing processes such as wastewater treatment. The use ofbacterial populations results in an ensemble average in the reportedactivity and the range of response of a given population can be an indicatorof the diversity of physiological states, morphological variance andgenetic heterogeneity of the members of that population. More sensitivedetection in the future may allow for the capture and measurement ofoutput from a single cell (Leveau and Lindow, 2002). As more newinformation about bacterial physiology is revealed in genome-wide analyzes,individual whole cells as well as populations of these biosensors willcontinue to be improved to assess environmental factors. Furthermore, it isconceivable that biofilms containing stress-responsive reporters will haveunique applications in flow chambers and related systems.While most cellular biosensors to date have been constructed with

regulatory circuits found in nature, future applications will not belimited by this constraint. Mutagenesis to change the specificity ofregulatory proteins will enable biosensor development for detectionof chemicals that are not natural inducers. For example, mutagenesisof DmpR, a regulatory protein for the phenol degradation pathway ofPseudomonas sp. strain CF600, was used to increase the range of phenolicmolecules detected (Wise and Kuske, 2000). Similarly, shuffling of effectorbinding domains of two related regulatory proteins, DmpR andXylR, yielded derivatives with broadened or narrowed response profiles(Skarfstad et al., 2000). Likewise, a combinatorial library made fromN-terminal domains of DmpR, XylR and TbuT in the XylR structure, was


found to contain members that responded to compounds that weresubstantially larger in size or with significantly different electronic propertiesthan the natural XylR effectors (Garmendia et al., 2001).As individual cellular biosensors, panels, and arrays are developed with

increasing utility for particular applications, the means to bring thesesensors to the field will also advance. Improvements in cellularimmobilization and lyophilization will result in better methods to couplewhole cells with sensing devices. Likewise, miniaturization of devices willfacilitate highly parallel, multimodal detection strategies. Miniaturizationand improved robustness will also enhance detector portability.Furthermore, remote signal sensing will allow widespread biosensordeployment.

ACKNOWLEDGMENTS

We wish to acknowledge the contributions of past and present membersof our respective research groups and our collaborators at SwarthmoreCollege and the DuPont Company. Work in A.C.V.’s laboratory has beensupported by grants from the National Science Foundation, the HowardHughes Medical Institute, the American Society for Microbiology,and Merck/AAAS.

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Bacterial Naþ- or Hþ-coupled ATP SynthasesOperating at Low Electrochemical Potential

Peter Dimroth1 and Gregory M. Cook2

1Institut fur Mikrobiologie, Eidgenossische Technische Hochschule, ETH-Zentrum,

Schmelzbergstrasse 7, CH-8092 Zurich, Switzerland2Department of Microbiology, Otago School of Medical Sciences, University of Otago,

P.O. Box 56, Dunedin, New Zealand

ABSTRACT

In certain strictly anaerobic bacteria, the energy for growth is derivedentirely from a decarboxylation reaction. A prominent example isPropionigenium modestum, which converts the free energy of thedecarboxylation of (S)-methylmalonyl-CoA to propionyl-CoA(�G� ¼�20.6 kJ/mol) into an electrochemical Naþ ion gradientacross the membrane. This energy source is used as a driving force forATP synthesis by a Naþ -translocating F1F0 ATP synthase. Accordingto bioenergetic considerations, approximately four decarboxylationevents are necessary to support the synthesis of one ATP. This uniquefeature of using Naþ instead of Hþ as the coupling ion has made thisATP synthase the paradigm to study the ion pathway across themembrane and its relationship to rotational catalysis. The membranepotential (� ) is the key driving force to convert ion translocationthrough the F0 motor components into torque. The resulting rotationelicits conformational changes at the catalytic sites of the peripheralF1 domain which are instrumental for ATP synthesis. Alkaliphilicbacteria also face the challenge of synthesizing ATP at alow electrochemical potential, but for entirely different reasons. Here, thelow potential is not the result of insufficient energy input from substrate



DOI 10.1016/S0065-2911(04)49004-3

degradation, but of an inverse pH gradient. This is a consequence ofthe high environmental pH where these bacteria grow and the necessityto keep the intracellular pH in the neutral range. In spite of thisunfavorable bioenergetic condition, ATP synthesis in alkaliphilic bacteriais coupled to the proton motive force (�mHþ ) and not to the muchhigher sodium motive force (�mNaþ ). A peculiar feature of the ATPsynthases of alkaliphiles is the specific inhibition of their ATP hydrolysisactivity. This inhibition appears to be an essential strategy for survivalat high external pH: if the enzyme were to operate as an ATPase,protons would be pumped outwards to counteract the low �mHþ , thuswasting valuable ATP and compromising acidification of thecytoplasm at alkaline pH.

1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1762. ATP synthesis in anaerobic bacteria at low electrochemical potential . . . . . . 179

2.1. Sodium ion cycles in bacteria and rationale for the presence ofa sodium-translocating F1F0 ATP synthase. . . . . . . . . . . . . . . . . . 180

2.2. �mNaþ generation by sodium-translocating decarboxylases . . . . . . . 1832.3. Sodium-translocating F1F0 ATP synthase. . . . . . . . . . . . . . . . . . . 187

3. Alkaliphilic bacteria growing at low �mHþ . . . . . . . . . . . . . . . . . . . . . 2003.1. Sodium and proton cycling in alkaliphiles . . . . . . . . . . . . . . . . . . 2013.2. pH Homeostasis in alkaliphilic bacteria and growth at low �mHþ . . . . . 2023.3. Mechanisms to synthesize ATP at low �mHþ using a

proton-coupled ATP synthase . . . . . . . . . . . . . . . . . . . . . . . . . 2033.4. Alkaliphilic-specific amino acid motifs in the atp operons of

mesophilic and thermoalkaliphilic bacteria . . . . . . . . . . . . . . . . . . 2053.5. The F1F0 ATP synthases from alkaliphilic bacteria show latent ATP

hydrolysis activity: a specific adaptation for growth at alkaline pH?. . . . 2073.6. Regulation of ATP hydrolysis activity by bacterial F1F0

ATP synthases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210

1. INTRODUCTION

Bacteria are remarkably versatile organisms degrading a wide varietyof organic substrates under diverse environmental conditions. Anaerobicbacteria, in general, gain much less energy from substrate degradationcompared to their aerobic counterparts, and in some species the degradation

176 PETER DIMROTH AND GREGORY M. COOK

of more than one substrate molecule is required to fulfill energeticrequirements for the synthesis of one ATP. A prominent example isPropionigenium modestum, which grows from the fermentation of succinateto propionate and CO2 (Schink and Pfennig, 1982; Dimroth and Schink,1998). The free energy of this reaction is about �20 kJ/mol whereasapproximately �70 kJ/mol is required to support ATP synthesis in growingbacteria (Thauer et al., 1977). To solve this apparent paradox, 3–4 succinatemolecules must be converted into propionate before one ATP moleculecan be synthesized. Accordingly, the stoichiometric formation of energy-rich precursors with a group transfer potential at least as high as that ofthe phosphoric anhydride bond of the ATP molecule itself is thermo-dynamically not feasible and chemiosmotic processes are thereforemandatory for energy conservation.The catabolism of succinate by P. modestum is shown in Fig. 1 and

described in more detail below. The only exergonic reaction in the wholepathway is the decarboxylation of (S)-methylmalonyl-CoA to propionyl-CoA and CO2. Not surprisingly, therefore, the decarboxylase is amembrane-bound enzyme that converts the free energy of the C–C bondcleavage into a transmembrane electrochemical gradient of sodium ions(�mNaþ ) (Hilpert et al., 1984; Bott et al., 1997; Dimroth, 1997). The iongradient subsequently drives ATP synthesis by a Naþ -translocatingF1F0 ATP synthase (Laubinger and Dimroth, 1988). This enzyme is aspecial member of the family of F1F0 ATP synthases which typically useprotons as their coupling ions. Due to its unique experimental options,the sodium-translocating ATP synthase has become the prototype forinvestigations on the ion path across the membrane-bound F0 motordomain. These studies have gained insight into the coupling ion bindingsites and their membrane-buried location on the c-subunit ring. It isproposed that two different channels connect the sites with the two differentsurfaces of the membrane. Glimpses into the torque-generating mechanismof this rotary motor have also been obtained and will be described inmore detail below.Alkaliphilic bacteria synthesize ATP by proton-coupled F1F0

ATP synthases (Hicks and Krulwich, 1990; Hoffmann and Dimroth,1990, 1991a; Cook et al., 2003). Like anaerobic bacteria, the alkaliphilessynthesize ATP at low free energy content of the proton motive force(�mHþ ), but for quite different reasons. These aerobic organisms havean abundant energy input from the degradation of the organic substratesand the limited energy supply of the �mHþ results from the necessity ofkeeping the internal pH near neutral in order not to compromisethe viability of the cells. At high external pH, a �pH of þ 120 mV may

LOW ELECTROCHEMICAL POTENTIAL 177

thus result, which opposes the membrane potential (� ). Hence, atmeasured membrane potentials of not more than �210 mV, the total �mHþ

may be as low as �80 mV, which makes ATP synthesis by a conventionalmechanism difficult to reconcile (Hoffmann and Dimroth, 1991b; Guffantiand Krulwich, 1992). Interestingly, the alkaliphiles maintain a largesodium motive force (�mNaþ > �250 mV) and use this energy sourcefor substrate uptake or the rotation of the flagella, but not for the synthesisof ATP. Models have been proposed for how these bacteria can synthesizeATP in spite of the low �mHþ , but these still await experimentalverification (Krulwich, 1995; Krulwich et al., 1998a). To cope with thebioenergetic challenge, the ATP synthases of alkaliphilic bacteria may

Figure 1 Energy metabolism and sodium ion cycling in P. modestum.(A) Succinate uptake system (the transport mechanism is unknown); (B) succinatepropionyl-CoA: CoA transferase; (C) methylmalonyl-CoA mutase; (D) methylmalonyl-CoA epimerase; (E) methylmalonyl-CoA decarboxylase; (F) Naþ translocating F1F0

ATP synthase.


have acquired unique structural and functional features. A peculiarity isthe specific blockade of the ATP synthase in the ATP hydrolysis direction(Hoffmann and Dimroth, 1990; Cook et al., 2003). This unusual propertyof the enzyme may be a necessity to survive at alkaline pH: if the ATPsynthase were to operate in reverse as an ATPase it would pump protonsoutwards, thereby raising the internal pH to potentially intolerable values.Moreover, at low �mHþ in alkaliphilic bacteria, the natural tendencyof a freely reversible enzyme will be to operate as an ATPase thus wast-ing valuable intracellular ATP. In this chapter, we will elaborate on thisproperty of the enzyme and propose a mechanism as to how the ATPhydrolysis activity may be regulated.

2. ATP SYNTHESIS IN ANAEROBIC BACTERIA AT LOWELECTROCHEMICAL POTENTIAL

Many anaerobic bacteria perform a chemiosmotic ATP synthesismechanism like their aerobic counterparts. In special cases when the energyfrom substrate degradation is not sufficient to support the synthesis ofstoichiometric amounts of ATP, the chemiosmotic ATP synthesismechanism is obligatory. The free energy derived from the degradation ofseveral substrate molecules is stored in the electrochemical ion gradientover the membrane which thus becomes sufficient to drive the synthesisof ATP. A prominent example for this type of energy metabolism isthe fermentation of succinate by P. modestum as shown in Fig. 1 (Hilpertet al., 1984; Dimroth and Schink, 1998). The catabolism starts with theuptake of succinate into the cells by an unknown mechanism.After activation to succinyl-CoA, this undergoes rearrangement of thecarbon skeleton and isomerization to yield (S)-methylmalonyl-CoA, whichis decarboxylated by a membrane-bound biotin-containing sodium ionpump to propionyl-CoA. The final step is the conversion of propionyl-CoA to propionate which is synchronized with the activation of a newsuccinate molecule to succinyl-CoA. The key enzyme for the bioenergeticsof these bacteria is the methylmalonyl-CoA decarboxylase Naþ pump(Hilpert and Dimroth, 1983; Bott et al., 1997). It belongs to the sodium iontransport decarboxylase family of enzymes which all utilize the energy of achemical decarboxylation reaction to pump Naþ ions across the membrane(Dimroth, 1997; Dimroth and Schink, 1998). The free energy changeof these reactions is comparatively small, but sufficient for conversion intoan electrochemical gradient of sodium ions.


The electrochemical sodium ion gradient thus established is the onlyenergy source for ATP synthesis in P. modestum. The Naþ ion cycle iscompleted during ATP synthesis by the Naþ -translocating F1F0 ATPsynthase (Laubinger and Dimroth, 1988). The number of decarboxylationevents required for the synthesis of one ATP can be estimated from thesodium ion stoichiometries of the decarboxylase and the ATP synthase.The ATP synthase operates at a Naþ to ATP stoichiometry of 3.7(see below). Methylmalonyl-CoA decarboxylase couples one decarboxyla-tion reaction to the electrogenic transport of one Naþ ion and theelectroneutral transport of a second Naþ ion (in exchange for a proton)across the membrane (Hilpert and Dimroth, 1991; Di Berardino andDimroth, 1996). As ATP synthesis requires the electrogenic uptake of3.7 Naþ ions, this number of succinate molecules has to be converted topropionate to synthesize one molecule of ATP. This calculation is based onthe provision that the ATP synthase is the only consumer of the membranepotential in these bacteria. The stoichiometry is close to that expectedfrom bioenergetic considerations. The free energy of methylmalonyl-CoAdecarboxylation is approximately �20 kJ/mol (Schink and Pfennig, 1982)and the free energy of ATP synthesis in a growing bacterium is estimatedto be about 70 kJ/mol (Thauer et al., 1977). Hence, on this basis,3.5 decarboxylation events would lead to the synthesis of one ATP. Theelectroneutrally translocated Naþ ions cannot be used to drive ATPsynthesis, but might energize other membrane reactions, e.g. the uptake ofsuccinate into the cells.

2.1. Sodium Ion Cycles in Bacteria and Rationale for the Presenceof a Sodium-Translocating F1F0 ATP Synthase

Sodium ion cycles in bacteria are widespread but primary sodium ionpumps and sodium-translocating ATP synthases are rare. Aerobic bacteriausually energize their membrane by a proton-extruding respiratorychain, and a sodium ion gradient is established as a secondary event viaa sodium/proton antiporter. These antiporters perform a number ofimportant physiological functions: they reduce the cytoplasmic Naþ

concentration to nontoxic levels, they regulate the cytoplasmic pH, andthey generate the sodium ion concentration gradient required for theuptake of certain substrates into the cells (Padan et al., 2001).Some Vibrio species living in environments of high Naþ content

synthesize a primary sodium pump which extrudes Naþ ions from thecells in the respiratory chain segment between NADH and ubiquinone


(Hayashi et al., 2001). This unique Naþ pump is called Naþ -NQR.Although the overall electron transfer from NADH to ubiquinone is thesame for the Naþ -NQR and for complex I and both are electrogenic ionpumps, these two enzymes are not related phylogenetically. The Naþ -NQRconsists of seven different subunits and contains FAD, FMN and one Fe/Scluster, while bacterial complex I has 14 subunits and contains FMN,several Fe/S clusters and tightly-bound quinones as prosthetic groups.The Naþ gradient generated by the Naþ -NQR is used for nutrient uptakesystems and for the rotation of the Naþ -dependent flagellar motor(Yorimitsu and Homma, 2001). The ATP synthase of Vibrio alginolyticusis a proton-coupled enzyme (Krumholz et al., 1990). This is notunreasonable, since in species like Vibrio cholerae the residual respiratorychain consists of the proton-translocating complexes bc1 and cytochromeoxidase. Therefore, the presence of a primary Naþ pump in a bacteriumdoes not necessarily imply that the ATP synthase is a Naþ -coupled enzyme.One should keep in mind, however, that the electrical component ofthe driving force established by the Naþ -NQR can be utilized by theATP synthase independent of whether this is a Naþ - or Hþ -dependentenzyme.Inspection of the genome sequence of Thermotoga maritima, a

hyperthermophilic anaerobic marine bacterium, reveals the presence ofa Naþ -NQR and of complex I. This bacterium performs anaerobicrespiration with Fe3þ as the terminal electron acceptor (Vargar et al.,1998) and, therefore, the �mNaþ generated by the Naþ -NQR (orcomplex I) is probably not complemented by an electrochemical Hþ

gradient formed by another respiratory pump. Hence, in order to drive ATPsynthesis by an Hþ -coupled F1F0 ATP synthase, the Naþ gradient wouldhave to be converted into a proton gradient. While typical Naþ /Hþ

antiporters convert an electrochemical Hþ gradient into a Naþ gradient viaproton uptake and Naþ extrusion (Padan et al., 2001), we are not aware ofa transporter operating in reverse (i.e. taking up Naþ and extruding Hþ ).Such an antiporter might be quite precarious for a bacterium exposedto varying Naþ concentrations in the environment. A sudden increasein the environmental Naþ concentration could lead to massive protonextrusion from the cells and an increase of the cytoplasmic pH tointolerable levels. Hence, a Naþ -coupled ATP synthase could bemandatory, if the only energy source for ATP synthesis is an electrochemicalNaþ ion gradient established by a primary Naþ pump. Recent workfrom our laboratory has demonstrated ATP-dependent 22Naþ uptake ininverted membrane vesicles of T. maritima that is sensitive to DCCD.Moreover, ATPase activity of inverted membrane vesicles is stimulated by


sodium ions and T. maritima harbors the Naþ binding signature on its csubunits suggesting it is a member of the family of Naþ -translocatingF1F0 ATP synthases (S. Ferguson, S. Keis, P. Dimroth and G.M. Cook,unpublished).In two other bacteria, the Naþ cycle is initiated by an oxaloacetate

decarboxylase Naþ pump. In Klebsiella pneumoniae, citrate is degradedanaerobically by the citrate fermentation pathway (Dimroth et al., 2001).The genes encoding the pertinent enzymes are clustered on the genome inthe cit operon and are induced via a two-component–regulatory-systemsensing citrate, Naþ , and anaerobic conditions (Bott et al., 1995;Bott, 1997). The proteins derived from these genes consist of a citratetransporter (CitS), citrate lyase and oxaloacetate decarboxylase. Alsoencoded by this operon are enzymes for the biosynthesis of thediphosphoribosyl dephospho-CoA prosthetic group of citrate lyase andfor its conversion into the catalytically active acetyl thioester derivative(Schneider et al., 2000a, 2000b). The uptake of citrate by CitS occurs asa cotransport with Naþ and Hþ (Lolkema et al., 1994; Pos and Dimroth,1996). Citrate is subsequently cleaved by citrate lyase to acetate and oxa-loacetate. The latter is decarboxylated by the oxaloacetate decarboxylaseNaþ pump to pyruvate and CO2 generating an electrochemical gradientof Naþ ions across the membrane (Dimroth, 1987, 1997). Pyruvate isfurther degraded by pyruvate/formate lyase to acetyl-CoA and formate.The conversion of acetyl-CoA to acetate is accompanied by ATPsynthesis via substrate level phosphorylation. In this organism, Naþ ionsare pumped outwards by the oxaloacetate decarboxylase and the Naþ cycleis completed by CitS catalyzing citrate uptake in symport with Naþ . It isnot surprising, therefore, that the ATP synthase of K. pneumoniae iscoupled to protons. The driving force for the ATP synthase derivesfrom the membrane potential generated by the oxaloacetate decarboxylaseNaþ pump and possibly the pH gradient arising from the electroneutralextrusion of the fermentation end products acetate and formate togetherwith Hþ .The fermentation of tartrate by Ilyobacter tartaricus is similar to that

of citrate by K. pneumoniae and differs mainly in the initial step bywhich the substrate is converted to oxaloacetate (Schink, 1984). It is ofinterest, therefore, that the F1F0 ATP synthase of I. tartaricus is coupledto Naþ ions (Neumann et al., 1998) while that of K. pneumoniae is coupledto Hþ . The Naþ -translocating ATP synthase of I. tartaricus utilizesthe electrochemical Naþ gradient generated by the oxaloacetate decarboxy-lase Naþ pump to drive ATP synthesis, similar to ATP synthesis in P.modestum. The additional Naþ ions, which are translocated electroneutrally


by the oxaloacetate decarboxylase (see above), cannot be used for ATPsynthesis but could recycle in a cotransport with the substrate tartrateinto the cell. It is interesting that enterobacteria like K. pneumoniae orEscherichia coli have the capacity to synthesize still another primaryNaþ pump, the Naþ -translocating complex I of the respiratory chain(Krebs et al., 1999; Steuber et al., 2000; Gemperli et al., 2002). This enzymespecifically pumps Naþ but not Hþ upon electron transfer from NADHto ubiquinone at a Naþ to electron stoichiometry of 1.0. As theenterobacterial complex I is expressed preferentially under anaerobicconditions (Tran et al., 1997), its Naþ pumping activity may be importantin anaerobic respirations, but the physiological significance of this Naþ

pump is still to be explored.Acetobacterium woodii is another anaerobic bacterium with a Naþ -

translocating F1F0 ATP synthase (Muller et al., 2001b). The presence ofthis enzyme in this acetogenic bacterium suggests that a �mNaþ generatingenzyme is also present in the cytoplasmic membrane to complete the Naþ

cycle. Although this has not yet been identified, a likely candidate isthe tetrahydromethanopterin coenzyme M methyltransferase which actsas a primary Naþ pump in methanogenic archaea (Gottschalk andThauer, 2001).Taken together, it appears that the presence of a Naþ -translocating

F1F0 ATP synthase is restricted to anaerobic bacteria harboring a primaryNaþ ion pump to act as a �mNaþ generator. This prediction seemsmandatory for bacteria like P. modestum, which gain all their ATP by achemiosmotic process. Bacteria like K. pneumoniae or I. tartaricus producepart of the ATP by substrate level phosphorylation and generate anelectrochemical Naþ ion gradient by the oxaloacetate decarboxylase.There is no rule for the presence of a Naþ -translocating or Hþ -translocating F1F0 ATP synthase in these bacteria because the enzymefrom Klebsiella is coupled to Hþ and that from Ilyobacter is coupled toNaþ . The ATP synthase seems to be proton-coupled if �mNaþ and �mHþ

generating pumps exist in parallel (e.g. the Naþ -NQR and proton pumpingrespiratory chain enzymes in V. cholerae).

2.2. DmNaþ Generation by Sodium-TranslocatingDecarboxylases

Oxaloacetate decarboxylase of K. pneumoniae was the first enzyme forwhich the energy conversion into an electrochemical Naþ ion gradient wasdemonstrated and has been the prototype for investigations of this ion


translocation mechanism (Dimroth, 1982; Dimroth et al., 2001). Othermembers of the family are methylmalonyl-CoA decarboxylase (Hilpertand Dimroth, 1983) (see above), glutaconyl-CoA decarboxylase ofAcidaminococcus fermentans and other glutamate-degrading anaerobicbacteria (Buckel and Semmler, 1983; Buckel, 2001), and malonatedecarboxylase of Malonomonas rubra (Hilbi et al., 1992; Dimroth andHilbi, 1997). Each of these decarboxylases catalyzes an essential step inthe respective degradation pathway and thereby pumps sodium ions out ofthe cell.As an example, Fig. 2 shows a cartoon on the subunit arrangement

within the oxaloacetate decarboxylase and their role in the reactionmechanism (Dimroth et al., 2001). The enzyme is composed of theperipheral a subunit (OadA), the membrane integral b subunit (OadB)and the small g subunit (OadG), which has a membrane anchor in theN-terminal tail and a hydrophilic C-terminal domain. OadG connectsOadA with OadB and has therefore an important role in the formation ofthe complex (Schmid et al., 2002b). OadA is composed of two domains:the N-terminal carboxyltransferase domain and the C-terminal biotindomain. These domains are connected by a short linker peptide consist-ing mostly of proline and alanine residues. This region of the protein is

Figure 2 Model showing the overall geometry of the oxaloacetate decarboxylase andfeatures of the catalytic events. B-H, biotin; B-CO�

2 , carboxybiotin; Lys, biotin bindinglysine residue. The carboxyltransfer reaction is catalyzed by the a subunit and thedecarboxylation reaction is catalyzed by the b subunit.


thought to be flexible to give the bound biotin residue the ability to movebetween the catalytic sites on OadA and OadB, respectively.The catalytic cycle starts with the transfer of the carboxylic

group from position 4 of oxaloacetate to the biotin prosthetic group onthe enzyme. The carboxybiotin thus formed switches from the carboxyl-transferase catalytic site on OadA to the decarboxylase site on OadB(Dimroth and Thomer, 1983; Dimroth and Thomer, 1988). Here thedecarboxylation takes place and the free biotin prosthetic group isregenerated. During the Naþ -dependent reaction, a periplasmically derivedproton is consumed and two sodium ions are translocated from thecytoplasm into the periplasm (Di Berardino and Dimroth, 1996).Essential residues for this coupled vectorial reaction have been

identified by site-specific mutagenesis of OadB (Jockel et al., 2000a,2000b). On the basis of these studies a model for the reaction mechanismwas proposed that is shown in Fig. 3 (Schmid et al., 2002a). Themodel predicts that a number of highly conserved and functionally indis-pensable residues on helices IV and VIII and region IIIa of OadB (Jockelet al., 1999) are involved in the ion translocation mechanism. In themechanistic model we propose that carboxybiotin formed at thecarboxyltransferase site of the enzyme switches to the decarboxylase siteon OadB where it forms a stable complex, possibly with the sidechain of R389 at the cytoplasmic surface of helix VIII. Evidence indicatesthat helix VIII aligns the Naþ and Hþ conducting channel (Jockel et al.,2000b; Schmid et al., 2002a; Wild et al., 2003). Evidently, the proton movingthrough this channel must reach the carboxybiotin to catalyzeits decarboxylation. A binding site near R389 would be well suited forthis purpose. According to our model (Fig. 3A) (Schmid et al., 2002a),the Naþ channel is initially open to the cytoplasm. In this conformation thetwo different sites are of high affinity (Km¼ 1 mM). The first Naþ isthought to bind at a site near the periplasmic surface (center I), whichincludes D203 and probably also N373. As the next step, we envisagebinding of the second Naþ ion to the Y229 and S382 including site(center II). As these residues are within the hydrophobic core of themembrane, the electroneutrality principle applies, which was developed forelectron transport complexes (Rich et al., 1995). Adopting this principleimplies that a Naþ ion would be tolerated at this position only after chargebalancing, requiring in this case the dissociation of a proton and itsremoval from the site. The phenolic hydroxyl group of Y229 is sufficientlyacidic to become dissociated upon Naþ binding. The dissociated protonis thought to move to the carboxybiotin, where it is consumed in thedecarboxylation of this acid-labile compound. Concomitantly, the biotin


Figure 3 Model for coupling Naþ and Hþ movements across the membrane to thedecarboxylation of carboxybiotin. The model shows the approximate location ofimportant residues of helix IV, helix VIII and of region IIIa of the b subunit.Also shown is the participation of these residues in the vectorial and chemical events ofthe Naþ pump. (A) shows the empty binding site region with enzyme-boundcarboxybiotin (B-COO�), exposing the Naþ binding sites toward the cytoplasm.(B) shows the situation where the first Naþ binding site at the D203/N373 pair (centerI) has been occupied and the second Naþ enters the Y229/S382 site (center II) with thesimultaneous release of the proton from the hydroxyl side chain of tyrosine 229. Thisdisplacement may be facilitated by R389 through lowering the pK of the tyrosinehydroxyl group. The proton is delivered to the carboxybiotin and catalyzes theimmediate decarboxylation of this acid-labile compound, involving a conformationchange (B to C) which exposes the Naþ binding sites toward the periplasm andsimultaneously decreases their Naþ binding affinities. The Naþ ions are subsequentlyreleased into this reservoir, while a proton enters the periplasmic channel and restoresthe hydroxyl group of Y229. In (D), the Naþ binding sites are empty and exposedtowards the periplasm and the biotin prosthetic group (B–H) is not modified. Uponcarboxylation of the biotin, the protein switches back into the conformation where theNaþ binding sites are exposed towards the cytoplasm (D to A).


prosthetic group leaves the site and OadB changes its conformation.This exposes the Naþ binding sites towards the periplasm and simul-taneously decreases their Naþ binding affinities. The Naþ ions are subse-quently released into this reservoir, while a proton enters the periplasmicchannel and restores the hydroxyl group of Y229. Hence, eachdecarboxylation event is coupled to the transport of two Naþ ions fromthe cytoplasm into the periplasm and the consumption of a periplasmically-derived proton.

2.3. Sodium-Translocating F1F0 ATP Synthase

As mentioned above, sodium-translocating ATP synthases are rareenzymes restricted to anaerobic bacteria which perform Naþ extrusion bya primary pump (e.g. a sodium-translocating decarboxylase). In structureand mechanism, however, sodium ion- and proton-translocating ATPsynthases appear to be closely related. The first Naþ translocating F1F0

ATP synthase was found in P. modestum (Laubinger and Dimroth, 1988).A representation of the overall organization and mechanism of the P.modestum ATP synthase is shown in Fig. 4. The enzyme consists of twostructurally and functionally distinct entities termed F1 and F0. Detailedknowledge on structure and function is available for the water-soluble F1

headpiece with the subunit composition a3b3gde (Abrahams et al., 1994).Alternating a and b subunits form a cylinder around subunit g. Part ofthe g subunit protrudes from the bottom of the cylinder and forms thecentral stalk together with the e subunit. At its foot, the stalk is connectedwith the oligomeric c-ring of the membrane-intrinsic F0 moiety. Theother F0 subunits of bacterial ATP synthases are a and b2, which abutthe c-ring laterally. Subunit a is an integral membrane protein which takesan active part in the ion translocation mechanism. The two b subunits areanchored with their N-termini within the membrane. The major partof the subunit b dimer is a-helical, making up the peripheral stalk thatconnects subunit a of F0 with subunit d of F1. The latter connection alsoinvolves the a subunit.While the structure of the water-soluble F1 part of the complex has

been solved to high resolution (Abrahams et al., 1994), structuralinformation on the membrane-intrinsic F0 moiety is more restricted. Mostinvestigations have been focused on subunit c. Structural informationby NMR was obtained for the monomeric unit in either chloroform/methanol/water or in SDS micelles. Subunit c of E. coli folds in theorganic solvent mixture as a helical hairpin with a different shape at


pH 5.5 or 8 (Girvin et al., 1998; Rastogi and Girvin, 1999). Subunit c of P.modestum does not fold into a stable three dimensional structure in eitherthe organic solvent mixture or in SDS, and the secondary structures aredistinct under both conditions (Matthey et al., 1999, 2002). This may

Figure 4 Model for the organization and mechanism of the Naþ translocating F1F0

ATP synthase of P. modestum. During ATP synthesis, the Naþ ions are envisaged toenter through the subunit a channel from the periplasmic side of the membrane.Approximately in the middle of the membrane they bind to an empty rotor site at thesubunit a/c interface. The next empty rotor site is attracted to the a subunit channel by themembrane potential and the previously occupied rotor site rotates out of the subunit a/cinterface. The bound Naþ is now accessible to the cytoplasmic reservoir by itsrotor intrinsic channel and may dissociate into this reservoir at very low externalNaþ concentrations. Under physiological conditions, however, the site remains occupieduntil it approaches the a subunit from the other side. The universally conservedarginine (R227 in P. modestum) facilitates the dissociation of Naþ from an approachingrotor site. (See colour plate section.)


indicate a high degree of structural flexibility in the monomeric unit.More detailed knowledge on the structure of this protein and itsorganization within the oligomeric ring was obtained by X-ray or electrondiffraction analyses of 3-D or 2-D crystalline samples. One prominentresult of these studies is the variation of the c-ring stoichiometry betweenspecies, being 10 for yeast mitochondria (Stock et al., 1999), 11 for twodifferent bacterial ATP synthases (Stahlberg et al., 2001; Meier et al., 2002),and 14 for spinach chloroplasts (Seelert et al., 2000). A C-a structuralmodel derived from electron densities of 2-D crystals of c11 from I. tartaricusis shown in Fig. 5 (Vonck et al., 2002). From this model it is evidentthat the oligomeric protein consists of two concentric rings of helicessurrounding a central cavity. In the inner ring the helices pack verytightly, leaving no room for side chains between neighbors. This allowedus to assign these to the N-terminal helices. These have the remarkablyconserved GxGxGxGx motif which must be responsible for this unusuallytight packing. Also seen on the map are the cytoplasmic loops connectingthe N-terminal inner helices with the C-terminal outer helices. Theouter helices are in staggered position to the inner helices of the ring.In this arrangement, there may be enough space between an inner helix

Figure 5 C-a structural model of the c-ring from I. tartaricus. (See colour platesection.)


and two outer helices for an ion access channel towards the binding sitewithin the middle of the membrane (see below).It has been shown by site-directed mutagenesis that residues Q32, E65,

and S66 are involved in Naþ binding (Kaim et al., 1997). Naþ is thephysiological coupling ion of the ATP synthase of P. modestum orI. tartaricus, but at low Naþ concentrations these enzymes may alsotranslocate Liþ or Hþ (Laubinger and Dimroth, 1989). For the binding ofLiþ , Q32 is dispensable and E65 is the only residue of the triplet requiredfor proton binding and translocation (Kaim et al., 1997). It is interestingin this context that P28, Q32, E65, and S66 provide a conserved motifin all c subunits of Naþ -translocating ATP synthases. This signature cantherefore be used to predict from the sequence whether an ATP synthasebelongs to the family of Naþ -translocating enzymes. The E65, or acorresponding aspartate residue, is universally conserved in all ATPsynthases, irrespective of whether they translocate Naþ or Hþ . It hastherefore been concluded that the proton-translocating ATP synthasesuse this conserved acidic residue as the ion binding site.Another well documented property of the conserved acidic residue

in subunit c is its specific reaction with dicyclohexylcarbodiimide(DCCD). The carbodiimide reacts specifically with the protonated form ofthis acid to yield an N-acylurea derivative (Kluge and Dimroth, 1993a).This highly specific covalent modification of the binding site residue hasbeen used to determine its membrane topology. For this purpose, cE65was modified with N-4-[3-(trifluoromethyl)-3H-diazirin-3-yl]benzyl-N0-cyclohexylcarbodiimide (diazirin-BCD), a photoactivatable derivative ofDCCD. After illumination of this modified ATP synthase reconstitutedinto phospholipid vesicles, photo crosslink products were specificallyformed with the fatty acid side chains of the phospholipids (von Ballmooset al., 2002a). This indicates that the acidic ion binding site residue is inclose contact to the fatty acid chains of the phospholipids and therefore nearthe center of the membrane. This topology of the binding site wascorroborated by fluorescence quenching studies. The ATP synthase ofI. tartaricus was specifically labeled at its c subunit sites with N-cyclohexyl-N0-(1-pyrenyl)carbodiimide (PCD), a fluorescent analogue of DCCD andthe enzyme was reconstituted into proteoliposomes containing spin-labeled phospholipids at different positions along their stearic acid chains.A quantitative analysis of the quenching of the fluorophore dependingon the position of the spin label indicated a deeply membrane-embeddedlocation of the fluorophore and, thus, of the binding site being 1.3� 2.4 Aapart from the center of the bilayer. For comparison, a similar investigationwas performed with the proton-translocating ATP synthase of E. coli


with the result that the binding site of this enzyme is at a conserved locationwithin the center of the membrane (von Ballmoos et al., 2002b).Additional evidence for the participation of cE65 of the P. modestum or

I. tartaricus ATP synthase in the Naþ binding site was obtained bymeasuring the kinetics of the modification of this residue by DCCD inrelation to Hþ and Naþ concentration. The second order rate constant ofthe reaction followed a titration curve with an inflection point at pH 7 inthe absence of Naþ , which represents the pK of cE65. The rate of themodification reaction dropped in the presence of Naþ ions and the pK wasshifted into the acidic range. These data suggest that Naþ and Hþ competefor binding to cE65 and that only the protonated species of this residuereacts with DCCD (Kluge and Dimroth, 1993a).The enhancement of the rate of modification of cE65 with DCCD by

Hþ and its inhibition by Naþ are direct evidence that this deeplymembrane-embedded site can be reached by these cations. This isindependent of whether the ATP synthase is dissolved in detergent orincorporated into the membrane and hence clear evidence that thesecations reach the site via specific access channels. Currently, there is muchdebate about the location of these channels. Both cytoplasmic andperiplasmic access channels have been placed in subunit a in a model ofthe E. coli enzyme, which requires experimental verification (Vik andAntonio, 1994; Junge et al., 1997). In contrast, the binding sites of the P.modestum or I. tartaricus ATP synthases are believed to be reached fromthe cytoplasm by rotor-intrinsic access channels and from the periplasmby the a subunit stator channel. This localization of the channels isstrongly supported by numerous biochemical data. Most convincingly,the subunit c ring is modified by DCCD or derivatives with the samerate in its isolated state or when incorporated into the ATP synthasecomplex and the modification is affected by Hþ or Naþ to a similar extent(Meier et al., 2002). The results of Fig. 6 show very slow labeling of rotorsites of the isolated c ring by PCD, the fluorescent analogue of DCCD,at pH 8.45 and a dramatic instantaneous increase of the reaction rate uponacidification to pH 6.1. Furthermore, the modification reaction stoppedimmediately by adding 15 mM NaCl (Meier et al., 2002). These resultsclearly show that Hþ and Naþ have rapid access to the membrane-embedded binding sites of the isolated rotor and therefore the accesschannels for these cations must exist within the rotor itself. According tothe structure of the c ring, the access channels could be aligned by an innerhelix and the two contiguous outer helices (Vonck et al., 2002) (see above).A remarkable feature of the c ring from the Naþ -translocating

ATP synthases is its extreme stability, resisting even boiling with SDS


(Laubinger and Dimroth, 1988). This observation indicates a rather rigidstructure of the c ring. Therefore, it appears unlikely that individual csubunits have been either lost or added during the isolation procedure.The identical mobility of the c ring on SDS-PAGE before or after isolationfrom the ATP synthase confirms this supposition (Meier et al., 2002).In addition, all c rings analyzed by atomic force microscopy (AFM)are composed of 11 monomeric units, except a minority of rings in which1–3 subunits are missing (Stahlberg et al., 2001; Muller et al., 2001a; Meieret al., 2002). Interestingly enough, the c rings with substoichiometricamounts of c subunits have the same diameter as the c11 rings andare therefore regarded as incompletely assembled. Hence, the number of csubunits seems to be determined by the folding of the monomeric unitsand there appears to be no variability of ring stoichiometries within onespecies. Interestingly, Naþ binding considerably contributes to thestability of the c11 rings. The rationale for this effect is cross bridging ofadjacent monomeric units via Naþ binding to the binding site residues atthe inner helix and at the two outer helices of neighboring monomericunits as depicted in Fig. 7 (Meier and Dimroth, 2002). This extremerigidity of the c ring assembly seems difficult to reconcile with hypotheses

Figure 6 Kinetics of the modification of E65 of the isolated c-ring from I. tartaricusby PCD in response to pH and the concentration of Naþ ions. The fluorescence increaseat 377 nm which is due to the formation of the covalent reaction product betweenPCD and cE65 was followed continuously. Indicated is the pH shift from pH 8.45 topH 6.1 which initiated the reaction and the addition of 15 mM Naþ which stopped thereaction.


in which the outer helices of each monomeric unit perform a significantturning movement each time they approach subunit a (Fillingame et al.,2000).Another interesting aspect of the c ring structure is the presence of

a phospholipid plug which closes the central cavity at the periplasmicside of the membrane (Meier et al., 2001). This plug has been observedby AFM of two dimensional arrays of the c ring embedded in aphospholipid bilayer. Since the reconstitution procedure involves the

Figure 7 Structural model showing details of the Naþ coordination site in the c11 ringof the sodium F1F0 ATP synthase. One subunit with C- and N-terminal a-helices is shownin light blue (subunit 1) and the C-terminal a-helix from the adjacent subunit is shownin light green (subunit 2). The Naþ binding pocket is coordinated by at least threeamino acid side chains. Subunit 1 provides glutamine 32 and serine 66 located on theN- and C-terminal helix, respectively. Glutamate 65 is located on the C-terminal a helixof subunit 2. (See colour plate section.)


addition of phospholipids, it cannot be concluded from these experimentswhether the plug is an artifact or an intrinsic property of the ATPsynthase in its native membrane environment. It is clear that the centralhole must be sealed in the membrane-bound enzyme to prevent uncontrol-led ion flux across the membrane. Phospholipids would be a perfectseal, but alternatively a protein or part of a protein might close thehole. To investigate this ambiguity, a photo cross linker was attachedto the N-terminal inner helix of the c ring of the membrane-boundATP synthase, and the cross-link products formed upon illuminationwere analyzed by MALDI mass spectrometry. Preliminary data indicateno cross-link formation of subunit c with another protein but show apeak of the size expected for a cross-linked product with a phospho-lipid (unpublished data). From these results, we hypothesize thatthe phospholipid plug at the periplasmic side of the c ring is not areconstitution artifact but a natural feature of the ATP synthase,where its role is to seal the central hole to prevent ion leakage alongthis route.

2.3.1. ATP Synthesis by Rotational Catalysis and its Dependenceon the Membrane Potential

Probably the most remarkable feature of the ATP synthase is therotational catalysis. It has been elegantly shown by direct observation witha microscope that a fluorescent actin filament attached to the g subunit ofthe F1 moiety rotates upon ATP hydrolysis (Noji et al., 1997). Themechanical driving forces for this rotation are bending movements of thethree b subunits in a coordinated fashion. At any one time, each of the threeb subunits is in a different conformation forming different contacts withthe asymmetrically bent g subunit in the interior lumen of the cylinder.These coordinated bending movements of the three b subunits enforce therotation of the g subunit (Abrahams et al., 1994, 1996; Oster and Wang,2003). Outside the cylinder the g subunit is connected with the e subunit andthese two are linked to the cytoplasmic loops of the c ring (Gibbons et al.,2000; Rodgers and Wilce, 2000). Hence, g, e and cn rotate together as aunit and are designated as the rotor, while the remaining subunits togetherform the stator (Capaldi and Aggeler, 2002).Most rotation experiments have been performed in the ATP hydrolysis

direction and with enzyme preparations that were either incomplete ornot fully coupled as evidenced from the lack of inhibition by DCCD(Sambongi et al., 1999; Panke et al., 2000; Yoshida et al., 2001). However,


more recently, rotation has also been reported for the coupled holoenzymein both ATP hydrolysis and ATP synthesis direction (Kaim et al., 2002).For this purpose, the Naþ -translocating ATP synthase of P. modestum wasreconstituted from individual subunits. On average, one fluorophore wasattached to one of the c subunits of the ring and served as the probe tomonitor rotation. ATP-driven rotation was strictly Naþ -dependent andinhibited by DCCD as expected for a coupled ATP synthase. To observethe rotation during ATP synthesis, the ATP synthase with anattached fluorophore at the c ring was reconstituted into proteoliposomesand the membrane energised by applying a Naþ concentration gradientand/or a membrane potential. The � component was obligatory for therotation. This result is in accord with various biochemical studiesdemonstrating the obligatory role of � as the driving force for theATP synthase (Kaim and Dimroth, 1998a, 1998c, 1999).Implicit to Mitchell’s chemiosmotic model is the assumption that

membrane potential and transmembrane ion gradients are thermodynami-cally equivalent:

Proton motive force ð�mHþÞ ¼ ð2:3RT=FÞ ��pHþ�

where R is the gas constant, T the absolute temperature and F the Faradayconstant. However, this equilibrium relationship does not take into accountthe kinetic driving forces of the ATP synthase under real working conditionsthat are far from equilibrium. This knowledge is crucial, however, tounderstand the mechanism of energy conversion within the F0 motorand thus of ATP synthesis.The essence of the membrane potential for the mechanism of F0

rotation was discovered more than 10 years ago during Naþ transportstudies with the P. modestum ATP synthase (Kluge and Dimroth, 1992).Sodium ion uptake into F0-containing liposomes was not observedin the presence of substantial Naþ ion concentration gradients (�pNaþ )and at � values <�40 mV. The Naþ ion transport rate increasesexponentially with increasing membrane potentials approaching satura-tion at � >�120 mV. Upon reversal of the membrane potential, thedirection of Naþ transport changes to the export of Naþ from the F0

liposomes. In the absence of a membrane potential, the F0 motor is in anidling mode performing oscillations of the rotor versus the stator in eitherdirection, which is characterized by sodium ion exchange across themembrane. Upon applying voltage (i.e. � ), the rotation is rectified andNaþ ions are transported unidirectionally across the membrane.Similarly, the reconstituted F1F0 complex is in the idling mode without


external energy sources and catalyzes sodium ion exchange across themembrane (Kaim and Dimroth, 1998c). This situation does not change inthe presence of a large �pNaþ , as this is not a suitable driving force toinduce unidirectional rotation and to generate torque. Upon applyingvoltage, however, the enzyme immediately switches from the idling into thetorque-generating mode, discontinuing sodium ion exchange acrossthe membrane.This informative result is consistent with ATP synthesis experiments

performed with reconstituted ATP synthases from P. modestum, E. coli, orspinach chloroplasts. In neither case is ATP synthesis observed if only anion gradient (�pNaþ or �pH) is applied. The rate of ATP synthesisincreases exponentially with increasing membrane potentials, approachingsaturation at � > 60 mV for the chloroplast ATP synthase and at � >120 mV for the P. modestum and E. coli ATP synthases, respectively (Kaimand Dimroth, 1998a, 1999). Interestingly, the dependence of the initialrate of ATP synthesis on the � by the P. modestum enzyme follows almostexactly the same profile as that of Naþ uptake by the F0 domain (Klugeand Dimroth, 1992) (see above). Accordingly, � driving Naþ transloca-tion through the F0 motor components generates the torque required tosynthesize ATP at the catalytic F1 sites.These data required us to revisit the classical acid-base transition

experiment performed almost four decades ago. When thylakoids areequilibrated with succinate buffer, pH 5.5, and then rapidly diluted intoTris–HCl buffer, pH 8.5, ATP is synthesized from ADP and phosphate.It was consequently concluded that ATP synthesis can be driven by �pHalone (Jagendorf and Uribe, 1966). This result is obviously contradictoryto the obligatory role of the membrane potential to drive ATP synthesis.This discrepancy is elegantly resolved by the generation of a large � under the conditions of the acid-base transition experiment (Kaim andDimroth, 1999). Succinate exists mainly as a monoanion at pH 5.5 and asa dianion at pH 8.5. Due to the concentration gradient of the succinatemonoanion and its membrane permeability, a diffusion potential ofapproximately 150 mV is generated which adds the essential electricaldriving force for the synthesis of ATP. The rationale for the permeabilityof the succinate monoanion is its folding into a ring structure in which thenegative charge is delocalized, since it is shared by both carboxylicgroups. Hence, maleinate (cis) which can also fold into a ring structureinduces a diffusion potential and supports ATP synthesis to the samedegree as succinate. Fumarate (trans) or monocarboxylates which areunable to form such ring structures were unable to induce ATP synthesis orto generate a diffusion potential.


2.3.2. Model of the F0 Motor

The fundamental question as to the mechanism of action of the ATPsynthase is how the electrical potential is used to allow ion flux through theF0 motor components and how this leads to the generation of torque.Unfortunately, structural knowledge on the entire F0 ensemble in general,and on subunit a in particular, is sparse. The F0 motor comprises acounter-rotating assembly of the c11 rotor domain and the stator asubunit that abuts the rotor laterally (Birkenhager et al., 1995; Singh et al.,1996; Takeyasu et al., 1996). From mutational analysis with the P.modestum enzyme, the a subunit was concluded to harbor an ion selectivechannel (stator channel) that connects the rotor site at the a/c interface withthe periplasmic surface of the membrane (Kaim and Dimroth, 1998b,1998c). As the binding sites are connected by rotor-intrinsic channels tothe cytoplasmic surface outside the a/c interface (Meier et al., 2002; Voncket al., 2002; von Ballmoos et al., 2002a), Naþ ions traverse the entiremembrane via the stator channel and one of the eleven rotor channels.A model of how this ion movement may be coupled to torque generation

is depicted in Fig. 8 (Dimroth et al., 1999). An important amino acid inthe process is the universally conserved arginine 227 within thepenultimate helix of subunit a (Lightowlers et al., 1987; Cain and Simoni,1989; Howitt and Cox, 1992; Hatch et al., 1995; Valiyaveetil and Fillingame,1997). Cross-linking experiments have identified the position of R227(the positive stator charge) at the interface with the outer C-terminalhelix of c11, approximately one helical turn from the binding site towardsthe cytoplasmic surface (Jiang and Fillingame, 1998). Let us first considerthe motor’s performance in the ATP synthesis direction. Without amembrane potential the rotor rocks against the stator within a narrowangle moving Naþ ions back and forth across the membrane (Kaim andDimroth, 1998c). At the physiological Naþ concentration inside a P.modestum cell (approximately 30 mM), the rotor sites are occupied most ofthe time while outside the a/c interface. Upon clockwise rotationas viewed from the periplasm, the site approaches the stator chargenear the a/c interface. This promotes the dissociation of the bound Naþ

which then diffuses through the pertinent rotor channel into the cytoplas-mic reservoir. Now negatively charged, the site is electrostatically attrac-ted by the positive stator charge. The rotor diffuses out of the potentialwell by thermal fluctuations. Without the membrane potential thisdiffusion occurs in either direction with equal probability. Upon applyingvoltage, however, the diffusion is biased towards the positive side dueto the electrostatic attraction of the negatively charged rotor site. Implicit


in this model is a horizontal component of the electric field at the positionof the rotor site. This is rationalized by the adjacent stator and rotorchannel, each reaching into the center of the membrane from a differentside. If these channels are aqueous the potential drop will be mainly in thehorizontal direction between these channels. Thus attracted, the sitecomes within reach of the stator channel where it quickly picks up a Naþ

ion. Now neutralized as a dipole it is no longer attracted backwards by

Figure 8 Model for torque generation by the sodium F0 motor. An occupiedrotor site approaches the subunit a/c interface with the conserved stator charge (R227in P. modestum). The positively charged arginine facilitates the dissociation of the sodiumion from the binding site into the rotor channel through which it escapes into thecytoplasmic compartment. The empty rotor site contains the negatively chargedglutamate and is attracted by the positive stator charge. Without an externaldriving force the site escapes from the potential well with equal probability into eitherdirection. In the presence of a membrane potential, however, the diffusion is biasedinto the direction of the stator channel. The rationale for this attraction is ahorizontal component of the membrane potential which forms between aqueous rotorand stator channels penetrating into the center of the membrane from the twoopposing sides. At the stator channel the rotor site quickly picks up a Naþ ionoriginating from the periplasmic reservoir and unidirectional rotation continues bythe attraction of the next empty rotor site through the membrane potential. (See colourplate section.)


the stator charge which therefore exerts a pull on the next empty rotorsite at the a/c interface. The occupied site simultaneously continues therotation through the hydrophobic portion of the stator and releasesthe bound Naþ ion into the cytoplasm as it comes close again to thestator charge.In the reverse mode, the ATP synthase operates as an ATP-driven

ion pump. Rotation is now in the opposite direction and transmittedfrom the F1 domain to the F0 domain. Hence, occupied rotor sites areforced to move first through the hydrophobic part and along the channelof the stator before they reach R227. Here, the Naþ dissociates andescapes through the stator channel to the periplasmic surface. Uponfurther rotation the empty site is moved out of the a/c interfacewhere it quickly picks up a Naþ ion from the rotor channel. Importantly,ATP hydrolysis or ATP-driven rotation of the c11 rotor ring in energy-coupled F1F0 are strictly Naþ -dependent, suggesting that in the hydroly-sis direction empty negatively-charged sites cannot enter the a/cinterface (Kluge and Dimroth, 1993a, 1993b). This observation is easilyreconciled with the model and further indicates that the electrostaticconstraints of moving a negative charge into the hydrophobic portion ofthe stator cannot be overcome by the torque elicited through ATPhydrolysis.The anticipated electrostatic attraction between the positive stator

charge and a negatively charged rotor site has been probed with mutantsof aR227. Changing this residue to lysine abolished ATP-driven Naþ

transport or 22Naþout/Naþ

in exchange at neutral pH, but not at pHvalues between 8 and 9 (Wehrle et al., 2002). This informative experimentindicates that lysine carrying a localized positive charge attracts thenegatively-charged rotor site more vigorously than arginine with amore delocalized charge. The strength of the ion pair between the statorlysine and the rotor site is so strong that it cannot be broken by the torquegenerated by ATP hydrolysis. At elevated pH values, where the lysineis deprotonated part of the time, the electrostatic force declines and thetorque created by the hydrolysis of ATP is now adequate for the rotationout of the potential well. Locking the rotor in an immobile positioncoincides with the conclusion from the E. coli ATP synthase that bychanging the conserved arginine to lysine any ATP-driven ion transportactivity is turned off (Lightowlers et al., 1987; Cain and Simoni, 1989; Hatchet al., 1995; Valiyaveetil and Fillingame, 1997). The pertinent experimentswith the E. coli enzyme were all performed at neutral pH and it wouldbe interesting to know whether the activity of this enzyme is also recoveredat elevated pH values.


Further insight into the function of R227 was obtained by mutating thisresidue to alanine (Wehrle et al., 2002). Intriguingly, the mutant enzymecatalyzed ATP synthesis, but only at Naþ concentrations in the outsidereservoir of the proteoliposomes, well below the dissociation constant ofthe binding sites (about 1 mM). From these and other results it wasconcluded that the critical role of R227 is to facilitate by its positive chargethe dissociation of rotor sites coming close to this residue. With theelectroneutral alanine in its place, rotor sites do not lose the ions at thea/c interface. They only dissociate from these sites outside subunit a andpenetrate through rotor channels into the external reservoir if the Naþ

concentrations in this environment are sufficiently low. Collectively, thesedata support an electrostatic mechanism for the F0 motor. The localelectrostatic field derived from the membrane potential in concert withoscillations between charged and neutral rotor sites, as ions move acrossthe membrane, drives the rotation of the F0 motor components andhence ATP synthesis. The man-made electrical motor depends onmagnetic forces instead of the electrostatic forces of the biologicalmotor, but operates otherwise on remarkably similar principles: aconstant field pulls a magnetized rotor component from a certain angle,whereupon it gets neutralized and the field exerts its pull on the next rotorcomponent. These oscillations of the rotor components between theenergized state, where they become attracted by the field, and the non-energized state, where they are silent, elicit the continuous rotation of rotorversus stator.

3. ALKALIPHILIC BACTERIA GROWING AT LOW DmHþ

Like anaerobic bacteria, alkaliphilic bacteria are also faced with thechallenge of synthesizing ATP at low �mHþ . Alkaliphilic bacteria growover the pH range 7.5–11.5 and can be divided into two groups: obligatealkaliphiles that grow between pH 9.0 and pH 11.5 (e.g. Bacillusalcalophilus, Bacillus firmus RAB) and facultative alkaliphiles that growbetween pH values of pH 7.5 and 11.2 (e.g. Bacillus pseudofirmus OF4and Bacillus halodurans C-125) (Krulwich and Guffanti, 1989). Recently, athermophilic facultative alkaliphile Bacillus sp. strain TA2.A1 which growsover the pH range 7.5–10.2 on non-fermentable carbon sources wasdescribed (Peddie et al., 1999; Olsson et al., 2003).Growth of alkaliphilic bacteria at pH 10.0 on non-fermentable

carbon sources is inhibited by the electrogenic protonophore carbonyl


cyanide m-chlorophenylhydrazone (CCCP) (Hoffmann and Dimroth,1991b; Olsson et al., 2003) and the electroneutral Naþ /Hþ antiportermonensin, suggesting that both an electrochemical gradient of protons(�mHþ ) and sodium ions (�mNaþ ) are required for growth at alkalinepH values. The roles of these respective chemical gradients in the growthand cellular bioenergetic processes (e.g. ATP synthesis) of alkaliphilicbacteria will be discussed.

3.1. Sodium and Proton Cycling in Alkaliphiles

It has been well documented that alkaliphilic bacteria exhibit a lowsodium dependency for growth when compared to marine bacteria(Horikoshi, 1991; Krulwich et al., 2001). Moreover, this requirement forsodium can vary depending on the pH for growth (Ito et al., 1997; Gilmouret al., 2000). Sodium is used as a coupling ion (in combination with � ) todrive both solute transport (Koyama et al., 1976; Krulwich et al., 1985;Peddie et al., 1999, 2000) and flagellar rotation for motility (Sugiyama et al.,1985, 1988; Imae et al., 1986). The maintenance of �mNaþ in alkaliphilicbacteria requires sodium extrusion activity mediated by electrogenicsecondary Naþ /Hþ antiporters (e.g. Mrp complex, NhaC) that facilitatenet proton accumulation with Naþ extrusion (see Krulwich et al. (2001)for a recent review).The respiratory chains of some alkaliphilic bacteria have been

described and their primary activity is proton extrusion (Hicks andKrulwich, 1995). The respiratory chain of B. pseudofirmus OF4 is branchedand terminates with two terminal oxidases, cytochrome caa3 andcytochrome bd-type (Quirk et al., 1991; Gilmour and Krulwich, 1997).Bacillus pseudofirmus contains a succinate dehydrogenase complex butlacks a terminal cytochrome bo complex (Gilmour and Krulwich, 1996).To date, no evidence has accumulated for the presence of a primaryrespiration system coupled to Naþ extrusion. Krulwich and coworkers(1998b) propose that primary sodium pumping without concomitant Hþ

uptake may be inhibitory to alkaliphilic bacteria due to the depletion ofcytoplasmic Naþ . Intracellular Naþ is crucial for sustained secondaryantiporter activity that catalyzes net proton accumulation during respira-tion for pH homeostasis. However, both solute transport and motilityserve as re-entry routes for the completion of the sodium cycle. The roleof these and other systems in Naþ uptake for pH homeostasis has beenreviewed (Sugiyama, 1995; Krulwich et al., 2001). Sodium extrusionmediated by a probable ABC transporter that is not coupled to proton


uptake has been reported in alkaliphilic bacteria (Wei et al., 1999).Presumably such systems operate at Naþ concentrations that may be toxicto the cell and keep the intracellular Naþ at a level that is congruent withNaþ /Hþ antiporter activity.Another mechanism to generate a �mHþ , employed primarily by

anaerobic bacteria, is by ATP hydrolysis coupled to proton pumping viathe membrane-bound F1F0 ATPase. Proton extrusion coupled to thehydrolysis of ATP would occur during growth on substrates that are notstrictly coupled to oxidative phosphorylation or under conditionswhere the �mHþ is low assuming that the ATP synthase can indeedoperate in the ATP hydrolysis direction under normal growth conditions(see below).

3.2. pH Homeostasis in Alkaliphilic Bacteria and Growthat Low DmHþ

The intracellular pH of alkaliphilic bacteria is maintained at values moreacidic (i.e. approximately 2 pH units) than their external environment(Hoffmann and Dimroth, 1991b; Sturr et al., 1994; Krulwich et al., 1997,1998b). Acidification of the cytoplasm is due to the activity of electrogenicsecondary Naþ /Hþ antiporters (e.g. Mrp complex, NhaC) that facilitatenet proton accumulation with Naþ extrusion (Krulwich et al., 1998a,1998b). Recently, we reported that the magnitude of the �pH generatedby the thermoalkaliphile Bacillus sp. strain TA2.A1 was only one pHunit (acid in) (Olsson et al., 2003). This lower pH gradient could be aresult of increased proton permeability that strain TA2.A1 exhibits at itsoptimum growth temperature (K. Olsson and G.M. Cook, unpublishedresults).Because the total �mHþ is the sum of the membrane potential

(� ; positive out) and the pH gradient (�pH; acid out in neutrophiles),the large �pH generated by alkaliphiles in the opposite direction (acid in)is adverse with respect to the magnitude of the total �mHþ . The obligatorydriving force for ATP synthesis, the � , ranges from �136 to �180 mV(pH 7.5 to 10.5) for B. pseudofirmus OF4 (Guffanti and Hicks, 1991), �206to �213 mV for B. alcalophilus (Hoffmann and Dimroth, 1991b) and �135to �150 mV for Bacillus sp. strain TA2.A1 (Olsson et al., 2003). Due to thelarge inverted pH gradient (approximately þ 120 mV) generated byalkaliphilic bacteria, the �mHþvalues (sum of � þZ�pH) wouldappear to be suboptimal for ATP synthesis (e.g. �50 to �78 mV)(Guffanti and Hicks, 1991; Krulwich, 1995; Olsson et al., 2003).


�mHþ values as high as �109 mV have been reported for B. alcalophilus,but even these values are near the lower threshold for ATP synthesis via aconventional chemiosmotic mechanism (Hoffmann and Dimroth, 1991b;Dimroth, 1992). It should be noted that this �mHþ value is similar to thatof most anaerobic bacteria.

3.3. Mechanisms to Synthesize ATP at Low DmHþ using aProton-Coupled ATP Synthase

Despite the apparent suboptimal �mHþ values for ATP synthesis at highpH and the increased energetic cost for pH homeostasis, the molar growthyields of alkaliphilic bacilli under these conditions are not compromisedwhen compared with near neutral pH growth values (Sturr et al., 1994;Olsson et al., 2003). In fact the molar growth yield of B. pseudofirmuson malate is higher at pH 10.5 than at pH 7.5 (Sturr et al., 1994).Furthermore, the phosphorylation potential (�Gp) values reported inalkaliphilic bacteria range from �418 to �500 mV at high pH values(Guffanti and Hicks, 1991; Hoffmann and Dimroth, 1991b; Sturr et al.,1994; Olsson et al., 2003) which are in good agreement with those valuesreported for conventional neutrophiles (Thauer et al., 1977). The ATPconcentration and ATP :ADP ratios are also comparable to other bacteriain which the �mHþ is high.No bioenergetic problem would exist if alkaliphilic bacteria used Naþ -

coupled processes for solute transport, motility, and ATP synthesisbecause �mNaþ and � are orientated in the same direction and thusadd driving force to one another. Indeed, ion/solute transport systemsand flagellar rotation depend on sodium as a coupling ion in alkaliphiles(see above). However, the F1F0 ATP synthase of mesophilic alkaliphilicB. pseudofirmus and B. alcalophilus have been shown to be exclusivelyproton-coupled enzymes (Hicks and Krulwich, 1990; Hoffmann andDimroth, 1990, 1991a). We have recently shown that the F1F0

ATP synthase of the thermoalkaliphilic Bacillus sp. strain TA2.A1 isalso a proton-coupled enzyme (Cook et al., 2003). It should be notedthat a number of putative mechanisms have been proposed to explainhow the ATP synthase in these bacteria remains proton-coupled atlow �mHþ , but these models are still to be experimentally proven(Krulwich, 1995).An elegant mechanism to synthesize ATP at a low �mHþ would be to

use a high stoichiometry of Hþ per ATP synthesised. In fact, a changein the calculated thermodynamic stoichiometry (�Gp/�mHþ

¼ Hþ /ATP)


of the ATP synthase, as the pH is increased from neutral to alkalinepH values, has been observed (Hoffmann and Dimroth, 1991b; Sturr et al.,1994; Olsson et al., 2003). At thermodynamic equilibrium forATP synthesis, the free energies of the �mHþ and phosphorylationpotential (�GP) are in balance and the equation Hþ /ATP��mHþ

¼�GP

applies. According to the work of Brusilow and colleagues, the numberof c subunits in the E. coli ATP synthase changes depending on thecarbon source used for growth (Schemidt et al., 1998). Molecular andbiochemical studies on the F1F0 ATP synthase of B. pseudofirmus haveshown, however, that there is no differential expression of the atp operonat pH 7.5 and 10.5, and the subunit properties and c subunit/holoenzymestoichiometry is constant at these pH values (Ivey et al., 1994).However, these studies were based on DCCD-labeling of invertedmembrane vesicles to quantify the amount of c monomer and thereforesubtle changes in the amount of c monomer per oligomeric c ring may havebeen overlooked.The above hypothesis is based on the dogma that the �mHþ is delo-

calized into the bulk phase. Krulwich and colleagues have proposed thata proton pathway generated by respiration-dependent proton pumpingand the ATP synthase may in fact be localized on or in the cytoplasmicmembrane (Krulwich, 1995). Based on this model, discrepancies betweenthe bulk �mHþ and the rate of steady state oxidative phosphorylationwould not need to be taken into account. If such a pathway was to exist,then the rate of ATP synthesis may be influenced by artificially imposeddriving forces from the bulk phase and those that involve the innateproton pathway (non-bulk or localized). Guffanti and coworkers addressedthis hypothesis by studying the rate of ATP synthesis by B. firmus RABwhen energized either by malate oxidation (natural respiratory process)or by a valinomycin-mediated potassium-diffusion potential (Guffanti et al.,1984, 1985). The authors report that a respiration-derived � throughmalate oxidation could drive ATP synthesis at pH 7.0 and 9.0, but avalinomycin-mediated potassium-diffusion potential was effective indriving ATP synthesis only at pH 7.0 but not at pH 9.0. The lack ofATP synthesis at pH 9.0 was not due to a lack of � as supported bytetraphenylphosphonium ion (TPPþ ) and a-aminoisobutyric acid (AIB)uptake (� -dependent processes). Based on these findings, it appears asthough respiration-derived energization is more effective for energizationof Hþ -coupled processes in a proton-depleted environment (pH 9.0)perhaps as a result of direct or localized coupling between proton pumpingand proton-translocating processes (e.g. ATP synthesis, Naþ /Hþ

antiporter activity). In contrast, the diffusion potential relies on protons


that are already in the bulk exterior phase to move inwards and this wouldbecome limiting as the pH increases. The pathway of proton movement isyet to be defined. It is important to consider that there is competitionbetween proton movement (via respiration) along the outer surface of themembrane versus proton equilibration with the bulk phase. This aspectmay be negligible for a bacterium growing at neutral pH in which theproton concentration in the bulk phase is similar to or even higher than inthe cytoplasm. However, it may become very significant if the protonconcentration in the bulk phase is much lower than the cytoplasm, asoccurs in alkaliphilic bacteria. The membrane potential (negative inside)can be regarded as an attractant for protons that are ejected by primarypumps on the outer surface of the cytoplasmic membrane resulting in alayer of high proton concentration. While the concentration of protons inthis layer and the bulk phase may not differ significantly in organismsgrowing at neutral pH, both concentrations would differ greatly in thecase of alkaliphilic bacteria. Based on this premise, the inverted pHgradient measured in alkaliphilic bacteria between bulk phase and thecytoplasm may be lower or may not even exist at the site of ATP syn-thesis, where only �pH between the surface layer and the cytoplasm areimportant. It could well be that the true value for �pH and �mHþ cannotbe precisely determined.

3.4. Alkaliphilic-Specific Amino Acid Motifs in the atpOperons of Mesophilic and ThermoalkaliphilicBacteria

The ability of the ATP synthases from alkaliphilic bacteria to work atlow or apparently suboptimal �mHþ has yet to be resolved. The atpoperons from the alkaliphiles B. pseudofirmus OF4 (Ivey and Krulwich,1991; Ivey and Krulwich, 1992), B. halodurans (Takami et al., 2000), andBacillus sp. strain TA2.A1 (Keis et al., 2004) have been cloned andsequenced. DNA sequence analysis reveals that the operon genearrangement and deduced primary structure of the gene products issimilar to other eubacterial operons which encode for proton-coupledF1F0 ATP synthases. The lack of the sodium ion binding signatureconsisting of residues Q32, E65, and S66 (P. modestum numbering) (Kaimet al., 1997; Kaim and Dimroth, 1998b) in the c subunit of alkaliphilicATP synthases further supports the notion that these enzymes are Hþ -coupled. The cloned ATP operons of B. pseudofirmus OF4 (Krulwichet al., 1998b) and Bacillus sp. strain TA2.A1 are unable to complement


atp mutants of E. coli for growth on succinate (Keis et al., 2004), eventhough membrane-bound ATPase activity exhibits characteristics of thealkaliphilic ATP synthase (i.e. stimulated by detergents).As reported by Krulwich and co-workers (Ivey et al., 1991; Ivey and

Krulwich, 1991, 1992), greatest variation from amino acid consensusis observed in the a and c subunits that are crucial for protontranslocation but no studies have been performed to determine the role(s)of these residues in alkaliphilic growth. The a subunit of F1F0-ATPsynthases is a hydrophobic protein, spanning the membrane five times.The E. coli enzyme is proposed to contain aqueous channels for protonsto gain access to the binding site in subunit c (Fillingame et al., 2002),while evidence from the Naþ -translocating ATP synthase of P. modestumindicates a periplasmic channel in the a subunit and 11 cytoplasmicaccess channels in the rotor ring (see above, Section 2.3). The universallyconserved stator charge arginine (R227 in P. modestum, R210 in E. coli)plays a critical role in the ion translocation mechanism (see Section 2.3)and this residue is also conserved in all alkaliphilic bacterial a subunits.An obvious deviation, however, as reported previously (Ivey et al., 1991;Ivey and Krulwich, 1992) is an invariant lysine residue (Lys218, E. colinumbering) located in membrane-spanning region 4 in the a subunit ofall alkaliphiles. Moreover, a glycine is also found in all alkaliphiles atposition 245 in membrane-spanning region 5 opposite Lys218. Hartzogand Cain (1994) reported that the effects of a single G218K mutation insubunit a of E. coli, which inhibits ATP synthase activity belowdetectable levels, are largely suppressed by a second aH245G mutationrestoring most of the lost activity. This would therefore suggest aninteraction of these residues but what role they play in the alkaliphilic asubunits has not been deduced. One potential explanation could be thatLys218 is at the entrance of the periplasmic channel where it capturesprotons from the alkaline environment and passes them via the channelonto the c subunit binding sites. In a recent study (Vonck et al., 2002),an alignment of c subunits from 38 different organisms showed that axGxGxGxGx motif in the N-terminal helix is highly conserved in allbacterial, chloroplast, and mitochondrial ATP synthases. This motif isbroken in the alkaliphiles and the glycines are replaced by bulkier aminoacids (viz. alanines and serines). Since the N-terminal helix is on theinside of the c subunit ring and glycines pack very tightly (Vonck et al.,2002), a more loose packaging of the inner helices could be envisaged inthe alkaliphiles. Site-directed mutagenesis studies are needed to definea role for these residues in alkaliphilic growth and ATP generationat low �mHþ .


3.5. The F1F0 ATP Synthases from Alkaliphilic Bacteria showLatent ATP Hydrolysis Activity: a Specific Adaptation forGrowth at Alkaline pH?

Another striking feature of the ATP synthases from alkaliphilic bacteriais the selective blockage of ATP hydrolysis but not ATP synthesis (Hicksand Krulwich, 1990; Hoffmann and Dimroth, 1990; Cook et al., 2003).So far, there have been no studies aimed at elucidating the molecularfeatures responsible for the specific blockage of ATP hydrolysis by theATP synthases from alkaliphilic bacteria and no rationale from aphysiological point of view has been provided. For a tenable explanation,the molecular features of the F1F0 ATP synthases from alkaliphilic bacteriamust be considered. Under conditions when the �mHþ drops below the�GP, the F1 motor hydrolyzes ATP, driving the F0 motor in reverse,whereupon it functions as a proton pump. Such a situation occursfrequently in anaerobic bacteria, where the main challenge is to keepthe membrane potential at a significant level. For this purpose, the ATPsynthase pumps protons outwards. Alkaliphilic bacteria are also confrontedwith situations in which the �mHþ is low (Krulwich et al., 1998a). This,however, is not caused by a low � but by the inverted proton gradientat high environmental pH. The main bioenergetic challenge for thesealkaliphilic bacteria is to maintain the cytoplasmic pH near neutral whengrowing at an external pH of 10.5 and therefore ATPase-dependentproton pumping, if the �mHþ drops below a critical level, may bedetrimental for this process. Thus, to block the ATP synthase of alkaliphilicbacteria in the ATP hydrolysis direction appears to be a necessaryadaptation for growth and survival in highly alkaline environmentswhere the �mHþ generated by these bacteria is low.

3.6. Regulation of ATP hydrolysis Activity by Bacterial F1F0

ATP Synthases

The ATPase activity of the ATP synthase enzyme is subject to regulation.In mitochondria under high oxygen tensions the ATP synthase operates inthe direction of ATP synthesis fueled by an electrochemical gradient ofprotons. When cells are limited for oxygen, the �mHþ collapses and theATP synthase switches from ATP synthesis to ATP hydrolysis fueledby ATP production from glycolysis (substrate level phosphorylation).In facultative and anaerobic bacteria the ATP synthase is used primarilyto generate a �mHþ and regulate pH homeostasis in the absence of oxygen.


Because the ATP synthase can hydrolyze ATP at low �mHþ , the potentialexists for ATP to be wasted under such conditions.In mitochondria, ATPase activity is regulated by the natural inhibitor

protein IF1 which binds to the ATP synthase in a pH-dependent manner.At mitochondrial matrix pH values below 7.0, the inhibitory action of IF1 isincreased (Panchenko and Vinogradov, 1985). Bovine IF1 has been shownto have two oligomeric states, tetramer and dimer, favored by pH valuesabove and below 6.5, respectively (Cabezon et al., 2000b). IF1 binds as adimer to form a stable complex with two F1 domains simultaneously(Cabezon et al., 2000a). At higher pH values IF1 is tetrameric and isinactive. Tetramer formation masks the inhibitory region of the proteinpreventing IF1 binding to the ATP synthase (Cabezon et al., 2001). Cross-linking studies suggest an interaction between IF1 and the C-terminal regionof the b-subunit (Jackson and Harris, 1988).The ATP synthase of chloroplasts is also subject to regulation of

ATPase activity mediated by a redox switch in the g subunit (intramoleculardisulfide bond S–S) (Nalin and McCarty, 1984; Richter et al., 1985). TheCF1F0 ATP synthase is a latent ATPase which can be stimulated tohydrolyze ATP in the presence of Ca2þ by dithiothreitol or heat, orthe presence of Mg2þ ions by methanol or octylglucoside treatment.Regulation of ATP hydrolysis is important to prevent ATP hydrolysis inthe dark in the absence of photophosphorylation. No homologue of IF1

has been found in either chloroplasts or bacteria.The e subunit from F1F0 ATP synthases is a two-domain protein

which consists of an N-terminal part that forms a flattened 10-strandedb-sandwich structure and a C-terminal domain that forms an a-helix-loop–a-helix structure. It has become evident that the e subunit acts as aninhibitor of ATP hydrolysis activity. Furthermore, it is apparent that mostor all of the inhibitory effect is caused by the C-terminal a-helical domain ofthis subunit. Two high resolution structural arrangements have beenrecently reported for the bovine mitochondrial central stalk in F1 and anE. coli complex of g and e (Gibbons et al., 2000; Rodgers and Wilce,2000). Tsunoda et al. (2001) have been able to trap the two conformationsof the e subunit in E. coli using cross-linking studies thus demonstratingthat both conformations of the e subunit exist in the enzyme complexof E. coli. In the e conformation with the C-terminal domain of the esubunit facing toward F1, ATP hydrolysis is strongly inhibited, but ATPsynthesis is not affected. In the other conformation of the e subunit withthe C-terminal domain of e towards the F0, the enzyme operates withequal efficiency in either direction (i.e. ATP synthesis or hydrolysis).It should be noted that the C-terminal domain of the e subunit of E. coli


is not required for growth on succinate indicating that the domain isdispensable for ATP synthesis. Some bacterial species lack this domain(Dunn, 1995). Suzuki et al. (2003) have investigated the factors thatinfluence the two transitions. In the ‘‘up-state’’, the two helices of e arefully extended and insert into F1. Without added nucleotide, e is in the up-state and is stabilized by the �mHþ . ATP addition induces the transitionto the down-state and ADP counteracts the action of ATP. Based onthese observations, the authors propose that increases in the �mHþ andADP concentration transform the e subunit into the up-state confor-mation thus gearing the enzyme towards ATP synthesis (Suzuki et al., 2003).Recent work suggests that the e subunit of some bacteria is capable ofbinding ATP and Yoshida and co-workers propose that the e subunit actsas a built-in cellular sensor of ATP concentration. An alternative hypo-thesis proposes that e is a coupling factor and undergoes conformationalchanges in response to both �mHþ and nucleotide occupancy, and thatthe resultant conformation allows rotation only in the direction thatwould result in coupling (Cipriano et al., 2002). In particular, the rotormust not be allowed to rotate in the direction of ATP synthesis if ADPand Pi are not bound. Preventing this may be e’s principle role.From a mechanistic point of view the most intriguing question

concerns the molecular details of how the F1F0 ATP synthases fromalkaliphilic bacteria manage to block the ATP hydrolysis direction whichmay become thermodynamically favorable under certain environmentalconditions. We have demonstrated that the preferential blockage of ATPhydrolysis activity and the uncoupling by LDAO was intrinsic to the F1

moiety (Cook et al. 2003). Based on these data, it is tempting to proposethat the e subunit of the ATP synthase from strain TA2.A1 is permanentlyfixed in a conformation in which the rotational movement in ATPhydrolysis direction is impaired. Perhaps the ATP synthases fromalkaliphilic bacteria represent a unique group of enzymes that have evolvedto work only in one direction to prevent wasteful ATP hydrolysis underlow energy conditions.

ACKNOWLEDGMENTS

Work in GMC’s laboratory was supported by a Marsden grant fromthe Royal Society of New Zealand, work in PD’s laboratory was supportedby the Swiss National Science Foundation and Research Commission ofETH Zurich.


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Dissimilatory Fe(III) and Mn(IV) Reduction

Derek R. Lovley*, Dawn E. Holmes and Kelly P. Nevin

Department of Microbiology, University of Massachusetts-Amherst,

Amherst, MA 01003, USA

ABSTRACT

Dissimilatory Fe(III) and Mn(IV) reduction has an importantinfluence on the geochemistry of modern environments, andFe(III)-reducing microorganisms, most notably those in theGeobacteraceae family, can play an important role in thebioremediation of subsurface environments contaminated with organicor metal contaminants. Microorganisms with the capacity to conserveenergy from Fe(III) and Mn(IV) reduction are phylogeneticallydispersed throughout the Bacteria and Archaea. The ability to oxidizehydrogen with the reduction of Fe(III) is a highly conservedcharacteristic of hyperthermophilic microorganisms and one Fe(III)-reducing Archaea grows at the highest temperature yet recorded forany organism. Fe(III)- and Mn(IV)-reducing microorganisms have theability to oxidize a wide variety of organic compounds, oftencompletely to carbon dioxide. Typical alternative electron acceptorsfor Fe(III) reducers include oxygen, nitrate, U(VI) and electrodes.Unlike other commonly considered electron acceptors, Fe(III) andMn(IV) oxides, the most prevalent form of Fe(III) and Mn(IV) inmost environments, are insoluble. Thus, Fe(III)- and Mn(IV)-reducingmicroorganisms face the dilemma of how to transfer electronsderived from central metabolism onto an insoluble, extracellular

*Corresponding author. Tel.: 413-545-9651; Fax: 413-545-1578;

E-mail: [email protected]



DOI 10.1016/S0065-2911(04)49005-5

electron acceptor. Although microbiological and geochemicalevidence suggests that Fe(III) reduction may have been the first form ofmicrobial respiration, the capacity for Fe(III) reduction appears tohave evolved several times as phylogenetically distinct Fe(III) reducershave different mechanisms for Fe(III) reduction. Geobacter species,which are representative of the family of Fe(III) reducers thatpredominate in a wide diversity of sedimentary environments, requiredirect contact with Fe(III) oxides in order to reduce them. In contrast,Shewanella and Geothrix species produce chelators that solubilizeFe(III) and release electron-shuttling compounds that transfer electronsfrom the cell surface to the surface of Fe(III) oxides not in directcontact with the cells. Electron transfer from the inner membrane to theouter membrane in Geobacter and Shewanella species appears toinvolve an electron transport chain of inner-membrane, periplasmic,and outer-membrane c-type cytochromes, but the cytochromes involvedin these processes in the two organisms are different. In addition,Geobacter species specifically express flagella and pili during growth onFe(III) and Mn(IV) oxides and are chemotactic to Fe(II) and Mn(II),which may lead Geobacter species to the oxides under anoxicconditions. The physiological characteristics of Geobacter speciesappear to explain why they have consistently been found to be thepredominant Fe(III)- and Mn(IV)-reducing microorganisms in avariety of sedimentary environments. In comparison with otherrespiratory processes, the study of Fe(III) and Mn(IV) reduction isin its infancy, but genome-enabled approaches are rapidlyadvancing our understanding of this environmentally significantphysiology.

1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2212. Environmental considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222

2.1. Environments in which Fe(III) and Mn(IV) reduction is or hasbeen important . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222

2.2. Sources of electron donors . . . . . . . . . . . . . . . . . . . . . . . . . . 2262.3. Forms of Fe(III) and Mn(IV) available for reduction and influence of

humic substances. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2273. Major groups of Fe(III)-and Mn(IV)-reducing microorganisms . . . . . . . . . . . 237

3.1. Microorganisms that do not conserve energy to support growthfrom Fe(III) reduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237

3.2. Microorganisms that conserve energy to support growth fromFe(III) reduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

220 DEREK R. LOVLEY, DAWN E. HOLMES AND KELLY P. NEVIN

4. Physiological diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2434.1. Alternative electron acceptors . . . . . . . . . . . . . . . . . . . . . . . . . 2444.2. Electron donors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2504.3. Temperature range, pH, and salinity ranges . . . . . . . . . . . . . . . . . 2534.4. Nitrogen fixation and autotrophy . . . . . . . . . . . . . . . . . . . . . . . 254

5. Mechanisms for Fe(III) and Mn(IV) reducution. . . . . . . . . . . . . . . . . . . . 2545.1. Strategies for Fe(III) oxide reduction – direct contact versus electron

shuttling or chelation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2555.2. Models for electron transfer to extracellular Fe(III) and Mn(IV) oxides . . . 258

6. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

1. INTRODUCTION

Dissimilatory Fe(III) and Mn(IV) reduction refers to the process in whichmicroorganisms reduce Fe(III) or Mn(IV) for purposes other thanassimilation of iron or manganese. The ability of microorganisms toreduce Fe(III) or Mn(IV) has been known since early in the 20th century(Harder, 1919; Allison and Scarseth, 1942). However, the capacity for somemicrobes to conserve energy to support growth via the oxidationof hydrogen (Balashova and Zavarzin, 1980) or organic compounds(Lovley et al., 1987; Lovley and Phillips, 1988b) was only discovered in the1980s. Of the known major forms of respiration in soils and sediments,which also include reduction of oxygen, nitrate, and sulfate as well asmethanogenesis (Reeburgh, 1983; Lovley and Chapelle, 1995), Fe(III) andMn(IV) reduction were the last to be discovered and have been theleast studied. However, as the importance of Fe(III) and Mn(IV)reduction to anaerobic degradation of organic matter (Lovley, 1991;Thamdrup, 2000) and the geochemistry of soils and sediments(Lovley, 1995b) has become apparent, interest in dissimilatory Fe(III) andMn(IV) reduction has grown. Investigations into this process have alsobeen stimulated by the possibility that Fe(III) reduction was one of thefirst forms of microbial respiration (Vargas et al., 1998; Lovley, 2000a,2003c; Tor et al., 2003). Interest in this form of respiration also stemsfrom practical applications such as the use of dissimilatory Fe(III) reducersin the bioremediation of subsurface environments contaminated withorganic and/or metal contaminants (Lovley, 1995a, 1997b, 2003b) and theharvesting of electricity from aquatic sediments and waste organic matter(Bond et al., 2002; Bond and Lovley, 2003; Chaudhuri and Lovley, 2003;Holmes et al., 2004b).

DISSIMILATORY Fe(III) AND Mn(IV) REDUCTION 221

As detailed below, there are now many species of Bacteria andArchaea that are known to grow with Fe(III) and/or Mn(IV) as the soleterminal electron acceptor. The availability of the full genome sequenceof a number of these organisms and the development of additionalmolecular tools for analyzing their physiology is greatly accelerating theunderstanding of dissimilatory Fe(III) and Mn(IV) reduction. The purposeof this review is to provide an overview of what is known about thephysiology of dissimilatory Fe(III) and Mn(IV) reduction. Most organismsthat reduce Fe(III) also reduce Mn(IV) and vice versa. Therefore, theprocess of Fe(III) and Mn(IV) reduction will be referred to as Fe(III)reduction for brevity, except when it is important to make a distinctionbetween the two.

2. ENVIRONMENTAL CONSIDERATIONS

2.1. Environments in which Fe(III) and Mn(IV) Reduction Is orHas Been Important

2.1.1. Pristine Sediments, Soils, and Subsurface Environments

One of the primary reasons for investigating the physiology of anyorganism is to better understand its influence on the environments in whichit is found and to gain insight into what environmental factors control itsgrowth and activity. Microbial Fe(III) and Mn(IV) reduction are importantprocesses in a diversity of anoxic environments in which organic matter and/or hydrogen as well as Fe(III) and Mn(IV) are available. For example,Fe(III) and Mn(IV) reduction are responsible for the anaerobic oxidationof substantial amounts of organic carbon is freshwater and marinesediments as well as submerged soils (Lovley, 1991, 1995b; Thamdrup,2000). The importance of Fe(III) and Mn(IV) reduction in processingorganic matter in aquatic sediments and submerged soils can be attributedto the abundance of iron and manganese in most of these environmentsand the fact that once Fe(II) and Mn(II) are produced from Fe(III) andMn(IV) reduction, Fe(III) and Mn(IV) are often rapidly regenerated(Thamdrup, 2000). This is because Fe(II) and Mn(II) are relatively solubleand tend to diffuse to the oxic/anoxic interface where they are oxidizedback to Fe(III) and Mn(IV). Bioturbation can greatly accelerate thisprocess. It has been estimated that in sediments each iron atom can go


through as many as 100 cycles of reduction and oxidation prior topermanent burial (Thamdrup, 2000).In a similar manner, when soil-feeding termites ingest soil, the organic

matter and Fe(III) enter the anaerobic intestinal system where substantialamounts of organic matter can be oxidized with the reduction of Fe(III)(Kappler and Brune, 2002). Presumably the reduced iron is reoxidized whenit is excreted back into the aerobic soil and thus can cycle many times inthe degradation of soil organic matter.Fe(III) and Mn(IV) reduction are also important processes in subsurface

environments. In pristine aquifers, which are often important sources ofdrinking water, Fe(III) and Mn(IV) reduction can generate substantialquantities of dissolved Fe(II) and Mn(II) (Lovley et al., 1990; Lovley,1997a). As drinking water is pumped to the surface and contacts oxygen,the Fe(III) and Mn(IV) oxides that are generated can plug plumbingand stain just about everything they contact. High concentrations ofFe(II) or Mn(IV) in groundwater is one of the most prevalent ground-water quality problems worldwide. However, Fe(III)-reducing micro-organisms outcompeting sulfate reducers (Lovley and Phillips, 1987a) inthe subsurface can prevent the production of toxic sulfides (Chapelle andLovley, 1992), which are even less desirable than highly dissolved iron(Lovley, 1997a).

2.1.2. Contaminated Environments

Fe(III) reduction is known to be an important process for the degradationof contaminants in groundwater polluted by petroleum, landfill leachates,or similar wastes (Lovley et al., 1989a; Anderson and Lovley, 1997;Lovley and Anderson, 2000; Bin et al., 2002; Roling and van Verseveld,2002). Enhanced microbial activity depletes the oxygen in such con-taminated environments, and Fe(III) is, in general, the most abundantelectron acceptor for organic matter degradation (Lovley, 1991, 1997b). Therate of contaminant degradation coupled to Fe(III) reduction in aquifersediments can be stimulated with the addition of compounds that makeFe(III) more accessible for microbial reduction such as Fe(III) chelators(Lovley et al., 1994, 1996b) or compounds such as humic substancesand other extracellular quinones which shuttle electrons from the cellsurface to the surface of Fe(III) oxides (Lovley et al., 1996a, 1998). Inthe presence of chelators and electron shuttles even compounds whichare often difficult to degrade under anoxic conditions, such as benzene(Lovley et al., 1994, 1996b; Anderson et al., 1998), chlorinated compounds


(Bradley et al., 1998) and methyl-tert-butyl ether (MTBE) (Finneran andLovley, 2000), are broken down.Fe(III) reduction is also an important process in aquifers undergoing

bioremediation for uranium. Uranium is soluble in the oxidized form,U(VI), but dissimilatory metal reducers can reduce U(VI) to U(IV), which isinsoluble and precipitates from groundwater (Lovley et al., 1991a; Lovleyand Phillips, 1992a; Lovley, 1995a). A successful strategy for removinguranium from contaminated groundwater is to add acetate to thesubsurface (Anderson et al., 2003b). The acetate stimulates the growthof Geobacteraceae species, which primarily grow via the reduction ofFe(III) oxides in the subsurface (Finneran et al., 2002a). However, theGeobacteraceae will simultaneously reduce U(VI) to U(IV) and effectivelyremove it from the groundwater (Finneran et al., 2002a; Anderson et al.,2003b).

2.1.3. Hot Environments and Fe(III) Reduction as the First Form ofMicrobial Respiration

Although less studied, Fe(III) reduction may also be an important processin some hydrothermal environments (Brock et al., 1976; Jannasch, 1995;Karl, 1995; Tor et al., 2003). Most hydrothermal fluids contain highconcentrations of Fe(II). As the fluids emerge into air in terrestrialenvironments, or cold, highly oxygenated waters at the bottom of theocean, the Fe(II) is rapidly oxidized to Fe(III). Rates of Fe(III) reductionin hydrothermal environments have yet to be adequately documented,but as discussed below, most, if not all, hyperthermophiles have thecapacity for Fe(III) reduction. Thus, there is significant potential forFe(III) reduction in hot environments. In addition to using hydrogen asan electron donor, hyperthermophilic Fe(III)-reducing microorganisms arecapable of using a wider range of organic electron donors than has beendocumented in hyperthermophiles using other electron acceptors (Torand Lovley, 2001; Tor et al., 2001). In fact, some hyperthermophiles thatuse multiple electron acceptors can utilize a wider range of electron donorsto support growth with Fe(III) than those with alternative electronacceptors (Vargas et al., 1998; Tor and Lovley, 2001). Fe(III) may also beavailable for microbial reduction in subsurface petroleum reservoirsfrom which thermophilic Fe(III) reducers have been recovered (Greeneet al., 1997; Slobodkin et al., 1999a). Large accumulations of ultrafine-grained magnetite, similar to that produced by many Fe(III)-reducingmicroorganisms have been found at depths as great as 6.7 km below


land surface (Gold, 1992). It has been suggested that this provides evi-dence for the activity of Fe(III)-reducing microorganisms in a deep, hotbiosphere on Earth, as well as possibly other planets (Gold, 1992).Microbiological and geochemical evidence also suggests that Fe(III)

was an important process on early Earth (Lovley, 2003c). The findingthat dissimilatory Fe(III)-reducing microorganisms produced copiousquantities of ultrafine-grained magnetite when oxidizing organic compoundsto carbon dioxide with the reduction of Fe(III) oxide (Lovley et al., 1987;Lovley, 1990) provided a potential explanation for the coincidentaccumulation of isotopically light carbonates associated with magnetiteaccumulations in PreCambrian Banded Iron Formations (Baur et al., 1985;Walker, 1987).Evidence for the importance of Fe(III) reduction even earlier in the

Earth’s history is more circumstantial, but a strong case can be madefor Fe(III) reduction preceding other commonly considered forms ofmicrobial respiration (Lovley, 2000a, 2003c). For example, the capacityfor Fe(III) reduction is much more universal among hyperthermophilicBacteria and Archaea than the ability to use other electron acceptors, suchas oxygen, nitrate, sulfur compounds, or carbon dioxide (Lovley, 2000a;Tor et al., 2003). This is significant because it is often proposed that lifeemerged on a hot, early Earth and that that the physiological characteris-tics of hyperthermophiles, which, among known organisms, are the mostclosely related to the last common ancestor(s), give insight into thephysiology of the last common ancestor(s) (Baross and Hoffman, 1985;Pace, 1991; Holm, 1992; Bock and Goode, 1996). Furthermore, analysisof possible geochemical conditions on pre-biotic Earth have suggestedthat conditions were ideal for the development of life forms that could takeadvantage of the availability of substantial quantities of hydrogen andFe(III) oxide and that life emerged from inorganic membranes thatcatalyzed hydrogen oxidation coupled to Fe(III) reduction (Russell et al.,1998; Russell and Hall, 2002). Of the other commonly considered electronacceptors, only carbon dioxide was also likely to have been readily availableas an electron acceptor on pre-biotic Earth. However, the complexbiochemistry required for hydrogen oxidation coupled to carbon dioxidereduction and the fact that neither methanogenesis or acetogenesis, theprimary pathways for this type of respiration, is a common form ofrespiration in the most deeply branching hyperthermophiles suggests thatcarbon dioxide reduction was not an early form of respiration (Lovley,2003c). Thus, at the present time the combination of microbiological andgeochemical evidence most strongly favors Fe(III) reduction as the firstform of microbial respiration. However, this should not be interpreted to


suggest that all modern Fe(III)-reducing microorganisms have a highlyconserved mechanism for Fe(III) reduction. Rather, as discussed below, itappears that phylogenetically distinct Fe(III) reducers have significantlydifferent strategies for Fe(III) reduction.

2.1.4. Energy Harvesting Electrodes in Sediments

A somewhat artificial environment in which Fe(III)-reducing microorgan-isms are important is on the surface of energy-harvesting electrodesplaced in aquatic sediments. As detailed below, a number of Fe(III)-reducing microorganisms can effectively transfer electrons to electrodesurfaces and conserve energy to support growth from this process.Therefore, when an anode is placed within anoxic sediments it will beheavily colonized with Fe(III)-reducing microorganisms, most notablyorganisms in the family Geobacteraceae (Bond et al., 2002; Holmes et al.,2004b).

2.2. Sources of Electron Donors

Understanding the source of electron donors available to supportmicrobial Fe(III) reduction is important in order to relate the physiologicalcharacteristics of these organisms to their potential environmentalrole. In most sedimentary environments the source of electron donors forFe(III) and Mn(IV) reduction is the complex organic matter depositedwithin the sediments (Lovley, 1991). Although there are Fe(III)-reducingmicroorganisms which can utilize sugars and amino acids as electrondonors (see below), they do not appear to be competitive with fermentativemicroorganisms in sedimentary environments (Lovley and Phillips, 1989).Current information suggests that microorganisms not directly involvedin substantial Fe(III) reduction break down the complex organicmatter to fermentation products, which are the primary electron donorsfor Fe(III) reduction (Lovley, 2000a). As in anoxic environments inwhich sulfate reduction is the predominant terminal electron-acceptingprocess, acetate is the primary fermentation intermediate, but other minorfermentation acids are also produced (Lovley and Phillips, 1989; Lovley andChapelle, 1995; Kusel et al., 2002b). The extent to which hydrogen servesas an intermediate in the oxidation of organic matter coupled to Fe(III)and Mn(IV) reduction is not clear, but hydrogen may be less importantthan in sulfate-reducing and methanogenic environments, because many


fermentative microorganisms have the ability to divert electron flow toFe(III) that might otherwise be used to produce hydrogen (see below).There are Fe(III)-reducing microorganisms (Table 1) that can metabolizethe monoaromatic compounds (Lovley and Lonergan, 1990; Coates et al.,1996) and long-chain fatty acids (Coates et al., 1995, 1999) that are releasedfrom complex organic matter and oxidize them to carbon dioxide withFe(III) serving as the sole electron acceptors.In hydrothermal environments, hydrogen, often present in high concen-

trations in hydrothermal fluids, may be an important electron donoras may sulfur. Recently, it has become apparent that there are hyper-thermophilic Fe(III) reducers which can oxidize important organic carbonsources such as acetate (Tor et al., 2001), monoaromatic compounds (Torand Lovley, 2001), and long-chain fatty acids (Kashefi et al., 2002b).In fact at the present time Fe(III)-reducing hyperthermophiles are theonly organisms available in pure culture known to be capable of metabo-lizing these compounds in hot (i.e.>80�C) environments.

2.3. Forms of Fe(III) and Mn(IV) Available for Reduction andInfluence of Humic Substances

Fe(III) and Mn(IV) are highly insoluble at non-acidic pH. Although itis clear that dissimilatory Fe(III)-reducing microorganisms can reducepoorly crystalline Fe(III) oxides (Lovley and Phillips, 1986; Phillips et al.,1993) and structural Fe(III) in clays (Kostka et al., 2002; Shelobolina et al.,2003a, 2003d), there has been considerable debate over whether highlycrystalline Fe(III) oxides can serve as an electron acceptor for microbialreduction in natural environments. A survey of the literature in this areademonstrates that studies which have suggested that crystalline Fe(III)oxides are an important electron acceptor have investigated Fe(III)reduction under highly artificial conditions which promote the reductionof crystalline Fe(III) oxides (Anderson et al., 2004; Glasauer et al., 2003).It has been suggested that culturing microorganisms in rich media maypermit them to produce reductases that are not synthesized under nutrient-poor conditions found in many environments (Glasauer et al., 2003).Furthermore, the addition of high concentrations of organic acids suchas lactate which chelate and solubilize Fe(III) may artificially promoteFe(III) oxide reduction. When conditions that more closely represent thosefound in sedimentary environments are employed then crystalline Fe(III)oxides are not reduced (Glasauer et al., 2003). Even more relevant is thefinding that crystalline Fe(III) oxides persist in sediments when natural


Table 1 Organisms known to conserve energy to support growth from Fe(III) reduction.

Organism Source Electron donors

oxidized with Fe(III)aOxidation

with Fe(III)bFe forms

reducedcOther electron

acceptorsdGrowth

Temp (�C)

Mor-

phology

Referencee

Acidimicrobium

ferroxidans TH3

Copper leach dump Glyc ND Fe(III)-sulfate O2 45 Rod Bridge and Johnson,

1998

Acidiphilium cryptum

JF-5

Acidic coal mine lake Cit, EtOH, Fum, Fru,

Glm, Glu, Glyc, H2,

Mal, Succ, Xyl

Complete Fe(III)–P O2 30 Rod Kusel et al., 1999

Acidithiobacillus

ferroxidans

Acid, bituminous coal

mine effluent

H2, S� ND Fe2(SO4)3 O2, S

� 30 Rod Das et al., 1992;

Pronk et al., 1992;

Ohmura et al., 2002

Aeromonas hydrophila Freshwater and

sewage

Glyc, Lac, Succ Complete PCIO, Fe(III)-Cit U(VI), Co(III), selenate,

nitrate, Fum, O2

37 Rod Knight and Blakemore,

1998

Aeronionas hydro-

phila strain 45/90

Microbial fuel cell Glu, Glyc, Pyr ND Fe(III)-cit electrode, O2,

nitrate, SO2�4

30 Rod Pham et al., 2003

Alicyclobacillus-like

Isolate Y004

Geothermal acidic site Glu ND Fe(III)-sulfate O2 55 Rod Johnson et al., 2003

Anaeromyxobacter

dehalogens

Freshwater sediment Ac Complete Fe(III)-cit, Fe(III)–P,

PCIO

Fum, nitrate, nitrite,

O2, OHP

30 Rod He and Sanford, 2003

Anaeromyxobacter

Strain FAc12

Drained rice field Ac Complete Fe(III)-cit, PCIO O2, nitrate 30 Rod Treude et al., 2003

Bacillus

arseniciselenatis

Mono Lake sediment Lac Incomplete Fe(III)-NTA As(V), Fum, nitrate,

Se(VI)

20 Rod Blum et al., 1998

Bacillus infernus Deep subsurface For, Lac Incomplete Fe(III)-Cl3 Mn(IV), nitrate, TMAO 60 Rod Boone et al., 1995

Bacillus subterraneus Deep subterranean

thermal waters

EtOH, Fru, Glyc, Glu,

Lac, Mat, starch,

Suc, Xyl, YE

ND PCIO Mn(IV), nitrate,

nitrite, Fur, O2

37 Rod Kanso et al., 2002

Clostridium beijerinckii Freshwater sediment Glu Incomplete Fe(III)(maitol) 3,

Fe(III)-cit

ND 37 Rod Dobbin et al., 1999b

Clostridium sp. EG3 Microbial fuel cell

using starch

processing wastewater

Glu Incomplete Fe(III)–P, PCIO Electrode 37 Rod Park et al., 2001

Deferribacter abyssi Hydrothermal vent Ac, Succ, H2 ND PCIO, Fe(III)-Cit Nitrate, S� 60 Rod Miroshnichenko et al.,

2003

Deferribacter

thermophilus

North Sea oil field Ac, CAA, H2, Lac,

Mal, Pept, Pyr,

Succ Try, Valr, YE

ND PCIO, Fe(III)-Cit Mn(VI), nitrate 60 Rod Greene et al., 1997

228

Desulfitobacterium

frappieri strain PCP-1

Methanogenic

consortium

Lac ND Fe(III)–P, PCIO As(V), Fum, Mn(IV),

PCP, Se(VI), S�, SO2�3 ,

S2O2�3 ,*-TCPg

38 Rod Bouchard et al., 1996;

Niggemyer et al.,

2001

D. frappieri strain G2 Subsurface smectite

bedding

Buty, BtOH, Cit,

EtOH, For, H2,

Lac, Mal, Pyr

Complete Fe(III)-cit, Fe(III)-

NTA, Fe(III)–P,

PCIO, smectite

AQDS, Fum, Nitrate,

PCE, SO2�3 , S2O

2�3 ,

TCE, U(VI)

30 Rod Shelobolina et al.,

2003d

Desulfitobacterium

hafniense

Municipal sludge Lac ND PCIO, Fe(III)–P As(V), Fum, Mn(IV)

nitrate, S�, SO2�3 ,

S2O2�3 , Se(VI), PCP,

*-TCPg

37 Rod Niggemyer et al., 2001

Desulfitobacterium

metallireducens

Uranium contaminated

aquifer sediment

Lac, For ND Fe(III)-Cit,

Fe(III)-NTA

AQDS, humics, S2O2�3 ,

Mn(IV), Cr(VI), S�,

Selenite, TCE, PCE,

3-chloro-4-HPE

30 Rod Finneran et al., 2002b

Desulfitobacterium

strain GBFH

Arsenic contaminated

sediments

Lac ND PCIO, Fe(III)–P As(V), Fum, Mn(IV), S�,

SO2�3 , S2O

2�3 , Se(VI)

37 Rod Niggemyer et al., 2001

Desulfobulbus

propionicus

Freshwater mud H2, Lac, Prop, Pyr Incomplete Fe(III)-cit,

Fe(III)-NTA,

Fe(III)–P, PCIO

Electrode, nitrate, nitrite,

O2, SO2�4

37 Rod Holmes et al., 2003

Desulfosporosinus

meridiei

Gasoline-contaminated

ground water

Lac Incomplete Fe(III) SO2�4 , SO2�

3 , S2O2�3 , S�,

DMSO

28 Rod Robertson et al., 2001

Desulfolomaculum

reducers

Heavy metal

contaminated

sediments

Buty, Lac, Valr Incomplete Fe(III)-cit Cr(VI), dithionite,

Mn(IV), SO2�4 , S2O

2�3 ,

S�, U(VI)

37 Rod Tebo and Obraztsova,

1998

Desulfovibrio

profundus

Deep undersea

sediment

Ac, H2, Pyr, Tol ND PCIO Fum, Lignosulfate,

nitrate, SO2�3 , S2O

2�4

25 Vibrio Bale et al., 1997

Desulfuromonas

acetexigens

Anoxic muds Ac Complete PCIO Mn(VI), S�, polysulfides,

Fum, Mal

30 Rod Coates et al., 1995

Desulfuromonas

acetoxidans

Marine sediments Ac, BtOH, EtOH

Prop, Pyr

Complete Fe(III)-Cit,

Fe(III)-NTA

Mn(IV) Glut, Mal, Furn 30 Rod Roden and Lovley,

1993

Desulfuromonas

chloroethenica

Freshwater sediments Ac, Pyr ND Fe(III)-NTA PCE, TCE, Fum, S2� 21–31 Rod Krumholz, 1997

Desulfuromonas

michiganensis

Freshwater sediment Ac, Fum, Lac, Mal,

Pyr, Succ

ND PCIO, Fe(III)-cit Fum, Mal, PCE, S�, TCE 25 Rod Sung et al., 2003

Desulfuromonas

palmitatis

Marine sediments Ac, Fum, Lac, Lau,

Pal, Ste, Succ

Complete PCIO, Fe(III)-Cit,

Fe(III)-NTA,

Fe(III)–P

Mn{IV), AQDS, S�, Fum 40 Rod Coates et al., 1995

Desulfuromusa bakii Marine and freshwater

muds

Ac Complete Fe(III)-NTA S�, Mal, Fum 25 Rod Lonergan et al., 1996

(Continued )

229

Table 1 Continued.





acceptorsdGrowth

Temp (�C)

Mor-

phology

Referencee

Desulfuromusa

kysingii

Freshwater anoxic

muds

Ac Complete Fe(III)-Cit,

Fe(III)-NTA

S�, Mal, Fum, DMSO,

nitrate

30 Rod Liesack and Finster,

1994; Lonergan

et al., 1996

Desulfuromusa

succnoxidans

Marine sediments Ac Complete Fe(III)-NTA S�, Mal, Fum 30 Rod Lonergan et al., 1996

Ferribacterium

limneticum

Mine-impacted lake

sediments

Ac Complete PCIO, Fe(III)–P Nitrate, Fum 25 Rod Cummings et al., 1999

Ferrimonas balearica Marine sediments Lac ND PCIO, F(III)-Cit Mn(IV), nitrate 37 Rod Rossello-Mora et al.,

1995

Ferroglobus placidus Hydrothermal vent Ac, Bz, Bzo, Cinn, Ph,

p-HB, p-HBz

Complete PCIO Nitrate, S2O2�3 85 Coccoid Tor and Lovley, 2001,

Tor et al., 2001

‘‘Geobacter argillaceus’’ Subsurface clay

beddings

Ac, BuOH, Buty,

EtOH, Glyc, Lac,

Pyr, Valr

ND Fe(III)-Cit,

Fe(III)-NTA,

Fe(III)–P, PCIO,

Smectite

AQDS, Fum, nitrate,

Mn(IV), S�30 Rod Shelobolina et al., 2004

‘‘Geobacter

bemidjiensis’’

Subsurface sediment Ac, Bzo, BuOH, Buyr,

EtOH, H2, IsoB,

Lac, Mal, Prop, Pyr

Succ, Valr

Complete Fe(III)-Cit,

Fe(IIl)-NTA,

Fe(III)–P, PCIO

AQDS, Fum, Mal,

Mn(IV)

30 Rod Shelobolina et al., 2004

Geobacter bremensis Freshwater ditch Ac, BtOH, Buty, Bzo,

EtOH, For, Fum,

H2, Lac, Mal, Prop,

PrOH, Pyr, Succ

ND PCIO Mn(IV), S�, Fum, Mal 30 Rod Straub and Buchholz-

Cleven, 2001

Geobacter chapellei Deep subsurface Ac, EtOH For, Lac Complete PCIO, Fe-NTA Mn(IV), AQDS, Fum 25 Rod Coates et al., 2000;

Lovley et al., 1990

Geobacter grbiciae Aquatic sediments Ac, Buty, EtOH, For,

H2, Tol, Prop, Pyr

Complete PCIO, Fe(III)-Cit AQDS 30 Rod Coates et al., 2000

Geobacter humireducens Contaminated wetland Ac, EtOH, For, H2,

Lac

Complete PCIO, Fe(III)-Cit Mn(IV), AQDS S�,

nitrate, Fum

30 Rod Coates et al., 1998b

Geobacter

hydrogenophilus

Contaminated aquifer Ac, Buty, Bzo, EtOH,

For, H2, Prop, Pyr,

Suc

Complete PCIO, Fe(III)-Cit AQDS, Fum 30 Rod Coates et al., 2000

230

Geabacter

metallireducens

Aquatic sediments Ac, Bz, BzOH, BtOH,

Buty, Bzo, BzOH,

p-CR, EtOH,

p-HBz, p-HBzo,

p-HBzOH, IsoB,

IsoV, Ph, Prop,

PrOH, Pyr, Tol,

Valr

Complete PCIO, Fe(III)-Cit,

Fe(III)-NTA

Mn(IV), Tc(VII)*, U(VI),

AQDS, humics, nitrate

30 Rod Lovley and Philips,

1988b; Lovley et al.,

1993a

Geobacter pelophilus Freshwater ditch Ac, EtOH, For,

Fum, H2, Mal,

Prop, PrOH, Pyr,

Succ

ND PCIO, Akaganeite Mn(IV), S�, Fum, Mal 30 Rod Straub and Buchholz-

Cleven, 2001

‘‘Geobacter pickeringii’’ Subsurface clay

beddings

Ac, BtOH, Buty,

EtOH, Glyc, H2,

Lac, MeOH, PrOH,

Pyr, Succ, Valr

ND Fe(III)-Cit,

Fe(III)-NTA,

Fe(III)–P, PCIO,

Smectite

AQDS, Fum, Nitrate,

mal, Mn(IV), S�, U(VI)

30 Rod Shelobolina et al.,

2004

Geobacter

sulfurreducens

Contaminated ditch Ac, For, Lac, H2 Complete PCIO, Fe(III)-Cit,

Fe(III)-P

Tc(VII)*, Co(III), AQDS,

S�, Fum, Mal

35 Rod Caccavo et al., 1994

‘‘Geogemma barossii’’ Hydrothermal vent For, H2 Complete PCIO None 105–107 Coccoid Kashefi et al., 2003a

‘‘Geogemma

hydrogenophila’’

Hydrothermal vent H2 Complete PCIO None 95 Coccoid Feinberg et al., 2003

Geoglobus ahangari Hydrothermal vent Ac, Arg, Asg, Buty,

For, Fum, Glm,

Glyc, H2, Isl, Mal,

Pal, Pept, Prop, Pyr,

Ser, Ste, Succ, Valr,

YE

Complete PCIO, Fe(III)-cit None 88 Coccoid Kashefi et al., 2002b

‘‘Geopsychrobacter

electriphilus

Strain A1’’

Anode surface from a

marine sediment

fuel cell

Ac, Ala, Buty, Bzo,

CAA, Cit, Fum,

Glu, Lac, Mal, Pept,

Prop, Pyr, Ste Succ,

Try, YE

Complete Fe(III)-cit,

Fe(III)-NTA,

Fe(III)-P, PCIO

AQDS, Mn(IV), S� 22 Rod Holmes et al., 2004d

‘‘Geopsychrobacter

electrediphlos’’

Strain AZ

Anode surface from a

marine sediment

fuel cell

Ac, Ace, Ala, Asp,

Bzo, CAA, Cit,

EtOH, Fum, Glu,

Gly, H2, Mal, Met,

Pyr, Pept, Succ, Ste

Try, YE

Complete Fe(III)-cit,

Fe(III)-NTA,

Fe(III)–P, PCIO

AQDS, Mn(IV), S� 22 Rod Holmes et al., 2004d

(Continued )

231

Table 1 Continued.





acceptorsdGrowth

Temp (�C)

Mor-

phology

Referencee

Geothermobacter

ehrlichii

Hydrothermal vent Ac, Arg, Asg, Buty,

CAA, EtOH, For,

Glm, His, Isl, Isop,

Mal, MtOH, MSG,

Pept, Prop, Pyr, Ser,

St, Try

Complete PCIO DMSO, Nitrate, Nitrite, 55 Rod Kashefi et al., 2003

Geothermobacterium

ferrireducens

Hot spring H2 Complete PCIO None 85 Rod Kashefi et al., 2002

Geothrix fermentans Contaminated

aquifier

Ac, Lac Complete PCIO, Fe(III)-Cit,

Fe-NTA

Mn(IV), AQDS, S� 30 Rod Coates et al., 1999

Geovibrio ferrireducens Contaminated ditch Ac, CAA, Fum, H2,

Lac, Pro, Prop,

Pyr, Succ, YE

Complete PCIO, Fe(III)-

citrate, F(III)–P

Co(III), S� 35 Vibrio Caccavo et al., 1996

Hyperthermus butylicus Solfataric sea

floor sediment

H2 Complete PCIO S� 99 Coccoid Kashefi et al., 2003

Pantaea agglomerarans Coastal marine

basin

Ac, H2 Complete Fe(III)–P,

Fe(III)-cit,

PCIO

AQDS, Cr(VI), Fum,

Mn(IV), nitrate, O2,

S�, TMA, CoE

30 Rod Francis et al., 2000

Pelabacter carbinolicus Marine sediments EtOH, H2 Incomplete Fe(III)-NTA S� 30 Rod Lovley et al., 1995

Pelobacter propionicus Lac Incomplete Fe(III)-NTA S� 30 Rod Lonergan et al., 1996

Pelobacter venetianus Freshwater sediments EtOH, For, H2 Incomplete Fe(III)-NTA S� 30 Rod Lonergan et al., 1996

‘‘Pseudomonas sp.’’ Swampy soil H2 PCIO Nitrate, O2 Rod Balashova and

Zavarzin, 1980

Pyrobaculum

aerophilum

Marine hydrothermal

waters

H2, Pept, YE ND PCIO, Fe(III)-cit Nitrate, nitrite, O2 100 Rod Kashefi and Lovley,

1999

Pyrobaculum

islandicum

Geothermal water H2, Pept, YE ND PCIO, Fe(III)-cit Mn(IV)*, U(VI)*,

Co(III)*, Tc(VII)*,

Cr(VI)*, Au(III)*,

Cyst, Glut, S�,

SO2�3 , S2O

2�3

100 Rod Kashefi and Lovley,

1999

Pyrodictium occultum Submarine

solfataric field

H2 Complete PCIO S� 105 Coccoid Kashefi et al., 2003

Rhodoferax

ferrireducens

Freshwater bay

sediment

Ac, Bzo, Fru, Glu,

Mal, Lac, Prop,

Pyr, Succ

Complete Fe(III)-NTA, Mn(IV), Fum, nitrate, O2,

electrode

25 Rod Finneran et al., 2003;

Chaudhuri and

Lovley, 2003

232

Shewanella algae Estuarine sediment H2, Lac Incomplete PCIO, Fe(III)-cit Mn(VI), U(VI)*, S2O2�3 ,

AQDS, TMAO,

Fum, O2

30 Rod Caccavo et al., 1992

Shewanella amazonensis Intertidal zone Lac ND PCIO Mn(IV), nitrate, nitrite,

S2O2�3

37 Rod Venkateswaran et al.,

1998

Shewanella frigidimarina Congelation Ice Cell, Glu, Mat, Suc,

Tre, Mann, Prop,

IsoB, Succ, Fum,

Mal, OAA, Leu,

Pro, Phe, Non, Lac

ND PCIO, Fe(III)-P Nitrate, TMAO 20 Rod Bowman et al., 1997

Shewanella gelidimarina Cangelation Ice Glu, Nag, Lac ND PCIO, Fe(III)-P Nitrate, TMAO 15 Rod Bowman et al., 1997

Shewanella olleyana Estuarine sediments Lac Incomplete Fe(III)-Cit, PCIO AQDS, Fum, Nitrate,

Mn(IV), O2, S2O2�3

20 Rod Skerratt et al., 2002

Shewanella oneidensis Aquatic sediments

and other diverse

environments

For, H2, Lac Pyr Incomplete PCIO, Fe(III)-cit Mn(VI), U(VI) S�,

S2O2�3 , AQDS, nitrate,

Furn, O2

30 Rod Myers and Nealson,

1988a; Lovley et al.,

1989b

Shewanella paeleana Accessory nidamental

gland of the squid

Loligo pealei

Lac Incomplete Fe(III)-cit Mn(IV), nitrate, O2, Fum,

S�, S2O2�3 , To

30 Rod Leonardo et al., 1999

Shewanella putrefaciens

CN32

Subsurface For, EtOH, H2,

Lac, Mal

Incomplete PCIO F(III)-cit Mn(IV), SO2�3 , nitrate,

Fum, TmaO

30 Rod Liu et al., 2002

Shewanella

saccharophilus

Aquatic sediments For, Glc, H2, Lac,

Pyr, Suc, YE

Incomplete PCIO, Fe(III)-cit,

Fe-NTA,

Fe(III)–P,

Fe(III)-EDTA

Mn(IV), U(VI)*, S�,

AQDS, S2O2�3 , nitrate,

Mal, Fum, O2

30 Rod Coates et al., 1998

Sulfobacillus acidophilus

strains

ALV and THWX

Self-heating coal

spoil heap

Glyc, Ttt ND Fe(III)-sulfate O2 45 Rod Bridge and Johnson,

1998

S. acidophilus YTF1 Thermal spring Glyc ND Fe(III)-sulfate O2 45 Rod Bridge and Johnson,

1998

Sulfobacillus-like Isolate

strains Y002, Y006,

Y008, Y0010, Y0012,

Y0013, Y0015,

Y0016, Y0017,

YD018

Geothermal acidic

site

Glu ND Fe(III)-sulfate O2 45 Rod Johnson et al., 2003

Sulfobacillus thermo-

sulfidoxidans TH1

Thermal spring Glyc, Ttt ND Fe(III)-sulfate O2 45 Rod Bridge and Johnson,

1998

(Continued )

233

Table 1 Continued.





acceptorsdGrowth

Temp (�C)

Mor-

phology

Referencee

Sulfurospirillum

barnesii

Freshwater marsh For, H2, Lac Incomplete PCIO, Fe(III)-cit Mn(IV), selenate,

arsenate, S2O2�3 ,

S�, nitrite, nitrate,

Fum, TMAO, O2

30 Vibrio Laverman et al.,

1995

Thernroanaerobacter

acetoethylicus

(strains SL26, SL28)

Deep subsurface

petroleum

reservoirs

H2, Pept ND PCIO ND 60 Rod Slobodkin et al.,

1999c

Thermoanaerobacter

brockii M739

Deep subsurface

petroleum

reservoirs


1999c

Thermanoerobacter

siderophilus

Hydrothermal vent H2 – PCIO AQDS, sulfate,

thiosulfate, S�, MnO2

70 Rod Slobodkin et al.,

1999b

Thermanoerobacterr sp.

(strains X513,

X514, X561)

Deep subsurface Ac, Glu, H2, Lac,

Pyr, Succ, Xyl

Incomplete PCIO, Fe(III)-cit Co(III), Cr(VI),

Mn(IV), U(VI)

60 Rod Roh et al., 2002

Thermococcus sp. strain

SN531

Deep sea

hydrothermal vent

Pept, YE ND PCIO ND 80 Coccoid Slobodkin et al.,

2001

Thermococcus sp.

(strains T642,

T739, T13044)

Deep subsurface

petroleum

reservoirs

H2, Pept ND PCIO S� 85 Coccoid Slobodkin et al.,

1999

Thermodesulfobacterium

commune

strain YSRA-1

Hydrothermal

environment

H2 Complete PCIO SO2�4 , S2O

2�3 70 Rod Kashefi et al., 2002

Thermoterrabacterium

ferrireducens

Hot springs

Yellowstone

H2, Glyc Incomplete PCIO, Fe(III)-Cit AQDS, S2O2�3 , Fum 65 Rod Slobodkin et al.,

1997

Thermotoga maritima Geothermally heated

sea floor

H2 ND Fe(III)-Cit S� 80 Rod Vargas et al., 1998

Thermatoga maritima

strain M12597

Deep subsurface

petroleum

reservoirs


1999a

Thermatoga subterranea

strain SL1

Continental oil

reservoirs

H2, Pept ND PCIO Cystine, S2O2�3 60 Rod Slobodkin et al.,

1999a

Thermovenabulum

ferriorganovorum

Terrestrial

hydrothermal

source

BE, CAA, Pept,

Pyr, St, YE

ND PCIO, Fe(III)-cit AQDS, Fum, Mn(IV),

nitrate, SO2�3 ,

S2O2�3 , S�

60 Rod Zavarzina et al.,

2002

234

Thermus strain SA-01 Deep gold-mine

groundwater

Lac ND Fe(III)-Cit,

Fe(III)-NTA

Mn(VI), Co(III)*,

Cr(VI)*, U(Vl)*, S�,

AQDS, nitrate, O2

65 Rod Kieft et al., 1999

Thermus scotoductus

strain NMX2 A.1

Neutral hot spring Lactate Complete Fe(III)-NTA Nitrate, O2, S� 65 Filamentous Kieft et al., 1999

Trichlorobacter thiogenes Anaerobic Soil Ac ND Fe(III)-NTA,

Fe(III)–P

Fum, S�, TCA 25 Rod Nevin et al., 2004

aAbbreviations for electron donors and acceptors: Acetate (Ac), Acetoin (Ace), N-acetylglucosamine (Nag), Anthraquinone-2,6-disulfonic acid (AQDS), Alanine (Ala),

Arginine (Arg), Asparagine (Asg), Aspartate (Asp), Beef Extract (BE), Benzaldehyde (Bz), Benzoate (Bzo), Benzylalcohol (BzOH), 1,2-butanediol (1,2-Bu), Butanol

(BtOH), Butyrate (Buty), Casamino acids (CAA), Casein (Cas), Cellobiose (Cell), t-Cinniminic Acid (Cina), Citrate (Cit), Co(III)-EDTA (CoE), Cystine (Cyst),

Dichlorophenol (DCP), Dimethylsulfoxide (DMSO), Ethanol (EtOH), Fructose (Fru), Formate (For), Fumarate (Fum), Gelatin (GE), Glucose (Glu), Glutamate (Glu),

Glutamine (Glm), Glutathione, oxidized (Glut), Glycerol (Glyc), Glycine (Gly), Histidine (His), p-hydroxybenzoate ( p-HB), p-hydroxybenzaldehyde ( p-HBz),

p-hydroxybenzylalcohol ( p-HBzOH), Hydroxyphenylacetate (HPE), p-cresol ( p-Cr), Hydrogen (H2), Inositol (Ino), Isobutyrate (IsoB), Isoleucine (Isl), Isopropanol

(Isop), Isovalerate (IsoV), Lactate (Lac), Laurate (Lau), Malate (Mal), Maltose (Mat), Maleate (Mle), Mannitol (Mann), Methanol (MtOH), Methionine (Met),

Nonanate (Non), Nitiloacetic acid (NTA), Ortho-substituted halophenols (OHP), Oxaloacetate (OAA), Palmitate (Pal), Pentachlorophenol (PCP) Peptone (Pept),

Phenol (Ph), Phenylalanine (Phe), Proline (Pro), Propanol (PrOH), Propionate (Prop), 1,2-propanediol (1,2-P), Pyruvate (Pyr), Ribose (Rib), Serine (Ser), Starch (St),

Stearate (Ste), Succinate (Succ), Sucrose (Suc), Tetrachloroethylene (PCE), Tetrachlorophenol (TTCP), Tetrathionate (Ttt), Trichloroethylene (TCE), Trimethylamine

N-oxide (TMA), Toluene (Tol), Trehalose (Tre), Trichloroacetic Acid (TCA), Trichlorophenol (TCP), Trimethylene oxide (To), Trimethylamine oxide (TMAO),

Tryptone (Try), Valerate (Valr), Yeast extract (YE), Xylose (Xyl).bComplete oxidation of multicarbon compounds to CO2, or incomplete, typically to acetate.cFe(III) forms: Poorly crystalline iron oxide (PCIO), ferric citrate (Fe(III)-cit), ferric nitriloacetic acid (Fe(III)-NTA), ferric pyrophosphate (Fe(III)-P), Fe(III) chloride

(Fe(III)-C13). Fe(III) ethylenediamine-tetraacetic acid, Not Specified (NS).dOrganism has the ability to reduce the metal but not determined whether energy to support growth is conserved from reduction of this metal.eReference in which the capacity to grow via Fe(III) reduction is described.fND¼Not determined.gChlorinated compounds used by Desulfitobacterium hafniense 2,4,5-TCP, 2,4,6-TCP, 2,4-DCP, 3,5-DCP, 3-Cl-4-OHPA, 2,3,4,5-TTCP, 2,3,4,6-TTCP, 2,3,4-TCP, 2,3,5-

TCP, 2,3,6-TCP, 3,4,5-TCP, 2,6-DCP. Chlorinated compounds used by Desulfitobacterium frappieri strain PCP-1,2,3,4,5-TTCP, 2,3,5,6-TTCP, 2,3,4-TCP, 2,3,5-TCP,

2,3,6-TCP, 2,4,5-TCP, 2,4,6-TCP, 3,4,5-TCP, 3,5-DCP, 2.6-DCP, 2,4-DCP.

235

communities have been examined in situ (Phillips et al., 1993; Andersonet al., 2004), and that the rate and extent of Fe(III) reduction is correlatedwith the availability of poorly crystalline Fe(III) oxide (Lovley and Phillips,1987b; Thamdrup, 2000). One reason for this is that at the concentrationsof acetate and hydrogen found in natural environments, the reduction ofcrystalline Fe(III) oxides is thermodynamically unfavorable (Thamdrup,2000; Anderson et al., 2004).Although insoluble Fe(III) oxides are the predominant form of Fe(III)

in most soils and sediments at circumneutral pH, soluble Fe(III),apparently chelated with organic ligands, has been detected in a variety ofanoxic sediments, submerged soils, and in groundwater (see Nevinand Lovley (2002b) for a review). The concentrations of Fe(III) aretypically low, ca. 5–50 mM, but oxidation of key electron donors for Fe(III)reduction, such as acetate and hydrogen, at environmentally relevantconcentrations (1 mM and 1 nM, respectively) can potentially yield enoughenergy to support cell growth (Nevin and Lovley, 2002b). It is notknown whether this soluble Fe(III) is actually available for microbialreduction as Fe(III) that is strongly chelated may not be readily reduced(Haas and DiChristina, 2002). Soluble Fe(III) could not be detected in anorganic-poor subsurface environment (Nevin and Lovley, 2002b), indicatingthat soluble Fe(III) is not universally available to Fe(III) reducers.Artificially increasing the concentration of soluble Fe(III) in sediments

with the addition of synthetic Fe(III) chelators can greatly acceleratethe metabolism of Fe(III)-reducing microorganisms (Lovley et al., 1994,1996b). The chelators increase the concentration of dissolved Fe(III)(Lovley and Woodward, 1996), thus making Fe(III) more available formicrobial reduction. Furthermore, chelated Fe(III) has a higher redoxpotential which may make Fe(III) reduction more thermodynamicallyfavorable (Thamdrup, 2000). Adding chelators is a potential strategy foraccelerating the degradation of organic contaminants in anoxic subsurfaceenvironments (Lovley et al., 1994, 1996b; Lovley, 1997a).Humic substances, an abundant form of organic matter in many soils

and sediments, may also enhance the availability of insoluble Fe(III)oxides as electron acceptors (Lovley et al., 1996a, 1998). As outlinedbelow, quinone moieties in humic substances can serve as electronacceptors in the respiration of Fe(III)-reducing microorganisms and oncereduced to the hydroquinone state, the humics can abiotically transferelectrons to Fe(III) oxides, producing Fe(II) and regenerating the oxi-dized form of the humic substances. Electron shuttling in this manner cangreatly accelerate the rate of both Fe(III) oxide reduction in aquifersediments and contaminant oxidation coupled to Fe(III) reduction (Lovley


et al., 1996a, 1996b; Bradley et al., 1998; Finneran and Lovley, 2000; Nevinand Lovley, 2000b). Electron shuttles may be particularly beneficial whenFe(III) oxides are occluded within pore spaces that are too small formicroorganisms to enter (Lovley et al., 1998). In addition to humicsubstances, other electroactive organic compounds, such as plant exudates,can stimulate Fe(III) reduction via electron shuttling (Nevin and Lovley,2000b). Electron shuttling compounds at concentrations high enough tostimulate microbial Fe(III) reduction have been detected in pore waters oforganic-rich sediments, but other environments lack significant quantitiesof electron shuttles (Nevin and Lovley, 2002b). Other compounds such asuranium, sulfur, and organic compounds with sulfhydryl groups may alsoserve as electron shuttles for Fe(III) reduction in pure cultures, but donot appear to be important in promoting Fe(III) reduction in sediments(Nevin and Lovley, 2000b).Electron shuttling via humic acids and other extracellular quinones

still leaves Fe(III)-reducing microorganisms with the difficulty oftransferring electrons to an extracellular electron acceptor. However, aswith chelated Fe(III), the reduction of these soluble extracellular electronacceptors is faster than the reduction of insoluble Fe(III) oxides. This mayreflect kinetic constraints in accessing the surface of the Fe(III) oxides.In summary, a detailed accounting of the relative importance of

natural chelators and extracellular quinones in Fe(III) reduction has yet tobe conducted. However, preliminary studies suggest that in all but themost organic-rich sediments, direct reduction of insoluble Fe(III) oxides islikely to be the most important mechanism for Fe(III) oxide reduction(Nevin and Lovley, 2002b).

3. MAJOR GROUPS OF Fe(III)- AND Mn(IV)-REDUCINGMICROORGANISMS

3.1. Microorganisms that Do Not Conserve Energy to SupportGrowth from Fe(III) Reduction

A wide phylogenetic diversity of microorganisms is known to reduce Fe(III)and Mn(IV) in a dissimilatory manner. Many of these microorganismsreduce Fe(III) as a minor side reaction in their metabolism but do notappear to conserve energy to support growth from this electrontransfer (Lovley, 1987, 1991). Extensive lists of such microorganisms areavailable in previous reviews (Lovley, 1987, 2000b). Many of the initial


concepts of the physiology of dissimilatory Fe(III) reduction, such as thepostulated requirement for contact between Fe(III) reducers and Fe(III)oxide for Fe(III) reduction and the inhibition of Fe(III) reduction in thepresence of nitrate and oxygen, were first developed from studies offermentative Fe(III)-reducing microorganisms (Ottow, 1970; Ottow andGlathe, 1971, Munch and Ottow, 1977, 1983). In some of these studies itcould be clearly demonstrated that Fe(III) reduction was an enzymaticprocess, but the mechanisms for Fe(III) reduction were never conclusivelydemonstrated. There has been little further investigation of Fe(III)reduction in fermentative Fe(III)-reducing microorganisms following thediscovery of microorganisms which use Fe(III) as a terminal electronacceptor in respiration because the latter are considered to be responsiblefor most of the Fe(III) and Mn(IV) reduction in soils and sediments.Two other types of microorganisms that reduce Fe(III), but have not

been definitively shown to conserve energy to support growth fromFe(III) reduction are some dissimilatory sulfate-reducing (Coleman et al.,1993; Lovley et al., 1993b) and methanogenic (Bond and Lovley, 2002)microorganisms. In both sulfate reducers and methanogens, hydrogen ismetabolized to a lower minimum threshold in the presence of Fe(III), tolevels that make sulfate reduction or methane production thermodynami-cally unfavorable. The mechanisms for Fe(III) reduction have not beenstudied in detail, but studies with Desulfovibrio vulgaris demonstrated thata c3 cytochrome was involved in the reduction of metals (Lovley, 1993;Lovley et al., 1993c; Lovley and Phillips, 1994). Diversion of electronflow to Fe(III) by sulfate reducers and methanogens, may be an importantcontributing factor in the inhibition of sulfate reduction and methaneproduction in the presence of Fe(III) in sediments (Coleman et al., 1993;Lovley et al., 1993b; Bond and Lovley, 2002).

3.2. Microorganisms that Conserve Energy to Support Growthfrom Fe(III) Reduction

Microorganisms which conserve energy to support growth from Fe(III)reduction are interspersed throughout the Bacteria and Archaea (Fig. 1)and the list of such microorganisms is rapidly growing as interest inthis process intensifies. The basic physiological characteristics of manyof these organisms are summarized in Table 1. Each of these organismsis interesting, but several phylogenetic and/or physiological clusters thathave been studied in significant detail deserve particular note.


3.2.1. Geobacteraceae

The first organisms found to conserve energy from the completeoxidation of organic compounds to carbon dioxide with Fe(III) servingas the sole electron acceptor are in the Geobacteraceae family inthe d-Proteobacteria (Lovley et al., 1987; Lovley and Phillips, 1988b).Many isolates in this family are available (Table 1). Furthermore, whenmolecular techniques which avoid culture bias are used to assess thecomposition of the microbial communities in environments in which Fe(III)reduction is an important process, Geobacteraceae are often the mostabundant microorganisms. This was observed in aquatic sediments(Stein et al., 2001), the Fe(III)-reducing zone of aquifers contaminatedwith petroleum (Anderson et al., 1998; Rooney-Varga et al., 1999;Snoeyenbos-West et al., 2000), or landfill leachate (Roling et al., 2001)as well as in subsurface sediments in which electron donors wereartificially added to stimulate dissimilatory metal reduction (Snoeyenbos-West et al., 2000; Holmes et al., 2002; Anderson et al., 2003b). In onestudy stimulation of Fe(III) reduction did not promote the growth ofGeobacteraceae, but this was because the salinity of the groundwater

Figure 1 Phylogenetic tree, based on 16S rRNA gene sequences, of microorganismsrepresenting genera containing microorganisms reported to be capable of conservingenergy to support growth from Fe(III) reduction.


was extremely high, 10-fold higher than that of seawater, a salinity toohigh for most mesohaline microorganisms, including known Geobacteraceae(Nevin et al., 2003a).The finding that microorganisms closely related to Geobacteraceae

available in pure culture predominate in a diversity of sedimentaryenvironments in which Fe(III) reduction is important suggests thatphysiological studies on the appropriate pure cultures of Geobateraceaecan provide insights into the factors controlling the rate and extentof dissimilatory Fe(III) reduction in sediments (Lovley, 2003a, 2003b). Thisis a rare opportunity in environmental microbiology because the moretypical finding is that the most environmentally relevant organismscannot be readily recovered in culture. Furthermore, of the few examplesof environmentally relevant organisms that are available in pure culture,many grow poorly making it difficult to mass culture them for biochemicalstudies and genetic systems for the organisms have not been developed.In contrast, growing cells of Geobacteraceae for biochemical investigationsis not difficult (Magnuson et al., 2000), and a genetic system for Geobactersulfurreducens, which is likely to be applicable to other members of theGeobacteraceae, is available (Coppi et al., 2001).One of the physiological characteristics that may lead to the

predominance of Geobacteraceae over other, more intensively studied,dissimilatory Fe(III)-reducing microorganisms is the ability of manyof the organisms in this family to use acetate as an electron donor(Table 1). Acetate is expected to be the key intermediate in the anaerobicdegradation of complex organic matter in a variety of sedimentaryenvironments (Lovley and Chapelle, 1995). Geobacter metallireducensand several other Geobacter species also have the ability to oxidize a varietyof aromatic hydrocarbons to carbon dioxide with Fe(III) serving as anelectron acceptor (Table 1), which as noted above, can be an importantprocess for removing aromatic contaminants from polluted aquifers. Theability of several Geobacteraceae to use chlorinated compounds as electronacceptors may also be important in some subsurface environments(Krumholz, 1997; Loffler et al., 2000; Sung et al., 2003).

3.2.2. Shewanella Species

Shewanella species in the c subclass of the Proteobacteria have been themost intensively studied Fe(III)-reducing microorganisms. Shewanella arefacultative organisms, grow rapidly, and are found in a diversity of envi-ronments, including as pathogens and in food spoilage (Venkateswaran,


1999; Heidelberg et al., 2002). Thus, pure cultures of Shewanella arerelatively easy to isolate and mass culture under aerobic conditions. Infact, in many studies of Fe(III) and Mn(IV) reduction in Shewanella species,the cells are first grown aerobically, then placed under anaerobic conditionsin order to study the mechanisms of Fe(III) and Mn(IV) reduction.The often stated justification for the study of Fe(III) reduction in

Shewanella species is that these microorganisms are likely to play animportant role in Fe(III) and Mn(IV) reduction in the environment as wellas in the bioremediation of metal and organic contaminants. This seemsunlikely. There is no evidence that these organism are substantialcontributors to Fe(III) and Mn(IV) reduction in the vast majority ofenvironments in which Fe(III) and Mn(IV) reduction are important.Although Shewanella species can be isolated from environments such asaquatic sediments (Myers and Nealson, 1988a; Caccavo et al., 1992) inwhich Fe(III) and Mn(IV) reduction take place, it is well-known that theability to culture an organism from a particular environment does notnecessarily indicate that it plays an important role in that environment. Thisis because microorganisms that are low in numbers and/or inactive in theenvironments of interest may be recovered in culture. In fact, detailedmolecular studies of a variety of environments in which Fe(III) and Mn(IV)reduction are important have yet to reveal an environment in whichShewanella are prevalent. This was true for aquatic sediments (Stein et al.,2001) and a diversity of subsurface environments (Rooney-Varga et al.,1999; Snoeyenbos-West et al., 2000; Roling et al., 2001; Holmes et al., 2002;Anderson et al., 2003b; Nevin et al., 2003a). Shewanella species couldnot be detected in these environments even when highly sensitivePCR primers designed specifically to detect Shewanella were employed(Snoeyenbos-West et al., 2000).Furthermore, Shewanella species have limited potential for bioremedia-

tion. Although the potential for Fe(III) reduction to be an importantprocess for the degradation of organic contaminants in subsurface envi-ronments is often mentioned in the justification for studying Shewanellaspecies, Shewanella species are not known to oxidize any importantgroundwater contaminants, such as aromatic hydrocarbons. Shewanellaspecies can reduce U(VI) (Lovley et al., 1991a), but have yet to be shownto have a role in U(VI) reduction in subsurface environments and otherorganisms, such as Desulfovibrio species (Lovley and Phillips, 1992a, 1992b),are probably better candidates for use in bioreactors treating uranium-containing wastes.These considerations suggest that it may be difficult to extrapolate the

results of physiological studies on Shewanella species to an understanding


of Fe(III) and Mn(IV) reduction in sedimentary environments. This iscompounded by the fact that the basic physiology of Shewanella speciesdiffers significantly from the Geobacteraceae that predominate in mostenvironments in which Fe(III) and Mn(IV) reduction are important. Forexample, as detailed below, Shewanella species are likely to reduceFe(III) oxides via mechanisms that are significantly different than theGeobacteraceae. Furthermore, the metabolism of organic electron donors byShewanella species is limited. Shewanella species are not known to utilizeacetate, the key electron donor supporting Fe(III) reduction in manyenvironments (Scott and Nealson, 1994). Multicarbon organic electrondonors that are utilized are limited to lactate and pyruvate, which areunlikely to be important free intermediates in the anaerobic degradationof organic matter (Lovley and Chapelle, 1995). Furthermore, thesemulticarbon substrates are only incompletely oxidized to acetate (Lovleyet al., 1989b). Thus, Shewanella species transfer less than half of theelectrons that could be potentially transferred to Fe(III) from thesesubstrates onto Fe(III), which is unlikely to be competitive with micro-organisms which can completely oxidize these substrates with Fe(III)reduction (Snoeyenbos-West et al., 2000).

3.2.3. Hyperthermophilic Microorganisms

All of the hyperthermophiles that have been evaluated have the abilityto oxidize hydrogen with the reduction of Fe(III) (Vargas et al., 1998;Lovley, 2003c; Tor et al., 2003). When Fe(III) reduction by hyperthermo-philes that had been isolated with electron acceptors other than Fe(III)was further investigated in detail, growth with Fe(III) as the sole electronacceptor was possible. These included Thermotoga maritima (Vargaset al., 1998) and Thermodesulfobacterium commune (Kashefi et al., 2002a),as representatives of the Bacteria, as well as the Archaea, Pyrobaculumislandicum (Kashefi and Lovley, 2000) and Ferroglobus placidus (Tor andLovley, 2001; Tor et al., 2001).Providing Fe(III) as an electron acceptor expands the metabolic

capabilities of some Fe(III) reducers beyond what is possible with otherelectron acceptors, such as sulfur compounds. For example, Thermotogamaritima was initially characterized as a fermentative microorganismwhich could divert a small portion of its electron flow to S� with noapparent increase in cell yield over growth in the absence of S�, but in thepresence of Fe(III) T. maritima can grow as a respiratory organismwith hydrogen as the sole electron donor and Fe(III) as the sole electron


acceptor (Vargas et al., 1998). Ferroglobus placidus, which was isolatedas an Fe(II)-oxidizing nitrate reducer (Hafenbradl et al., 1996), wasconsidered to be unable to use organic electron donors to supportgrowth until studies with Fe(III) as the electron acceptor were conducted.In the presence of Fe(III), F. placidus is capable of oxidizing acetate(Tor et al., 2001) and monoaromatic compounds (Tor and Lovley,2001) to carbon dioxide with the reduction of Fe(III). This metabolismyields energy to support cell growth. Along with Geoglobus ahangari(Kashefi et al., 2002b), F. placidus represents the first hyperthermo-phile of any kind definitely shown to oxidize acetate as an electrondonor (Tor et al., 2001). Ferroglobus placidus was also the first hyper-thermophile and Archaea known to oxidize aromatic compounds (Tor andLovley, 2001).Fe(III) appears to be the only electron acceptor for some Fe(III)-reducing

hyperthermophiles (Kashefi et al., 2002a, 2002b; Kashefi and Lovley, 2003).When Fe(III) was used as the electron acceptor for enrichment and isola-tion of microorganisms from a variety of hot environments, some novelmicrobes were discovered. For example, studies in a Yellowstone hot springyielded Fe(III) reducing microorganisms in culture with 16S rRNA genesequences that were closely related to 16S rRNA sequences that wereprevalent in the spring, but the microorganisms had not yet been cultured(Kashefi et al., 2002a). Strain 121 (‘‘Geogemma barossii’’ – Table 1), thefirst organism found to grow at temperatures as high as 121�C and tosurvive exposure at 130�C, was isolated from a hydrothermal vent samplewith Fe(III) as the sole electron acceptor (Kashefi and Lovley, 2003).The physiology of Fe(III) reduction in hyperthermophilic Fe(III)-

reducing microorganisms is of special interest due to the fact that, asdiscussed above, reduction of Fe(III) by organisms living in hot envi-ronments, may have represented the first form of microbial respiration.However, investigations into the mechanisms for Fe(III) reduction inhyperthermophiles has been very limited (Childers and Lovley, 2001).

4. PHYSIOLOGICAL DIVERSITY

There are significant differences in the metabolic capabilities of dissimilatoryFe(III) reducers, which are important to consider in relating the physio-logy of these organisms to their function in the environment and inpredicting the activity of Fe(III) reducers under various environmentalconditions.


4.1. Alternative Electron Acceptors

4.1.1. Oxygen

As noted above, some hyperthermophilic dissimilatory Fe(III)-reducingmicroorganisms are not known to use any electron acceptors other thanFe(III). However, most characterized Fe(III) reducers have the abilityto use one or more alternative electron acceptors (Table 1). For exam-ple, Fe(III) reducers such as Shewanella, Panatoea, Acidiphiilum, andRhodoferax species are facultative organisms and grow as well or betterwith oxygen as the electron acceptor as with Fe(III). Fe(III) reducers thatgrow at circumneutral pH are expected to preferentially reduce oxygenover Fe(III) due to the low solublity and redox potential of Fe(III), butat low pH when Fe(III) is soluble and at a redox potential comparableto that of oxygen, oxygen and Fe(III) may be reduced simultaneously(Kusel et al., 1999, 2002a).Although Fe(III)-reducing microorganisms in the Geobacteraceae

have previously been classified as strict anaerobes, sequencing the genomesof several Geobacteraceae has brought this into question. For example,in the complete genome of G. sulfurreducens (Methe, 2003) there aregenes not only for tolerating oxygen exposure, but also for a putativecytochrome oxidase that might function in oxygen respiration. Moredetailed investigations of this organism have revealed that under theappropriate culturing conditions G. sulfurreducens can grow with oxygenas the sole electron acceptor at concentrations as high as 10% oxygen inthe headspace (Lin et al., 2004). The ability to use oxygen as an electronacceptor is an obvious advantage to Fe(III)-reducing microorganismsbecause Fe(III) will often be most abundant near the oxic–anoxicinterface where Fe(III) reducers are likely to be intermittently exposed tooxygen.

4.1.2. Other Metals

As noted above, most microorganisms that can reduce Fe(III) can alsoreduce Mn(IV). Fe(III)-reducing microorganisms can also potentiallyreduce Mn(IV) indirectly via Fe(III) reduction (Lovley and Phillips,1988a). Fe(II) produced from Fe(III) reduction can abiotically reduceMn(IV) to Mn(II), recycling the Fe(II) to Fe(III). Thus, in the presenceof both Fe(III) and Mn(IV), microorganisms that cannot enzymaticallyreduce Mn(IV) can still have a net electron transfer to Mn(IV).


It is unknown how much of the Mn(IV) in sedimentary environmentsis reduced in this manner, but hydrogen measurements suggest thatdirect Mn(IV) reduction is a likely process, at least in some sediments(Lovley and Goodwin, 1988).In addition to Fe(III) and Mn(IV), many Fe(III) reducers can utilize

other metals as electron acceptors. For example, several dissimilatoryFe(III)-reducing microorganisms have been reported to reduce U(VI)(Lovley et al., 1991a; Lovley and Phillips, 1992b; Kashefi and Lovley, 2000;Anderson and Lovley, 2002; Shelobolina et al., 2003b; Shelobolinaet al., 2003c) and at least two, G. metallireducens and S. oneidensis (for-merly Alteromonas putrefaciens) have been shown to grow with U(VI) asthe sole electron acceptor (Lovley et al., 1991a). Microbial reduction ofU(VI) to U(IV) has environmental implications because U(VI) is highlysoluble in most natural waters and waste streams whereas U(IV) is highlyinsoluble. Reductive precipitation of uranium via microbial U(VI) reductionhas been shown to be effective in removing uranium from a variety ofcontaminated waters in laboratory reactors (Gorby and Lovley, 1991b;Lovley and Phillips, 1992a). However, the broader application for micro-bial U(VI) reduction may be in the in situ treatment of contaminatedgroundwater. Stimulating the growth of Geobacteraceae by adding acetateto groundwater can effectively precipitate uranium in the subsurfaceand prevent its further migration (Anderson et al., 2003b). In a similarmanner, vanadium which G. metallireducens can utilize as the sole elec-tron acceptor to support growth, is removed from groundwater when thegrowth of Geobacteraceae is stimulated in the subsurface (Ortiz-Bernad andLovely, 2003).Other metals that one or more dissimilatory Fe(III)-reducing micro-

organisms can reduce include oxidized forms of cobalt, technetium,chromium, and gold (Lovley and Coates, 2000; Kashefi et al., 2001).However, in most instances, the ability to grow with these metals as asole electron acceptor has not been adequately demonstrated. Althoughof physiological interest, the ability of these metals to support growth maybe of little environmental consequence because even in heavily contaminatedenvironments the concentrations of these metals is likely to be muchless than that of Fe(III), which will typically be the primary electronacceptor supporting growth. In each case the reduction of these metalsmakes them less soluble. Some of these metals, such as technetium andchromium, can also be abiotically reduced by Fe(II) that is produced byFe(III) reducers (Lloyd et al., 2000). A few Fe(III) reducers such asSulfospirillum barnesi can also use metalloids such as selenate and arsenateas electron acceptors (Stolz and Oremland, 1999).


4.1.3. Extracellular Quinones

Many Fe(III)-reducing microorganisms can grow with extracellularquinones as the sole electron acceptor (Lovley et al., 1996a, 1998, 2000;Lovley, 2000b). In natural environments, humic substances may be themost abundant source of extracellular quinones for Fe(III) reducers (Lovleyet al., 1996a; Scott et al., 1998). Humic substances can be technically diff-icult to work with and expensive; therefore, many studies have been con-ducted with the humic substances analog anthraquinone-2,6-disulfonate(AQDS). The quinone moieties in these compounds are reduced to thehydroquinone state (Scott et al., 1998). As noted above, in environmentswhere Fe(III) is available the hydroquinones react with Fe(III), reducing itto Fe(II) and regenerating the quinone state. Therefore, even inenvironments in which the concentrations of soluble humic substancesand other extracellular quinones are low, there can be substantial electronflow through humic substances as each molecule goes through multiplecycles of reduction and oxidation.

4.1.4. Sulfur Compounds

Many dissimilatory Fe(III)-reducing microorganisms have the ability touse S� as an electron acceptor (Table 1). This may reflect the fact thatFe(III) and S� are often found in the same sediment intervals. For exam-ple, as sulfide produced within the sulfate reduction zone of aquatic sedi-ments diffuses into Fe(III)-containing sediments, it is abiotically oxidizedto S�. Around hydrothermal vents, the concurrent oxidation of Fe(II) andsulfide as anoxic hydrothermal fluids interact with cooler aerobic watersmay contribute to a similar co-occurrence of Fe(III) and S�.Although a number of dissimilatory sulfate-reducing microorganisms

can reduce Fe(III) (Coleman et al., 1993; Lovley et al., 1993b), most donot appear to conserve energy to support growth from Fe(III) reduction.Only two organisms, Desulfotomaculum reducens (Tebo and Obraztsova,1998) and Desulfobulbus propionicus (Holmes et al., 2004a) have beenreported to conserve energy to support growth from both Fe(III) andsulfate reduction.

4.1.5. Nitrate

Nitrate is a relatively common alternative electron acceptor for Fe(III)-reducing microorganisms (Table 1). When nitrate and Fe(III) are provided


simultaneously, nitrate is typically reduced prior to net Fe(III) reduction.This is not necessarily due to transcriptional regulation in whichthe presence of nitrate represses the expression of genes involved inFe(III) reduction as nitrate-grown cells can retain the capacity for Fe(III)reduction (Gorby and Lovley, 1991a). Electrons may be preferentiallydiverted to the nitrate and/or nitrite reductase in the presence of nitrate(DiChristina, 1992). However, an alternative explanation for the lack of netFe(III) reduction, at least in G. metallireducens, is that any Fe(II) that isproduced is rapidly reoxidized to Fe(III) with nitrate serving as the electronacceptor (Finneran et al., 2002c). Inhibition of net Fe(III) reduction withnitrate appears to require the expression of genes that are more highlyexpressed in the presence of nitrate, as nitrate inhibited net Fe(III) reductionin washed cell suspensions of nitrate-grown cells, but not Fe(III) grown cells(Finneran et al., 2002c). This may reflect the fact that components necessaryfor nitrate reduction are only expressed during growth on nitrate (Gorbyand Lovley, 1991a). More detailed evaluation of the influence of nitrate ongene expression in G. metallireducens will be possible now that wholegenome DNA microarrays for this organism are available.

4.1.6. Fumarate

A number of Fe(III)-reducing microorganisms are capable of usingfumarate as the sole electron acceptor to support growth (Table 1).Although fumarate is unlikely to be an abundant electron acceptor in mostsedimentary environments, fumarate is an excellent soluble electronacceptor for culturing some dissimilatory Fe(III) reducers in order togenerate biomass that has been grown under anaerobic conditions forbiochemical or other investigations that require substantial quantities ofcells. Fumarate respiration has been studied in G. sulfurreducens andShewanella species. In Shewanella species the fumarate reductase is a novelperiplasmic, flavocytochrome (Myers and Myers, 1997b). The fumaratereductase in G. sulfurreducens is an inner membrane protein withhigh homology to the fumarate reductase found in Wolinella succinogenes(Butler and Lovley, 2004). Knocking out frdA, predicted to encode forthe catalytic flavoprotein of the fumarate reductase, eliminated the ability ofG. sulfurreducens to use fumarate, but not Fe(III), as the sole electronacceptor. The respiratory fumarate reductase in G. sulfurreducens alsofunctions as the succinate dehydrogenase, the first time that the sameenzyme has been documented to function both in fumarate respiration andas a part of the TCA cycle.


Geobacter sulfurreducens has a three-fold higher growth yield withfumarate as the electron acceptor than with Fe(III) (Esteve-Nunez et al.,2004b). Yet when both Fe(III) and fumarate are provided to culturesgrowing in chemostats, Fe(III) is preferentially reduced (Esteve-Nunezet al., 2004a). This is associated with a substantial reduction in levels offrdA mRNA, suggesting that transcription of the fumarate reductasegenes is downregulated in the presence of Fe(III). Fe(II) has no impacton levels of the fumarate reductase mRNA. This is a rare instance inwhich Fe(III), but not Fe(II), regulates gene expression in microorganisms.A potential explanation for G. sulfurreducens regulating its respirationto favor Fe(III) reduction over fumarate reduction, even though fuma-rate reduction yields more energy, is that in most sedimentary environ-ments Fe(III) is abundant, but organic electron donors are limited.Thus, G. sulfurreducens regulates its respiration to utilize fumarate as anelectron donor rather than as an electron acceptor (Esteve-Nunez et al.,2004a).

4.1.7. Chlorinated Compounds

It is becoming increasingly apparent that a substantial number ofdissimilatory Fe(III)-reducing microorganisms can also conserve energy tosupport growth from electron transport to chlorinated compounds.For example, members of the Geobacteraceae, Desulfuromonas chloroenthe-nica (Krumholz et al., 1996; Krumholz, 1997) and Desulfuromonasmichiganensis (Sung et al., 2003) could reductively dechlorinate tetrachlor-oethene (PCE) to cis-1,2-dichloroethene (cis-DCE) with acetate as theelectron donor and another member of this family, ‘‘Trichlorobacterthiogenes’’, is also capable of dechlorination (De Wever et al., 2000). OtherFe(III) reducers capable of dechlorination include Anaeromyxobacterdehalogens (He and Sanford, 2003), a member of the Myxobacteriaand several Desulfitobacterium species (Lovley et al., 1998; Niggemyeret al., 2001; Finneran et al., 2002b; Shelobolina et al., 2003d). The capacityfor dechlorination may be more widespread amongst dissimilatoryFe(III)-reducing microorganisms than currently recognized because evalua-tion of the ability to reduce a range of chlorinated compounds has notbeen part of the characterization of many Fe(III) reducers. The presence ofFe(III) oxide does not appear to inhibit reductive dechlorination, suggestingthat Fe(III) reduction and dechlorination may take place simultaneouslyin contaminated subsurface environments (He and Sanford, 2003; Sunget al., 2003).


4.1.8. Electrodes

Another extracellular electron acceptor that at least some dissimilatoryFe(III)-reducing microorganisms can utilize are current-harvestinggraphite electrodes. Quantitative electron transfer from the oxidation oforganic compounds to electrodes was first noted in studies investigatingthe potential for harvesting electricity from marine sediments. When agraphite electrode was buried in anoxic marine sediments and electricallyconnected to a graphite electrode in the overlying, aerobic water, there wasa flow of electrons (Reimers et al., 2001). Further investigation of thisphenomenon revealed that the surface of the graphite electrodes buriedin the anoxic sediments was highly enriched in microorganisms inthe family Geobacteraceae (Bond et al., 2002; Tender et al., 2002; Holmeset al., 2003b). In marine sediments the Geobacteraceae on the electrodefell primarily in the Desulfuromonas cluster, whereas electrodes buriedin freshwater sediments were primarily colonized by Geobacter species.This reflects the general preference of Desulfuoromonas species formarine salinities and Geobacter species for freshwater. Pure cultures ofDesulfuromonas acetoxidans, G. sulfurreducens, and G. metallireducens, aswell as ‘‘Geopsychrobacter’’ sp., which were isolated from an electrode, werecapable of conserving energy to support growth by oxidizing acetateto carbon dioxide with an electrode serving as the sole electron acceptor(Bond et al., 2002; Bond and Lovley, 2003; Holmes et al., 2004). Thestoichiometry of electricity production and acetate consumption demon-strated that electron transfer to the electrode was quantitative. Geobactermetallireducens is also capable of oxidizing aromatic compounds in thismanner (Bond et al., 2002). Further studies with G. sulfurreducensdemonstrated that the microorganisms form a near mono-layer over theelectrode surface and it is these attached cells which account for the currentproduction (Bond and Lovley, 2003).In one marine deployment of sediment batteries, microorganisms

closely related to known Desulfobulbaceae species predominated on theelectrode instead of Geobacteraceae (Holmes et al., 2003b). Subsequentstudies demonstrated that like the previously studied Geobacteraceae, a pureculture of Desulfobulbus propionicus could oxidize organic compoundswith an electrode serving as the sole electron acceptor (Holmes et al., 2003a).It could also oxidize S� in this manner. This is significant because sulfide,which is found in high concentrations in some marine sediments canabiotically react with electrodes to form S� (Tender et al., 2002), whichmight then serve as an electron donor for organisms like D. propionicus onthe electrode (Holmes et al., 2003a).


In fact, the ability to transfer electrons onto an electrode surface may bea general characteristic of dissimilatory Fe(III)-reducing microorganisms,possibly because the mechanisms for transferring electrons onto insolu-ble, extracellular Fe(III) oxides might also permit electron transfer toextracellular, insoluble electrodes. For example, Shewanella oneidensiscan transfer electrons to electrodes (Kim et al., 2002), but it is not yetclear how efficient this process is or whether S. oneidensis can conserveenergy to support growth from this electron transfer. Geothrix fermen-tans (D. Bond, unpublished data) and Rhodoferax ferrireducens (Chaudhuriand Lovley, 2003) can conserve energy to support growth from electrontransfer to electrodes with an apparent direct, highly efficient electrontransfer similar to that observed in Geobacteraceae and Desulfobulbaceaespecies.The electron transfer to electrodes carried out by Fe(III)-reducing

microorganisms has several potential advantages over previouslydescribed microorganisms that have been evaluated for the developmentof microbial fuel cells (Chaudhuri and Lovley, 2003). One majordifference is that microorganisms that are not Fe(III) reducers typicallyrequire the presence of a mediator which can facilitate electron transferbetween the cell and the electrode. In contrast, the Fe(III) reducers that havebeen examined in detail have the ability to directly transfer electrons tothe electrode surface without the need for an electron-shuttlingmediator. Furthermore, the Geobacteraceae that have been evaluated, aswell as R. ferrireducens and G. fermentans, transfer more than 80% of theelectrons available in their organic substrates to electricity. In contrast,previous prototype microbial fuel cells have employed microorganismswith a fermentative metabolism that allows transfer of less that 10% ofthe electrons available in the organic substrate, even in the presence ofmediators. Therefore, most of the electrons remain in organic metabolicend products. The higher efficiency of Fe(III)-reducing microorgan-isms and the lack of a requirement for mediator compounds suggeststhat they may be the organisms of choice for engineering systems forharvesting electricity from waste organic matter and renewable biomass.

4.2. Electron Donors

4.2.1. Organic Compounds and Hydrogen

Most Fe(III) reducers that conserve energy to support growth fromFe(III) reduction are restricted by the organic electron donors that they


can utilize. The most common electron donors are organic acids. Forexample, as noted above, a common physiological feature of Geobacterspecies is their ability to grow with acetate as the sole electron donor. Thisis significant because acetate is likely to be the most important electrondonor for Fe(III) reduction in many sedimentary environments.Experimental evidence (Champine and Goodwin, 1991; Mikoulinskais

et al., 1999; Galushko and Schink, 2000) and metabolic modeling fromgenome sequences (Radhakrishnan et al., 2004) indicate that Geobacterspecies oxidize acetate via the TCA cycle. A unique aspect of the GeobacterTCA cycle is the citrate synthase gene. The citrate synthase genes in allof the Geobacteraceae that have been evaluated are most closely related tocitrate synthase genes found in eukaryotes (Bond et al., 2003; Methe, 2003).This is an apparent rare instance of lateral gene transfer from a eukaryote toa prokaryote (Bond et al., 2003). The citrate synthase in G. sulfurreducens isa dimer, like those found in eukaryotes rather than a hexamer as found inprokaryotes (Mester et al., 2003).A recent surprise was the finding that several hyperthermophilic

Fe(III)-reducing microorganisms are capable of acetate oxidation (Toret al., 2001). Prior to this study, there was no verifiable proof thathyperthermophiles of any kind could oxidize acetate, leading to theconcept that acetate, produced from fermentation or abiotic processes inhot (i.e. >80�C) environments would have to diffuse into cooler zonesprior to being metabolized (Slobodkin et al., 1999c). However, two hyper-thermophiles, Geoglobus ahangari and Ferroglobus placidus, were foundto conserve energy to support growth from the oxidation of acetate tocarbon dioxide coupled to the reduction of Fe(III), indicating that acetatemay be anaerobically oxidized in hot environments, at least those thatcontain Fe(III).Hydrogen was the first electron donor found to support the growth of

dissimilatory Fe(III)-reducing microorganisms (Balashova and Zavarzin,1980). Many mesophilic Fe(III)-reducing microorganisms includingsome of the known Geobacter species and most, if not all, Shewanellaspecies can oxidize hydrogen with the reduction of Fe(III) (Table 1).As noted above, the capacity for hydrogen oxidation coupled to Fe(III)reduction is highly conserved among hyperthermophilic Archaea andBacteria and some are only known to grow via this form of respiration.Two operons in the G. sulfurreducens genome appeared to code for

periplasmic respiratory hydrogenases (Coppi et al., 2003). Mutationalanalysis has indicated that one of these, Hyb, is required for hydrogen-dependent reduction of Fe(III), AQDS, and fumarate. The role of theHya, is not yet apparent because deletion of hya had no discernable


impact on the hydrogen-dependent reduction of these electron acceptors(Coppi et al., 2003).A few Fe(III)-reducing microorganisms available in pure culture can

oxidize aromatic compounds (Table 1). Geobacter metallireducens wasthe first organism of any kind found to oxidize an aromatic hydrocarbonin the absence of oxygen (Lovley et al., 1989a; Lovley and Lonergan,1990). This metabolism serves as a model for the oxidation of aromaticcontaminants coupled to the reduction of Fe(III) in subsurface environ-ments, which can be an important process for the removal of thesecontaminants from polluted groundwater (Anderson and Lovley, 1997;Anderson et al., 1998; Lovley and Anderson, 2000). Analysis of thecomplete genome sequence of G. metallireducens indicates that thepathway for the anaerobic degradation of aromatic compounds islikely to be similar to those previously described in the nitrate-reducer,Thauera aromatica.The ability of anaerobes to metabolize sugars with the reduction of

Fe(III) has been known since the early investigations into microbial Fe(III)reduction (see for example, Roberts (1947)). However, as previouslyreviewed in detail (Lovley, 1987), microorganisms in those early studies hada primarily fermentative metabolism and only reduced Fe(III) as a sidereaction. However, more recently, microorganisms that can conserve energyto support growth from the oxidation of sugars coupled to the reductionof Fe(III) have been described. In some instances metabolism is onlyincomplete to acetate (Coates et al., 1998a), but in others the sugars arecompletely oxidized to carbon dioxide with Fe(III) serving as the soleelectron acceptor (Kusel et al., 1999; Chaudhuri and Lovley, 2003).A number of Fe(III) reducers have been grown with peptides and/orindividual amino acids as the electron donor for Fe(III) reduction(Table 1), but with a few exceptions (Caccavo et al., 1996) it has not beendetermined whether the amino acids were completely oxidized to carbondioxide with Fe(III) serving as the sole electron acceptor.

4.2.2. Fe(II), Hydroquinones, and Electrodes

Under the appropriate conditions, reduced end products of the respirationof Fe(III)-reducing microorganisms can serve as electron donors. Forexample, Acidithiobacillus ferrooxidans, which is well known for its abilityto grow as an Fe(II) oxidizer at acidic pH, is also a Fe(III) reducer(Pronk et al., 1991; Das et al., 1992; Ohmura et al., 2002) as are anumber of other acidophilic Fe(II) oxidizers (Blake II and Johnson, 2000).


The hyperthermophilic Fe(III) reducer Ferroglobus placidus (Tor andLovley, 2001) was first isolated as a Fe(II)-oxidizing nitrate reducer(Hafenbradl et al., 1996). The Fe(III) reducer Desulfitobacterium frappierican utilize not only soluble Fe(II), but the structural Fe(II) in clay as anelectron donor for nitrate reduction (Shelobolina et al., 2003d). Fe(II) mayserve as an electron donor for the reduction of nitrate in G. metallireducens,but it was not determined whether this reaction yields energy to support cellgrowth (Finneran et al., 2002c). Yet to be determined in any of theseorganisms is to what degree the pathways for Fe(II) oxidation and Fe(III)reduction share similar components.A wide phylogenetic diversity of Fe(III)-reducing microorganisms can

oxidize reduced humic substances and/or the reduced humic substancesanalog, anthrahydroquinone-2,6-disulfonate (AHQDS) with nitrate and/orfumarate as the electron acceptor (Lovley et al., 1999). Those that wereevaluated in detail, including Geobacter and Shewanella species, werecapable of conserving energy to support growth from the oxidation ofAHQDS. The hydroquinone moieties in the reduced humic substances andAHQDS were shown to be the electron donor.Electrodes can serve as an electron donor for the reduction of nitrate

and fumarate by Geobacter species when the electrodes are maintainedat a sufficiently negative potential. When a graphite electrode poisedat the appropriate potential was placed in a nitrate-containingmedium inoculated with sediment, the nitrate was reduced over time(Gregory et al., 2003). The surface of the electrode was heavily colonizedwith microorganisms with 16SrRNA gene sequences closely relatedto those of known Geobacter species. Cultures of G. metallireducensand G. sulfurreducens attached to electrode surfaces could reducenitrate or fumarate respectively, with the electrode serving as the soleelectron donor.

4.3. Temperature Range, pH, and Salinity Ranges

Pure cultures of Fe(III)-reducing microorganisms have been reported thatcan grow at temperatures as low as 4�C (Finneran et al., 2003; Holmes,2004d) and, as noted above, an Fe(III) reducer currently holds the recordfor highest temperature (121�C) known to support the growth of a pureculture (Kashefi and Lovley, 2003). Several Fe(III) reducers, suchas A. ferrooxidans (Ohmura et al., 2002) and Acidiphilium cryptum (Kuselet al., 1999) grow at low pH at which Fe(III) is primarily available as asoluble ion. An arsenate-reducing Bacillus species was capable of Fe(III)


reduction at pH 9 (Blum et al., 1998). Fe(III) reduction has been noted inaquifer sediments with a salinity 10-fold higher than seawater (Nevin et al.,2003a). Analysis of 16S rRNA gene sequences indicated that thepredominant organisms during Fe(III) reduction were Pseudomonas orDesulfosporsinus species, but the high-salt tolerant organisms have not yetbeen recovered in culture.

4.4. Nitrogen Fixation and Autotrophy

A number of hyperthermophilic Fe(III)-reducing microorganisms arecapable of growing in the absence of organic carbon with hydrogenas the electron donor (Kashefi et al., 2002a, 2002b; Kashefi and Lovley,2003). This may be an adaptation to growth near hydrothermal fluidswhich may be high in hydrogen, but have limited organic content.Examination of over 30 Geobacteraceae available in pure culture

demonstrated that they all contained genes for nitrogen fixation (Holmeset al., 2004) and those cultures that have been examined furtherhave had the ability to grow in the absence of fixed nitrogen (Bazylinskiet al., 2000; Coppi et al., 2001). The ability to fix nitrogen may be acompetitive advantage in nutrient-poor subsurface environments. Forexample, petroleum contamination of sandy aquifers may providesignificant organic carbon to support microbial Fe(III) reduction, butlittle fixed nitrogen (Bazylinski et al., 2000). This hypothesis was supportedby the fact that genes for nitrogen fixation (nifD) were expressed byGeobacteraceae in Fe(III)-reducing petroleum contaminated sediments(Holmes et al., 2004e).

5. MECHANISMS FOR Fe(III) AND Mn(IV) REDUCTION

Unlike commonly considered electron acceptors such as oxygen, nitrate,sulfate, or carbon dioxide, Fe(III) and Mn(IV) are highly insoluble in mostenvironments at circumneutral pH. Soluble electron acceptors can diffuseinto cells in order to be reduced whereas Fe(III) and Mn(IV) reducersface the challenge of how to transfer electrons onto an insoluble,extracellular, electron acceptor. This is also a challenge for investigatorsof this process as working with insoluble electron acceptors raise a numberof technical difficulties.


The study of Fe(III) and Mn(IV) oxide reduction is further complicatedby the many types of Fe(III) and Mn(IV) oxides that can be found in soilsand sediments. There are no generally accepted standards for which Fe(III)oxides are the most suitable for such studies. If the goal is to elucidate themechanisms involved in Fe(III) reduction in soils and sediments atcircumneutral pH, then studies with poorly crystalline Fe(III) oxides willprobably be more informative than those with highly crystalline Fe(III)forms because, as noted above, the poorly crystalline forms are the primarysource of Fe(III) oxides available for microbial reduction in theseenvironments.In evaluating studies on mechanisms for Fe(III) reduction, it is important

to note that many of these have focused on electron transport tosoluble, chelated Fe(III). Chelated Fe(III) may enter the periplasm priorto reduction (Dobbin et al., 1995, 1996), and thus be reduced by electrontransfer components that are not directly involved in the reductionof Fe(III) oxides. Alternatively, soluble Fe(III) may be adsorbed to thecells prior to reduction (Haas and DiChristina, 2002). If this is the case,reduction of soluble and insoluble Fe(III) may follow more similarmechanisms.

5.1. Strategies for Fe(III) Oxide Reduction – Direct Contact VersusElectron Shuttling or Chelation

The discovery that exogenous Fe(III) chelators and electron shuttles couldgreatly stimulate Fe(III) oxide reduction, led to the question of whetherFe(III) reducers might, themselves, release Fe(III) chelators or electronshuttles. Until recently it was generally considered that microorganismsreducing insoluble Fe(III) oxides had to directly contact the oxides inorder to reduce them. This concept was evident in the pioneering studies ofOttow and co-workers. Fe(III) reducers which were separated from Fe(III)oxides with a semi-permeable membrane failed to reduce the Fe(III)(Munch and Ottow, 1983). Subsequent studies with a diversity ofFe(III) reducers had similar results (Tugel et al., 1986; Arnold et al., 1988;Lovley and Phillips, 1988b; Lovley et al., 1991b; Caccavo et al., 1992).These results were interpreted as evidence that Fe(III)-reducing micro-organisms did not release chelators that could solubilise Fe(III) orelectron shuttles that could carry electrons from the cell surface to thesurface of the Fe(III) oxide. This was because it was assumed that thesechelators or shuttles would be able to diffuse through semi-permeablemembranes and release soluble Fe(III) or permit electron transfer across


the membranes. However, the validity of these early studies waschallenged when it was found that the appropriate positive controls hadnot been conducted in any of these previous studies and that an electronshuttling compound or an Fe(III) chelator did not freely transfer electronsor Fe(III) between G. metallireducens and Fe(III) retained within a dialysismembrane with the largest pore size available (Nevin and Lovley, 2000a).Subsequent studies have demonstrated that some, but not all,

dissimilatory Fe(III)-reducing microorganisms do release electron shuttlingcompounds and Fe(III) chelators to promote Fe(III) oxide reduction.For example, it was observed that S. oneidensis released a diffusiblecompound, thought to be a quinone, that would rescue the growth of aS. oneidensis mutant deficient in menaquinone biosynthesis and it wassuggested that this same compound might provide an electron shuttlefor Fe(III) oxide reduction (Newman and Kolter, 2000). Subsequentstudies with the closely related Shewanella algae demonstrated that itcould in fact reduce Fe(III) oxides that it could not directly contact(Nevin and Lovley, 2002b). The Fe(III) oxides were sequestered withinmicroporous alginate beads designed to permit the entry of molecules up toca. 12 kDa, but exclude direct contact between the cells and the Fe(III)oxide. Unlike the semi-permeable membranes described above, the alginatebeads allow electron shuttling compounds to enter, contact andreduce Fe(III) oxides, and then exit the bead to be reduced again by thecells (Nevin and Lovley, 2000a, 2002b). The Fe(II) produced from reduc-tion of Fe(III) remained in the beads, indicating that the Fe(III) hadbeen reduced within the beads. The compound(s) responsible for electronshuttling in those studies has yet to be identified. However, in other studiesS. algae was found to produce melanin which could serve as a solubleelectron shuttle to promote Fe(III) reduction (Turick et al., 2002). Melaninassociated with the cell surface could also promote Fe(III) oxide reduction(Turick et al., 2003). These studies clearly demonstrate that S. algae can useelectron shuttling to effectively reduce Fe(III) oxides that it is not directlycontacting.Further evidence consistent with Shewanella species reducing Fe(III) via

a soluble electron shuttle is the observation that Fe(III) oxides can bereduced at locations that are at significant distances from where the cellsare attached (Rosso et al., 2003). In the absence of electron shuttling,Fe(III) reduction would be expected to be localised at the point of cellattachment (Rosso et al., 2003). Furthermore, an adhesion-deficient strainof S. algae reduced Fe(III) oxide as well as wild type despite the fact thatthe number of cells attached to Fe(III) oxide in this strain was less thanhalf that of wild type (Caccavo et al., 1997).


In addition to using an electron shuttle(s) to alleviate the need for cell-Fe(III) oxide contact, S. algae also solubilised Fe(III) from Fe(III) oxide(Nevin and Lovley, 2002b). The Fe(III) chelator(s) have not been identi-fied, but cultures of S. algae growing on Fe(III) oxide contained as muchas 450 mM dissolved Fe(III) (Nevin and Lovley, 2002b). This contrasts,with soluble Fe(III) concentrations of less than 3 mM in uninoculatedmedia or in cultures of other Fe(III)-reducing microorganisms that donot solubilise Fe(III) during Fe(III) oxide reduction (Nevin and Lovley,2000a).Shewanella algae is not the only organism with strategies that allow

growth on insoluble Fe(III) oxide without direct contact. Cell-free filtratesof Fe(III) oxide-grown cultures of G. fermentans greatly stimulated Fe(III)oxide reduction in washed cell suspensions of G. fermentans (Nevin andLovley, 2002a), as was seen with S. algae (Nevin and Lovley, 2002b).Furthermore, as with S. alga, G. fermentens could reduce Fe(III) oxidesthat it could not contact, sequestered in microporous alginate beads.Culture filtrates contained an electron shuttling capacity equivalent to ca.25 mM quinone (Nevin and Lovley, 2002a) and analysis of the filtrateswith thin layer chromatography suggested that the electron shuttle hadcharacteristics similar to a water-soluble quinone.It was initially proposed that G. sulfurreducens might also reduce Fe(III)

via electron shuttling (Seeliger et al., 1998). The hypothesized mechanismfor electron shuttling was not via quinones, as proposed for Shewanellaand Geothrix species, but rather via release of a small (9.6 kDa) c-typecytochrome (Seeliger et al., 1998). However, further studies demonstratedthat, G. sulfurreducens did not in fact release this cytochrome and that evenif exogenous cytochrome was added to cultures, it did not function as anelectron shuttle (Lloyd et al., 1999). The original proposers of this modelfor electron shuttling by G. sulfurreducens have subsequently rescindedthis hypothesis (Straub and Schink, 2003).Not only does G. sulfurreducens not produce a soluble cytochrome

electron shuttle, but further evaluation of the mechanisms involved inFe(III) oxide reduction in Geobacter species have suggested that it doesnot produce any electron shuttles or Fe(III) chelators. In contrast toShewanella and Geothrix species, G. metallireducens was unable to reduceFe(III) oxide sequestered in microporous alginate beads (Nevin and Lovley,2000a) or agar (Straub and Schink, 2003). However, when the artificialelectron shuttle, AQDS was added, the Fe(III) within the beads was readilyreduced. Furthermore, Fe(III) was not solubilised by G. metallireducensduring growth on Fe(III) oxide (Nevin and Lovley, 2000a). These resultsindicate that G. metallireducens has to directly contact Fe(III) oxides in


order to reduce them. As detailed in the next section, further studies withG. metallireducens and G. sulfurreducens have demonstrated that Geobacterspecies have highly regulated special adaptations to permit them to accessinsoluble Fe(III) oxides in order to reduce them.The finding that phylogenetically distinct Fe(III)-reducing microorgan-

isms have different mechanisms for growing on insoluble Fe(III) oxideshas important implications for extrapolating from pure culture results tomechanisms of Fe(III) oxide reduction in soils and sediments. It can nolonger be assumed that studies with any Fe(III) reducer that canconveniently be cultured are appropriate. For example, given theirsignificantly different strategies for Fe(III) oxide reduction it would beinappropriate to apply results from studies with S. algae or G. fermentansto the many subsurface environments in which Geobacter species are thepredominant Fe(III)-reducing microorganisms.The different physiological approaches to Fe(III) oxide reduction

may also have a significant impact on which environmental nichea microorganism might inhabit. In a typical subsurface environmentwhere the population densities of Fe(III)-reducing microorganisms are notexpected to be high (Holmes et al., 2002), it may not be beneficial to releasean electron shuttle and/or chelator. Such a compound has to recycled manytimes to recoup the energetic cost of biosynthesis, yet the released com-pound is unlikely to return to the cell, but is more likely to be lost viadiffusion and advection (Childers et al., 2002; Nevin and Lovley, 2002b).In contrast, in environments with high densities of Fe(III)-reducingmicroorganisms, the members of the community might mutually benefitfrom the release of an electron shuttle or a chelator (Nevin and Lovley,2002b). Such environments have yet to be documented, but the surfaceof corroding steel pipelines where Shewanella species may congregate(Westlake et al., 1986; Semple et al., 1989) is a potential example(Hernandez and Newman, 2001; Nevin and Lovley, 2002b).

5.2. Models for Electron Transfer to Extracellular Fe(III) andMn(IV) Oxides

Whether dissimilatory Fe(III)-reducing microorganisms are reducingsoluble Fe(III), electron-shuttling quinones, or insoluble Fe(III) oxides,they need to transfer electrons derived from central metabolism to asite of reduction somewhere outside the inner membrane. Althoughcytoplasmic proteins which will reduce Fe(III) in vitro can be found indissimilatory Fe(III) reducers, as detailed below, reduction of even soluble


Fe(III) is likely to take place either in the periplasm or at the surface ofthe outer membrane. Just as phylogenetically distinct Fe(III) reducersappear to have different strategies for transferring electrons onto Fe(III)oxides, the proteins involved in electron transfer to the periplasm and theouter membrane in these different organisms are similar in function, but notclosely related in those Fe(III) reducers that have been most intensivelystudied. This further suggests that strategies for dissimilatory Fe(III)reduction have evolved independently several times.Electron transport proteins involved in Fe(III) and Mn(IV) reduction

have been studied most intensively in Shewanella and Geobacter species.In both cases it is generally considered that electron transfer proteins orquinones in the inner membrane transfer electrons to electron transferproteins, primarily c-type cytochromes, in the periplasm, and then on toother c-type cytochromes in the outer membrane (Figs. 2 and 3). One ormore of these c-type cytochromes in the outer membrane are then involvedin electron transfer directly to Fe(III) or Mn(IV) or to soluble electronshuttling compounds. Studies to date have identified some of thecomponents involved in this electron transfer process.

Figure 2 Summary of components suggested to be involved in electron transfer toFe(III) and Mn(IV) oxides in Shewanella species.


When evaluating the role of various cell components in electron transportto Fe(III) it is important to recognize that many redox proteins andquinones are capable of reducing Fe(III) in vitro. For example, it hasbeen emphasized that most of the proteins which have been investigated asassimilatory Fe(III) reductases are actually flavin reductases and thatthe flavins that have been included in the assays for these assimilatoryFe(III) reductases can reduce Fe(III) (Fontecave et al., 1994). In asimilar manner, there are soluble proteins in the cytoplasm of dissimilatoryFe(III) reducers that can reduce Fe(III) in vitro (Childers and Lovley, 2001;Kaufmann and Lovley, 2001) but it is unlikely that they are involved indissimilatory Fe(III) reduction in vivo.Even when proteins are located in the periplasm or outer membrane

where they could conceivably have access to Fe(III), care must be taken inassigning the role of a Fe(III) reductase to such proteins based solelyupon their ability to reduce Fe(III) in vitro. For example, most, if not all, ofthe c-type cytochromes in Shewanella and Geobacter species willreduce Fe(III) in vitro, yet it is unlikely that they are all terminal Fe(III)reductases. Thus, more detailed evaluations of in vivo function are

Figure 3 Summary of components suggested to be involved in electron transfer toFe(III) and Mn(IV) oxides in Geobacter species.


required. To date, several important components in electron transfer toFe(III) and Mn(IV) reduction have been identified in Shewanella andGeobacter species.

5.2.1. Shewanella Species

As emphasised above, the principal mechanism for electron transferto Fe(III) oxide in Shewanella species may not be direct electron transferfrom the cell to the Fe(III) oxide surface. Rather, chelators releasedfrom the cells may first solubilize Fe(III) and/or electron shuttles that thecells produce may be the actual electron carriers to Fe(III) oxide.In any event, most studies on the biochemical mechanisms for Fe(III)reduction in Shewanella have focused on the reduction of soluble,chelated Fe(III).Shewanella species appear to be specially adapted for the reduction

of extracellular electron acceptors. Most notable is the specific localizationof c-type cytochromes to the outer membrane in anaerobically growncells (Myers and Myers, 1992, 1997c) and localization of Fe(III) reductaseactivity in the membrane fraction (Myers and Myers, 1993a; Dobbin et al.,1995). Knocking out a gene in a type II secretory system significantlydiminished the capacity for Fe(III) and Mn(IV) reduction withoutimpacting the rate of reduction of soluble electron acceptors (DiChristinaet al., 2002). This was associated with failure of the mutant to localizea 91 kDa heme-staining protein with Fe(III) reductase activity to theouter membrane. It is possible that other outer-membrane proteins alsorequired for the reduction of Fe(III) and Mn(IV) were also not properlylocalized. Differences in the attractive force between Shewanella and Fe(III)oxide under aerobic and anaerobic conditions have been suggested toprovide evidence for a Fe(III) reductase in the outer membrane (Loweret al., 2001).As outlined below, there have been detailed genetic investigations into

specific proteins involved in Fe(III) and Mn(IV) reduction in Shewanellaspecies. However, in many of the mutational studies it is difficult to assessthe true role of some of the proteins in Fe(III) or Mn(IV) oxide reduc-tion because the cells were not grown in an anaerobic defined medium withFe(III) or Mn(IV) oxide as the electron acceptor. In many instances, growthof mutants under investigation on Fe(III) oxide has not been evaluated orreported (Beliaev and Saffarini, 1998; Gordon et al., 2000). Alternatively,cells have been pregrown to high densities aerobically in an organic-richmedium and then used as an ‘‘innoculum’’ to evaluate the potential for


Mn(IV) and Fe(III) reduction (Myers and Myers (2002b) and referencestherein). This approach also does not evaluate the effect of the mutationson growth on Fe(III) or Mn(IV) oxide and often the amounts of Fe(III) orMn(IV) oxide reduced, even in the wild type, are low. One reason for thisapproach may be due to the fact that growing some strains of Shewanella onFe(III) oxide is very difficult, especially in the absence of a Fe(III) chelatoror electron shuttle (Nevin and Lovley, 2002b). However, better approachesare required because growth of cells on alternative electron acceptors maysignificantly influence the biochemical composition and Fe(III) reductioncapacity of Shewanella species (Blakeney et al., 2000).Components that have been identified to be important in electron

transfer to Fe(III) and/or Mn(IV) in Shewanella species are summarizedbelow.

5.2.1.1. CymA. CymA is a tetra-heme, 21 kDa c-type cytochromeassociated with the inner membrane and periplasmic fraction of S.oneidensis which is required for the reduction of Fe(III), fumarate, nitrate,nitrite, and DMSO (Myers and Myers, 1997a, 2000). Based upon itslocation, it is assumed to be involved in transfer of electrons from theinner membrane to electron carriers or acceptors in the periplasm (Schwalbet al., 2003). The source of the electrons for CymA may be menaquinonesthat are required for electron transfer to Fe(III) and Mn(IV) (Myersand Myers, 1993b). A similar cytochrome is found in S. frigidimarina(Field et al., 2000).

5.2.1.2. MtrA. MtrA is a 32 kDa decaheme c-type cytochromepredicted to be located in the periplasm (Beliaev et al., 2001; Pitts et al.,2003). This gene is part of an operon which includes mtrA, mtrB, and mtrC(Beliaev et al., 2001). It has been suggested that MtrA is required for Fe(III)reduction, but proper complementation studies have yet to be conducted toconfirm this. It is possible that MtrA accepts electrons from CymA andtransfers these electrons to an acceptor in the outer membrane (Beliaev et al.,2001). Alternatively, it may directly reduce soluble Fe(III) that enters theperiplasm because the addition of chelated Fe(III) forms results in theoxidation of MtrA expressed in E. coli (Pitts et al., 2003).

5.2.1.3. IfcA. IfcA is a 63.9 tetraheme, periplasmic flavocytochromeexpressed in cultures of S. frigidimarina during growth on soluble Fe(III),but not other soluble electron acceptors (Dobbin et al., 1999a). This wasassociated with a substantially higher potential for Fe(III) reduction incells grown on Fe(III). Expression during growth on insoluble Fe(III) orMn(IV) oxides was not evaluated. The cytochrome contains non-covalently


bound FAD and is capable of catalyzing fumarate reduction. Deletion ofifcA did not have an impact on soluble Fe(III) reduction, possibly becauseother periplasmic cytochromes were produced in higher levels in the mutant(Dobbin et al., 1999a).

5.2.1.4. Cytochrome c3. Another periplasmic tetraheme c-type cyto-chrome that appears to be involved in reduction of soluble Fe(III) inS. frigidimarina has been referred to as cytochrome c3 (Gordon et al.,2000). Knocking out the gene for this cytochrome reduced the capacity forthe reduction of Fe(III) citrate without affecting the reduction of othersoluble electron acceptors (Gordon et al., 2000). A similar cytochrome isfound in S. oneidensis and structural analysis has indicated that electrontransfer partners may interact with any of the hemes in this cytochrome,suggesting that it optimized efficient intermolecular electron transfer inserving as an electron shuttle in the periplasm (Leys et al., 2002).

5.2.1.5. MtrB. MtrB is predicted to be a ca. 76 kDa outer membraneprotein (Beliaev and Saffarini, 1998) and is required for reduction ofFe(III) (Beliaev and Saffarini, 1998) as well as the humic acid analog, AQDS(Shyu et al., 2002), in S. oneidensis. Due to an apparent metal-bindingmotif, it was initially proposed to function in binding Fe(III) prior toreduction (Beliaev and Saffarini, 1998). However, subsequent studieshave demonstrated that MtrB is required for proper localization ofOmcA and OmcB, and possibly other cytochromes, to the outer membrane(Myers and Myers, 2002a). It has been suggested that cytochromelocalization, rather than metal binding is the true function of MtrB(Myers and Myers, 2002a).

5.2.1.6. OmcA. OmcA is a 83 kDa decaheme c-type cytochrome locatedin the outer membrane of Shewanella species (Myers and Myers, 1998).Protease and immunofluorescent localization studies have indicated thatOmcA is exposed on the outer surface of S. oneidensis (Myers and Myers,2003a). When omcA was deleted in S. onedensis, there was no impacton the reduction of a variety of soluble electron acceptors (Myers andMyers, 2001). However, the mutant reduced Mn(IV) oxide at a rate45% lower than the wild type. Overexpressing omcB could compensate forthe lower rate of Mn(IV) reduction in the omcA mutant, suggesting thateither omcA or omcB can independently fulfill similar roles in Mn(IV)reduction (Myers and Myers, 2003b). The omcA mutant reduced Fe(III)oxide as well as the wild type (Myers and Myers, 2001). This result mightnot have been expected since omcA is the most abundant cytochrome inthe outer member of S. onedensis (Myers and Myers, 2001), and given that


Fe(III) oxide is typically a more abundant electron acceptor than Mn(IV)oxide, it might have been assumed that the most abundant c-typecytochrome involved in metal oxide reduction would have a role in Fe(III)oxide reduction. However, the capacity for Fe(III) reduction was evaluatedwith cells that had been grown aerobically in an organic rich medium andthus rates and mechanisms for Fe(III) oxide reduction may have beensignificantly different than if the cells had been grown with Fe(III) oxide asthe sole electron acceptor.There does appear to be controversy over the role of OmcA in Mn(IV)

reduction as a subsequent study reported that insertional inactivationof omcA had no impact on Mn(IV) reduction (Beliaev et al., 2001).Clearly, more investigation into the role of this abundant cytochrome iswarranted.

5.2.1.7. OmcB. OmcB (previously known as mtrC (Beliaev et al., 2001))is also an outer membrane decaheme c-type cytochrome (Myers and Myers,2001, 2003b). Like OmcA, OmcB appears to be involved in electrontransfer to Mn(IV). It also appears to be exposed on the outer surface, butpossibly to a lesser degree than OmcA (Myers and Myers, 2003a). Itsrole in electron transfer to Fe(III) is as yet uncertain, as it has beenimplicated in Fe(III) reduction in some studies (Beliaev et al., 2001), but thelatest studies suggest it is not required for Fe(III) reduction (Myers andMyers, 2003b).

5.2.1.8. Summary of electron transfer to extracellular electron acceptors inShewanella species. The available evidence is beginning to provide insightinto mechanisms by which Shewanella species may transfer electronsfrom the inner membrane to reductases in the outer membrane (Fig. 2). Themodel presented here is consistent with the suggested function of theseelectron transport components in one or more publications, but this modelcannot accommodate still existing differences in the proposed functionof some of these proteins. The genome of S. oneidensis (Heidelberg et al.,2002) suggests that there are genes for other electron transport componentswhose function have yet to be evaluated and further study of these is likelyto complete this developing picture.

5.2.2. Geobacter Species

Early biochemical studies on the mechanisms for electron transportto Fe(III) in Geobacter species suggested that cytochromes and otherredox-active proteins were involved in this process (Gorby and Lovley,


1991a; Lovley et al., 1993a; Gaspard et al., 1998; Magnuson et al., 2000;Kaufmann and Lovley, 2001; Magnuson et al., 2001), but the in vivofunction of these various proteins was impossible to verify until the recentdevelopment of a genetic system for G. sulfurreducens (Coppi et al., 2001).Components for which there is strong evidence for involvement in Fe(III)reduction are discussed below.

5.2.2.1. OmcB. OmcB is a 12-heme, outer-membrane, c-type cyto-chrome with an estimated molecular weight of ca. 87 kDa (Leang et al.,2003). It is now recognized that OmcB and the highly similar, OmcC wereboth probably present in the c-type cytochrome preparation previouslyreferred to as ‘‘FerA’’ (Magnuson et al., 2001). OmcB and OmcC arepredicted to have such similar size and charge properties (Leang et al., 2003)that they would not have been separated in the purification procedureused to isolate ‘‘FerA’’ (Magnuson et al., 2001). The ‘‘FerA’’ fraction wasfound to be a component of a NADH-dependent Fe(III) reductase com-plex purified from the membrane fraction of G. sulfurreducens (Magnusonet al., 2000) and the cytochrome(s) in the ‘‘FerA’’ fraction could transferelectrons to Fe(III) (Magnuson et al., 2001). These studies suggested that‘‘FerA’’ might be a terminal Fe(III) reductase in G. sulfurreducens.When omcB was deleted from G. sulfurreducens, it could no longer grow

with Fe(III) as the electron acceptor (Leang et al., 2003). Reduction ofsoluble as well as insoluble Fe(III) oxide was inhibited, whereas themutation had no impact on fumarate reduction. Deletion of omcC didnot affect either Fe(III) or fumarate reduction. Further evidence that omcBis involved in Fe(III) reduction was the finding that, in chemostat culturesin which growth rates could be reproducibly controlled, levels of mRNAfor omcB were substantially higher in cells grown with Fe(III) as theelectron acceptor than in fumarate-grown cells (Chin et al., 2004). As ratesof Fe(III) reduction were increased in the chemostats, there was a direct,proportional increase in levels of omcB mRNA. In contrast, expression ofomcC was upregulated during growth on fumarate in comparison withgrowth on Fe(III) (Chin et al., 2004).The finding that two c-type cytochromes that are 73% similar in their

predicted amino acid sequence have such different functions demonstratesthe potential pitfalls in attempting to predict gene function from sequencedata alone. The omcB and omcC genes appear to be components of a sharedgene duplication event with the two genes now evolving significantlydifferent roles (Leang et al., 2003).The outer membrane location of OmcB, and the finding that the

cytochrome fraction which presumably contained OmcB could transfer


electrons to Fe(III), suggest that omcB could be a terminal Fe(III)reductase in G. sulfurreducens (Leang et al., 2003). However, it has not yetbeen determined whether OmcB has active sites exposed on the outsideof the cell which could access Fe(III) in vivo. Thus, an alternative rolefor omcB that cannot yet be ruled out is that it is an intermediaryelectron carrier that transfers electrons to another, as yet unidentifiedFe(III) reductase (Leang et al., 2003). In accordance with the suggesteddifferent mechanisms for Fe(III) reduction in Shewanella and Geobacterspecies, no homologs of omcB could be found in the S. oneidensis genome.

5.2.2.2. PpcA. Ppc A is a tri-heme, 9.6 kDa c-type cytochrome that waspurified from the periplasm of G. sulfurreducens (Lloyd et al., 2003). It issimilar in sequence to the 9.1 kDa cytochrome of another member of theGeobacteraceae, Desulfuromonas acetoxidans (Correia et al., 2002), and a9.7 kDa c-type cytochrome in G. metallireducens (Afkar and Fukumori,1999; Champine et al., 2000; Schwalb et al., 2003). As discussed above, itwas originally proposed that G. sulfurreducens released PpcA as a solubleelectron shuttle to promote Fe(III) oxide reduction (Seeliger et al., 1998).However, it was later demonstrated that this was incorrect as PpcA wasnot released into the medium and PpcA added to cultures did not serve asan effective electron shuttle (Lloyd et al., 1999).When ppcA was deleted from G. sulfurreducens it grew much slower

in acetate-Fe(III) medium than the wild type (Lloyd et al., 2003). Themutation had no impact on growth with fumarate as the electronacceptor. Cell suspensions of fumarate-grown cells had rates of Fe(III)reduction with acetate as the electron donor that were 60% of wild type.Rates of reduction of U(VI) and the humic substances analog, AQDS, wereonly 20% and 5% of wild type, respectively. In contrast, rates of hydrogen-dependent reduction for all three electron acceptors were equivalent to wildtype. These results suggest that ppcA is an intermediary electron carrier forelectrons derived from acetate metabolism in the cytoplasm to Fe(III),U(VI), and AQDS. Hydrogen-dependent reduction may not be affected dueto the presence of a hydrogenase in the periplasm, which may not requirePpcA to transfer electrons to the appropriate carriers in the periplasm orouter membrane. Furthermore, genes for four other periplasmic c-typecytochromes of ca. 10 kDa have been found in the G. sulfurreducens genome.Preliminary results indicate that these genes as well as ppcA are differen-tially expressed depending upon growth conditions, and that the relativeimportance of each of these cytochromes to electron transfer varies withthe electron acceptor being reduced. As with omcB, there is no cytochromein S. oneidensis with high sequence identity to PpcA (Lloyd et al., 2003).


5.2.2.3. PpcB. PpcB is a 36 kDa, di-heme, c-type cytochrome predictedto be localized in the periplasm of G. sulfurreducens, possibly looselyassociated with the inner membrane (Butler and Lovley, 2003). A com-parison of protein and mRNA levels in cells grown on fumarate or Fe(III)indicated that expression of ppcB was up-regulated in Fe(III)-respiringcells. Deleting ppcB from G. sulfurreducens greatly impaired its ability togrow with Fe(III) as the electron acceptor, but had no impact on growthwith fumarate. Rates of Fe(III) reduction in cell suspensions of fumarate-grown cells were only 10% of wild type with acetate as the electron donorand 17% of wild type with hydrogen. These results suggest that PpcB is akey intermediary electron carrier in electron transfer to Fe(III).

5.2.2.4. OmcD and OmcE. Two cytochromes, designated OmcD andOmcE, were readily sheared from the outer membrane of intact cells ofG. sulfurreducens grown on Mn(IV) oxide (Mehta et al., 2003a). OmcD ispredicted to be a 48 kDa tetra-heme c-type cytochrome, whereas OmcE hassix hemes. When either omcD or omcE were deleted, cells grew on all solubleelectron acceptors, including Fe(III) citrate, but did not grow on Fe(III)oxide or Mn(IV) oxide. Given the loose association of the cytochromes withthe outer membrane and the fact that they are required for Fe(III) andMn(IV) reduction, it is possible that these cytochromes serve as terminalreductases for the reduction of Fe(III) and Mn(IV) (Mehta et al., 2003a).

5.2.2.5. Fro1. When NADPH rather than the NADH that had beenprovided in previous studies (Magnuson et al., 2000) was used as theelectron donor for recovering proteins capable of reducing Fe(III) inG. sulfurreducens, a cytoplasmic protein capable of reducing Fe(III) at highrates with NADPH as the electron donor was purified (Kaufmann andLovley, 2001). However, further study demonstrated that this protein,subsequently designated Fro1, was unlikely to be involved in dissimilatoryFe(III) reduction in vivo. For example, spheroplasts of G. sulfurreducensdid not reduce Fe(III) whereas they retained the ability to reduce fumarate,suggesting that the Fe(III) reductase was not localized in the cytoplasm orinner membrane (Coppi et al., 2004). Furthermore, acetate-dependentreduction of fumarate and AQDS, as well as Fe(III), was inhibited in amutant in which Fro1 had been deleted whereas there was little impact onhydrogen-dependent reduction of either electron acceptor (Coppi et al.,2004). The mutant could be adapted over time to slowly grow with acetateas the electron donor. Analysis of gene expression with a whole-genomeDNA microarray indicated that the suite of genes that were up-regulatedand down-regulated in the adapted mutant was consistent with expectedchanges in gene expression if a NADPH dehdyrogenase complex had been


eliminated (Coppi et al., 2004). Thus, the more likely in vivo role of Fro1is as a NADPH dehydrogenase donating electrons for the potential reduc-tion of many electron acceptors. These results further emphasise theimportance of evaluating the role of putative Fe(III) reductases with agenetic approach because of the ability of many redox-active proteins tonon-specifically reduce Fe(III) in vitro.

5.2.2.6. Proteins for localizing and attaching to Fe(III) oxides. Theapparent need for Geobacter species to directly contact Fe(III) oxidesin order to reduce them is associated with a number of adaptations foraccessing this insoluble electron acceptor. Early studies on motility inGeobacter species, which primarily focused on cells grown on solubleelectron acceptors, indicated that Geobacter species were non-motile.However, the discovery of genes for flagella in the genome sequence ofG. sulfurreducens led to a more intensive evaluation of the possibility formotility. Studies with G. metallireducens demonstrated that it specificallyproduced flagella when grown on insoluble Fe(III) or Mn(IV) oxides(Childers et al., 2002). There was little or no flagella production duringgrowth on soluble electron acceptors, including chelated Fe(III). One roleof the flagella may be to permit G. metallireducens to swim to Fe(III) andMn(IV) oxides. Geobacter metallireducens is chemotactic to Fe(II) andMn(II) (Childers et al., 2002) and a gradient of Fe(II) and Mn(II) emanatingfrom Fe(III) and Mn(IV) oxides under anaerobic conditions may leadG. metallireducens to the oxides. Another potential role of the flagella,which has yet to be evaluated in detail, is in the initial attachment to theFe(III) and Mn(IV) oxides.Pili are also specifically produced during growth on Fe(III) and Mn(IV)

oxides and are localized on one side of the cell, the same side as theflagella (Childers et al., 2002). The genome of G. sulfurreducenscontains genes with a high homology to genes in other organisms whichare involved in the production and utilization of type IV pili (Mehta et al.,2003b). When the putative pilA, the gene for the structural pilin protein,was deleted in a G. sulfurreducens mutant, the mutant could no longerreduce Fe(III) oxide, while it reduced soluble Fe(III) as well as the wildtype. Furthermore, the mutant did reduce Fe(III) oxides if an electronshuttle (AQDS) or a Fe(III) chelator (NTA) was added to alleviate theneed for direct contact with the Fe(III) oxides. These results are consistentwith the concept that pili are also required for G. sulfurreducens to accessinsoluble Fe(III) oxides.

5.2.2.7. Other outer membrane proteins. When oxpG, one of the genesin the apparent operon for a putative secretory system in G. sulfurreducens


was deleted, the mutant could not longer grow on insoluble Fe(III) orMn(IV) oxide, but did grow on soluble electron acceptors, includingchelated Fe(III) (Mehta et al., 2003b). The secretory system has highhomology to a novel type II GSP-related pathway that is required forsecretion of a manganese-oxidizing factor by Pseudomonas putida. Thus, itseems likely that this secretory system in G. sulfurreducens is involved inexporting one or more proteins to the outer membrane that are required forthe reduction of Fe(III) and Mn(IV) oxides. Comparisons of proteinlocalization in the mutant and the wild type have identified several proteinsthat are not properly translocated in the mutant, and mutation of one ofthese was found to specifically inhibit Fe(III) oxide reduction.

5.2.2.8. Summary of mechanisms of electron transport to Fe(III) inGeobacter species. The study of electron transport to Fe(III) in Geobacterspecies is clearly in its infancy. However, a preliminary model, takinginto account all the current information, can be constructed (Fig. 3). As inShewanella, it is predicted that c-type cytochromes play an important rolein shuttling electrons from the inner membrane to the outer membraneand that outer membrane cytochromes are important in the electrontransfer to Fe(III), and may themselves possibly serve as the terminalreductase.

5.3. CONCLUSIONS

The current understanding of the physiology of dissimilatory Fe(III)-and Mn(IV)-reducing microorganisms suggests that some of theseorganisms are well adapted for survival and growth in a diversity ofenvironments in which organic matter and/or hydrogen are available aselectron donors where Fe(III) or Mn(IV) is present. In addition to playingan important role in the natural cycle of carbon and metals, Fe(III)- andMn(IV)-reducing microorganisms appear to be useful tools for thebioremediation of contaminated subsurface environments and may beharnessed to harvest electricity from aquatic sediments and waste organicmatter. The understanding of the factors controlling the rate and extentof Fe(III) and Mn(IV) reduction in environments of interest is currentlyrudimentary, at best. However, the availability of the complete genomesequence of a number of dissimilatory Fe(III) and Mn(IV) reducers,coupled with the appropriate genome-scale physiological studies, arelikely to greatly expand the understanding of the physiology these


organisms, and the ability to model predictively their metabolism, underdifferent environmental conditions, in the near future.

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Subject Index

A. nidulans, 25, 27, 30, 37, 48, 50

Absorption spectroscopy, 103–106

Acetobacterium woodii, 183

Acidaminococcus fermentans, 184

Acremonium chrysogenum, 18, 46

Aequorea victoria, 138

Aging and cell death of fungi, 48

Alkaliphiles

proton cycling in, 201–202

sodium cycling in, 201–202

Alkaliphilic bacteria, 207

growing at low �mHþ, 200–209

pH homeostasis in, 202–203

Alkaliphilic-specific amino acid motifs,

205–206

Amino acid motifs, alkaliphilic-specific,

205–206

Amino acid sequence analysis, 96–101

�-(L-a-aminoadipyl)-L-cysteinyl-D-valine

(ACV), 44

Antioxidant defence systems, 24

Arabidopsis, 87

Archaebacteria, 6

Archaeoglobus fulgidus, 95, 101

Aspergillus niger, 11

Atomic force microscopy (AFM),

192–193

ATP hydrolysis, 194–195, 199, 202, 207

regulation of, 207–209

ATP synthase, 199, 207–209

Hþ-coupled, 175–218

Naþ-coupled, 175

proton-coupled, 203–205

sodium-translocating, 187

ATP synthesis, 200, 209

at low �mHþ, 203–205

by rotational catalysis and its dependence

on membrane potential, 194–196

in anaerobic bacteria at low

electrochemical potential, 179–200

Aureobasidium pullulans, 18–19

Bacillus alcalophilus, 200, 202–203

Bacillus firmus, 200, 205

Bacillus halodurans, 200, 205

Bacillus pseudofirmus, 200–204

Bacillus subtilis, 82, 145, 159–160

Bacterial gene circuits, 133

Biosensors as environmental monitors,

131–174

Bioton, carboxylation of, 186

C. boidinii, 40

C. pasteurianum, 112

Candida albicans, 15, 18

Candida glabrata, 36

Carbon starvation, 39

Carbon-starvation response, 151

Carboxybiotin, decarboxylation of, 186

Carboxylation of bioton, 186

Cellular biosensors, 135–136

future trends, 162–163

Chlorinated compounds as electron

acceptors, 248

Class A enzymes, 93–95

Class B enzymes, 95–96

Class B FD-NOR, redox properties of

C-terminal Rd-domain, 112–113

Class B rubredoxin domain, 100

Class C enzymes, 96

Class C NAD(P)H:flavin oxidoreductase

domain, 100–101

Clostridium acetobutylicum, 94

Clostridium perfringens, 93

cobA, 140

Contaminated environments, 223–224

Core domains, 97–100

Crithidia fasciculata, 6

CymA, 262

L-Cysteine, biosynthesis and transport,

153–154

L-Cysteinylglycine dipeptidase (CG), 17

Cytochrome c3, 263

Decarboxylation of carboxybiotin, 186

Decarboxylation reaction, 175

Desiccation, 34

Desulfobibrio vulgaris, 238

Desulfobulbus propionicus, 246

Desulfotomaculum reducens, 246

Desulfovibrio, 241

Desulfovibrio gigas, 88–91, 93, 97,

104, 108

Detoxification, 37

GSH-dependent processes, 40–46

of xenobiotics, 42–44

Dicyclohexylcarbodiimide (DCCD),

190–191, 194–195

Diiron centre, 90

Dissimilatory Fe(III) and Mn(IV)

reduction, 219–286

alternative electron acceptors, 244–250

components involved in electron

transfer, 260

direct contact versus electron shuttling

or chelation, 255–258

electron donors, 250–253

environmental considerations, 222–237

forms of Fe(III) and Mn(IV) available,

227–237

Geobacter species, 264–269

influence of humic substances, 227–237

major groups of reducing

microorganisms, 237–243

mechanisms, 254–269

methals other than Fe(III) and Mn(IV),

244–245

models for electron transfer to

extracellular Fe(III) and Mn (IV)

oxides, 258–269

overview, 221–222

pH range, 253–254

physiological diversity, 243–254

salinity range, 253–254

Shewanella species, 261–264

sources of electron donors, 226–227

temperature range, 253–254

Dithiothreitol (DTT), 21

L-djenkolic acid, 154

DNA arrays, 158–160

DNA damage, 144

dsRed, 139–140

Electrodes, as electron donors, 252–253

Electron acceptors, alternative, 244

Electron donors, 250–253

sources of, 226–227

Electron transfer chains, 93–96

Electron transfer models, 258–269

Electron transport proteins, 259

Energy harvesting electrodes in

sediments, 226

Entamoeba histolytica, 21, 23, 97

Environmental monitoring, 135–136

EPR spectroscopy, 106–108, 112

Erwinia chrysamthemi, 85, 119

Escherichia coli, 82, 86–88, 95, 104, 116,

135, 138, 144–145, 147, 150–152,

155–160, 183, 187, 196, 199,

206, 208

Escherichia coli flavorubredoxin, 106, 114

diiron centre in ß-lactamase module, 112

EPR spectra, 107

NO reductase activity, 115

Extracellular quinones, 246

F0 motor model, 197–200

Fe(II) as electron donor, 252–253

Fe(III) oxides, proteins for localizing and

attaching to, 268

298 SUBJECT INDEX

Fe(III) reduction, 219–286

microorganisms that conserve energy to

support growth from, 238–243

microorganisms that do not conserve

energy to support growth from,

237–238

organisms known to conserve energy to

support growth from, 228–235

see alsoDissimilatory Fe(III) and Mn(IV)

reduction

Ferroglobus placidus, 242–243

Flavodiiron proteins, 77–129

biochemical properties, 102

classification, 92–93

dendrogram, 99

enzymatic studies, 114–116

function of, 114–119

modular arrangement, 92

molecular genetics studies, 116–119

physicochemical properties, 101–113

redox properties, 108–113

spectroscopic studies, 103–108

UV-visible spectra, 105

Flavodoxin domain, 91

Flavohemoglobins, 85

FMN pocket, structural modelling, 103

Fro1, 267

Fumarate as electron acceptor, 247–248

Fumarate Nitrate Regulator (FNR), 84

Gene expression, reporters of, 136–142

Gene fusion arrays, 160–161

Genotoxicity, 142–145

Geobacter, 220

Geobacter metallireducens, 240, 245, 247

Geobacter sulfurreducens, 240, 244, 247–248

Geobacteraceae, 239–240

Geothrix, 220

gfp, 138–139

GFP, detection of, 139

Giardia lamblia, 97

Globin family of proteins, 84–87

Glutaconyl-CoA decarboxylase, 184

g-glutamyl cycle, 9–10

Glutaredoxin thioredoxin, oxidation

systems, 28–29

Glutathione (GSH), 1–76

biology of, 4

biosynthesis, 7, 9–12, 20

degradation, 7, 9–10

degradation and recycling, 15–17

detoxification, 8

dispensability in microorganisms, 20

extracellular functions, 17–18

in cell differentiation and development,

18–19

in stress responses, 23–40

in unstressed cells, 19–23

metabolism, 4

metabolism in fungi, 9–23

metabolism under unstressed conditions,

11–13

overview, 6–9

oxidation systems, 28–29

principal enzyme and transport systems

associated with, 7

production and consumption, 5

reasons for studying, 4–5

redox balance, 7

regulation of biosynthesis, 12–13

regulator of b-lactam antibiotic synthesis,

44–46

role in aging and autolysis, 48–51

synthesis, 27

transport, 9

transport and metabolism in

Saccharomyces cerevisiae, 15

uptake and storage, 13–15

see also GSH/GSSG

Glutathione (GSH)-conjugates, uptake and

storage, 13–15

Glutathione (GSH) synthases, 25

Glutathione disulfide (GSSG), 6, 34

see also GSH/GSSG

Glutathione-glutaredoxin-thioredoxin

system, 26

Glutathione reductase (GR), 6, 17

Glutathione S-transferase (GST), 8

Glyoxalases (GLO) I and II, 41

Graphite electrodes as electron acceptor,

249–250

GS-X, transport and metabolism in


GSH/GSSG redox balance/imbalance, 17,

19, 48

GSH/GSSG redox ratio, 22, 27, 50

GSH/GSSG redox signaling, 18

gGT, 15–17

SUBJECT INDEX 299

Hþ-coupled ATP synthase, 175–218

Hansenula polymorpha, 34, 40

Heat shock, 33

Heat shock response, 145–146

Heavy metal stress, 35–38

Helicobacter pylori, 87

High cell density cultures, 34–35

Histoplasma capsulatum, 18

Hot environments, 224–226

Humic substances, influence of, 227–237

Hydrogen as electron donor, 251

Hydroquinones as electron donors, 252–253

Hyperthermophilic microorganisms,

242–243

IfcA, 262

Ilyobacter tartaricus, 182–183, 189–192

inaZ, 138

Klebsiella pneumoniae, 182–183

b-lactam antibiotic synthesis, 44–46

Lactobacillus fermentum, 82

lacZ, 138

luc, 140

luxAB, 141–142

luxCDABE, 141–142

Macromolecular damage, 142–150

Magnaporthe grisea, 25, 27, 30

MALDI mass spectrometry, 194

Malonomonas rubra, 184

Membrane damage, 147

Mesophilic bacteria, 205–206

Metallo b-lactamase domain, 90–91

Methanocaldococcus janaschii, 95, 97

Methanosarcina acetovorans, 95

Methanothermobacter thermoacetica, 101,

104, 108

Methanothermobacter thermoautotrophicus,

88, 103

Methylglyoxal, 41

Methylmalonyl-CoA decarboxylase, 184

Microbial respiration, 224–226

Mn(IV) reduction, 219–286

see alsoDissimilatory Fe(III) and Mn(IV)

reduction

Moorella thermoacetica, 95

Mossbauer spectroscopy, 108

MtrA, 262

MtrB, 263

Multi-nutrient starvation response,

154–155

Multi-stress responses to toxic metals,

147–150

Mycobacterium tuberculosis, 85, 158

Naþ-coupled ATP synthase, 175

Nematoloma frowardii, 18

Neurospora crassa, 15, 18, 25, 27, 30

Nitrate as electron acceptor, 246

Nitric oxide (NO)

and microbes, 81–87

biological chemistry, 80–81

chemistry of, 78–80

Nitric oxide (NO) detoxification, 77–129

Nitrite reductase (Nrf), 84

Nitrogen fixation and autotrophy, 254

Nitrogen species, oxidation state diagram

for, 79

Nitrogen starvation, 38

Nitrogen-starvation response, 152

Nitrosative stress

genetic responses, 83–84

prokaryotic defence systems against,

84–87

Nutrient deprivation stress, 38–40

Nutrient limitation/imbalance, 150–155

OmcA, 263

OmcB, 264–266

OmcD, 267

OmcE, 267

Organic electron donors, 250

Osmotic shock, 33

Oxidation state diagram for nitrogen

species, 79

Oxidative stress, 23–33

Oxidative-stress responsive, 146–147

OxyR, 83, 86

P. putida, 150, 153

Paxillus involutus, 37

Penicillium chrysogenum, 18, 30, 38, 40, 43,

46–47, 49–50

pH homeostasis in alkaliphilic bacteria,

202–203

Phosphate-starvation response, 152–153

300 SUBJECT INDEX

Photinus pyralis, 140

Photobacterium phosphoreum, 142

Photorhabdus luminescens, 142, 156

PpcA, 266

PpcB, 267

Pristine sediments, 222–223

Prokaryotic defence systems against

nitrosative stress, 84–87

Propionibacterium freudenreichii cobA, 140

Propionigenium modestum, 175, 177–180,

182–183, 187–188, 190–191, 195–197,

205–206

Proteins, globin family of, 84–87

�-Proteobacteria, 239

Proton-coupled ATP synthase, 203–205

Proton cycling in alkaliphiles, 201–202

Protonophore side-chain precursors, 47

Pseudomonas, 146, 162

Pseudomonas fluorescens, 150–151, 153–154

Pseudomonas syringae, 138

Pyrobaculum islandicum, 242

Pyrophorus plagiophthalamus, 140

Ralstonia eutropha, 116

Reactive oxygen species (ROS), 23–33, 48

Renilla reniformis, 141

Rhodobacter capsulatus, 84, 88, 104, 106

Ribonucleotide reductases (RNR), 90

ruc, 141

S-nitroglutathione (GSNO), 80

Saccharomyces cerevisiae, 4, 10–11, 14–15,

17–18, 21–22, 30, 35–36, 38–39,

48–49, 87

Salmonella typhimurium, 86–87, 156, 160

Schizosaccharomyces pombe, 10–12, 15, 17,

22–23, 25, 36–37

Sediments, energy harvesting electrodes

in, 226

Shewanella, 220, 240–242, 247

Sodium cycling in alkaliphiles, 201–202

Sodium ion cycles in bacteria, 180–183

Sodium-translocating decarboxylases,

�mNaþ generation by, 183–187

Sodium-translocating F1F0 ATP synthase,

180–183, 187

Soils, 222–223

SoxRS, 83, 86

Staphylococcus aureus, 82

Stress response

definition and scope, 133–134

specificity and sensitivity, 134–135

Stress responsive bacteria, 131–174

Stress responsive gene fusions,

156–158

Subsurface environments, 222–223

Sulfospirillum barnesi, 245

Sulfur, transport and metabolism in


Sulfur compounds as electron

acceptor, 246

Sulfur starvation, 39

Sulfur-starvation response, 153–154

Superoxide dismutase (SOD)/catalase

enzymes, 24

Synechococcus, 152

Synechocystis, 100, 106

Synechocystis sp. PCC6803, 104, 116

Synechocystis SsATF573, 101

Thermoalkaliphilic bacteria, 205–206

Thermodesulfobacterium commune, 242

Thermosynechococcus elongatus, 100

Thermotoga maritima, 181–182, 242

Thiol group, 6

Toxic metabolites, elimination of, 40–42

Toxic metals, multi-stress responses to,

147–150

Trypanosoma cruzi, 6

UV-visible spectra, 112

Vibrio alginolyticus, 181

Vibrio cholerae, 181

Vibrio fischeri, 141–142, 156

Vibrio harveyi, 142

Vitreoscilla, 85–86

Xenobiotics, detoxification of, 42–44

Zygosaccharomyces bailii, 49

SUBJECT INDEX 301

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