Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes

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9 Microbiology Monographs Series Editor: Alexander Steinbüchel

Microbiology Monographs Volumes published in the series

Inclusions in Prokaryotes Jessup M. Shively (Editor) Volume 1 (2006) Complex Intracellular Structures in Prokaryotes Jessup M. Shively (Editor) Volume 2 (2006) Magnetoreception and Magnetosomes in Bacteria Dirk Schüler (Editor) Volume 3 (2007) Predatory Prokaryotes – Biology, Ecology and Evolution Edouard Jurkevitch (Editor) Volume 4 (2007) Amino Acid Biosynthesis – Pathways, Regulation and Metabolic Engineering Volker F. Wendisch (Editor) Volume 5 (2007) Molecular Microbiology of Heavy Metals Dietrich H. Nies and Simon Silver (Editors) Volume 6 (2007) Microbial Linear Plasmids Friedhelm Meinhardt and Roland Klassen (Editors) Volume 7 (2007)

Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes Volume Editor: Jan Tachezy

123

Volume Editor: Prof. Jan Tachezy, Ph. D. Department of Parasitology Faculty of Science Charles University Viniˇcna 7 12844 Prague 2 Czech Republic e-mail: [email protected] Series Editor: Professor Dr. Alexander Steinbüchel Institut für Molekulare Mikrobiologie und Biotechnologie Westfälische Wilhelms-Universität Corrensstraße 3 48149 Münster Germany e-mail: [email protected]

ISBN 978-3-540-76732-9 DOI 10.1007/978-3-540-76733-6

e-ISBN 978-3-540-76733-6

Microbiology Monographs ISSN 1862-5576 Library of Congress Control Number: 2007940388 c 2008 Springer-Verlag Berlin Heidelberg This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, speciﬁcally the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microﬁlm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a speciﬁc statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover Design: WMXDesign GmbH, Heidelberg, Germany Printed on acid-free paper 9876543210 springer.com

Preface

Hydrogenosomes represent a group of double membrane-bound organelles that have the production of molecular hydrogen and ATP in common. Such a well deﬁned feature is unknown for another type of double membrane-bound organelles called by various names such as mitosomes, cryptons, remnant mitochondria or mitochondria-like organelles. These organelles neither produce hydrogen nor synthetize ATP and we know little about their functions, except the biogenesis of iron-sulfur clusters found in some of them. The common denominator for both hydrogenosomes and mitosomes is their evolutionary origin. It is generally accepted that both hydrogenosomes and mitosomes evolved from mitochondria or an ancestral pre-mitochondrial endosymbiont (discussed by Bill Martin in this volume). With our growing knowledge about hydrogenosomes and mitosomes it is also becoming apparent that it will be difﬁcult to draw a clearly deﬁned line between mitochondria, hydrogenosomes and mitosomes. As we believe and as the title of this volume indicates, all these organelles are a kind of mitochondria in a broad sense, and the reason for their different names is primarily historical. In line with this hypothesis, the name “amitochondriates” should no longer be used for the organisms possessing hydrogenosomes and mitosomes, since this term refers to an alleged absence of mitochondria. We would like to thank all the authors who contributed outstanding chapters about hydrogenosomes and mitosomes to this Microbiology Monographs volume summarizing current knowledge about evolution, biogenesis, structure and function of these exciting organelles. We highly appreciate the time and effort that the authors invested in the preparation of their manuscripts. Although this form of publication is usually not scored by grant agencies, it is essential in providing comprehensive information to readers working in other ﬁelds of biology and in attracting young scientists and students interested in the ﬁeld of protist cell biology. We would like to dedicate this volume to Miklós Müller and Jaroslav Kulda ˇ erkasovová, who together with their colleagues (Donald Lindmark, Apolena C ˇ Jiˇrí Cerkasov) offered the ﬁrst description of hydrogenosomes in the early 1970s. Although various new hydrogenosomal pathways and components have since been revealed (especially by recent genomic and proteomic studies), the

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basic metabolic pathways uncovered in pioneering biochemical studies in their laboratories in New York and Prague remain a backbone of hydrogenosomal metabolism in trichomonads. Prague and Münster, December 2007

Jan Tachezy Alexander Steinbüchel

Introduction

,,Es ist schlimm genug,” rief Eduard, ,,dass man jetzt nicht mehr für sein ganzes Leben lernen kann. Unsere Vorfahren hielten sich an den Unterricht, den sie in ihrere Jugend empfangen; wir aber müssen jetzt alle fünf Jahre umlernen, wenn wir nicht ganz aus der Mode kommen wollen.” Johann Wolfgang Goethe, Die Wahlverwandschaften, 1807 “It is bad enough,” said Eduard, “that our learning does not last for our whole life. Our ancestors could hold onto the learning that they acquired in their youth; we have to relearn everything every ﬁve years, just to remain fashionable.” (Author’s translation) This volume summarizes our knowledge on a group of diverse and “unusual” cell organelles. It is published at a most appropriate time when it seems that a consensus is emerging about the nature of these entities. Evidence accumulating over the past few years has brought home the prodigal sons of the mitochondrial family. Hydrogenosomes and mitosomes are now within the fold of the well-known and respected organelle family that includes the “powerhouses” of the aerobic eukaryotic world. Chapters in this book describe in detail these organelles which exhibit a bewildering array of characteristics but according to an emerging consensus of most biologists all are mitochondria in different guises. They are assumed to represent a monophyletic group of biological entities that originated from an ancestral endosymbiotic event that gave rise to the protomitochondrion. Of course the emerging consensus does not mean that there are no dissenters in many quarters of the biology community. What convincing arguments today might not turn out to be red herrings tomorrow? The history of the past 30 or so years gives ample warning. Our views and theories related to this ﬁeld have changed drastically so many

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times during these years that we cannot be certain that future surprises are not lurking around the corner. At any rate, this book takes stock of the status quo. It presents a cast consisting of double-membrane bounded organelles of eukaryotic protists and fungi, which live under anaerobic or hypoxic conditions, as well as certain intracellular parasites of aerobic cells. The common denominator of these organelles is that they lack the aerobic energy conservation system of typical mitochondria, i.e. the cytochrome-dependent electron transport chain and oxidative phosphorylation. Another common denominator is that—one curious exception disregarded—they are the site of a process, the synthesis of iron–sulfur clusters, which is regarded today as the only really indispensable function of eukaryotic mitochondria. These organelles exhibit, however, properties and characteristics in so many different combinations that all generalizations of the group (beyond the just-mentioned major negative and positive hallmarks) would be futile, i.e. any general deﬁnition would require more qualiﬁcations than unequivocal statements. As evident from the individual chapters, the major members of this family are the mitosomes, small organelles without a known role in energy metabolism, and the hydrogenosomes, organelles of the approximate size of mitochondria, characterized by their ability to produce molecular hydrogen as a metabolic end product. Recent evidence, however, has revealed that the latter organelles can be quite different, anywhere from typical mitochondria to organelles that do not resemble mitochondria at all. The individual differences within these groups are so great, and the boundaries between them so undeﬁned, that no generally acceptable nomenclature has yet emerged. Each investigator uses a different designation for his/her organelle. I wish to emphasize the perhaps most signiﬁcant aspect of the emerging consensus. The distribution of these organelles in the living world is broad. They are found in certain members of almost all of the currently recognized major evolutionary lineages of eukaryotes. Their presence seems to be primarily correlated with the ecology of the organisms, which live in an anoxic habitat or in a nutrient-rich intracellular niche that permits life without the efﬁcient energy-generating system of typical mitochondria. This distribution argues strongly for the hypothesis that diverse mitosomes and hydrogenosomes arose independently, and repeatedly, during the diversiﬁcation of the eukaryotes. Convergent evolution in action! It also argues strongly against the earlier assumption that the presence of these organelles could represent an ancestral, pre-mitochondrial state. As mentioned above, the road leading up to the present state of affairs was convoluted, and often terminated in dead-end streets. We had to change our views and hypotheses repeatedly, had to relearn everything, just as Eduard was required to relearn everything two centuries ago. This foreword is not the place to recount the history of our views of these organelles. I gave an account of the events leading up to the ﬁrst description of a hydrogenosome in another

Introduction

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place (Müller, 2007), and so I hope that someone will tell us the story as it has unfolded during the intervening years. I wish to add some personal remarks here. I was very pleased when the editor of this volume, Honza Tachezy, asked me to write a brief foreword and I thank him very much. I am certain that this book will play a major role in the future development of this exciting area of comparative and evolutionary biology. Little did Don Lindmark and I think in 1973 that our description of an unusual cell organelle from a group of odd parasites would give the ﬁrst push to the unfolding of what turned out to be a most exciting story in evolutionary cell biology (Lindmark and Müller, 1973). It is a great personal satisfaction to me, and I am sure, to all those who were with me at various times, to see where our endeavors have led. I look forward to seeing what the future holds. Lastly, I wish to acknowledge the unfailing support and friendship of my mentor Christian de Duve. It was in his Department at Rockefeller University where Don and I performed the initial experiments, and where I have spent many productive years. I would like to thank all the past members of my group who were responsible for all their own work on these organelles. I also thank all those colleagues working in diverse parts of the world who believed in the promise this ﬁeld held and carried the torch further. I received much moral support from them, however, they are too numerous to be named here. I also acknowledge the uninterrupted support of our original work at the Rockefeller University for almost 30 years by the same NIH grant (AI 11942) and also by several grants from NSF. The Rockefeller University, New York and Collegium Budapest, Budapest, September 2007

Miklós Müller

References Lindmark DG, Müller M (1973) Hydrogenosome, a cytoplasmic organelle of the anaerobic ﬂagellate, Tritrichomonas foetus, and its role in pyruvate metabolism. J Biol Chem 248:7724–7728 Müller M (2007) The road to hydrogenosomes. In: Martin WF, Müller M (eds) Origin of mitochondria and hydrogenosomes. Springer, Berlin Heidelberg New York, pp 1–11

Contents

Anaerobic Eukaryotes in Pursuit of Phylogenetic Normality: the Evolution of Hydrogenosomes and Mitosomes W. Martin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

Protein Import into Hydrogenosomes and Mitosomes S. D. Dyall · P. Doleˇzal . . . . . . . . . . . . . . . . . . . . . . . . . . . .

21

Structure of the Hydrogenosome M. Benchimol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

75

Hydrogenosomes of Anaerobic Ciliates J. H. P. Hackstein · R. M. de Graaf · J. J. van Hellemond · A. G. M. Tielens

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Metabolism of Trichomonad Hydrogenosomes I. Hrdý · J. Tachezy · M. Müller . . . . . . . . . . . . . . . . . . . . . . . 113 Hydrogenosomes of Anaerobic Chytrids: An Alternative Way to Adapt to Anaerobic Environments J. H. P. Hackstein · S. E. Baker · J. J. van Hellemond · A. G. M. Tielens . . 147 The Proteome of T. vaginalis Hydrogenosomes K. Henze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Hydrogenosome: The Site of 5-Nitroimidazole Activation and Resistance J. Kulda · I. Hrdý . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 Mitosomes in Parasitic Protists J. Tachezy · O. ˇSmíd . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 The Mitochondrion-Related Organelle of Cryptosporidium parvum J. S. Keithly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231

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Mitochondrial Remnant in Blastocystis N. Yarlett · K. S. W. Tan . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 Possible Mitochondria-Related Organelles in Poorly-Studied “Amitochondriate” Eukaryotes V. Hampl · A. G. B. Simpson . . . . . . . . . . . . . . . . . . . . . . . . 265 Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_108/Published online: 8 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Anaerobic Eukaryotes in Pursuit of Phylogenetic Normality: the Evolution of Hydrogenosomes and Mitosomes William Martin Institute of Botany, University of Düsseldorf, Universitätsstr. 1, 40225 Düsseldorf, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2

Older Views of How and Why the Mitochondrion Become Established . .

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Anoxic and Sulﬁdic Oceans up to ∼ 580 MY Ago . . . . . . . . . . . . . .

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Canﬁeld Oceans Give Anaerobic Eukaryotes Room to Breathe . . . . . . .

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Anoxic, Fine, and What About Sulﬁdic? . . . . . . . . . . . . . . . . . . . .

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6

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract The evolutionary relationship of hydrogenosomes and mitosomes to mitochondria is no longer an issue of contention for specialists who work on the organelles. Hydrogenosomes and mitosomes are mitochondria in the evolutionary sense in that they descend from one and the same eubacterial endosymbiont, but the evolutionary signiﬁcance of eukaryotic anaerobes that possess hydrogenosomes, mitosomes, and anaerobically functioning mitochondria is an issue of some contention. This chapter serves to further develop the thesis that the role of oxygen in eukaryote evolution needs to be reconsidered and viewed in light of what geologists have discovered relatively recently regarding oxygen in Earth history. According to newer ﬁndings from geochemical studies, there existed during a protracted period of Earth ocean history, during which the oceans were mostly anoxic and sulﬁdic (“Canﬁeld” oceans). This period started about 2.3 billion years ago and only came to an end about 580 million years ago. This was the time during which eukaryotes arose and diversiﬁed into their major lineages. In light of this, anaerobic eukaryotes with mitochondria are not, in an evolutionary sense, strange, obscure, unexpected, or otherwise out of the ordinary, hence no special or unusual mechanisms are required to explain their origin. They are normal in every respect, and so are their mitochondria.

1 Introduction Hydrogenosomes and mitosomes are mitochondria in the evolutionary sense in that they descend from one and the same eubacterial endosymbiont, but

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W. Martin

they differ from the mitochondria that most people know from textbooks in that neither hydrogenosomes nor mitosomes studied to date have been found to possess mechanisms of oxidative phosphorylation. Hydrogenosomes and mitosomes occur among eukaryotes that have oxygen-independent ATP synthesis. Accordingly, they occur among various protists that inhabit anaerobic environments and among various protists that have a parasitic lifestyle. The discovery and study of both organelle types has had a substantial impact on our understanding of eukaryotic evolution. It is probably fair to state that everyone who currently works on hydrogenosomes and mitosomes agrees that these organelles are mitochondria in the evolutionary sense. The list of those who agree on this issue would surely include those who discovered hydrogenosomes (Lindmark and Müller 1973), those who discovered mitosomes (Tovar et al. 1999), many authors of this volume, and the authors of another volume that recently appeared on the topic (Müller 2007; Lane 2007; Allen et al. 2007; Sapp 2007; Tielens and van Hellemond 2007; Tachezy and Dolezal 2007; Hackstein et al. 2007; Cavalier-Smith 2007; Emelyanov 2007; Barbera et al. 2007; Tovar 2007). The evidence to support that view has been the subject of numerous recent topical reviews and minireviews (Embley et al. 1997; Biagini et al. 1997; van der Giezen et al. 2005; van der Giezen and Tovar 2005; Hackstein et al. 2006; Embley and Martin 2006) and various facets thereof are summarized in the other chapters of this volume. The evidence from the genome sequences of organisms that possess hydrogenosomes, Trichomonas (Carlton et al. 2007), or mitosomes— Encephalitozoon (Katinka et al. 2001) and Entamoeba (Loftus et al. 2005)—is also fully consistent with the view that both organelles share a common ancestry with mitochondria. Hence there is no need to recite here once more the evidence in support of this view, which is based in gene sequence comparisons (van der Giezen et al. 2005; Embley and Martin 2006) shared mechanisms and components of protein import (Dolezal et al. 2006), shared enzymes (Tielens et al. 2002; Müller 2003; see chapters by Henze and Müller in this volume), and, in one case (Boxma et al. 2005), shared organelle DNA sequences. However, the circumstance that everyone who works on hydrogenosomes and mitosomes agrees on the mitochondrial nature of these organelles does not mean that everyone agrees with that view, hence there remain a few prominent dissenting opinions on the issue. The most outspoken critics of common ancestry for mitochondria and hydrogenosomes tend to either overlook the evidence indicating the common ancestry for these organelles, to dismiss it as equivocal, or to marginalize its possible evolutionary signiﬁcance (Margulis et al. 2006; de Duve 2007). Other critics of common origin favor unspeciﬁed lateral gene transfer (LGT) mechanisms to explain hydrogenosome origin (Kurland and Andersson 1999; Andersson et al. 2003). Yet a different camp attributes the ability of eukaryotes to survive without oxygen to piecemeal LGT mechanisms, while avoiding the issue of the organelles altogether (Andersson et al. 2007). Most current evidence for LGT comes from the study of trees and branch

Evolution of Eukaryotic Anaerobes

3

lengths, and at present nobody knows which, what kind of, or how many phylogenetic trees we can really trust (see Dyall et al. 2004 vs. Hrdy et al. 2004, for a particularly clear example of different trees for the same data). There are thus two kinds of issues to consider: (1) the evolutionary relationship of hydrogenosomes and mitosomes to mitochondria, which is no longer really an issue for any specialists and hence will not be dealt with here, and (2) the phylogenetic status and evolutionary signiﬁcance of eukaryotic anaeorobes that possess hydrogenosomes, mitosomes, and anaerobically functioning mitochondria in the geological context of life’s history on Earth. The latter issue is of some interest. This chapter serves to further develop the thesis, presented to some extent in earlier papers (Martin et al. 2003; Martin 2007), that (i) the role of oxygen in eukaryote evolution needs to be reconsidered and viewed in light of what geologists are telling us about oxygen in Earth history, and that (ii) mitochondriate anaerobes are altogether normal eukaryotes with no special kind of evolutionary rank or status. It will be argued that the common ancestry of ATP-synthesizing forms of mitochondria among eukaryotic aerobes and anaerobes and the presence of mitochondria in eukaryotes that synthesize their ATP in the cytosol are most readily interpreted in light of evolutionary specializations of the mitochondrion. Conversely, the strictly oxygen-dependent forms of the organelle that we know from organisms the live above the soil line (or above the sediment in aquatic environments) are also most easily interpreted as evolutionary specializations of the organelle from a more generalized ancestral state—facultatively anaerobic—that existed during a protracted, anaerobic period of Earth (n.b.) ocean history. According to newer ﬁndings that geologists have been reporting for about 10 years, this period, in which the oceans were mostly anoxic and sulﬁdic (Canﬁeld oceans), started about 2.3 billion years ago and only came to an end about 580 million years ago. A main goal of this chapter will be to convey to the readers of this book (mainly biologists) the new view of oxygen accumulation during Earth history that has emerged over the last 10 years. The newer view of oxygen in Earth history would render anaerobic eukaryotes with mitochondria a natural observation to be expected in many disparate lineages. Under this view, anaerobic eukaryotes with mitochondria are not, in an evolutionary sense, strange, obscure, or otherwise out of the ordinary, hence no special or unusual mechanisms are required to explain their origin. They are normal in every respect, and so are their mitochondria.

2 Older Views of How and Why the Mitochondrion Become Established Current concepts relating to the origin of mitochondria are most readily understood before the backdrop of how these concepts arose. Since about 1967,

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mainstream reasoning on the rationale behind mitochondrial origins has focussed on oxygen, ATP, and improved energy yield from glucose breakdown through oxidative phosphorylation. There are exceptions to this rule (Blackstone 1995), but the exceptions prove the rule. In order for there to be a biochemical rationale behind mitochondrial origins, there has to be a null hypothesis about the nature of the host, so that one can explicate the possible nature of host–symbiont interactions that lead to a stable symbiosis and hence, in turn, to specify the selective advantages for either partner during the transition from endosymbiont to organelle. The host has always been the obscure partner in endosymbiotic theory as it concerns the origin of mitochondria. Following the rise and fall endosymbiotic theory in the ﬁrst half of the 20th century (Merschkowsky 1905; Sapp 1994), Margulis (named Sagan in 1967) suggested that the host that acquired the mitochondrial endosymbiont (hereafter just called “the host”) was an anaerobic, heterotrophic, fermenting, cell wall-lacking (amoeboid, in the broad sense) prokaryote perhaps similar to modern Mycoplasma (Sagan 1967; Margulis 1970). This host corresponded, in terms of cell topology, to the nucleocytoplasmic component of eukaryotes. Margulis believed that the selective advantage that the mitochondrion conferred upon its assumedly fermenting host was improved ATP yield from glucose breakdown by virtue of oxygen respiration and oxidative phosphorylation. Sagan (1967, p 229) wrote: “The anaerobic breakdown of glucose to pyruvate along the Embden–Meyerhof pathway occurred in the soluble cytoplasm under the direction of the host genome. [...] The greater amounts of energy available after the incorporation of the mitochondrion resulted in large cells with amoeboid and cyclotic movement”. This idea was highly compatible with another view that emerged at the time, namely that the origin of eukaryotes (and their mitochondria) corresponded temporally and causally to the global rise in atmospheric oxygen levels ∼2 billion years ago. For example, Sagan (1967, p 225) wrote: “The subsequent evolution of aerobic metabolism in prokaryotes to form aerobic bacteria (protoﬂagella and protomitochondria) presumably occurred during the transition to the oxidizing atmosphere”. A main point of this chapter, to which we will return to shortly, concerns the last word of that quote, because microbial evolution did not take place in the atmosphere. That dual assumption of Margulis’s original versions of mitochondrial origins, which one can call the “oxygen/ATP” argument (Martin 2007), was accepted by many subsequent authors, most notably in the present context by those who contest the common ancestry of mitochondria, hydrogenosomes, and mitosomes. However, most people have always rejected the part of Margulis’s theory suggesting that ﬂagella (and in later formulations, also the nucleus) stem from spirochaete symbionts. Similarly, most people have always rejected the part of de Duve’s (1969, 2007) modiﬁcation of Margulis’s (Sagan 1967; Margulis et al. 2006) theory in which peroxisomes, instead of spirochaetes, are the additional endosymbiont. Possibly as a consequence

Evolution of Eukaryotic Anaerobes

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of that and other early controversies about endosymbiosis in cell evolution (Cavalier-Smith 1975), a new view emerged around 1980 that placed the oxygen/ATP advantage in the context of a modiﬁed form of Margulis’s theory that (i) lacked the spirocheate endosymbiont and that (ii) assumed the newly discovered archaebacteria to be relatives of the host lineage, ﬂanked by the assumption that the host was a phagocytotic, anaerobic, fermenting eukaryote (possessing a nucleus and other salient eukaryotic features). The 1980 view assumed that the prokaryote-to-eukaryote transition occurred via gradualist mechanisms such as point mutation and hence did not involve symbiosis at all (van Valen and Maiorana 1980; Doolittle 1980) and culminated with a cell that possessed a nucleus, but lacked mitochondria. This is what Doolittle (1998) has called the “standard model”. In this view, mitochondria are interpreted as a small tack-on to, and mechanistically unrelated to, the process that made eukaryotic cells nucleated and complex (Cavalier-Smith 2002). In the standard model, mitochondria (and chloroplasts) are descended from endosymbionts, but the nuts-and-bolts of the prokaryote-to-eukaryote transition (the origin of eukaryote-speciﬁc traits) was seen as having occurred independently from, and prior to, the origin of mitochondria. The paper by van Valen and Maiorana (1980) expresses this view in clear physiological terms: the host was assumed to be an amoeboid, anaerobic, fermenting cell related to archaebacteria, the advantage of the mitochondrial endosymbiont was to supply ATP. The most important sentence of this section probably got lost in the foregoing, hence it is repeated here: The 1960s idea that oxygen and ATP drove the origin of mitochondria was highly compatible with another 1960s view, namely that the origin of eukaryotes (and their mitochondria) corresponded temporally and causally to the global rise in atmospheric oxygen levels ∼2 billion years ago. In that 1960s view, there was no room for anaerobic eukaryotes and especially no room for their organelles. The evolutionary signiﬁcance of hydrogenosomes and mitosomes is, today, still marginalized by proponents of theories that have one assumption in common: The origin of eukaryotes and their mitochondria was viewed as mechanistically related to global oxygen change, but this idea might be wrong, so it is worthwhile to see if geochemical evidence tells a different story about the global history of oxygen in 2007 than in 1970. This is the subject of the next section. We will see that eukaryotic anaerobes ﬁt much better in modern views of global oxygen change.

3 Anoxic and Sulfidic Oceans up to ∼580 MY Ago The older, traditional model of global oxygen history arguably starts off with a paper (Cloud 1968) that appeared at the time when Margulis was reviving endosymbiotic theory. Further developed and supported by much evidence

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W. Martin

into the 1990s, the model went more or less like this: The early Earth was devoid of O2 , O2 in the atmosphere stems from photosynthesis, cyanobacteria produced several millions of billions of tonnes of O2 , much of which served to oxidize Fe(II) in the early oceans, and once that titration process was complete, the planet was oxidized and O2 started accumulating in the atmosphere about 2–2.3 billion years ago as evidenced by the disappearance of particular uranium minerals and the appearance of redbeds (iron oxidized on continents) at that time (Holland and Beukes 1990; Kasting 1993). In this model, the transition from anoxic oceans to oxic oceans occurred in a very narrow window of time (< 100 Myr) and coincided with the appearance of atmospheric O2 (Kasting 1993). In this (now outdated, see next paragraph) view of global oxygen history, one could make sense of eukaryotes in a model that went more or less like this: Eukaryotes are a very ancient lineage, the most primitive lineages branched off early and lack mitochondria, and with the advent of O2 , mitochondria and the lineage leading to higher eukaryotes arose (Knoll 1992). This is also the essence of the recent contributions by Margulis et al. (2006) and de Duve (2007), who cast doubt upon the common ancestry of mitochondria and hydrogenosomes, who disregard mitosomes, and who derive eukaryotes via their own versions of endosymbiotic theory that entail hypothetical symbioses (ﬂagella vs. peroxisomes, respectively) preceding that which gave rise to the mitochondrion. The main biological interpretation of these models, and other models that have no room for hydrogenosomes, mitosomes, or anaerobes among evolutionarily advanced eukaryotic lineages is: Anaerobic eukaryotes without mitochondria arose early (and are somehow poorly understood or mysterious), aerobic eukaryotes with mitochondria arose late, the border between early and late coincides with O2 accumulation. The ﬁrst chinks in the old geological model came from unexpectedly late occurrences and reinterpretation of particular deposits called banded iron formations (BIFs) (Holland 1999), but the model still stood. Over about the last 10 years, evidence obtained primarily from the geochemical sulfur isotope record has given rise to a fundamentally new model of ocean geochemistry (Canﬁeld 1998; Anbar and Knoll 2002). The new model of global oxygen history starts off like the old one: The ancient Earth was devoid of O2 , O2 stems from photosynthesis, and O2 started accumulating in the atmosphere about 2.3 billion years ago. So far so good. The big change concerns the time between ∼2.3 and ∼0.58 billion years ago, and it goes more or less like this: About 2.3 billion years ago, the O2 that was accumulating in the atmosphere started to oxidize continental sulﬁde deposits via weathering, carrying very substantial amounts of sulfate into the oceans, providing the substrate required for sulfate-reducing prokaryotes, hence marine biological sulfate reduction (BSR) (Poulton et al. 2004). Marine BSR then started going great guns on a global scale as evidenced by the sedimentary sulfur isotope record (Canﬁeld et al. 2000; Anbar and Knoll 2002; Johnston

Evolution of Eukaryotic Anaerobes

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et al. 2005). Marine BSR produces marine sulﬁde, and that sulﬁde did two things: it precipitated marine transition metals, and it kept the oceans largely anoxic. This means that the photic zone (surface water) was producing oxygen during this time, but below the photic zone, the oceans were anoxic and sulﬁdic (Shen et al. 2003; Hurtgen 2003; Arnold et al. 2004; Brocks et al. 2005). This condition—called Canﬁeld oceans or the intermediate oxidation state—remained more or less stable until ∼580 MY ago, when the deep ocean water also became oxic, Canﬁeld oceans came to an end, and the oceans started to look, geochemically speaking, much like they do today. Two relatively recent reports provide evidence suggesting the end of Canﬁeld oceans to have occurred ∼580 MY, at the same time as the ﬁrst animal macrofossils appear in the geological record (Fike et al. 2006; Canﬁeld et al. 2007), the interpretation being that oxygenation of the oceans allowed preexisting and diversiﬁed animal lineages to increase in size. Of course, a model entailing anoxic oceans up until about ∼580 MY has dramatic consequences for our understanding of metazoan evolution and for our understanding of anaerobic energy metabolism in protist and mitochondrial evolution, the topic of the next section.

4 Canfield Oceans Give Anaerobic Eukaryotes Room to Breathe If we accept the fossil evidence to indicate that eukaryotes are at least 1.45 Gyr old (Javaux et al. 2001), which this author does, then the majority of eukaryote evolution (about the ﬁrst two-thirds, at least) occurred at a time when the photic zone was oxic, but the rest of the oceans were anoxic and sulﬁdic. Under the new model of ocean geochemistry (Canﬁeld oceans), hydrogenosomes, mitosomes, and anaerobic eukaryotes in general would ﬁt quite naturally into the overall scheme of Earth and life history, without any kind of special explanation required to account for anaerobic lifestyles. We might assume that eukaryote evolution only occurred in the photic zone, but there is no reason to make such an assumption, because eukaryotes from many diverse lineages thrive today in anaerobic environments (Fenchel and Finlay 1995; Bernhard et al. 2000). Why should that have been any different in the past? What Canﬁeld oceans probably should be telling biologists, particularly those who work on anaerobic eukaryotes, is that globally widespread oxic (aerobic) marine environments are a very late arrival on the evolutionary scene, almost as late as life on land. In this sense, the ecological specialization to strictly oxic environments is a comparatively late evolutionary development, something that arose in the last ∼580 Myr. This statement stands in direct opposition to much traditional argumentation regarding the origin and evolution of mitochondria (de Duve 2007; Margulis et al. 2006; Cavalier-Smith 2004), so let’s contrast it to some readily available observations to see how it fares.

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We can start with vertebrates. Vertebrates are specialized to oxic environments, because their mitochondria are specialized to oxic environments. Vertebrates go back maybe 500 Myr in the fossil record (Benton and Donoghue 2007), so this makes sense. Vertebrates arose after the oceans were oxic and moved onto land at a time when they had evolved lungs and were able to crawl. Let’s move back in evolution just one single step more, into the invertebrates. The mitochondria of most marine invertebrates (worms, mollusks, and crustaceans) can also use oxygen, but can also endure sustained anoxia without any problems whatsoever, during which their mitochondria function anaerobically, excreting acetate, propionate, succinate, and CO2 as end products of mitochondrial ATP synthesis, without the participation of oxygen (reviewed extensively in Grieshaber et al. 1994). The biochemical steps involved in the synthesis of the main end products of mitochondrial ATP synthesis—succinate, propionate, acetete, and alanine (Schöttler and Bennet 1991; de Zwaan 1991; Grieshaber et a. 1994)—are the same in various freeliving marine invertebrates as those found in various parasitic worms that afﬂict humans and livestock (Tielens et al. 2002; van Hellemond et al. 2003). The steps involve fumarate reductase (FRD), rhodoquinone instead of ubiquinone at the FRD step (Tielens et al. 2002), and an acetate : succinate CoA transferase (ASCT) as occurs in hydrogenosomes (Müller 2003). The molecular identity of the hydrogenosomal ASCT has only just now been reported (van Grinsven et al. 2007). The methylmalonyl-CoA route to propionyl-CoA in anaerobic mitochondrial energy metabolism of invertebrates is nothing special, it is simply a reversal of the pathway used by the mammalian digestive tract to incorporate propionate into glycogen (Schöttler and Bennet 1991), and it is also used by Euglena mitochondria (Schneider and Betz 1985), which contain rhodoquinone as required for the FRD step (Hoffmesiter et al. 2004), during anaerobic respiration leading to synthesis of odd-chain fatty acids and alcohols during Euglena’s wax ester fermentation (Tucci et al. 2007). Succinate production in anaerobic metazoan mitochondria is not a process of substrate level phosphorylation, it involves rhodoquinone-dependent proton-pumping at the FRD step and chemiosmotic ATP synthesis (Tielens et al. 2002; van Hellemond et al. 2003), and could thus be described as fumarate respiration, the term commonly used when discussing energy metabolism in prokaryotes (Fenchel and Finlay 1995), although the term malate dismutation is also used (Tielens and van Hellemond 2007). The typical method for cultivating the marine invertebrates to characterize their anaerobic mitochondrial metabolism is to incubate them in N2 -saturated water over longer periods of time (Grieshaber et al. 1994). For very small metazoans (less than ∼2 mm in size; meiofauna) biochemical data about anaerobic metabolism are scarce at best, but it is known that many of them live in the anoxic layer of marine sediment (Nicholas 1991). The circumstance that mitochondria of marine invertebrates (metazoans, higher eukaryotes in any sense of the word) can readily do their job of synthe-

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sizing ATP for the animal without the help of oxygen is not very well known or well appreciated in the evolutionary community, even though the biochemistry and physiology of invertebrate anaerobic mitochondria has been studied in considerable detail. Grieshaber et al. (1994, p 64) wrote: “Meanwhile, details of the anaerobic energy metabolism have been elucidated in all major phyla of invertebrates and even in some insect species (Zebe 1991) which are usually considered as completely oxygen dependent”. Their citation of Zebe (1991) refers to a book entitled Metazoan life Without Oxygen, which contains excellent access to the literature on the topic; for example the chapter by Runnegar (1991) concludes with the sentence: “It was the rise of large, muscular and/or mineralized animals that marks the end of the Precambrian and the beginning of the Phaenerozioc, and the best explanation for their appearance may be a signiﬁcant increase in the oxygen content of the atmosphere”. That is almost exactly what Fike et al. (2006) and Canﬁeld et al. (2007) are now saying, except one would need to replace the word “atmosphere” with “ocean”. The widespread occurrence of anaerobic mitochondria among marine invertebrates might seem like an odd adaptation of some sort if we think that mitochondria arose because of oxygen, but it makes sense if we think that mitochondria arose at a time when there was little oxygen available and that the low availability of oxygen persisted throughout most of eukaryotic history in the oceans. Metazoan mitochondria that deal with low or no oxygen are easily interpreted as a preserved relic of Canﬁeld oceans within the metazoan lineage. If we dig a bit deeper into eukaryotic phylogeny and look at the fungi, for example, that are often thought of as the sister group to animals (Baldauf et al. 2000), then we already arrive at the anaerobic chytridiomycetes, a basal group of true fungi that inhabit anaerobic habitats and have fully ﬂedged hydrogenosomes (Yarlett et al. 1986; Yarlett 1994; Biagini et al. 1994; Boxma et al. 2004). The chytrid hydrogenosomes are anaerobic mitochondria that produce ATP without an electron transport chain (without chemisomosis) and that produce H2 . The main end products of ATP synthesis in hydrogenosomes of Piromyces sp. E2 are formate and acetate, with a comparatively small amount of H2 ; additional end products of energy metabolism are succinate, lactate, formate and ethanol produced in the cytosol (Boxma et al. 2004). The formate is produced by an enzyme called pyruvate : formate lyase (PFL), which is a comparatively rare metabolic enzyme among most eukaryotes studied so far, but the same PFL enzyme occurs in other chytridiomycetes and in the green alga Chlamydomonas reinhardtii (Gelius-Dietreich and Henze 2004; Atteia et al. 2006), suggesting the presence of PFL already in the eukaryote common ancestor, in light of current views concerning the position of the root in the eukaryotic tree (Embley and Martin 2006; Simpson et al. 2006). The ethanol produced by Piromyces is generated from acetyl-CoA by the action of bifunctional aldehyde/alcohol dehydrogenase (ADHE), which is found in a variety of anaerobic protists (Boxma et al. 2004), in particular many possessing mitosomes (Müller 2003), and which is also found in Chlamydomonas

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(Atteia et al. 2006) and its colorless chlorophyte relative, Polytomella sp. (Atteia et al. 2004). The presence of ADHE in the eukaryote common ancestor also seems likely (Atteia et al. 2004). The hydrogenase of chytrid hydrogenosomes is an iron-only hydrogenase ([Fe]-Hyd), the same type as occurs in hydrogenosomes of other anaerobic eukaryotes, and also in Chlamydomonas (Horner et al. 2002; Embley et al. 2003). The ATP-synthesizing enzyme of the chytrid hydrogenosome, succinate thiokinase (also called succinyl-CoA synthase), is an enzyme of the citric acid cycle and is the same one (Brondijk et al. 1996; Dacks et al. 2006) as is found in other hydrogenosomes and in mitochondria that have an electron transport chain (Lahti et al. 1994). Thus, the small repertoire of a few enzymes involved in anaerobic energy metabolism in chytridiomycete hydrogenosomes and their cytosol reveals nothing unique to the group, it entails the same enzymes that are found in other unicellular eukaryotes that deal with anaerobiosis permanently or on a regular basis, and as are also found in eukaryotes that produce oxygen. In light of Canﬁeld oceans and a predominantly anaerobic past, this should not be surprising in the slightest. The reader might wonder how Chlamydomonas could encounter anaerobiosis, because it is often regarded as a oxygenic producer, but it is a generalist that commonly inhabits soil (Sack et al. 1994) and avidly grows heterotrophically on acetate (Heifetz et al. 2001), a common currency of metabolic end product and carbon source among microbes (Wolfe 2005). The example of the chytrid hydrogenosomes uncovers many links of shared common ancestry for enzymes regarded as “characteristic” for eukaryotic anaerobes, but the same enzymes also occur among oxygen-adapted forms like Chlamydomonas. This makes sense in the context of Canﬁeld oceans. So if we look around in other eukaryotic groups, do we see anything really special among the anaerobes? In the trichomonads, the group where hydrogenosomes were discovered, the map of metabolism is slightly different from the chytrids, because the trichomonad hydrogenosomes studied so far possess pyruvate-ferredoxin oxidoreductase (PFO), an enzyme far more widespread among eukaryotic anaerobes (Müller 2003, see also chapter by Müller this volume) than PFL. The end products of energy metabolism in trichomonad hydrogenosomes are equimolar amounts of H2 , CO2 and acetate (Steinbüchel and Müller 1986). The H2 is generated by the same type of typical [Fe]-HYD as found in chytrids and Chlamydomonas (Horner et al. 2002). The same kind of PFO is found in some ciliate hydrogenosomes (Ellis et al. 1994) but also in the mitochondria of the photosynthetic protist Euglena (Rotte et al. 2001) and, at least as an expressed sequence in Chlamydomonas (Atteia et al. 2006). Some ciliate hydrogenosomes even still have a genome (Boxma et al. 2005). In anaerobic eukaryotes that perform all of their ATP synthesis in the cytosol, like Entamoeba or Giardia, both of which have mitosomes (Tovar et al. 1999, 2003; Mai et al. 1999), several of the enzymes otherwise found in mitochondria or hydrogenosomes, for example PFO and [Fe]-HYD (Müller 2003)

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have been relocalized to the cytosol. Anaerobic eukaryotes that perform all of their ATP synthesis in the cytosol, like Entamoeba or Giardia, tend to produce acetate and ethanol, or a mixture of both, as their main metabolic end products (Müller 2003), although Giardia also produces H2 under highly anaerobic conditions (Lloyd et al. 2002). The main enzymes involved are PFO and ADHE, which were discussed above as not being speciﬁc to the anaerobes, and an acetyl-CoA synthase (ADP-forming) (Sanchez et al. 2000), that, like PFL, seems to be a rare enzyme in eukaryote energy metabolism, but the corresponding gene also occurs in the malaria parasite Plasmodium falciparum (Sanchez et al. 2000), which uses the mitochondrial electron transport chain in some stages of the life cycle (Painter et al. 2007), so again there is no case to be made that the enzyme is genuinely speciﬁc to anaerobes. In some highly reduced parasitic (but not anaerobic) eukaryotes that possess mitosomes (Williams et al. 2002), it is not clear that the parasite actually produces its own ATP, so particularly among energy parasites, the presence of highly reduced mitochondria need not correlate with the anaerobic lifestyle. Although numerous functions link mitochondria and hydrogenosomes (see the chapter by Henze in this volume), the only function discovered so far that links mitosomes to mitochondria and hydrogenosomes is iron-sulfur cluster assembly (Tovar et al. 2003), which seems to be seen as evolutionarily signiﬁcant by most specialists in the ﬁeld, as discussed in other chapters in this volume. Iron-sulfur cluster assembly is a notoriously O2 -sensitive process, so it is not surprising that in oxygen-addicted vertebrates like humans, the process occurs in the mitochondrial matrix, which is generally oxygenpoor, similar to the situation in Bradyrhizobium, where N2 -ﬁxation in the cytosol takes place in the presence of active O2 -respiration at the surrounding plasma membrane (Preisig et al. 1996). To sum up this section, if we look around among the eukaryotic anaerobes that possess hydrogenosomes and mitosomes in search of enzymes or other biochemical attributes that set them distinctly apart from eukaryotes with mitochondria specialized to intermittently oxic or to permanently oxic environments, we come up with an empty set, hence the older ideas of that there are some eukaryotic lineages that diverged after a ﬁrst endosymbiosis entailing spirochaetes (ﬂagella) or some unspeciﬁed bacterium (peroxisomes) but before the origin of mitochondria should probably be abandoned; in 37 years no molecular evidence has come forth in support of either idea (Jekely and Arendt 2006; Gabaldon et al. 2006).

5 Anoxic, Fine, and What About Sulfidic? Canﬁeld oceans entail globally widespread anoxic and sulﬁdic marine environments, whereby the sulﬁde comes from biological sulfate reduction, the

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sulfate coming in turn from O2 -dependent weathering of continental sulﬁdes to sulfate following O2 -accumulation (Canﬁeld 1998; Poulton et al. 2004). From this it would follow that eukaryotes should also be able to deal with sulﬁde. This is also the case, and it is the mitochondria that do it. The mitochondria of many marine invertebrates can detoxify sulﬁde, and in some cases the electrons from sulﬁde oxidation are fed into the mitochondrial electron transport chain for ATP synthesis (Grieshaber and Völkel 1998; Doeller et al. 2001). The biochemical details of eukaryotic sulﬁde oxidation are still poorly understood (Yong and Searcy 2001; Theissen et al. 2004; Searcy 2006), but it is clear that the central enzyme for eukaryotic sulﬁde oxidation studied so far, sulﬁde:quinone oxidoreductase (SQR), transfers electrons from sulﬁde to a quinone acceptor in the inner mitochondrial membrane, similar to the situation in prokaryotes (Theissen 2006). Mitochondrial sulﬁde oxidation is conserved as a function from marine invertebrates to vertebrates (Yong and Searcy 2001; Theissen et al. 2003), and its presence among protists has only recently been reported (Searcy 2006), but the biochemical mechanisms involved are still mostly unknown. Eukaryotes are hardly running out of biochemical questions in terms of core energy metabolism and environmental interactions, a prime example being the recently discovered denitrifying forams (Risgaard-Petersen et al. 2006). If we consider the animals as gutless tubeworms that live at highly sulﬁdic hydrothermal vents (discussed, for example, in Vetter et al. 1991), then we see that there are many forms that depend upon sulﬁde and we are faced with the question: did they invent this basic biochemistry to handle sulﬁde de novo or is it a holdover from earlier times?

6 Conclusion The common ancestry of mitochondria, hydrogenosomes, and mitosomes does not seem to be an issue anymore, at least for those who work on the latter two manifestations of the organelle. But views concerning the origin, age, and polarity of the ability of eukaryotes to thrive in anaerobic habitats remains an issue of some debate. There are some who would possibly prefer the idea that the enzymes of anaerobic energy metabolism in eukaryotes are acquisitions via LGT in response to the adaptation to anaerobic habitats (Andersson et al. 2007), that the ancestral state of the eukaryotic lifestyle is aerobic, period (Cavalier-Smith 2004), or that eukaryotic anaerobes do not have mitochondria (Margulis et al. 2006; de Duve 2007). But if we actually look at the anaerobes, they are altogether normal eukaryotes, possessing no special sorts of attributes that would set them apart from the aerobes. If all of us had learned the biochemistry of mitochondria in college using the example of marine invertebrates, then we would see mitochondria as organelles that produce ATP under aerobic and anaerobic conditions. But we learned mi-

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tochondrial biochemistry with the example of rat liver mitochondria. Rats do not survive without oxygen, marine invertebrates do. More robust characters, like the presence or absence of genes and enzymes, the presence of organelles, or the intermingling of aerobic and anaerobic lineages, as for example among the ciliates (Embley et al. 1995) or higher eukaryotic groups (Embley and Martin 2006), suggest that there is no phylogenetic distinction at all between eukaryotic aerobes and eukaryotic anaerobes. Similarly, there is no phylogenetic distinction between eukaryotes that possess or lack ﬂagella (missing among many amoebae, all red algae, and most fungi), and no phylogenetic distinction between eukaryotes that possess or lack peroxisomes (typically lacking in anaerobes). That is, however, not to say that there are no phylogenetic distinctions among eukaryotes at all, for there clearly is a phylogenetic distinction between eukaryotes that possess or lack primary plastids, and the same is true for many other characters as well (see Adl et al. 2006 for a compilation). But for some obviously signiﬁcant eukaryotic characters, such as those shared among the excavate taxa (Simpson et al. 2006), the phylogenetic distinctions are not yet clear, whereby one should note that among the excavate taxa many aerobic and anaerobic lineages are found. In light of Canﬁeld oceans, eukaryotic anaerobes should hardly be surprising. The brunt of eukaryote evolution appears now to have occurred in anaerobic and sulﬁdic marine environments. What do evolutionary analyses of sequenced genomes say about these issues? A recent study by Pisani et al. (2007) comparing trees for all singlecopy genes in available sequenced genomes found that, at face value, the results were compatible with the predictions of only two among the dozens of competing theories for the origin of mitochondria: one entailing anaerobic hydrogen production by the ancestral mitochondrial endosymbiont and one entailing anaerobic sulfur metabolism of the ancestral mitochondrial endosymbiont. That need not mean that either model is correct, but it suggests that the anaerobic hydrogen and sulfur metabolism of mitochondria deserve further study, and that the larger context of geological history provides a good framework for such endeavors. Debate about the origin of mitochondria is still ongoing. With the recognition that all of the eukaryotes that were once thought to lack mitochondria do indeed have a form, the organelle or nuclear genes that betray its past presence, the issue of whether the origin of mitochondria and the origin of the eukaryotic lineage coincide to the same event stands as much in the foreground as ever (Davidov and Jurkevitch 2007; Martin et al. 2007). Some authors even think that the origin of mitochondria could have mechanistically precipitated the origin of the nucleus (Koonin and Martin 2006). The ﬁrst time this author stumbled across the new model of ocean geochemistry was 2002 through a review that appeared in Science (Anbar and Knoll 2002). It had a profound impact on my thinking about early evolution: many of the things that I had learned in college and always believed about

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oxygen and Earth geochemistry turned out probably to be wrong; there is an alternative model that accounts for the same observations relating to oxygen in Earth history more fully, and then some. This is exciting! Canﬁeld oceans have geologists excited, so much so that they are now writing about it in biological journals (Dietrich et al. 2006). The topic of Canﬁeld oceans has been the subject of geochemical contributions in Nature and Science at least 12 times since 1998 (Canﬁeld 1998; Canﬁeld et al. 2000, 2007; Anbar and Knoll 2002; Shen et al. 2003; Hurtgen 2003; Kah et al. 2004; Arnold et al. 2004; Brocks et al. 2005; Poulton et al. 2004; Johnston et al. 2005; Fike et al. 2006), about the same number as for hydrogenosomes and mitosomes during the same time. But there is still very little, if anything, in the way of visible interaction between geologists and biologists towards making joint progress on the issue of oxygen in eukaryotic evolution. Finally, it is of some interest to note that at the same time as Margulis was repopularizing endosymbiotic theory and as Cloud was forging the old model of oxygen accumulation in Earth history, and as people were pondering the idea that mitochondrial origins and oxygen might somehow correlate, Fenchel and Riedl (1970) published their seminal paper on the occurrence of a globally widespread anoxic and sulﬁdic ecosystem in marine sediments. In the decades since this time, the interactions between those studying anaerobic protists, those studying endosymbiosis, and those studying Earth history probably could have been closer. I hope that this chapter serves to promote interaction between geologists and biologists that leads to progress in understanding the largely anaerobic phase of eukaryotic evolution prior to ∼580 Myr ago.

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_105/Published online: 27 November 2007 © Springer-Verlag Berlin Heidelberg 2007

Protein Import into Hydrogenosomes and Mitosomes Sabrina D. Dyall1 (u) · Pavel Dolezal2 1 School

of Biomedical Sciences, University of Nottingham Medical School, Queen’s Medical Centre, Nottingham NG7 2UH, UK [email protected]

2 Department

of Biochemistry and Molecular Biology, Bio21 Institute, University of Melbourne, 3010 Parkville, Australia

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2.1 2.2 2.3

Protein Trafﬁcking in Eukaryotes The Nucleus . . . . . . . . . . . . The Endoplasmic Reticulum . . . The Mitochondrion . . . . . . . .

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Evolution of the Mitochondrial Protein Import Machinery . . . . . . . . .

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4 4.1 4.2

Studying Hydrogenosomal and Mitosomal Protein Import . . . . . . . . . Laboratory Techniques and Tools . . . . . . . . . . . . . . . . . . . . . . . Mining Genome Sequence Data . . . . . . . . . . . . . . . . . . . . . . . .

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5 5.1 5.2 5.2.1 5.2.2 5.2.3 5.2.4 5.2.5 5.2.6 5.3

Organellar Targeting Signals . . . . . . . . . . . . . . . . . . . . . . . . . . Mitochondrial Targeting Signals . . . . . . . . . . . . . . . . . . . . . . . . Signals on Precursors of Soluble Hydrogenosomal and Mitosomal Proteins Trichomonas Hydrogenosomes . . . . . . . . . . . . . . . . . . . . . . . . . Neocallimastix Hydrogenosomes . . . . . . . . . . . . . . . . . . . . . . . . Cryptosporidium Mitosomes . . . . . . . . . . . . . . . . . . . . . . . . . . Entamoeba Mitosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Giardia Mitosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Microsporidian Mitosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . Signals on Hydrogenosomal and Mitosomal Membrane Proteins . . . . . .

34 34 35 35 41 42 42 43 44 45

6 6.1 6.1.1 6.1.2 6.2 6.3 6.3.1 6.3.2 6.3.3

Crossing the Organellar Membranes . . The Outer Membrane . . . . . . . . . . . The General Import Pore . . . . . . . . . Sorting and Assembling β-Barrel Proteins The Intermembrane Space Chaperones . The Inner Membrane . . . . . . . . . . . The TIM22 Complex . . . . . . . . . . . The TIM23 Complex . . . . . . . . . . . The PRAT Family . . . . . . . . . . . . .

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The Protein Import Motor . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Preprotein Processing Peptidases . . . . . . . . . . . . . . . . . . . . . . . The Mitochondrial Processing Peptidase . . . . . . . . . . . . . . . . . . . The Inner Membrane Protease . . . . . . . . . . . . . . . . . . . . . . . . .

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Folding Newly Imported Soluble Proteins . . . . . . . . . . . . . . . . . .

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Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract In the past decade, studies on protein targeting to hydrogenosomes and mitosomes have revealed several characteristics in common with mitochondrial protein targeting. Proteins from one system can readily be imported into another, strongly suggesting that targeting signals on hydrogenosomal, mitosomal and mitochondrial preproteins are conserved. By extension, these observations, together with the proposed common origin of hydrogenosomes, mitosomes and mitochondria, led to the proposition that components of the respective protein import machineries for these organelles are conserved. With the advent of complete genome sequence databases for diverse eukaryotes, we are now in a better position to examine this proposition. In this review, we report and integrate the latest experimental and bioinformatics data on the state of protein import in hydrogenosomes, mitosomes and mitochondria.

1 Introduction Eukaryotic cells have internal membranes deﬁning sub-cellular compartments, or organelles, each of which has discrete metabolic and/or biosynthetic functions. To fulﬁl such functions, speciﬁc sets of proteins must be precisely targeted to, and quantitatively imported and localized within these organelles in a timely fashion. Of the approximately 5000 proteins encoded by the nuclear genome of Saccharomyces cerevisiae, about 1000 proteins are targeted to the endoplasmic reticulum (ER), and from there subsequently localized throughout the endomembrane system. A similar number of proteins are targeted to mitochondria, while the rest (∼ 3000 proteins) are folded in the cytoplasm, with some of them retargeted to the nucleus, or to peroxisomes. In general, a sophisticated system of membrane translocases with associated propelling machines recognizes the address on individual protein molecules. In most cases, trafﬁcking of preproteins is fuelled by the hydrolysis of either ATP or GTP. Additionally, proteins travelling through or into mitochondrial inner membranes require an electrochemical membrane potential generated by the mitochondrial electron transport chain (Wickner and Schekman 2005). Despite these general similarities, the respective mechanisms responsible for protein trafﬁcking to the nucleus, the ER and the mitochondrion differ fundamentally, with each employing distinct molecular machines. While the molecular machines operating in the ER were recruited during the evolution of the eukaryotic cell from an ancestral prokaryote, and adapted towards current needs, nuclear and mitochondrial protein import systems seem to have been almost entirely created de novo by the eukaryotic cell (Dyall et al. 2004a; Wickner and Schekman 2005; Dolezal et al. 2006; Stewart 2007).

Protein Import into Hydrogenosomes and Mitosomes

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Some eukaryotes lack bona ﬁde mitochondria but have unusual organelles such as hydrogenosomes in Trichomonas and Nyctotherus, and mitosomes in Entamoeba, Giardia, Cryptosporidium and microsporidia. Despite their different morphologies and several non-mitochondrial functional pathways, hydrogenosomes and mitosomes contain a subset of mitochondrial-type proteins. Phylogenetic analyses of these proteins provide strong evidence that mitochondria, mitosomes and hydrogenosomes originate from a single eubacterium that formed an endosymbiotic association with a eukaryotic or pre-eukaryotic cell over two billion years ago (Embley 2006). Critical events during the conversion of the mitochondrial endosymbiont into an organelle would have involved endosymbiotic gene transfer to the host nucleus, and the concomitant evolution of an organellar apparatus to import nuclearencoded proteins. Recent studies have demonstrated that there are several compatible mechanistic and structural features between protein import into hydrogenosomes, mitosomes and mitochondria (Burri and Keeling 2007). These observations suggest that the molecular machines that import proteins into these organelles are of a similar nature and/or origin. In this chapter, we shall examine what is known about mitochondrial, hydrogenosomal and mitosomal protein import, and shall discuss the evolution of protein import mechanisms in these organelles in relation to eubacterial and other organellar systems.

2 Protein Trafficking in Eukaryotes Protein trafﬁcking has been extensively studied in fungi and mammals, and a number of elaborate machines have been described that speciﬁcally import certain proteins into typical eukaryotic organelles such as the nucleus, the endoplasmic reticulum and the mitochondrion. 2.1 The Nucleus The nuclear envelope is perforated with huge macromolecular assemblies of ∼30 different proteins that form nuclear pore complexes with a central channel of 25–30 nm in diameter. This channel allows proteins smaller than 30 kDa to passively traverse the outer and inner nuclear membranes. Larger proteins are actively transported across the nuclear envelope and contain nuclear localization signal (NLS) sequence motifs. These signals consist of one or two clusters of four or ﬁve basic residues localized usually within the polypeptide chain. The import of proteins with NLS through the channel is facilitated by the carrier heterodimer of importin-α : β (Gorlich and Kutay 1999; Pemberton and Paschal 2005). Upon passing through the nuclear pore, the interaction of the complex

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with RanGTP initiates the release of cargo protein from the importins. The whole process of translocation is regulated by the nucleotide state of Ran, which accordingly cycles between the nucleus and the cytoplasm (Stewart 2007). 2.2 The Endoplasmic Reticulum In contrast to cytosolic, nuclear and most mitochondrial proteins that are synthesized on free ribosomes, mRNA transcripts encoding ER-destined proteins are translated on ribosomes tightly bound to the ER membrane. Nascent luminal proteins are equipped with an N-terminal signal sequence that consists of a basic amino-terminus followed by a stretch of 8–14 non-polar residues, and a cleavage motif for the signal peptidase (Blobel and Dobberstein 1975a,b; von Heijne 1990). Membrane proteins usually contain internal topological signals instead of the N-terminal signal peptide. Translation and translocation are coordinated by the signal recognition particle (SRP), a complex of 7S RNA and six protein subunits (Pool 2005). Initially, SRP binds the signal peptide emerging from the ribosome. Translation slows down until SRP is recognized by its ER-bound receptor, whereupon translocation can resume following the binding of GTP to both SRP and its receptor. The passage through the membrane is formed by the Sec61 translocon consisting of a Sec61α channel and two accessory subunits β and χ (Wickner and Schekman 2005). According to studies on the orthologous archaeal SecYEG complex, the actual membrane translocase is a dimer of two Sec complexes allowing both the translocation of soluble proteins across the membrane, and the lateral insertion of membrane proteins (Mitra et al. 2005). The passage of substrate protein through the channel is then driven by the action of lumenal Hsp70 (Osborne et al. 2005). Analogously, in bacteria, the SecYEG translocon is used for secretion of proteins across the plasma membrane, and the signal peptides of secreted proteins share similar characteristics with ER proteins (Wickner and Schekman 2005). While protein import into ER requires nucleotide triphosphates, bacteria need an additional membrane electrochemical gradient to export proteins across the plasma membrane. After translocation into the ER, the signal peptide is cleaved from the precursor polypeptide by the signal peptidase. This step is necessary for releasing the protein from the membrane lipid bilayer to which it is bound via the hydrophobic signal peptide. The ER signal peptidase is a membrane protein that shares ancestry with both the bacterial signal peptidase and the inner membrane protease complex in mitochondria (Dalbey et al. 1997). 2.3 The Mitochondrion About 99% of the mitochondrial proteins are nuclear-encoded and are synthesized in the cytosol, from where they are imported into mitochondria

Protein Import into Hydrogenosomes and Mitosomes

25

(Pfanner and Geissler 2001). Protein import into mitochondria is mostly posttranslational, although recent data suggest that mRNA sub-populations are organized in the proximity of the mitochondrial outer membrane (GarciaRodriguez et al. 2007). Besides having to be targeted to the organelle, these proteins have to be internally sorted to either of four distinct subcompartments: the outer membrane, the intermembrane space (IMS), the inner membrane and the mitochondrial matrix. Proteins destined to mitochondria possess N-terminal and/or internal targeting signals that ensure their correct delivery to the organelle. A large majority of mitochondrial proteins are synthesized with N-terminal cleavable presequences of variable length, ranging from 10 to 80 amino acids (Pfanner and Geissler 2001). The presequence is not generally conserved at the primary sequence level among the different preproteins, but it is rather the α-helical structure engaged by the presequence upon interaction with the outer membrane receptor Tom20 that appears to be the common factor (Schatz and Dobberstein 1996; Abe et al. 2000). This α-helix is amphipathic, containing patches of positively charged and hydrophobic amino acids, respectively, on opposite surfaces of the theoretical cylinder. The presequence is usually processed by the mitochondrial processing peptidase (MPP) and the mature protein is sorted to either the matrix, or to the inner membrane if it bears a hydrophobic stop-transfer sequence. Some mitochondrial proteins, mostly destined to the membranes, do not have cleavable N-terminal presequences but have internal targeting signals that are not well characterized (Pfanner and Geissler 2001). Translocation across mitochondrial membranes is carried out by several molecular machines. These machines consist of core transmembrane translocases complemented by additional components that provide speciﬁcity. Such a modular structure enables the independent evolution and function of each molecular machine (Dolezal et al. 2006). The translocase of the outer membrane (TOM) complex constitutes the central recognition point and gate for all nuclear-encoded mitochondrial proteins (Fig. 1). Tom70 and Tom20 are receptor subunits that recognize the precursor proteins and release them subsequently into the translocation channel. This transfer is assisted by Tom22, which together with Tom40 and Tom5, represent the core and essential part of the TOM complex (Meisinger et al. 2001). Two other small proteins, Tom6 and Tom7 participate in the maintenance of the complex. The translocation pore is formed by several subunits of Tom40, which is most likely a β-barrel protein (Hill et al. 1998). After passing through the TOM complex, preproteins may interact with either of three distinct molecular machines (Fig. 1), depending on their ﬁnal destination. The insertion and assembly of β-barrel outer membrane proteins, including Tom40, are assisted by the sorting and assembly machinery (SAM) complex (Fig. 1). The SAM complex consists of four subunits, the core translocase Sam50, which is itself a putative β-barrel protein (Kozjak et al. 2003; Paschen et al. 2003; Gentle et al. 2004), and the additional proteins Sam35,

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S.D. Dyall · P. Dolezal

Fig. 1 Mitochondrial protein import machinery as deﬁned in S. cerevisiae. TOM translocase of the outer mitochondrial membrane; SAM sorting and assembly machinery; TIM translocase of the inner mitochondrial membrane; MIA mitochondrial IMS import and assembly machine; PAM presequence translocase associated motor; IMP inner membrane protease; MPP mitochondrial processing peptidase. The numbers on the individual Tom, Sam, Tim or Pam components represent their approximate molecular masses in kDa. See text for mechanistic details. Adopted from Dolezal et al. 2006

Sam37 and Mdm10 (Bohnert et al. 2007). Recently, a distinct complex of Mdm10/Mdm12/Mmm1 has been shown to function in β-barrel biogenesis downstream of the SAM core complex, but the exact role of all these components is still unclear (Meisinger et al. 2007). The structure of β-barrel precursors does not allow their lateral insertion into the lipid bilayer directly from the TOM channel, and the precursors must ﬁrst be released into the IMS. The passage of β-barrel precursors from the TOM complex to the SAM complex is assisted by the so-called small translocase of the inner membrane (TIM) chaperone complexes (small Tims). These soluble complexes are trimeric assemblies of either Tim9/Tim10 heterodimers, or of Tim8/Tim13 heterodimers, and protect the exposed hydrophobic epitopes of some membrane proteins from the aqueous environment of the IMS (Hoppins and Nargang 2004; Koehler 2004a; Wiedemann et al. 2004). Precursors destined for the IMS have a sorting signal at the N-terminus. The inner membrane protease (IMP) complex is responsible for the maturation of these proteins. IMP comprises the two proteases Imp1 and Imp2, and the regulatory subunit Som1. Some of the precursors contain bipartite presequences consisting of a matrix-targeting signal followed by an IMS-sorting signal (Gakh et al. 2002). Recently, a speciﬁc pathway for the import and assembly of a subset of IMS preproteins has been discovered in S. cerevisiae. The mitochondrial IMS import and assembly (MIA) machinery consist of

Protein Import into Hydrogenosomes and Mitosomes

27

IMS components Mia40, Erv1 and Hot13p (Fig. 1). Substrates for this pathway are precursors that contain one of the cysteine motifs that are characteristic of some IMS proteins, such as the twin CX3 C motif in the small Tims (Mesecke et al. 2005; Gabriel et al. 2007). According to the proposed model, Mia40 binds the precursors in the IMS via disulﬁde bridges, thereby trapping them after their entrance through the Tom40 pore. Further isomerization of disulﬁde bridges releases the precursors from Mia40, which is subsequently oxidized by Erv1 (Mesecke et al. 2005). In this process, Hot13p might perform a reducing action on the precursors (Curran et al. 2004). Some precursors that are to be integrated in the inner membrane, such as the mitochondrial carrier family (MCF) proteins, are inserted by the TIM22 machine (Fig. 1), which is built around a Tim22 subunit and contains another two membrane-integral subunits Tim54 and Tim18 (Rehling et al. 2003a). The precursors are shuttled from the TOM to the TIM22 complex by the Tim9/Tim10 chaperone through the IMS. Tim12, peripherally associated with TIM22, serves as a docking site for the Tim9/Tim10 complex that detaches from the precursor upon contact with the TIM22 complex (Koehler 2004a). Soluble matrix-destined preproteins, usually synthesized with a cleavable N-terminal presequence, are passed from the TOM complex to the distinct TIM23 machine through interaction with the IMS domain of Tim50 (Fig. 1). In yeast, TIM23 consists of Tim23, Tim17, Tim50, and Tim21 (Bohnert et al. 2007). Tim23 forms a protein-conducting channel that is regulated by the action of Tim50, and Tim17 (Martinez-Caballero et al. 2006; Meinecke et al. 2006). The TIM23 complex is also capable of inserting preproteins into the inner mitochondrial membrane (Koehler 2004a). These inner membrane preproteins are synthesized with a short hydrophobic sorting sequence downstream of the N-terminal presequence (Glick et al. 1992; Stuart 2002). To discriminate between, and to coordinate the dual translocation and insertion activities, TIM23 either interacts with Tim21, or associates with the presequence translocase associated motor (PAM) complex (Fig. 1). This cycle is regulated by Tim17, which recruits the PAM complex to the TIM23 complex (Chacinska et al. 2005). The PAM complex, together with the membrane potential, drives translocation of the precursor through the TIM23 complex (Rehling et al. 2003a). First, the negatively-charged matrix face of the inner membrane generates an electrophoretic force on the predominantly positively-charged presequence and mediates preprotein insertion into the Tim23 channel. Consequently, the PAM complex completes the translocation of the bulk polypeptide in an ATP-dependent manner (Voos and Rottgers 2002). The central component of the PAM complex is the molecular chaperone Hsp70 (Fig. 1). The activity of Hsp70 is regulated by two inner membrane, J-domain containing proteins Pam18 and Pam16, and the soluble nucleotide exchange factor Mge1. The PAM complex is tethered to the translocation channel, probably by the peripheral membrane protein Tim44. Upon translocation into the matrix, the N-terminal presequence of precur-

28

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sors is processed by MPP and the mature protein is thereafter folded into its native conformation. Some preproteins with an octapeptide-containing presequence require sequential processing of the targeting presequence by MPP followed by the mitochondrial intermediate peptidase (MIP), which removes the octapeptide sequence (Gakh et al. 2002). After being driven in by the PAM complex and having their presequence processed, some newly-imported matrix proteins require further assistance from molecular chaperones. First, soluble Hsp70 and its co-chaperone Mdj1 accost the substrate proteins to partially fold them. Two populations of Hsp70 thus exist in mitochondria: (1) a Tim44-bound membrane-associated form serving as the protein import driver and (2) a matrix-soluble chaperone that assists protein folding (Horst et al. 1997). After release from Hsp70, the protein is passed along to an Hsp60/Hsp10 system, which is the major chaperone system for protein folding in the matrix (Manning-Krieg et al. 1991). A homooligomer of Hsp60 provides a protected cavity for protein folding, while Hsp10 regulates the ATPase cycle of Hsp60 and the behaviour of individual subunits (Martin et al. 1991a).

3 Evolution of the Mitochondrial Protein Import Machinery Mitochondria are of endosymbiotic origin and have descended from eubacteria. Extensive sequence analyses have shown that mitochondria form a monophyletic group, and have demonstrated strong afﬁnities between mitochondrial genomes and present-day α-proteobacterial genomes, particular those of the Rickettsiales (Andersson et al. 1998; Gray et al. 1999). Consequently, the endosymbiotic theory for the origin of mitochondria purports that the mitochondrion originates from a single eubacterium, possibly an ancestor of Rickettsia, which formed a symbiotic relationship with a preeukaryotic or a primitive eukaryotic cell around two billion years ago. Over time, the endosymbiont lost its capacity to function and reproduce as an independent organism, and its fate was sealed within the host as it transferred the bulk of its genome to the host nucleus, or simply discarded some of it. The possible reasons or driving forces behind the symbiosis, and the subsequent loss of the endosymbiont genome and how that happened, are beyond the scope of this chapter and are comprehensively covered in Chap. 2 (this volume), and in reviews by Adams and Palmer (2003) and by Timmis et al. (2004). The outcome of, or perhaps the support for, the endosymbiont transferring its genes to the nucleus was the evolution of new machinery in the eukaryotic cell to send the nuclear-encoded proteins back to the degenerate endosymbiont to allow the latter to function. Moreover, it is of note that the large majority of extant mitochondrial proteins are not of endosymbiotic or α-proteobacterial origin. These proteins have either been recruited

Protein Import into Hydrogenosomes and Mitosomes

29

from other eubacterial sources, or have been invented de novo by the evolving eukaryote (Andersson et al. 2003; Gabaldon and Huynen 2003, 2004). All these proteins would have had to develop targeting signals whilst the eukaryote was inventing a new machine, either from scratch or by tinkering existing protein targeting components, to intake nuclear-encoded precursors into the proto-mitochondrion. Of the ﬁve protein import machines characterized to date in yeast mitochondria, namely TOM, SAM, TIM22, TIM23 and MIA, only the SAM complex bears a component, Sam50, that is clearly related to a bacterial translocase (Gentle et al. 2004, 2005). Thus, the majority of translocase components are a product of eukaryotic invention. Much can be inferred about the evolution of mitochondrial biogenesis by examining these translocases and their features, as we discuss later in this chapter. It is of particular interest whether hydrogenosomes and mitosomes use phylogenetically similar translocases as mitochondria. The majority of mitochondrial proteins have an N-terminal presequence that is both necessary and sufﬁcient to target a passenger protein to mitochondria (von Heijne et al. 1989). How and when these presequences were initially acquired, and how they have been distributed to genes on different loci are intriguing questions. It has been demonstrated that synthetic mitochondrial presequences can penetrate either artiﬁcial or bacterial lipid bilayers (Roise et al. 1986; Maduke and Roise 1993; Neupert 1997). It is therefore plausible that presequences were developed prior to the existence of speciﬁc receptors or pores on the outer surface of the proto-mitochondrion, and that the latter were later evolved to enhance the efﬁciency of translocation. Hermann (2003) proposed that a primitive PAM might have driven the presequence and the mature polypeptide into the proto-mitochondrion in the absence of any TOM or TIM component. Sequences with presequencelike features are commonly found in genomes, and the odds of a transferred gene landing in such a locus may have been quite good (Baker and Schatz 1987; Lucattini et al. 2004). Since some presequences are distributed over several exons in some nuclear-encoded mitochondrial genes, exon shufﬂing and alternative splicing have been proposed as mechanisms for presequence generation (McFadden 1999). Much insight on these aspects has been gained by studying mitochondrial gene transfer processes in ﬂowering plants (Adams et al. 2000; Adams and Palmer 2003). In those species, mitochondrial gene transfer is an ongoing process, such that functional copies of some genes for mitochondrial proteins can be found in: (1) both the nucleus and the mitochondrion of one species, or (2) in the nucleus of one species, but only in the mitochondrion of a sister species. Thus, the changes required to target the newly transferred genes to the mitochondrion can be examined. Productive gene transfer not only involves the evolution of a targeting signal, but is primarily dependent on the acquisition of gene expression signals. It was shown that mitochondrial copies of

30

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freshly transferred genes remain active for some indeterminate period and do not immediately get shut off (Choi et al. 2006). Transferred genes can acquire presequences by simply integrating into the locus for a duplicated nuclearencoded mitochondrial gene (Sandoval et al. 2004; Murcha et al. 2005b; Choi et al. 2006). Curiously, some mitochondrial presequences have independently been acquired from the same donor gene, e.g. from mt-hsp70 (Adams et al. 2000; Choi et al. 2006). Genes for non-mitochondrial proteins have also acted as donors for gene control regions and for fortuitous presequence-like stretches (Murcha et al. 2005b). Or sometimes, the transferred gene does not gain a presequence but uses internal signals from the mature polypeptide (Murcha et al. 2005b; Choi et al. 2006). Additionally, changes in local hydrophobicity in the protein sequence of certain transferred genes have been implied in enhancing protein import (Daley et al. 2002). Therefore, a plethora of tricks exists to append presequences to, or to create internal signals within, mitochondrial proteins and, though the precise mechanisms are shady, it appears that these tricks occur repeatedly and independently.

4 Studying Hydrogenosomal and Mitosomal Protein Import It has been hypothesized that the process of inventing a protein import machine for mitochondria would have been so intricate and critical that it is unlikely to have occurred more than once (Cavalier-Smith 1987). By extension, similarities found between mitosomal and hydrogenosomal and mitochondrial protein import have been presented as strong support that all these organelles use common components for import, and are therefore one and the same. These observations have prompted a number of experimental and bioinformatics studies to shed light on the constitution and evolution of the protein import machineries of hydrogenosomes and mitosomes. 4.1 Laboratory Techniques and Tools Unfortunately, a limited set of tools is available to study the hydrogenosomal or mitosomal species. In yeast, much has been deduced about mitochondrial biogenesis through extensive genetic manipulation, and a variety of mutants can readily be obtained that can be used to assess the function of individual components in the biogenetic pathway (Bonnefoy et al. 2007). These studies are complemented with a wealth of highly-honed biochemical techniques such as in organello import, membrane separation, creation of mitoplasts, generation of protein import intermediates, to list but a few (Stojanovski et al. 2007). Nonetheless, these techniques provide a basis for developing new methods to study protein import in hydrogenosomal or mitosomal species.

Protein Import into Hydrogenosomes and Mitosomes

31

Among the species under study, T. vaginalis is currently the most experimentally tractable. It can be genetically transformed to express endogenous or exogenous proteins (Delgadillo et al. 1997), or to delete genes by homologous recombination (Land et al. 2004). Of all the mitochondria-related organelles, only T. vaginalis hydrogenosomes (∼ 0.8 µm in diameter) have been isolated to high purity, on a Percoll gradient, and have been demonstrated to be protein-import competent (Bradley et al. 1997). The development of an assay for importing precursor proteins into isolated T. vaginalis hydrogenosomes has revealed several requirements that appear to be in common with mitochondrial protein import (Bradley et al. 1997). For this assay, recombinant precursor proteins are either metabolically radiolabelled in Escherichia coli for detection by autoradiography, or puriﬁed with a Cterminal hexahistidine (His6) tag for detection by Western analysis. The isolated organelles and the precursor are incubated in an isotonic import buffer supplemented with ATP and crude T. vaginalis cytosol. In mitochondrial import systems, the precursor protein is synthesized and radiolabelled in rabbit reticulocyte lysate or in wheat germ extract; these extracts contain cytosolic chaperones that support mitochondrial protein import. However, hydrogenosomal protein import is absolutely dependent on T. vaginalis crude cytosol that cannot be substituted for by either rabbit reticulocyte lysate or wheat germ extract (Bradley et al. 1997; our unpublished data), emphasizing the speciﬁcity of the cytosolic factor(s). Successful import is measured as follows: resistance of the imported protein to externally added protease and presequence cleavage, as detected by faster electrophoretic migration on SDS-PAGE. The import of a matrix precursor protein, ferredoxin, has been shown to be linear and saturable, and dependent on a protease-sensitive component(s) on the outer hydrogenosomal surface, indicating the presence of a speciﬁc receptor (Plumper et al. 2000). Precursor ferredoxin import has been shown to be dependent on the presence of a speciﬁc presequence, and on ATP, a weak electrochemical potential and temperature (Bradley et al. 1997), which are all requirements for import into mitochondria (Schleyer et al. 1982). Import studies have also been carried out to explore the functional conservation of import pathways between hydrogenosomes and mitochondria (Dyall et al. 2000, 2003), or between hydrogenosomes and mitosomes (Dolezal et al. 2005). The second most studied hydrogenosomal species is Neocallimastix, although not many techniques are available for thorough studies. An enriched hydrogenosomal fraction can be prepared by differential centrifugation of disrupted cells of this species, and can be used for rudimentary sub-organellar fractionation studies (Marvin-Sikkema et al. 1993). However, most of the studies on Neocallimastix sp. hydrogenosomal proteins have been done in heterologous fungal systems (van der Giezen et al. 1998, 2002, 2003). A high-speed differential centrifugal fraction for Giardia mitosomes has been generated that has been successfully used to reconstitute Fe–S cluster

32

S.D. Dyall · P. Dolezal

formation (Tovar et al. 2003). Further enrichment of these fractions was obtained on sucrose gradients that yielded organelles of ∼ 150 nm in diameter (Dolezal et al. 2005). These organelles have been used for localization studies, but there has been no report of a successful protein import assay as yet. The proteome of mitosomes is far from complete and only a few proteins have been physically localized in the mitosomes by speciﬁc polyclonal antibodies, or by detection of their tagged recombinant versions. Genetic transformation techniques exist for both E. histolytica and G. intestinalis, and these have been applied to localize mitosomal proteins, and to investigate putative targeting signals (Mai et al. 1999; Tovar et al. 1999; Dolezal et al. 2005; Regoes et al. 2005). Ribozyme-based RNA silencing has also been developed for G. instestinalis, although the technique has not been widely applied in the ﬁeld (Dan et al. 2000). Microsporidia and Cryptosporidium cannot be genetically manipulated as yet, and putative mitosomal proteins and associated targeting signals have been investigated in fungi (Slapeta and Keithly 2004), and/or in the apicomplexan species Toxoplasma gondii (Slapeta and Keithly 2004; Lagier et al. 2003). The likelihood of mitosomes ever being puriﬁed from these organisms is not very high as the organelles are extremely small, ranging from 70 nm for the microsporidian Trachiopleistosphora (Williams et al. 2002) to 150–300 nm for the C. parvum mitosome (Riordan et al. 2003; Putignani et al. 2004). Moreover, the single C. parvum mitosome is entangled by the rough ER (Riordan et al. 2003; Putignani et al. 2004), which will render any disruption technique quite tricky. Despite these limitations, recent studies of protein import into mitosomes of G. intestinalis (Dolezal et al. 2005; Regoes et al. 2005), E. histolytica (Mai et al. 1999; Tovar et al. 1999) and of the two microsporidian species Encephalitozoon and Antonospora (Nosema) locustae (Burri et al. 2006) brought exciting insight into the degree of functional similarity between mitosomes, hydrogenosomes and mitochondria, and also into the degree of adaptation of mitosomes within the microsporidia (Burri and Keeling 2007). Although mitosomes are likely to have arisen independently and repeatedly, and in the case of microsporidia and Entamoeba, are most probably reduced forms of a more complex mitochondrial compartment, the molecular basis of the reduced mitosomal protein import machinery may offer clues as to the composition of the original sets of translocases installed in the membranes of proto-mitochondria. 4.2 Mining Genome Sequence Data Thanks to the recently completed, or to the ongoing genome sequencing projects for hydrogenosomal and mitosomal species (McArthur et al. 2000; Katinka et al. 2001; Abrahamsen et al. 2004; Xu et al. 2004; Loftus et al. 2005;

Protein Import into Hydrogenosomes and Mitosomes

33

Carlton et al. 2007), we have a tremendous amount of information about the biology and evolution of these organisms at hand. Having all these data available, we are presented with the difﬁcult issue of efﬁcient data mining. Our attempts to identify possible homologous sequences in the genomes of evolutionary diverse species are very often faced with the danger of false negative results, and therefore of incorrect conclusions. The widely used approach is BLAST, based on pair-wise sequence analyses (Altschul et al. 1990, 1997). BLAST searches are sometimes inefﬁcient simply because a particular query may be too divergent to pick the target sequence from genome databases. In the ﬁeld of mitochondrial protein import, queries originate primarily from S. cerevisiae or other fungal sequences. While pair-wise sequence analyses were sufﬁcient to identify equivalent components in animals, they work less well on plants, and often fail to identify homologous sequences in other phylogenetic groups, especially protists (Hoogenraad et al. 2002). If the whole family of proteins instead of a single sequence is available, a search based on the hidden Markov model (HMM) offers a signiﬁcantly more sensitive mining method when compared with BLAST (Krogh et al. 1994; Eddy 1996, 1998). In practice, analyses based on HMMs represent a reversed search of the protein family (PFAM) database (Bateman et al. 2004). Instead of comparing one query sequence with all the available HMMs in PFAM, a single HMM is used to search the genome database. Although HMMs were ﬁrst designed for speech recognition, they can be applied to a variety of problems, where hidden parameters need to be determined from obvious parameters, such as sequence alignment of homologous proteins. The parameters that are extracted from the sequence alignment, for instance the probability of occurrence of certain amino acids in a particular position, can be then used for mining data from the conceptual translation of a genome sequence. Depending on the selection of sequences for the alignment, HMMs can even pick structural information otherwise hidden in the primary sequence (Dolezal et al. 2006). For example, the alignment of some homologous β-barrel proteins might provide enough information to ﬁnd any β-barrel protein in the examined genome. Importantly, newly identiﬁed homologous sequences can be included into the alignment used for the building of new HMM, thus providing a reﬁned and more sensitive tool for the next round of searches. Usually several cycles of reﬁnement are used to craft a reliable HMM that is powerful enough to pick very divergent homologous sequences, but that is insensitive to unrelated sequences. The freely available HMMER software (http://hmmer.janelia.org/) enables the building of tailored HMMs based on the user’s protein sequence alignment. However, efﬁcient searches using single or several HMMs in a large database are time-consuming, and certainly a challenge for the typical bench-work biologist. Collectively, in silico studies and laboratory studies have begun to reveal much about hydrogenosomal and mitosomal targeting signals, translocases, chaperones and processing peptidases.

34

S.D. Dyall · P. Dolezal

5 Organellar Targeting Signals Targeting signals contain the minimal information necessary for a protein precursor to be recognized by targeting machinery, and to be directed to the correct compartment in a cell. These sequences are both “necessary and sufﬁcient” to target a passenger protein to a given organelle. Other sequence stretches within the precursor may be necessary to target the protein to the correct sub-organellar location (Neupert 1997). Several categories of targeting signals have been deﬁned for mitochondrial precursors, and to a lesser extent, for hydrogenosomal and mitosomal precursors. 5.1 Mitochondrial Targeting Signals The majority of mitochondrial precursors are synthesized with N-terminal cleavable presequences. Typical presequences contain 10–80 amino acid residues, many of which are positively charged, hydrophobic and hydroxylated (Pfanner and Geissler 2001). Negatively charged amino acid residues are notoriously absent in most presequences (von Heijne et al. 1989). Generally, the amino acid residues on presequences are disposed in an amphipathic α-helix, which has one hydrophobic surface opposed by a positively charged surface (Roise et al. 1986; Abe et al. 2000). These contrasting surfaces make contact sequentially with Tom and Tim translocases as the presequencecontaining precursor traverses both mitochondrial membranes (Pfanner and Geissler 2001). For instance, structural studies have shown that the hydrophobic surface of the α-helical presequence makes contact with the Tom20 binding groove (Abe et al. 2000). Subsequently, the precursor is passed over to Tom22 through interaction of the positive surface of the α-helix with negative charges on Tom22 (Brix et al. 1997), and continues through a “binding chain” by contacting the various translocases (Pfanner and Geissler 2001). Eventually, as the precursor reaches TIM23, the positive charges on the presequence are acted upon by the membrane potential that draws the precursor into the channel (Martin et al. 1991b). Upon translocation through TIM23, the presequence is generally cleaved by MPP from the majority of proteins (Gakh et al. 2002), but there are exceptions where presequences remain an integral part of the mature protein (Rospert et al. 1993). Curiously, one case of a cleavable presequence at the C-terminus of a matrix protein has been reported, where the precursor is translocated in a C- to N-terminal rather than the common N- to C-terminal orientation (Lee et al. 1999). Following cleavage, if any, the precursors are then either imported for further folding into the matrix, or released by TIM23 into the inner membrane if they additionally possess a hydrophobic stop-transfer signal (Glick et al. 1992; Beasley et al. 1993; Bomer et al. 1997). Some inner membrane and IMS preproteins con-

Protein Import into Hydrogenosomes and Mitosomes

35

tain a bipartite presequence that comprises an N-terminal positively charged matrix-targeting sequence, and a downstream sorting signal that is similar to sorting signals found on bacterial and ER secretory proteins. These preproteins are ﬁrst processed by MPP, and then undergo a second cleavage by IMP (Schneider et al. 1991; Nunnari et al. 1993). Other types of inner membrane proteins have internal targeting signals as a combination of transmembrane hydrophobic segments together with positively charged loops (Folsch et al. 1996; Davis et al. 1998). Multiple internal targeting signals that act cooperatively have been characterized for the inner membrane ADP/ATP carrier (AAC). No consensus sequence has been computed for these signals, but each segment of about ten amino acid residues can be recognized individually by the Tom70 receptor (Brix et al. 2000; Wiedemann et al. 2001). In general, internal targeting signals for hydrophobic proteins are poorly characterized. Some outer membrane, monotopic proteins, like Tom70, have a non-cleavable presequence that directs the precursor to mitochondria, and drives the insertion of a downstream hydrophobic stretch that acts as a membrane anchor (Hahne et al. 1994). Signals within other types of outer membrane proteins such as the β-barrel proteins have not yet been characterized. 5.2 Signals on Precursors of Soluble Hydrogenosomal and Mitosomal Proteins Characterization of targeting signals within the hydrogenosomal and mitosomal proteins has mainly focused on putative N-terminal cleavable presequences. These are relatively straightforward to detect, by experimentally determining the N-terminal sequences on isolated endogenous organellar proteins and comparing those with the conceptual translation of the corresponding genes. However, this exercise has been done on only one protein each from hydrogenosomes of N. frontalis, or from mitosomes of G. intestinalis and E. histolytica, and on 14 proteins from T. vaginalis hydrogenosomes (Table 1). Most of the putative N-terminal presequences on hydrogenosomal and mitosomal proteins have been either predicted by programs that have been devised to search for mitochondrial presequences, or by sequence comparison with eubacterial homologues. In many cases, the ability of these putative presequences to function as genuine targeting signals has been tested by assessing their efﬁciency in conducting passenger proteins to mitochondria, or to the relevant organelle. 5.2.1 Trichomonas Hydrogenosomes A hydrogenosomal N-terminal cleavable presequence was ﬁrst noted in T. vaginalis ferredoxin, a matrix protein, when puriﬁed endogenous ferredoxin was found to lack eight amino acid residues at the N-terminus, relative to the

Ferredoxin b Succinyl coA synthetase, α-subunit 1 Succinyl coA synthetase, α-subunit 2 Succinyl coA synthetase, α-subunit 3 Succinyl coA synthetase, β-subunit Adenylate kinase Chaperonin60 Pyruvate:ferredoxin oxidoreductase subunit A, PFORA PFORB Malic enzyme A Malic enzyme B Malic enzyme C Malic enzyme D Isd11 c [Fe]-hydrogenase maturase HydG d [Fe]-hydrogenase maturase HydF [Fe]-hydrogenase maturase HydE Ferredoxin e Iron-sulfur cluster assembly protein IscS-1 IscS-2

Trichomonas vaginalis, hydrogenosomal matrix

MLSSFLSRTF>ANESVMANLRES MLASLSRS[]YGKLRADVSKTL MLTSIGRY[]FAKKGNDLPRTH MSHDHIVRL[]LNPRTKDEIDA MLSQCSPLRF()GSVTVTKGGA MLTNLYNKA()FHGHYLDAQATSI MLGSVSRS()YFKGHYLDTQATSV

MLSQVCRF^GTITAVKGGVKK MLAGDFSRN^LHKPLLFIDKD MLSSSFERN^LHQPLLFIDKD MLSSSFERN^LHQPLLFIDKD MLSSSFARN^FNILEWQSKEI MLSTLAKRF^ASGKKDRMVVF MSLIEAAKHFTRAF^AKARDL MLRSF^GKRIPGDGNTAATSV MLRNF^SKRVPGDGNTAATSV MLTSSVSVPVRN^ICRAKVPT MLTSSVNFPARE^LSRNVRPT MLTSVSYPVRN^ICRSKLPLA MLTSVSLPVRN^ICRSKLPVA

N-terminal presequence

Table 1 Presequences of precursors to hydrogenosomal and mitosomal proteins a

Johnson et al. (1990) Lahti et al. (1994) Lahti et al. (1994) Lahti et al. (1994) Lahti et al. (1992) Lange et al. (1994) Bui et al. (1996) Hrdy and Muller (1995a) Hrdy and Muller (1995a) Hrdy and Muller (1995b) Hrdy and Muller (1995b) Hrdy and Muller (1995b) Hrdy and Muller (1995b) Richards and van der Giezen (2006) Putz et al. (2006) Putz et al. (2006); our unpublished data Putz et al. (2006); our unpublished data Dolezal et al. (2005) Tachezy et al. (2001); Sutak et al. (2004) Tachezy et al. (2001); Sutak et al. (2004)

Refs.

36 S.D. Dyall · P. Dolezal

Dyall et al. (2000) Our unpublished data; Acc EAX95270g Our unpublished data; Acc EAY15971g Our unpublished data; Acc EAY00062g

MAQPAEQILIAT^SPKPSLSP MATEADKVLIAT∗ SPNGALPT MKPADKILIAT∗ SPSDAKLKP MKIKFSFGQKQKKDNL∗ SPVQ

MLATFARNF∗ AAKKVTIKPLG MSIVNKF∗ VEKALSLPTYAKA MSIISRY∗ AVPQISKLSNGVRV()

MLSSVARSTSSLFSRG()FAAG

DnaK/Hsp70

Chaperonin 10 Pam18 Mitochondrial processing peptidase β-MPP Trichomonas vaginalis, hydrogenosomal membrane Mitochondrial carrier family protein Hmp31 Hmp31-a Hmp31-b Hmp31-d

Dolezal et al. (2005) Dyall et al. (2004b); Hrdy et al. (2004) Dyall et al. (2004b); Hrdy et al. (2004) Mukherjee et al. (2006a) Mukherjee et al. (2006a) Mukherjee et al. (2006b) Dyall et al. (2004b) Dyall et al. (2004b) Bui and Johnson (1996) Bui and Johnson (1996); our unpublished data Bui et al. (1996); Germot et al. (1996); Dyall et al. (2003) Bui et al. (1996); our unpublished data Dolezal et al. (2005) Dolezal et al. (2005)

Refs.

MLAAVSRS()SALNMMKPLGIM MLAAYGHRF()QTKFLDPKDRI MLASVNTSRF()FARLNKKS>VL MITSCFTRA∗ AKQYSKDHLWF MISTLCNCSRNF∗ TKLYAKT>H MLKNVFHRF∗ SSSWILSEKVL MLTSVSLPVRN∗ ICRSKLPVA MLTSSVSLPARE∗ LSRKVLPT MLASSATAMKGFANSLRM∗ KD MLASSSRAAANIRW()VDTSHN

N-terminal presequence

Iron-sulfur cluster assembly protein IscU Complex I protein Ndh51/NuoF Complex I protein Ndh24/NuoE Glycine cleavage H protein H1 f Glycine cleavage H protein H2 Serine hydroxymethyl transferase SHMT Malic enzyme G Malic enzyme H [Fe]-hydrogenase A [Fe]-hydrogenase B

Table 1 (continued)

Protein Import into Hydrogenosomes and Mitosomes 37

Antonospora locustae putative mitosome mtG3PDH MINKRTYTYAFAAIGTGVLGYVGHRYYRHRK∗

Burri et al. (2006)

Clark and Roger (1995); Mai et al. (1999); Tovar et al. (1999)

MLRVLSENRF∗ PLSLVAGVVA MLIRD()IVPGALPS>ATVVFSG

MLSSSSHYNGKLLSLNIDCRE^NVL

Pam18 β-MPP Entamoeba histolytica Chaperonin 60

Tovar et al. (2003); Dolezal et al. (2005) Nixon et al. (2002a); Nixon et al. (2002b); Dolezal et al. (2005) Dolezal et al. (2005) Dolezal et al. (2005)

MTSLQLSSTSLLQSVARF^LTKKTSSDEVYSE MSLLSSIRRFITFRV()VQQ>

Giardia intestinalis IscU Ferredoxin

van der Giezen et al. (2003) van der Giezen et al. (2003)

Voncken et al. (2002)

MSMLSSVLNKAVVNPKLTRSLATAAEKMVNISING RKF()QV

MLSARSLICKSMIKSGFRRAVAPSVAMAASSMTLTARRNY∗ SSKY MFLSTLAKKSTTFGVSNVVKNALSSKVMRTTPRMFQRF∗ ESSK

van der Giezen et al. (1997) Brondijk et al. (1996)

Refs.

MLAPIQTIARPVSSILPATGALAAKRT^FFA MLANVTRSTSKAAPALASIAQTAQKRF∗ LSV

N-terminal presequence

Neocallimastix patriciarum Chaperonin 60 DnaK/Hsp70

Malic enzyme Succinyl coA synthetase, β-subunit Neocallimastix sp. L2 [Fe]-hydrogenase

Neocallimastix frontalis

Table 1 (continued)

38 S.D. Dyall · P. Dolezal

MLLRSGINLYKSVEGSIGLRSAAIRFGMRYISSGKE>()LSF MSMIINSSFNGVVNSSGIAARILKRSLPLVFSRY[]MSSK MIVHRYCRQWAPSVVRGISKLAFFSSMSSIAKKRPAY>[]FDY MLQLRQLIDKRILIKKCVPICQRLFYS>DTV

N-terminal presequence

Riordan et al. (2003) Slapeta and Keithly (2004) LaGier et al. (2003) LaGier et al. (2003)

Refs.

a This survey only shows proteins that have been shown to localize to hydrogenosomes, mitosomes or heterologous mitochondria, as reported in the literature, or from unpublished proteomics data b Experimentally determined presequences are shown in bold with the cleavage site marked with ^. Arg residues at the – 2 or – 3 position relative to the cleavage site are underlined c Cleavage sites predicted by MitoProtII (Claros and Vincens 1996) are indicated by > d Cleavage sites predicted by iPSORT (Bannai et al. 2002) are indicated by [] e Cleavage sites predicted by PSORTII (Nakai and Horton 1999) are indicated by () f Cleavage sites suggested by authors or suggested here are indicated by ∗ g GenBank accession number

Cryptosporidium parvum Chaperonin 60 DnaK/Hsp70 IscS IscU

Table 1 (continued)

Protein Import into Hydrogenosomes and Mitosomes 39

40

S.D. Dyall · P. Dolezal

conceptual gene translation (Johnson et al. 1990). This presequence (Table 1) has an overall positive charge and is signiﬁcantly shorter than typical mitochondrial targeting sequences, which range from 10 to 80 amino acid residues (Pfanner and Geissler 2001). Using an in organello import assay, Bradley et al. (1997) demonstrated that deletion of this eight-amino acid sequence abolishes binding to, and thus translocation of the protein into the hydrogenosome (Bradley et al. 1997). This result has been conﬁrmed in vivo, when ferredoxin that lacked residues 2 to 8 was expressed in T. vaginalis, and was found to reside exclusively in the crude cytosolic fraction (Dyall et al. 2000). This suggests an important role for the presequence in binding to any receptor and/or pore that promotes entry into the hydrogenosome. In the last 10 years, dozens of T. vaginalis hydrogenosomal presequences have been characterized, or predicted for proteins involved in various pathways (Table 1). The emerging picture is that, unlike mitochondrial and plastidic presequences, the T. vaginalis hydrogenosomal presequences are highly conserved at primary sequence level. This is even more striking upon examination of over 100 soluble protein sequences identiﬁed during proteomic studies, where about 75% of the translated gene sequences have N-terminal sequences that closely resemble those shown in Table 1 (our unpublished data). How these presequences have been appended and are so well conserved remains a mystery, but it is possible that, as is the case for newly-transferred angiosperm mitochondrial genes (Adams et al. 2000; Choi et al. 2006), a small subset of hydrogenosomal protein genes has preferentially been used as presequence donor. The T. vaginalis hydrogenosomal presequences are generally short, ranging from 5 to 14 amino acid residues for those that have been proven experimentally, and up to 17 residues for the predicted presequences (Table 1). The presequences are enriched in the amino acid residues Ser (20%), Leu (14%), Arg (11%), Ala (8%), Phe (7%), Val (6%), Thr (6%) and Asn (5%). The other amino acids are signiﬁcantly under-represented. Incidentally, or accidentally, the three amino acids most commonly found in these presequences, Ser, Leu and Arg, are the ones that are each encoded by six codons. This may have been relevant in the evolution of these presequences. The mitochondrial matrix N-terminal presequences are enriched in Arg (14%), Leu (12%), Ser (11%) and Ala (14%). On the other hand, chloroplast leader peptides have a different amino acid composition with 19% Ser and 9% Thr (von Heijne et al. 1989). Markedly under-represented in hydrogenosomal presequences are the acidic residues, as in the case of both mitochondrial and plastidic presequences (von Heijne et al. 1989). Three of the frequently occurring amino acid residues in hydrogenosomal presequences are positionally conserved as well. Of the 13 hydrogenosomal matrix preproteins for which presequence cleavage sites have been experimentally determined, 12 have Leu at position 2 of the presequence, and the exception has a Leu residue at position 3 (Table 1). Thus, not only the presence, but also the position of the Leu residue is conserved. This is even more striking when we

Protein Import into Hydrogenosomes and Mitosomes

41

examine predicted N-terminal presequences for a further 20 proteins that have been localized to hydrogenosomes, where 85% have Leu at position 2. Mutation of the Leu residue at position 2 in the ferredoxin presequence disrupted binding of the protein precursor to hydrogenosomes (Bradley et al. 1997), suggesting that this particular residue plays a critical role in binding. The Arg residue occurs at the – 2 or – 3 position relative to the cleavage site in all the experimentally determined presequences, with 77% at the – 2 position. Phe residues can be frequently found in the vicinity of the Arg residue. Interestingly, many, but not all, mitochondrial N-terminal presequences likewise contain Arg at – 2 or – 3 from the cleavage site (Gavel and von Heijne 1990) but the role of the residue in specifying the cleavage site for MPP is unclear (Gakh et al. 2002). The exact role of these conserved residues, i.e. whether they are important for binding, for translocation, or for cleavage, is not known. Nonetheless, some of these conserved features were applied to devise consensus sequences that were used to screen the T. vaginalis genome sequence database. A genome-wide search using the consensus sequences M-L-(S/T/A)-x(1 – 15) -R-(N/F/E/xF), MS-L-x(1 – 15) -R-(N/F/xF) or M-L-R-(S/N)-F, picked out 138 sequences with 67% showing similarity to known proteins involved in metabolic pathways, electron transport, protein import, protein folding and oxygen scavenging pathways (Carlton et al. 2007). There are undoubtedly variations on these consensus, as have been found during proteomic studies (our unpublished data). Apart from a similar amino acid enrichment, a common feature of these hydrogenosomal presequences and of the mitochondrial N-terminal presequences is their ability to form amphipathic α-helices (Johnson et al. 1990; Lahti et al. 1992; Pfanner et al. 1997; Dolezal et al. 2005). The amphipathic α-helical structure within mitochondrial N-terminal presequences has been shown to be critically important for sequential electrostatic or hydrophobic interaction with various translocases (Pfanner and Geissler 2001). The hydrogenosomal presequence may be interacting with hydrogenosomal translocases using a similar “binding chain” mechanism. To date, it has not been demonstrated that the typical T. vaginalis soluble preprotein presequence is sufﬁcient for translocating the protein into hydrogenosomes, although it has been shown to be necessary (Bradley et al. 1997; Dyall et al. 2000). It is possible that there are additional downstream signals in the mature part of the protein that participate in translocation at stages beyond binding. It has been shown, however, that hydrogenosomal presequences can target a passenger protein into Trypanosoma and S. cerevisiae mitochondria, but at very low efﬁciency (Hausler et al. 1997). 5.2.2 Neocallimastix Hydrogenosomes Although there have been several reports of putative Neocallimastix sp. hydrogenosomal proteins in silico, few have actually been localized to hydro-

42

S.D. Dyall · P. Dolezal

genosomes or to heterologous mitochondria (Brondijk et al. 1996; van der Giezen et al. 1997, 2003; Voncken et al. 2002). These proteins have quite similar N-terminal extensions (Table 1), but only one of them has been experimentally conﬁrmed (van der Giezen et al. 1997). The extensions range from 27 to 37 amino acid residues, and are within the range for typical mitochondrial N-terminal presequences, with similar amino acid enrichment and characteristics. Indeed, when expressed in yeast, the hydrogenosomal malic enzyme was targeted to mitochondria in a presequence-dependent fashion (van der Giezen et al. 1998). The predicted N-terminal presequences on N. patriciarum Cpn60 and Hsp70 were sufﬁcient to target the green ﬂuorescent protein (GFP) to mammalian mitochondria, although some non-speciﬁc targeting was observed for the Cpn60 presequence, suggesting that additional signals are present in the mature part of the protein (van der Giezen et al. 2003). 5.2.3 Cryptosporidium Mitosomes Since the discovery of the single mitochondrial relict organelle in C. parvum (Riordan et al. 2003; Putignani et al. 2004), only four proteins have been localized to the organelle. Cpn60 has a putative 38-amino acid N-terminal presequence (Table 1) that does not follow the Arg-2 rule, but the N-terminal 57-amino acid portion of Cpn60 was necessary and sufﬁcient to target GFP into yeast mitochondria (Riordan et al. 2003). Likewise, the predicted N-terminal extensions on mitosomal IscU and IscS (Table 1) were both sufﬁcient to target GFP to yeast mitochondria (LaGier et al. 2003). The predicted 34-amino acid presequence of Hsp70 closely resembles typical mitochondrial presequences with a predicted amphipathic α-helical domain, similar enrichment in amino acids, and an Arg-2 cleavage site motif (Gavel and von Heijne 1990; Slapeta and Keithly 2004). This predicted presequence could speciﬁcally deliver GFP into yeast and Toxoplasma mitochondria, and it was shown that the speciﬁc presequence region critical for targeting included the predicted amphipathic α-helical domain (Slapeta and Keithly 2004). 5.2.4 Entamoeba Mitosomes Not many E. histolytica mitosomal proteins have been identiﬁed, leaving us with very little information on protein targeting signals. The only experimental data come from the analysis of Cpn60 that has an N-terminal extension of 15 amino acids (Table 1) shown to be important for mitosomal targeting (Mai et al. 1999; Tovar et al. 1999). This presequence, like most of the T. vaginalis presequences, has a Leu residue at position 2 and is highly enriched in Ser residues. While deletion of the extension leads to the accumulation of Cpn60 in the cytosol, the swapping of the extension with the N-terminal presequence

Protein Import into Hydrogenosomes and Mitosomes

43

from mitochondrial Hsp70 of Trypanosoma delivers the protein back into the enriched mitosomal fraction (Tovar et al. 1999). Similar N-terminal sequences were found on Hsp70 and pyridine nucleotide transhydrogenase, but neither protein has been localized to mitosomes as yet (Clark and Roger 1995; Arisue et al. 2002; Bakatselou et al. 2003). Recently, Cpn10 was localized to the mitosomal cellular fraction but it lacked any recognizable N-terminal extension (van der Giezen et al. 2005). No further functional data are currently available on the processing of targeting presequences in E. histolytica, and although complete genome sequence data are available (Loftus et al. 2005), the identiﬁcation of other putative mitosomal proteins has been without success so far. 5.2.5 Giardia Mitosomes In contrast to Entamoeba mitosomes, several proteins have been successfully localized to G. intestinalis mitosomes (Tovar et al. 2003; Dolezal et al. 2005; Regoes et al. 2005). The import of giardial homologues of IscU and [2Fe-2S]ferredoxin (Table 1) was shown to be dependent on the N-terminal targeting sequence (Dolezal et al. 2005; Regoes et al. 2005) as their truncated versions were mislocalized and/or degraded in the cytosol. These two presequences are enriched in Ser, Thr, Leu and Arg and are very similar to the T. vaginalis presequences. The N-terminal sequences of IscU and [2Fe-2S]-ferredoxin, extending beyond the respective predicted presequence cleavage sites, were sufﬁcient to target GFP into mitosomes. The increased electrophoretic mobility of the fusion protein in organellar fractions suggested that the N-terminal presequences were removed upon targeting (Regoes et al. 2005). These two targeting sequences consist of 15–18 amino acid residues that can be projected to form amphipathic α-helices (Dolezal et al. 2005). Interestingly, the gene coding for [2Fe-2S] ferredoxin was demonstrated to contain a spliceable intron just between the exons coding for the N-terminal targeting sequence and the mature ferredoxin (Nixon et al. 2002b). Other soluble proteins that have been localized in mitosomes are the homologues of Cpn60, Hsp70, Pam18, the β-subunit of MPP (β-MPP) and IscS (Tovar et al. 2003; Dolezal et al. 2005; Regoes et al. 2005). Of these, only Pam18 and β-MPP contain recognizable N-terminal presequences (Table 1), while the import of others may rely on internal signals. This is quite unusual since these mitochondrial proteins typically contain cleavable presequences in other studied organisms. Indeed, it was shown that if G. intestinalis IscS is expressed as a 202-amino acid N-terminal polypeptide and a 232-amino acid C-terminal polypeptide, respectively, both truncated proteins could be successfully delivered to organelles (as tested on T. vaginalis hydrogenosomes) showing that targeting information is found in multiple loci within the protein (Dolezal et al. 2005). Deletion of the ﬁrst ﬁve amino acids on G. intestinalis Cpn60 did not affect the targeting of the protein to mito-

44

S.D. Dyall · P. Dolezal

somes (Regoes et al. 2005). Thus, G. intestinalis mitosomes display both presequence-dependent and presequence-independent targeting for soluble preproteins (Dolezal et al. 2005; Regoes et al. 2005). The targeting information on mitosomal proteins can be recognized and processed by the heterologous systems of human and yeast mitochondria, as well as T. vaginalis hydrogenosomes (Dolezal et al. 2005; Regoes et al. 2005). The [2Fe-2S]-ferredoxin N-terminal presequence was sufﬁcient to deliver a passenger protein into human mitochondria (Regoes et al. 2005), and T. vaginalis hydrogenosomes can speciﬁcally import G. intestinalis [2Fe-2S]ferredoxin, IscU, IscS, Pam18 and β-MPP (Dolezal et al. 2005). Furthermore, the presequence on IscU was sufﬁcient to efﬁciently target a passenger protein into T. vaginalis hydrogenosomes. The N-terminal presequence of IscU can be processed by S. cerevisae mitochondrial extract and also by puriﬁed recombinant rat MPP (Dolezal et al. 2005). Altogether, these results strongly suggest that targeting information on G. intestinalis mitosomal proteins can be crossrecognized by the respective protein import machineries of mitochondria and hydrogenosomes. 5.2.6 Microsporidian Mitosomes So far, the only protein that has been localized in situ in microsporidian mitosomes is the homologue of Hsp70 from T. hominis (Williams et al. 2002). Like G. intestinalis Cpn60, Hsp70 and IscS (Tovar et al. 2003; Dolezal et al. 2005; Regoes et al. 2005), this protein sequence does not contain a recognizable N-terminal presequence. Based on the genome sequence of E. cuniculi, the conceptual metabolic map of its putative mitosome was created using a set of 21 predicted mitosomal proteins, some of which appear to have N-terminal presequences with an Arg-2 motif (Katinka et al. 2001; Gakh et al. 2002; Williams and Keeling 2005). However, no mitochondrion-like organelle has been physically detected in this species as yet. Nor has one been detected in the related species A. locustae. The ongoing genome sequencing project for A. locustae (Antonospora Genome Project, Marine Biological Laboratory at Woods Hole, MA, USA) has provided some more interesting insight and surprising differences in mitosomal protein import mechanisms of various microsporidia. Of the identiﬁed microsporidian mitosomal proteins, only a handful has amphipathic N-terminal presequences, and others do not appear to have any extensions, nor have many characteristics in common (Burri et al. 2006). As no genetic transformation technique has been developed as yet for microsporidia, the targeting information on these proteins was investigated by expressing the full-length and truncated versions of these proteins in S. cerevisiae as fusions with GFP (Burri et al. 2006). Of the 16 proteins under investigation, only six, most from A. locustae, could direct GFP to mitochondria. Deletion of the N-terminal predicted extensions from four of these fusion

Protein Import into Hydrogenosomes and Mitosomes

45

proteins disrupted targeting to mitochondria, showing that the extensions are necessary for cross-organellar targeting. The other two proteins, including mitochondrial glycerol-3-phosphate dehydrogenase (mtG3PDH), could still be delivered to mitochondria, suggesting that internal targeting signals are sufﬁcient for targeting. However, the N-terminal sequence of mtG3PDH (Table 1) was also found to be sufﬁcient to deliver GFP to yeast mitochondria. The Nterminal extensions from the other proteins were not sufﬁcient to target GFP to mitochondria. This ﬁnding undermines the exclusive role of the N-terminal sequence in organellar protein targeting. It is apparent that a combination of N-terminal, and mainly internal signals seems to fulﬁl the targeting role in microsporidian mitosomes (Burri et al. 2006). 5.3 Signals on Hydrogenosomal and Mitosomal Membrane Proteins Targeting signals on membrane proteins are generally poorly characterized. Not only do membrane protein precursors require targeting, membrane sorting and insertion signals, but they also require a means of protection against premature folding or aggregation in the hydrophilic environments they encounter during transport to the organelle membrane. A variety of membrane proteins are targeted to mitochondria: β-barrels, tail-anchored, α-helical polytopic and monotopic proteins have been characterized. Given this diversity in structure, speciﬁc but sometimes overlapping pathways are utilized for their insertion (Rehling et al. 2003b; Koehler 2004a; Bohnert et al. 2007). Most of the data available on membrane protein insertion has been generated for members of the mitochondrial carrier family (MCF), particularly for AAC, the model precursor. The T. vaginalis hydrogenosomal Hmp31 precursor protein, a member of the MCF, was found to have a cleavable 12-amino acid N-terminal presequence. Although this sequence is predicted to be mostly α-helical, it does not have an amphipathic disposition, and has an overall negative charge. This presequence was found not to be necessary for targeting and integration of mature Hmp31 in the membrane, suggesting that Hmp31 utilizes internal targeting signals, like virtually all MCF proteins. However, the presequence was necessary, and sufﬁcient, to target a passenger protein to the soluble hydrogenosomal fraction, and as such acted as a targeting signal. Thus, the Hmp31 precursor has internal targeting signals and a functional N-terminal targeting signal (Dyall et al. 2000). Four more Hmp31 orthologues were found during proteomic analyses (our unpublished data) and two of those, Hmp31-a and Hmp31-b, were each found to have a similar N-terminal extension (Table 1). Another orthologue, Hmp31-d, however, had a putative N-terminal extension that resembles the matrix-targeting N-terminal presequence, with an overall positive charge. The fourth one does not appear to have an N-terminal extension. However, none of those novel Hmp31 ortho-

46

S.D. Dyall · P. Dolezal

logues have had their N-termini experimentally determined as yet. No such presequence has been found on either the Neocallimastix hydrogenosomal AAC (van der Giezen et al. 2002) or on the Entamoeba mitosomal AAC (Chan et al. 2005). Although most MCF proteins are synthesized without N-terminal extensions, a subset of precursors has cleavable presequences. Plant mitochondrial AACs are synthesized with long N-terminal presequences, but these are both not necessary and not sufﬁcient to target passenger proteins to mitochondria and are therefore not acting as targeting signals (Glaser et al. 1998; Murcha et al. 2005a). Another MCF protein, the mammalian phosphate carrier, bears a presequence that may act as an enhancer for translocation but is not strictly necessary, though it was marginally sufﬁcient to target a passenger protein to mitochondria (Zara et al. 1992). Recently, it has been suggested that the dispensable N-terminal presequences of mammalian and ﬁsh citrate carriers may in fact act as chaperones to increase the solubility of the preprotein in the cytosol through electrostatic interaction (Zara et al. 2007). All MCF members that have been characterized in hydrogenosomes or mitosomes have been successfully imported into yeast mitochondria (Dyall et al. 2000; van der Giezen et al. 2002; Chan et al. 2005). Therefore, all three precursors must have targeting signals that are compatible with the speciﬁc mitochondrial pathway used for mitochondrial carriers (Rehling et al. 2003b). Indeed, T. vaginalis Hmp31 import into mitochondria was found not only to be dependent on membrane potential, but also on the presence of the small TIM chaperone complex (Fig. 1) that is essential for proper mitochondrial AAC translocation (Dyall et al. 2000). Conversely, mitochondrial AAC was efﬁciently targeted to T. vaginalis hydrogenosomes, showing that targeting signals are compatible between the two systems (Dyall et al. 2000). The targeting of β-barrel membrane proteins is conserved between hydrogenosomes and mitochondria, as a unique hydrogenosomal β-barrel protein, Hmp35, could be targeted to mitochondrial membranes where it associated with, or assembled into, a high molecular weight complex (Dyall et al. 2003). It is notable that βbarrel precursors from eubacteria and plastids can be successfully imported and assembled into mitochondria as well (Rohl et al. 1999; Muller et al. 2002). Thus, targeting and insertion pathways for β-barrel proteins appear to be conserved between eubacteria, mitochondria, plastids and hydrogenosomes. T. vaginalis hydrogenosomes may be using a conserved SAM-like pathway (Fig. 1) for insertion of β-barrel proteins, as a homologue of Sam50 has recently been discovered in the T. vaginalis genome (Dolezal et al. 2006).

6 Crossing the Organellar Membranes All hydrogenosomes and mitosomes examined to date appear to have double membranes, which implies the presence of an intermembrane space.

Protein Import into Hydrogenosomes and Mitosomes

47

However, no technique has been developed as yet to selectively disrupt the organelles to separate the membranes, and thus isolate any putative IMS proteins. To date, no hydrogenosomal or mitosomal protein import component has been functionally characterized, but some putative players have been identiﬁed in the genomes of the protists through sequence comparison with mitochondrial translocases from various species. Given that mitosomal and hydrogenosomal preproteins bear signals that are recognized by the mitochondrial protein import machinery, it is likely that at least some components are phylogenetically and/or functionally conserved between these organelles. More insight into potential import processes can be gained by examining in greater detail how mitochondrial preproteins interact with translocases to cross organellar membranes. 6.1 The Outer Membrane Two major protein import machineries have been characterized to date in the mitochondrial outer membrane: the TOM and the SAM complexes (Fig. 1). 6.1.1 The General Import Pore Almost all preproteins enter mitochondria through the TOM complex. In yeast mitochondria, this complex consists of an approximately 400 kDa stable core, referred to as the general import pore (GIP), that comprises Tom40, Tom22, Tom5, Tom6 and Tom7. The preprotein receptors Tom70 and Tom20 are only loosely associated with the GIP (Pfanner and Geissler 2001; van der Laan et al. 2006a; Bohnert et al. 2007). Tom70 is the preferred receptor for hydrophobic preproteins with or without presequences (Wiedemann et al. 2001; Chan et al. 2006), although Tom20 also participates in binding (Brix et al. 1997). A typical substrate for Tom70 is the precursor to AAC. AAC has multiple internal targeting signals that are recognized by several Tom70 dimers, which probably act to prevent aggregation of these hydrophobic precursors (Brix et al. 2000; Wiedemann et al. 2001). Preproteins with N-terminal presequences initially make contact with Tom20 (Sollner et al. 1989). This interaction occurs through the hydrophobic surface of the amphipathic helix formed by the presequence, as demonstrated by structural studies (Abe et al. 2000). Thereafter, the two surfaces of the presequence are differentially recognized by binding domains of increasing afﬁnity within the downstream translocases (Pfanner and Geissler 2001). Preproteins from both Tom20 and Tom70 are subsequently transferred to Tom22, which acts both as a docking point for Tom20 and Tom70 and as a central receptor for preproteins within the GIP (Bolliger et al. 1995; Honlinger et al. 1995; Brix et al. 1997; van Wilpe et al. 1999).

48

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The cytosolic domain of Tom22 interacts with the positively-charged surface of the amphipathic helix formed by N-terminal presequences (Brix et al. 1997). Next, the small protein Tom5 transfers preproteins from Tom22 to the Tom40 channel for translocation across the outer membrane (Dietmeier et al. 1997; Hill et al. 1998; Kunkele et al. 1998). It is thought the GIP houses two to three such channels, which are each made up of two molecules of the β-barrel protein Tom40, the only essential component of the TOM complex (Baker et al. 1990; Dekker et al. 1998; Kunkele et al. 1998; Ahting et al. 1999; van Wilpe et al. 1999; Becker et al. 2005). Besides making up the channel, Tom40 also has a binding site for presequences (Hill et al. 1998). After they pass through the channel, presequence-containing precursors bind to the IMS domain of Tom22 through the positive surface of the presequence, and are subsequently sorted to the TIM23 complex. Therefore, a typical N-terminal presequence is recognized at least ﬁve times by Tom proteins, through either hydrophobic or ionic interactions (Pfanner and Geissler 2001; Bohnert et al. 2007). Following passage through the Tom channel, other types of preproteins are sorted into their respective specialized biogenesis pathways. Precursors to outer membrane β-barrel and inner membrane carrier proteins are guided by the small TIM chaperone complexes to their respective SAM or TIM22 pathway. Precursors to the small Tims and to other IMS proteins are taken up into the MIA pathway for further processing (Bohnert et al. 2007). Given the intricacy and speciﬁcity displayed by the yeast mitochondrial protein import machinery, one might expect that the outer membrane translocases, or Tom proteins, would be conserved across species. Moreover, the demonstrated ability to successfully and speciﬁcally import mitosomal and hydrogenosomal preproteins into mitochondria led many to infer that similar and phylogenetically related receptors were present in hydrogenosomes and mitosomes as in mitochondria. This inference has in turn been used as supporting evidence for a common origin for mitochondria and related organelles. However, recent sequence surveys of complete genome databases have taught us that, to start with, not all Tom proteins are conserved across all mitochondrial species (Macasev et al. 2004; Likic et al. 2005; Chan et al. 2006; Perry et al. 2006), let alone mitosomal or hydrogenosomal species. Indeed, a comprehensive survey of available completed eukaryotic genomes revealed that only Tom7, Tom22 and Tom40 sequences are conserved amongst the majority of eukaryotes, including animals, plants, fungi and some protists (Macasev et al. 2004). Other components such as Tom20 and Tom70 have only been found in the genomes of animals and fungi so far, but not in plants or protists (Likic et al. 2005; Chan et al. 2006). Although a “Tom20” has been named and functionally characterized in plants (Heins and Schmitz 1996; Werhahn et al. 2001), it bears no primary sequence similarity to the fungal and animal Tom20 sequences, and is likely to be of independent origin (Likic et al. 2005; Perry et al. 2006). Strikingly, though, the plant Tom20 has similar, but oppositely orientated structural domains to the

Protein Import into Hydrogenosomes and Mitosomes

49

fungal Tom20, which appear to fulﬁl similar functions (Abe et al. 2000; Likic et al. 2005; Perry et al. 2006). These observations, taken together, have led to the hypothesis that the mitochondrial ancestor to eukaryotes had invented a core TOM complex consisting of Tom40, Tom22 and Tom7, and that other components subsequently evolved independently in the descendants as they progressively tweaked their respective mitochondrial protein import apparatuses (Macasev et al. 2004; Dolezal et al. 2006). Within the completed genome sequences of T. vaginalis, G. intestinalis, E. histolytica, Cryptosporidium and E. cuniculi, putative Tom40 homologues were only found in Cryptosporidium (Abrahamsen et al. 2004; Xu et al. 2004) and in E. cuniculi, which also has a putative Tom70 homologue (Katinka et al. 2001). A possible Tom70 homologue was reported for G. intestinalis (Regoes et al. 2005), but closer examination of the sequence revealed that it did not possess sequence signatures speciﬁc to Tom70 (Chan et al. 2006). It is possible that highly divergent homologues to the “ancestral” set of Tom40, Tom22 and Tom7 (Macasev et al. 2004; Dolezal et al. 2006) may be discovered using more sensitive genome mining methods such as the HMM. However, the positions of T. vaginalis and G. intestinalis in the eukaryotic tree are largely unresolved, and their appellation of “early-diverging” has been generally rescinded. Therefore, it is difﬁcult to hypothesize what translocases “should” be or have been in their respective genomes. 6.1.2 Sorting and Assembling β-Barrel Proteins Upon entering mitochondria through the TOM channel, precursors to β-barrel proteins such as Tom40, porin and Mdm10 are directed by the small TIM chaperone complexes to the SAM pathway for correct sorting and insertion into the outer membrane (Bohnert et al. 2007). The exact function of each of the four components of the core SAM complex is still unclear, but Sam50 (Kozjak et al. 2003; Paschen et al. 2003; Gentle et al. 2004) and Sam35 (Milenkovic et al. 2004; Waizenegger et al. 2004) are essential for cell viability, whereas Mdm10 (Meisinger et al. 2004) and Sam37 (Wiedemann et al. 2003), though involved in β-barrel biogenesis, are not essential components. Recently, more players such as Mim1 and Mdm12/Mmm1 have been characterized that act downstream of the core SAM complex (Ishikawa et al. 2004; Waizenegger et al. 2005; Meisinger et al. 2007). Some components appear to be important only for the biogenesis of sub-categories of β-barrels, such that further speciﬁc pathways may be uncovered in the near future. The insertion of β-barrel precursors is one of the two translocation processes, besides the sorting of inner membrane and IMS proteins, that are clearly derived from a eubacterial translocation system. β-Barrel proteins are exclusively found in the outer membranes of Gram-negative bacteria and of endosymbiotic organelles such as mitochondria and plastids (Wim-

50

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ley 2003). The discovery that Sam50, a protein of eubacterial ancestry, played a critical role in the insertion of mitochondrial β-barrel proteins allowed several parallels to be drawn between the eubacterial and mitochondrial βbarrel biogenesis pathways (Paschen et al. 2005; Dolezal et al. 2006). Sam50 is itself a β-barrel protein that is homologous to the β-barrel bacterial protein Omp85, which is found in all bacteria that have an outer membrane. Omp85 is essential for bacterial viability and has been shown to be involved in the insertion of β-barrel protein precursors into the outer membrane of Neisseria (Voulhoux et al. 2003). Phylogenetic analyses revealed that the sam50 gene is widely distributed among eukaryotes, and probably derived from an α-proteobacterial-like bacterium, possibly the mitochondrial endosymbiont (Gentle et al. 2004). Another parallel crops up between the small TIM chaperone complexes and the chaperones Skp and SurA that assist β-barrel precursors as they navigate through the bacterial periplasmic space. In effect, the mitochondrial IMS represents the periplasmic space of the mitochondrial endosymbiont. Though the two chaperone systems are phylogenetically unrelated, they presumably function to prevent aggregation of the substrates according to similar principles (Paschen et al. 2005; Dolezal et al. 2006). With the help of HMM analyses, homologous sequences to Sam50 have been found in the genomes of virtually all eukaryotes with complete genome sequences. These putative translocases all have features common to mitochondrial Sam50, and possibly share a common ancestor though no phylogenetic analyses have been performed on the more recently discovered sequences (Dolezal et al. 2006). The distribution of other components of the SAM complex has not yet been thoroughly investigated amongst mitochondrial eukaryotes, but some components are limited to fungi. No convincing homologues to Sam35, Sam37, Mdm10, Mdm12, Mmm1 or Mim1 have been found by BLAST searches of any of the complete genomes of hydrogenosomal or mitosomal species. 6.2 The Intermembrane Space Chaperones The small TIM chaperones have been shown to convey “complicated” substrates like β-barrel and polytopic hydrophobic membrane protein precursors across the hydrophilic environment of the IMS (Koehler 2004b; Bohnert et al. 2007). The small Tims – Tim8, Tim9, Tim10, Tim12, Tim13 – are about 10 kDa in size and are characterized by a C-x3 -C-x11–16 -C-x3 -C motif. Of the small Tims, only Tim9, Tim10 and Tim12 are essential, and Tim8 and Tim13 appear to be dispensable for yeast (Koehler 2004b). However, both Tim9–Tim10 and Tim8–Tim13 complexes can bind AAC or β-barrel precursors (Gentle et al. 2007). Recent examination of the distribution of these small proteins using HMM analyses revealed that the small Tims have no prokaryotic homologues

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and may be eukaryotic inventions devised to assist membrane protein import. One or more small Tim proteins are diversely distributed amongst eukaryotes but the only occurrence of small Tim-like homologues in the hydrogenosomal or mitosomal species occurs in the genome of C. parvum. The occurrence of the small tim genes in diverse eukaryotes suggest an early origin for these genes (Gentle et al. 2007). It is conceivable that the other hydrogenosomal and mitosomal species developed or acquired independent chaperones. As we saw earlier, there are other types of proteins such as Skp and SurA that undertake similar chaperoning functions in bacteria. The dependence of the hydrogenosomal inner membrane protein Hmp31 on the small TIM chaperone complex when imported into mitochondria suggests that this precursor is likely to be sensitive to the IMS, just like its mitochondrial counterparts (Dyall et al. 2000). Therefore, by extension, a similar but unrelated chaperoning system may exist in T. vaginalis hydrogenosomes. 6.3 The Inner Membrane The two complexes that import cytosolic proteins through the mitochondrial inner membrane, TIM22 and TIM23 split the import pathways of hydrophobic inner membrane proteins from that of presequence-containing preproteins (Fig. 1). 6.3.1 The TIM22 Complex The Tim9–Tim10 chaperone complex delivers MCF proteins such as AAC from the TOM to the TIM22 complex. Tim12, which is peripherally associated to TIM22, acts as a docking receptor for the chaperone complex. The TIM22 complex contains twin pores built from Tim22 that form a voltage-activated channel that is sensitive to synthetic peptides bearing AAC internal targeting signals, but insensitive to synthetic N-terminal presequences. The passage of the substrate through the channel is entirely dependent on the membrane potential, and not on ATP hydrolysis (Kovermann et al. 2002; Rehling et al. 2003a; Koehler 2004a). The roles of the other TIM22 components, Tim18 and Tim54 are as yet undetermined. However, like Tim22 (Sirrenberg et al. 1996) and Tim12 (Jarosch et al. 1997), Tim54 is essential (Kerscher et al. 1997), whereas Tim18 is not (Kerscher et al. 2000; Koehler et al. 2000). All components of the yeast TIM22 complex appear to be restricted to fungi, except for Tim22 which is widely distributed among eukaryotes (Rassow et al. 1999; Dolezal et al. 2006). Among the mitosomal and hydrogenosomal species, only a putative Tim22 sequence was reported in the genome of E. cuniculi (Katinka et al. 2001).

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6.3.2 The TIM23 Complex The TIM23 complex, which intakes presequence-containing precursors, is better characterized than the TIM22 complex, and exhibits the most intricate import mechanisms (Bohnert et al. 2007). Within the complex, Tim23 forms a cation-selective, voltage-gated, protein-conducting, possibly twinpore, channel that is speciﬁcally sensitive to synthetic presequence peptides (Truscott et al. 2001; Martinez-Caballero et al. 2007). Tim17, though homologous in sequence and secondary structure to Tim23, does not form part of the channel, but modulates its activity (Meier et al. 2005; Martinez-Caballero et al. 2007). Tim21 makes direct contact with the TOM complex by interacting with the IMS domain of Tom22, where it promotes precursor release by competing with presequence binding (Chacinska et al. 2005). Tim50 has a dual role in acting as a receptor for presequences, and in regulating the closure of the TIM23 channel (Geissler et al. 2002; Yamamoto et al. 2002; Meinecke et al. 2006). Tim21 also regulates the interaction between PAM and TIM23 by associating with TIM23. This complicated interaction serves to generate two types of TIM23 complexes: one that is matrix-import competent and one that is competent to sort and insert the presequence-carrying inner membrane proteins (Chacinska et al. 2005; van der Laan et al. 2006a,b). All components of TIM23, except for Tim21 (Chacinska et al. 2005), are essential (Dekker et al. 1993; Emtage and Jensen 1993; Maarse et al. 1994; Ryan et al. 1994; Geissler et al. 2002; Yamamoto et al. 2002; Mokranjac et al. 2003a). This is quite surprising, given the central role played by Tim21 at various levels. Again, this attests to the ﬂexibility of the mitochondrial protein import machinery. Genes homologous to tim23 and tim17 have been found in most mitochondrial eukaryotes (Rassow et al. 1999; Dolezal et al. 2006), and tim21 homologues can be found in animal, plant and fungal genomes (Chacinska et al. 2005) but not in protists (our unpublished observations). Tim50 contains a LIM domain commonly found in proteins of diverse functions and no profound sequence analyses have yet been performed to assess its distribution among various species. Sequences related to tim17 and tim23 have been detected in the respective genomes of T. vaginalis and C. parvum (Abrahamsen et al. 2004; Henriquez et al. 2005; Dolezal et al. 2006), thus a core TIM23 complex could exist in the organelles of these organisms. No clear homologues to any TIM23 component could be detected in the complete genome sequences of either E. histolytica (our unpublished observations) or G. intestinalis (Regoes et al. 2005). 6.3.3 The PRAT Family Interestingly, signiﬁcant sequence homology exists for Tim17 and Tim23 from the TIM23 complex, and Tim22 from the TIM22 complex, which all form

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the core of their respective complexes. Thus, it appears that these proteins derive from a common ancestor. An outer envelope protein of the chloroplast, OEP16, which forms an amino acid-selective channel, was also found to share homology with these Tim proteins (Rassow et al. 1999). These proteins all have four transmembrane helices separated by hydrophilic loops, and contain the motif [G/A]-x2 -[F/Y]-x10 -R-x3 -D-x6 -[G/A/S]-G-x3 -G within the transmembrane domains. A similar motif is present in a region of the E. coli LivH protein, which is an amino acid permease. It has been suggested that the eukaryotic proteins may have evolved from similar permeases or amino acid transporters (Rassow et al. 1999). The PRAT family of preprotein and amino acid transporters has been deﬁned to include the structural similarities and the functions of the Tim17/22/23, OEP16 and LivH proteins (Rassow et al. 1999). Eukaryotic PRAT sequences are quite widely distributed and the proteins have been localized to either mitochondria or chloroplasts (Rassow et al. 1999; Ferro et al. 2003; Murcha et al. 2007), but no systematic analyses have been performed as yet to categorize these sequences. In the case of mitosomal and hydrogenosomal species, the only homologues found to membrane-integral Tims are to the PRAT Tims (Katinka et al. 2001; Abrahamsen et al. 2004; Henriquez et al. 2005; Dolezal et al. 2006). Given their ubiquitous distribution, the PRAT Tims may have had an early origin during the conversion of the proto-mitochondrion. However, these putative translocases have yet to be localized, and no phylogenetic analyses have been generated to assess their afﬁnities.

7 The Protein Import Motor The ﬁnal step of the journey of the mitochondrial matrix-targeted preprotein across the membranes involves the participation of an ATP-driven protein import motor, PAM, which pulls the preprotein from the Tim23 channel into the matrix (Fig. 1). In yeast, the core component of the PAM complex is Ssc1, or mt-Hsp70, which is assisted by Mge1, Pam18, Pam16, Pam17 and Tim44 (van der Laan et al. 2006a; Bohnert et al. 2007). With the exception of Pam17 (van der Laan et al. 2005), all PAM components identiﬁed so far are essential for yeast viability (Craig et al. 1987; Maarse et al. 1992; Bolliger et al. 1994; D’Silva et al. 2003; Mokranjac et al. 2003b; Truscott et al. 2003; Frazier et al. 2004; Kozany et al. 2004; Li et al. 2004). Mt-Hsp70 is a member of the Hsp70 chaperone family that is distributed in all domains of life. The bacterial cytoplasmic homologues are called DnaK, and various types of hsp70 genes are found in eukaryotes, with the products localizing to the cytosol, the ER, the mitochondrion or a plastid compartment (Gupta and Singh 1994; Bukau and Horwich 1998). Phylogenetic analyses show strong afﬁnity and conserved signature sequences between

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mt-Hsp70 and α-proteobacterial DnaK, supporting the endosymbiotic origin of mitochondria from an α-proteobacterial-like ancestor (Boorstein et al. 1994; Falah and Gupta 1994; Gupta 2000). The Hsp70 proteins are the central part of protein folding machines that are powered by ATP. Generally, Hsp70 molecules have a highly conserved amino-terminal region with an ATPase domain, and a carboxy-terminal region with a peptide-binding domain. The extensively studied chaperone system in E. coli revealed much about the mechanism of action of DnaK, which is assisted in its function by the nucleotide exchange factor GrpE, and the J-protein DnaJ that enhances ATPase activity (Bukau and Horwich 1998). A similar system can be found operating with a likewise mechanism at the matrix side of the TIM23 complex. In this situation, however, mt-Hsp70 is not involved in protein folding per se, but its properties are put to use to bind a largely unfolded incoming preprotein and to drive it completely into the mitochondrial matrix in an action regulated by ATP hydrolysis and co-chaperones. A fraction of mt-Hsp70 docks onto the TIM23 complex through the essential peripheral membrane protein Tim44 (Voos and Rottgers 2002; van der Laan et al. 2006a). Genes homologous to tim44 have been found in all the completed genome sequences of mitochondrial eukaryotes, and also of α-proteobacteria, where the putative functions of the homologues are unknown (Dolezal et al. 2006). As the freshly imported preprotein enters the mitochondrial matrix, it is bound and/or pulled in by mt-Hsp70, which is assisted by the soluble matrix protein Mge1 (Bolliger et al. 1994; Voos and Rottgers 2002) and the inner membrane protein Pam18 (D’Silva et al. 2003; Mokranjac et al. 2003b; Truscott et al. 2003). These are the respective homologues of bacterial GrpE and DnaJ. Pam18 has a matrix-oriented J-domain with which it can stimulate mt-Hsp70 ATPase activity (Truscott et al. 2003). Pam18 is tightly bound to Pam16, which contains a degenerate J-domain and acts as a regulatory peripheral inner membrane protein within the motor (Frazier et al. 2004; Kozany et al. 2004; Li et al. 2004). The role of the ﬁnal and nonessential component of PAM, Pam17, is unclear, but it is necessary for the stable modular association of Pam16 and Pam18 with TIM23 (van der Laan et al. 2005). This particular component appears to be fungi-speciﬁc as convincing homologues could not be found in other mitochondrial species, or in eubacteria. Mitochondrial-type Hsp70 is the only PAM component that has homologues in all mitosome- or hydrogenosome-containing species examined to date, namely T. vaginalis (Bui et al. 1996; Germot et al. 1996), G. intestinalis (Morrison et al. 2001; Arisue et al. 2002), E. histolytica (Arisue et al. 2002), C. parvum (Slapeta and Keithly 2004), E. cuniculi (Peyretaillade et al. 1998), A. locustae (Germot et al. 1997), T. hominis (Williams et al. 2002), N. ovalis (Boxma et al. 2005) and N. patriciarum (van der Giezen et al. 2003). So far, putative components of the PAM complex found in T. vaginalis genome are homologues to genes encoding mt-Hsp70/DnaK (Bui et al. 1996; Germot et al.

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1996), Pam18 (Dolezal et al. 2005), Tim44 (Dolezal et al. 2006), Mge1/GrpE (our unpublished data). All these components have been localized to the hydrogenosome (Bozner 1997; Dyall et al. 2003; Dolezal et al. 2005; our unpublished data), suggesting that a PAM-like system functions in the organelle. The same complement of genes has been found in the genome sequence of C. parvum (Abrahamsen et al. 2004), but only mt-Hsp70 has been localized to its mitosome so far (Slapeta and Keithly 2004). In Giardia, only mt-Hsp70 (Morrison et al. 2001; Arisue et al. 2002) and Pam18 (Dolezal et al. 2005) have been reported, and BLAST searches could not identify homologues of the other proteins. Both proteins could be targeted to mitosomes (Dolezal et al. 2005; Regoes et al. 2005). In both E. histolytica and E. cuniculi complete genomes, homologues to only mt-hsp70 were found (Katinka et al. 2001; Arisue et al. 2002) but neither gene product has been localized to the mitosome, although in the case of E. cuniculi, no mitosome has been reported as yet. The putative origins of most of the mitosomal and hydrogenosomal mtHsp70 homologues have been thoroughly pursued through phylogenetic analyses where most sequences group with the mitochondrial homologues with fairly strong support, except in the case of G. intestinalis mt-Hsp70, which is divergent (Morrison et al. 2001; Arisue et al. 2002). Generally, it is assumed that the mt-Hsp70 homologues originate from the α-proteobacterial-like endosymbiont that gave rise to the mitochondrion.

8 Preprotein Processing Peptidases Upon translocation into the matrix, the N-terminal presequence of preproteins is processed by MPP and the mature protein is thereafter folded into its native conformation (Fig. 1). Some preproteins have a bipartite presequence that is processed in two steps, the ﬁrst part by MPP, and the second part, which includes an octapeptide motif, by the mitochondrial intermediate peptidase (MIP). Precursors destined for the IMS have an IMS-sorting signal at the N-terminus. The IMP complex is responsible for the maturation of these proteins. Some of the precursors contain bipartite presequences consisting of a matrix-targeting signal followed by an intermembrane space-sorting signal (Gakh et al. 2002). 8.1 The Mitochondrial Processing Peptidase The mitochondrial processing peptidase is an essential zinc-dependent metallopeptidase (Yaffe et al. 1985; Luciano and Geli 1996; Gakh et al. 2002; Nomura et al. 2006). It cleaves the N-terminal presequence from precursors to matrix-targeted proteins, and from precursors destined for the inner mem-

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brane or the IMS. Through sequence comparisons, Gavel and von Heijne (1990) deﬁned four cleavage site motifs for MPP and MIP: 1. R-2 motif: x-R-x^x-(S/x). 2. R-3 motif: x-R-x-(Y/x)^(S/A/x)-x. 3. R-10 motif: x-R-x^(F/L/I)-x-x-(S/T/G)-xxxx^ , where the second cleavage site is for MIP. 4. R-none motif: x-x^x-(S/x). Surveys of mitochondrial presequences showed that, though quite common, these above motifs are not found in all of them, and that the primary sequence for the cleavage site is quite degenerate. The role of the Arg at the – 2 or – 3 position is unclear and may be presequence-dependent as studies on a variety of precursors revealed that mutating the Arg results in cleavage inhibition or modiﬁcation in some cases, but not in others. It may be that the structure, rather than the primary sequence composition of the presequence and perhaps of the mature protein, determines the MPP cleavage site (Gakh et al. 2002). Generally, the enzyme consists of two core subunits each of about 50 kDa in size: α-MPP and β-MPP, which are widely distributed amongst mitochondrial eukaryotes. α-MPP and β-MPP are homologous to each other with up to about 30% identical residues in some species. The catalytic unit is β-MPP, which contains the conserved and critical zinc-binding motif H-xx-H-x76 -E. This motif is characteristic of the pitrilysin protease family that includes bacterial proteases (Rawlings and Barrett 1995). The α-subunit is not involved in processing, but may be involved in substrate recognition and interaction through a highly-conserved glycine-rich loop. However, both subunits are required for processing the presequence in mitochondria (Geli et al. 1990). MPP has long been thought to have evolved from a bacterial protease of the pitrilysin family (Gakh et al. 2002). Recently, a putative peptidase has been characterized from the α-proteobacterial parasitic bacterium Rickettsia and related species, and was found to have domains typical of both subunits of MPP (Kitada et al. 2007). Strikingly, the N-terminal domain of this rickettsial putative peptidase (RPP) resembles the N-terminal region β-MPP with an H-x-x-H-x76 -E motif, and the C-terminal domain of RPP resembles the C-terminal region of α-MPP, minus the glycine-rich loop. Unlike β-MPP, monomeric recombinant RPP was shown to have proteolytic activity on its own, cleaving basic synthetic peptides preferentially. RPP was able to cleave mitochondrial presequence peptides at speciﬁc sites in some cases, albeit at reduced efﬁciency compared with MPP. However, when tested on mitochondrial preproteins with short and long presequences, respectively, RPP was inactive on its own. Processing of the short presequence only occurred when RPP was stoichiometrically mixed with yeast β-MPP, and it was demonstrated that β-MPP was involved in the catalytic activity, and not RPP. Thus, RPP be-

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haved like α-MPP as an activator of β-MPP. The long presequence was not processed by either RPP/β-MPP or RPP/α-MPP, and mutational studies on MPP indicated that this could be due to the absence of a glycine-rich loop on RPP (Kitada et al. 2007). Given the close relationship between mitochondria and Rickettsia (Andersson et al. 1998), these ﬁndings indicate that RPP may represent an ancestral form of both α-MPP and β-MPP, derived from the α-proteobacterial-like mitochondrial endosymbiont. Homologues to β-MPP, both with conserved catalytic motifs, were discovered recently in the genomes of T. vaginalis and G. intestinalis. The Giardia β-MPP homologue was localized to mitosomes and N-terminal sequencing of mitosomal IscU conﬁrmed the cleavage site of its presequence at the position suggested by the PSORT prediction program (Table 1). Together, these data strongly suggest that N-terminal presequence processing in Giardia is MPP-mediated (Dolezal et al. 2005). Curiously, no α-MPP homologue has been found in either species. It is possible that hydrogenosomal and mitosomal β-MPP have proteolytic activity on their own, like RPP, but unlike mitochondrial β-MPP. In C. parvum as well, a homologue to only β-MPP, but not α-MPP has been reported (Abrahamsen et al. 2004; Henriquez et al. 2005). In E. histolytica, one presequence has been shown to be cleaved at a site predicted for MPP (Mai et al. 1999; Tovar et al. 1999), and a possible candidate sequence for β-MPP is found in the genome (our unpublished observations). No MPP homologue was found in E. cuniculi (Katinka et al. 2001), but given the occurrence of presequence-independent protein import in microsporidia (Burri et al. 2006), they may have dispensed with processing peptidases during their reductive evolution. The only hydrogenosomal or mitosomal species reported to have a homologue to α-MPP is N. ovalis (Boxma et al. 2005). There have been no reports of MIP-like proteins, nor of any R-10 motif on protein precursors in any of the hydrogenosomal or mitosomal species. 8.2 The Inner Membrane Protease Anchored on the outer face of the inner membrane, the mitochondrial IMP complex consists of two proteases Imp1 and Imp2 and a regulatory subunit Som1 (Fig. 1). The two proteases have distinct speciﬁcities for IMS protein precursors. Some of the precursors contain bipartite presequences consisting of a matrix-targeting signal followed by an intermembrane spacesorting signal for sequential cleavage by MPP and IMP. Imp1 and Imp2 show signiﬁcant similarity to bacterial type I leader peptidases that cleave the N-terminal signal of precursors that traverse the bacterial membrane (Gakh et al. 2002). The import route of mtG3PDH into microsporidian mitosomes seems to follow the stop-transfer pathway in S. cerevisiae, during which the translo-

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cation of mtG3PDH is stopped at the TIM23 complex, where the precursor remains in the membrane without release into the matrix (Esser et al. 2004). However, the processing step is different in A. locustae and E. cuniculi preproteins. In A. locustae, as in S. cerevisiae, the precursor seems to be processed by IMP that cleaves off the presequence at the position following the ﬁrst transmembrane segment (Esser et al. 2004; Burri et al. 2006). In contrast, in E. cuniculi, the N-terminal domain is retained within the mature protein. S. cerevisiae IMP could process the A. locustae mtG3PDH precursor, and an IMP2 homologue is present in the A. locustae genome. Together, these data suggest that A. locustae has retained an IMP proteolytic processing pathway, but that the related microsporidian species E. cuniculi may have discarded both MPP and IMP processing (Burri and Keeling 2007; Burri et al. 2006). Currently, there is no evidence for IMP processing in any of the other mitosomal or hydrogenosomal species.

9 Folding Newly Imported Soluble Proteins Newly imported proteins enter mitochondria in an extended or only partly folded conformation. Two main chaperone systems have been characterized in mitochondria that fold these incoming proteins into a native state that permits them to perform their function. Mitochondria have inherited these efﬁcient and intricate folding systems from their bacterial progenitor/s: one involving mt-Hsp70, and the other with Cpn60/Cpn10 or Hsp60/Hsp10 (Neupert 1997; Voos and Rottgers 2002). Besides its role in preprotein translocation across the inner membrane through TIM23 and PAM, mt-Hsp70 can also act as a protein folding chaperone. Indeed, mt-Hsp70 in yeast mitochondria is either found in a membraneassociated complex with Tim44 and PAM, or in a soluble state in association with co-chaperones Mdj1 and Mge1. Mdj1 is a highly conserved non-essential mitochondrial homologue of bacterial DnaJ, and was shown not to be involved in translocation, but to be important for protein folding in association with the homologues of GrpE and DnaK (Neupert 1997; Voos and Rottgers 2002). The manner in which the mt-Hsp70 chaperone functions is very similar to that of bacterial DnaK and the system is likely to have been inherited from the bacterial progenitor of mitochondria (Hartl et al. 1994; Stuart et al. 1994; Szabo et al. 1994). As we reported in Sect. 7, homologues to mt-Hsp70/DnaK have been found in the genomes of all mitosomal or hydrogenosomal species examined to date, and homologues to Mge1/GrpE have been found in T. vaginalis (Carlton et al. 2007) and C. parvum (Abrahamsen et al. 2004). Homologues to Mdj1/DnaJ have been reported in T. vaginalis, E. cuniculi and N. ovalis (Katinka et al. 2001; Boxma et al. 2005; Carlton et al. 2007). All the components of the DnaK-type machinery have been localized

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to T. vaginalis hydrogenosomes (Bozner 1997; Dyall et al. 2003; our unpublished data), suggesting that a similar protein folding mechanism occurs in these organelles. The mitochondrial Cpn60/Cpn10 or Hsp60/Hsp10 chaperone system participates in the folding of the majority of newly imported matrix proteins (Neupert 1997; Voos and Rottgers 2002). This system functions downstream of the mt-Hsp70 system, but both systems are likely to co-operate in protein folding (Manning-Krieg et al. 1991). Cpn60 and Cpn10 derive from bacterial homologues GroEL and GroES, respectively, and phylogenetic and comparative analyses of both protein sequences show a robust relationship between the respective monophyletic mitochondrial groups and α-proteobacteria. Since the progenitor of mitochondria is likely to have been an ancestor of extant α-proteobacteria, these ﬁndings support the notion that Cpn60 and Cpn10 originate from the endosymbiont that gave rise to mitochondria (Gupta 1995). In bacteria, including α-proteobacteria, groel and groes genes are found on a single operon, such that the eukaryotic genes are likely to have a common origin (Gupta 1995). Much has been learnt about the mechanism of protein folding in bacteria through the structure of the bacterial GroEL/GroES complex. In E. coli, the GroEL proteins form a double-ring structure comprising two apposed heptameric rings that form a central cavity that binds protein folding intermediates of up to 50 kDa and facilitates folding to the native state. The chaperonin cavity switches from a binding to a folding state through conformational changes induced by ATP. This action is regulated by a saucer-shaped heptameric complex of GroES, which modulates both the ATPase cycle and the conformation of GroEL monomers (Rye et al. 1997; Xu et al. 1997; Bukau and Horwich 1998). Both Cpn60 and Cpn10 are encoded by essential genes in yeast, and are likely to function similarly to their bacterial homologues (Cheng et al. 1989; Rospert et al. 1993), but not all mitochondrial proteins require Cpn60 for folding (Rospert et al. 1996). Homologues to Cpn60 that show high afﬁnity to mitochondrial Cpn60 have been found in E. histolytica (Clark and Roger 1995), T. vaginalis (Bui et al. 1996; Horner et al. 1996; Roger et al. 1996), G. intestinalis (Roger et al. 1998), C. parvum (Riordan et al. 2003; Putignani et al. 2004), N. patriciarum (van der Giezen et al. 2003) and N. ovalis (Boxma et al. 2005). These putative chaperones have been localized to either hydrogenosomes or mitosomes in T. vaginalis (Bui et al. 1996; Bozner 1997), E. histolytica (Mai et al. 1999; Tovar et al. 1999), G. intestinalis (Regoes et al. 2005), C. parvum (Riordan et al. 2003; Putignani et al. 2004) and N. patriciarum (van der Giezen et al. 2003). The only exception is N. ovalis but the Cpn60 has a putative N-terminal presequence predicted by MitoProtII (Claros and Vincens 1996; Boxma et al. 2005). So far, homologues to Cpn10 have been reported in the genomes of both T. vaginalis and E. histolytica, but phylogenetic relationships with either mitochondrial or α-proteobacterial sequences could not be convincingly inferred (Bui et al. 1996; van der Giezen et al. 2005). Still, a common an-

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cestry is assumed with its interacting partner Cpn60, and by extension, an origin from the mitochondrial endosymbiont is inferred (Bui et al. 1996; van der Giezen et al. 2005). Surprisingly, no homologue to either cpn60 or cpn10 was found in the only complete microsporidian sequence available to date (Katinka et al. 2001). It may be that microsporidian mitosomal proteins do not require Cpn60/Cpn10 for protein folding, as has been noted for a subset of mitochondrial matrix proteins (Rospert et al. 1996). It is plausible that, for the sake of economy, the highly reduced microsporidian mitosomes utilize homologues to the multifunctional mt-Hsp70 protein to both translocate and fold newly imported proteins, and have dispensed with the energetically expensive Cpn60 machinery. The absence of cpn10-like sequences in the completed genomes of G. intestinalis and two Cryptosporidium species (Abrahamsen et al. 2004; Xu et al. 2004) may likewise represent an ongoing reductive process for this folding system.

10 Perspectives It is evident that protein import mechanisms are conserved between hydrogenosomes, mitosomes and mitochondria. Although no protein import pathway has been functionally deciphered for hydrogenosomes and mitosomes, we have started to get a glimpse of some putative mitochondrial-like components that may be involved in importing, processing and folding preproteins during biogenesis. Of all the relevant studied species, only four species that have been shown to harbour either mitosomes or hydrogenosomes have had their genomes completely sequenced, and thus offer us an opportunity to examine their putative mitochondrial protein import complement. Mitosomes of C. parvum and hydrogenosomes of T. vaginalis potentially house the most mitochondrial-like components, though many of them have not as yet been localized. C. parvum mitosomes potentially contain Tom40, Sam50, small Tims, members of the PRAT family, Tim44, Hsp70, Pam18, Mge1, β-MPP and Cpn60. Thus, most of the major mitochondrial protein import machines are represented, except for the Tom receptors Tom20, Tom70 and Tom22. Since the presequence on a C. parvum mitosomal protein could act as a mitochondrial targeting signal, it is likely that functionally similar receptors are present on the C. parvum mitosome, but that they escape detection by sequence analyses. T. vaginalis has homologues to Sam50, Tim17, Tim23, Tim44, mt-Hsp70, Pam18, Mge1, β-MPP, cpn60 and cpn10. In theory, T. vaginalis hydrogenosomes could have mitochondrial-like SAM, TIM23 and PAM machines, and mitochondrial-like preprotein processing and folding. It is quite surprising that T. vaginalis has no obvious homologues to Tom proteins, given the demonstrated compatibility of protein import between hydrogenosomes and mitochondria. In particular, homologues to

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translocases comprising the speciﬁc pathway for the import of MCF preproteins are absent in the T. vaginalis genome, but a mitochondrial MCF protein could be speciﬁcally targeted to hydrogenosomal membranes. Are these translocases present in the genome but too divergent to be ﬁshed out by sequence comparison? Or have they arisen independently to undertake similar functions? These two questions come up repeatedly as we examine the distribution of mitochondrial-like components across the mitosomal species. In the case of G. intestinalis mitosomes, only four mitochondrial-type protein import components have been identiﬁed and localized: Pam18, Hsp70, β-MPP and Cpn60. E. histolytica has homologues to Hsp70, β-MPP, Cpn60 and Cpn10. There are undoubtedly more components involved in organellar protein import in these systems, and again these components are likely to use similar principles as their mitochondrial counterparts, given the seamless ease and efﬁciency with which hydrogenosomal and mitosomal precursors are imported into heterologous mitochondria. For all we know, the protein import machineries in these organelles may be as intricate and as sophisticated as the mitochondrial equivalent. We seem to be reaching the limits of how much we can assimilate and conclude from genome sequence analyses. These have been invaluable in identifying some putative protein translocases and chaperones. More sensitive searches like HMM may indeed deliver further putative candidates for mitochondrial-type translocases from the genome sequence databases of the hydrogenosomal and mitosomal species. However, we shall need to go back to the bench to demonstrate their localization and investigate their involvement in organellar protein trafﬁcking. We now have speciﬁc questions to tackle. For instance, what are the equivalents of the Tom receptors on the outer surface of hydrogenosomes and mitosomes? What is the equivalent of the TIM23 channel in G. intestinalis and E. histolytica mitosomes? How do T. vaginalis hydrogenosomes and E. histolytica mitosomes import MCF proteins? Do mitosomes have β-barrel proteins? How are β-barrel proteins inserted? Some of these questions can be answered with the limited set of tools we have at hand, but it is imperative to develop new techniques if we want to dissect these pathways. Once these questions are answered, we shall be in a better position to formulate hypotheses on how these fantastic machines have evolved. By comparing protein import mechanisms and examining the structure of translocases between hydrogenosomal, mitosomal, mitochondrial, plastidic and eubacterial systems, it is likely that we discover common principles for protein targeting. We can ask further and broader impact questions. For instance, how do the intricacy of the protein targeting machines correlate with proteome size? How do targeting signals and translocases co-evolve? How hard is it for an endosymbiont to build a protein import machine?

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_104/Published online: 5 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Structure of the Hydrogenosome Marlene Benchimol Universidade Santa Úrsula, Rua Jornalista Orlando Dantas 59, CEP 222-31-010 Rio de Janeiro, Brazil [email protected] 1

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Abstract Hydrogenosomes are very interesting organelles found in nonmitochondrial organisms. They display similarities and differences with mitochondria. Hydrogenosomes are spherical or slightly elongated organelles, although very elongated hydrogenosomes are also found. They measure between 200 and 1000 nm, but under stress conditions can reach 2 µm. Hydrogenosomes divide in three different ways, like mitochondria: segmentation, partition, and the heart form. They may divide at any phase of the cell cycle. Nucleoid or electron-dense deposits are not considered part of the normal structure of the hydrogenosome. Hydrogenosomes are surrounded by two closely apposed membranes and present a granular matrix. Hydrogenosomes have one or multiple peripheral vesicles, which incorporate calcium. The peripheral vesicle can be isolated from the hydrogenosomal matrix and is considered a distinct hydrogenosomal compartment. Dysfunctional hydrogenosomes are removed by an autophagic process and further digested in lysosomes.

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1 Introduction During the evolutionary process, eukaryotic microorganisms appeared presenting special cytoplasmic organelles (Fig. 1a–d). One example was the hydrogenosome initially found in protozoa of the Trichomonadida Order, which contains enzymes that participate in the metabolism of pyruvate formed during glycolysis and was the site of molecular hydrogen and ATP formation (reviewed in Müller 1993). In trichomonads, hydrogenosomes have been recognized by light microscopists for a long time, as paraxostylar and paracostal granules, due to their proximity to the axostyle (bundle of microtubules) and the costa, a periodic proteinaceous structure (Fig. 1c,d). Only biochemistry revealed their functional signiﬁcance, which showed molecular hydrogen production as a metabolic end product. Consequently, they were named hydrogenosomes by Lindmark and Müller, in 1973. Hydrogenosomes present an unusual function: under anaerobic conditions they produce molecular hydrogen by oxidizing pyruvate to malate (Müller 1993).

2 Where are Hydrogenosomes Found? The hydrogenosome is an unusual organelle found in some anaerobic fungi (Fig. 1a) (Yarlett et al. 1986; Marvin-Sikkema et al. 1992; Benchimol et al. 1997), in rumen ciliates (Fig. 1b) (Yarlett et al. 1981, 1984; Snyers et al. 1982), in protozoa of the Trichomonadida Order (Fig. 1c) (Lindmark and Müller 1973; Kulda et al. 1987), and also in some free-living ciliates (van Bruggen et al. 1984). The hydrogenosome-containing microorganisms do not present typical mitochondria (Embley et al. 2003). Until today, hydrogenosomes have not been found in multicellular animals or plants, nor in other anaerobic protists, such as amoebas and giardias. The most extensive studies of this organelle have been carried out in the trichomonad species. The distribution of hydrogenosomes in different Phyla raised questions concerning their evolutionary origin. Fig. 1 General view of different cells that possess hydrogenosomes (H): a the anaero- bic fungus Neocallimastix, b a rumen ciliate, and c,d Tritrichomonas foetus as seen by transmission electron microscopy (a–c) and high voltage electron microscopy (d). In d, the plasma membrane of T. foetus was removed and the preparation was critical point dried and observed in a high voltage electron microscope with 1000 kV of electron acceleration. Note that hydrogenosomes are preferentially located along the axostyle (Ax) and costa (C). ER, endoplasmic reticulum; GL, glycogen granules; F, anterior ﬂagella; RF, recurrent ﬂagellum; N, nucleus; P, pelta; V, vacuoles. Bars: a = 2 µm; b = 300 nm; c = 500 nm; d = 1 µm. (Fig. 1a from Benchimol et al. 1997; Fig. 1b–d, Benchimol, unpublished)

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3 Hydrogenosome Shape In some fungi such as Neocallimastix (Figs. 1a, 4, 5b) (Yarlett et al. 1986; Benchimol et al. 1997; van der Giesen 1997), in rumen ciliates (Fig. 1b) (Yarlett et al. 1981, 1984), in trichomonads (Figs. 1c,d, 2, 5, 8, 9a, 16) (Benchimol et al. 1996a), and in free-living ciliates (Finlay and Fenchel 1989) hydrogenosomes are spherical or slightly elongated granules. Using freezeetching, ﬁlaments are seen connecting hydrogenosomes to cytoskeletal structures such as the axostyle and costa structures (Benchimol et al. 2000). In some cells, hydrogenosomes are not spherical but are very elongated structures, such as those found in Monocercomonas (Diniz and Benchimol 1998), which can reach 2.0 µm in length (Fig. 2c).

Fig. 2 Routine preparation of a hydrogenosome (H) in Trichomonas vaginalis (a), Tritrichomonas foetus (b), and Monocercomonas sp. (c). Note that in T. foetus the hydrogenosome is spherical, enveloped by a double membrane (arrows), and presents a single peripheral vesicle, whereas in T. vaginalis several peripheral vesicles are seen surrounding the organelle (arrows) and in Monocercomonas the hydrogenosome is very elongated. ER, endoplasmic reticulum; N, nucleus. Bars = 100 nm. (From Benchimol 2001; Diniz and Benchimol 1998)

4 Hydrogenosome Size Hydrogenosome sizes vary according to the species or if the cell is under stress or drug treatment (Figs. 3, 13) (Benchimol 1999, 2001; Madeiro and Benchimol 2004). In trichomonads without drug treatment (Figs. 1, 2, 5a, 8, 9a, 16, 18), hydrogenosomes present an average diameter of 300 nm (Figs. 2a,b, 5a), but may reach 2 µm in Monocercomonas (Diniz and Benchimol 1998) (Fig. 2c) or when trichomonads are subjected to drug treatment. In this last situation when drugs such as metronidazol, hydroxyurea, cytochalasin, and others were used, abnormal or bizarre shapes and sizes were

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Fig. 3 Abnormal hydrogenosomes. Note that the hydrogenosomes (H) are not spherical as in the routine preparations. They are giant and present internal compartments and abnormal peripheral vesicles. Dense spots are seen in the hydrogenosomal matrix, which represent calcium deposits. When cells are submitted to stress conditions, such as incubation with ﬁbronectin (Benchimol 2001) or other drugs (Madeiro and Benchimol 2004), the hydrogenosomes present such abnormal shapes and sizes, reaching 2 µm. Enlarged vesicles and internal membranes are also seen (asterisks). Bars = 150 nm. (From Benchimol et al. 1996a)

found (Figs. 3, 13) (Benchimol 1999, 2001; Ribeiro et al. 2002; Mariante et al. 2003).

5 Hydrogenosome Components 5.1 The Hydrogenosome Envelope It has been shown that trichomonads (Figs. 2a,b, 5a, 7c, 15b) and fungal hydrogenosomes are enveloped by two unit membranes (Figs. 4, 5b) (Benchimol and De Souza, 1983; van der Giezen et al. 1997; Benchimol et al. 1997). These membranes are very thin and very closely apposed to each other (Figs. 2, 4, 5, 15b). As a general rule no space is observed between the two membranes. Each membrane has a thickness of 6 nm and presents a certain undulation (Benchimol and De Souza, 1983; Benchimol et al. 1996a). The presence of CaCl2 in the ﬁxation solution, as well as the use of reduced osmium, is important for good visualization of the two membranes (Benchimol and De Souza 1983; Benchimol et al. 1996a) (Figs. 2, 4, 5). When the sections are thicker the two membranes are hardly visualized. Conventional freeze-fracture replicas revealed the presence of the two membranes enveloping the hydrogenosomes presenting a different number and distribution of intramembranous particles (Fig. 7a–c) (Benchimol et al. 1996a; Benchimol 2001). Four fracture faces were identiﬁed: two concave faces representing the P faces of the outer and the inner membranes and two con-

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Fig. 4 Thin sections of the fungal hydrogenosome (H) from Neocallimastix frontalis. In a there is one hydrogenosome in the process of division and the inner hydrogenosomal membrane is in process of septum formation. Note that there is another hydrogenosome with a double membrane, which is not in process of division. In b all the hydrogenosomes are dividing, presenting internal septa. Bars = 100 nm. (From Benchimol et al. 1997)

vex faces representing the E faces of the two membranes (Fig. 7a,c). The P and E faces of the outer membrane were frequently found. Although no quantitative analyses were carried out, the P face seems to have a higher particle density than the E face. The E face of the inner membrane was observed only in a few cases (Benchimol et al. 1996a). Special arrangements of intramembranous particles (rosettes) were found only when isolated hydrogenosomes were freeze-fractured, after cell fractionation (Fig. 7b) (Benchimol 2000). Invaginations of the hydrogenosome membrane delimiting inner compartments were observed when trichomonads where drug treated (Figs. 2b, 3). Some of the compartments had the same morphology and electron density as the hydrogenosomal matrix, while others had a lower density and presented tubular structures (Benchimol et al. 1996a). Ciliate hydrogenosomes in Metopus and Cyclidium present internal membranes and look like mitochondria (Fenchel and Finlay 1995). In addition, these

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Fig. 5 Thin sections of a T. foetus (a) and a fungal (b) hydrogenosome (H) from Neocallimastix frontalis. Both hydrogenosomes are clearly enveloped by a double membrane (arrow in (a), arrowheads in (b)). Invaginations of the hydrogenosome membrane were occasionally observed (arrow in (a)). The black spot in (a) is a calcium deposit in the peripheral vesicle (asterisks). CW, cell wall. Bars = 100 nm. (Fig. 5a from Benchimol, unpublished; Fig. 5b, from Benchimol et al. 1997)

hydrogenosomes are also calcium stores and display a membrane potential, which are similar to features found in mitochondria (Biagini et al. 1997). 5.2 The Peripheral Vesicle A special compartment is found at the periphery of most hydrogenosomes (Figs. 2a,b, 5a, 6b, 7b,c, 8b,c, 9, 14a,c, 15). T. foetus hydrogenosomes present one or two peripheral vesicles (Figs. 2b, 5a), whereas T. vaginalis hydrogenosomes exhibit several vesicles at the organelle periphery (Fig. 2a). Thus, the number of these compartments can be useful in taxonomic studies (Benchimol, unpublished). The hydrogenosome peripheral vesicle varies in size and electron density. Morphometric analysis showed that it represents 8.6% of the whole organelle in T. foetus. The peripheral vesicle is surrounded along its full extension by two closely apposed unit membranes. The inner portion of this compartment is completely distinct from the hydrogenosomal matrix (Figs. 8c, 9b). In some preparations it appears empty (Fig. 6b), whereas in others it presents a certain content, such as when the cells are ﬁxed in a solution containing calcium (Figs. 2a,b, 5a, 9a, 14a,c, 15a,b) or when the cells are submitted to freeze-fracture and deep-etching (Fig. 8b). This compartment is occupied by electron-dense reaction products after various cytochemical detections for calcium (Figs. 2a,b, 5a), phosphatases such as acid phosphatase (Fig. 11b), Mg++ -ATPase, or 5 -nucleotidase (Queiroz et al. 1991).

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Fig. 6 Carbohydrates in hydrogenosomes. Monocercomonas sp. after the Thiéry technique (a). The hydrogenosomal membranes are positive for carbohydrates. b T. foetus cryosection labeled with gold-conjugated WGA. Hydrogenosome (H) shows that the membrane lining the peripheral vesicle, but not other portions of the organelle, is labeled. Bars = 100 nm. (Fig. 6a from Diniz and Benchimol 1998; Fig. 6b from Benchimol et al. 1996a)

Fig. 7 Freeze-fracture images of hydrogenosomes from T. foetus. a Fractured cell showing a prominent Golgi (G) with several lamellae and fenestrae, as well proﬁles of endoplasmic reticulum (ER) in close proximity (arrows) with hydrogenosomes (H). Bar = 100 nm. b Hydrogenosomes from an isolated fraction observed by freeze-fracture. Note the clusters of intramembranous particles forming rosettes (arrow). The peripheral vesicle is smooth and does not present clusters of particles or rosettes. Bar = 50 nm. (From Benchimol et al. 2001). c Two freeze-fractured hydrogenosomes exhibiting different fracture planes (arrows). Bar = 100 nm. (From Benchimol et al. 1996a)

Carbohydrates were cytochemically detected in hydrogenosome membranes using the periodic acid–thiosemicarbazide–silver proteinate technique (Fig. 6a) and gold-labeled lectins, such as WGA (Fig. 6b) (Benchimol

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Fig. 8 Different views of T. foetus hydrogenosomes (H) after ﬁeld-emission scanning electron microscopy (FESEM) (a) and freeze-etching (b,c). An isolated hydrogenosome obtained from T. foetus observed by FESEM, where details of its surface can be seen. b A calcium deposit in the peripheral vesicle (asterisk); c shows that the peripheral vesicle (arrow) presents a smooth surface, distinct from the organelle body. Bars = 50 nm. (From Benchimol 2000)

et al. 1996a; Benchimol and Bernardino 2002). The membrane of the peripheral vesicle compartment is intensely labeled by WGA-gold, indicating the presence of N-acetyl glucosamine (Fig. 6b). Interestingly, the membrane lining the outer portion of the hydrogenosome is not labeled (Benchimol et al. 1996). The membrane surrounding the peripheral vesicle is smooth when analyzed in freeze-fracture and deep-etched cells (Figs. 7c, 8c) and thus

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Fig. 9 a Hydrogenosomes isolated by Percoll–sucrose density centrifugation. The hydrogenosomes (H) are seen as spherical organelles presenting a peripheral vesicle. Bar = 300 nm. b Hydrogenosome peripheral vesicles (P) isolated from a pure hydrogenosome fraction (a), show that they form a distinct subcompartment. Bar = 100 nm. (Courtesy of Dr. José Andrés Morgado Díaz)

rather distinct from the whole organelle, which is rough (Fig. 8a,c). With ﬁeld-emission scanning electron microscopy (FESEM), the hydrogenosomal surface exhibits a distinct morphology with a rough surface (Fig. 8a) (Benchimol 2000). In a study by Díaz and De Souza (1997), the authors were able to purify a hydrogenosomal fraction (Fig. 9a) and then another hydrogenosomal subfraction containing only the peripheral vesicles (Fig. 9b), showing that it is a distinct compartment. The authors used Tritrichomonas and showed that a further treatment with proteinase K solubilized the matrix components, leaving a pure peripheral vesicle fraction (Fig. 9b). The isolated peripheral vesicles maintained their ﬂattened morphology, suggesting that each individual vesicle has its own inherent structural framework (Fig. 9b). After hydrogenosomal membrane solubilization, the hydrogenosome matrix appeared attached to the peripheral vesicle. SDS-PAGE showed that proteins of 66, 45, and 32 kDa were localized in the peripheral vesicle (Díaz and De Souza 1997). Western blot analysis revealed the presence of glycoproteins, with a major one of 45 kDa in the peripheral vesicle of the hydrogenosome. The authors concluded that the peripheral vesicle is a distinct hydrogenosomal compartment (Díaz and De Souza 1997). 5.3 The Matrix of the Hydrogenosome The hydrogenosome matrix is homogeneous, presenting a granular appearance, which is different from the cytoplasmic matrix (Figs. 1, 2, 5, 8b). The hydrogenosome matrix was described in previous studies as homogeneously granular, occasionally presenting a dense amorphous or crystalline core, also

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known as nucleoid (Fig. 10a) (Honigberg and Brugerolle 1990). Observations indicated that this core is not a usual structure, appearing however either when the protozoa are incubated in the presence of drugs or when good ﬁxation is not achieved (Benchimol et al. 1996). The electron-dense core is frequently seen in not-well-preserved cells, a situation in which the hydrogenosome proteins could coagulate and precipitate, leading to the formation of the core (Fig. 10a). In healthy or well-preserved cells (Figs. 1, 2, 5), as well as in no drug-treated cells, it is very unusual to ﬁnd this electron-dense amorphous core and thus it is not considered as a hydrogenosomal structure any more (Benchimol et al. 1996a). The hydrogenosome matrix exhibited basic proteins when cytochemistry for ammoniacal silver and PTA (phosphotungstic acid) (Fig. 10b) were used (Benchimol et al. 1982a). The granular structure of the hydrogenosome matrix may be clearly visualized in replicas of quick-frozen, freeze-fractured, deep-etched, and rotary-replicated cells (Figs. 8b, 18b). When the fracture plane exposed the internal portion of the hydrogenosome matrix a large number of particles were seen. Most of them had a diameter of 6 nm. Some, however, were larger, with a diameter of 20 nm. These particles were not randomly distributed; a certain orientation in their array was noted.

Fig. 10 a T. foetus showing a dense material in the hydrogenosomes (H) matrix, named core or nucleoid (arrow) in early literature. The nucleoid is considered a ﬁxation artifact or is a result of a disturbance in hydrogenosome metabolism, being a sign of organelle death (Benchimol, unpublished). Bar = 300 nm. b Cytochemistry for basic proteins (PTA), showing positive reaction in the hydrogenosomes matrix (H), in the axostyle (Ax), costa (C), and pelta (P). Bar = 300 nm. (From Benchimol et al. 1982a)

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Fig. 11 a T. foetus after cytochemistry for calcium using potassium pyroantimonate, showing an intense labeling in the hydrogenosome matrix (arrows). N, nucleus. Bar = 300 nm. (From Benchimol et al. 1982b). b Cytochemical detection of acid phosphatase presented positive reactions not only in lysosomes (L) but also in the peripheral vesicles of the hydrogenosomes (H). Bar = 100 nm. (From Benchimol et al. 2001)

Calcium deposits are seen as electron-dense spots in the matrix of some hydrogenosomes when cells are incubated in the presence of calcium ions in the ﬁxative (Fig. 3a) or processed cytochemically for the localization of Ca++ (Fig. 11a), such as with the pyroantimonate technique (Benchimol et al. 1982b). When cells are stressed with drugs, the hydrogenosomal matrix, which normally appears ﬁnely granular and homogeneous, frequently contains internal elements, such as concentric membranes, internal subcompartments, vesicles, grains, and ﬁbrils (Fig. 3). A large, dense precipitate or nucleoid (Fig. 10a) is sometimes observed in the hydrogenosome matrix, which means that the organelle is dysfunctional (Benchimol et al. 1996a).

6 Fungal Hydrogenosomes The hydrogenosomes of the anaerobic fungus Neocallimastix are round or elongated structures, always enveloped by two distinct but tightly apposed membranes (Figs. 4a,b, 5b) (van der Giezen et al. 1997; Benchimol et al. 1997). N. frontalis hydrogenosomes were studied in routine preparations for transmission electron microscopy, freeze-fracture, and immunocytochemistry. In addition, images of organelle division (Fig. 4b) were very similar to those observed in trichomonad protozoa (Fig. 15a). These observations suggested that hydrogenosomes are homologous organelles in unrelated species, weakening a previous hypothesis of a polyphyletic origin (Marvin-Sikkema et al. 1992)

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and reinforcing the hypothesis that fungal and trichomonad hydrogenosomes are derived from an ancestral endosymbiont.

7 Proximity with Other Cellular Structures Glycogen particles, although distributed throughout the protozoan, are concentrated in the region where hydrogenosomes are located (Figs. 1, 4b, 14a, 16a,b). 7.1 Hydrogenosomes and Endoplasmic Reticulum Not only continuity but also chemical similarities between hydrogenosomes and the endoplasmic reticulum (ER) have been demonstrated. The ER is closely associated, though not necessarily continuous, with mitochondria, peroxisomes (Frederick et al. 1968), and plastids. Data indicated that certain mitochondrial phospholipids were formed in ER and then transferred to the mitochondrion (Jungalwala and Dawson 1970). Since the hydrogenosome has been considered a modiﬁed mitochondria (Embley et al. 1997) and presents several similarities to this organelle, it has been suggested that at least the peripheral vesicle of the hydrogenosomes originated from the ER. The continuities or close associations between hydrogenosomes and the rough or smooth ER are often observable (Figs. 7a,c, 12a,b, 13a,b, 17a,b). Some hydrogenosomes present membranous cisternae projecting to the cytoplasm, conferring bizarre images of the organelle (Fig. 12a). In some cases, the hydrogenosomal outer membrane displayed attached granules, similar to ribosomes, when cells were treated with cytochalasin B. An intimate association of the hydrogenosome with the ER has been described (Benchimol et al. 1996a, Benchimol 1999). It has been proposed that the ER could provide new membranes for hydrogenosome growth, since this

Fig. 12 Thin sections of T. foetus showing that proﬁles of endoplasmic reticulum (ER) are seen in continuity (arrow) to the hydrogenosomes (H). Bars = 100 nm

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organelle enlarges before its division. The ER could participate by providing the membrane lipids. Three-dimensional reconstruction of a T. foetus in division (not shown) and in interphase demonstrated a close association of the membranous proﬁles of the ER and hydrogenosomes, in a similar way to mitochondria and peroxisomes (Franke and Kartenbeck 1971). Also it has been suggested that membranous structures seen in close contact with or in the vicinity of the hydrogenosomes might be a source of membrane lipids for hydrogenosome growth (Benchimol et al. 1996a).

8 Hydrogenosome Autophagy T. foetus under serum deprivation, drug treatment (hydroxyurea, zinc sulfate), and also in normal conditions presents autophagy (Benchimol 1999). Apparently normal hydrogenosomes, and also giant, abnormal hydrogenosomes and those presenting internal membranes, were observed in the autophagic process (Fig. 13). The ﬁrst event observed was the presence of cisternae of the rough ER surrounding and enclosing the altered hydrogenosome, forming an isolation membrane (Fig. 13a–c). The hydrogenosome is ﬁrst sequestered from the remaining cytoplasm (Fig. 13b,c) and then degraded within lysosomes (Fig. 13d). The autophagic vacuole is limited by double (Fig. 13c) or multiple concentric membranes and may contain a recognizable hydrogenosome (Fig. 13b–d), probably in the preliminary steps of degradation. Lysosomes fuse with the autophagic vacuole, forming a degradative structure bounded by a single membrane (Fig. 13d) and containing the hydrogenosome in various stages of degeneration. Hydrogenosomes have been found within lysosomes, partially degraded, forming hydrogenosomal remnants (Benchimol 1999).

9 Hydrogenosome Division Hydrogenosomes, as almost all other organelles, always grow by proliferation of preexisting organelles. Thus, each daughter cell will receive a complete set of organelles during cell division. Morphological evidence was presented showing that trichomonad hydrogenosomes, like mitochondria, may divide by three distinct processes: (1) segmentation (Fig. 14a,b), (2) heart-form division (Fig. 14c), and (3) partition (Fig. 15a,b). In the segmentation process, the hydrogenosome grows, becoming elongated with the appearance of a constriction in the central portion (Fig. 14a,b) (Benchimol et al. 1996b). Microﬁbrillar structures appear to help the furrowing process, ending with a total ﬁssion of the organelle. In the partition process, rounded hydrogenosomes, in a bulky form, are further separated by a membranous internal

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Fig. 13 Hydrogenosomes in process of autophagy. a View of a hydrogenosome from a T. foetus cell grown in a culture medium containing 4 mM hydroxyurea for 15 h. One giant hydrogenosome presenting an enlarged peripheral vesicle (PV) and internal membranes (arrows) is seen. Proﬁles of the endoplasmic reticulum (ER) are seen close to and surrounding the organelle. Bar = 80 nm. b Thin section of a hydrogenosome (H) enclosed by the rough endoplasmic reticulum (ER). Bar = 50 nm. c Observation of a double-layered autophagic vacuole (arrows), containing an intact hydrogenosome (H) after high-pressure freezing and freeze-substitution. The T. foetus cell was grown under normal conditions. Arrows point to the double membrane of the enclosing vacuole. Bar = 200 nm. d Routine preparation of T. foetus without any drug treatment. Several hydrogenosomes (H) are seen all around, while one of them (asterisk) is seen inside a vacuolar lysosome-like structure (V), suggesting an autophagic process. Bar = 200 nm. (From Benchimol 1999)

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Fig. 14 Views of the segmentation process (a,b) of a dividing hydrogenosome (H). a The organelle is elongated showing a constriction in the central region (arrows). Bar = 100 nm. (From Benchimol et al. 1996b). b Freeze-etching of a T. foetus hydrogenosome after quick freezing of living cells. Arrows point to a linear array of particles at the septum constriction. Bar = 200 nm. c T. foetus showing a hydrogenosome in the process of heart division. Bar = 100 nm. (From Benchimol and Engelke 2003)

septum (Fig. 15a,b). The division begins by an invagination of the inner hydrogenosome membrane, forming a transversal septum, separating the organelle matrix into two compartments (Benchimol et al. 1996b). A necklace of intramembranous particles delimiting the outer hydrogenosomal membrane in the region of organelle division was observed by freeze-etching (Fig. 14b). In the hydrogenosome heart-shaped process (Benchimol and Engelke 2003), the organelle gradually presents a membrane invagination on one side, leading to the organelle division (Fig. 14c). In this case, the hydrogenosome grows anteriorly and the process of division starts at one of the organelle poles, which

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Fig. 15 a Thin section of two hydrogenosomes (H) dividing via the partition process. The hydrogenosome (H) becomes larger and an invagination of the inner hydrogenosomal membrane is observed (arrowheads), gradually dividing the hydrogenosomal matrix in two compartments. Figure b shows that initially the inner membrane separates the hydrogenosome (H) in two compartments, but they are still joined by the outer hydrogenosomal membrane (arrows). Bars = 100 nm. (From Benchimol et al. 1996b)

becomes larger than the remaining organelle. Gradually, the membrane at this pole is seen inwarding. Although the organelle division begins with an inward furrowing, no septum is formed. The heart-shaped process is also distinct from segmentation, since in this process an elongation of the organelle occurs ﬁrst, giving a sausage shape and a progressive attenuation of its mid-region. In the heart-shaped process, the organelle neither forms a septum nor elongates and in this way is considered a new organellar division process. The most common form of division observed in T. foetus was the segmentation process, whereas partition is the most unusual division process observed in this protist. However, in the hydrogenosomes of the fungus Neocallimastix the most common division process was the partition (Benchimol, unpublished observations). On the other hand, in Trichomonas the heart shape was the most frequently observed division process (Benchimol, unpublished observations). It is important to point out that all three forms of division could be found during any phase of the cell cycle, and that the segmentation and heart form could be observed in the same cell. In addition, the organellar division is not synchronized (Benchimol and Engelke 2003). Interestingly, these three modes of division and timing were also described previously in mitochondria (Tandler and Hoppel 1973). It is not known why and how the cell chooses which form of hydrogenosomal division to perform. The mitochondrial division process has been compared with the division in bacteria, since the mitochondrial inner membranes are likely to be of bac-

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terial descent. In bacteria a FtsZ ring adheres to the inside of the bacterial membrane and constricts it to mediate division. The use of FtsZ homologues (van der Bliek 2000) and dynamin has been shown to participate in mitochondrial division (Koch et al. 2005). However, these proteins have not yet been studied in dividing hydrogenosomes. In hydrogenosomes, the process of division of the inner membrane is unknown, whereas the division of the outer membrane appears to be mediated by membranous proﬁles, probably of endoplasmic reticulum origin. These structures could aid the hydrogenosome division or participate in another function, such as providing membranes for hydrogenosome growth.

10 Hydrogenosome Behavior in the Cell Cycle Dividing hydrogenosomes can be found in all phases of the trichomonads cell cycle, similar to mitochondria (Suzuki et al. 1994). They were even observed during the mitotic process (Benchimol and Engelke 2003). During the inter-

Fig. 16 General aspect of T. foetus in interphase (a) and under division (b–d). Notice that in interphase the hydrogenosomes are aligned on the costa and axostyle (a), whereas during the division they are close to the nucleus (b–d). Bars = 300 nm. (Fig. 16a from Benchimol, unpublished; Fig. 16b,c from Benchimol and Engelke 2003; Fig. 16d from Ribeiro et al. 2000)

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phase the hydrogenosomes in trichomonads are distributed mainly along the axostyle and costa (Figs. 1c,d, 16a) and at mitosis onset the hydrogenosomes migrate to and around the nucleus (Fig. 16b,c). In nondividing cells, small hydrogenosomes (100–200 nm in diameter) were found together with normal sized organelles (about 300 nm). They were located mainly along the duplicated axostyle (Fig. 16b). During karyokinesis, hydrogenosomes follow the axostyle (Fig. 16d) (Benchimol and Engelke 2003; Ribeiro et al. 2000).

11 Hydrogenosomes Connection to Microtubules Delicate bridges were observed connecting the outer hydrogenosomal membrane to the microtubules of the axostyle. These bridges were also seen when fast-freezing and freeze-etching were used and could explain the hydrogenosome alignment along the axostyle. In addition, close proximity to free microtubules was also seen (Fig. 17), both in thin sections (Fig. 17a) and after quick-freezing followed by freeze-etching (Fig. 17b) (Benchimol, unpublished).

Fig. 17 Association of hydrogenosomes and microtubules in T. foetus. a Thin section of hydrogenosomes (H) in close proximity with microtubules (asterisks). b Freeze-etching after quick-freezing and rotatory shadowing showing a hydrogenosome in close association with cytoskeletal structures, probably microtubules. Bars = 200 nm. (Benchimol, unpublished)

12 Immunolabeling Hydrogenosomes can be labeled when speciﬁc antibodies anti-hydrogenosomal proteins are used, such as anti-malic enzyme (Fig. 18) or other proteins found in the hydrogenosomal matrix. Interestingly, similar labeling is also ob-

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Fig. 18 Immuno-electron microscopy showing detection of malic enzyme in the hydrogenosomes (H) of T. vaginalis. N, nucleus; bar = 100 nm. (Benchimol, unpublished)

served when antibodies anti-AP65, an anti-adhesin protein, is used. Alderete et al. (2001) explain this result as an example of molecular mimicry and functional diversity. Acknowledgements This work was supported by CNPq (Conselho Nacional de Desenvolvimento Cientíﬁco e Tecnológico), PRONEX (Programa de Núcleo de Excelência), FAPERJ (Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro), FENORTE (Fundação Estadual do Norte Fluminense), and AUSU (Associação Universitária Santa Úrsula).

References Alderete JF, Millsap KW, Lehker MW, Benchimol M (2001) Enzymes on microbial pathogens and Trichomonas vaginalis: molecular mimicry and functional diversity. Cell Microbiol 3:359–370 Benchimol M (1999) Hydrogenosome autophagy in Tritrichomonas foetus: an ultrastructural and cytochemical study. Biol Cell 91:165–174 Benchimol M (2000) Ultrastructural characterization of the isolated hydrogenosome in Tritrichomonas foetus. Tissue Cell 32:1–9

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Benchimol M (2001) Hydrogenosome morphological variation induced by ﬁbronectin and other drugs in Tritrichomonas foetus and Trichomonas vaginalis. Parasitol Res 87:215–222 Benchimol M, Bernardino MV (2002) Ultrastructural localization of glycoconjugates in Tritrichomonas foetus. Parasitol Res 88:134–143 Benchimol M, De Souza W (1983) Fine structure and cytochemistry of the hydrogenosome of Tritrichomonas foetus. J Protozool 30:422–425 Benchimol M, Engelke F (2003) Hydrogenosome behavior during the cell cycle in Tritrichomonas foetus. Biol Cell 95:283–293 Benchimol M, Elias CA, De Souza W (1982a) Tritrichomonas foetus: ultrastructural localization of basic proteins and carbohydrates. Exp Parasitol 54:135–144 Benchimol M, Elias CA, De Souza W (1982b) Ultrastructural localization of calcium in the plasma membrane and in the hydrogenosome of Tritrichomonas foetus. Exp Parasitol 54:277–284 Benchimol M, Almeida JCA, De Souza W (1996a) Further studies on the organization of the hydrogenosome in Tritrichomonas foetus. Tissue Cell 28:287–299 Benchimol M, Johnson PJ, De Souza W (1996b) Morphogenesis of the hydrogenosome: an ultrastructural study. Biol Cell 87:197–205 Benchimol M, Durand R, Almeida J (1997) A double membrane surrounds the hydrogenosomes of the anaerobic fungus Neocallimastix frontalis. FEMS Microbiol 154:277–282 Benchimol M, Diniz JAP, Ribeiro K (2000) The ﬁne structure of the axostyle and its associations with organelles in trichomonads. Tissue Cell 32:178–187 Biagini GA, Hayes AJ, Suller MTE, Winters C, Finlay BJ, Lloyd D (1997) Hydrogenosomes of Metopus contortus physiologically resemble mitochondria. Microbiology 143:1623– 1629 Díaz JAM, De Souza W (1997) Puriﬁcation and biochemical characterization of the hydrogenosomes of the ﬂagellate protist Tritrichomonas foetus. Eur J Cell Biol 74:85–91 Diniz JA, Benchimol M (1998) Monocercomonas sp. cytochemistry and ﬁne structure of freeze-fractured membranes. J Eukaryot Microbiol 45:314–322 Embley TM, Horner DA, Hirt RP (1997) Anaerobic eukaryote evolution: hydrogenosomes as biochemically modiﬁed mitochondria? Trends Ecol Evol 12:437–441 Embley TM, van der Gienzen M, Horner DA, Hirt RP, Dyal PL, Bell S, Foster PG (2003) Hydrogenosomes, mitochondria and early eukaryotic evolution. IUBMB Life 55:387– 395 Fenchel T, Finlay BJ (1995) Ecology and evolution in anoxic worlds. Oxford University Press, Oxford Finlay BJ, Fenchel T (1989) Hydrogenosomes in some anaerobic protozoa resemble mitochondria. FEMS Microbiol Lett 65:311–314 Franke WW, Kartenbeck J (1971) Outer mitochondrial membrane continuous with endoplasmic reticulum. Protoplasma 73:35–41 Frederick SE, Newcomb EH, Vigil EL, Wergin WP (1968) Fine-structural characterization of plant microbodies. Planta 81:229–252 (Berlin) Honigberg MB, Brugerolle G (1990) Structure. In: Honigberg BM (ed) Trichomonads parasitic in humans. Springer, New York, pp 5–35 Jungalwala FB, Dawson RMC (1970) Phospholipid synthesis and exchange in isolated liver cells. Biochem J 117:481–490 Koch A, Yoon Y, Bonekamp NA, McNiven MA, Schrader M (2005) A role for Fis1 in both mitochondrial and peroxisomal ﬁssion in mammalian cells. Mol Biol Cell 16:5077– 5086

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Kulda J, Nohýnková E, Ludvik J (1987) Basic structure and function of the trichomonad cell. Acta Univ Carol Biol 30:181–198 Lindmark DG, Müller M (1973) Hydrogenosome, a cytoplasmic organelle of the anaerobic ﬂagellate, Tritrichomonas foetus, and its role in pyruvate metabolism. J Biol Chem 248:7724–7728 Madeiro RF, Benchimol M (2004) The effect of drugs in Tritrichomonas foetus. Parasitol Res 92:159–170 Mariante RM, Guimarães CA, Linden R, Benchimol M (2003) Hydrogen peroxide induces caspase activation and programmed cell death in the amitochondrial Tritrichomonas foetus. Histochem Cell Biol 120:129–141 Marvin-Sikkema FD, Lahpor GA, Kraak MN, Gottschal JC, Prins R (1992) Characterization of an anaerobic fungus from llama faeces. J Gen Microbiol 138:2235–2241 Müller M (1993) The hydrogenosome. J Gen Microbiol 139:2879–2889 Queiroz RC, Santos LM, Benchimol M (1991) Cytochemical localization of enzyme markers in Tritrichomonas foetus. Parasitol Res 77:561–566 Ribeiro KC, Monteiro-Leal LH, Benchimol M (2000) Contributions of the axostyle and ﬂagella on the division process of Tritrichomonas foetus. J Eukaryot Microbiol 47:481– 492 Ribeiro KC, Vetö Arnholdt AC, Benchimol M (2002) Tritrichomonas foetus: induced synchrony by hydroxyurea. Parasitol Res 88:627–631 Rosa IA, Einicker-Lamas M, Bernardo RR, Previatto LM, Mohana-Borges R, Díaz JAM, Benchimol M (2006) Cardiolipin in hydrogenosomes: evidence of symbiotic origin. Eukaryot Cell 5:784–787 Snyers S, Hellings P, Bovy-Kesler C, Thines-Sempoux D (1982) Occurrence of hydrogenosomes in the rumen ciliates Ophryoscolecidae. FEBS Lett 137:35–39 Suzuki K, Ehara T, Osafune T, Kuroiwa H, Kawano S, Kuroiwa T (1994) Behavior of mitochondria, chloroplasts and their nuclei during the mitotic cycle in the ultramicroalga Cyanidioschyzon merolae. Eur J Cell Biol 63:280–288 Tandler B, Hoppel L (1973) Division of giant mitochondria during recovery from cuprizone intoxication. J Cell Biol 56:266–272 van Bruggen JJA, Zwart KD, van Assema RM, Stumm CK, Vogels GD (1984) Methanobacterium formicium, an endosymbiont of the anaerobic ciliate Metopus striatus McMurrich. Arch Microbiol 139:1–7 van der Bliek AM (2000) A mitochondrial division apparatus takes shape. J Cell Biol 151:F1–4 van der Giezen M, Sjollema KA, Artz RR, Alkema W, Prins RA (1997) Hydrogenosomes in the anaerobic fungus Neocallimastix frontalis have a double membrane but lack an associated organelle genome. FEBS Lett 408:147–150 Yarlett N, Hann AC, Lloyd D, Williams AG (1981) Hydrogenosomes in the rumen protozoan Dasytricha ruminantum Schuberg. Biochem J 200:365–372 Yarlett N, Coleman GS, Williams AG, Lloyd D (1984) Hydrogenosomes in known species of rumen entodiniomorphid protozoa. FEMS Microbiol Lett 21:15–19 Yarlett N, Orpin CG, Munn EA, Yarlett NC, Greenwood CA (1986) Hydrogenosomes in the rumen fungus Neocallimastix patriciarum. Biochem J 236:729–739

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_109/Published online: 8 January 2008 © Springer-Verlag Berlin Heidelberg 2008

Hydrogenosomes of Anaerobic Ciliates Johannes H. P. Hackstein1 (u) · Rob M. de Graaf1 · Jaap J. van Hellemond2,3 · Aloysius G. M. Tielens2,3 1 Department

of Evolutionary Microbiology, Faculty of Science, Radboud University Nijmegen, Toernooiveld 1, 6525 ED Nijmegen, The Netherlands [email protected] 2 Department of Biochemistry and Cell Biology, Faculty of Veterinary Medicine, Utrecht University, PO Box 80176, 3508 TD Utrecht, The Netherlands 3 Department of Medical Microbiology and Infectious Diseases, Erasmus MC, University Medical Center Rotterdam, PO Box 2040, 3000CA Rotterdam, The Netherlands 1

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Abstract Ciliates are highly complex unicellular eukaryotes. Most of them live in aerobic environments and possess mitochondria. However, in several orders of ciliates, anaerobic species evolved that contain “hydrogenosomes”, organelles that produce hydrogen and ATP. These hydrogenosomes of ciliates have not been studied in the same detail as those of trichomonads and chytrid fungi. Therefore, generalizations on the characteristics of hydrogenosomes of ciliates are somewhat premature, especially since phylogenetic studies suggest that hydrogenosomes have arisen independently several times in ciliates. In this chapter, the hydrogenosomes of the anaerobic, heterotrichous ciliate Nyctotherus ovalis from the hindgut of cockroaches will mainly be described as these are the ones that are, at the moment, the most thoroughly studied. Thus far, this is the only hydrogenosome known to possess a genome and this genome is clearly of mitochondrial origin. In fact, the hydrogenosome of N. ovalis unites typical mitochondrial features such as a genome and an electron-transport chain with the most characteristic hydrogenosomal property, the production of hydrogen. The hydrogenosomal metabolism of N. ovalis will be compared with that of two other ciliates that have been studied in less detail, i.e. the holotrichous rumen ciliate Dasytricha, and the free-living plagiopylid ciliate Trimyema. All studies combined indicate that it is likely that the various types of hydrogenosomes in ciliates evolved by modiﬁcations of aerobic mitochondria when the different ciliates

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adapted to anaerobic or micro aerobic environments. Furthermore, it is clear that the hydrogenosomes of anaerobic ciliates are different from those of chytrid fungi and from the well-studied ones in trichomonads.

1 Introduction Ciliates represent an extremely species-rich, monophyletic group of highly complex unicellular eukaryotes. They are characterized by a nuclear dimorphism and rather complex patterns of morphologically distinct, cortical cilia. Most of the ciliates thrive in aerobic environments and possess mitochondria, but anaerobic species evolved in at least 8 of the 22 orders of ciliates as classiﬁed by Corliss (1979), (Fenchel and Finlay 1995). Certain ciliates in seven of these 8 orders possess “hydrogenosomes”, organelles that produce hydrogen and ATP (Müller 1993; Hackstein et al. 1999, 2001; Embley et al. 2003; Dyall et al. 2004; Embley and Martin 2006, Embley 2006). However, the identiﬁcation of these hydrogenosomes was frequently based solely on the presence of symbiotic methanogenic archaea. Such an association is indicative of an inter-species hydrogen transfer and could reveal the presence of intracellular hydrogen sources, i.e. hydrogenosomes (Hackstein et al. 2002). The development of ﬂuorescence microscopy, electron microscopy, cytobiochemistry, and techniques for cellular fractionation soon allowed the discovery of hydrogenosomes in free-living anaerobic ciliates such as Plagiopyla and Trimyema (Plagiopylea), and Metopus (Armophorea) (van Bruggen et al. 1983, 1984, 1986; Goosen et al. 1988, 1990; Zwart et al. 1988; Finlay and Fenchel 1989; Fenchel and Finlay 1995; Biagini et al. 1997). Ciliates with hydrogenosomes were also identiﬁed in the gastro-intestinal tract of ruminants and marsupials (e.g. Isotricha, Dasytricha, Epidinium, Eudiplodinium, Polyplastron, Amylovorax; Trichostomatia) (Vogels et al. 1980; Snyers et al. 1982; Yarlett et al. 1981, 1982, 1983, 1984, 1985; Lloyd et al. 1989; Paul et al. 1990; Ellis et al. 1991a,b,c; Cameron and O’Donoghue 2002a) and in the hindgut of cockroaches (Nyctotherus; Armophorea) (Gijzen et al. 1991). See Fig. 1

Fig. 1 Neighbour-joining phylogenetic tree of 18S ribosomal RNA of ciliates. Ribosomal RNA sequences were aligned using Clustal X (Jeanmoughin et al. 1998) and phylogenetic trees were prepared by neighbour-joining (Saitou and Nei 1987). The accession numbers of used sequences and the bootstrap values for 1000 independent analyses are shown. Shaded boxes indicate anaerobic ciliates with hydrogenosomes, whereas all other ciliates contain mitochondria and function aerobically. The natural habitat of the hydrogenosome-containing ciliates is indicated by the following abbreviations: F, freeliving, HG, hindgut, R, rumen. Ciliate species that might possess mitosomes instead of hydrogenosomes are indicated by an asterisk. The length of the branches represents the relative divergence in nucleotide sequence. The bar represents 2% diversity

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for the distribution of hydrogenosomes and mitochondria in various orders of ciliates. All of these hydrogenosomes are surrounded by a double membrane, and under optimal ﬁxation conditions in a number of ciliates species, cristae-like protrusions can be seen in these organelles. In such a way, they

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Fig. 2 Electron micrograph of N. ovalis (a) with a hydrogenosome (b). The scale bar in (a): represents 10 micrometers, the scale bar in (b): represents 0.5 micrometers. H: hydrogenosomes, Ma: macronucleus, Mi: micronucleus, Cs: cytostome, PV: pulsating vacuole

clearly resemble mitochondria (Fig. 2). The hydrogenosomes of ciliates have not been studied in the same detail as those of trichomonads and chytrid fungi because culturing of these ciliates is difﬁcult (Müller 1993; Hackstein et al. 2001).

2 Nyctotherus ovalis Nyctotherus species (Armophorea) are anaerobic, heterotrichous ciliates with hydrogenosomes that thrive in the intestinal tract of cockroaches, millipedes, frogs and reptiles. N. ovalis from the hindgut of cockroaches is the only species that has been studied in more detail (van Hoek et al. 1998, 1999, 2000b). Notably, the presence of a mitochondrial genome has been demonstrated in the hydrogenosomes of N. ovalis (Akhmanova et al. 1998; van Hoek et al. 2000a, Boxma et al. 2005). This genome, which has been identiﬁed by immunocytochemistry, ﬂuorescence in situ hybridization, Southern blotting, cDNA analysis and DNA sequencing, is a typical mitochondrial genome (Boxma et al. 2005). Since the organelle unites the hallmark features of mitochondria (a DNA genome and an electron transport chain) and those of hydrogenosomes (hydrogen production), one has to conclude that this organelle represents a true “missing link” connecting the aerobic and anaerobic forms of mitochondrion-like organelles which are now recognized to form a rainbow of organelles with a wide spectrum of metabolic properties (Tie-

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lens et al. 2002; Martin 2005; Embley and Martin 2006; Hackstein et al. 2006). Since the phylogenetic analysis of the hydrogenosomal genome of N. ovalis unequivocally revealed a ciliate origin, its descent from an ancestral ciliate mitochondrion can be taken for sure. This ciliate origin is reinforced by the analysis of some 80 nuclear-encoded genes that encode mitochondrial proteins (Boxma et al. 2005; see below) 2.1 The Metabolism of N. ovalis In order to investigate their energy metabolism, isolated N. ovalis cells were incubated in the presence of tracer amounts of radioactive labelled glucose (labelled in all (C1 to C6) positions, or at position C6 only). These studies revealed that a small part of the glucose was degraded to typical end products of a glycolytic fermentation, as approximately 24% of the degraded glucose was excreted as lactate and 5% as ethanol (Boxma et al. 2005). The major part of the glucose was degraded via the hydrogenosomes as approximately 60% of the degraded glucose was excreted as acetate and 12% as succinate. Carbon dioxide was only produced in the incubations with uniformly labelled glucose and not in the incubations with [6-14 C]glucose. These results prompt several important conclusions. First, it implies that N. ovalis does not use a complete citric acid cycle for the degradation of glucose, as 14 C-labelled CO2 is released from [6-14 C]glucose exclusively by successive decarboxylations in subsequent rounds in the citric acid cycle (Tielens et al. 1992). Secondly, because formate is not one of the end products, N. ovalis does not use pyruvate formate lyase (PFL) activity in its pyruvate metabolism, as is the case in hydrogenosomes of anaerobic chytrids (see Hackstein, this volume). Thirdly, the product of glycolysis in the cytosol, pyruvate, is apparently either converted into lactate or ethanol, or is transported into the hydrogenosome to be converted into acetate or succinate. Fourthly, for the production of acetate, this pyruvate is decarboxylated, which is conﬁrmed by the release of 14 C-labelled carbon dioxide from incubations with uniformly labelled glucose. In principle, this decarboxylation could be performed either by a pyruvate dehydrogenase complex (PDH), as in “normal” mitochondria (see below), or by a pyruvate:ferredoxin oxidoreductase (PFO) that occurs in certain other types of hydrogenosomes. Fifthly, the excretion of signiﬁcant amounts of succinate indicates that endogenously produced fumarate is used as a terminal electron acceptor. Protons act as another hydrogenosomal electron acceptor, which results in the formation of hydrogen. Fumarate reduction is most likely catalysed by a membranebound fumarate reductase (an anaerobically-functioning variant of complex II), coupled to complex I of the electron transport chain via quinones. The reduction of fumarate requires the presence of rhodoquinone, which has a lower redox potential than ubiquinone, the quinone used by aerobic mito-

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chondria for the reverse reaction, the acceptance of electrons from succinate (Tielens et al. 2002). Consistent with these biochemical/biophysical requirements, small amounts of rhodoquinone 9 and menaquinone 8 were detected, whereas ubiquinone 7 and 8 (which are found in large amounts in the aerobic ciliates Euplotes and Tetrahymena, respectively) were not detected in N. ovalis (Boxma et al. 2005). 2.2 In Silico Reconstruction of the Basal Hydrogenosomal Metabolism of N. ovalis The signiﬁcance of these experimental (metabolic) data might be circumstantial without molecular (DNA sequence) data support (Boxma et al. 2005). Notably, no gene similar to PFO or PFL could be detected. However, genes for all three subunits of a PDH are present and are expressed. In addition, a gene was detected for acetyl-CoA synthase, an enzyme for the produc-

Fig. 3 Speculative metabolic schemes of the main pathways in carbohydrate metabolism in N. ovalis Abbreviations: AcCoA, acetyl-CoA, CI, complex I, Citr, citrate, FRD, fumarate reductase, FUM, fumarate, Hyd, hydrogenase, α-KG, α-ketoglutarate, MAL, malate, OXAC, oxaloacetate, PDH, pyruvate dehydrogenase, PEP, phosphoenolpyruvate carboxykinase, PYR, pyruvate, RQ, rhodoquinone, SUCC, succinate, SUCC-CoA, succinyl-CoA

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tion of acetate from acetyl-CoA, and also several genes, which are predicted to encode enzymes of the citric acid cycle, i.e. malate dehydrogenase, succinate dehydrogenase (2 subunits), succinyl-CoA synthetase, alphaketoglutarate dehydrogenase (2 subunits). Thus, basically the core energy (pyruvate) metabolism of a typical (ciliate) mitochondrion was detected, albeit in an anaerobic version (Fig. 3). Finally, although up to now no homologs of an Fo F1 -ATP synthase have been discovered, the hydrogenosome of N. ovalis has retained certain metabolic traits of anaerobic mitochondria. Therefore, it is likely that these anaerobic hydrogenosomes gain their energy by the generation of a PMF through proton-pumping by the mitochondrial complex I, just like anaerobic mitochondria. Currently, from the subunits of a mitochondrial complex I, 11 out of the 14 “core” ones were cloned and sequenced. This far, there is no evidence for genes encoding components of mitochondrial complexes III and IV, but these complexes are also absent in the electrontransport chains of anaerobic mitochondria (Tielens et al. 2002). Accordingly, imaging studies using inhibitors and ﬂuorescent dyes not only demonstrated the presence of a functional complex I as the source of a proton gradient in these hydrogenosomes, but also indicated the absence of functional complexes III and IV, and the absence of the plant-like alternative terminal oxidase (Boxma et al. 2005).

3 The Hydrogenosomes of Other Ciliates Metabolic studies have also been carried out on hydrogenosomes of rumen ciliates such as Dasytricha, Isotricha, Epidinium and Eudiplodinium. All rumen ciliates form a monophyletic group (Fig. 1; Strüder-Kypke et al. 2006), but not all of them possess hydrogenosomes, e.g. Entodinium simplex, Diploplastron afﬁne, Ophryoscolex caudatus, Eremoplastron bovis and Ostracodinium obtusum bilobum (Yarlett et al. 1984, 1985). Many rumen ciliates utilize cellulose and starch – besides being predators of bacteria and smaller protozoa. Glucose is the major monosaccharide liberated by degradation of these plant polymers and can be used by these rumen protozoa as fermentation substrate. The main end products of the metabolism of exogenously added glucose as well as of intracellular amylopectine of rumen ciliates with hydrogenosomes are hydrogen, acetate, lactate, butyrate and CO2 (Yarlett et al. 1985; Ellis et al. 1991a–c). The ratio in which these end products are formed is inﬂuenced by O2 and CO2 at those concentrations present in the rumen. The investigated rumen ciliates are able to use oxygen as terminal electron acceptor. The nature of this terminal oxidase is still unknown, but cytochromes do not appear to be involved. Dasytricha ruminantium is the most thoroughly studied rumen ciliate, however, even knowledge regarding

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Fig. 4 Speculative metabolic schemes of the main pathways in carbohydrate metabolism in Dasytricha sp. (after Ellis et al. 1991) Abbreviations: AcCoA, acetyl-CoA, Hyd, hydrogenase, PEP, phosphoenolpyruvate carboxykinase, PFO, pyruvate : ferredoxin oxidoreductase, PYR, pyruvate, Xox, red , unknown electron carrier

the metabolism of this rumen ciliate is still far from complete. The enzyme used for the degradation of pyruvate to acetyl-CoA in this protozoon is suggested to be PFO, which has been identiﬁed tentatively in the hydrogenosomal fraction (Yarlett et al. 1981, 1982, 1985). This acetyl-CoA is the substrate for the hydrogenosomal formation of acetate, but seems also to be exported from the hydrogenosomes for the formation of butyrate (Yarlett et al. 1985; Ellis et al. 1991b). A hypothetical scheme for the hydrogenosomal metabolism of the holotrich (rumen) ciliate Dasytricha that is based on various studies is shown in Fig. 4. The scheme is remarkable as it requires the export of acetyl-CoA from the hydrogenosome for the formation of butyryl-CoA. This butyryl-CoA is then used for the production of butyrate, which is accompanied by the production of ATP (Yarlett et al. 1985; Ellis et al. 1991a–c). These aspects make this hydrogenosome of rumen ciliates very different from that of Nyctotherus (Fig. 3) and also different from the hydrogenosomes of Trichomonas (see Hrdý et al., this volume). The only other published metabolic study on hydrogenosomes of ciliates deals with the free-living Plagiopylid ciliate Trimyema (Goosen et al. 1988, 1990). This free-living ciliate produces neither butyrate nor signiﬁcant amounts of succinate, as rumen cilates and N. ovalis, respectively, do.

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Fig. 5 Speculative metabolic schemes of the main pathways in carbohydrate metabolism in Trimyema compressum (after Goosen et al. 1990). End products are in boxes. Abbreviations: AcCoA, acetyl-CoA, Hyd, hydrogenase, PEP, phosphoenolpyruvate carboxykinase, PFL, pyruvate : formate lyase, PFO, pyruvate : ferredoxin oxidoreductase, PYR, pyruvate, Xox, red , unknown electron carrier

Trimyema consumes oxygen under micro-aerobic conditions and is reported to produce formate under such conditions as the major end product with minor amounts of acetate and lactate (Goosen et al. 1990). Under those micro-aerobic conditions, hydrogen and ethanol are not produced. Under strictly anaerobic conditions, however, ethanol is the main end product, while acetate, lactate, formate and hydrogen are then formed in minor amounts (Goosen et al. 1990). For easy comparison with the metabolism of the other hydrogenosome-containing ciliates (Fig. 3 and 4), a very speculative scheme of the carbohydrate metabolism of Trimyema is shown in Fig. 5. This pattern of anaerobic fermentation products resembles the one found in anaerobic chytridiomycete fungi from the gastro-intestinal tract of large herbivores (Boxma et al. 2004). These fungi perform a bacterial type mixed acids fermentation using PFL for the degradation of pyruvate instead of PDH or PFO, which is used by N. ovalis and Trichomonads, respectively. Even though no additional biochemical data are available and no cell fractionation studies have been performed, it is likely that the plagiopylids evolved a type of hydrogenosome that is clearly different from those of Nyctotherus and Dasytricha.

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4 Can the Methanogenic Symbionts Tell us More About the Origin and Function of Ciliate Hydrogenosomes Additionally, the nature of the methanogenic (endo)symbionts supports the conclusion that ciliates host different types of hydrogenosomes. While Nyctotherus and Metopus, but also Plagiopyla and Trimyema, host endosymbiotic methanogens, certain rumen ciliates seem to host episymbiotic methanogens. Whether this episymbiotic association is speciﬁc, and whether there is any rumen ciliate (except Dasytricha and Isotricha) with symbiotic methanogens is still a matter of debate (Fenchel and Finlay 1995; Tokura et al. 1999; Regensbogenova et al. 2004). Because the methanogens (regardless of being endo- or episymbiotic) rely on substrates provided by the host, the properties of the endosymbiont might provide some information about the metabolic characteristics of the host. The Vogels and Stumm group succeeded in cultivating a number of putative methanogenic endosymbionts from the anaerobic ciliates Metopus striatus, Metopus contortus, and Plagiopyla nasuta, from the amoeboﬂagellate Psalteriomonas vulgaris and the giant amoeba Pelomyxa palustris (van Bruggen et al. 1984, 1986, 1988; Goosen et al. 1988; see Fenchel and Finlay 1995 for more references and discussion). The conclusion from these studies was that many endosymbionts were similar if not identical to well-known free-living methanogens, e.g. Methanobacterium formicicum. Only the putative endosymbiont from Metopus contortus seemed to represent a new type of methanogen, i.e. Methanoplanus endosymbiosus. The latter host, Metopus, belongs to the same taxon as N. ovalis, which makes it likely that this ciliate (Metopus) possesses a similar mode of pyruvate metabolism. The metabolic properties of the methanogenic endosymbiont M. formicium suggested that these might be capable of using other substrates besides hydrogen and CO2 , e.g. formate (Dong et al. 1994). This argues again for metabolic diversity among ciliate hydrogenosomes. This metabolic diversity could provide hard arguments for multiple origins of the hydrogenosomes, but unfortunately, metabolic data of both hosts and symbionts are scarce.

5 Evolutionary Aspects There exists a rather broad agreement that the anaerobic ciliates evolved secondarily from aerobic ancestors since several ciliate taxa comprise both aerobic and anaerobic species. Phylogenetic studies indicate that hydrogenosomes have arisen independently at least three to four times in ciliates (Fig. 1: Clarke et al. 1993; Embley and Finlay 1994; Embley et al. 1995, 2003; Fenchel and Finlay 1995; Hirt et al. 1998; Hackstein et al. 2001, 2002). The existence in

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N. ovalis of a “missing link”, an organel with characteristics of anaerobic mitochondria as well as of hydrogenosomes, demonstrates that hydrogenosomebearing ciliates can evolve from mitochondriate ciliates (Martin 2005). Albeit that the patchy distribution of hydrogenosomes alone is not sufﬁcient to prove multiple, independent origins of ciliate hydrogenosomes, the existence of such a missing link like the hydrogenosome of N. ovalis provides a clear scenario for the evolution of hydrogenosomes from mitochondria. Apparently, hydrogenosomes in ciliates can evolve “easily” by evolutionary tinkering from mitochondria in the course of the adaptation of their hosts to anaerobic/microaerobic environments. This happened several times independently in the evolution of ciliates – at least 3 independent origins are supported by the existence of 3 different types of hydrogenosomes in the ciliates that have been studied so far. It has remained unclear until now whether or not all anaerobic ciliates possess hydrogenosomes, in particular those anaerobes that do not possess endosymbiotic methanogens. Theoretically, anaerobic ciliates might possess anaerobic mitochondria, hydrogenosomes or they could even have lost ATPgenerating organelles completely. In this case, they most likely host mitochondrial remnants, mitosomes, just like Giardia and Entamoeba spp., which are completely dependent on cytosolic reactions for the production of ATP (Fig. 6; see, e.g. Hackstein et al. 2006 for discussion). However, the presence of these elusive organelles has not been studied systematically and in more detail in ciliates thus far – with a few remarkable exceptions to be discussed below. It has already been addressed that (at least) among the rumen ciliates, species with mitosomes might exist since there is evidence that certain rumen ciliates, such as for example Entodinium simplex, Entodinium caudatum, Diploplastron afﬁne, Ophryoscolex caudatus, Eremoplastron bovis, and Ostracodinium obtusum bilobum, did not exhibit detectable hydrogenase activity in the particulate cell-fraction, and therefore, by deﬁnition do not contain hydrogenosomes (Yarlett et al. 1984). Also, electron microscopy did not reveal the presence of mitochondrial-shaped organelles or typical hydrogenosomes in certain species of rumen and marsupial gut ciliates; a systematic search for mitosomes, however, has not been performed (Williams and Coleman 1992, Cameron and O’Donoghue 2002b). The observation that also PFO and malate dehydrogenase (decarboxylating) activities (Yarlett et al. 1984) are not enhanced in the particulate cell – fraction, together with a low cytoplasmic hydrogenase activity might argue for the absence of hydrogenosomes, but potentially for the presence of mitosomes. However, until now, there are no additional data that could support this speculation. The adaptation to an anaerobic lifestyle with the aid of hydrogenosomes required the acquisition of an (oxygen-sensitive) hydrogenase. The evolution of fumarate respiration in N. ovalis shows that an adaptation to life in anaerobic environments can occur in steps – by evolutionary tinkering.

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Fig. 6 Cartoon interpreting the evolution of mitochondria and related organelles (H hydrogenosomes, MS mitosomes, MR mitochondrial remnants, MM modiﬁed mitochondria). The solid lines represent the evolutionary descent based on the presence of an organellar genome, the broken lines indicate the absence (loss) of a genome. The monophyly of the mitochondria has been derived from the phylogenetic analysis of more than 41 alpha-proteobacterial and 816 mitochondrial genomes (February 2006). Since the branching order is not yet resolved, the cartoon solely indicates the common origin by the “origin” in the centre. A later loss of the organellar genome is only indicated if additional data argue for a loss of the organelle genome after the diversiﬁcation of the hosts. The phylogenetic relationships between the hosts are still discussed controversially. Therefore, the smallest common denominator is used to display an unrooted “tree” of the most basic taxonomic arrangement agreed by most biologists: plantae, uniconta (animals and fungi), rhizaria, chromalveolata, excavata. No attempts were made to root this tree, nor to give any indication of a potential branching order. Certain green algae possess “normal” mitochondria, but express a plastidic hydrogenase under anaerobic conditions (“chlorophytes”). Substantial differences in metabolism have been established for the hydrogenosomes of Trichomonas, Piromyces/Neocallimastix, and the various ciliates. From Hackstein et al. 2006, modiﬁed

Once anaerobiosis could be tolerated by the invention of fumarate respiration, it became possible to acquire a hydrogenase. The [FeFe] hydrogenase of N. ovalis most likely has been obtained by lateral gene transfer from anaerobic (sulfate-reducing) bacteria. The peculiar 24 and 51 kD subunits of this complex hydrogenase are paralogous to the corresponding proteins of the mitochondrial complex I (which is functional in N. ovalis), and have a different (most likely beta proteobacterial) origin (Akhmanova et al. 1998; Boxma et al.

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2005). The acquisition of this hydrogenase obviously allows a ﬁne tuning of the NADH pool, which is crucial for the maintenance of homeostasis under anaerobic conditions. Thus, N. ovalis not only turns out to be a missing link, it also demonstrates that the adaptation to anaerobic environments can involve several steps to allow the evolution of multiple levels for the control of homeostasis. In recent years it has become clear that there are not only mitochondria that function as described in biochemical textbooks, but that many different types of mitochondria exist, which exhibit a large variety in metabolic properties (Tielens et al. 2002). It is now also clear that the same holds true for hydrogenosomes. Firstly, there are obvious metabolic differences between the hydrogenosomes of Trichomonads and those of chytrid fungi and of anaerobic ciliates. Secondly, substantial differences in metabolism also exist within the large community of anaerobic ciliates (Fig. 3, 4, and 5).

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_110/Published online: 5 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Metabolism of Trichomonad Hydrogenosomes Ivan Hrdý1 (u) · Jan Tachezy1 · Miklós Müller2,3 1 Department

of Parasitology, Faculty of Science, Charles University in Prague, Viniˇcná 7, 12844 Prague 2, Czech Republic [email protected]

2 The

Rockefeller University, 1230 York Avenue, New York, NY 10021, USA

3 Collegium

Budapest, Szentharomsag utca 2, 1014 Budapest, Hungary

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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The Hydrogenosomal Membrane . . . . . . . . . . . . . . . . . . . . . . .

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Energy Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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4 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8 4.9

Proteins of the Core Catabolic Pathway Pyruvate : Ferredoxin Oxidoreductase . Ferredoxin . . . . . . . . . . . . . . . . Hydrogenase . . . . . . . . . . . . . . . Malic Enzyme . . . . . . . . . . . . . . NADH Dehydrogenase . . . . . . . . . Succinyl-CoA : Acetate CoA Transferase Succinyl-CoA Synthetase . . . . . . . . Adenylate Kinase . . . . . . . . . . . . Metabolic Enzymes Not Detected . . .

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Amino Acid and Polyamine Metabolism . . . . . . . . . . . . . . . . . . . Glycine Decarboxylase Complex and Serine Hydroxymethyltransferase . . Polyamine Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Trichomonad hydrogenosomes are probably the best studied organelles of their kind to date. Their role in energy metabolism has been ﬁrmly established and their indispensable function in the assembly of iron–sulfur centers, the vital cofactors of certain proteins, has been discovered rather recently. In this chapter, we summarize the knowledge of trichomonad hydrogenosome biochemistry, particularly the energy-linked

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pathway, the oxygen-related biochemistry, and the iron–sulfur cluster assembly machinery. The structure of the proteins constituting the core pathway is dealt with in some detail. We also attempt to accommodate the ﬁndings of the Trichomonas vaginalis genome sequencing project into the metabolic scheme of the hydrogenosome.

1 Introduction Hydrogenosomes are deﬁned as eukaryotic organelles that produce molecular hydrogen, a deﬁnition that reﬂects a single biochemical property and does not imply any particular morphology or other common biochemical traits for the organelles from different organisms. Hydrogenosomes have been detected in various protists that live in anoxic and hypoxic habitats. The history of the evolutionary appearance of hydrogenosomes in different organisms is still a much debated topic that is discussed in other chapters of this volume (see Martin, this volume). Hydrogenosomes were ﬁrst detected in anaerobic or microaerophilic parabasalid ﬂagellates, trichomonads. The ﬁrst species to be explored was the cattle parasite, Tritrichomonas foetus (Lindmark and Müller 1973). Soon thereafter, these organelles were detected in the human parasite, Trichomonas vaginalis (Lindmark et al. 1975), and in a reptile parasite, Monocercomonas sp. (Lindmark and Müller 1974). Hydrogenosomes of other protists and fungi share the ability to form hydrogen as a metabolic end product, but differ from the trichomonad organelles in many aspects of their metabolism (see Hackstein et al., this volume). This circumstance warrants a separate discussion of metabolism and enzymology of trichomonad hydrogenosomes in this chapter. Although earlier studies concentrated more on T. foetus, most recent work was performed on the medically important T. vaginalis. In the present chapter, we also focus on T. vaginalis and refer to T. foetus only occasionally. This bias is also due to the fact that the genome of T. vaginalis has been completed recently, providing much additional, often unexpected, information on the putative metabolic properties of its hydrogenosome. Other trichomonad species have been studied much less extensively and will be disregarded in our presentation.

2 The Hydrogenosomal Membrane Trichomonad hydrogenosomes are bounded by two unit membranes that are more closely spaced than the two membranes surrounding typical mitochondria (Benchimol and De Souza 1983). The properties of this membrane have not been studied in detail. An early work reported the presence of phos-

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phatidylethanolamine, phosphatidylserine, phosphatidylcholine, and cardiˇ erkasovová et al. 1976). The olipin in the hydrogenosomes of T. foetus (C discovery of cardiolipin in the hydrogenosomal membrane was regarded as having particular signiﬁcance, since this phospholipid is present almost exclusively in bacterial and inner mitochondrial membranes, where it plays important functional and structural roles. This is considered additional evidence for the endosymbiotic origin of mitochondria. However, the presence of cardiolipin in the hydrogenosomes was later refuted (Paltauf and Meingassner 1982) and this conclusion became generally accepted. It was not until 2006 that the presence of cardiolipin in the hydrogenosomes of T. foetus was again reported (Rosa et al. 2006), but its potential function in hydrogenosomes remains unclear. No further information exists on the lipid composition of the hydrogenosomal membrane. Ubiquinone, a lipid-soluble coenzyme mainly participating in mitochondrial electron transport but also having additional functions has been identiﬁed at low levels (up to two orders of magnitude lower than in mitochondria-possessing cells) in extracts of T. foetus, but neither the subcellular localization nor the possible role of this coenzyme in T. foetus has been explored (Ellis et al. 1994a). The hydrogenosomal membrane displays selective permeability, thereby presenting an effective barrier to pyridine nucleotides and coenzyme A ˇ erkasovov et al. 1978; Lindmark and Müller 1973; Müller 1973; Steinbüchel (C and Müller 1986). Transport of metabolic substrates and products across the hydrogenosomal membrane remains to be studied, but isolated T. foetus hydrogenosomes were shown to readily accumulate both radiolabeled pyruvate and malate (our unpublished data). Data on the protein composition of T. vaginalis hydrogenosome membranes are restricted to two integral membrane proteins. One is Hmp31, a member of the mitochondrial carrier family (Dyall et al. 2000), which was shown to act as an ADP/ATP carrier in the related species Trichomonas gallinae. Unlike mitochondrial transporters with the same function, the trichomonad protein is insensitive to bongkrekic acid, the inhibitor of mitochondrial ATP/ADP carriers (Tjaden et al. 2004). The second membrane protein is Hmp35, a β-barrel protein with no known homologues and a putative pore-forming function (Dyall et al. 2003).

3 Energy Metabolism The only metabolic activity that has been for decades ascribed to the trichomonad hydrogenosome was related to energy metabolism. It has been unequivocally shown that, under anaerobic conditions, isolated intact hydrogenosomes produced roughly equimolar amounts of acetate, CO2 , and hydrogen from pyruvate in a process accompanied by substrate-level phos-

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phorylation of ADP to ATP (Fig. 1). These ﬁndings conﬁrmed the metabolic role of the organelle deduced from the enzymatic activities found in the organelle (Müller 1993; Steinbüchel and Müller 1986), and are now fully supported by the results of the T. vaginalis genome sequencing project. All but one gene coding for enzymes of the core pathway were identiﬁed, mostly in multiple copies and with clearly recognizable hydrogenosomal targeting signals at the amino termini of the translated gene products (Carlton et al. 2007). The single exception to the complete annotation of the pyruvatecatabolizing pathway is the acetate : succinate CoA transferase (see below). Moreover, using immunological, mass spectroscopy, and microsequencing methods, most enzymes of the core pathway now could be identiﬁed on SDS– polyacrylamide gels, showing that these proteins are the most prominent of all hydrogenosomal proteins (Fig. 2). This is not to say that hydrogenosomes do not have other, and indispensable, functions. In fact, their role in energy metabolism may not be the one which cannot be missed. Hydrogenosomes lacking their typical activities involved in ATP generation persist in trichomonad cells that exhibit extreme levels of resistance to the antitrichomonad drug metronidazole (induced in laboratory conditions by long cultivation of trichomonads under increasing drug pressure—see Kulda and Hrdý, this volume), a drug that is activated to its cytotoxic form in this organelle in wild-type cells. The fact that hydrogenosomes of such resistant organisms may not be energetically “dead” was suggested on the basis that they contain alternative 2-keto acid oxidoreductases (Brown et al. 1999). This issue has not been resolved yet. The true indispensable role of hydrogenosomes may be connected to their iron–sulfur (FeS) cluster assembly system (see below) that is located in them and that is shared between hydrogenosomes, mitochondria, and certain mitosomes. Other functions of the hydrogenosome are expected to be uncovered when data from the T. vaginalis genome project are complemented with biochemical studies. The substrates for the core catabolic pathway of hydrogenosomes originate from glycolysis in the cytosol. The main substrate is pyruvate. Malate is also utilized after it is oxidatively decarboxylated to pyruvate by malic enzyme inside hydrogenosomes. While pyruvate could be produced in T. vaginalis from phosphoenolpyruvate (PEP) by conventional pyruvate kinase (Mertens et al. 1992), pyruvate, orthophosphate dikinase (Slamovits and Keeling 2006) (www.tigr.org/tdb/e2k1/tvg/), and cytosolic NADP-speciﬁc malic enzyme (Doleˇzal et al. 2004) (apart from the activity of aminotransferases), malate is formed by reduction of oxaloacetate arising from PEP by the activity of PEP carboxykinase. The relative carbon ﬂux through the reactions starting from PEP is unknown (and may vary depending on conditions), as is the contribution of the two substrates to hydrogenosomal catabolism. It has been shown that malate can be excreted, rather than consumed by the organelle. This observation was made during a study of isolated hydrogenosomes under

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Fig. 1 Proposed metabolic map of the T. vaginalis hydrogenosome. Orange rectangles represent enzymes known to be involved in energy metabolism. Ferredoxins (Fdx, yellow ovals) donate electrons to hydrogenases which subsequently synthesize molecular hydrogen. In the presence of metronidazole (Mz, pink), Fdx transports electrons derived via oxidative decarboxylation of malate or pyruvate to the drug, resulting in its activation. Predicted protein activities, mostly uncovered by analysis of the genome sequence, are color-coded as follows: pink, iron–sulfur cluster assembly machinery and hydrogenase maturation; blue, oxygen scavenging system; yellow, amino acid metabolism; and green, protein translocation and maturation. Open circles located in the membrane indicate unidentiﬁed transporters that likely facilitate substrate and metabolite transport. Key to abbreviations: AK, adenylate kinase; Fdx, ferredoxin; GDC, glycine decarboxylase complex; Hy, hydrogenase; Hy?, putative fusion hydrogenase with NAD-binding domain; PFOR, pyruvate : ferredoxin oxidoreductase; SOD, superoxide dismutase; STK, succinate thiokinase (succinate CoA ligase, succinyl-CoA synthetase); SHMT, serine hydroxymethyl transferase; Trx, thioredoxin; TrxP, thioredoxin peroxidase; TrxR, thioredoxin reductase; THF-CH2 , N 5 ,N 10 -methylene tetrahydrofolate; ASCT, succinyl-CoA:acetate CoA transferase; oxidase?, A-type ﬂavodiiron protein may catalyze this reaction; HydG, HydF, and HydE, auxiliary maturases of Fe hydrogenase; IscS, cysteine desulfurase; IscU, FeS scaffold protein; IscA and Nfu, alternative FeS scaffold proteins; Hsp70, Hsp60, Hsp10, heat shock proteins; Jac1 and Mdj1, J-domain containing cochaperones; Mge, nucleotide exchange factor; HPP, hydrogenosomal processing peptidase; Hmp35, hydrogenosomal integral membrane protein; Tom?, putative translocase of the outer membrane; Tim23? the core translocase of the inner membrane; Sam50, the core translocase of the sorting and assembly machinery complex; Pam18, J-domain containing protein of the presequence translocase associated motor complex; MCF, mitochondrial carrier family protein (Hmp31, ATP/ADP carrier). Dashed lines represent presumed reactions that await experimental veriﬁcation. Modiﬁed from Carlton et al. 2007

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Fig. 2 SDS–polyacrylamide gel electrophoresis of T. vaginalis hydrogenosomes puriﬁed by isopycnic centrifugation on a Percoll gradient. 1 PFOR; 2 malic enzyme, 64-kDa hydrogenase, and Cpn 60; 3 succinyl CoA synthetase β subunit; 4 Hmp35; 5 succinyl CoA synthetase α subunit; 6 Hmp31 (ATP/ADP carrier); 7 adenylate kinase; 8 thiol peroxidase. Well-resolved but unmarked bands mostly belong to malic enzyme fragments or unknown proteins. Proteins were identiﬁed by mass spectroscopy. The 12% gel is stained with Coomassie Brilliant Blue R 250 (authors’ original)

an elevated concentration of CO2 , accounting for the lowered production of hydrogen (Steinbüchel and Müller 1986). Similarly, a high CO2 concentration was observed to lower the hydrogenosomal acetate and hydrogen production in T. vaginalis cultures and it was suggested that reductive carboxylation of pyruvate to malate occurs in the hydrogenosome under these conditions (Paget and Lloyd 1990). On the other hand, the thermodynamically favorable oxidative decarboxylation of malate by malic enzyme in the hydrogenosomes occurs in cultures of T. vaginalis strains that were forced to develop high resistance to metronidazole. These cells lack the enzymatic activities necessary to reductively activate the drug, primarily the activity of pyruvate : ferredoxin oxidoreductase (PFOR). However, this activity is not the only one in the hydrogenosomes that is able to reduce metronidazole. It has been shown that another pathway of metronidazole activation exists in T. vaginalis hydrogenosomes which is dependent on the activity of NADH dehydrogenase (see below) that reduces ferredoxin (the electron transporter directly involved

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in metronidazole reduction; see Kulda and Hrdý, this volume) and that uses NADH resulting from the activity of malic enzyme as a reductant. To develop high-level metronidazole resistance, T. vaginalis strains not only eliminate PFOR but also malic enzyme, showing that the enzyme functions in the direction of malate decarboxylation and toward the production of reducing power in the form of NADH (Hrdý et al. 2005; Rasoloson et al. 2002). Thus, it seems that, in vivo, the reaction of malic enzyme proceeds in both directions and is affected by the concentration of reactants, probably by the actual concentration of CO2 . Within the hydrogenosome, pyruvate is oxidatively decarboxylated to acetyl coenzyme A and CO2 (Fig. 1). This reaction is catalyzed by an iron– sulfur protein, PFOR, an enzyme typical of certain anaerobic and nitrogenﬁxing bacteria that otherwise occurs only in a few anaerobic eukaryotes. Electrons released from pyruvate are transferred to ferredoxin, a low molecular weight electron carrier protein. This is also an iron–sulfur protein, with a [2Fe2S] cluster. Reduced ferredoxin is reoxidized by [FeFe] hydrogenase in a reaction that reduces protons to molecular hydrogen. The energy of the thioester bond of acetyl-CoA resulting from the activity of PFOR is conserved in one molecule of ATP that is formed in two consecutive steps. First, acetate : succinate CoA transferase releases acetate and transfers the CoA moiety to succinate. The succinyl-CoA thus formed serves as a substrate for energy-conserving succinyl-CoA synthetase (ADP/GDP-forming, also known as succinate thiokinase (STK)) that catalyzes the phosphorylation of ADP (GDP) to ATP (GDP) while releasing CoA-SH and succinate, which reenter the catalytic cycle. Succinyl-CoA synthetase is the only enzyme of the mitochondrial Krebs cycle that is shared by the two organelles. The ubiquitous enzyme of energy metabolism, ATP : AMP phosphotransferase (adenylate kinase), is also present in T. vaginalis hydrogenosomes (Declerck and Müller 1987). When malate is used as a hydrogenosomal substrate, it is oxidatively decarboxylated to pyruvate and CO2 by malate dehydrogenase (decarboxylating; malic enzyme) (Fig. 1). This preferentially NAD-speciﬁc enzyme (Drmota et al. 1996) is by far the most abundant hydrogenosomal protein (Fig. 2). Utilization of malate results in the production of reduced pyridine nucleotide coenzyme NADH. In order to maintain the redox balance, the NADH needs to be reoxidized; however, the enzyme that would enable this has remained elusive for a long time. The activity that reduces the artiﬁcial electron acceptor methyl viologen at the expense of NADH, and presumably the same activity that transfers electrons between reduced ferredoxin and NAD+ , was identiﬁed in the mid-1980s (Thong and Coombs 1987; Steinbüchel and Müller 1986), but the enzyme itself was isolated only in 2004 with a surprising result. The NADH : ferredoxin (and methyl viologen, ferricyanide, and other electron acceptors) oxidoreductase activity was found to belong to a heterodimeric protein whose subunits turned out to be homologous to the 51-

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and 24-kDa subunits of respiratory complex 1 (also called NADH dehydrogenase or NADH : ubiquinone oxidoreductase) of mitochondria and aerobic bacteria. These two subunits are part of the hydrophilic peripheral arm of this most complicated respiratory complex and function as an electron-input site to the respiratory chain (Hrdý et al. 2004). The unexpected ﬁnding of complex 1 core subunits in trichomonad hydrogenosomes points to another trait shared by mitochondria and trichomonad hydrogenosomes (in addition to the similar mode of protein import and processing, see Dyall and Doleˇzal, this volume, and iron cluster assembly machinery, see below): NADH is recycled by homologous proteins in a similar way by the two organelles. The fate of electrons derived from NADH is markedly different, however. In mitochondria, the transfer of electrons to ubiquinone by respiratory complex 1 (and eventually to oxygen in complex 4) is linked to the formation of a proton gradient across the inner mitochondrial membrane that serves as a driving force for ATP synthesis by membrane-anchored F1 F0 ATP synthase. Hydrogenosomes lack the electron transport chain-driven extrusion of protons and ATP synthase. Instead, in addition to its role in the oxidation of pyruvate, they apparently recruit ferredoxin also as an acceptor of electrons derived from NADH, and may thus link the NADH oxidation to production of hydrogen by hydrogenase, for which reduced ferredoxin is a physiological electron donor. In addition, NADH may serve directly as the source of reducing power for hydrogen formation, since hydrogenase homologues with a carboxy-terminally located NAD(P)H-binding domain have been identiﬁed in the T. vaginalis genome. To our knowledge, the ability of complex 1 to transfer electrons to ferredoxin has not been reported for any other organism.

4 Proteins of the Core Catabolic Pathway The few proteins involved in the core catabolic pathway of hydrogenosomes represent a curious patchwork of enzymes that can be found only in a very limited number of eukaryotes and of enzymes with an almost ubiquitous distribution in eukaryotes. Of course, it is the presence of enzymes with limited distribution that makes the trichomonad hydrogenosome a hydrogenosome. The proteins in question have all been studied with enzymological methods and, with one exception, they have been sequenced. It has to be mentioned that earlier studies have already revealed the presence of multiple genes coding for most of these proteins, a point that was further supported by the analysis of the complete genome of T. vaginalis (Carlton et al. 2007). For some proteins, these multiple genes code for very similar isoforms, but in the case of others, the putative products are quite dissimilar and may have different functions.

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4.1 Pyruvate : Ferredoxin Oxidoreductase Pyruvate : ferredoxin oxidoreductase (PFOR, EC 1.2.7.1) is a hallmark enzyme of hydrogenosomes, together with hydrogenase. The protein catalyzes the reversible, CoA-dependent oxidative decarboxylation of pyruvate, releasing acetyl-CoA, CO2 , and two electrons. In T. vaginalis, the electron acceptor is [2Fe2S] ferredoxin. The reaction catalyzed by PFOR is formally identical to that of the pyruvate dehydrogenase complex (PDH) of mitochondria and aerobic bacteria (however, the reaction catalyzed by PDH is irreversible), but structurally the trichomonad enzyme has little in common with the multienzyme dehydrogenase complex. It is an oxygen-sensitive, 120-kDa-subunit dimeric protein that is relatively tightly associated with the hydrogenosomal membrane. It contains noncovalently bound thiamin pyrophosphate cofactor and [4Fe4S] clusters (Williams et al. 1987). The number of FeS clusters has not been determined for the trichomonad enzyme, but the homologous protein with identically spaced critical cysteine residues from Desulfovibrio africanus has six clusters per dimer (Chabriere et al. 1999). In addition to the homologues of similar structure that are present in other anaerobic eukaryotes, another homologue can be found in two other eukaryotes. This is the pyruvate : NADP oxidoreductase (PNO) from the mitochondria of Euglena gracillis (Inui et al. 1987) and from the cytosol of the parasitic apicomplexan Cryptosporidium ˇ trnáctá et al. 2006; Rotte et al. 2001). In these two organisms, parvum (C PNO possesses an additional carboxy-terminal NADPH-cytochrome P450 reductase domain that enables transfer of electrons from pyruvate to NADP+ . Annotation of the T. vaginalis genome identiﬁed seven different PFOR genes (one truncated at the carboxy terminus) (Carlton et al. 2007). At least two of them, PFOR A and PFOR B1, are transcribed. The hydrogenosomal targeting sequences of these two proteins are very short, consisting of only ﬁve amino acid residues, with a conserved arginine at the –3 position relative to the processing site (typically, the conserved arginine is in the –2 position) (Hrdý and Müller 1995a). Based on overall similarity, these seven sequences could be divided into two groups: (1) PFOR A, B1, B2, and E and (2) PFOR C and F, plus the relatively divergent and somewhat longer PFOR D (Carlton et al. 2007). Neither the expression status nor the physiological activity of homologues other than A and B1 is known. Some may possess the 2-keto acid oxidoreductase activity that has been noted in wild-type as well as metronidazole-resistant trichomonads (Brown et al. 1999). A thorough phylogenetic analysis revealed monophyly of all eukaryotic PFORs and clearly showed that closer homologues are eubacterial rather then archaebacterial proteins; however, no donor eubacterial group from which the eukaryotic sequences originated could be identiﬁed with sufﬁcient support (Horner et al. 1999). A pharmacological implication of the presence of PFOR in T. vaginalis should be brieﬂy mentioned. PFOR is the key enzyme in the activation of

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antimicrobial drugs from the group of 5-nitroimidazoles, such as metronidazole. These drugs are reduced and activated into their cytotoxic form by the enzymatic systems that utilize low redox potential electron transporters, as are FeS proteins like PFOR and ferredoxins. Such pathways are absent from aerobic organisms, which makes these drugs both highly selective and effective. Short-lived radicals resulting from the reduction of the nitro group of 5-nitroimidazoles are very reactive, inﬂicting cellular damage and subsequent cell death (Kulda, 1999; and see Kulda and Hrdý, this volume). 4.2 Ferredoxin The low molecular weight iron–sulfur protein, ferredoxin, is the key electron transport component of hydrogenosomes. Trichomonad ferredoxin is a small, approximately 10-kDa, soluble protein. It is the univalent electron transporter that links PFOR and NADH dehydrogenase with hydrogenase. This protein was the ﬁrst hydrogenosomal protein to be puriﬁed and characterized (Marczak et al. 1983), the ﬁrst whose gene was cloned and sequenced (Johnson et al. 1990), and the only one whose crystal structure has been determined (Crossnoe et al. 2002). It was also the ﬁrst hydrogenosomal protein on which the presence of a short amino-terminal targeting signal has been recognized and its function suggested (Johnson et al. 1990). When the primary structure of T. vaginalis ferredoxin was established, it came as a surprise that it belonged to [2Fe2S]-type ferredoxins, which are usually found in aerobic bacteria and mitochondria. It was expected that the hydrogenosomal ferredoxin would be of the 2[4Fe4S] type, the one that is common in hydrogen-producing anaerobic bacteria and is also present in Entamoeba histolytica (Huber et al. 1988; Reeves et al. 1980) and Giardia intestinalis (Townson et al. 1994). The latter species, however, possesses multiple ferredoxins of both types (http://gmod.mbl.edu/perl/site/giardia14). For a long time it was thought that T. vaginalis possessed only a single ferredoxin gene. This view was challenged when it was found that the ferredoxin knockout did not result in decreased susceptibility of transformed trichomonads to metronidazole, since ferredoxin is believed to play a critical role in drug activation (Land et al. 2004). Shortly thereafter, seven ferredoxin genes were recognized in the T. vaginalis genome, all with predicted hydrogenosomal target peptides (Carlton et al. 2007). Dominantly expressed is ferredoxin 1, the one that was puriﬁed from hydrogenosomes (Gorrell et al. 1984) and the one ﬁrst sequenced (Johnson et al. 1990). Based on overall similarity, T. vaginalis ferredoxins could be assigned to two groups, ferredoxin 1–3 and ferredoxin 4–6, plus the more distant and somewhat larger ferredoxin 7. Ferredoxin 7 is the only other Trichomonas ferredoxin for which the expression status is known: it is not only expressed (albeit apparently at a lower level) along with ferredoxin 1 in wild-type trichomonads, it is

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also expressed (even at a somewhat higher level) in extremely metronidazoleresistant cells, suggesting that this particular ferredoxin plays an important (and different from ferredoxin 1) role in the metabolism of hydrogenosomes and is not directly involved in metronidazole activation (our unpublished data). Recent phylogenetic analysis showed that ferredoxins most similar to T. vaginalis ferredoxins 1–3 are found in Tritrichomonas foetus and in the oomycete Phytophthora soaje. Proteins from G. intestinalis, red algae of genus Cyanidioschyzon, and also from the hyperthermophilic eubacterium Aquifex are also similar to T. vaginalis ferredoxins (our unpublished data). 4.3 Hydrogenase The second hallmark enzyme of hydrogenosomes is the hydrogenase (hydrogen : ferredoxin oxidoreductase, EC 1.12.7.2). This enzyme catalyzes the reversible reduction of protons yielding molecular hydrogen. Based on the metal composition of their diatomic active site, the hydrogenases can be divided into two broad classes: [NiFe] hydrogenases possessing one nickel and one iron atom, and [FeFe] or Fe-only hydrogenases possessing two atoms of iron in the active site. A third class of enzymes has a different active site composition and reaction mechanism, and has a very limited distribution among a few methanogenic Archaea. Representatives of the ﬁrst two classes occur in a wide variety of microorganisms, the more prevalent [NiFe] enzymes being found in both Bacteria and Archaea, while the fewer representatives of [FeFe] hydrogenases are limited to Bacteria, a few anaerobic eukaryotes, and green algae (Meyer, 2007). A CO- and oxygen-sensitive 64-kDa hydrogenase was partially puriﬁed from T. vaginalis with the conclusion that the protein belongs to the family of [FeFe] hydrogenases (Payne et al. 1993). Subsequently, three hydrogenase genes have been identiﬁed and two of them sequenced, describing the putative proteins as being 63% identical, approximately 51.5- and 50-kDa [FeFe] hydrogenases. Western blotting demonstrated the localization of at least the larger one (but likely both) in hydrogenosomes (Bui and Johnson 1996). While the quaternary structure has not been determined for any of these proteins, it seems reasonable to expect that they are monomers, as are the majority of [FeFe] hydrogenases characterized so far (Meyer 2007). Analysis of the T. vaginalis genome revealed an extraordinarily large set of up to nine [FeFe] hydrogenase homologues, six of them with conserved all cysteine residues that are known to constitute the hydrogen-activating center consisting of the [4Fe4S] cluster and binuclear [2Fe] center (H cluster) (Meyer 2007; Vignais et al. 2001). All genes possess signature motifs implicating the presence of additional FeS clusters (two to four, depending on the particular gene) upstream of the H cluster, but not all of them have a clearly recog-

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nizable hydrogenosomal targeting sequence. Of the hydrogenases identiﬁed in early studies, the 64-kDa protein is encoded by two similar genes and possesses binding motifs for one [2Fe2S] and three [4Fe4S] clusters in the amino-terminal part of the sequence. The approximately 50-kDa hydrogenase is also encoded by two genes and likely binds two auxiliary [4Fe4S] clusters. These four genes possess predicted hydrogenosomal targeting presequences. In three sequences out of nine, the cysteine 1 of the H cluster, believed to act as an acid/base near the active site (Meyer 2007), is missing, raising the question of the possible function of these putative proteins. Interestingly, two of the sequences lacking cysteine 1 possess an additional, carboxy-terminally located domain with FAD and NAD(P) binding motifs (Carlton et al. 2007), which is similar to NADPH-cytochrome P450 reductase and to the carboxyterminal domain of pyruvate : NADP oxidoreductases of Euglena gracillis and ˇ trnáctá et al. 2006; Rotte et al. 2001). The number Cryptosporidium parvum (C and structural diversity of T. vaginalis hydrogenases suggests that the production of hydrogen by this parasite could be a much more complex process than originally thought. While the assembly of accessory FeS clusters of these hydrogenases likely requires the activity of a more universal Isc system (Meyer 2007; and see below), assembly and maturation of the complex active site of [FeFe] hydrogenases depends on the activity of additional specialized proteins. Hydrogenase maturases, as they are called, or Hyd proteins, have been described to catalyze the maturation of [FeFe] hydrogenase in the green alga Chlamydomonas reinhardtii and are invariably found in all prokaryotes possessing [FeFe] hydrogenases (Posewitz et al. 2004). All of these proteins, HydE, HydF, and HydG, have recently been identiﬁed in T. vaginalis hydrogenosomes (Pütz et al. 2006). Characterization of the maturases from T. maritima revealed that HydE and HydG are radical-SAM proteins and that HydF is a GTPase; they all contain FeS clusters (Rubach et al. 2005). Comparative evaluation of the domain structure and phylogenetic reconstruction of the sequences of [FeFe] hydrogenases still leave the challenging question of the origin and history of this enzyme in eukaryotic organisms wide open (Meyer 2007). 4.4 Malic Enzyme Malic enzyme (malate dehydrogenase (decarboxylating), EC 1.1.1.39) catalyzes reversible oxidative decarboxylation of malate to pyruvate. The enzyme uses NAD+ as an electron acceptor, but it is also able to utilize NADP+ with lower afﬁnity (Drmota et al. 1996). With a subunit size of approximately 63 kDa, the Trichomonas hydrogenosomal malic enzyme belongs to the family of large, eukaryotic type of malic enzymes. In contrast, the approximately 40-kDa-subunit malic enzyme, located in the cytosol, belongs

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to the prokaryotic-type malic enzymes and was apparently acquired by lateral gene transfer (Doleˇzal et al. 2004). Hydrogenosomal malic enzyme is a membrane-associated tetramer (Drmota et al. 1996). T. vaginalis encodes nine similar hydrogenosomal malic enzyme genes (three sequences in the database are truncated) (Carlton et al. 2007), of which at least two (Drmota et al. 1996; Hrdý and Müller 1995b), but likely more (our unpublished proteomic data), are expressed. Malic enzyme is the most abundant hydrogenosomal protein (Fig. 2) and since its activity is easy to follow by a spectrophotometric assay, it is used as a marker protein for trichomonad hydrogenosomes. Because the membrane of hydrogenosome is impermeable to NAD+ , the isolated and osmotically protected hydrogenosomes usually display rather low malic enzyme activity in vitro. Upon detergent treatment or after several cycles of freezing and thawing, the membrane is permeabilized and the total activity can be measured. The ratio of this liberated malic enzyme activity to the activity of intact hydrogenosomes is called latency and is taken as a measure of the physiological state of the hydrogenosomal membrane. 4.5 NADH Dehydrogenase NADH dehydrogenase (NADH : ubiquinone oxidoreductase, complex 1, EC 1.6.5.3) is the most complicated multisubunit complex of the respiratory chain. It consists of 13–14 subunits in bacteria and of over 40 subunits in typical mitochondria (Gabaldon et al. 2005). The complex catalyzes the oxidation of NADH coupled to the reduction of membrane-soluble ubiquinone (coenzyme Q). The transfer of electrons along the redox centers (FMN, FeS clusters) within the complex is linked to the extrusion of protons that form an electrochemical gradient across the inner mitochondrial (or bacterial) membrane. For a long time, it was assumed that trichomonad hydrogenosomes did not contain any components of the classical respiratory chain. It was only recently demonstrated that two proteins of complex 1 are present in T. vaginalis hydrogenosomes. These are a 47-kDa protein (homologue of mitochondrial 51-kDa subunit or NuoF in bacterial terminology), with a primary structure that is suggestive of the presence of one [4Fe4S] cluster and an FMN cofactor, and a 22-kDa protein (homologue of 24-kDa mitochondrial subunit or NuoE) that likely binds one [2Fe2S] cluster (Dyall et al. 2004; Hrdý et al. 2004). The T. vaginalis complex puriﬁes as an active heterodimer. The genome analysis did not reveal any other subunits of complex 1 with signiﬁcant conﬁdence, making the Trichomonas enzyme the most reduced remnant of complex 1 seen so far. Experimental data suggest that the physiological electron acceptor is likely a [2Fe2S] ferredoxin, although the activity of the complex 1 homologue with other electron transporters like FeS ﬂavoproteins cannot be currently ruled out (Hrdý et al. 2004). The trichomonad protein is insensitive

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to rotenone, a complex 1 inhibitor, which is not surprising as the rotenonebinding subunits are not present. T. vaginalis encodes for two similar copies of the 47-kDa subunit, but expression has only been conﬁrmed for one so far. The 22-kDa subunit is encoded by a single gene (Carlton et al. 2007). No subunits of other respiratory complexes were found in the T. vaginalis genome. Interestingly, a somewhat less reduced complex 1 has been observed in hydrogen-producing mitochondria of the ciliate Nyctotherus ovalis, with seven identiﬁed subunits (Boxma et al. 2005). 4.6 Succinyl-CoA : Acetate CoA Transferase Succinyl-CoA : acetate CoA transferase (acetate/succinate CoA transferase (ASCT), EC 2.8.3.8) catalyzes the transfer of the CoA moiety between acetate and succinate and produces the hydrogenosomal end product, acetate. This activity was ﬁrst detected in the hydrogenosomes of T. foetus in the mid-1970s (Lindmark 1976) and subsequently in T. vaginalis as well (Steinbüchel and Müller 1986). However, the enzyme has not been puriﬁed or characterized in any detail, nor has it been sequenced. During the preliminary analysis of the T. vaginalis genome, no genes were annotated as ASCT, but four similar genes were annotated as putative members of the acetyl-CoA hydrolase/transferase family, with easily recognizable hydrogenosomal targeting sequences on their amino termini. It remains to be determined whether these putative proteins are in fact acetyl-CoA hydrolases or CoA transferases. The closer homologues of these proteins appear to be hydrolase-type enzymes. Since the presence of a transferase activity instead of the energy-wasting hydrolase activity is characteristic of the hydrogenosome, these genes could in fact code for the transferases. It is expected that this issue will soon be resolved, as the protein in question is under investigation. At any rate, ASCT is the last enzyme of the principal hydrogenosomal catabolic pathway that awaits assignment to a particular gene. The presence of enzymes with similar function has been recognized in anaerobic mitochondria of kinetoplastid protists and helminths (Tielens et al. 2002), but little is known about their overall distribution. 4.7 Succinyl-CoA Synthetase Succinyl-CoA synthetase (SCS), also known as succinate thiokinase (STK) or succinate CoA ligase (EC 6.2.1.4–5), is so far the only known hydrogenosomal enzyme directly involved in energy conservation. The protein catalyzes the reversible, substrate-level phosphorylation of ADP or GDP to the respective triphosphate at the expense of the high-energy thioester bond of succinylCoA. Succinate and CoA are released in the reaction. The T. vaginalis enzyme

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consists of two subunits, α (32.5 kDa) and β (43 kDa), which constitute the active heterotetramer located in the hydrogenosomal matrix. The enzyme is apparently able to utilize both ADP and GDP as phosphate (Jenkins et al. 1991; Lahti et al. 1992, 1994). Three similar genes encoding either subunit are present in the T. vaginalis genome; all three encoding the α subunit and at least two of the β subunit genes are expressed (Lahti et al. 1992, 1994). 4.8 Adenylate Kinase Adenylate kinase (ATP : AMP phosphotransferase, EC 2.7.4.3) is a ubiquitous central enzyme of energy metabolism, catalyzing the reversible transfer of a phosphate group between two molecules of ADP, forming ATP and AMP. An adenylate kinase has been puriﬁed and characterized from T. vaginalis hydrogenosomes (Declerck and Müller 1987). The enzyme shares some properties with cytosolic and some with mitochondrial homologues. The T. vaginalis genome encodes several (ten, one truncated at the amino terminus) adenylate kinase homologues (Carlton et al. 2007). Based on the presence of a stretch of 27 amino acid residues in the carboxy-terminal half of the proteins, only two of these sequences belong to the longer, mitochondrialtype eukaryotic adenylate kinases, while the rest belong among the shorter, cytosolic-like ones. One of the two longer sequences is equipped with a typical hydrogenosomal targeting signal, and this sequence corresponds to the protein that was previously puriﬁed and characterized (Lange et al. 1994). 4.9 Metabolic Enzymes Not Detected During the biochemical characterization of trichomonad hydrogenosomes, certain components characteristic of mitochondria and peroxisomes were looked for but the results were negative within the detection limits of the methods used. These components were cytochromes (Lloyd et al. 1979b) and the F1 F0 ATPase (Lloyd et al. 1979a). These negative results supported the conclusion that trichomonad hydrogenosomes and typical mitochondria markedly differ in their energy metabolism.

5 Interaction with Oxygen T. vaginalis is usually regarded as an anaerobic organism for which oxygen concentrations higher than those encountered in situ in the vagina (above ≈60 µM) are toxic (Ellis et al. 1994b). On the other hand, it was observed that very low oxygen concentrations (less than 0.25 µM) actually

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have a positive effect on the growth of trichomonad cultures (manifested as shorter doubling time), and it was therefore suggested to regard T. vaginalis as a microaerophile (Paget and Lloyd 1990). This stimulatory effect of traces of oxygen might be possibly explained on the basis of cytosolic redox balance: NADH produced during glycolysis would be reoxidized by the oxygen-reducing NADH oxidase and not through the activity of lactate dehydrogenase, which would allow more pyruvate to enter the hydrogenosome and provide additional ATP. Despite being fermentative organisms that lack cytochromes, mitochondrial-type electron transport, and therefore the ability to carry out oxidative phosphorylation, trichomonads do take up oxygen at a high rate when it is present in their environment. If the oxygen concentration is high enough to saturate the cytosolic oxygen reductases, the gas diffuses into the hydrogenosome and instead of protons it assumes the role of terminal electron acceptor in the hydrogenosomal metabolism. Production of hydrogen stops (Lloyd and Kristensen 1985), but the composition of metabolic end products remains largely the same as under anaerobic conditions; only the ratio of hydrogenosomal and cytosolic end products may shift in response to actual oxygen and CO2 concentrations. It was observed that acetate prevails under microaerobiosis, while glycerol and lactate formed in the cytosol become dominant when, together with traces of oxygen, CO2 is present at a high (ca. 5 mM) concentration (Paget and Lloyd 1990). Two pyridine nucleotide-speciﬁc dehydrogenases are responsible for oxygen reduction in the cytosol: a highly active NADH oxidase that reduces oxygen to water (Linstead and Bradley, 1988; Tanabe, 1979) and a minor NADPH oxidase that produces hydrogen peroxide (Linstead and Bradley 1988). The situation in the hydrogenosome is less clear. The fact that hydrogenosomes are able to utilize oxygen as a terminal electron acceptor with high activity was already recognized in the pioneer times of the 1970s. When incubated under aerobic conditions, hydrogenosomes isolated from T. foetus respired actively when supplied with α-glycerophosphate, the main hydrogenosomal substrate pyruvate, or malate together with ˇ erkasovov et al. 1978; Müller 1973). While the respiration with NAD+ (C α-glycerophosphate was not studied any further, respiration with pyruvate was found to be dependent on the availability of ADP, CoA, and a catalytic amount of succinate, all compounds necessary in the oxidative decarboxylation of pyruvate coupled to substrate-level phosphorylation in the principal hydrogenosomal catabolic pathway. That the observed respiration was not due to the activity of the standard respiratory chain was obvious, since rotenone and cyanide, the inhibitors of mitochondrial electron transport and respiration, had no effect. Hydrogenosomal respiration differed from that of mitochondria in another important aspect: its rate declined rapidly and the oxygen uptake soon became irreversibly inhibited, indicating damage to ˇ erkasovov et al. 1978; Müller components of the oxygen-reducing pathway (C

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1973; Müller and Lindmark 1978). The putative terminal oxidase has not been identiﬁed, but involvement of a ﬂavoprotein that produces harmful hydroˇ erkasovov et al. 1978). However, as has gen peroxide has been proposed (C already been suggested (Docampo et al. 1987), it is quite possible that autooxidation of iron–sulfur clusters of proteins involved in pyruvate oxidation, namely PFOR and/or ferredoxin, is responsible for the high respiration rate at the atmospheric oxygen concentration. The resulting products of incomplete oxygen reduction, such as superoxide anion, hydrogen peroxide, and the highly toxic hydroxyl radical formed from superoxide anion and hydrogen peroxide in the presence of iron ions through Fenton chemistry, could account for the inactivation of sensitive hydrogenosomal components, and could be responsible for the toxicity of elevated nonphysiological levels of oxygen to trichomonads. The key hydrogenosomal enzymes hydrogenase and PFOR are sensitive proteins prone to inactivation by oxygen (Lindmark and Müller 1973; Lloyd and Kristensen 1985). Obviously, hydrogenosomes need to be equipped with defense mechanisms to combat the damage caused by oxygen and reactive oxygen species formed either enzymatically or upon contact of oxygen with reduced ﬂavines and FeS clusters. The almost ubiquitous superoxide dismutase (SOD), which converts two molecules of superoxide radical into hydrogen peroxide and oxygen, is present in the cytosol and, to a lesser extent, in the hydrogenosomes of T. foetus (Lindmark and Müller 1974), and the same, albeit lower, activity has also been detected in T. vaginalis (Ellis et al. 1994b). However, in the human parasite the subcellular localization of the enzyme has not been studied. Seven genes encoding iron-containing SODs (FeSODs) have been found in T. vaginalis by classical molecular methods. The T. vaginalis SOD is a dimeric protein composed of 22-kDa subunits, which is similar to the homologues from parasitic protists like Trypanosoma or Entamoeba and to the proteobacterial sequences (Viscogliosi et al. 1998). The number of SOD genes was conﬁrmed by the T. vaginalis genome project (Carlton et al. 2007); however, none of the genes was found to contain the typical amino-terminal hydrogenosomal targeting sequence. Nevertheless, a SOD protein spot has recently been identiﬁed by mass spectrometry on 2D gels of carefully puriﬁed hydrogenosomes, conﬁrming the presence of this defense protein in the hydrogenosomes (Pütz et al. 2005). The presence of a SOD certainly accounts for the observed hydrogen peroxide formation in the hydrogenosomes under aerobic conditions, possibly together with reduced ﬂavines (Chapman et al. 1999). However, a catalase that removes hydrogen peroxide has been detected in T. foetus, but not in T. vaginalis (Page-Sharp et al. 1996). Proteomic studies of isolated organelles and data mining of the genome show the presence of additional enzymatic systems that could play a role in the defense of hydrogenosomes against oxygen and reactive oxygen species. This plethora of putative defense systems is of great interest in view of the known relative oxygen tolerance of T. vaginalis, a property that is difﬁcult to

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ascribe to the presence of SOD alone. It has to be stressed that biochemical conﬁrmation of these activities and their cellular function is still outstanding, thus the whole area of oxygen-related biochemistry and oxidative stress management in the hydrogenosome is still open to future exploration. A possible means to cope with the deleterious effects of peroxide buildup was proposed recently, following the ﬁnding of two putative peroxidereducing enzymes, rubrerythrin and peroxiredoxin, by a proteomic approach in T. vaginalis hydrogenosomes (Pütz et al. 2005). The ﬁrst of these, rubrerythrin, is a diiron-center and rubredoxin-like center-containing protein (Jin et al. 2002) with peroxidase-like activity, so far known only from anaerobic prokaryotes and E. histolytica, where a homologous sequence is present (Pütz et al. 2005). Several rubrerythrin genes have been identiﬁed in the T. vaginalis genome database; however, none of them possesses the typical amino-terminal hydrogenosomal targeting sequence. The amino terminus of the mature, hydrogenosome-located protein has not been determined, so it is unknown whether it is processed upon import into the organelle. Nevertheless, at least one of the proteins is localized in the organelle, as veriﬁed by immunoﬂuorescence microscopy using a hemagglutinin epitope on overexpressed recombinant rubrerythrin (Pütz et al. 2005). The second candidate peroxide-reducing protein of T. vaginalis hydrogenosomes is bacterial-type thiol-dependent peroxidase, a member of the ubiquitous peroxiredoxin family of proteins that contain active cysteine residues and participate in peroxide detoxiﬁcation and sensing (Chae et al. 1994; Wood et al. 2003). They likely play a major role in peroxide protection in anaerobic organisms that lack catalase and glutathione peroxidase, including helminths, trypanosomatids, and Plasmodium falciparum (McGonigle et al. 1998; Müller et al. 2003). The function of peroxiredoxin is dependent on its reduction by a small protein thioredoxin that itself is reduced by NADPH-dependent thioredoxin reductase (McGonigle et al. 1998). These three proteins form the typical peroxide detoxifying peroxiredoxin system. The presence of this system has been demonstrated previously in the cytosol of T. vaginalis (Coombs et al. 2004) and recent evidence shows its possible presence in hydrogenosomes. A thiol peroxidase homologue that is distinct from the reported cytoplasmic protein has been identiﬁed by mass spectroscopy in hydrogenosomes and two similar genes coding for this protein are present in the T. vaginalis genome. Similar to SOD and rubrerythrin, this thiol peroxidase does not feature a characteristic hydrogenosomal target signal (Pütz et al. 2005), nor is it known whether the amino terminus of any of these proteins is processed. The cysteine-regenerating thioredoxin has been identiﬁed by mass spectroscopy in the hydrogenosome as well. In this instance, the corresponding sequence does contain the amino-terminal extension with strong similarity to known hydrogenosomal targeting sequences. The ﬁrst protein of the catalytic cascade, the thioredoxin reductase, has not yet been found in hydrogenosomes, but ﬁve open reading frames

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coding for a low molecular weight-type thioredoxin reductase are present in the T. vaginalis genome. Comparison to the cytosolic thioredoxin reductase (Coombs et al. 2004) shows that three of these contain amino-terminal extensions which, however, do not resemble typical hydrogenosomal target signals (Pütz et al. 2005). Another protein potentially involved in oxidative stress protection, which was identiﬁed in the hydrogenosomes by a proteomic approach, shows similarity to the OsmC and Ohr proteins that so far have only been detected in bacteria. These proteins are involved in organic hydroperoxide detoxiﬁcation. Similar to the structurally unrelated peroxiredoxins, these proteins directly use highly reactive cysteine thiol groups to reduce hydroperoxide substrates (Lesniak et al. 2002, 2003). The function of the hydrogenosomal protein is unknown, but the crucial catalytically active cysteine residues are conserved in all four putative homologues found in the T. vaginalis genome. The amino termini of these proteins show similarity to hydrogenosomal target peptides and the protein isolated from hydrogenosomes is processed at the predicted site at position –2 relative to the conserved arginine residue (our unpublished data). A fourth protein detected in T. vaginalis hydrogenosomes and likely involved in their protection is a member of the ﬂavodiiron protein superfamily. These proteins are widespread among anaerobic or facultative anaerobic Eubacteria as well as Archaea. They were ﬁrst proposed to provide protection against oxygen by safely reducing it to water (Chen et al. 1993), but it was later recognized that they also possess considerable nitric oxide reductase activity and therefore are likely important in protection against nitrosative stress (Gomes et al. 2002). These proteins are functional dimers that share a common structural core, consisting of an amino-terminal β-lactamase-like domain with a diiron center and of a carboxy-terminal ﬂavodoxin domain that noncovalently binds FMN. The simplest and most common representatives, designated as Class A ﬂavodiiron proteins, consist of only this two-domain core; Class B and Class C proteins possess additional domains in the carboxyterminal part of the molecule that participate in electron transfer (Saraiva et al. 2004). The protein identiﬁed in T. vaginalis hydrogenosomes belongs to the Class A subfamily; it is a dimer and binds FMN like its prokaryotic counterparts. Its amino terminus displays a typical hydrogenosomal targeting signal peptide that is cleaved from the mature protein (our unpublished data). The electron donor for the T. vaginalis ﬂavodiiron protein remains to be identiﬁed. The rubredoxin that donates electrons to the ﬁrst characterized member of this group, the rubredoxin : oxygen oxidoreductase from Desulfovibrio vulgaris (Chen et al. 1993), is not present in T. vaginalis (Carlton et al. 2007). It has been suggested that an FeS ﬂavoprotein with presumed electron transporting function may be involved in the reduction of the ﬂavodiiron protein in Methanosarcina acetivorans (Saraiva et al. 2004). Since putative FeS ﬂavoproteins with predicted hydrogenosomal target peptides have also

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been identiﬁed in the T. vaginalis genome (www.tigr.org/tdb/e2k1/tvg/), it is possible that some of these are linked to the ﬂavodiiron protein. It seems reasonable to expect that the function of T. vaginalis ﬂavodiiron protein is either oxygen or NO (or both) reduction, as seen in its prokaryotic homologues. Either function would be highly relevant. In the case of oxygen reduction, it would provide a safe way of oxygen disposal; in the case of NO reductase activity, it would provide protection against nitrosative stress that is likely to be encountered by trichomonads in situ upon confrontation with the host immune system. It is noteworthy that genes coding for homologous proteins could be detected in the genomes of other anaerobic unicellular parasites, like Giardia (http://gmod.mbl.edu/perl/site/giardia14), Spironucleus (Andersson et al. 2007), and Entamoeba (Andersson et al. 2006; Loftus et al. 2005), and a free-living relative of Entamoeba, Mastigamoeba balamuthi (Andersson et al. 2006).

6 Iron–Sulfur Cluster Assembly Machinery Iron–sulfur clusters are ubiquitous inorganic cofactors which are required for the biological function of a number of proteins in various cell compartments. Although the formation of some FeS clusters in apoproteins could be achieved by noncatalytic chemical reconstitution in the living cell, this process is mediated by a complex FeS cluster (ISC) assembly machinery. Eukaryotes inherited this machinery from the bacterial endosymbiont which gave rise to mitochondria and related organelles, such as hydrogenosomes and mitosomes. In fact, formation of FeS clusters is the only common function which is shared by mitochondria, hydrogenosomes, and mitosomes, supporting their common origin (see Tachezy and ˇSmíd, this volume). Several recent reviews summarize mechanisms of FeS cluster assembly in eukaryotes and bacteria (Johnson and Dean 2004; Lill and Mühlenhoff 2005, 2006; Rouault and Tong 2005). In principle, FeS clusters are ﬁrst assembled on a scaffold protein and they are subsequently transferred to acceptor apoproteins. The ﬁrst step requires the activity of the IscS/Isd11 complex, which is a cysteine desulfurase. This enzyme provides sulfur for the scaffold protein IscU. Another component, IscA, may serve either as an alternative scaffold protein or for maturation of more complex clusters. Delivery of iron for FeS cluster assembly, as well as for heme synthesis, is most likely facilitated by mitochondrial frataxin. The chaperone system, including Hsp70, J-type cochaperone, and ADP/ATP nucleotide exchange factor, was suggested to participate in the second step in which FeS clusters are transferred to apoproteins. The assembly of an FeS cluster also requires reducing equivalents which are provided by [2Fe2S] ferredoxin. Genes coding for all of the components of ISC machinery were identiﬁed in the T. vaginalis genome (Carlton et al. 2007); however,

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experimental studies on the hydrogenosomal FeS cluster assembly machinery are rather limited. Initially, two genes coding for IscS were characterized in T. vaginalis and hydrogenosomal localization of the gene products was predicted based on the presence of amino-terminal targeting presequences (Tachezy et al. 2001). The hydrogenosomal localization of the iscs gene product in hydrogenosomes was later conﬁrmed in isolated organelles as well as by immunoﬂuorescence microscopy (Sutak et al. 2004). This ﬁnding strongly suggested that ISC assembly operates in hydrogenosomes (Fig. 1). Indeed, the ability of isolated organelles to catalyze assembly and insertion of an FeS cluster into apoprotein was demonstrated (Sutak et al. 2004). Frataxin is another protein which was shown to be targeted into hydrogenosomes (Doleˇzal et al. 2007). Importantly, T. vaginalis frataxin can, in part, functionally replace mitochondrial frataxin, as demonstrated by its ability to partially restore defects in FeS cluster assembly in Saccharomyces cerevisiae ∆yfh1 mutants and frataxin-deﬁcient Trypanosoma brucei (our unpublished data). In yeast, T. vaginalis frataxin also partially restored defects in heme synthesis, although neither heme-containing proteins nor components involved in heme synthesis have been identiﬁed in T. vaginalis (Doleˇzal et al. 2007). Interestingly, the transcription of genes coding for hydrogenosomal IscS as well as frataxin was markedly upregulated under iron deﬁciency, which might reﬂect an increased demand for the synthesis of new FeS clusters (Doleˇzal et al. 2007; Sutak et al. 2004). This is in direct contrast to what is found with yeast mitochondrial frataxin (yfh1), the expression of which is strongly stimulated by iron (Santos et al. 2004). The observed differences may reﬂect differences in the function of frataxin in various organisms. For example, Gakh et al. (2002) proposed that frataxin may serve as a mitochondrial ironstorage molecule, for which upregulation of transcription by iron could be expected. According to our transcriptional data, T. vaginalis frataxin is unlikely to have this function in hydrogenosomes. As mentioned above, sequencing of the complete T. vaginalis genome revealed the presence of seven [2Fe2S] ferredoxin homologues. It is likely that some of them serve as electron donors for FeS cluster biogenesis, while others are required for electron transport associated with the energy metabolism of hydrogenosomes. The mitochondrial ISC assembly machinery is essential not only for maturation of FeS proteins within mitochondria, but also for extramitochondrial FeS proteins (Lill and Kispal 2000). Although the nature of compounds exported from mitochondria is not known, the formation of extramitochondrial FeS clusters is dependent on ISC export machinery, which involves the ABC transporter Atm1, sulfhydryl oxidase Erv1, and glutathione, and the cytosolic components P-loop NTPases Cfd1, Nbp35, hydrogenase homologue Nar (Narf), and WD40 repeat domain protein (Lill and Mühlenhoff 2006). It is noteworthy that no candidates for a member of the ISC export machinery

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were identiﬁed in the T. vaginalis genome, although homologues of cytosolic components are present. Thus, it would be interesting to investigate whether, similarly to mitochondria, hydrogenosomes are involved in the maturation of FeS proteins outside of hydrogenosomes and, if so, which membrane components are involved in this process.

7 Amino Acid and Polyamine Metabolism Analysis of the T. vaginalis genome (Carlton et al. 2007) provided a number of unexpected results. One of these was the detection of two enzymes with a possible role in amino acid metabolism. Amino acid metabolism in T. vaginalis was regarded as essentially restricted to arginine decarboxylation and some transaminations. Hydrogenosomes were assumed to have no enzymes involved in amino acid metabolism. Surprisingly, genes encoding proteins involved in the interconversion of glycine and serine, glycine decarboxylase complex (GDC) and serine hydroxymethyltransferase (SHMT), were detected. 7.1 Glycine Decarboxylase Complex and Serine Hydroxymethyltransferase GDC is a multienzyme complex composed of four loosely associated component enzymes: the P protein (a homodimer containing pyridoxal phosphate), the H protein (a monomeric lipoamide-containing protein), the T protein (a monomeric protein requiring tetrahydrofolate cofactor), and the L protein, a dihydrolipoamide dehydrogenase (a homodimer containing FAD). In the T. vaginalis genome (www.tigr.org/tdb/e2k1/tvg/), only genes coding for the H protein (two genes) and a single gene for the L protein have been identiﬁed. Amino termini of both proteins bear similarity to hydrogenosomal target peptides. The messages for all three proteins are transcribed, both H and L proteins were detected in the hydrogenosomes of the wildtype trichomonads by Western blotting, and the overexpressed, plasmidencoded, and hemagglutinin-tagged proteins have been shown to localize to hydrogenosomes of transformed trichomonads. The recombinant L protein could utilize either H protein as a substrate in vitro (Mukherjee et al. 2006a). In T. vaginalis, the single gene coding for SHMT (Carlton et al. 2007) has a presequence that is somewhat different from the typical hydrogenosomal targeting peptides, but nevertheless possesses a mitochondrial-like processing site. The closest homologues of T. vaginalis SHMT are of the mitochondrial type, but the T. vaginalis enzyme was found to be a dimer, based on sucrose density gradient analysis of overexpressed protein, like its bacterial homologues rather than a tetramer as is observed in mitochondria (Mukherjee et al. 2006b). In addition to identifying the presence of overexpressed and

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tagged SHMT in T. vaginalis hydrogenosomes by immunoﬂuorescence, the corresponding (albeit low) enzymatic activity was also detected in the hydrogenosomes of wild-type trichomonads (Mukherjee et al. 2006b). In other organisms, the function of these two enzymes is the interconversion of serine and glycine, an essential and ubiquitous step in primary metabolism. It not only produces serine from glycine (and vice versa), both precursors for amino acid and phospholipid biosynthesis, but, most importantly, it provides the activated one-carbon units bound to tetrahydrofolate that are utilized in a number of biosynthetic pathways, such as the biosynthesis of methionine, pyrimidines, and purines. In other words, the role of the GDC–SHMT pathway is to interconnect the metabolism of one-, two-, and three-carbon compounds (Bauwe and Kolukisaoglu 2003). Eukaryotic GDC is an exclusively mitochondrial enzyme which catalyzes the oxidative decarboxylation and deamination of glycine, thereby producing CO2 and NH3 and concomitantly reducing NAD+ coenzyme to NADH. The remaining methylene carbon of glycine is transferred to tetrahydrofolate, forming N 5 ,N 10 -methylene tetrahydrofolate, which serves as a donor of a one-carbon unit in the reaction of SHMT leading to the formation of serine from another molecule of glycine. SHMT is a pyridoxal phosphate-dependent enzyme that catalyzes reversible interconversion of serine and tetrahydrofolate to glycine and N 5 ,N 10 -methylene tetrahydrofolate, the carbon carrier (Schirch and Szebenyi 2005). The two enzymes are interconnected through a common soluble pool of tetrahydrofolate. The presence of these two enzymes in T. vaginalis hydrogenosomes indicates that this organelle plays a so far unexpected role in amino acid metabolism. Two aspects of these results present a major conundrum, however. GDC, as a typical decarboxylase complex, consists of several enzymes that are necessary for its function; however, no genes were detected that encode for two of the obligate components, the P enzyme that is the glycine decarboxylating subunit and the T enzyme that transfers the methylene group to tetrahydrofolate. Of course, the two proteins might be sufﬁciently divergent to escape recognition, but, if they are really absent, the putative function of the complex is difﬁcult to visualize. Moreover, dihydrofolate reductase, the enzyme necessary to convert the imported folate to tetrahydrofolate, the active coenzyme, could not be detected in the T. vaginalis genome database, making the usual functional interconnection of SHMT with GDC through the tetrahydrofolate pool and the activity of SHMT itself a problem that will require future research. 7.2 Polyamine Metabolism One reaction of arginine metabolism also apparently takes place in the hydrogenosomes. Arginine is a precursor in the biosynthesis of polyamines, the

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ubiquitous polycations that participate in pivotal cellular functions, such as macromolecule biosynthesis and cell growth and differentiation (Pegg 1986). Due to their positive charge at physiological pH, polyamines are thought to electrostatically bind to phospholipids, nucleic acids, and proteins and affect their stability and conformation (Feuerstein et al. 1990). In trichomonads and Giardia, arginine is metabolized to the polyamine putrescine by a bacterial-type arginine dihydrolase pathway in which arginine is ﬁrst converted to citrulline by arginine deiminase. Citrulline is then converted to ornithine and carbamylphosphate by catabolic ornithine carbamyl transferase and putrescine is formed from ornithine by ornithine decarboxylase. Carbamylphosphate is a macroergic molecule that is broken down to bicarbonate and ammonia by carbamate kinase, which concomitantly phosphorylates one molecule of ADP to ATP (Yarlett et al. 1994). This pathway is probably a signiﬁcant source of ATP to the parasite in situ, as evidenced by rapid replacement of arginine with putrescine in the vaginal ﬂuid of infected patients (Chen et al. 1982), but when abundant glucose is available, as in in vitro cultures, the arginine dihydrolase pathway adds only about 1% of extra ATP to that generated from carbohydrates (Yarlett et al. 1996). Localization of enzymes of the arginine dihydrolase pathway has been studied in T. vaginalis as well as in T. foetus. Almost half of the arginine deiminase activity was found to localize to the sedimentable fraction, while the other enzymes of the pathway were present predominantly in the cytosol. Further fractionation experiments identiﬁed the arginine deiminase in a subcellular fraction with a density distinct from both typical lysosomes and hydrogenosomes. It was suggested that this fraction may consist of fragmented cytoplasmic membranes, with which at least part of the arginine deiminase activity could be associated (Yarlett et al. 1994). However, analysis of the T. vaginalis genome sequence revealed the presence of three arginine deiminase genes, all with amino termini strongly resembling known hydrogenosomal target signals (www.tigr.org/tdb/e2k1/tvg/), and subsequent experiments with hemagglutinin-tagged, episomally overexpressed arginine deiminase conﬁrmed the colocalization of the protein in the organelles labeled with the hydrogenosomal marker protein, malic enzyme. The association of arginine deiminase with hydrogenosomes was further supported by mass spectroscopic analysis of puriﬁed hydrogenosomes (our unpublished results). It may be that the protein is predominantly localized in a subpopulation of hydrogenosomes that differ in density from the majority of the organelles. The reason why only one and moreover the ﬁrst enzyme of the arginine dihydrolase pathway is localized in the organelle while the other members of the pathway are cytosolic is unknown. Polyamines are of another signiﬁcance for the hydrogenosomes: inhibition of putrescine-synthesizing enzyme ornithine decarboxylase by 1,4-diamino2-butanone (DAB) resulted in markedly decreased levels of polyamines in T. foetus cells and ultimately led to reduction of hydrogenosome numbers and

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to alterations of hydrogenosomal ultrastructure in growth-inhibited cultures (Reis et al. 1999). Since polyamines have been implicated in protection against oxidative stress (Tadolini 1988) and in membrane stability (Tabor and Tabor 1976), it was suggested that damage to hydrogenosomes in DAB-treated and polyamine-depleted trichomonads could be, at least in part, caused by membrane disorganization and by reactive oxygen species generated within hydrogenosomes under aerobic conditions (Reis et al. 1999).

8 Other Predicted Hydrogenosomal Proteins Availability of the T. vaginalis genome database has the promise to provide us with a complete inventory of the protein composition of the hydrogenosome. Searches for potential hydrogenosomal matrix proteins utilizing the consensus hydrogenosomal targeting motifs (see Dyall and Doleˇzal, this volume; Carlton et al. 2007) identiﬁed 138 candidate genes. BLAST searches showed 67% of these to be similar to known proteins. Among these were most of the already identiﬁed hydrogenosomal proteins (see Dyall and Doleˇzal, this volume; Carlton et al. 2007); however, these searches missed proteins that possess divergent amino-terminal extensions or that do not possess a recognizable targeting motif at all, yet have already been isolated from hydrogenosomes or were identiﬁed by proteomic studies, e.g., the membrane proteins Hmp31 (member of mitochondrial carrier family) (Dyall et al. 2000), the Hmp35 pore-forming membrane protein (Dyall et al. 2003), rubrerythrin, thiol peroxidase, superoxide dismutase (Pütz et al. 2005), a member of inner membrane translocase complex Pam 18 (Doleˇzal et al. 2005), and hydrogenase maturase HydE (Pütz et al. 2006). When variations of the targeting motif were designed, based on the sequences of proteins shown to localize to hydrogenosomes (e.g., Doleˇzal et al. 2005; Pütz et al. 2006), and used to search the database, between approximately 300 and 800 putative hydrogenosomal proteins were identiﬁed in the T. vaginalis genome database (our unpublished data). While the higher number is certainly an overestimate, 200–300 predicted hydrogenosomal proteins is probably a more realistic number. As revealed by BLAST searches, the majority of these putative proteins are hypothetical proteins without known function. The rest includes diverse proteins that are similar to enzymes, chaperones, or electron transport proteins. The list includes fructokinase, xanthine oxidase, D-isomer-speciﬁc 2-hydroxyacid dehydrogenase, organic acid CoA ligase (ADP forming), aminotransferases, protein kinases, numerous ﬂavodoxin fold containing proteins, iron–sulfur ﬂavoproteins, and others. Until the expression status and true cellular localization of these proteins are veriﬁed and until they are biochemically characterized, however, it is futile to speculate on their roles in the hydrogenosome.

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9 Technical Note: Isolation of Hydrogenosomes for Biochemical Experiments The initial studies describing trichomonad hydrogenosomes utilized isopycnic centrifugation to obtain highly puriﬁed organelles (Lindmark and Müller 1973). The hydrogenosome-enriched fraction (also called large granule fraction) used as starting material for further subfractionation was obtained by differential centrifugation of cell homogenate. Separation of hydrogenosomes from contaminating fractions was achieved by isopycnic centrifugation on a sucrose gradient, where hydrogenosomes typically sediment at an equilibrium density of 1.24 (Lindmark and Müller 1973; Müller 1973). While this procedure provided almost pure hydrogenosomes, the organelles were nevertheless affected by the hyperosmotic concentrations of sucrose necessary to achieve the separation. In experiments where hydrogenosomes with intact membranes were needed, only the large granule fraction, osmotically protected with 250 mM sucrose and yielding hydrogenosomes contaminated ˇ erkasovov et al. 1978; Müller and Lindmostly with lysosomes, was used (C mark 1978; Steinbüchel and Müller 1986). The current standard for isolation of pure and mostly intact (as evidenced by the preserved latency of malic enzyme) hydrogenosomes is subfractionation of the large granule fraction in a self-forming gradient of buffered 45% Percoll with 250 mM sucrose using ultracentrifugation in a vertical rotor (Bradley et al. 1997; Sutak et al. 2004).

10 Perspectives While our knowledge of hydrogenosome biochemistry has increased considerably over last three decades, we are far from a complete understanding of the workings of the trichomonad hydrogenosome. The core hydrogenosomal pathway and the role of the organelle in the biosynthesis of iron–sulfur centers are reasonably well deﬁned. The T. vaginalis genome project provided us with a wealth of data that indicate, however, the possible presence of previously unknown functions. Even a number of relatively reliably identiﬁed proteins cannot be accommodated into our current biochemical schemes. Classical biochemistry combined with recombinant molecular methods will be necessary to clarify the roles of numerous hydrogenosomal hydrogenases, electron transport proteins, ﬂavoproteins, membrane proteins, and putative enzymes of carbohydrate metabolism, to name just a few. That there is still a long way to go is evident from the fact that predicted and even wellidentiﬁed hydrogenosomal proteins with unknown functions represent a very high percentage of all proteins present in the organelle. Large-scale proteomic studies will have to complement the genomic data, just to deﬁne as many gen-

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uine hydrogenosomal proteins as possible. Establishing the functions of these proteins will be a major and nontrivial task, especially since we still do not have the methodology to selectively silence the genes of interest. Development of such systems is a challenge for the near future. Acknowledgements We thank Petr Jedelský, M.Sc., for mass spectroscopic determination of hydrogenosomal proteins. The excellent technical assistance of Ms. Míˇsa Marcinˇciková and Dr. Helena Kulíková is gratefully acknowledged. Part of the original research presented in this chapter was supported by the grant 204/06/0944 of the Czech Science Foundation to I.H. and by the Ministry of Education, Youth, and Sports of the Czech Republic MSM0021620858 and LC07032 and the Grant Agency of the Academy of Sciences of the Czech Republic IAA501110631 to J.T.

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_111/Published online: 9 January 2008 © Springer-Verlag Berlin Heidelberg 2008

Hydrogenosomes of Anaerobic Chytrids: An Alternative Way to Adapt to Anaerobic Environments Johannes H. P. Hackstein1 (u) · Scott E. Baker2 · Jaap J. van Hellemond3,4 · Aloysius G. M. Tielens3,4 1 Department

of Evolutionary Microbiology, Faculty of Science, Radboud University Nijmegen, Toernooiveld 1, 6525 ED Nijmegen, The Netherlands [email protected] 2 Chemical & Biological Processes Development Group, Paciﬁc Northwest National Laboratory, Richland, WA USA 3 Department of Biochemistry and Cell Biology, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands 4 Department

of Medical Microbiology and Infectious Diseases, Erasmus MC, University Medical Center Rotterdam, Rotterdam, The Netherlands

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Hydrogenosomal Metabolism of Piromyces and Neocallimastix . . . . . .

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The Role of the Hydrogenosomes in the Energy Metabolism of Piromyces sp. E2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Evidence for a Mitochondrial Origin of Chytrid Hydrogenosomes . . . . .

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Abstract Fungi form a very diverse group of eukaryotes. The majority of investigated fungi contain mitochondria and are capable of oxidative phosphorylation. On the other hand, anaerobically functioning chytridiomycete fungi, found as symbionts in the gastrointestinal tract of many herbivorous mammals, contain hydrogenosomes. These organelles are found in multiple classes of protozoa and catabolize glycolytic end products and produce hydrogen and ATP by substrate-level phosphorylation. However, in contrast to the hydrogenosomes of trichomonads and anaerobic ciliates, the hydrogenosomes of the anaerobic chytrids Neocallimastix and Piromyces lack pyruvate dehydrogenase (PDH) and pyruvate-ferrodoxin oxidoreductase (PFO) and instead contain pyruvate-formate lyase (PFL). The function in carbohydrate metabolism of these hydrogenosomes of anaerobic chytridiomycete fungi and their evolutionary relation to fungal mitochondria is discussed.

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1 Introduction Fungi comprise a monophyletic, but morphologically and biochemically extremely diverse eukaryotic kingdom. The taxon Fungi includes not only some of the best-studied eukaryotic model organisms, i.e. the baker’s yeast Saccharomyces cerevisiae, the ﬁssion yeast Schizosaccharomyces pombe and the moulds Neurospora crassa and Aspergillus nidulans, but also poorly understood, uncultured fungi from soil and aquatic environments, or symbiotic fungi such as, for example mycorrhiza, and elusive anaerobic fungi from the gastro-intestinal tract of herbivorous mammals (Trinci 1994; Schadt et al. 2003; Strack et al. 2003; Anderson and Cairney 2004; Luo et al. 2005). Anaerobic chytridiomycete fungi are important symbionts in the gastrointestinal tract of herbivorous mammals. A ﬂagellated rumen-dwelling organism, Neocallimastix frontalis, was described in 1975 by Colin Orpin of Australia’s CSIRO (Orpin 1975). Two years later, Orpin published a report showing that N. frontalis and two other anaerobes had cell walls that contained chitin, indicating that these rumen dwelling organisms are fungi (Orpin 1977). Over the next three decades, a body of research was generated covering the physiology, ultrastructure and taxonomy, ecology, cell biology, biochemistry and metabolism and genomics of anaerobic chytrids. Monocentric chytrids are deﬁned by a single thallus (zoospore containing structure) and center of growth from which rhizoids, ﬁne anucleate ﬁlaments involved in substrate anchorage and nutrient absorption, radiate. In contrast, polycentric chytrids, have multiple thalli and centers of growth. While not morphologically described, there is evidence that at least one species of anaerobic chytrids can undergo meiosis (Wubah et al. 1991). Zoospores of anaerobic chytrids can have one or multiple ﬂagella depending on the species. The diversity of the anaerobic chytridiomycete fungi is large and they are found in the gastro-intestinal tract of nearly all large herbivores, ranging from ruminants such as cattle, sheep, goat, deer and antelopes to the foregut-fermenting marsupials and camelids on the one hand, and hindgutfermenting species such as horse, elephant, rhinoceros, mara (Patagonian hare) and capybara (“water pig”, the worlds largest rodent) on the other hand. Anaerobic chytridiomycetes can be isolated from rumen ﬂuid or faeces and they are maintained in anaerobic culture, most of them as pure axenic cultures. In the rumen of cattle or sheep, these anaerobic fungi can be as frequent as 7.6 × 108 thallus-forming units; in the faeces there are still 4.2 × 104 units per g dry weight (Trinci et al. 1994). Anaerobic chytrids are not truly host-speciﬁc since it is possible to transfaunate various host animals with isolates from different hosts. On the other hand, the various isolates are not the same, even if collected from the same host species and assigned to the same chytrid species. The patterns of utilization of substrates, and the metabolic properties are different from isolate to isolate (Trinci et al. 1994).

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We now know that the anaerobic chytrids comprise many species that are integral in the rumen ecosystem, and crucial in the digestion of plant material to more metabolizable substrates such as simple sugars. Moreover, they produce hydrogen, needed for the growth of methanogenic bacteria (reviewed in Williams et al. 1994). With regard to degradation of plant material, the rumen chytrids produce a wide variety of glycosyl hydrolases needed for the breakdown of plant biomass (reviewed in Chen et al. 1995). Because of their intimate association with the plant biomass being degraded, chytrid fungi were often discarded with the solid gut material leaving a paucity of culturable material behind, which kept them from being more thoroughly characterized earlier (Bauchop 1979). Later on, they were frequently studied, mainly for biotechnological applications of their cellulolytic enzymes. From both an applied and basic science perspective, anaerobic chytrids are interesting and important organisms. For that reason, the US Department of Energy Joint Genome Institute (JGI) has active genome sequencing projects for the monocentric Piromyces and polycentric Orpinomyces. An early study of the DNA content of a Neocallimastix species found that its genome was extremely A + T rich (up to 82%, Brownlee 1989). Subsequent studies on other anaerobic chytrids produced similar results (Brownlee 1994). These results were later conﬁrmed with a limited gene sequencing study in Orpinomyces (Nicholson et al. 2005). It has been further speculated, that rDNA constitutes up to 25% of the genome (Brownlee 1989, 1994; Nicholson et al. 2005). A number of studies in which genes have been cloned from a variety of anaerobic chytrid species further has shown that the protein coding sequences contain more “normal” levels of A + T (40–60%) (see for example Nicholson et al. 2005). Thus, it is likely that introns and intragenic spaces are responsible for the A + T richness of anaerobic chytrid genomes.

2 Mitochondria Versus Hydrogenosomes The majority of the cultured fungi, which belong to the taxa Ascomycota, Basidiomycota, and Zygomycota contain mitochondria. These mitochondria host a genome of varying size, which characteristically encodes only a handful of proteins (Bullerwell and Lang 2005). This implies that the vast majority of the 700–800 mitochondrial proteins (Sickmann et al. 2003) are nuclear encoded, synthesized in the cytoplasm and imported into the organelles. Interestingly, certain cultivars of mitochondriate species are able to maintain mitochondria in the absence of a mitochondrial genome. Such yeasts are known as “petites”; they are viable but respiration deﬁcient and incapable of growing on non-fermentable substrates (Contamine and Picard 2000). In this respect they are similar to the genome-less chytrid hydrogenosomes to be discussed in detail below.

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On the other hand, two natural isolates of ﬁssion yeasts, Schizosaccharomyces japonicus var. japonicus and S. japonicus var. versatilis lack detectable cytochromes and are respiration deﬁcient, but nevertheless retained fully functional mtDNA (Bullerwell and Lang 2005). These ﬁssion yeasts are considered to be an intermediate evolutionary stage in between respiratorycompetent fungi and those that completely lack mitochondrial DNA. The mitochondria of these yeast species might be similar to the “hydrogenosomes” of the ciliate Nyctotherus ovalis (see Hackstein et al., 2007, in this volume), and therefore, might represent an evolutionary intermediate between the classical mitochondria of most fungi and the “hydrogenosomes” of the chytridiomycete fungi described below. Notably, the members of the phylum chytridiomycota such as, for example, Piromyces (Fig. 1) and Neocallimastix, which possess hydrogenosomes, lack both mitochondria and a mitochondrial genome (van der Giezen et al. 1997). These hydrogenosomes of chytrid fungi are membrane-bounded compartments up to 1 micrometer in size that produce ATP by substrate-level phosphorylation together with hydrogen as an end product allowing growth under anoxic conditions (Marvin-Sikkema et al. 1993, 1994; Hackstein et al. 2001; Voncken et al. 2002b). We will discuss here that these “hydrogenosomes” are specialized mitochondria, just like all other hydrogenosomes, but that these chytrid hydrogenosomes are nevertheless different from those of trichomonads (see Hrdý et al., 2007, this volume)

Fig. 1 Epiﬂuorescence micrograph of Piromyces sp. E2 originally isolated from the faeces of an Indian elephant. Magniﬁcation about ×400. The organism was vitally stained with rhodamine 123. S: young sporangia

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as well as from those of anaerobic ciliates (see Hackstein et al., 2007, in this volume).

3 Anaerobic Chytrids Possess Hydrogenosomes and Perform a (Bacterial-Type) Mixed Acid Fermentation The metabolism of anaerobic chytrids has not been studied in great detail, but it is known that most anaerobic chytrids studied so far produce formate, acetate, succinate, lactate and ethanol besides hydrogen and carbon dioxide when growing on cellulose, glucose or fructose as a carbon source (Julliand et al. 1998). Such a mixed acid fermentation is very similar to bacterial mixed acid fermentations that are, for example, well known for facultative anaerobic enteric bacteria, such as Escherichia coli. Many bacterial mixed-acid fermentations do not produce ethanol from pyruvate via pyruvate decarboxylase and alcohol dehydrogenase as in the alcoholic fermentation of yeast, but by the successive action of pyruvateformate lyase (PFL) and alcohol dehydrogenase E (ADHE). The latter enzyme combines aldehyde dehydrogenase (ALDH) and alcohol dehydrogenase (ADH) activities, and acetyl-CoA is the substrate (Kessler et al. 1991; Arnau et al. 1998; Fontaine et al. 2002). Until now, ADHE has been found exclusively in eubacteria, with the remarkable exception of certain eukaryotic “type I” anaerobes such as Giardia, Spironucleus, Entamoeba and Mastigamoeba, which do not possess mitochondria or hydrogenosomes, but most likely possess mitosomes or other mitochondrial remnants (Bruchhaus and Tannich, 1994; Sánchez 1998; Dan and Wang 2000; Field et al. 2000; Andersson et al. 2003). However, these eukaryotic anaerobes do not perform a bacterialtype mixed acid fermentation, they do not exhibit PFL activity, and, notably, they lack energy-generating organelles such as mitochondria or hydrogenosomes (Müller 1998). Apparently, the presence of hydrogenosomes is essential for the performance of mixed acid fermentations in eukaryotes.

4 Hydrogenosomal Metabolism of Piromyces and Neocallimastix The hydrogenosomal metabolism has been studied in more detail in the chytridiomycetes Piromyces and Neocallimastix. Notably, the hydrogenosomes of these organisms are clearly different from those known of trichomonads and anaerobic ciliates, structurally and metabolically. Most importantly, the hydrogenosomes of Neocallimastix sp. L2 and Piromyces sp. E2 contain PFL as key enzyme, and not PDH (as in N. ovalis) or PFO (as in Trichomonas vaginalis) (Fig. 2).

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Fig. 2 Energy metabolism of Piromyces sp. E2. Shown is a scheme of the metabolic pathways involved in the production of the major end products. The numbers indicate the following enzymes: (1) hexokinase, glucose-6-phosphate isomerase, phosphofructokinase, aldolase and triose phosphate isomerase; (2) glyceraldehyde 3-phosphosphate dehydrogenase; (3) phosphoglycerate kinase, phosphoglycerate mutase and enolase; (4) phosphoenolpyruvate carboxykinase; (5) malate dehydrogenase; (6) fumarase; (7) fumarate reductase; (8) pyruvate kinase; (9) lactate dehydrogenase; (10) cytosolic pyruvate:formate lyase; (11) alcohol dehydrogenase E; (12) pyruvate import into hydrogenosomes; (13) malic enzyme; (14) hydrogenase; (15) hydrogenosomal pyruvate : formate lyase; (16) acetate:succinate CoA-transferase; (17) succinyl-CoA synthethase; (18) ADP/ATP carrier. Abbreviations; AcCoA, acetyl-CoA; EtOH, ethanol; FUM, fumarate; G3P, glyceraldehyde3-phosphate; MAL, malate; OXAC, oxaloacetate; PEP, phosphoenolpyruvate; PYR, pyruvate; SUCC, succinate. (From Boxma et al. 2004)

As discussed above, these chytridiomycetes perform a mixed acid fermentation, where carbohydrate degradation results in the production of formate, acetate, succinate, lactate, ethanol, hydrogen and carbon dioxide (Fig. 2). However, the ratio of these excreted end products is not constant, as it was shown that growth of Piromyces sp. E2 in the presence of increasing concentrations of fructose is accompanied by changes in the fermentation pattern (Boxma et al. 2004). Increasing the fructose concentration from 0.1 to 0.5% resulted in a three-fold increase in degradation of this substrate to end-

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products. It is remarkable that the relative ﬂuxes of fructose degradation through the various pathways were not constant during changing fructose concentrations (Fig. 3). Although the absolute amounts of hydrogen formed in the incubations during growth at these increasing concentrations of fructose remained constant, the relative ﬂux of malate into the hydrogenosomes and hence the relative ﬂux to hydrogen decreased from 47 to 15% (Fig. 3, Boxma et al. 2004). In contrast, the relative ﬂuxes in the formation of the cytosolic end-products lactate, ethanol and succinate increased several-fold. These observations show that increasing amounts of a fermentable carbon source result in an increased metabolism without an increased production of hydrogen and cause a relative shift from hydrogenosomal carbon metabolism to a cytosolic one (Fig. 3). Experiments using [6-14 C]-glucose and [U-14 C]-glucose indicated that an incomplete TCA cycle operates in the reductive mode allowing the formation of succinate from oxaloacetate via a malate intermediate (Fig. 2). Since the formation of signiﬁcant amounts of labelled CO2 could be excluded while formate and acetate plus ethanol were formed in a 1 : 1 ratio, it must be concluded that PFL and not pyruvate : ferredoxin oxidoreductase (PFO) or pyruvate dehydrogenase (PDH) play the central role in the hydrogenosomal metabolism. The activity of the latter enzymes would have generated one molecule of labelled carbon dioxide per molecule of pyruvate degraded. However, less than 1% of the expected amount could be measured (Boxma et al. 2004). Moreover, Piromyces sp. E2 and Neocallimastix sp. L2 exhibit ADHE activity, which is characteristic for bacterial mixed acid fermentations. In E. coli, where PFL is only expressed under anaerobic growth conditions, ADHE can act as a PFL inactivase, thereby protecting PFL against irreversible damage by oxygen (Kessler et al. 1991, 1992; Sawers and Watson 1998). The situation is likely to be comparable in anaerobic chytrids, and, notably, anaerobic chytrids also possess a typical PFL-activating enzyme (Gelius-Dietrich and Henze 2004). There has been a controversy about the presence or activity of pyruvate : ferredoxin oxidoreductase in anaerobic chytrids: in a few cases PFO activity instead of PFL activity has been reported to be responsible for the hydrogenosomal pyruvate degradation. A low enzymatic activity attributed to PFO has been measured in Neocallimastix sp. L2 and N. patriciarum (Yarlett et al. 1986; Marvin-Sikkema et al. 1993, 1994), while in another species, N. frontalis, the absence has been reported of any detectable PFO activity, measured as pyruvate synthase using ferredoxin as the electron acceptor (O’Fallon et al. 1991). Notably, the observation of putative PFO activity in anaerobic chytrids has not been substantiated by characterization of the enzyme nor by identiﬁcation of a PFO gene in these organisms. Furthermore, the observed formate production in all metabolic studies strongly suggests the participation of an active PFL in the metabolism of anaerobic chytrids, as there is no other fermentative pathway known to result in formate produc-

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Fig. 3 Changes in metabolic ﬂuxes in the energy metabolism of Piromyces induced by changes in the fructose concentration in the medium. The thickness of the arrows is proportional to the calculated ﬂuxes in the presence of 0.1% fructose (Panel A) or 0.5% fructose (Panel B). Abbreviations are as in Fig. 3. (Calculated from the data in Boxma et al. 2004)

tion. On the other hand, PFO is not completely unknown in the fungal world, as it has been shown that Saccharomyces and other fungi contain enzymes involved in methionine biosynthesis that are fusion proteins of PFO domains and fragments of redox enzymes (Horner et al. 1999).

5 The Role of the Hydrogenosomes in the Energy Metabolism of Piromyces sp. E2 The observation that the hydrogenosomal PFL and the cytoplasmic ADHE are the key enzymes in the degradation of carbohydrates by anaerobic chytrids reveals that the metabolism of these hydrogenosomes is fundamentally different from the hydrogenosomal metabolism in both Trichomonads and N. ovalis-like ciliates. Obviously, anaerobic chytrids chose their own way to adapt to anaerobic environments by evolutionary tinkering. The metabolic scheme displayed in Fig. 2 shows a generalized metabolism. A quantitative analysis revealed that (i) PFL must be present, and (ii) that under certain conditions, hydrogen formation can become marginal (Fig. 3, Boxma 2004). The evolutionary strategy of chytrids apparently tends to avoid the formation of reduction equivalents by using PFL instead of PFO or PDH, (Akhmanova et al. 1999; Hackstein et al. 1999, 2006; Voncken 2001). At higher fructose concentrations, the major ﬂow through the chytrid hydrogenosome involves pyruvate, which is split by PFL into formate and acetate without the generation of reduction equivalents (Figs. 2, 3). Thus, these reactions do not contribute to hydrogen formation by its [FeFe] hydrogenase (Davidson et al. 2002; Voncken 2001; Voncken et al. 2002b). Hydrogen formation depends on the import of malate into the hydrogenosome, where malate is decarboxylated by malic enzyme, which provides the electrons for the reduction of H+ to hydrogen (Marvin-Sikkema et al. 1994). However, the latter reaction seems to represent only a minor pathway of the anaerobic energy metabolism in this type of hydrogenosome and might indicate a role in controlling or ﬁne tuning of the intra-hydrogenosomal environment. While at a low fructose concentration of 0.1% nearly 50% of the glycolytic ﬂux leads to malate formation with a nearly complete ﬂow of this malate through the hydrogenosome with the concomitant production of hydrogen, less than 30% of the initial hydrogen formation is observed at a fructose concentration of 0.5% (Fig. 3, Boxma et al. 2004).

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The major role of the chytrid hydrogenosomes seems to be the generation of ATP by substrate-level phosphorylation. The presence of PFL in the absence of hydrogenosomal ADHE most probably directs all organellar pyruvate into substrate level ATP formation. A possible presence of ADHE inside the hydrogenosomes would compromise this function of the hydrogenosome as an energy-generating organelle. In the cytoplasm, however, ADHE might allow regulation of PFL activity, thus saving pyruvate (and its metabolites) for anapleurotic pathways. A partial TCA cycle with links to the anapleurotic pathways operates in the cytoplasm (Akhmanova et al. 1998). This hypothesis is supported by the observation that several mitochondrial enzymes, which are involved in anabolic reactions, e.g. malate dehydrogenase, aconitase, isocitrate dehydrogenase and acetohydroacid reductoisomerase, have been retargeted to the cytoplasm in Piromyces sp. E2 (Akhmanova et al. 1998; Hackstein et al. 1999). Consequently, compartmentalization of the energy metabolism seems to enhance the possibilities for regulation of the metabolic pathways of this organism.

6 Evidence for a Mitochondrial Origin of Chytrid Hydrogenosomes Many different data, which will be discussed below, are consistent with a mitochondrial origin of the hydrogenosomes of anaerobic chytrids: their morphology, the mitochondrial-type targeting signals that are used to import proteins, the ADP/ATP carriers and mitochondrial-type chaperones, and all this supplemented by genomic analyses of separate hydrogenosomal proteins. Using 18S rDNA phylogenies, or the phylogenies of mitochondrial genes from aerobic chytrids, a monophyletic origin of all chytrids becomes evident (Bullerwell and Lang 2005). However, as already mentioned, anaerobic chytrids lack mitochondria, and the sistergroup relationships with other fungal groups are less evident: in most cases only moderate support is obtained. Nevertheless, there is no doubt about a fungal origin of the chytrids— regardless as to whether they are thriving in oxic or anoxic environments. The aerobic representatives possess mitochondria: phylogenetic analysis of their nuclear and mitochondrial genomes reinforces their fungal origin (Bowman et al. 1992; cf. Paquin et al. 1995; Paquin and Lang 1996). Also an analysis of biochemical and morphological traits consistently establishes a close relationship between chytrids and other fungi (Ragan and Chapman 1978), and Akhmanova et al. (1998) demonstrated that several enzymes of mitochondrial origin, which lack putative targeting signals, were retargeted to the cytoplasm (in active form) in the hydrogenosome-bearing chytrid Piromyces. Chytrid hydrogenosomes look rather different from the pictures of mitochondria in textbooks (Fig. 4). However, serial sectioning followed by electron microscopical analysis revealed a structure resembling the ultrastructure

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Fig. 4 Serial sectioning of the hydrogenosomes of Neocallimastix sp. L2. A–D Bar = 0.5 micrometer; h, hydrogenosomes; r, ribosome globules (Munn et al. 1988). Asterisk, internal vesicular structures. From Voncken et al. 2002a, modiﬁed

of mitochondria from particular diseased human patients (Frey and Mannella 2000; Hackstein et al. 2001; Voncken et al. 2002a). Also, the relict mitochondrion of Cryptosporidium parvum, looks very similar (Keithly et al. 2005). Apparently, in these cases the inner membrane undergoes a derangement in the mechanism that normally stabilizes the crista junctions (Mannella 2006). Experimental approaches failed to demonstrate the presence of a genome in chytrid hydrogenosomes (van der Giezen et al. 1997). Therefore, a straightforward proof for the mitochondrial ancestry will be difﬁcult. All proteins found in the hydrogenosomes must be encoded by the nucleus, translated in the cytoplasm and transported into the hydrogenosomes. These nuclearencoded “hydrogenosomal” genes underwent changes in codon usage, might have experienced other evolutionary constraints than before or might have been replaced by genes from other cellular or foreign sources (Timmis et al. 2004; Galbadon and Huynen 2004; Gabaldon et al. 2006). All these effects are likely to contribute to bias in a phylogenetic analysis. Consequently, many nuclear encoded hydrogenosomal genes might be less suitable for a phylogenetic analysis than others. Because of their intrinsic function in the organelle, ADP/ATP carriers (AACs) and chaperonins are the best indicators for the phylogenetic analysis of an organelle of putative mitochondrial origin. Phylogenetic analysis of the AACs and chaperonins of anaerobic chytrids revealed unequivocally a fungal mitochondrial ancestry (Voncken 2001, Voncken et al. 2002a; van der Giezen et al. 2002, 2003). Moreover, the spectrum of responses against the various inhibitors is quite speciﬁc and differentiates these AACs clearly from other adenine transporters—regardless as to

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whether these transporters are from mitochondrial or hydrogenosomal origin (Hackstein et al. 2006). While the AACs are eukaryotic “inventions” that allowed the exploitation of the ATP formed inside the organelle after the organelle formation, the chaperonins tend to trace the ancestry of the organelle back to the endosymbiont that gave rise to the mitochondrion. Also, the phylogenetic analysis of HSP 60 and the HSP 70 clearly reveals a clustering with their fungal mitochondrial relatives and not with the alpha-proteobacterial cluster (Hackstein et al. 1999; Voncken et al. 2002a; van der Giezen et al. 2003). Genomic analyses of the hydrogenosomal enzymes succinyl-CoA synthetase (Dacks et al. 2006) and of two hydrogenosomal enzymes involved in arginine biosynthesis (Gelius-Dietrich et al. 2007) further conﬁrm the fungal mitochondrial origin of the Neocallimastix hydrogenosome. Most of the other hydrogenosomal genes have not been identiﬁed so far, but a genome project for Piromyces is in progress and it is likely that these data will provide new insights on the evolutionary history of its hydrogenosomes (http://www.jgi.doe.gov/sequencing/why/CSP2006/piromyces.html). From the metabolic and phylogenetic studies it is clear that the hydrogenosomes of anaerobic chytrids are substantially different from the hydrogenosomes of trichomonads (see Hrdý et al. 2007, in this volume) and also from those of ciliates (see Hackstein et al. 2007, in this volume). From a metabolic point of view the main difference is probably their pyruvate metabolism. Whereas anaerobic chytrids use PFL activity to produce acetyl-CoA, in the hydrogenosomes of trichomonads acetyl-CoA is produced by PFO and in the hydrogenosomes of the anaerobic ciliate N. ovalis this is performed by the pyruvate dehydrogenase complex. In contrast to PFO and PDH, the PFL reaction does not generate reduced equivalents, and therefore, the role of the hydrogenase is in the hydrogenosomes of anaerobic chytrids smaller than in the two other types of hydrogenosomes.

References Akhmanova A, Voncken FGJ, Harhangi H, Hosea KM, Vogels GD, Hackstein0 JHP (1998) Cytosolic enzymes with a mitochondrial ancestry from the anaerobic chytrid Piromyces sp. E2. Mol Microbiol 30:1017–1027 Akhmanova A, Voncken FGJ, Hosea KM, Harhangi H, Keltjens JT, den Camp HJMO, Vogels GD, Hackstein JHP (1999) A hydrogenosome with pyruvate formate-lyase: anaerobic chytrid fungi use an alternative route for pyruvate catabolism. Mol Microbiol 32:1103–1114 Andersson JO, Sjogren AM, Davis LAM, Embley TM, Roger AJ (2003) Phylogenetic analyses of diplomonad genes reveal frequent lateral gene transfers affecting eukaryotes. Curr Biol 13:94–104 Anderson IC, Cairney JWG (2004) Diversity and ecology of soil fungal communities: increased understanding through the application of molecular techniques. Environ Microbiol 6:769–779

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_106/Published online: 1 December 2007 © Springer-Verlag Berlin Heidelberg 2007

The Proteome of T. vaginalis Hydrogenosomes Katrin Henze Institut für Botanik III, HUH Düsseldorf, Universitätsstrasse 1, 40225 Düsseldorf, Germany [email protected] 1

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A Preliminary Status of Trichomonad Hydrogenosomal Proteomes . . . .

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The Origins of Hydrogenosomal Proteins . . . . . . . . . . . . . . . . . . .

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Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract The occurrence of hydrogenosomes in independent eukaryotic lineages implies that they evolved from mitochondrial predecessors which most likely were already highly diverse in functional range and protein content. In contrast to mitochondria, where multiple large-scale genome and proteome analyses provide a constantly increasing coverage of organellar protein content and functions, hydrogenosomes are mostly described by small-scale studies on single proteins or pathways in a very small number of organisms to date. Several independent analyses can be drawn upon to compile a preliminary inventory of the hydrogenosomal proteome in trichomonads: multiple experimental studies on single proteins or pathways, gene products predicted by a computer algorithm to contain an N-terminal targeting peptide for import into hydrogenosomes, and proteins identiﬁed by a proteomic approach based on 2D electrophoresis of puriﬁed hydrogenosomes. On the basis of these observations the proteome of T. vaginalis hydrogenosomes can be tentatively estimated to consist of at least about 200 different proteins, but certainly signiﬁcantly less than the 700 proteins predicted for yeast mitochondria. This estimate puts the trichomonad organelles in the range of the highly reduced mitochondria in parasitic organisms or mitosomes.

1 The Mitochondrial Background Hydrogenosomes are anaerobic, hydrogen-producing relatives of mitochondria with multiple independent origins in the diverse eukaryotic lineages like the parabasalids, the ciliates, or the chytridiomycetes (Embley 2006). Comprehensive proteome comparisons of hydrogenosomes from diverse organisms would help elucidate the multiple pathways from mitochondria to hydrogenosomes, but unlike the mitochondria, where multiple genome comparisons and proteome analyses have been carried out and have uncovered

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an unexpected range of complexity and functional diversity, similar data on hydrogenosomes are scarce. Mitochondria and their relatives, hydrogenosomes and mitosomes, are descendants of an α-proteobacterial endosymbiont. (Esser et al. 2004; Embley 2006). This endosymbiont, the proto-mitochondrion, was estimated to have contained a minimum of 630 different proteins (Gabaldon and Huynen 2003). Modern mitochondria vary tremendously in their protein content. Numbers predicted in silico by the presence of putative transit sequences range between 187 proteins for the highly reduced organelles of the parasite Plasmodium, 1082 for Saccharomyces, 2957 for Arabidopsis, and 4251 for human mitochondria. Only a tiny fraction of these proteins is still encoded on the mitochondrial genomes, e.g. three in P. falciparum, 28 in yeast, 85 in A. thaliana and 13 in humans (Richly et al. 2003). During transformation of the proto-mitochondrion into the many diverse variants of modern mitochondria, extensive protein loss has occurred, either through endosymbiotic gene transfer to the nucleus and relocation of the respective gene products (and functions) to other compartments, or through complete loss of proteins no longer needed at all. In addition, numerous new proteins were recruited from various origins, either through lateral gene transfer from other prokaryotes to the host or as new eukaryotic inventions (Gabaldon and Huynen 2004). Among the latter are major components of the protein import machinery (Dolezal et al. 2006), components of the electron transfer chain complexes (Berry 2003) or the mitochondrial ATP/ADP translocases (Kuan and Saier 1993; Kunji 2004). As a result, less than 20% of mitochondrial proteins can still be traced back to the α-proteobacterial endosymbiont (Gabaldon and Huynen 2004, 2005). Some of the new acquisitions, e.g. ATP/ADP translocators and protein import machineries, are common to all mitochondria and most likely happened before the divergence of the major eukaryotic lineages, but a signiﬁcant portion of mitochondrial proteins, ranging from 69% in the pufferﬁsh Fugu rubripes to 35% in Schizosaccharomyces, is unique to the speciﬁc organisms (Richly et al. 2003), reﬂecting a varying degree of mitochondrial specialization in different organisms, such as the wax ester production in facultatively anaerobic mitochondria of Euglena (Hoffmeister et al. 2005) for example. How do the highly reduced anaerobic relatives of mitochondria, the hydrogenosomes, compare to this variety? The occurrence of hydrogenosomes in independent eukaryotic lineages like parabasalids, ciliates and fungi is indicative of a multiple independent origin of these organelles (Embley 2006), and implies that they evolved from mitochondrial predecessors which most likely were already highly diverse in functional range and protein content. Accordingly, hydrogenosomes should be diverse too, unless their mitochondrial ancestors were all reduced to the same set of proteins and had acquired the same new proteins in adaptation to the anaerobic lifestyles of their host organisms. A scenario that is highly unlikely and has already

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been disproven by the discovery of several signiﬁcant differences between hydrogenosomes in trichomonads, anaerobic chytridiomycetes and anaerobic ciliates, such as the splitting of pyruvate to formate and acetyl-CoA by pyruvate:formate lyase (PFL) (Akhmanova et al. 1999; Gelius-Dietrich and Henze 2004) in chytridiomycete organelles rather than oxidative decarboxylation by pyruvate:ferredoxin oxidoreductase (PFOR) observed in trichomonads (Müller 1993) or the presence of steps of the arginine biosynthesis in the hydrogenosomes of Neocallimastix which are absent from trichomonad organelles (Yarlett et al. 1994; Gelius-Dietrich et al. 2007). Hydrogenosomes in the anaerobic ciliate Nyctotherus differ from those in both trichomonads and chytridiomycetes in that they possess a genome and ribosomal proteins as well as the mitochondrial membrane complexes I and II, traits which make these organelles appear to be a missing link between bona ﬁde mitochondria and hydrogenosomes (Akhmanova et al. 1998; Hackstein et al. 2006).

2 A Preliminary Status of Trichomonad Hydrogenosomal Proteomes How do hydrogenosomes compare to mitochondria in terms of proteome complexity? In contrast to mitochondria, where multiple large-scale genome and proteome analyses provide a constantly increasing coverage of organellar protein content and functions, hydrogenosomes are mostly described by small-scale studies on single proteins or pathways in a very small number of organisms to date. No comprehensive proteome analysis of hydrogenosomes has yet been published, mainly because only two groups of hydrogenosomebearing organisms, trichomonads and chytridiomycetes, can be cultured axenically and grown to cell densities that allow cell fractionation and hydrogenosome puriﬁcation with yields sufﬁcient for proteomic analyses using 2D electrophoresis and mass spectrometry. In addition, the complete genome sequence, an indispensable prerequisite for the identiﬁcation of the proteins in large-scale proteomic approaches, has been determined for only one hydrogenosome-containing organism, T. vaginalis (Carlton et al. 2007). Thus, the hydrogenosomes of T. vaginalis are the ﬁrst, and so far the only ones, for which reasonably comprehensive proteome characterization is feasible. How complex can the proteome of hydrogenosomes in T. vaginalis be expected to be? Trichomonad hydrogenosomes have lost many standard metabolic capacities of mitochondria like—and consequently most of the proteins involved in—the tricarboxylic cycle, membrane-bound electron transport and ATP-production (Müller 1993), or fatty acid synthesis (Beach et al. 1990). Because of the absence of a genome (Clemens and Johnson 2000) the complex machineries of DNA replication and repair, gene transcription and protein synthesis are also absent from these organelles. On the other hand, experimental evidence exists for only a small number of metabolic

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functions, which are hydrogen-forming pyruvate metabolism (Müller 1993), FeS cluster formation (Tachezy et al. 2001; Sutak et al. 2004; Pütz et al. 2006), a few steps of amino acid metabolism (Mukherjee et al. 2006a,b; Pütz et al. 2006), oxygen scavenging (Lindmark and Müller 1974; Pütz et al. 2005), and protein import and processing (Bozner 1997; Bradley et al. 1997; Dolezal et al. 2005). Annotation of the genome sequence did not add any complex new functions to this list (Carlton et al. 2007). Several independent analyses can be drawn upon to compile a preliminary inventory of the hydrogenosomal proteome in trichomonads (Table 1): multiple experimental studies on single proteins or pathways, gene products predicted by a computer algorithm to contain an N-terminal targeting peptide (TP) for import into hydrogenosomes (Carlton et al. 2007), and proteins identiﬁed by a proteomic approach based on 2D electrophoresis of puriﬁed hydrogenosomes from T. vaginalis and Tritrichomonas (Fig. 1) and mass spectrometry (MS) (unpublished data). Computer-based search of the genome sequence for putative N-terminal targeting sequences predicted a hydrogenosomal localization for 138 proteins, these included 86 proteins with functional annotations, most of which were involved in known hydrogenosomal processes like energy metabolism, iron metabolism, or oxygen scavenging, and 53 hypothetical proteins (Carlton et al. 2007). Partial proteome analysis via 2D electrophoresis and mass spectrometry led to the identiﬁcation of 61 proteins, 55 of which had known functions. Thirty-four of the proteins identiﬁed by 2D electrophoresis/MS were not represented in the TP prediction data. Among 24 randomly identiﬁed proteins from the hydrogenosomes of the related Tritrichomonas foetus 21 were also present in the T. vaginalis 2D data and/or in the TP prediction data, while three were not found in either of those two data sets. And ﬁnally, at least 14 proteins are encoded in the genome, some by multiple genes, which are not represented in the TP search- and 2D-data but can be predicted to be hydrogenosomal based on their putative function and/or have been experimentally shown to be hydrogenosomal. Altogether 189 proteins are linked to trichomonad hydrogenosomes by the data summarized in Table 1. Of these 115 are identiﬁed by only one approach and await conﬁrmation by independent experimental evidence, and several will most likely turn out to be false positives like for instance the ribonuclease-related extracellular protein, putative cell wallassociated hydrolase or calcium motive P-type ATPase. The remaining 74 proteins are present in at least two of the independent data sets and/or their hydrogenosomal localization has been proven by experimental evidence. The list of hydrogenosomal proteins in Table 1 is by no means complete, as neither the bioinformatic nor the proteomic approach are capable of covering the complete proteome. TP prediction does not reliably recognize all import signals, as is demonstrated by the failure of the TP prediction algorithm to recognize proven hydrogenosomal presequences like those on HydG and HydE for instance (Table 1, Pütz et al. 2006). Likewise, 2D electrophoretic

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Fig. 1 2D SDS-PAGE analyses of hydrogenosomal proteins from A Trichomonas vaginalis and B Tritrichomonas foetus. 1 mg of protein from puriﬁed organelles was separated by isoelectric focussing on immobilized continuous pH 3–10 gradients in the ﬁrst dimension and 12% SDS-PAGE Gels in the second dimension. Proteins were stained with Coomassie blue R340. PFOR pyruvate : ferredoxin oxidoreductase, MDH decarboxylating malate dehydrogenase, β-SCS β-subunit of succinyl CoA synthase, SOD superoxide dismutase, Rbr rubrerythrin

Pam18 Putative SAM50 Putative TIM23 Putative TIM44 Ferredoxin Succinyl-CoA synthase alpha chain Succinyl-CoA synthase beta chain Malic enzyme subunit A

Co-chaperone GrpE Cpn60 hydrogen. Hsp70 Hsp10 DnaJ homolog Hsp20 Hydrogenosomal processing peptidase Hmp35 Hmp31 (MCF)

Putative function

– – – – + (2) + (3) + (3) + (3)

+ (3)

+ (4)

– + (2)

– –

– – – – + (3) + (3)

+ (2) + (2) + (2) + (2) – + (1) –

+ (1)

+ (1)

– – – – – + (1)

– + (1)

– – –

– + (1) + (2)

Identiﬁed by proteomics in (number of different gene products) T. vaginalis T. foetus

+ (2) + (2) + (3) + (4) + (2) – –

Predicted TP∗ (number of genes predicted)

–

–

+ + + + – –

+ –

– – – – – – +

Gene(s) present, hydrogenosomal localization predicted on function

– (Bozner 1997) (Bozner 1997) – – – (Bradley et al. 1997; Dolezal et al. 2005) (Dyall et al. 2003) (Dyall et al. 2000; Tjaden et al. 2004) (Dolezal et al. 2005) – – – (Suchan et al. 2003) (Lahti et al. 1994; Brugerolle et al. 2000) (Lahti et al. 1992; Brugerolle et al. 2000) (Drmota et al. 1996; Brugerolle et al. 2000)

Experimental evidence for hydrogenosomal localization in trichomonads

Table 1 Proteins linked by genome analysis, proteome analysis or experimental evidence to the hydrogenosomes of Trichmonas vaginalis

168 K. Henze

+ (1) + (1) + (1) + (1) + (1) – –

+ (1)

–

–

+ (1) –

+ (2)

+ (1)

+ (1) + (1)

+ (1)

+ (2)

+ (2)

– –

–

–

–

– –

+ (1)

+ (1) + (1)

+ (2)

+ (2)

+ (5)

64 kDa iron hydrogenase Iron hydrogenase hydA Iron hydrogenase hydB Pyruvate : ferredoxin oxidoreductase A Pyruvate : ferredoxin oxidoreductase B put. Acetate : succinate CoA transferase Adenylate kinase NADH dehydrogenase 24 kDa subunit NADH dehydrogenase 51 kDa subunit Dihydrolipoamide acyltransferase (E2) Aminomethyltransferase (H-protein of the glycine cleavage system) put. Alanine aminotransferase Serine hydroxymethyl transferase

+ (2)

Identiﬁed by proteomics in (number of different gene products) T. vaginalis T. foetus

+ (1) – + (1) + (2)

+ (2)

Predicted TP∗ (number of genes predicted)

Malic enzyme subunit B

Putative function

Table 1 (continued)

– +

–

–

–

– –

–

–

– + – –

–

Gene(s) present, hydrogenosomal localization predicted on function

– (Mukherjee et al. 2006b)

(Mukherjee et al. 2006a)

(Steinbüchel and Müller 1986) (Lange et al. 1994) (Dyall et al. 2004; Hrdy et al. 2004) (Dyall et al. 2004; Hrdy et al. 2004) (Mukherjee et al. 2006a)

(Williams et al. 1987)

(Drmota et al. 1996; Brugerolle et al. 2000) – (Bui and Johnson 1996) (Bui and Johnson 1996) (Williams et al. 1987)

Experimental evidence for hydrogenosomal localization in trichomonads

The Proteome of T. vaginalis Hydrogenosomes 169

+ (1) + (2) + (3) – – – – + (1) – – – –

– – + (2) – – + (1) + (1) – + (1)

Rubrerythrin Thiolperoxidase Thioredoxin Thioredoxin reductase OsmC-like protein hydrogenosomal oxygen reductase Glyoxylate reductase Glyoxalase Alpha-glucosidase II

Predicted TP∗ (number of genes predicted)

Arginine deiminase NifU-like protein HesB-like protein (IscA) Frataxin IscU IscS HydG HydF HydE Mge Jac1 Superoxide dismutase

Putative function

Table 1 (continued)

+ (1) + (1) + (1) – + (1) + (1) + (1) + (1) –

– – + (1) – – – + (1) – – – – + (1) + (2) – – – – + (1) – – –

– – – – – – – – – – – + (1)

Identiﬁed by proteomics in (number of different gene products) T. vaginalis T. foetus

– – – + – – – – –

– – – + + + – – + + + –

Gene(s) present, hydrogenosomal localization predicted on function

– – – – – (Tachezy et al. 2001) (Pütz et al. 2006) (Pütz et al. 2006) (Pütz et al. 2006) – – (Lindmark and Müller 1974) (Pütz et al. 2005) (Pütz et al. 2005) – – – – – – –

Experimental evidence for hydrogenosomal localization in trichomonads

170 K. Henze

Sugar kinase (YjeF like) Fructokinase Phosphofructokinase (bacterial type) Glycerol kinase family protein Flavin–nucleotide binding protein NADH : ﬂavin oxidoreductase FeS Flavoprotein (WrbA) Multimeric ﬂavodoxin domain related MRP protein (NifH superfamily) Nudix family protein Predicted CoA–binding protein Xanthine dehydrogenase Cathepsin L-like cysteine protease 26/29 kDa protease Serine/threonine kinase Casein kinase II subunit alpha Protein phosphatase 2C Protein phosphatase PP2A Similar to Bardet–Biedl

Putative function

Table 1 (continued)

+ (1) + (1) + (4) – – + (3) + (1) – – – – – – – – – – – –

+ (1) + (2)

– – + (5)

+ (2) + (1) + (1) + (1) + (3) + (1) + (1) + (1) + (1) + (1) + (1)

– – – – – – – – – – –

– – –

– –

– + (1) + (1)

Identiﬁed by proteomics in (number of different gene products) T. vaginalis T. foetus

– + (1) –

Predicted TP∗ (number of genes predicted)

– – – – – – – – – – –

– – –

– –

– – –

Gene(s) present, hydrogenosomal localization predicted on function

– – – – – – – – – – –

– – –

– –

– – –

Experimental evidence for hydrogenosomal localization in trichomonads

The Proteome of T. vaginalis Hydrogenosomes 171

syndrome 4 MYB21 Extracellular ribonuclease-related SMC protein Ribophorin I Putative maltose O-acetyltransferase N-acetyltransferase Hybrid cluster protein (HCP) Hydroxylamine reductase Glucosyl ceramidase precursor Putative cell wall-associated hydrolase Calcium motive P-type ATPase Hydroxylacylglutathione hydrolase Hypothetical proteins total

Putative function

Table 1 (continued)

– – – – – – – – – – – – 6 61 (34 not predicted by TP analysis)

– – – + (1) + (1)

+ (1) + (1) 53 138

– – – – –

– – – – –

Gene(s) present, hydrogenosomal localization predicted on function

– – – – – – 24 (3 not in TP and 14 T. vaginalis analyses)

+ (1) + (1) + (1) – –

– – – – –

Identiﬁed by proteomics in (number of different gene products) T. vaginalis T. foetus

+ (1) + (1) + (1) + (1) + (1)

Predicted TP∗ (number of genes predicted)

– – –

– – – – –

– – – – –

Experimental evidence for hydrogenosomal localization in trichomonads

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techniques are usually biased against rare proteins and proteins of very large or very small molecular weights. Intrinsic membrane proteins in particular are under-represented in both data sets, as they do not possess N-terminal targeting sequences identiﬁable by TP prediction programmes, and they are usually highly hydrophobic and thus difﬁcult to solubilize for isoelectric focussing during 2D electrophoresis. Accordingly, with the exception of the putatively pore-forming but functionally uncharacterized Hmp35 (Dyall et al. 2003) no membrane-bound components attributable to the protein import machinery were detected by either TP prediction or 2D-electrophoresis, although Hidden Markov-model pattern searching of the T. vaginalis genome identiﬁed putative homologs of the Tim23/Pam complex and the outer membrane sorting and assembly (SAM) complex subunit Sam50 (Dolezal et al. 2006). These observations together with the previously identiﬁed soluble components of protein import like Hsp70, Pam18 and hydrogenosomal processing peptidase suggest that a core import machinery similar to that proposed for the proto-mitochondrion is present in the hydrogenosomal envelope (Dolezal et al. 2005, 2006). The most elusive group of hydrogenosomal membrane proteins are the transporters. Although several metabolite- and substrate-carrying transporters must undoubtedly be present in the organellar membranes, only Hmp31, a member of the mitochondrial carrier family that is represented by ﬁve genes in the genome, has so far unequivocally been assigned to the hydrogenosomal membrane (Dyall et al. 2000). On the basis of these observations and the data summarized in Table 1— and corroborated by the relatively low complexity of trichomonad hydrogenosomal proteomes observed in 2D analyses (Fig. 1)—the proteome of T. vaginalis hydrogenosomes can be tentatively estimated to consist of at least about 200 different proteins, but certainly signiﬁcantly less than the 700 proteins predicted for yeast mitochondria (Prokisch et al. 2004). This estimate puts the trichomonad organelles in the range of the highly reduced mitochondria in parasitic organisms like Plasmodium (187) or mitosomes of Encephalitozoon (156) (Richly et al. 2003).

3 The Origins of Hydrogenosomal Proteins The proteins of trichomonad hydrogenosomes can be largely split into those which are present in mitochondria and those which are not. The ﬁrst group can safely be assumed to have been contributed to the hydrogenosomal proteome by the α-proteobacterial endosymbiont/mitochondrial predecessor the hydrogenosome evolved from. Examples are Hsp70 and Cpn60 (Bui et al. 1996), NADH dehydrogenase (24 kDa and 51 kDa subunits) (Hrdy et al. 2004), the FeS cluster synthesis component IscS (Tachezy et al. 2001; Richards and van der Giezen 2006), or succinyl coA synthase as the only remaining enzyme

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of the citric acid cycle in the hydrogenosomes (Schnarrenberger and Martin 2002), which are unequivocally of alpha-proteobacterial/protomitochondrial origin. Other likely members of the “mitochondrial origin” group of hydrogenosomal proteins are putatively eukaryotic additions to the organellar proteomes like the protein translocase components Tim23 and Pam18 (Dolezal et al. 2006). The second group of hydrogenosomal proteins are those with functional annotations which are not known from any mitochondria. Their origins are much more difﬁcult to pinpoint: They could either be contributions of the α-proteobacterial endosymbiont which were speciﬁcally retained by hydrogenosomes and lost from mitochondria, or they may have been acquired in the course of hydrogenosomal evolution through lateral gene transfer. Interestingly, this group contains not only the proteins involved in the deﬁning anaerobic pyruvate metabolism and hydrogen production of hydrogenosomes, but also proteins putatively responsible for the removal of reactive oxygen species and peroxides. PFOR and [Fe]-hydrogenase are the key enzymes of anaerobic hydrogenosomal energy metabolism and typical for anaerobic bacteria, in their bacterial/hydrogenosomal forms these enzymes are unknown from extant mitochondria. Accordingly, their origin could also be the key to understanding the evolution of hydrogenosomes from mitochondrial ancestors, but it has turned out that neither is truly unique to hydrogenosomes in eukaryotes. PFOR is also present in the cytosol of Entamoeba and the diplomonads (Rodriguez et al. 1996; Brown et al. 1998). A fusion protein of PFOR and NADPH-cytochrome P450 reductase were detected in the facultatively anaerobic mitochondria of Euglena as well as Cryptosporidium cells (Rotte et al. 2001). [Fe]-hydrogenases are also present in Entamoeba and the diplomonads (Nixon et al. 2003), as well as the chloroplasts of green algae (Happe et al. 1994; Florin et al. 2001). In addition, the closely related protein Narf, which is reportedly involved in various processes like prelamin binding in the nucleus (Barton and Worman 1999) and FeS cluster synthesis in the cytosol (Balk et al. 2005), but does not have hydrogenase activity, is widely distributed among eukaryotes. Phylogenetic analyses of both PFOR and [Fe]hydrogenase, and their widespread distribution, suggested their presence in early eukaryotes but did not allow them to be unequivocally linked to the mitochondrial ancestor (Embley 2006). Similarly inconclusive are the origins of [Fe]-hydrogenase maturases HydG, E and F, which are essential for the synthesis/insertion of the speciﬁc FeS cluster (H-cluster) of [Fe]-hydrogenase (Posewitz et al. 2004) and thus are likely to have been acquired by eukaryotes together with the [Fe]-hydrogenase. These three proteins have recently been detected in the chloroplasts of green algae and the hydrogenosomes of T. vaginalis (Posewitz et al. 2004; Pütz et al. 2006), but are suspiciously absent from E. histolytica and G. intestinalis genomes. Algal and trichomonad HydG and E cluster together and are clearly of eubacterial origin, but again there is

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no evidence for an α-proteobacterial/mitochondrial origin (Pütz et al. 2006). Thus, the phylogenetic origin of the deﬁning enzymes of hydrogenosomal metabolism remains unresolved. A part of hydrogenosomal metabolism that is strongly characterized by enzymes not present in modern mitochondria is the decomposition of peroxides. Hydrogenperoxide is probably reduced by the FeS-protein rubrerythrin (Pütz et al. 2005) and a thioredoxin-linked peroxiredoxin, a homolog of bacterial periplasmatic Tpx, that is only distantly related to the cytosolic thiolperoxidase of T. vaginalis (Coombs et al. 2004; Pütz et al. 2005). A third protein, OsmC (Table 1), has alkylperoxidase activity in bacteria (Lesniak et al. 2003) and may have the same function in T. vaginalis hydrogenosomes. These three proteins are wide-spread among anaerobic and facultatively anaerobic eubacteria but, with the exception of a ruberythrin in Entamoeba, have no known homologs in any other eukaryotes. Phylogenetic analyses of rubrerythrin and Tpx sequences unequivocally showed that these proteins have a eubacterial origin (Pütz et al. 2005), thus, rubrerythrin, Tpx and OsmC may represent lateral acquisitions from bacterial sources during the evolution of the trichomonad hydrogenosomes. A comprehensive study of the phylogenetic history of the hydrogenosomal proteome is not yet available, but the analyses of single proteins described above suggest a mixed origin of hydrogenosomal proteins from mitochondria, that is proteins contributed by the α-proteobacterial endosymbiont or proteins acquired by the protomitochondrion before evolution of the hydrogenosomes, and proteins that may have been acquired from bacterial sources by lateral gene transfer.

4 Outlook The era of proteomics is only just beginning for hydrogenosomes, but it could contribute signiﬁcantly to our understanding of hydrogenosome function and evolution. An ongoing genome project on Piromyces sp. will soon facilitate the proteome analysis of the divergent hydrogenosomes from chytridiomycetes. Comparison with the trichomonad hydrogenosomal proteome will allow a ﬁrst assessment of the variability and complexity of hydrogenosomes in comparison with mitochondria and provide valuable insight into the way hydrogenosomes evolve from mitochondrial ancestors. In this context, characterization of the proteomes of the “intermediate” organelles of Nycthotherus and the hydrogenosomes from free-living organisms like the heterolobosean Psalteriomonas or anaerobic ciliates would be highly desirable, and would make worthwhile the considerable effort of setting up genome sequencing projects for these organisms and the development of techniques to make these organisms and their organelles accessible for proteomics.

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Dyall SD et al. (2004) Non-mitochondrial complex I proteins in a hydrogenosomal oxidoreductase complex. Nature 431:1103–1107 Embley TM (2006) Multiple secondary origins of the anaerobic lifestyle in eukaryotes. Philos Trans R Soc Lond B Biol Sci 361:1055–1067 Esser C et al. (2004) A genome phylogeny for mitochondria among alpha-proteobacteria and a predominantly eubacterial ancestry of yeast nuclear genes. Mol Biol Evol 21:1643–1660 Florin L, Tsokoglou A, Happe T (2001) A novel type of iron hydrogenase in the green alga Scenedesmus obliquus is linked to the photosynthetic electron transport chain. J Biol Chem 276:6125–6132 Gabaldon T, Huynen MA (2003) Reconstruction of the proto-mitochondrial metabolism. Science 301:609 Gabaldon T, Huynen MA (2004) Shaping the mitochondrial proteome. Biochim Biophys Acta 1659:212–220 Gabaldon T, Huynen MA (2005) Lineage-speciﬁc gene loss following mitochondrial endosymbiosis and its potential for function prediction in eukaryotes. Bioinformatics 21(2):ii144–ii150 Gelius-Dietrich G, Henze K (2004) Pyruvate formate lyase (PFL) and PFL activating enzyme in the chytrid fungus Neocallimastix frontalis: a free-radical enzyme system conserved across divergent eukaryotic lineages. J Eukaryot Microbiol 51:456–463 Gelius-Dietrich G, Braak MT, Henze K (2007) Mitochondrial Steps of Arginine Biosynthesis are Conserved in the Hydrogenosomes of the Chytridiomycete Neocallimastix frontalis. J Eukaryot Microbiol 54:42–44 Hackstein JH, Tjaden J, Huynen M (2006) Mitochondria, hydrogenosomes and mitosomes: products of evolutionary tinkering. Curr Genet 50:225–245 Happe T, Mosler B, Naber D (1994) Induction, localization and metal content of hydrogenase in the green alga Chlamydomonas reinhardtii. Eur J Biochem 222:769–774 Hoffmeister M, Piotrowski M, Nowitzki U, Martin W (2005) Mitochondrial trans-2-enoylCoA reductase of wax ester fermentation from Euglena gracilis deﬁnes a new family of enzymes involved in lipid synthesis. J Biol Chem 280:4329–4338 Hrdy I et al. (2004) Trichomonas hydrogenosomes contain the NADH dehydrogenase module of mitochondrial complex I. Nature 432:618–622 Kuan J, Saier MHJ (1993) The mitochondrial carrier family of transport proteins: structural, functional, and evolutionary relationships. Crit Rev Biochem Mol Biol 28:209–233 Kunji ERS (2004) The role and structure of mitochondrial carriers. FEBS Lett 564:239–244 Lahti CJ, Bradley PJ, Johnson PJ (1994) Molecular characterization of the alpha-subunit of Trichomonas vaginalis hydrogenosomal succinyl CoA synthetase. Mol Biochem Parasitol 66:309–318 Lahti CJ, d’Oliveira CE, Johnson PJ (1992) Beta-succinyl-coenzyme A synthetase from Trichomonas vaginalis is a soluble hydrogenosomal protein with an amino-terminal sequence that resembles mitochondrial presequences. J Bacteriol 174:6822–6830 Lange S, Rozario C, Müller M (1994) Primary structure of the hydrogenosomal adenylate kinase of Trichomonas vaginalis and its phylogenetic relationships. Mol Biochem Parasitol 66:297–308 Lesniak J, Barton WA, Nikolov DB (2003) Structural and functional features of the Escherichia coli hydroperoxide resistance protein OsmC. Protein Sci 12:2838–2843 Lindmark DG, Müller M (1974) Superoxide dismutase in the anaerobic ﬂagellates Tritrichomonas foetus and Monocercomonas sp. J Biol Chem 249:4634–4637 Mukherjee M, Brown MT, McArthur AG, Johnson PJ (2006a) Proteins of the glycine decarboxylase complex in the hydrogenosome of Trichomonas vaginalis. Eukaryot Cell 5:2062–2071

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Mukherjee M, Sievers SA, Brown MT, Johnson PJ (2006b) Identiﬁcation and biochemical characterization of serine hydroxymethyl transferase in the hydrogenosome of Trichomonas vaginalis. Eukaryot Cell 5:2072–2078 Müller M (1993) The hydrogenosome. J Gen Microbiol 139:2879–2889 Nixon JE et al. (2003) Iron-dependent hydrogenases of Entamoeba histolytica and Giardia lamblia: activity of the recombinant entamoebic enzyme and evidence for lateral gene transfer. Biol Bull 204:1–9 Posewitz MC, King PW, Smolinski SL, Zhang L, Seibert M, Ghirardi ML (2004) Discovery of two novel radical S-adenosylmethionine proteins required for the assembly of an active [Fe]-hydrogenase. J Biol Chem 279:25711–25720 Prokisch H et al. (2004) Integrative analysis of the mitochondrial proteome in yeast. PLoS Biol 2:e160 Pütz S, Dolezal P, Gelius-Dietrich G, Bohacova L, Tachezy J, Henze K (2006) [Fe]-hydrogenase maturases in the hydrogenosomes of Trichomonas vaginalis. Euk Cell 5:579–586 Pütz S, Gelius-Dietrich G, Piotrowski M, Henze K (2005) Rubrerythrin and peroxiredoxin: two novel putative peroxidases in the hydrogenosomes of the microaerophilic protozoon Trichomonas vaginalis. Mol Biochem Parasitol 142:212–223 Richards TA, van der Giezen M (2006) Evolution of the Isd11-IscS complex reveals a single alpha-proteobacterial endosymbiosis for all eukaryotes. Mol Biol Evol 23:1341–1344 Richly E, Chinnery PF, Leister D (2003) Evolutionary diversiﬁcation of mitochondrial proteomes: implications for human disease. Trends Genet 19:356–362 Rodriguez MA, Baez-Camargo M, Delgadillo DM, Orozco E (1996) Cloning and expression of an Entamoeba histolytica NAPD+(–)dependent alcohol dehydrogenase gene. Biochim Biophys Acta 1306:23–26 Rotte C, Stejskal F, Zhu G, Keithly JS, Martin W (2001) Pyruvate: NADP+ oxidoreductase from the mitochondrion of Euglena gracilis and from the apicomplexan Cryptosporidium parvum: a biochemical relic linking pyruvate metabolism in mitochondriate and amitochondriate protists. Mol Biol Evol 18:710–720 Schnarrenberger C, Martin W (2002) Evolution of the enzymes of the citric acid cycle and the glyoxylate cycle of higher plants. A case study of endosymbiotic gene transfer. Eur J Biochem 269:868–883 Steinbüchel A, Müller M (1986) Anaerobic pyruvate metabolism of Tritrichomonas foetus and Trichomonas vaginalis hydrogenosomes. Mol Biochem Parasitol 20:57–65 Suchan P et al. (2003) Incorporation of iron into Tritrichomonas foetus cell compartments reveals ferredoxin as a major iron-binding protein in hydrogenosomes. Microbiology 149:1911–1921 Sutak R et al. (2004) Mitochondrial-type assembly of FeS centers in the hydrogenosomes of the amitochondriate eukaryote Trichomonas vaginalis. Proc Natl Acad Sci USA 101:10368–10373 Tachezy J, Sanchez LB, Müller M (2001) Mitochondrial type iron-sulfur cluster assembly in the amitochondriate eukaryotes Trichomonas vaginalis and Giardia intestinalis, as indicated by the phylogeny of IscS. Mol Biol Evol 18:1919–1928 Tjaden J, Haferkamp I, Boxma B, Tielens AG, Huynen M, Hackstein JH (2004) A divergent ADP/ATP carrier in the hydrogenosomes of Trichomonas gallinae argues for an independent origin of these organelles. Mol Microbiol 51:1439–1446 Williams K, Lowe PN, Leadlay PF (1987) Puriﬁcation and characterization of pyruvate: ferredoxin oxidoreductase from the anaerobic protozoon Trichomonas vaginalis. Biochem J 246:529–536 Yarlett N, Lindmark DG, Goldberg B, Moharrami MA, Bacchi CJ (1994) Subcellular localization of the enzymes of the arginine dihydrolase pathway in Trichomonas vaginalis and Tritrichomonas foetus. J Eukaryot Microbiol 41:554–559

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_112/Published online: 8 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Hydrogenosome: The Site of 5-Nitroimidazole Activation and Resistance Jaroslav Kulda (u) · Ivan Hrdý Department of Parasitology, Faculty of Science, Charles University in Prague, Viniˇcná 7, 128 44 Prague 2, Czech Republic [email protected] 1

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Activation of Metronidazole in Hydrogenosomes . . . . . . . . . . . . . . Classical Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alternative Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Metronidazole Resistance . . . . . . . . . . . . . . . . . . . . Aerobic Type of Metronidazole Resistance . . . . . . . . . . Anaerobic Type of Metronidazole Resistance . . . . . . . . . Development of Anaerobic Resistance . . . . . . . . . . . . . Changes in Metabolism and Protein Expression . . . . . . . Compensation of Hydrogenosomal Insufﬁciency . . . . . . . Hydrogenosomes of Metronidazole-Resistant Trichomonads Postgenomic Challenge . . . . . . . . . . . . . . . . . . . . . Multiple Genes . . . . . . . . . . . . . . . . . . . . . . . . . . Other Potential Mechanisms of Metronidazole Resistance . .

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Abstract Derivatives of 5-nitroimidazole, such as metronidazole or tinidazole, are the drugs of choice for treatment of sexually transmitted infections of humans caused by the parasitic protist Trichomonas vaginalis. These drugs with selective activity against anaerobic and microaerophilic microorganisms enter the trichomonad cell and accumulate in hydrogenosomes, where their antimicrobial properties are activated. In this chapter we discuss metabolic pathways of hydrogenosomes involved in metronidazole activation. We also summarize present knowledge on the development and biochemical mechanisms of metronidazole resistance in T. vaginalis and the related cattle parasite Tritrichomonas foetus. Implications of data from the T. vaginalis genome project suggesting the presence of novel mechanisms of drug resistance are also considered.

1 Introduction Derivatives of 5-nitroimidazole are potent drugs that are active against anaerobic and microaerophilic microbial pathogens of both eukaryotic and

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Fig. 1 Metronidazole, 1-(2-hydroxyethyl)-2-methyl-5-nitroimidazole, and side chains in positions 1 and 2 of related compounds tinidazole and ornidazole. Metronidazole and tinidazole are approved by the FDA for treatment of trichomoniasis, and ornidazole is used for this purpose outside the USA. Both tinidazole and ornidazole have been used with variable success in attempts to cure metronidazole-resistant infections

prokaryotic origin. Metronidazole, the ﬁrst compound of this group introduced into clinical use in 1959, still remains the most widely used 5-nitroimidazole (Fig. 1). Related compounds (tinidazole, ornidazole, nimorazole, secnidazole, etc.), although differing in side-chain structure and some pharmacokinetic properties, all possess a common mechanism of action. Their antimicrobial properties are activated by metabolic reduction within susceptible target cells, resulting in the formation of cytotoxic nitro anion radicals. Since clinically used 5-nitroimidazoles possess low redox potential (e.g., –486 mV for metronidazole), they act selectively against organisms equipped with low redox potential systems of electron generation and transport that are able to mediate one-electron step transfer to the 5-nitro group of the drug (Edwards 1993a). In trichomonads, activation of 5-nitroimidazoles takes place in hydrogenosomes with the involvement of a key enzyme of the hydrogenosomal pyruvate catabolism, pyruvate : ferredoxin oxidoreductase (PFOR) and [2Fe-2S] ferredoxin, the PFOR-linked electron carrier. Under physiological conditions the hydrogenosomal ferredoxin transports electrons to protons in a reaction catalyzed by the [FeFe] hydrogenase that produces molecular hydrogen (see Hrdý et al., this volume, for an overview of basic data on the metabolism of hydrogenosomes).

2 Activation of Metronidazole in Hydrogenosomes The metabolism of metronidazole has been studied in both species of pathogenic trichomonads that are responsible for sexually transmitted urogenital diseases in humans (Trichomonas vaginalis) and cattle (Tritrichomonas foetus). Metronidazole enters the cell as an inactive prodrug by

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simple diffusion (Müller and Lindmark 1976; Müller and Gorrell 1983) and the same mode of entry into the hydrogenosome is assumed. Two pathways of the drug activation have been identiﬁed in trichomonads. 2.1 Classical Pathway In this pathway, which is present in both T. vaginalis and T. foetus, electrons for the drug reduction are generated by PFOR during the course of oxidative decarboxylation of pyruvate and transferred to ferredoxin. Metronidazole acts within the hydrogenosome as a high afﬁnity electron acceptor that effectively competes with protons for electrons. Consequently, the production of hydrogen is suppressed (Lloyd and Kristensen 1985) and ferredoxin-mediated electron transport is directed to the drug (Fig. 2). In agreement with the general concept of activation of 5-nitroimidazoles (Edwards 1993a), reduction of the prodrug nitro group proceeds in hydrogenosomes as a one-electron step transfer resulting in the release of reactive intermediates. Generation of metronidazole anion radicals in intact cells, cell homogenates, or hydrogenosomal fractions of both T. foetus and T. vaginalis exposed to the drug has been repeatedly demonstrated by EPR spectroscopy (Moreno et al.

Fig. 2 Scheme of metronidazole activation in the hydrogenosome: the pyruvate pathway. In the presence of metronidazole, electrons generated by pyruvate : ferredoxin oxidoreductase (PFOR) are transferred by ferredoxin (Fd) to the prodrug form of metronidazole (MTZox ) and not to their natural acceptor hydrogenase (HY). The cytotoxic nitro free radicals R-NO2 – are generated as intermediate products of the metronidazole reduction (MTZred )

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Fig. 3 Signal of metronidazole anion radicals generated by hydrogenosomes of T. vaginalis in the presence of 45 mM metronidazole, pyruvate, CoA, and ferredoxin (activity of PFOR). Recorded by EPR spectroscopy at 25 ◦ C with 20 mW of microwave power, a frequency of 9.64 GHz, and a modulation amplitude of 0.19 mT. The hyperﬁne coupling constants were N aNO2 = 1.565 mT, H a4 = 0.542 mT, and H aCH3 = 0.229 mT

1983, 1984; Chapman et al. 1985; Yarlett et al. 1987; Rasoloson et al. 2002) (Fig. 3). Involvement of reductive activation in the antitrichomonad action of metronidazole was further conﬁrmed by studies on laboratory-developed drug-resistant strains that lack the electron generating enzymes and are thus incapable of activating the drug (see below). 2.2 Alternative Pathway An alternative hydrogenosomal pathway of metronidazole activation, which is independent of PFOR activity, was discovered in T. vaginalis (Hrdý et al. 2005). In this pathway the electrons for the drug reduction are generated by the oxidative decarboxylation of malate catalyzed by the NAD-dependent malic enzyme (malate dehydrogenase (decarboxylating)). The NADH produced by this reaction is reoxidized by an enzyme with NADH : ferredoxin oxidoreductase activity that has been recently identiﬁed as a homologue of the NADH dehydrogenase (NDH) module of the mitochondrial respiratory complex I (Hrdý et al. 2004; and see Hrdý et al., this volume). The T. vaginalis NDH transfers electrons to ferredoxin, which serves as a terminal electron carrier donating the electrons to the drug (Fig. 4). Attention to this pathway was drawn by earlier metabolic studies on the biochemical background of drug resistance development in T. vaginalis (Rasoloson et al. 2002), in particular by a need to explain the partial susceptibility of PFOR-deﬁcient T. vaginalis lines to metronidazole (see below). Hrdý et al. (2005) provided direct evidence that the pathway is functional in both wild-

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Fig. 4 Scheme of alternative pathway of metronidazole activation in hydrogenosomes of T. vaginalis: the malate pathway. Electrons generated by hydrogenosomal malic enzyme (ME) reduce NAD+ to NADH, NADH dehydrogenase (NDH) recycles NADH and transfers electrons to ferredoxin (Fd) which, in a ﬁnal step, donates electrons for metronidazole reduction. MTZ: metronidazole, R-NO2 – : metronidazole anion radicals. Experimental conditions as in Fig. 3

Fig. 5 EPR spectra demonstrating generation of metronidazole anion radicals in T. vaginalis hydrogenosomes by the alternative pathway. Trichomonads were exposed to 43 mM metronidazole. a Malic enzyme reaction. Signal generated in the presence of malate, ferredoxin, and NAD+ . b NADH dehydrogenase activity. Signal generated in the presence of NADH and ferredoxin. c Absence of the pyruvate pathway in PFOR-deﬁcient strain TV1002-MR5 with functional alternative pathway of metronidazole activation. No signal generated in the presence of pyruvate, ferredoxin, and CoA. Experimental conditions as in Fig. 3

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type and PFOR-deﬁcient trichomonads by monitoring metronidazole anion radical release with the aid of EPR spectroscopy. They demonstrated the ability of malic enzyme to generate electrons required for drug reduction in isolated T. vaginalis hydrogenosomes supplied with NAD+ , malate, ferredoxin, and metronidazole (Fig. 5a), and proved NDH involvement in the electron transfer by using NADH as a single electron donor in the same assay system (Fig. 5b). To prove that the enzyme involved in the alternative pathway of metronidazole reduction is indeed the T. vaginalis NDH, they tested radical production in an assay mixture containing puriﬁed T. vaginalis enzyme, recombinant T. vaginalis ferredoxin, NADH, and metronidazole. Radical formation was detected and found to be dependent on the presence of ferredoxin in the reaction mixture. These results show that NDH is involved in electron transfer to metronidazole but cannot donate electrons directly to the drug. Thus, ferredoxin acts as an essential electron donor in both pyruvate- and malate-dependent pathways of metronidazole activation.

3 Metronidazole Resistance It has been generally accepted that trichomonads can develop two distinct types of metronidazole resistance called “aerobic” and “anaerobic” according to the conditions at which the resistance is manifested. Both types of resistance have been demonstrated in T. foetus as well as T. vaginalis. Aerobic resistance in T. vaginalis typically occurs in clinical isolates from treatmentrefractory patients and apparently results from impaired oxygen scavenging and subsequent interference of intracellular oxygen with the reductive activation of metronidazole. It should be noted that the natural habitat of T. vaginalis is not anaerobic, as the parasites are exposed to ﬂuctuating oxygen levels at the vaginal surface in the range of 15–56 µM (Wagner and Levin 1978). The PFOR pathway responsible for activation of the drug remains functional in the strains with aerobic resistance; accordingly, they are susceptible to metronidazole under anaerobic conditions. Anaerobic resistance, as deﬁned in laboratory-developed strains, results from the gradual loss of PFOR and other hydrogenosomal proteins leading to elimination of the drug-activating pathway. Both types of resistance have been repeatedly reviewed (Edwards 1993b; Kulda 1999; Upcroft and Upcroft 2001; Dunne et al. 2003; Cudmore et al. 2004). Surprisingly, monitoring of changes in metabolism and protein expression that accompany in vitro development of metronidazole resistance in T. vaginalis (Rasoloson et al. 2002) revealed that the two types of resistance belong to a common multistep process, with aerobic resistance occurring at its earliest stage.

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3.1 Aerobic Type of Metronidazole Resistance The phenomenon of aerobic resistance was ﬁrst recognized in a T. foetus strain obtained by subcurative treatment of experimentally infected mice (Meigassner et al. 1978). It was later conﬁrmed in a clinical isolate of T. vaginalis from treatment-refractory patients (Meingassner and Thurner 1979). The aerobic type of resistance in T. vaginalis was also induced in vitro (Tachezy et al. 1993; Rasoloson et al. 2002) by cultivation of a drug-susceptible strain with low concentrations of metronidazole (1–3 µg/ml) for a relatively short period of time (∼50 daily transfers in drug-containing medium). With the introduction of improved assay systems that allow access of suitable oxygen concentrations to the parasites, resistant T. vaginalis isolates related to treatment failures in patients have been reported worldwide (see Müller et al. 1988; Sobell et al. 1999; Meri et al. 2000; Dunne et al. 2003, and other references in these papers). Most resistant clinical isolates showed values of minimal lethal concentration (MLC) within the range 25 to >400 µg/ml metronidazole in assays based on the Meingassner protocol (see Müller et al. 1988) conducted under aerobic conditions. An MLC of 100 µg/ml (588 µM) was reported most frequently. The clinical resistance of this type is incomplete and in most (but not all) cases can be overcome by elevated doses of the drug (Lossick 1986). There is general agreement that aerobic resistance is caused by a defect in oxygen scavenging that permits more effective futile cycling between the prodrug form of metronidazole and reactive products of its reduction (Fig. 6). Lloyd and Pedersen (1985) examined the generation of metronidazole anion radicals using suspensions of the metronidazole-resistant clinical isolate and drug-susceptible strain exposed to various partial pressures of oxygen. Using EPR spectroscopy they demonstrated a signiﬁcantly faster decrease in intensity and an eventual disappearance of the radical signal in the resistant strain in response to increasing O2 level. Accordingly, the MLC levels for aerobically resistant trichomonads increased from 3 to 400 µg/ml (7.6 µM to 2.35 mM) metronidazole with increasing oxygen content in the assay system (1 to 20% O2 ), while the effect on the drug-susceptible strain was negligible (MLC 3 to 6 µg/ml) (Tachezy et al. 1993). To determine whether the increased quenching of the metronidazole radical in strains displaying the aerobic type of resistance was indeed due to increased intracellular concentrations of oxygen, Yarlett et al. (1986a) examined oxygen afﬁnities of two susceptible and three resistant strains using mass spectrometric methods. Respiratory rates were measured in intact organisms and subcellular fractions. The KM values for O2 in the resistant strains were six- to tenfold higher than those in susceptible strains, and the lowered oxygen afﬁnity was associated with the hydrogenosomal fraction. It has been repeatedly conﬁrmed that the changes of cytosolic oxidoreductase activities are not involved in aerobic resistance (Müller and Gorrell 1983; Rasoloson et al. 2001), but the hydrogenosomal terminal oxi-

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Fig. 6 Scheme of futile cycling of metronidazole in the presence of oxygen. Nitro anion radicals are converted back to the inactive reoxidized form. By-products of this conversion are superoxide radicals

dase assumed to be responsible for this phenomenon has, thus far, not been identiﬁed. Although isolated hydrogenosomes consume oxygen with high activity ˇ erkasov et al. 1978), this respiration could potentially be at (Müller 1973; C least partially ascribed to autooxidation of FeS clusters, namely in PFOR and ferredoxin (Docampo 1978). Nevertheless, it is likely that organelles harboring oxygen-sensitive enzymes, such as PFOR and hydrogenase (Lindmark and Müller 1973; Lloyd and Kristensen 1985), possess oxygen-scavenging enzymes. One candidate for such an oxygen reductase could be a member of the ﬂavodiiron protein superfamily. These proteins are commonly found in anaerobic prokaryotes and are believed to function as oxygen and/or nitric oxide reductases (Saraiva et al. 2004). The ﬂavodiiron protein homologue has been identiﬁed in T. vaginalis hydrogenosomes (our unpublished data), and subsequently a corresponding single gene with a predicted hydrogenosomal targeting sequence has been found in the T. vaginalis genome (Carlton 2007). The trichomonad enzyme, belonging to the Class A subfamily of ﬂavodiiron proteins, is a ﬂavin mononucleotide containing dimer and is expected to catalyze the safe reduction of oxygen to water, similar to its bacterial homologues for which the function has been determined. However, the electron donor of the trichomonad enzyme is unknown and the expression of this protein in metronidazole-resistant strains has not yet been studied. Consistent with defective oxygen scavenging, trichomonad strains with aerobic-type resistance show decreased tolerance to oxygen, as demonstrated by various viability assays comparing drug-susceptible and drug-resistant strains (Ellis et al. 1994; Rasoloson et al. 2001). Thus, the presence of intracellular oxygen, although beneﬁcial to resistant parasites when they interact with metronidazole, can in turn decrease their viability under oxidative stress. The phenotype of aerobic resistance also includes increased activity of iron-containing superoxide dismutase (Fe-SOD), as observed in both clinical isolates and laboratory-induced T. vaginalis strains (Ellis et al. 1994; Rasoloson et al. 2001). Increased activity of the enzyme and upregulation of Fe-SOD gene expression has also been observed in Entamoeba histolytica selected in

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vitro for metronidazole resistance that appears to be of the aerobic type (Samarawickrema et al. 1997; Wassmann et al. 1999). It seems unlikely that the enzyme participates directly in mechanisms of resistance. Rather its increased activity in drug-resistant parasites could be a response to the presence of intracellular oxygen and its reactive metabolites. However, mechanisms of antioxidant defense of T. vaginalis that lacks catalase and glutathione-cycling enzymes remain to be elucidated. Although candidates for peroxide-reducing proteins have recently been identiﬁed (see Hrdý et al., this volume) no speciﬁc data on their function are yet available. Upregulation of peroxiredoxins have been reported in resistant Entamoeba (Wassmann et al. 1999). Altered ferredoxin function or an insufﬁcient amount of ferredoxin caused by defective transcription has been proposed to participate in mechanisms of aerobic resistance in T. vaginalis (Yarlett et al. 1986b; Quon et al. 1992). Downregulated transcription of one of the two genes for ferredoxin was also reported in laboratory-induced metronidazole-resistant E. histolytica (Wassmann et al. 1999). Our data do not conﬁrm involvement of ferredoxin deﬁciency in aerobic metronidazole resistance. No signiﬁcant differences in ferredoxin levels between the drug-susceptible T. vaginalis strain and its aerobically resistant derivative were found by Western blot analysis (Rasoloson et al. 2001, 2002). Also, identical signals for functional ferredoxin iron sulfur centers were detected in the drug-susceptible strain, its aerobically resistant derivative, and a clinical isolate displaying the aerobic resistance (Rasoloson et al. 2001). Moreover, all aerobically resistant strains of T. vaginalis examined thus far show ferredoxin-dependent hydrogenosomal PFOR pathway activity, both under aerobic and anaerobic conditions (Müller and Gorrell 1983; Ellis et al. 1992; Tachezy et al. 1993). Decreased expression of the ferredoxin gene (ehfd1) in Entamoeba was observed only in cells grown under microaerophilic conditions with 40 µM (6.8 µg/ml) metronidazole, the highest drug concentration at which the parasites can multiply. These cells survived prolonged exposure to 200 µM (∼34.2 µg/ml) metronidazole and could well be in a transition state toward anaerobic resistance. Downregulation of ferredoxin typically accompanies development of anaerobic-type resistance (see below). 3.2 Anaerobic Type of Metronidazole Resistance This type of resistance, detectable in vitro under anaerobic conditions, is known from laboratory-induced strains that lack drug activating pathways. 3.2.1 Development of Anaerobic Resistance Anaerobic resistance was developed in both pathogenic species by prolonged cultivation of drug-susceptible strains or clones under increasing pressure of

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metronidazole in vitro (Kulda et al. 1984, 1993; Brown et al. 1999). The resulting derivatives with fully developed resistance showed extremely high tolerance to metronidazole under anaerobic conditions (MLC around 1000 µg/ml, i.e., ∼5.9 mM in both species) and were able to multiply in culture in the presence of 100 µg/ml metronidazole (∼588 µM). The uptake of metronidazole by these strains was negligible or undetectable, indicating that the drug was not metabolized (Kulda et al. 1989). Accordingly, the signal for nitro free radicals was absent in the EPR spectra of strains with fully developed anaerobic resistance (Fig. 7c). In both species resistance was induced in a stepwise process including a sequence of stages that differed in levels and stability of resistance as well as metabolic properties. In T. vaginalis, a transient stage of aerobic resistance preceded development of anaerobic resistance (Rasoloson et al. 2002). At the ﬁnal stage the hydrogenosomal pathways responsible

Fig. 7 EPR spectra demonstrating generation of metronidazole radicals by hydrogenosomes of T. vaginalis of different susceptibility to metronidazole. Hydrogenosomes were obtained from the drug-susceptible strain TV 10-02 (a) and its derivatives at early phase of anaerobic resistance development TV 10-02 MR 5 (b) and with fully developed anaerobic resistance TV 10-02 MR 100 (c) . All samples were exposed to 43 mM metronidazole. a Drug-susceptible strain. Signal produced in the presence of pyruvate, CoA, and ferredoxin. b Early anaerobic resistance. Signal generated in the presence of malate, ferredoxin, and NAD+ . This strain does not produce a signal in the PFOR reaction (see Fig. 5c). c Fully developed anaerobic resistance. Absence of signal in the presence of malate, ferredoxin, and NAD+ . Experimental conditions as in Fig. 3

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for drug activation were eliminated and reshufﬂing of pyruvate metabolism in the cytosol compensated for energy loss caused by nonfunctional pyruvate catabolism in the hydrogenosomes. Full resistance was easier to induce in T. foetus where it developed faster (3–7 months) and could be attained by exposure of drug-susceptible parasites to constant and relatively low concentrations of metronidazole (3–5 µg/ml) (Kulda et al. 1984). In T. vaginalis full resistance took 1–2 years to develop and the process required stepwise selection by gradually increasing the pressure of the drug. In both species freshly acquired resistance to high concentrations of drug (growth at 100 µg/ml metronidazole) was usually unstable in the absence of drug pressure, and additional cultivation with 100 µg/ml metronidazole was necessary to obtain stable resistance. Unstable lines, however, did not revert to complete susceptibility but retained moderate levels of the resistance (growth at ≤10 µg/ml). 3.2.2 Changes in Metabolism and Protein Expression Development of anaerobic resistance in both species was accompanied by a progressive decrease and eventual loss of PFOR and hydrogenase activities and ferredoxin levels. For development of anaerobic resistance in T. foetus, ˇ erkasovová et al. the decrease in PFOR activity was of crucial importance (C 1984) being strictly paralleled by a decrease in metronidazole uptake and an increase in resistance levels (Kulda et al. 1989). In T. vaginalis, PFOR activity was lost at the early stage of anaerobic resistance development when parasites were still susceptible to moderate concentrations of drug (MLC <15 µg/ml, i.e., ∼88 µM). It was later found that the malic enzyme-dependent pathway of metronidazole activation is likely to be involved in the residual susceptibility, as indicated by persisting activities of malic enzyme, NDH (referred to as NADH : ferredoxin oxidoreductase) and by the presence of ferredoxin in hydrogenosomal extracts at this stage of resistance development (Rasoloson et al. 2002). These hydrogenosomes also generated metronidazole anion radicals as detected by EPR spectroscopy. However, the signal showed a lower amplitude than that of the parent, drug-susceptible strain and was only detectable in the presence of malate, ferredoxin, and NAD+ needed for malic enzyme activity, but not in the presence of pyruvate, CoA, and ferredoxin required for activity of PFOR (Hrdý et al. 2005) (Fig. 7b). Parasites with fully developed anaerobic resistance lost all of these activities and metronidazole anion radicals were not produced in their hydrogenosomes (Fig. 7c) (Kulda et al. 1989; Rasoloson et al. 2002; Hrdý et al. 2005). The activity of hydrogenase also decreased with increasing resistance in both species but the decrease was slower and the activity disappeared later than that of PFOR (Kabíˇcková et al. 1988; Rasoloson et al. 2002). However, the causal relationship between the gradual reduction in the hydrogenase activity and the development of metronidazole resistance seems unlikely and the phenomenon is more likely attributed to

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metabolic feedback. Examination of the protein spectra of T. vaginalis during development of resistance by Western blotting (Fig. 8) revealed that the decrease and ultimate loss in PFOR and malic enzyme activities directly reﬂected lowered levels or absence of the respective proteins (Rasoloson et al. 2002). Ferredoxin was detected until the early stage of anaerobic resistance when it was needed as an essential component of the alternative pathway of metronidazole activation, but disappeared in more advanced stages of resistance development (Fig. 8). As shown by Northern blotting, the decrease of steady-state levels of mRNA for PFOR and malic enzyme paralleled the pattern of decreasing protein expression (Rasoloson et al. 2002) (Fig. 9). Brown et al. (1999) also detected loss of mRNA for PFOR and ferredoxin by Northern blot analysis in another T. vaginalis strain induced for high-level anaerobic resistance. Transcription of T. vaginalis genes for PFOR and malic enzyme were further examined in the strain with fully developed anaerobic resistance and its drug-susceptible parent by the run-on assay for synthesis of nascent RNA (Rasoloson et al. 2002). Results revealed that PFOR deﬁciency is caused by downregulation of gene transcription. No marked changes were found in the transcription of malic enzyme genes, indicating that the expression of this protein is controlled at the mRNA level (Rasoloson et al. 2002). Land et al. (2001) examined the molecular basis of hydrogenosomal deﬁciency in T. foetus. It was also demonstrated in this species that a decrease or loss of enzyme

Fig. 8 SDS-PAGE (a) and Western blot (b) analysis of the puriﬁed hydrogenosomal fractions isolated from the metronidazole-susceptible T. vaginalis strain TV 10-02 (P) and its metronidazole-resistant derivatives MR-3, MR-5, MR-30, MR-50, and MR-100 displaying the aerobic (3), early anaerobic (5), advanced anaerobic (30, 50), and fully developed anaerobic resistance (100) to metronidazole. Numbers in the designation of MR strains indicate the concentrations of metronidazole in µg/ml at which the organisms multiply in culture. About 10 µg protein was loaded per line. PFOR: pyruvate : ferredoxin oxidoreductase, α-STK: α subunit of succinate thiokinase (hydrogenosomal enzyme not involved in metronidazole resistance; used as control), Fdx: ferredoxin. From Rasoloson et al. (2002) by courtesy of the Society of General Microbiology

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Fig. 9 Comparison of steady-state concentrations of mRNA encoding PFOR and hydrogenosomal malic enzyme in the drug-susceptible T. vaginalis strain TV 10-02 and its metronidazole-resistant derivatives with increasing level of resistance (see legend to Fig. 8). The amount of total RNA loaded onto each line was standardized according to the level of β-tubulin. From Rasoloson et al. (2002) by courtesy of the Society of General Microbiology

activities was associated with the decrease or absence of particular proteins. Northern blot analysis showed a reduction in steady-state levels of mRNA for PFOR, ferredoxin, hydrogenase, and malic enzyme. Quantitation of differences in mRNA levels between the parent strain and its derivative with fully developed resistance by phosphorimaging showed a reduction of between 60 (PFOR) and 98% (malic enzyme). The run-on transcription assay using nuclei from the drug-sensitive parent strain and its resistant derivative with fully developed anaerobic resistance showed decreased transcription of PFOR and malic enzyme genes in the resistant derivative. 3.2.3 Compensation of Hydrogenosomal Insufficiency As revealed by the above studies, both T. vaginalis and T. foetus acquire full anaerobic resistance to metronidazole at the cost of losing the hydrogenosomal pathway of pyruvate metabolism. In T. vaginalis the malate metabolizing pathway is eliminated as well. Consequently, the end products of the hydrogenosomal metabolism, acetate, H2 , and CO2 , are not formed and generation of hydrogenosomal ATP is abolished. To compensate for the hydrogenosomal insufﬁciency, both species increase the rate of glycolysis and adjust cytosolic catabolism of pyruvate to the production of a single dominant end product (Kulda et al. 1989). Metabolic balances of T. vaginalis with fully developed anaerobic resistance showed that over 92% of glucose is conˇ erkasovová et al. 1988). Accordingly, the activity of lactate verted to lactate (C dehydrogenase increased progressively with development of anaerobic resistance, exceeding sevenfold the original activity of the drug-susceptible parent strain (Rasoloson et al. 2002). The T. foetus strain with fully developed resistance converted over 90% of consumed glucose to ethanol, while only 10% of glucose was converted to this end product by its drug-susceptible par-

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ent. Enhanced ethanol fermentation was associated with a marked increase ˇ erkasovová et al. 1984). Although the in pyruvate decarboxylase activity (C glycolytic rate is increased in resistant trichomonads, the T. foetus strain with fully developed anaerobic resistance lacks activity of pyruvate kinase. It was found, however, that the high glycolytic ﬂux observed in these parasites is maintained exclusively through the malate bypass catalyzed by phosphoenolpyruvate carboxykinase, malate dehydrogenase, and malic enzyme. Accordingly, two overexpressed isoforms of cytosolic, NADP-dependent malic enzyme were detected and characterized and shown to provide the NADPH required for activity of the terminal enzyme in ethanol production, the alcohol dehydrogenase (Hrdý et al. 1993). 3.2.4 Hydrogenosomes of Metronidazole-Resistant Trichomonads The fact that trichomonads with fully developed anaerobic resistance can survive and multiply in the absence of the core hydrogenosomal catabolic pathway raised the question as to whether this organelle persists in trichomonads maintained for prolonged periods of time under high metronidazole pressure. Electron microscopic observations showed that hydrogenosomes are present (Kulda et al. 1987, 1989; Land et al. 2001). In addition, activities or proteins of some hydrogenosomal enzymes not directly involved in drug resistance (e.g., succinyl-CoA synthetase and ATP : AMP phosphotransferase) were detected in highly resistant strains of both T. vaginalis and ˇ erkasovová et al. 1984; Land et al. 2001; Rasoloson et al. 2002). T. foetus (C Electron microscopy revealed that the number of hydrogenosomes per cell in drug-resistant T. foetus is about the same as that in its drug-susceptible parent strain; their size, however, is smaller (Land et al. 2001). Our observations (Kulda et al. 1989) revealed marked changes in the ultrastructure of hydrogenosomes from drug-resistant T. foetus. While the organelles of the parent strain were spherical, those of the resistant derivatives had irregular shapes and contained remarkably enlarged electron-dense cores. In lines maintained for a long period under high drug pressure, hydrogenosomes extended into tubular protrusions interconnected with a reticulum of smooth membranes (Fig. 10). However, the functional signiﬁcance of these modiﬁcations is not known. The persistence of hydrogenosomes deﬁcient in pyruvate metabolism in resistant trichomonads is not yet fully understood. Mutants lacking hydrogenosomes did not appear in any of the T. vaginalis and T. foetus lines despite the high selective pressure imposed on them by cultivation with 100 µg/ml metronidazole for 1 to 3 years. This suggests that hydrogenosomes perform other functions essential for the parasite. Data from the T. vaginalis genome project offer a rich basis for speculations on this topic, but actual functional pathways in hydrogenosomes of metronidazoleresistant trichomonads remain to be resolved. Our preliminary results point

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Fig. 10 Ultrastructure of hydrogenosomes of the T. foetus strain KV1-MR100 with fully developed metronidazole resistance (a,b) and of its drug-susceptible parent clone KV1 (c,d). Note the irregular shape, enlarged electron-dense core, and presence of tubular extensions (arrows) in hydrogenosomes of the drug-resistant strain. Bars: 0.2 µm in a,b,c and 0.4 µm in d. Original electron micrographs from the author (JK)

to FeS center assembly. Components of this system have been identiﬁed in hydrogenosomes and some of them are upregulated in metronidazole-resistant T. vaginalis (unpublished data). In addition, an alternative 2-keto acid oxidoreductase activity that could utilize a broad range of 2-keto acids including pyruvate as a substrate was reported in the hydrogenosomes from both wildtype and resistant T. vaginalis (Brown et al. 1999). This activity could account for the residual production of acetate (more than one order of magnitude lower than in wild-type cells) observed in metronidazole-resistant lines (Rasoloson et al. 2002). The relationship between 2-keto acid oxidoreductase and PFOR homologues identiﬁed in the T. vaginalis genome (Carlton et al. 2007; Hrdý et al., this volume) remains to be determined. 3.3 Postgenomic Challenge Analysis of the data from the T. vaginalis genome sequencing project indicates that the mechanisms involved in acquisition of metronidazole resistance by this parasite might be more complex than currently understood.

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3.3.1 Multiple Genes Multiple genes encode the enzymes involved in metronidazole reduction, and the expression of their corresponding proteins during resistance development is only known for the dominant isoforms. The potential role of other isoforms of malic enzyme, PFOR, and ferredoxin in metronidazole activation is unknown. It was found that the dominant ferredoxin (ferredoxin 1), one of seven ferredoxin genes identiﬁed in the T. vaginalis genome (Carlton et al. 2007), is not indispensable for metronidazole activation since T. vaginalis cells did not acquire the resistant phenotype after the ferredoxin 1 gene was removed by homologous recombination (Land et al. 2004). 3.3.2 Other Potential Mechanisms of Metronidazole Resistance T. vaginalis was found to code for numerous enzymes that could play a role in metronidazole sensitivity and resistance in a way different from the known mechanisms. T. vaginalis possesses several NAD(P)H nitroreductases, likely cytosolic as indicated by the amino termini of the putative proteins which do not conform to hydrogenosomal targeting signals. Related proteins belonging to the nitroreductase family are implicated in metronidazole resistance in microaerophilic bacteria including the important human pathogen Helicobacter pylori, in which inactivation of the respective proteins is linked to drug resistance (Mendz and Mégraud 2002 and references therein). Proteins coded by the Nim gene family were also shown to confer 5-nitroimidazole resistance in certain Bacteroides fragilis group strains (Haggoud et al. 1994; Leiros et al. 2004 and references therein). The resistance mechanism appears to differ from that related to NAD(P)H nitroreductases, where the lack or inactivation of this enzyme is implicated in resistance. In the case of Nim proteins, it is the presence of Nim genes that is correlated with metronidazole resistance. Comparison of the metabolism of a 5-nitroimidazole-susceptible B. fragilis strain with the same strain harboring a plasmid coding for a NimA sequence revealed that the classical reduction of nitroimidazole to its nitro radical anion occurred in the sensitive strain, while the drug was reduced to a nontoxic amine derivative in the resistant, Nim-bearing strain. It was therefore proposed that Nim proteins function as 5-nitroimidazole reductases (Carlier et al. 1997). The crystal structure of the NimA nitroimidazole reductase from Deinococcus radiodurans has been deduced and a reaction mechanism involving covalently bound pyruvate has been proposed, in which two-electron reduction of metronidazole prevents formation of the toxic nitro radical anion (Leiros et al. 2004). T. vaginalis was found to code for three NimA homologues, two of which contain the typical hydrogenosomal targeting sequences on the amino ter-

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mini of translated gene products (one gene might be truncated) (Carlton et al. 2007). All T. vaginalis genes possess a conserved histidine residue that was found to bind to the catalytically important pyruvate in D. radiodurans (Leiros et al. 2004). The expression of the Nim homologues, which may have been acquired by lateral gene transfer from enteric bacteria, in T. vaginalis strains has not yet been studied, but the tagged, episomally expressed NimA homologue is targeted into hydrogenosomes of transformed trichomonads (our unpublished data).

4 Conclusions Hydrogenosomes, the sites of activation of the 5-nitroimidazole drugs, are also the cell compartments where the processes leading to development of drug resistance take place. Research focused on the biochemical mechanisms of metronidazole resistance have led to the discovery of an alternative pathway of metronidazole activation involving metabolic processes of general biological interest. It has been shown that the key enzyme in the alternative pathway of metronidazole activation is a remnant of mitochondrial-type complex 1, which possesses an unusual ability to reduce low redox potential electron transporter ferredoxin at the expense of NADH. The unexpected presence of such an enzyme in hydrogenosomes further supported the view of common evolutionary ancestry of mitochondria and trichomonad hydrogenosomes. The presence of hydrogenosomes in highly resistant cells shows that these organelles are probably not indispensable energy generators, but rather that they have other functions important for trichomonad cells. One of these functions is almost certainly a pathway forming FeS clusters, which has been identiﬁed in hydrogenosomes and may be the least common functional denominator of mitochondria, hydrogenosomes, and certain mitosomes. Other functions of hydrogenosomes (and we mean “resistant ones” as well) are likely to be uncovered when the data from the T. vaginalis genome sequencing project are completed with proteomic and biochemical studies. In addition, the phenomenon of anaerobic resistance itself points to a remarkable ability of unicellular organisms to selectively transform major pathways of their metabolism in response to external conditions. Of the two types of resistance identiﬁed, aerobic resistance is of clinical importance as most clinical isolates from treatment-refractory patients show resistance of this type. However, ﬁndings regarding the stepwise development of resistance, and continuity of resistance development from the aerobic to the anaerobic type, point to the possibility that T. vaginalis strains at the early stage of anaerobic resistance could also appear in the ﬁeld. In fact, some isolates reported from Upcroft’s laboratory (Dunne al. 2004) and occasional isolates from surveys (e.g., Lossick et al. 1987) displaying high aerobic MLC

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(up to 1000 µg/ml) show also increased anaerobic MLC (16–20 µg/ml, i.e., 94–147 µM metronidazole), compatible with the phenotype of early anaerobic resistance. Such isolates are usually associated with severe therapeutic problems. Although metronidazole resistance is not yet an alarming problem, it should be watched closely. All clinically available drugs are derivatives of 5-nitroimidazole with identical mechanisms of action and proven crossresistance (Meingassner and Mieth 1976; Upcroft et al. 1999). Clearly the development of new drugs with different antimicrobial mechanisms is desirable. Acknowledgements Numerous coworkers participated in the work on metronidazole resistance done at the Department of Parasitology, Charles University in Prague. In parˇ ticular, the cooperation of Jiˇrí and Apolena Cerkasov, Jan Tachezy, Hana Kabíˇcková, Eva Tomková, Dominique Rasoloson, and ˇ Stˇepánka Vaˇ náˇcová is gratefully acknowledged. This work was supported in part by grants from the Ministry of Education of the Czech Republic (OCB9.10, OCB22.001 (JK), MSM0021620858) and from the Czech Science Foundation (204/97/0263 (JK), 204/06/0944 (IH)).

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Land KM, Delgadillo-Correa MG, Tachezy J, Vaˇ náˇcová S, Hsieh CL, ˇ Sut’´ak R, Johnson PJ (2004) Targeted gene replacement of a ferredoxin gene in Trichomonas vaginalis does not lead to metronidazole resistance. Mol Microbiol 51:115–122 Leiros HKS, Kozielski-Stuhrmann S, Kapp U, Terradot L, Leonard GA, McSweeney SM (2004) Structural basis of 5-nitroimidazole antibiotic resistance. J Biol Chem 279:55840–55849 Lindmark DG, Müller M (1973) Hydrogenosome, a cytoplasmic organelle of the anaerobic ﬂagellate Tritrichomonas foetus, and its role in pyruvate metabolism. J Biol Chem 248:7724–7728 Lloyd D, Kristensen B (1985) Metronidazole inhibition of hydrogen production in vivo in drug-sensitive and resistant strains of Trichomonas vaginalis. J Gen Microbiol 131:849–853 Lloyd D, Pedersen JZ (1985) Metronidazole anion radical anion generation in vivo in Trichomonas vaginalis: oxygen quenching is enhanced in a drug-resistant strain. J Gen Microbiol 131:87–92 Lossick JG, Müller M, Gorrell TE (1986) In vitro drug susceptibility and doses of metronidazole required for cure in cases of refractory vaginal trichomoniasis. J Infect Dis 153:948–955 Meingassner JG, Mieth H (1976) Cross-resistance of trichomonads to 5-nitroimidazole derivatives. Experientia 32:183–184 Meingassner JG, Thurner J (1979) Strain of Trichomonas vaginalis resistant to metronidazole and other 5-nitroimidazoles. Antimicrob Agents Chemother 15:254–257 Meingassner JG, Mieth H, Czok R, Lindmark DG, Müller M (1978) Assay conditions and the demonstration of nitroimidazole resistance in Tritrichomonas foetus. Antimicrob Agents Chemother 13:1–3 Mendz GL, Mégraud F (2002) Is the molecular basis of metronidazole resistance in microaerophilic organisms understood? Trends Microbiol 10:370–375 Meri T, Jokiranta TS, Suhonen L, Meri S (2000) Resistance of Trichomonas vaginalis to metronidazole: report of the ﬁrst three cases from Finland and optimization of in vitro susceptibility testing under various oxygen concentrations. J Clin Microbiol 38:763–767 Moreno SN, Mason RP, Muniz RP, Cruz FS, Docampo R (1983) Generation of free radicals from metronidazole and other nitroimidazoles by Tritrichomonas foetus. J Biol Chem 258:4051–4054 Moreno SN, Mason RP, Docampo R (1984) Distinct reduction of nitrofurans and metronidazole to free radical metabolites by Tritrichomonas foetus hydrogenosomal and cytosolic enzymes. J Biol Chem 259:8252–8259 Müller M (1973) Biochemical cytology of trichomonad ﬂagellates. I. Subcellular localization of hydrolases, dehydrogenases, and catalase in Tritrichomonas foetus. J Cell Biol 57:453–474 Müller M, Gorrell T (1983) Metabolism and metronidazole uptake in Trichomonas vaginalis isolates with different metronidazole susceptibilities. Antimicrob Agents Chemother 24:667–673 Müller M, Lindmark DG (1976) Uptake of metronidazole and its effect on viability in trichomonads and Entamoeba invadens under anaerobic and aerobic conditions. Antimicrob Agents Chemother 9:696–700 Müller M, Lossick JG, Gorrell TE (1988) In vitro susceptibility of Trichomonas vaginalis to metronidazole and treatment outcome in vaginal trichomoniasis. Sex Transm Dis 15:17–24 Quon DV, d’Oliveira CE, Johnson PJ (1992) Reduced transcription of the ferredoxin gene in metronidazole-resistant Trichomonas vaginalis. Proc Natl Acad Sci USA 89:4402–4406

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Rasoloson D, Tomková E, Cammack R, Kulda J, Tachezy J (2001) Metronidazole-resistant strains of Trichomonas vaginalis display increased susceptibility to oxygen. Parasitology 123:45–56 Rasoloson D, Vaˇ náˇcová S, Tomková E, Rázga J, Hrdý I, Tachezy J, Kulda J (2002) Mechanisms of in vitro development of resistance to metronidazole in Trichomonas vaginalis. Microbiol-SGM 148:2467–2477 Samarawickrema NA, Brown DM, Upcroft JA, Thammapalerd N, Upcroft P (1997) Involvement of superoxide dismutase and pyruvate : ferredoxin oxidoreductase in mechanisms of metronidazole resistance in Entamoeba histolytica. J Antimicrob Chemother 40:833–840 Saraiva LM, Vicente JB, Teixeira M (2004) The role of the ﬂavodiiron proteins in microbial nitric oxide detoxiﬁcation. Adv Microb Physiol 49:77–129 Sobel JD, Nagappan V, Nyirjesi P (1999) Metronidazole resistant vaginal trichomoniasis— an emerging problem. N Engl J Med 341:292 Tachezy J, Kulda J, Tomková E (1993) Aerobic resistance of Trichomonas vaginalis to metronidazole induced in vitro. Parasitology 106:31–37 Upcroft P, Upcroft JA (2001) Drug targets and mechanisms of resistance in the anaerobic protozoa. Clin Microbiol Rev 14:150–164 Upcroft JA, Campbell RW, Benakli K, Upcroft P, Vanelle P (1999) Efﬁcacy of new 5-nitroimidazoles against metronidazole-susceptible and -resistant Giardia, Trichomonas and Entamoeba spp. Antimicrob Agents Chemother 43:73–76 Wagner G, Levin R (1978) Oxygen tension of the vaginal surface during sexual stimulation in humans. Fertil Steril 30:50–53 Wassman C, Hellberg A, Tannich E, Bruchhaus I (1999) Metronidazole resistance in protozoan parasite Entamoeba histolytica is associated with increased expression of iron-containing superoxide dismutase and peroxiredoxin and decreased expression of ferredoxin 1 and ﬂavin reductase. J Biol Chem 274:26051–26056 Yarlett N, Yarlett NC, Lloyd D (1986a) Metronidazole resistant clinical isolates of Trichomonas vaginalis have lowered oxygen afﬁnities. Mol Biochem Parasitol 19:111–116 Yarlett N, Yarlett NC, Lloyd D (1986b) Ferredoxin-dependent reduction of nitroimidazole derivatives in drug-resistant and susceptible strains of Trichomonas vaginalis. Biochem Pharmacol 35:1703–1708 Yarlett N, Rowlands CC, Evans JC, Yarloett N, Lloyd D (1987) Nitroimidazole and oxygen radicals detected by electron spin resonance in hydrogenosomal and cytosolic fractions from Trichomonas vaginalis. Mol Biochem Parasitol 24:255–261

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_113/Published online: 8 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Mitosomes in Parasitic Protists Jan Tachezy (u) · Ondˇrej ˇSmíd Department of Parasitology, Faculty of Science, Charles University in Prague, Vinicna 7, 12844 Prague 2, Czech Republic [email protected] 1

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Biogenesis of Mitosomes . . . . . . . . . . . . . The Genome . . . . . . . . . . . . . . . . . . . . Protein Targeting, Translocation, and Maturation Division and Segregation of Mitosomes . . . . . Division . . . . . . . . . . . . . . . . . . . . . . . Segregation . . . . . . . . . . . . . . . . . . . . .

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Physiological Functions . . . . . . . . . . . . . . . . . . . . . . . . . Iron–Sulfur Cluster Assembly Machinery . . . . . . . . . . . . . . . . The Mitochondrial Model . . . . . . . . . . . . . . . . . . . . . . . . The Role of Mitochondria in the Maturation of Extramitochondrial FeS Proteins . . . . . . . . FeS Cluster Assembly in Mitosomes . . . . . . . . . . . . . . . . . . Requirements for ATP, Membrane Potential, and Electron Transport Other Putative Functions . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Mitosomes are simple, mitochondrion-derived organelles, which were recently found in various “amitochondrial” protists, including the human parasites Entamoeba histolytica, Giardia intestinalis, Cryptosporidium parvum, and microsporidians. Similar organelles might also be present in some free-living protists. Although all these organisms underwent different evolutionary histories, they arrived at common life strategies for which oxygen-dependent ATP synthesis is not required: they inhabit either an oxygenpoor environment, such as the intestinal tract of their hosts, or they are adapted to intracellular parasitism. Consequently, the majority of their mitochondrial functions were permanently lost with concomitant loss of the organellar genome, and mitochondria gradually transformed into their highly reduced forms named mitosomes. The common features of mitosomes, which were retained and pointed to their mitochondrial origin, are a double membrane surrounding the organellar matrix, conserved mechanisms of protein import and processing, and the biosynthesis of iron–sulfur (FeS) clusters. Finding the latter function in mitosomes supports the notion that FeS cluster assembly is

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the only essential function of mitochondria necessary for the maturation of cellular FeS proteins. Only in the mitosomes of E. histolytica was the mitochondrion type of FeS cluster assembly machinery not conserved, and their function remains enigmatic. Unlike hydrogenosomes, another type of mitochondrion-derived organelle, mitosomes do not synthesize ATP and hydrogen. Many more investigations are required to elucidate the biology of mitosomes and the evolutionary paths leading to the formation of the various mitochondrion-derived organelles, of which mitosomes are the most simpliﬁed.

1 Introduction A parasitic lifestyle leads to the evolutionary adaptation of the parasitic organisms to the new niches of the host environment. As a result, the parasites can regulate their biochemical capabilities during their often complicated life cycles to optimize their metabolism within different hosts and host tissues. Such changes lead to various functional and consequently morphological modiﬁcations of the parasitic cells, including alterations in the functions and structure of mitochondria, the principal energy-generating organelles of eukaryotes. A textbook example is the mitochondrion of Trypanosoma. This organelle undergoes dramatic metabolic and structural changes during the trypanosome switch between the blood of the mammalian host and the digestive tract of the tsetse ﬂy. An excess of glucose in the mammalian host permits the bloodstream stage to mainly employ glycolysis for ATP generation. Consequently, the mitochondrion is repressed (van Dooren et al. 2005), lacking the ability to produce ATP by oxidative phosphorylation (Clayton and Michels 1996). In the insect vector, trypanosomes encounter a nutrient-poor environment, where they can only survive upon the activation of the mitochondrion. Structurally, these biochemical changes are reﬂected by the tubular cristae mitochondrion in its bloodstream forms (Fig. 1), which transforms to the platelike cristae-containing mitochondrion in its vector stages (Vickerman 1985). Some other parasites, including certain parasitic helminthes, remain within the same host; however, they migrate through different host tissues for part of their life cycle, entering the oxygen-poor environment of the host’s intestinal tract. These organisms adapted their mitochondria for either aerobic or anaerobic respiration according to the availability of oxygen (Komuniecki et al. 1993). The function of mitochondria is often altered in intracellular parasites adapted to life within the host cell. For example, the mitochondria of the species Plasmodium lack the pyruvate dehydrogenase complex converting pyruvate to acetyl-CoA, a substrate of the TCA cycle. Although genes encoding all components of the TCA cycle as well as ATP synthase were found in the Plasmodium genome, it is believed that most if not all ATP is produced by glycolysis (van Dooren et al. 2006). In more extreme cases, the adaptive and reductive course of evolution led to permanent rejection of a number of metabolic pathways, including

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Fig. 1 Morphology of mitochondria, hydrogenosomes, and mitosomes in parasitic protists. A Ciliate mitochondrion with platelike and tubular cristae. B Mitochondrion of bloodstream stage of Trypanosoma brucei with tubular cristae. C Hydrogenosome from Trichomonas vaginalis. The arrow points to the intramembrane vesicle. D–G Mitosomes from D Mastigamoeba balamuthi, E Entamoeba histolytica, F Giardia intestinalis, and G Trachipleistophora hominis. Arrows and arrowheads indicate outer and inner membranes, respectively. C Kindly provided by J. Kulda, Charles University in Prague, Czech Republic. Figures reproduced with permission from the International Journal for Parasitology, Elsevier Limited (B), the American Society for Microbiology (E), Nature Publishing Group, Macmillan Limited (F, G). Scale bars: B–E 100 nm, F, G 50 nm

those residing in mitochondria, and consequently to the transformation of mitochondria to organelles known as hydrogenosomes and mitosomes (syn. cryptons, mitochondrial remnants). Hydrogenosomes have been found in unicellular eukaryotes (protists), parasites, or comensals, typically inhabiting the intestinal or urogenital tract, as well as in some free-living protists inhabiting oxygen-poor environments. They evolved repeatedly, being found in phylogenetically distant protists, including trichomonads, anaerobic ciliates, fungi, and heteroloboseans (Embley et al. 1995; Yarlett and Hackstein 2005). The common feature of hydrogenosomes is the production of molecular hydrogen, which is coupled with ATP synthesis by substrate-level

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phosphorylation. They are unable to generate ATP by oxidative phosphorylation and a number of mitochondrial functions have been lost (see Hrd´y et al., this volume). Yet the most reduced mitochondria-derived organelles are mitosomes, in which only a single biosynthetic pathway mediating the formation of FeS clusters has so far been found. This chapter reviews the current status of knowledge in mitosome research comparing the morphology, biogenesis, and function of mitosomes in various lineages of parasitic protists.

2 Mitosomes, “Novel” Organelles in “Ancient” Protists? Mitosomes have been found in two types of unrelated parasites employing considerably different living strategies: (1) anaerobic parasites invading the intestinal tract—Giardia (Tovar et al. 2003), Entamoeba (Mai et al. 1999; Tovar et al. 1999), and (2) intracellular parasites invading various cell tissues— apikomplexan Cryptosporidium (Riordan et al. 1999), microsporidia (Williams et al. 2002). Moreover, it is likely that mitosomes are also present in some anaerobic free-living protists (see Hampl and Simpson, this volume). Though all these organisms underwent different evolutionary paths, they arrived at common life strategies, for which oxygen-dependent ATP synthesis is not required. Consequently, the majority of mitochondrial functions were permanently lost. However, until recently, the absence of “canonical” mitochondrial pathways and cellular structures conforming to typical mitochondria in several groups of protists was interpreted as the absence of mitochondria, and such organisms were labeled “amitochondriates”. The lack of mitochondria was considered to be a result of two possible scenarios. (1) According to the Archezoa hypothesis, contemporary amitochondriates were the direct descendents of primitive amitochondrial proto-eukaryotes, which separated from the main eukaryotic trunk before the endosymbiotic acquisition of mitochondria. Based on this attractive hypothesis, four amitochondrial groups including Archamoebae (for example Entamoeba histolytica), Metamonads (Giardia intestinalis), Microsporidia (Encephalitozoon cuniculi), and Parabasala (Trichomonas vaginalis) were incorporated into the taxon named Archezoa to indicate their ancient origin (Cavalier-Smith 1987a,b). Two types of amitochondriates were later distinguished: Type I amitochondriates, in which energy metabolism is not compartmentalized (Giardia, Entamoeba), and Type II amitochondriates possessing ATP-generating hydrogenosomes (trichomonads) (Martin and Müller 1998). (2) According to the second scenario, the amitochondriates lost mitochondria secondarily due to their speciﬁc adaptation to anaerobic or oxygen-restricted conditions. The second scenario obviously explains their amitochondrial status in organisms which belong to the monophyletic taxons, together with organisms possessing typical mitochondria such as the apicomplexan Cryptosporidium. Indirect evidence

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supporting the secondary loss of mitochondria also in E. histolytica came from the phylogenetic analysis of its small subunit ribosomal RNA, which places this parasite after the branch of mitochondrion-harboring protists such as kinetoplastids (Sogin 1991). Similarly, a phylogenetic analysis of tubulins (Keeling and Doolittle 1996) and the largest subunit of the RNA polymerase II (Hirt et al. 1999) of microsporidians revealed the afﬁnity of these protists to fungi, which typically possess mitochondria. It is generally accepted that the majority of genes of the endosymbiotic ancestor, which gave rise to mitochondria, was either lost or transferred into the host nucleus. In order to provide more direct evidence of the secondary loss of mitochondria, further investigations were focused on uncovering genes of mitochondrial origin, which would have been transferred from the mitochondrial to the nuclear genome if mitochondria were once present and which might be retained in contemporary amitochondriates. Indeed, various genes regarded as mitochondrial in origin, including mitochondrial-type heat shock protein 70 (mtHSP70) (Arisue et al. 2002; Bui et al. 1996; Germot et al. 1996, 1997; Morrison et al. 2001), chaperonin 60 (CPN60) (Bui et al. 1996; Clark and Roger 1995; Horner et al. 1996; Roger et al. 1998), valyltRNA synthetase (Hashimoto 1998), pyridine-nucleotide transhydrogenase (PNT) (Clark and Roger 1995), cysteine desulfurase IscS (Tachezy et al. 2001), and other components of the FeS cluster assembly machinery (Katinka et al. 2001), were identiﬁed in the genomes of all members of Archezoa groups. The next logical step, which led to the discovery of mitosomes, was to determine the cellular localization of proteins coded by genes of mitochondrial origin. The ﬁrst mitosomes (cryptons) were described in E. histolytica (Mai et al. 1999; Tovar et al. 1999). Two research groups showed in parallel studies that amoebic CPN60 was expressed and strongly induced at elevated temperature in this parasite. Western blotting using anti-CPN60 antibody revealed the presence of the protein in microsomal cellular fractions. The organellar localization of CPN60 was conﬁrmed by immunoﬂuorescence microscopy in wild-type cells as well as in Entamoeba transfected with a recombinant vector expressing CPN60 tagged with a c-myc epitope. Subsequently, transmission electron microscopy of the mitosome-enriched fraction showed organelles bound by a double membrane (Ghosh et al. 2000). Analysis of the Encephalitozoon cuniculi genome sequence, which revealed the presence of several genes coding for proteins with putative mitochondrial function, led to a prediction of mitosome existence in microsporidia (Katinka et al. 2001). Direct evidence came from studies of Trachipleistophora hominis, an intracellular parasite causing severe myositis in patients with AIDS. Williams et al. (2002) demonstrated the localization of Trachipleistophora mtHSP70 to a number of double-membrane-bound organelles by means of immunoﬂuorescence and immunoelectron microscopy. Preliminary evidence of a mitosome (relict mitochondrion) in Cryptosporidium was provided by an electron microscopic study that revealed the

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presence of a single double-membrane-bound organelle in this apicomplexan (Riordan et al. 1999). Subsequently, CPN60 (Riordan et al. 2003) and HSP70 (Slapeta and Keithly 2004) were detected within this organelle by immunomicroscopy. The discovery of the FeS cluster assembly as an essential mitochondrial function provided new markers for tracing mitochondrion-related organelles, such as cysteine desulfurase IscS and the molecular scaffold protein IscU (Tachezy and Dolezal 2007). These two markers were used for the identiﬁcation of mitosomes in G. intestinalis (Tovar et al. 2003). Speciﬁc antibodies against Giardia IscS and IscU uncovered their localization in double-membranebound organelles using immunomicroscopy and cell fractionation. In subsequent studies, Giardia CPN60 and mtHSP70 were detected within the same cellular compartment as IscS and IscU (Regoes et al. 2005). G. intestinalis was for a while considered one of the most solid candidates to represent the deepest diverging lineages of amitochondrial eukaryotes (Adam 2001; Best et al. 2004). Thus, the ﬁnding that Giardia also possesses a mitochondrion-like organelle strongly contributed to the corrosion of the amitochondriate concept. The current evidence indicates that neither Type I nor Type II amitochondriates are truly amitochondrial organisms. They both possess organelles, respectively mitosomes or hydrogenosomes, which could be viewed as highly modiﬁed mitochondria (see Martin, this volume).

3 Morphology of Mitosomes Mitosomes are tiny ovoid or elongated organelles that measure less than 0.5 µm in length. In electron micrographs, the organelles appeared to be limited by two membranes, which envelop a granular electron-dense matrix. Mitosomes do not form cristae, the inner membrane infoldings, which are typically present in respiring mitochondria. Morphologically, mitosomes are reminiscent of the mitochondria found in yeast cells grown anaerobically, “acristae” mitochondria in Plasmodium merozoits (Fry and Beesley 1991), as well as hydrogenosomes found in some anaerobic protists (Fig. 1). The identiﬁcation of the double-limiting membrane in mitosomes was critical to the argument for their relationship with mitochondria (Ghosh et al. 2000; Riordan et al. 1999; Tovar et al. 2003; Williams et al. 2002). The detailed morphology of mitosomal double membranes, however, is not identical for the mitosomes observed in different species. Electron micrographs of the mitosomal membranes in Entamoeba and Trachipleistophora and in the mitosome-like organelles in Mastigamoeba showed that they are formed by two closely opposed membranes without an apparent intramembrane space (Fig. 1; Hampl and Simpson, this volume). The membranes are visible as three lipidic layers: the outer layer of the outer membrane, the inner layer of the inner membrane, and the central

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lipidic bilayer. This arrangement is very similar to that observed in hydrogenosomes. In these organelles, the central bilayer separates around calcium-rich intramembrane vesicles, which are surrounded by two complete membrane units (Benchimol, this volume). However, such intramembrane vesicles have not been observed in mitosomes so far. An intramembrane space reminiscent of mitochondria was observed in Giardia mitosomes and mitosome-like organelles in some microsporidians. Also, the size and number of mitosomes varied in different cells. In initial studies of E. histolytica a single mitosome was observed in most cells using immunoﬂuorescence microscopy, but a few cells contained two, and rarely three, organelles (Mai et al. 1999; Tovar et al. 1999). The size of mitosomes was estimated to be 0.5–1.0 µm based on electron microscopy micrographs. However, in a later study, confocal microscopy images revealed that mitosomes are rather abundant organelles of over 150 mitosomes per Entamoeba trophozoit with an estimated size of less than 0.5 µm for most organelles (Leon-Avila and Tovar 2004). Giardia mitosomes are also abundant structures, ranging from 25 to 100 per cell (Doleˇzal et al. 2005; Regoes et al. 2005; Tovar et al. 2003). However, they are considerably smaller, with a size of less than 0.2 µm (an average of 184×140 nm). The majority of Giardia mitosomes are randomly distributed throughout the cytoplasm, often in lateral and posterior regions of the trophozoits. Interestingly, some mitosomes form a distinct rodlike structure between the two Giardia nuclei in close proximity to the basal bodies. This structure, which is invariably present in all cells, consists of several attached mitosomes organized between the axonemes of caudal ﬂagella (Fig. 2). It was suggested that this central structure might be involved in the biogenesis and inheritance of Giardia mitosomes (Regoes et al. 2005). The microsporidian Trachiplestophora hominis contains between 7 and 47 mitosomes, with an average of 28 mitosomes per cell observed throughout the cytoplasm. With a size of 50×90 nm estimated by electron microscopy, these structures are the smallest mitosomes to have been observed thus far (Williams et al. 2002). Similar structures have been observed in other microsporidian species, including the meronts and sporonts of Vavraia culicis from Spodoptera exigue, Amblyospora sp. from Cyclops strenuus, Vairimorpha sp. from Lymantria dispar, and Marssoniella elegans parasitizing Cyclops vicinus (Vávra 2005). Unlike in Trachiplestophora, only a few mitosome-like organelles are freely scattered in the cytoplasm of the other microsporidia. Typically, they form a group of organelles which are associated with a spindle plaque, situated in a depression of the nuclear membrane (Fig. 3). The mitosome-like organelles associated with a spindle plaque probably correspond to the “polar vesicles” previously reported in various ultrastructural studies of microsporidia (Vávra 1976; Youssef and Hammond 1971). The mitosomes in these species are oval, about 0.15–0.5×0.1–0.25 µm in size. The apicomplexan Cryptosporidium contains a single mitosome (relict mitochondrion) which is

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Fig. 2 Mitosomes of Giardia intestinalis localized between axonemes (a) of caudal ﬂagella close to basal bodies (n, nucleus). Scale bar = 500 nm. Kindly provided by E. Noh´ ynková, Charles University in Prague, Czech Republic

Fig. 3 Mitosome of Vavraia culicis associated with a spindle plaque (n, nucleus; s, spindle plaque; m, mitosome). Scale bar = 200 nm. Kindly provided by J. Vávra, Charles University in Prague, Czech Republic

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invariably localized to the posterior part of the cell between the nucleus and crystalloid body. The size of the organelle estimated by electron microscopy ranges between 0.2 and 0.5 µm in diameter (Putignani et al. 2004; Riordan et al. 2003). The organelle is enveloped by a rough endoplasmic reticulum that extends from the outer nuclear membrane. A tomographic reconstruction showed that the inner mitosomal membrane of Cryptosporidium is highly folded without a cristae-like connection, or it forms independent vesicles (Keithly, this volume).

4 Biogenesis of Mitosomes Mitosomal biogenesis, as well as the biogenesis of mitochondria, is a complex process of maintenance and multiplication of the organelle in the cell. It also includes segregation of the organelles during the cell cycle. 4.1 The Genome No evidence has been found for the mitosomal genome in any organism so far. An earlier study on E. histolytica reported the presence of DNA in Entamoeba mitosomes (Ghosh et al. 2000) and synonymized these organelles with DNA-containing structures named kinetoplast-like organelles (EhKO) (Orozco et al. 1997). However, work by Leon-Avila and Tovar (2004) later showed that EhKO and mitosomes are not related structures. Moreover, in situ nick translation coupled with immunoﬂuorescence microscopy failed to detect the presence of DNA in Entamoeba mitosomes (Leon-Avila and Tovar 2004). In contemporary mitochondria, the genome of the original endosymbiotic ancestor is retained to various extents that range from 97 genes in the protist Reclinomonas (Lang et al. 1997) to ﬁve genes in Plasmodium (Wilson and Williamson 1997). Even though many mitochondrial genomes contain only a few genes, no mitochondrion lost its genome completely. Several hypotheses have been proposed to explain why: (1) the import of highly hydrophobic proteins from cytosol to mitochondria is not efﬁcient; (2) certain proteins coded in the mitochondrial genome are toxic when present in the cytosol; (3) the retention of genes, whose products are involved in electron transport and energy metabolism, may reﬂect the need for precise regulation of their expression by the redox state of the mitochondrion; and (4) in some eukaryotes, mitochondrial genes may be trapped in the mitochondrial genome by a nonstandard genetic code usage in the organelle (Adams and Palmer 2003). Interestingly, two genes, coding for cytochrome b (cyt b) and subunit I of cytochrome c oxidase (cox1), are present in all mitochondrial genomes known

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to date. These genes may represent core components of the mitochondrial genome whose transfer to the nucleus is not possible, for any of the reasons mentioned above. Both cyt b and cox1 are components of the respiratory chain, which is linked to ATP production by oxidative phosphorylation. Although these pathways are typically present in mitochondria, they are absent in mitosome-bearing organisms. Thus, the loss of a cytochrome-dependent respiratory chain might be the reason why the mitosomal genome was not required any more for the functioning of the organelles, and consequently the genome was completely eliminated. 4.2 Protein Targeting, Translocation, and Maturation Like the mitosome, most organelles in a eukaryotic cell do not possess a genome and translation machinery. Consequently, proteins must be imported to these compartments from the cytosol. Yet the membranes separating these organelles from the cytosol are not freely permeable for large hydrophilic molecules of proteins. As a result, the protein translocation is mediated by membrane transporters in a complex energy-consuming process. The mode of protein translocation across these barriers is typical for each compartment. Although mitochondria harbor DNA, the majority of their genetic information has been transferred to the nucleus. The information for the import of mitochondrial proteins synthesized in the cytosol is carried either within their N-terminal extension (targeting presequence of matrix proteins) or within the polypeptide chain (internal targeting signals of membrane proteins). The primary structure of the targeting signals is not conserved. However, the secondary structure of N-terminal presequences exhibits a consensus conformation of an amphipathic alpha helix with positively charged and apolar sides. Mitochondrial import signals are recognized by receptors on the outer mitochondrial membrane. These receptors are clustered together with a general import pore to the complex called TOM (translocase of the outer mitochondrial membrane). Once proteins pass through the outer membrane, they follow one of three major import pathways. (1) Presequence-carrying proteins are directed to the translocase of the inner membrane called the TIM23 complex. The pore within TIM23 cooperates with mitochondrial HSP70 and other proteins that form the presequence translocase-associated motor (PAM) that pulls the imported protein to the matrix. Once the protein reaches the matrix, the targeting sequence is cleaved off by a matrix-localized mitochondrial processing peptidase. (2) Proteins of the inner membrane are guided by small chaperones through the intermembrane space to the TIM22 complex. The pore of TIM22 inserts these proteins into the mitochondrial inner membrane. Only a few proteins follow an alternative route, being exported to the inner membrane from the mitochondrial matrix by the OXA

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complex. (3) Outer membrane proteins are integrated into the membrane by the sorting and assembly machinery (SAM) complex (for more details see Wiedemann et al. 2004; Dyall and Dolezal, this volume). Mitosomes of G. intestinalis have been demonstrated to share a common mode of protein targeting with hydrogenosomes and mitochondria (Dolezal et al. 2005; Regoes et al. 2005; see also Dyall and Dolezal, this volume). Targeting sequences resembling the N-terminal extensions of mitochondrial proteins are required for the efﬁcient import of some proteins into the Giardia mitosome. Namely, the role of the targeting sequences of the iron–sulfur cluster assembly protein IscU and mitosomal ferredoxin has been demonstrated. Moreover, the IscU and ferredoxin targeting sequences were able to direct proteins into the hydrogenosomes of T. vaginalis (Dolezal et al. 2005) and the mitochondria of human kidney cells (Regoes et al. 2005). However, the mitosomal matrix proteins IscS, CPN60, and mtHSP70 lack N-terminal targeting sequences and import into the mitosome is driven by an internal signal. A characterization of the translocase complexes responsible for mitosomal protein import in Giardia is under way. An iterative BLAST search of the G. intestinalis genome was employed for the identiﬁcation of the import machinery components. The only candidate to have been found so far is GiPAM18, a mitosomal protein homologous to mitochondrial presequence translocase-associated motor 18 (Doleˇzal et al. 2005). After translocation into the mitosome, the N-terminal presequences of mitosomal proteins are cleaved off, most likely by a peptidase that is homologous to the mitochondrial processing peptidase (MPP). The MPP is a matrix-localized metallopeptidase with a zinc binding motif His-X-X-GluHis that is conserved in the putative mitosomal processing peptidase reported in G. intestinalis (Doleˇzal et al. 2005). Recently, a mitosomal protein import machinery has been described in microsporidia (Burri et al. 2006). Some mitosomal proteins of microsporidia carry short N-terminal targeting presequences. In some cases, when expressed in the yeast Saccharomyces cerevisiae, these presequences are able to direct proteins into mitochondria. Other proteins possess an internal targeting signal or both a presequence and an internal signal. Even though the mitosomal translocase machinery of microsporidia is homologous to that of mitochondria, it is highly reduced. Only two components of the TOM complex are present (Tom40 and Tom70). TIM22 and TIM23 complexes are reduced to a single protein each (Tim22 and Tim17, respectively). The mitosomal presequences of matrix proteins are not processed by a homologue of the mitochondrial processing peptidase that is absent in microsporidia. As these presequences are short, they probably do not interfere with the function of matrix proteins. Intermembrane space proteins require the processing of another type of protease, the inner membrane peptidase. A homologue of this enzyme has been demonstrated in the microsporidia Antonospora, while it is missing in Encephalitozoon. It has been hypothesized that the mitosome

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of the latter organism represents a further stage of reductive adaptation of the pristine fungal mitochondrion (Burri et al. 2006). 4.3 Division and Segregation of Mitosomes Structures of mitosomes resembling dividing mitochondria have been occasionally observed on some electron micrographs (Keithly et al. 2005; our observations). However, no information on the molecular mechanisms of mitosome division is available and only little is known about their segregation during cytokinesis. 4.3.1 Division Bacteria, the progenitors of mitosomes, mitochondria, and hydrogenosomes, employ the ring-forming protein FtsZ in their division (Errington et al. 2003). Some eukaryotes retained FtsZ (Cyanidioschyzon merolae of Rhodophyta, Cyanophora paradoxa of Glaucophyta, Mallomonas splendens of Heterokontophyta, and Dictyostelium of Amoebozoa) to be employed in the division of mitochondria (Kuroiwa et al. 2006). In most eukaryotic organisms, including parasitic protists, FtsZ has been lost and mitochondrial ﬁssion depends on a dynamin-related protein along with other, as yet unknown, proteins that form a complex mitochondrial division apparatus (Kuroiwa et al. 2006). Dynamins are GTPases with various functions that include the formation of clathrin-coated endocytic vesicles. Interestingly, T. brucei, a kinetoplastid protist, exploits a single dynamin-related protein for both endocytosis and mitochondrial division (Chanez et al. 2006). However, the only dynaminrelated protein of G. intestinalis is involved in the scission of budding vesicles, but not in the division of mitosomes. This ﬁnding supports the hypothesis that the original function of dynamin-related proteins was endocytosis (Chanez et al. 2006; Kuroiwa et al. 2006); however, the molecular mechanisms of Giardia mitosome division remains unknown. Similarly, Spironucleus, a diplomonad protist related to G. intestinalis, and E. cuniculi of Microsporidia possess a single dynamin-related protein (Andersson et al. 2007). However, whether this protein is involved in division of their mitosomes remains to be elucidated. 4.3.2 Segregation In the highly organized eukaryotic cell, organelles including mitochondria are not randomly distributed within the cytosol but their position is determined by speciﬁc interactions with one or more of the three cytoskeletal

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systems (microtubules, microﬁlaments, and intermediate ﬁlaments). Mechanisms that control the movement of mitochondria are highly diverse, even within a group of closely related organisms. For example, the migration of mitochondria in the cells of S. cerevisiae depends on actin ﬁlaments (Boldogh et al. 2005), while the related fungus Schizosaccharomyces pombe uses microtubule tracks to control mitochondrial motility (Yaffe et al. 2003). Precise segregation of mitochondria into daughter cells during cytokinesis is essential, since the organelles cannot be formed de novo. This process is mediated by the same cytoskeletal components that are involved in mitochondrial movement during interphase (Boldogh et al. 2005; Yaffe et al. 2003). Considering the origin of mitosomes, the mitosomal segregation during cytokinesis may be analogous to the segregation of mitochondria. For example, a recent electron microscopic study of Microsporidia showed the association of mitosomes with electron-dense depressions of the nuclear membrane. Since these structures are mitotic spindle organization centers of microsporidia, they may be responsible for the distribution of mitosomes into daughter cells (Vávra 2005). Much less is known about the movement and segregation of mitochondria in the cells of other protists. As for the Excavata group, the protist T. brucei harbors a single mitochondrion. The organelle is connected with the ﬂagellar basal body by ﬁlaments that span both mitochondrial membranes and bind mitochondrial DNA. Replication of the basal body and its segregation into daughter cells is responsible for the precise segregation of mitochondria during cell division (Ogbadoyi et al. 2003). In another excavate protist, G. intestinalis, two groups of mitosomes can be distinguished: peripheral mitosomes that are dispersed throughout the cell and the so-called central mitosomes that are in contact with the basal bodies of ﬂagellar axonemes. The latter were reported to divide together with ﬂagellar basal bodies before mitosis. A connection between daughter basal bodies and mitosomes has been proposed to be responsible for the segregation of mitosomes into daughter cells (Regoes et al. 2005). However, the basal bodies of the eight ﬂagella of G. intestinalis do not replicate before the start of mitosis, as previously thought, but during the complex process of parental ﬂagella migration (Noh´ynková et al. 2006). Therefore, precise mitosomal segregation must be achieved by a different mechanism. According to our observations, the central mitosomes separate during mitosis and are transported to daughter cells by association with the axonemes of caudal ﬂagella. These are the only ﬂagella that neither change their function nor position in daughter cells (Fig. 4). C. parvum is a protist related to the apicomplexans P. falciparum and Toxoplasma. P. falciparum and T. gondii possess the apicoplast organelle, a remnant plastid acquired by secondary symbiosis. It has been shown that the T. gondii apicoplast is segregated into daughter cells by microtubules of mitotic spindle poles (Striepen et al. 2000). As the apicoplast is in contact with the mitochondrion, it has been hypothesized that mitochondrial inheritance

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Fig. 4 Immunoﬂuorescence microscopy of Giardia intestinalis showing segregation of mitosomes during mitosis. A Interphase, B prophase, and C telophase cells. Nuclei were stained by DAPI (blue), mitosomes were detected by an antibody raised against GiiscU (red), and axonemes were visualized by the antibody AXO 49 recognizing polyglycylated carboxy-terminal peptides of α- and β-tubulin (green). Note the proximity of mitosomes (white arrows) to axonemes

is mediated indirectly via the apicoplast (van Dooren et al. 2005). It is not yet known how mitosomal inheritance is controlled in C. parvum. However, as C. parvum is the apicomplexan which lacks an apicoplast, direct contact between the mitosomes and mitotic spindle might be expected. There is still no information available on the movement and segregation of the mitosomes of E. histolytica.

5 Physiological Functions The selection pressure orchestrating the mitochondria-to-mitosome transition was apparently different in different organisms. Nevertheless, it resulted in the formation of similar, albeit not identical, organelles, which completely lost their energy metabolism and the majority of other mitochondrial functions over the course of reductive evolution. Unlike mitochondria and hydrogenosomes, mitosomes are not able to synthesize ATP. There is no evidence of the pyruvate dehydrogenase complex producing acetyl-CoA, components of the tricarboxylic acid cycle generating electrons, or of the cytochrome-dependent respiratory chain required for oxidative phosphorylation. Pyruvate:ferredoxin oxidoreductase, which generates acetyl-CoA for substrate level phosphorylation in hydrogenosomes as well as hydrogenase, also seems to be absent in mitosomes. The activity of the former enzyme was detected in nonsedimentable fractions of mitosome-bearing protists (Lindmark 1980; Reeves et al. 1977), although its association with some cellular structures cannot be excluded (Ellis et al. 1993; Rodriguez et al. 1998). There is no evidence of mitochondrial biosynthetic pathways, including the biosynthesis of heme, biotin, and cardiolipin, as well as for the urea cycle, fatty acid degradation, and metabolism of amino acids and nucleotides. The only

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known mitochondrion function to be retained in the vast majority of mitosomes is the biosynthesis of FeS clusters. This recently discovered pathway seems to represent the only essential function operating in mitochondria, hydrogenosomes, and mitosomes (Tachezy and Doleˇzal 2007). 5.1 Iron–Sulfur Cluster Assembly Machinery FeS clusters represent one of the most ancient ubiquitous prosthetic groups, and are essential to the function of a number of proteins involved in numerous distinct metabolic pathways (Johnson and Dean 2004). Proteins containing FeS clusters in their active sites (FeS proteins) mediate electron transport, oxygen sensing, iron homeostasis, and various enzymatic catalyses. In bacteria, there are at least three independent systems responsible for FeS cluster assembly. The ISC (iron–sulfur cluster) assembly machinery provides FeS clusters for the maturation of various “housekeeping” FeS proteins (Zheng et al. 1998). The NIF (nitrogen-ﬁxing) system is primarily dedicated to forming speciﬁc FeS clusters in the nitrogenase of nitrogen-ﬁxing bacteria (Kennedy and Dean 1992). Finally, the SUF (sulfur mobilization) system most likely repairs FeS clusters under oxidative stress and iron-restricted conditions (Takahashi and Tokumoto 2002). In eukaryotes, FeS cluster assembly is considered to be the only essential function of mitochondria (Lill et al. 1999). The machinery mediating this process was shown to be of an ISC type. Phylogenetic analyses indicate that the mitochondrial ISC machinery was inherited from the proteobacterial endosymbiont, which is consistent with the proposed origin of mitochondria (Tachezy et al. 2001). Components of the SUF system were found in plastids and in the apicoplast of apicomplexan such as P. falciparum (Wilson et al. 2003). The SUF machinery was most likely inherited from cyanobacteria, the ancestors of plastids (Tachezy et al. 2001). Entamoebids and related protists are the only eukaryotes which acquired components of the NIF system (Ali et al. 2004; van der Giezen et al. 2004). 5.1.1 The Mitochondrial Model The current model for FeS cluster formation is based mainly on studies of Saccharomyces and human cells (reviewed in Lill and Mühlenhoff 2006; Rouault and Tong 2005; Tachezy and Doleˇzal 2007). The pyridoxal phosphatedependent cysteine desulfurase (IscS) is a central component catalyzing the production of sulfur from l-cysteine (Li et al. 1999; Schwartz et al. 2000). The released sulfur is transferred to IscU which constitutes a scaffold for the formation of a transient FeS cluster (Agar et al. 2000). Subsequently, the FeS cluster is transferred to mitochondrial apoproteins with the assistance of the

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HSP70 (Ssq1) chaperone, J-type (Jac1) cochaperone, and the nucleotide exchange factor Mge1 (Dutkiewicz et al. 2003). The process is dependent on the donation of reducing power, which is likely facilitated by [2Fe2S] ferredoxin (Yah1p) and ferredoxin reductase (Arh1p). There are several more components that are associated with FeS cluster assembly, however their role is still uncertain. Frataxin, a mitochondrial protein whose impaired expression is associated with Friedreich’s ataxia in humans, interacts with the IscS/IscU complex (Gerber et al. 2003; Shan et al. 2007). It was suggested that frataxin provides iron for the FeS cluster assembly and reparation, although the function of this protein is still controversial (Campuzano et al. 1996). Frataxin interacts with the Isd11 protein, which also contributes to the formation of the IscS/IscU complex (Shan et al. 2007). It was suggested that glutaredoxin (Grx5) is involved in the regulation of the redox state of important cysteine residues in IscS and/or IscU (Alves et al. 2004; Rodriguez-Manzaneque et al. 2002). Two proteins, IscA and Nfu, were proposed to serve as an alternative scaffold for FeS cluster assembly (Ollagnier-de-Choudens et al. 2001; Schilke et al. 1999; Wu et al. 2002). Interestingly, both IscU and Nfu are homologous to different domains of NifU, which functions in the speciﬁc assembly of FeS clusters in nitrogenase as a part of the NIF system. 5.1.2 The Role of Mitochondria in the Maturation of Extramitochondrial FeS Proteins Experiments on S. cerevisiae mutant strains as well as T. brucei deﬁcient in mitochondrial FeS cluster assemblies indicated that the mitochondrial pathway is also crucial for the biogenesis of cytosolic and nuclear FeS proteins (Kispal et al. 1999; Smid et al. 2006). Three mitochondrial membrane components were found to be essential for the biogenesis of extramitochondrial FeS proteins: Atm1, an inner mitochondrial membrane transporter (Kispal et al. 1999); Erv1, a sulfhydryl oxidase present in the intermembrane space of mitochondria (Lange et al. 2001); and the tripeptide glutathione, which represents a major free thiol pool (Sipos et al. 2002). As the defect in glutathione synthesis is synergic with Atm1 deﬁciency, it was hypothesized that FeS clusters may be chelated by glutathione and thus stabilized during their transport from mitochondria to the cytosol. However, the nature of the compound exported from mitochondria is not known. In addition to mitochondrial components, four cytosolic factors were characterized that contribute speciﬁcally to the maturation of extramitochondrial FeS proteins in S. cerevisiae. Cfd1 (Roy et al. 2003) and Nbp35 (Hausmann et al. 2005) are essential soluble P-loop NTPases. They form a complex which acts as a scaffold for the formation of transient FeS clusters (Netz et al. 2007). A WD40 repeat protein named Cia1 then facilitates the incorporation of preassembled FeS clusters into their target proteins (Balk et al. 2005a), which

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is dependent on interaction with Nar1 (human Narf), a homologue of [Fe] hydrogenases (Balk et al. 2005b; Horner et al. 2000). While in unicellular eukaryotes the mitochondria were shown to play a central role in cellular FeS cluster assembly, metazoan ISC machinery might not be exclusively localized to mitochondria. It has been found that key ISC components, including IscS, IscU, Nfu, and frataxin, are located in the cytosol and nucleus as well as in mitochondria, and their participation in de novo FeS cluster synthesis outside of mitochondria was suggested (Tong and Rouault 2000, 2006). It would be interesting to investigate at which step of eukaryote evolution the transfer of ISC machinery to other cell compartments occurred and to clarify its extramitochondrial function. 5.1.3 FeS Cluster Assembly in Mitosomes The central role of mitochondria in mediating the biosynthesis of FeS clusters in unicellular eukaryotes is noteworthy. It is this physiological function which may provide a critical selective pressure to retain highly reduced mitochondria over the course of the protist’s adaptation, either to a parasitic lifestyle or to inhabiting oxygen-poor or anaerobic niches. 5.1.3.1 Giardia intestinalis and Other Diplomonads The identiﬁcation of the IscS gene in G. intestinalis provided the ﬁrst evidence for the presence of the ISC machinery in this parasite (Tachezy et al. 2001). A phylogenetic reconstruction revealed that G. intestinalis IscS is related to its mitochondrial homologues (Emelyanov 2003; Tachezy et al. 2001). The absence of a putative targeting signal at the end of the N terminus of giardial IscS, as is typical for all known mitochondrial IscS proteins, led to the suggestion that Giardia is an amitochondrial organism, with secondary loss of the organelle (Tachezy et al. 2001). However, putative N-terminal targeting sequences were later identiﬁed in giardial [2Fe2S] ferredoxin (Nixon et al. 2002) and IscU (Tovar et al. 2003). The cell localization of IscU and IscS by immunoﬂuorescence and immunoelectron microscopy then led to the discovery of mitosomes that both proteins are distributed to. The presence of [2Fe2S] ferredoxin in the same compartment was later demonstrated by expression of the recombinant protein. Gene mining revealed the presence of an IscA homologue and an ongoing proteomic analysis identiﬁed the presence of glutaredoxin 5 in mitosome-enriched cell fractions together with IscS, IscU, and HSP70 (our unpublished results). In summary, the giardial mitosomes contain all key components of the FeS cluster assembly machinery, thus following the general mitochondrial scheme (Table 1). Importantly, the ability of the mitosome-rich fraction to form an FeS cluster

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Table 1 Components of FeS cluster assembly in Saccharomyces cerevisiae, Escherichia coli, and “amitochondriate” eukaryotes; +: high sequence similarity; ?: limited sequence similarity; –: no similarity found. mtHSP70 (Ssq1) proteins cannot be distinguished from other mtHSP70 isoforms (reanalyzed after Lill and Mühlenhoff 2005) S. cerevisiae

E. coli T. vaginalis G. intestinalis E. histolytica E. cuniculi C. parvum

Mitochondrial type of iron–sulfur cluster assembly machinery IscS + Isd11 – IscU1/IscU2 + IscA + [2Fe2S] Ferredoxin+ Ferredoxin + reductase Frataxin + Nfu – Glutaredoxin 5 ? mtHSP70 (Ssq1) + Jac1 + Mge1 +

+ + + + + –

+ – + + + –

NIFS – NIFU – + –

+ ? + – + +

+ – + – + +

+ + – + + +

– – + + + ?

– – – + ? ?

+ – ? + + –

+ – ? + ? +

Export machinery in mitochondrial membrane Atm1 Erv1

– –

– –

– –

– –

? +

? ?

+ + + +

? + + ?

+ + + +

+ + + +

+ + + +

Cytosolic components Nar1 Cfd1 Nbp35 Cia1

– – – –

in apoprotein was demonstrated. However, some of the factors mediating important steps in mitochondrial FeS cluster synthesis were not found in G. intestinalis, including Isd11, frataxin, and ferredoxin-speciﬁc reductase. An intriguing question is identifying the source of ATP and reducing equivalents, which are required for FeS cluster assembly, since no ATP-generating pathway that could supply the ISC assembly machinery with electrons has so far been found in the mitosomes. Whether this is because of our incomplete information on the mitosomal metabolism or whether these compounds are speciﬁcally transported into the organelles from the cytosol remains to be elucidated. In addition to Giardia, it is likely that mitosomes are present in other related diplomonads, such as the ﬁsh parasite Spironucleus. A recent genomic survey of this organism indicated the presence of IscS in this parasite, which might be localized in putative mitosomes (Andersson et al. 2007). However,

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cell localization studies are required to determine which cell compartment the IscS of S. salmonicida is targeted to. Interestingly, homologues of the mitochondrial membrane transport system (Atm1, Erv1) have not been identiﬁed in the giardial genome and the formation of glutathione was not detected in this organism (Brown et al. 1993). However, cytosolic components which are required for maturation of extramitochondrial FeS proteins are likely present (Table 1). Thus, further investigations should be focused on conﬁrming whether the mitosomal FeS cluster assembly is required for the maturation of extramitosmal FeS proteins, and which components of mitosomal membranes are involved in the export from the mitosomes of products of the mitosomal FeS cluster assembly machinery. 5.1.3.2 Microsporidia Analysis of the Encephalitozoon genome sequence revealed eight genes coding for putative mitosomal proteins involved in FeS cluster assembly: the two central components IscS and IscU, the HSP70 chaperone, the iron chaperone frataxin, the electron transporter [2Fe2S] ferredoxin, ferredoxin NADPH oxidoreductase (FNR), the inner membrane transporter Atm1, and Erv1 (Katinka et al. 2001). These genes displayed signiﬁcant similarity to their S. cerevisiae homologues and phylogenetic analysis of ﬁve of them indicated that they are related to α-proteobacterial genes, the bacterial group from which mitochondria are believed to have evolved. Genes coding for the mitochondrial type of HSP70 were found in all microsporidia investigated so far, and immunolocalization experiments on Trachipleistophora hominis showed the presence of HSP70 in mitosomes (Williams et al. 2002). More recently, some components of the FeS cluster assembly machinery of Encephalitozoon (IscU) and another microsporidian Antonospora (ferredoxin, frataxin, FNR) were expressed in S. cerevisiae to demonstrate that these components could be delivered into yeast mitochondria (Burri et al. 2006). All these data support the proposal that microsporidian mitosomes are involved in FeS cluster assembly. However, there are currently no experimental data providing direct evidence of this function. 5.1.3.3 Cryptosporidium parvum Similarly to microsporidia, our knowledge of the FeS cluster assembly in the “amitochondrial” apicomplexan C. parvum is based mainly on the identiﬁcation of genes coding for components of the FeS cluster assembly machinery in the parasite’s genome and reconstruction of the hypothetical pathway in the mitosomes (Table 1). The presence of cryptosporidian IscS and IscU was

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not directly localized within the mitosome, although both proteins contain N-terminal targeting signals efﬁcient for their translocation into yeast mitochondria (LaGier et al. 2003). Several other components, such as frataxin (unpublished data), [2Fe2S] ferredoxin (LaGier et al. 2003), and Narf (Stejskal et al. 2003), were shown to be expressed in C. parvum. 5.1.3.4 Entamoebids Entamoebids are so far the only eukaryotic lineage in which homologues of ISC machinery have not been found. Instead of an ISC system, these organisms including E. histolytica possess components of a nonredundant NIF system. Two components, NifS and NifU, were shown to be expressed and active in this parasite (Ali et al. 2004). In addition to E. histolytica, homologues of NIF components are present in the genomes of other species of the Entamoeba genus, such as the reptile parasite E. invadens and freeliving E. moshkovskii (http://www.sanger.ac.uk), and most likely in the related free-living protist Mastigamoeba (our unpublished data). A phylogenetic reconstruction of E. histolytica NifS and NifU indicated that both genes were acquired by horizontal gene transfer from the group of non-diazotrophic ε-proteobacteria, with the closest relationship to the Campylobacter and Helicobacter species (van der Giezen et al. 2004). When expressed in a strain of Escherichia coli with deletions of both the ISC and SUF systems, NifU and NifS of E. histolytica complement the growth of the mutant bacteria (Ali et al. 2004). This result suggests that NIF components can catalyze general FeS cluster assembly in E. histolytica, as was shown for some non-diazotrophic bacteria (Olson et al. 2000). Importantly, the analysis of entamoebid NifS and NifU did not reveal the presence of a putative mitochondria-like targeting signal that would predict their localization within the mitosomes, and their cytosolic localization was suggested (van der Giezen et al. 2004). If so, the entamoebid mitosomes are the only mitochondria-related organelles that lost the ability to synthesize FeS clusters and their function is currently unknown. 5.2 Requirements for ATP, Membrane Potential, and Electron Transport In mitosome-harboring organisms, the generation of ATP is exclusively dependent on cytosolic energy metabolism. Mitosomes are unable to synthesize ATP by either oxidative or substrate level phosphorylation. However, ATP hydrolysis is required for the functioning of mtHSP70 and CPN60, which are present in mitochondria as well as all related organelles, including mitosomes. In mitochondria, they are involved in protein import, protein maturation, and the formation of FeS clusters. It is likely that similar demands for ATP are required for corresponding functions in mitosomes. If so, ATP

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has to be imported from the cytosol to the organelles to satisfy mitosomal needs. Indeed, genes for ATP/ADP carriers have been identiﬁed in C. parvum and E. cuniculi (Abrahamsen et al. 2004; Katinka et al. 2001), though not in G. intestinalis. A unique ADP/ATP carrier has been found and characterized in some detail in E. histolytica (Chan et al. 2005). Unlike mitochondrial carriers, the Entamoeba carrier is not sensitive to the classic inhibitors carboxyatractyloside and bongkrekic acid, and also exhibits differences in its substrate preferences. In addition to ATP and ADP, which are transported by mitochondrial carriers, the Entamoeba carrier also transports AMP. Mitochondrial protein import is dependent on the membrane potential, which is mainly generated by cytochrome-dependent electron transport. In mitosome-carrying protists, however, neither cytochromes nor other components of the membrane-associated respiratory chain have been found and it is not clear whether, and if so how, the mitosomal membrane potential is generated. The only candidate that might be involved in mitosomal transmembrane proton translocation is pyridine nucleotide transhydrogenase (PNT). This proton pump, typically present in the inner mitochondrial membrane, catalyzes direct transfer of the hydride ion between NAD(H) and NADP(H) providing intramitochondrial NADPH (Hateﬁ and Yamaguchi 1996). However, in helminths such as Hymenolepis that enter the anaerobic phase of their life cycle, PNT is the principal proton pump generating a membrane potential. It operates in the reverse direction, translocating protons from the matrix to the intramembrane mitochondrial space (Mercer et al. 1999). The genes coding for PNT have been found in the genomes of E. histolytica and C. parvum (Putignani et al. 2004; Yu and Samuelson 1994). Catalytic activity was demonstrated in the former organism, in which the activity was associated with a sedimentable cellular fraction (Harlow et al. 1976). Together with the observation that the amino terminus of E. histolytica PNT possesses an extension similar to targeting presequences, these ﬁndings suggest that PNT may operate in mitosomes. However, mitosomal localization and the functionality of PNT remain to be elucidated in both E. histolytica and C. parvum. In Giardia, the presence of membrane-bound structures displaying membrane potential and electron transport was tested using the cationic dye rhodamine 123 and 5-cyano-2,3-ditolyl tetrazolium chloride (CTC), respectively (Lloyd et al. 2002). Rhodamine 123 was accumulated in spherical and oval structures adjacent to the inner face of the giardial plasma membrane. CTC is an artiﬁcial electron acceptor, which reduces to ﬂuorescent insoluble formazan. The formation of formazan was observed in several membrane structures under the giardial plasma membrane. However, the CTC-labeled structures were visibly distinct from those labeled with rhodamine 123. Although both types of labeling were considered evidence of the presence of the membrane system with mitochondrion-like functions in Giardia, the speciﬁcity of the labeling in Giardia and the character of the observed structures

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needs to be validated. First, it is clear that the rhodamine 123-labeled structures are distinct from mitosomes: they are considerably larger than mitosomes (1.3–2 µm) and their number (4–5 per cell) and cellular distribution are also different. The speciﬁcity of CTC labeling is questionable, as the reduction of tetrazolium salts is mediated by a number of nonmitochondrial enzymes and electron carriers and formazan might be also unspeciﬁcally formed by superoxide radicals (Bernas and Dobrucki 2000; Breeuwer and Abee 2000). Moreover, neither rhodamine 123 nor CTC labeled the mitosomal rodlike structure, which is invariably observed between Giardia nuclei (Doleˇzal et al. 2005; Regoes et al. 2005; Tovar et al. 2003). Thus, colocalization studies with speciﬁc markers are required to clarify the relationship between rhodamine 123- and CTC-labeled structures and mitosomes. Indirect evidence of some electron transport in mitosomes indicates the prescence of [2Fe2S] ferredoxin. This electron carrier is typically present in mitochondria and hydrogenosomes, providing electrons for various pathways including FeS cluster formation. Indeed, [2Fe2S] ferredoxin colocalizes with IscS and IscU in Giardia mitosomes. This type of ferredoxin was also found in the genomes of Encephalitozoon cuniculi and C. parvum, and its mitosomal localization was suggested. However, the source of reducing equivalents which could be transported by the ferredoxins within mitosomes is unknown. Interestingly, [2Fe2S] ferredoxin is absent in Entamoeba histolytica, which is consistent with the replacement of ISC by the NIF type of FeS cluster assembly machinery in this parasite. In the NIF machinery, the function of [2Fe2S] ferredoxin is likely substituted by a ferredoxin-like module of NifU (Agar et al. 2000). 5.3 Other Putative Functions More information about the functions of mitosomes was expected to be obtained from an analysis of the complete genomes of mitosome-bearing protists. However, very few genes coding for putative mitosomal proteins have been uncovered so far. In E. cuniculi, genes coding for the alpha and beta subunits of a putative E1 component of a mitochondrial pyruvate dehydrogenase complex have been identiﬁed (Katinka et al. 2001). The E1 component is the pyruvate dehydrogenase itself that catalyzes oxidative decarboxylation of pyruvate. In mitochondria, two other components of the PDH complex, E2 and E3, are required for the subsequent production of acetyl coenzyme A. These components are absent in E. cuniculi. In addition, glycerol-3-phosphate dehydrogenase, an enzyme which is involved in electron transport from the cytosol to mitochondria, and manganese-containing superoxide dismutase, which forms part of the system that protects against oxidative stress, were suggested to be localized in the mitosomes of E. cuniculi and A. locustae (Burri et al. 2006).

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The genome of C. parvum revealed the presence of genes coding for an alternative oxidase, which transfers electrons to molecular oxygen. This enzyme, together with thioredoxin reductase, glutathione peroxidase, and iron superoxide dismutase, might be involved in the scavenging of toxic oxygen species (Putignani et al. 2004). Interestingly, this parasite also possesses a gene coding for the F1β subunit of ATP-synthase (mitochondrial complex V). Further experimental work is required to elucidate the physiological functions and metabolic context of these currently isolated mitosomal proteins, which are, in mitochondria, components of various complex metabolic pathways.

6 Perspectives Although the ﬁrst mitosomes were discovered in 1999, we are still at the beginning of the process of understanding their physiological functions and importance to their cells. The most exciting question is what the essential function is that forces various organisms to retain these organelles over the course of evolution. Such an essential function might not be identical in all mitosomes, as they evolved in organisms distributed over a large spectrum of distinct taxonomic groups living under different environmental conditions. Indeed, while the formation of FeS clusters might be the essential function, common to mitosomes in Giardia, microsporidians, and Cryptosporidium, no evidence for mitosomal FeS cluster assembly was observed in E. histolytica. It was speculated that the compartmentalization of this function might be essential to avoid the toxicity of ferrous iron and sulﬁde, which are needed for FeS cluster assembly (Tachezy and Dolezal 2007). Moreover, although anaerobic protists harboring mitosomes or hydrogenosomes have considerably higher nutritional demands for iron than aerobic cells, they do not possess ferritin, a key iron storage protein in bacteria and metazoa. Thus, mitosomes may also serve as FeS clusters and/or Fe storage organelles in these organisms. It is still not clear, however, what characters of compounds are exported from mitosomes, as well as from hydrogenosomes and mitochondria to the cytosol for the maturation of FeS proteins in other cellular compartments. A different physiological function for mitosomes might be expected in E. histolytica, which has completely lost the components of the mitochondrial-type ISC assembly machinery and assembles FeS clusters using NIF machinery. If the NIF system operates in the entamoebid cytosol, which needs to be unequivocally proved, it is not clear how the cell avoids the toxicity of the intermediate compounds. Many more functional studies are required to clarify these questions and to decipher mitosomal functions that are still unknown. In this respect, large-scale comparative proteomic analyses of highly puriﬁed organelles are of particular interest.

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Investigations on mitosomes are highly attractive in terms of gaining new information relevant to understanding the evolutionary paths leading to the formation of various mitochondria-derived organelles, of which mitosomes are the most simpliﬁed. Moreover, the biochemical simplicity of mitosomes nominate these organelles to be a unique model for the detailed study of identiﬁed pathways such as FeS cluster assembly, which are, in mitochondria, operating in a complex biochemical context. A particularly attractive subject of current investigations is the mitosomal protein import system, which seems to be highly divergent and simpliﬁed compared to that known in mitochondria. Finally, further comparative studies of mitosomes and other mitochondrion-derived organelles may contribute to our knowledge of how parasitic protists adapted to the host environment, whether the parasitic lifestyle drove the changes leading to the organelles’ simplicity or whether preadaptation of free-living protists to an oxygen-poor environment provided them with an advantage for subsequent invasion of potential hosts. Acknowledgements We thank our colleagues who contributed to the research on mitosomes at Charles University in Prague: Pavel Doleˇzal, Robert ˇ Sut’ák, Pavla T˚ umová, Petr Rada, Zuzana Zubáˇcová, Ivan Hrd´ y, and Jaroslav Kulda. We also thank Michaela Marcinˇciková for excellent technical help. Research in the J.T. laboratory is supported by the Ministry of Education, Youth, and Sports of the Czech Republic MSM0021620858 and LC07032, Czech Science Foundation (206/06/0947), and Grant Agency of the Academy of Sciences of the Czech Republic (IAA501110631).

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van der Giezen M, Cox S, Tovar J (2004) The iron–sulfur cluster assembly genes iscS and iscU of Entamoeba histolytica were acquired by horizontal gene transfer. BMC Evol Biol 4:7 van Dooren GG, Marti M, Tonkin CJ, Stimmler LM, Cowman AF, McFadden GI (2005) Development of the endoplasmic reticulum, mitochondrion and apicoplast during the asexual life cycle of Plasmodium falciparum. Mol Microbiol 57:405–419 van Dooren GG, Stimmler LM, McFadden GI (2006) Metabolic maps and functions of the Plasmodium mitochondrion. FEMS Microbiol Rev 30:596–630 Vávra J (1976) Structure of the microsporidia. In: Bulla LA Jr, Cheng TS (eds) Comparative pathobiology, vol 1. Biology of the Microsporidia. Plenum, New York, pp 1–86 Vávra J (2005) Polar vesicles of microsporidia are mitochondrial remnants (mitosomes)? Folia Parasitol 52:193–195 Vickerman K (1985) Development cycles and biology of pathogenic trypanosomes. Br Med Bull 41:105–114 Wiedemann N, Frazier AE, Pfanner N (2004) The protein import machinery of mitochondria. J Biol Chem 279:14473–14476 Williams BAP, Hirt RP, Lucocq JM, Embley TM (2002) A mitochondrial remnant in the microsporidian Trachipleistophora hominis. Nature 418:865–869 Wilson RJ, Williamson DH (1997) Extrachromosomal DNA in the Apicomplexa. Microbiol Mol Biol Rev 61:1–16 Wilson RJM, Rangachari K, Saldanha JW, Rickman L, Buxton RS, Eccleston JF (2003) Parasite plastids: maintenance and functions. Philos Trans R Soc Lond B Biol Sci 358:155–162 Wu SP, Wu G, Surerus KK, Cowan JA (2002) Iron–sulfur cluster biosynthesis. Kinetic analysis of [2Fe-2S] cluster transfer from holo ISU to apo Fd: role of redox chemistry and a conserved aspartate. Biochemistry 41:8876–8885 Yaffe MP, Stuurman N, Vale RD (2003) Mitochondrial positioning in ﬁssion yeast is driven by association with dynamic microtubules and mitotic spindle poles. Proc Natl Acad Sci USA 100:11424–11428 Yarlett N, Hackstein JHP (2005) Hydrogenosomes: one organelle, multiple origins. Bioscience 55:657–667 Youssef NN, Hammond D (1971) The ﬁne structure of the developmental stages of the microsporidian Nosema apis Zander. Tissue Cell 3:283–294 Yu Y, Samuelson J (1994) Primary structure of an Entamoeba histolytica nicotinamide nucleotide transhydrogenase. Mol Biochem Parasitol 68:323–328 Zheng L, Cash VL, Flint DH, Dean DR (1998) Assembly of iron–sulfur clusters. Identiﬁcation of an iscSUA-hscBA-fdx gene cluster from Azotobacter vinelandii. J Biol Chem 273:13264–13272

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_115/Published online: 8 December 2007 © Springer-Verlag Berlin Heidelberg 2007

The Mitochondrion-Related Organelle of Cryptosporidium parvum Janet S. Keithly The Wadsworth Center, New York State Department of Health, Division of Infectious Diseases, David Axelrod Institute for Public Health, 120 New Scotland Avenue, Albany, NY 12208, USA [email protected] 1

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Abstract The mitochondrion-related organelle of Cryptosporidium parvum is structurally distinguished from the hydrogenosomes and mitosomes of anaerobic protists by its (1) close association with the crystalloid body, an organelle unique to this apicomplexan and the function of which is currently unknown; (2) close association with the outer nuclear membrane and possibly nuclear pores; (3) envelopment by rough endoplasmic reticulum and in some cases an apparent direct tethering to ribosomes; and (4) atypical internal membranous compartments that lack well-deﬁned crista junctions with the mitochondrial inner membrane, a characteristic that deﬁnes most aerobic eukaryotic mitochondria. Like most hydrogenosome- and mitosome-bearing anaerobic protists, however, C. parvum lacks a mitochondrial genome, i.e. proteins are encoded by the nucleus and targeted back to the mitochondrion-like organelle. As a consequence of this reductive evolution, there are no genes for electron transport or oxidative phosphorylation, and the only function so far ascribed to this tiny organelle is one common to all

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eukaryotic mitochondria, the assembly and maturation of iron sulfur clusters. The ultrastructure and tomography of the relic mitochondrion and crystalloid body, as well as their probable functions, are the primary topics herein. An overview of iron sulfur cluster biosynthesis, the likely mechanisms for import into and export from the mitochondrion, as well as core carbohydrate and energy metabolism are discussed. Similarities and differences in the structure and function of both organelles with anaerobic protists in general, as well as with other apicomplexans speciﬁcally, are described.

1 Introduction Cryptosporidium parvum belongs to the diverse group of intracellular apicomplexans that includes species of human (Babesia microti, Plasmodium falciparum, Toxoplasma gondii) and veterinary (Eimeria tenella, Neospora caninum) importance (Ellis et al. 1998; Fayer 1997; Thompson et al. 2005). While a single mitochondrion is present in most Apicomplexa (McFadden 2003), its existence in C. parvum (Aji et al. 1991; Beyer et al. 2000; Fayer 1997; Riordan et al. 1999; Tetley et al. 1998), and in other species of the genus (Alvarez-Pellitero 2004; Uni et al. 1987), had remained an open question. The evidence for a mitochondrion-like organelle in C. parvum sporozoites was ﬁrst conﬁrmed by ultrastructural and phylogenetic analyses (Riordan et al. 1999). These initial data showed that cells of C. parvum possessed a double membrane-bounded single organelle enveloped by rough endoplasmic reticulum (RER) to which C. parvum Cpn60 (CpCpn60) (Riordan et al. 2003) and mitochondrial (mt) Hsp70 (CpmtHsp70) were immunolocalized (Riordan et al. 2003; Slapeta and Keithly 2004). This organelle was also in intimate contact with the nucleus and sandwiched between it and the crystalloid body (CB), a cryptosporidium-speciﬁc organelle of unknown function. Mitochondrionlike structures had already been seen in C. parvum (Beyer et al. 2000) and C. muris (Uni et al. 1987) from infected rats and mice, respectively. Initially, the ultrastructure of the organelle in sporozoites of C. parvum observed posterior to the nucleus was thought to more closely resemble the apicoplast, a chloroplast remnant observed in most apicomplexans (Eimeria, Neospora, Plasmodium, Toxoplasma), rather than a mitochondrion (Tetley et al. 1998). Upon further investigation, this supposition could not be sustained. Both transmission electron microscopy (TEM) and electron tomography showed that the organelle in C. parvum was bounded by two membranes (Keithly et al. 2005; Putignani et al. 2004; Riordan et al. 2003; Slapeta and Keithly 2004), rather than the 3 to 4 observed in apicoplasts (reviewed in McFadden 2003; Williams and Keeling 2003). In addition, tomography indicated that the inner membrane (IM) of the C. parvum organelle formed membranous subcompartments (Keithly et al. 2005) and were therefore quite distinct from the infoldings of plastid inner membranes (McFadden 2003). Indeed, although there did not appear to be “crista junctions” with the IM of

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the organelle, the internal morphology of the mitochondrion-like organelle was quite similar to that observed for yeast and mammalian mitochondria in a highly “condensed” state when the matrix is contracted (Mannella et al. 2001; reviewed in Frey et al. 2006 and Medalia et al. 2005). Finally, subsequent ultrastructural studies have indicated that C. parvum lacks an apicoplast (Keithly et al. 2005; Putignani et al. 2004; Riordan et al. 1999, 2003; Slapeta and Keithly 2004), and that both human species of Cryptosporidium lack an apicoplast genome (Abrahamsen et al. 2004; Cai et al. 2003; Xu et al. 2004; Zhu et al. 2000). Taken together, these data indicated that the double-membrane bounded organelle in C. parvum was not an apicoplast, but rather, an unusually compacted organelle of reduced size that most closely resembled the mitochondria (Krungkrai et al. 2000; Melo et al. 2000), mitosomes (Clark and Roger 1995; Leon-Avila and Tovar 2004; Tovar et al. 1999; reviewed in Tovar 2006), and hydrogenosomes (reviewed in Barbera et al. 2006; McFadden 2003; Müller 1973, 1975; van der Giezen et al. 2005; van Hoek et al. 2000) of other anaerobic parasitic protists, in essence, a relic mitochondrion.

2 Ultrastructural Morphology Cryptosporidium parvum is distinguished from mitosome- and hydrogenosome-bearing protists by being an intracellular parasite that contains a mitochondrion-like organelle that is drastically reduced in size and structure, and that lacks a mitochondrial genome. Plasmodium and Toxoplasma, its nearest apicomplexan relatives, are also intracellular parasites with mitochondria reduced in structure and function. Unlike C. parvum, however, the mitochondria of P. falciparum and T. gondii still possess small mitochondrial genomes which permit them to express during certain parts of the life cycle cytochrome oxidase b and subunits I and III of the cytochrome c oxidase, the universal mitochondrial genes retained in the mitochondrial genome of all respiring eukaryotes (Feagin and Drew 1995; Kita et al. 2002; Mi-Ichi et al. 2003). In addition, the nuclear genome of the sexual stages of the malaria parasite is also able to transfer reducing equivalents to oxygen using the classical respiratory pathways, and to fumarate using a complex II quinol-fumarate reductase (Kita et al. 2002). It is thought that these variations in mitochondrial structure and function amongst the Apicomplexa have occurred in response to differences in the host cell environment. 2.1 Local Environment Initial TEM and tomographic reconstruction of chemically ﬁxed sporozoites of C. parvum have revealed a complex arrangement of membranes both

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around this organelle and within it (Figs. 1, 2). Speciﬁcally, the relic mitochondrion is enveloped by multiple segments of RER that appear to extend from the outer nuclear envelope (Fig. 1b,c; see Figs. 4–9, Keithly et al. 2005). In tomographic reconstructions of ﬁxed sporozoites, a single, highly folded inner membrane (Fig. 2), or multiple internal subcompartments (see Figs. 12–15, Keithly et al. 2005), were visualized within the organelle. As mentioned before, the infoldings of the IM appeared to lack tubular crista

Fig. 1 Transmission electron microscopy of Cryptosporidium parvum sporozoite organelles (reprinted from Figs. 1–3 of Keithly et al. 2005 with permission of the publishers). a Longitudinal section of sporozoite at low magniﬁcation. The mitochondrion-like organelle (∗ ) is between the nucleus (N) and the crystalloid body (CB). A typical apical complex is evident (X), and the Golgi (G) is anterior to the nucleus. Shown in detail in the inset. Other organelles include the micronemes (M), single rhoptry (R), and dense granules (D) for host cell invasion, and amylopectin (A), a plant-like storage granule (bar = 0.5 µm). b Higher magniﬁcation showing the intimate relationship between the nucleus, its membranes (arrows), the CB, and the mitochondrial remnant (∗ ). The mitochondrion-like organelle is spherical, 150–300 nm in diameter, contains internal crista-like membranes (C), and is enveloped by rough endoplasmic reticulum (RER). The vesicular CB is closely apposed to the mitochondrion (bar = 0.1 µm). c A higher magniﬁcation of the nucleus, its membranes, the CB, and the relic mitochondrion. A larger arrow denotes the lumen of the nuclear envelope, the outer membrane of which is studded with ribosomes (black/white arrow). The mitochondrion-related organelle is surrounded by two limiting membranes (small arrow). Scale bar = 0.2 µm

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Fig. 2 Electron tomographic reconstruction of a Cryptosporidium parvum mitochondrionlike organelle (reprinted from Figs. 16–19 of Keithly et al. 2005 with permission of the publishers). a Projection image of the untilted specimen showing the nucleus (N), crystalloid body (CB), mitochondrion-like organelle (∗ ), and the colloidal gold particles used for alignment. Original section thickness approximately 140 nm (scale bars = 0.1 µm). b Slice (3.2 nm thick) from the center of the tomogram, in the plane normal to the electron beam showing the mitochondrion-like organelle surrounded by the prominent RER. c Identical to b, but contours have been added to show the outer (red) and inner membrane proﬁles (green). d Four views of a three-dimensional model of the membrane surfaces in the reconstructed mitochondrion-like organelle showing the outer (red) and one large inner membrane compartment (green). The model represents a volume approximately 130 nm thick at the center of the reconstruction

junctions typical for metazoan, fungal, and protist mitochondria. These observations for the internal membranes of the C. parvum relic mitochondrion are consistent with the loss, through reductive evolution, of the normal oxidative phosphorylation machinery in C. parvum, and the loss of its mitochondrial genome (Abrahamsen et al. 2004; Keeling 2004). 2.2 Internal Organization The organization of the IM of the C. parvum mitochondrion-related organelle is quite distinct from other amitochondriate protists. With the exception of

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the anaerobic free-living ciliates e.g. Metopus and Plagiopyla (Fenchel et al. 1977), the parasitic ciliate Nyctotherus ovalis (Boxma et al. 2005) and the stramenopile (syn. heterokont) Blastocystis hominis (Nasirudeen and Tan 2004; Tan and Suresh 2006; Yarlett, 2007, in this volume), all hydrogenosome- and mitosome-bearing anaerobic protists and fungi lack mitochondrial cristae. These include the parabasalids Trichomonas vaginalis and Tritrichomonas foetus (reviewed in Babera et al. 2006); archaeamoebid Entamoeba histolytica (Clark and Roger 1995; Tovar et al. 1999; reviewed in Tovar 2006); diplomonads Giardia intestinalis and Spironucleus barkhanus (Tovar et al. 2003; Simpson and Roger 2004); chytrid and microsporidian fungi Neocallimastix and Piromyces species (reviewed in Williams and Keeling 2003), Encephalitozoon cuniculi (Katinka et al. 2001) and Trachipleistophora hominis (Williams et al. 2002) (reviewed in Barbera et al. 2006, Keeling and Fast 2002). The infoldings of the mitochondrial IM also differ from those of most Apicomplexa, in which the type and number of cristae vary considerably during the life cycle, ranging from tubular in T. gondii tachyzoites (Melo et al. 2000) and P. falciparum gametocytes, to “acristate” in asexual P. falciparum and P. knowlesi (Fry and Beesley 1991; Krungkrai et al. 1999; Krungkrai et al. 2000). Ultrastructural analyses of the C. parvum mitochondrion-derived organelle indicate that although it has an outer and an inner boundary membrane, the presence of membranous subcompartments sets it apart from these near relatives (Riordan et al. 1999, 2003). Indeed, tomographic reconstructions of ﬁxed sporozoites suggest that junctions which typically connect cristae to the mitochondrial IM (Frey and Mannella 2000; Mannella et al. 2001) are absent (or greatly reduced) in the remnant mitochondrion of C. parvum. 2.2.1 Variations in Cristae It is reasonably well established that the morphology of cristae can vary as a result of osmotic adjustments in the matrix volume (Frey et al. 2006; Lucic et al. 2005; Mannella et al. 2001). Most mammalian mitochondria observed in situ are in an “orthodox” conformation, with a large matrix volume that pushes the inner boundary membrane against the outer membrane leaving an ∼8 nm space between them. Nevertheless, under conditions of stress or loss of genetic function, the IM of mammalian and yeast mitochondria becomes condensed (matrix contracted) (Perkins et al. 1998; reviewed in Frey et al. 2006). In this state, the number of cristae are reduced and their size expands. This is congruent with tomographic reconstructions in chemically ﬁxed cells of the internal membrane organization of the C. parvum relic mitochondrion, i.e. subcompartments are few and expanded (Fig. 2d). Recent observations from cryo-electron tomography of C. parvum sporozoites visualized in their “native state” following rapid freezing and freeze substitution

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further support this view (C. Hsieh, M. Marko, J.S. Keithly, C.A. Mannella, unpublished results). Both of these observations in ﬁxed and native cells on the lack of typical internal compartmentation in the C. parvum relic mitochondrion are consistent with a structural reﬂection of the secondary loss of oxidative phosphorylation during the reductive evolution of this organelle (Keeling 2004; Williams and Keeling 2004).

3 Cell Biology 3.1 Iron Sulfur [FeS] Cluster Biosynthesis As in the anaerobic protists Giardia and Trichomonas (Suchan et al. 2003; Sutak et al. 2004; Tielens and van Hellemond 2006; Tovar et al. 2003), and Microsporidia (Katinka et al. 2001; Keeling and Fast 2002; Williams et al. 2002), it was suggested that the relic mitochondrion of C. parvum has been retained for the assembly of FeS clusters (LaGier et al. 2003; reviewed in Tachezy and Dolezal 2007). In fact, proteins of the ISC system are present in the genomes of both C. parvum and C. hominis (Abrahamsen et al. 2004; Xu et al. 2004), and all of them possess mitochondrial targeting presequences (Dolezal et al. 2006; LaGier et al. 2003; reviewed in Tachezy and Dolezal 2007). These include frataxin, the candidate iron donor for FeS assembly (Dolezal et al. 2007), IscS, a pyridoxal phosphate-dependent cysteine desulfurase for the production of inorganic sulfur, IscU which is the scaffold for transient FeS cluster formation, a [2Fe2S] ferredoxin, as well as a Nar/Narf (nuclear prelaminin A recognition factor) hydrogenase-like protein (Stejskal et al. 2003) for accepting electrons, and an Nbp35 P-loop NTPase, for the maturation of cytosolic and nuclear FeS proteins (Hausmann et al. 2005; reviewed in Tachezy and Dolezal 2007). Phylogenetic analyses of mtHsp70, Cpn60, and all of the Cryptosporidium assembly sequences cluster together with homologs from mitochondrionbearing eukaryotes, including G. intestinalis, T. vaginalis, and other Apicomplexa (reviewed in Barbera et al. 2006; LaGier et al. 2003; Riordan et al. 2003; Slapeta and Keithly 2004). Furthermore, the presequences of CpCpn60, as well as both CpIscS and CpIscU, two genes associated with mitochondrial FeS cluster biogenesis, can target green ﬂuorescent protein (GFP) to the mitochondrial network of the yeast Saccharomyces cerevisiae (LaGier et al. 2003). In addition, the presequence of C. parvum mtHsp70 can target GFP not only to yeast mitochondria, but also to the single mitochondrion of the apicomplexan Toxoplasma gondii (Slapeta and Keithly 2004). Fluorescent- and immunogold-labeled polyclonal antibodies to CpmtHsp70 localize within the mitochondrion-related organelle of both C. parvum free

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sporozoites and intracellular meronts (Slapeta and Keithly 2004), targeting the same organellar compartment where antibodies to CpCpn60 are localized (Riordan et al. 2003). Taken together, these data strongly suggest that, like other mitochondrion-bearing eukaryotes, the C. parvum relic organelle performs FeS cluster biosynthesis. 3.2 Protein Import and Export The Cryptosporidium mitochondrion-derived organelle also appears to possess a fairly complete mechanism for protein import and export. As mentioned previously, both Cpn60 and Hsp70 are targeted into the organelle where they ensure proper protein folding (LaGier et al. 2003; Riordan et al. 2003; Slapeta and Keithly 2004). In addition, the genomes of C. hominis and C. parvum include components of the Translocase Outer Membrane (TOM) and Inner Membrane (TIM) import systems (Abrahamsen et al. 2004; Xu et al. 2004), as well as both the α and β subunits of a mt Matrix Processing Peptidase for removing the targeting signal after entry across the inner membrane (Abrahamsen et al. 2004; Xu et al. 2004; reviewed in Tachezy and Dolezal 2007). All eukaryotes examined so far contain lineage-speciﬁc TOM, TIM22, and Sorting and Assembly Machinery (SAM) complexes (Gentle et al. 2007; Dolezal et al. 2006). Among the Apicomplexa, this includes several species of Cryptosporidium, Plasmodium and Theileria. The genomes of these three apicomplexans have been recently found to possess “Tiny TIMS” which collect substrate proteins from the TOM complex for delivery either to the TIM22 complex on the inner mitochondrial membrane, or to the SAM complex in the outer membrane (Gentle et al. 2007). Both species of Cryptosporidium encode a single, hybrid complex of Tim8 and Tim13 that is known to preferentially bind the last transmembrane segment of Tim22. In addition, both ATM1 and Erv1, which are required for the maturation of extra-mitochondrial (cytosolic) proteins and that are part of the mt export machinery, have been identiﬁed in the C. parvum genome (reviewed in Barbera et al. 2006). The former is an IM protein of the ABC transporter family that faces the mt matrix (Decottignies and Goffeau 1997), and the second is an intermembrane sulhydryl oxidase whose function in cytosolic FeS protein biosynthesis is still unknown (reviewed in Barbera et al. 2006). When combined with glutathione, which may help stabilize proteins being exported from the mitochondrion into the cytosol (reviewed in Tachezy and Dolezal 2007), these components facilitate export of proteins from eukaryotic mitochondria. Two cytosolic factors identiﬁed in the C. parvum genome are also thought to help cytosolic and nuclear FeS proteins mature (Abrahamsen et al. 2004; Templeton et al. 2004). Nucleotide binding protein of 35 kDa (Nbp35) is an essential soluble P-loop ATPase (Hausmann et al. 2005), that together with a Narf -like protein (Stejskal et al. 2003) may assist either in transferring FeS

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clusters to apoproteins after they exit the relic mitochondrion, and/or play a role in the de novo synthesis of cytosolic FeS proteins by using some FeS clusters assembled within the relic mitochondrion (Dong et al. 2004; Lill and Mühlenhoff 2005; reviewed in Tachezy and Dolezal 2007). Together this evidence suggests that C. parvum has most of the necessary machinery to import proteins into, and export them from, the relic mitochondrion. 3.3 Association of the Mitochondrion with the RER The origin of all mitochondrion-derived organelles is generally accepted to be the endosymbiosis of an α-proteobacterium (reviewed in Barbera et al. 2006; Brown and Doolittle 1997; Gray and Lang 2001; Rotte et al. 2000; van der Giezen et al. 2005), whereas the nucleus, endoplasmic reticulum, lysosomes and apical organelles of the apicomplexan endomembrane system are thought to be derived from the infoldings of the ancestral plasma membrane (Brown and Doolittle 1997; Cavalier-Smith 1991). As previously mentioned, TEM conﬁrmed the presence of a roughly spherical 150–300 nm organelle with two closely apposed boundary membranes, the outer membrane of which is closely associated with the RER, and which appears to originate from the outer envelope of the nearby nucleus (Fig. 1). In fact, the relic mitochondrion is almost completely enveloped by protrusions of the RER (Fig. 1c), an association that was dramatically visualized by tomographic reconstruction (see Fig. 10, Keithly et al. 2005). These initial observations have recently been conﬁrmed by cryo-electron tomography (Csordas et al. 2006; reviewed in Frey et al. 2006; Lucic et al. 2005; Medalia et al. 2002) of C. parvum sporozoites in their “native state” which not only shows envelopment by the RER, but the possible intimate association of the RER with nuclear pores (C. Hsieh, M. Marko, J.S. Keithly, C.A. Mannella, unpublished results). This is an important observation in view of the recent discovery of a novel and essential FeS protein found in the cytosol of all eukaryotes, RNase L inhibitor 1 (Rli1), which is required for the maturation and export of ribosome subunits from the nucleus, as well as for the efﬁcient initiation of translation (Dong et al. 2004; Kispal et al. 2005; Lill et al. 2005). Indeed, a homolog of this protein has been found in both species of Cryptosporidium, as well as in all hydrogenosome- and mitosome-bearing protists (reviewed in Tachezy and Dolezal 2007). Because Rli1 homologs have been found only in archaebacteria (reviewed in Tachezy and Dolezal 2007), it is hypothesized that Rli1 and other components of cytosolic translation machinery are derived from them. Therefore, we propose that the intimate association of the relic mitochondrion, of eubacterial origin, with its associated membranes is a structural reﬂection of the cell’s attempt to facilitate efﬁcient ribosome biogenesis and translation initiation following reductive evolution of the organelle (Keeling 2004;

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Williams and Keeling 2002). We also suggest that this close association of the mitochondrion-like organelle and endomembrane system facilitates the assembly of cytosolic Rli1, the only known essential FeS protein for yeast viability (Lill et al. 2005; reviewed in Tachezy and Dolezal 2007). These data are consistent with previous hypotheses that the primary function of the mitochondrion-like organelle in C. parvum is the assembly of FeS clusters in order to provide mature FeS proteins to all cellular compartments, including the cytosol, mitochondrion, and nucleus.

4 Carbohydrate Metabolism Although the carbohydrate metabolism of C. parvum is only partially understood, this apicomplexan primarily relies upon glycolysis for energy production (Abrahamsen et al. 2004; Entrala and Mascaro 1997). In fact, its core metabolism differs signiﬁcantly from its near relatives Plasmodium and Toxoplasma (reviewed in Crawford et al. 2003), resembling instead that of the microsporidian Encephalitozoon cuniculi (Katinka et al. 2001; Keeling and Fast 2002), and the microaerophilic protists Giardia intestinalis, and Entamoeba histolytica (reviewed in Barbera et al. 2006 and Müller 2003). All of these microorganisms lack both a functional TCA cycle and oxidative phosphorylation. 4.1 Pyruvate : NADP+ Oxidoreductase (PNO) In C. parvum, pyruvate is converted to acetyl-CoA by a novel pyruvate : NADP+ oxidoreductase (CpPNO) (Rotte et al. 2001) containing two distinct domains that are fused together: an N-terminal pyruvate:ferredoxin oxidoreductase (PFO), similar to that found in the anaerobic protists Entamoeba histolytica, Giardia intestinalis, Trichomonas vaginalis, and a C-terminal NADPHcytochrome P450 reductase (CPR). A nearly identical PFO-CPR fused protein has also been described from the euglenozoan Euglena gracilis (Inui et al. 1987; Nakazawa et al. 2000; Rotte et al. 2001), a distant relative of the Alveolata to which all apicomplexans belong. More recently, a similar fusion has been identiﬁed within the genome of the stramenopile Thalassiosira pseudonana, a marine centric diatom (cited in Ctrnacta et al. 2006). Unlike Euglena PNO, which is a mitochondrial protein (Rotte et al. 2001), CpPNO lacks a mitochondrial targeting presequence and does not localize within the relic organelle (Ctrnacta et al. 2006). In fact, sporozoites of C. parvum visualized both by confocal immunoﬂuorescence (Fig. 3) and immunogold-labelled TEM (Fig. 4) conﬁrm that CpPNO has an unique compartmentalization: ﬁrstly within the cytosol as expected, but secondly within

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Fig. 3 Confocal microscopic immunolocalization of pyruvate : NADP+ oxidoreductase (CpPNO) in Cryptosporidium parvum sporozoites (reprinted from Figs. 4–7 of Ctrnacta et al. 2006 with permission of the publishers). Scale bars = 5.0 µm. a Control localization of cytosolic fatty acid phosphopantetheinyl transferase (CpPPTase) using Alexa-488 donkey anti-rabbit secondary antibody shows diffuse green ﬂuorescence throughout the cytoplasm. b Using Alexa-633 donkey anti-goat secondary antibody against goat IgG antiCpPFO, both the CpPNO in the cytosol and the crystalloid body (arrows) exhibit red ﬂuorescence. c The sporozoites are stained with Alexa-488 donkey anti-rabbit secondary antibody against rabbit IgG anti-CpPFO. An identical pattern of ﬂuorescence in both the cytosol and crystalloid body (arrow) is seen in this ﬁgure and that of b. The nucleus is stained with DAPI (blue). d When a and b are merged, co-localization of CpPNO and CpPPTase within the cytosol is observed (yellow). The higher concentration of CpPNO within the crystalloid body (red) is readily apparent. The nucleus is stained with DAPI

the crystalloid body (Figs. 3b,d and 4, arrows), a cryptosporidium-speciﬁc organelle posterior to the nucleus that is intimately associated with the relic mitochondrion (Ctrnacta et al. 2006; Keithly et al. 2005). Although the role of

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Fig. 4 Transmission electron microscopy of a longitudinal section of the posterior end of a Cryptosporidium parvum sporozoite showing immunogold localization of pyruvate : NADP+ oxidoreductase (CpPNO). The mitochondrion-like organelle (∗ ) is posterior to the nucleus, and lies between the nucleus and the CB. It is labeled by mitochondrionspeciﬁc 15-nm gold anti-CpCpn60 particles. Small 6-nm gold goat anti-CpPFO particles (arrows) show the localization of CpPNO. There are no 6-nm particles localized within the mitochondrion-like organelle (reprinted from Fig. 12 of Ctrnacta et al. 2006 with permission of the publishers)

CpPNO in carbohydrate metabolism is not yet known, a Narf -like hydrogenase has been identiﬁed in both C. parvum (Stejskal et al. 2003; Abrahamsen et al. 2004) and C. hominis (Xu et al. 2004), which may function to oxidize the NADPH produced by PNO during pyruvate decarboxylation. Unlike other amitochondriate protists (Entamoeba, Giardia, Trichomonas), neither of these cryptosporidia possesses an [FeFe]-hydrogenase capable of transferring electrons produced during the oxidation of PFO (Horner et al. 2000). It is proposed that the acetyl-CoA resulting from the decarboxylation of pyruvate in C. parvum may then be converted to malonyl-CoA (Templeton et al. 2004), an important precursor for fatty acid and polyketide biosynthesis (Zhu et al. 2002; Zhu et al. 2004), both of which play an essential role in core fatty acid metabolism of this human pathogen (reviewed in Zhu 2004). However, it is unlikely that fatty acids serve as an energy source in C. parvum because so far only one enzyme of the fatty acid oxidative pathway, 3-hydroxyacyl-CoA dehydrogenase, has been identiﬁed in this apicomplexan (Cai et al. 2005). 4.2 Predicted End Products of Glycolysis Substrate level phosphorylation in C. parvum predicts that several organic end products, including acetate, lactate, malate, and ethanol will be produced, and that as in other anaerobic protists, one molecule of ATP will result when acetate is generated from acetyl-CoA (Crawford et al. 2005; reviewed in Müller 2003; Templeton et al. 2004). The enzymes essential for yielding

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these products have been identiﬁed in the genomes of both C. parvum and C. hominis (Abrahamsen et al. 2004; Xu et al. 2004), and several, including alcohol-, lactate- and malate-dehydrogenases have been biochemically and phylogenetically analyzed (Madern et al. 2004). In addition, C. parvum lactate dehydrogenase (LDH) has been crystallized both in the absence and presence of its substrates and cofactors, whereas pyruvate kinase and glyceraldehyde 3-phosphate dehydrogenase have been crystallized only as apoproteins (Senkovich et al. 2005). These studies conﬁrm the essential role of these enzymes in the core carbohydrate metabolism of C. parvum. Interestingly, LDH has recently been localized within the refractile bodies (RB) of the coccidian E. tenella (de Venevelles et al. 2005), indicating the presence of a metabolic pathway that could generate energy for the invasion of host cells. The possible signiﬁcance of this observation for C. parvum sporozoite entry into host cells is discussed in Sect. 6.

5 Energy Metabolism As mentioned in the previous section, the core energy metabolism of C. parvum is primarily fermentative (Abrahamsen et al. 2004; Entrala and Mascaro 1997), resembling that of the microaerophilic protists G. intestinalis, E. histolytica, Nyctotherus ovalis, and T. vaginalis (reviewed in Crawford et al. 2005 and Müller 2003). Although C. parvum lacks a mitochondrial genome (Abrahamsen et al. 2004; Keeling 2004; Templeton et al. 2004) and oxidative phosphorylation typical of aerobic mitochondria, this apicomplexan does possess a mitochondrial-derived, cyanide-insensitive alternative oxidase (AOX) which has been proposed to transfer electrons from complex I through ubiquinone (Ellis et al. 1994; Roberts et al. 2004; Suzuki et al. 2004). Phylogenetic analyses place the AOX of Cryptosporidium with that of other eukaryotes and α-proteobacteria (Roberts et al. 2004; Suzuki et al. 2004). Although the C. parvum AOX targeting presequence predicts that this protein may serve as an alternative terminal electron acceptor for respiration by the organelle (Henriquez et al. 2005; Roberts et al. 2004), this hypothesis remains to be tested. The presence of AOX in C. parvum is thought to be an evolutionary adaptation for coping with oxidative stress in the host intestine (Putignani et al. 2004). Furthermore, although ATP synthesis is thought to occur primarily by substrate level phosphorylation in the cytosol of C. parvum, both α and β subunits of an F0 F1 ATP synthase (complex V) containing mitochondrial targeting peptides have been found in this parasite (Abrahamsen et al. 2004; Henriquez et al. 2005; Putignani et al. 2004), as well as two genes encoding the ATP/ADP transporter pyridine nucleotide transhydrogenase (PNT) (Abrahamsen et al. 2004). Therefore, ATP may also be generated by proton-gradient-coupled oxidative phosphorylation in the C. parvum

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mitochondrion-like organelle. In fact, in both Plasmodium and Cryptosporidium, it has been recently proposed that the membrane potential could be generated through a combined action of the matrix-located F0 F1 of ATP synthase and one (or both) of these membrane-located ATP/ADP transporters (Painter et al. 2007). Brieﬂy, since only a few ions are required for movement across the membrane, ATP might be hydrolyzed by the F0 F1 ATP synthase, and then the ADP generated by it exchanged by one of the two PNT transporters (Painter et al. 2007). Since this tiny organelle lacks both mtDNA and an electron transport chain, this would certainly help answer two frequently asked questions, i.e. how does C. parvum generate a membrane potential for exporting FeS proteins from the relic mitochondrion into the cytoplasm for FeS cluster assembly (Kispal et al. 2005; Lill et al. 2005), and for taking up mitotracker dyes into the organelle (Ctrnacta et al. 2006; Kayser et al. 2002). A novel PNT transporter has also been recently characterized from the mitosome of the microaerophilic protist E. histolytica (Chan et al. 2005).

6 The Crystalloid Body 6.1 Ultrastructural Morphology One of the most puzzling structures within C. parvum sporozoites, is the crystalloid body (CB), which is also in intimate contact with the relic mitochondrion, outer nuclear membrane, and RER (Fig. 1) (Keithly et al. 2005). Although it is still unclear whether this organelle is surrounded by a limiting membrane, or is simply a complex of closely packed membrane-bounded vesicles, it has been shown that like the relic mitochondrion, the CB takes up mitotracker dyes (Ctrnacta et al. 2006; Kayser et al. 2002; Keithly et al. 2005). This observation suggests that the CB also has a mechanism for generating a membrane potential (see Sect. 5). In addition, its total volume in sporozoites is at least equal to that of the nucleus, which it appears to wrap around (Figs. 1, 3) (Ctrnacta et al. 2006; Keithly et al. 2005). Together these data suggest that the CB plays an essential role during the life cycle of this parasite. Ultrastructural data have indicated that the CB of C. parvum differs markedly from the paracrystalline CB seen in ﬁsh coccidia (Lukeˇs 1992), and the randomly or regularly arrayed non-membranous particles observed amongst certain haemogregarines, haemoprotozoa (Leucocytozoon) and archigregarines (Schrevel 1971a,b; Siddall and Desser 1992; Treﬁak and Desser 1973; Vivier and Schrevel 1966). Furthermore, although the C. parvum CB showed some indication of long-range order in computed diffraction patterns (i.e. Fourier power spectra), these were much less obvious than the cubic

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paracrystalline patterns of mitochondrial IM observed in starved free-living amoebae (Deng et al. 1999). The vesicular nature of the CB in sporozoites visualized in situ by us (Keithly et al. 2005) was nearly identical to that of subcellular fractions from partially disrupted sporozoites collected by sucrose density centrifugation and viewed by negative staining (Petry and Harris 1999). One striking feature of the isolated CB we noted, however, was the structural similarity of the negatively stained vesicles to those of ferritin, a non-toxic iron storage molecule typical of many eukaryotes (Harris and Schefﬂer 2002; Johnson et al. 2005). Therefore, it seemed reasonable to test the hypothesis that the CB might be a compartmentalized reservoir of soluble iron near the relic mitochondrion where FeS clusters are assembled. Preliminary studies, however, using various ferrous- and ferric-speciﬁc dyes showed no localization within the organelle (M. Behr, H. Johnson, A. Fillion, J.S. Keithly, unpublished results). This hypothesis has yet to be vigorously tested. 6.2 Putative Functions Although neither the function nor the origin of this enigmatic organelle is yet known, the three-dimensional structure of the CB visualized by tomographic reconstruction suggested that the closely packed vesicles observed by TEM might actually be a series of interconnected channels (see Fig. 20, Keithly et al. 2005). The completion of the C. parvum (Abrahamsen et al. 2004) and C. hominis (Xu et al. 2004) genomes, combined with a recent proteomic analysis of Eimeria tenella isolated RB (de Venelles et al. 2006) are providing additional insights into the possible functions of the C. parvum CB. Within the genome of C. parvum, for example, there has been a lineage-speciﬁc expansion of several protease families (Abrahamsen et al. 2004; Aravind et al. 2000; Templeton et al. 2004), including aspartyl proteases, cryptopain-like cysteine proteases, a Plasmodium facilysin-like insulin degrading protease, and a single subtilisin-like protease. The most interesting of these are the aspartyl proteases which include homologs of proline- and valine-rich articulins. These are an ancient cytoskeletal feature of the clade Alveolata, to which C. parvum belongs, and that form part of the subpellicular network typical of all apicomplexans (Mann and Beckers 2001; Morisette and Sibley 2002). Recent cryo-electron tomography has conﬁrmed the presence of articulin cytoskeletal ﬁbers surrounding the CB of C. parvum sporozoites suggesting that these may form a supporting scaffold for the organelle (C. Hsieh, M. Marko, J.S. Keithly, C.A. Mannella, unpublished results). These data are congruent with the “cup-like” subpellicular network previously observed in the extreme posterior end of T. gondii tachyzoites (Morrisette and Sibley 2002). In addition, it has been recently documented that the RB of E. tenella synthesize several aspartyl proteases that are later trafﬁcked to the apical

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organelles and secreted during host cell invasion (de Venelles et al. 2006; Fetterer et al. 2007). Monoclonal antibodies to the two most abundant RB aspartyl proteases, eimepsin and SO7 inhibit sporozoite entry into chick intestinal epithelial cells (de Venevelles et al. 2006). The CB of C. parvum and the RB of E. tenella are differentially expressed during the life cycle of these parasites, i.e. both organelles are found only in sporozoites and ﬁrst generation meronts (de Venelles et al. 2006; reviewed in Fayer 1997 and Thompson et al. 2005). After invasion, these organelles disappear, and reappear only in newly shed oocysts. For that reason, it has been suggested that like the RB of E. tenella, the CB of C. parvum may also synthesize enzymes that promote invasion of intestinal and bile duct epithelial cells (Coombs and Müller 2002; reviewed in Crawford et al. 2005, Fayer 1997, and Thompson et al. 2005). Probably the most compelling evidence that the CB of C. parvum may be involved in the invasion of host cells is the observation that CpPNO is concentrated within this organelle (Figs. 3a–c, 4; Ctrnacta et al. 2006). As mentioned previously, it is well known that oxygen-sensitive PFO is the key enzyme for anaerobic core energy metabolism in parasitic protists, and that it may be compartmentalized either into the cytosol (Giardia Entamoeba), or within the hydrogenosome (Trichomonas, Nyctotherus) (Embley et al. 2003; Müller 2003). Signiﬁcantly, the PNO of C. parvum is localized not only in the cytosol, where substrate level phosphorylation generates two molecules of ATP (reviewed in Crawford et al. 2005; Katinka et al. 2001; Williams et al. 2004), but also within the cryptosporidium-speciﬁc CB. Therefore, it is reasonable to think that the CB might generate additional molecules of ATP to power the invasion of host cells.

7 Concluding Remarks and Future Perspectives Research in the last few years, including the completion of the genome projects for Cryptosporidium parvum and C. hominis, has provided a wealth of information on the structure and likely functions of the relic mitochondrion, its associated membranes and organelles. Nevertheless, research is still hampered by the absence of an in vitro system for cultivating these parasites, which would permit comparisons of gene expression during the life cycle. Although most of the genes for mitochondrial and cytosolic iron sulfur cluster biosynthesis, protein import and export into the mitochondrionlike organelle, and an alternative oxidase system occur within the genome of human cryptosporidia, these functions still remain to be experimentally determined. The same may be said for the core carbohydrate and energy metabolism. The novel compartmentalization of the fusion enzyme CpPNO into both the cytosol and CB suggests that a highly specialized type of core energy metabolism may exist in C. parvum. Intriguing clues about the struc-

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ture and function of the crystalloid body are beginning to emerge as a result of cryo-electron microscopy and tomography, and it is likely that in the near future the isolation of the CB and mitochondrion-like organelle, combined with proteomics, will ﬁnally unravel the mystery of their association with the RER and with one another. Further exploration of this unusual compartmentalization might also provide insights into the evolutionary position of the cryptosporidia amongst mitosome- and hydrogenosome-bearing anaerobic eukaryotes, as well as eventually leading to better strategies for the development of new drugs against human cryptosporidiosis. Acknowledgements I wish to thank all of my coworkers at the Wadsworth Center who have made critical contributions to this evolving story. The following were invaluable in this research: Guan Zhu, Christina Riordan, Michael J. LaGier, as well as Fogarty Fellows Frantisek Stejskal, Jan Slapeta, and Vlasta Ctrnacta. To them, I express my sincere thanks. For excellence in electron microscopy and tomography, I wish to thank all of the members of the Resource for the Visualization of Biological Complexity (RVBC), but most especially to Carmen Mannella and Karolyn Buttle, who have opened my eyes to the nuances of mitochondrial membrane structure and function. Finally, to my friend and colleague Susan Langreth at Uniformed Services University, a special debt of thanks for her excellent electron microscopy and collaborating support through the years. This research was generously supported by the National Institutes of Health (NIH) Allergy and Infectious Disease Opportunistic Infections Branch, Training Grants from the NIH Fogarty International Center, and the National Center for Research Resources.

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Suchan P, Vyoral D, Petrak J, Sutak R, Rasoloson D, Nohynkova E, Dolezal P, Tachezy J (2003) Incorporation of iron into Tritrichomonas foetus cell compartments reveals ferredoxin as a major iron-binding protein in hydrogenosomes. Microbiol-UK 149:1911–1921 Sutak R, Dolezal P, Fiumera HL, Hrdy I, Dancis A, Delgadillo-Correa M, Johnson PJ, Müller M, Tachezy J (2004) Mitochondrial-type assembly of FeS centers in the hydrogenosomes of the amitochondriate eukaryote Trichomonas vaginalis. Proc Natl Acad Sci USA 101:10368–10373 Suzuki T, Hashimoto T, Yabu Y, Kido Y, Sakamoto K, Nihei C, Hato M, Suzuki S, Amano Y, Nagai K, Hosokawa T, Minagawa N, Ohta N, Kita K (2004) Direct evidence for cyanide-insensitive quinol oxidase (alternative oxidase) in the apicomplexan parasite Cryptosporidium parvum: phylogenetic and therapeutic implications. Biochem Biophys Res Commun 313:1044–1052 Tachezy J, Dolezal P (2007) Iron-sulfur proteins and iron-sulfur cluster assembly in organisms with hydrogenosomes and mitosomes. In: Martin WF, Müller M (eds) Origin of Mitochondria and Hydrogenosomes. Springer, Berlin Heidelberg New York, pp 105– 133 Tan TC, Suresh KG (2006) Amoeboid form of Blastocystis hominis—a detailed ultrastructural insight. Parasitol Res 99:737–742 Templeton TJ, Iyer LM, Anatharam V, Enomoto S, Abrahante JE, Subramanian GM, Hoffman SL, Abramsen MS, Aravind L (2004) Comparative analysis of Apicomplexa and genomic diversity in eukaryotes. Genome Res 14:1686–1695 Tetley L, Brown SMA, McDonald V, Coombs GH (1998) Ultrastructural analysis of the sporozoite of Cryptosporidium parvum. Microbiol-UK 144:3249–3255 Thompson RC, Olson ME, Zhu G, Enomoto S, Abrahamsen MS, Hijjawi NS (2005) Cryptosporidium and cryptosporidiosis. Adv Parasitol 59:77–158 Tielens AGM, van Hellemond JJ (2006) Anaerobic mitochondria: properties and origins. In: Martin WF, Müller M (eds) Origin of Mitochondria and Hydrogenosomes. Springer, Berlin Heidelberg New York, pp 86–103 Tovar J (2006) Mitosomes of parasitic protozoa: biology and evolutionary signiﬁcance. In: Martin WF and Müller M (eds), Origin of Mitochondria and Hydrogenosomes. Springer, Berlin Heidelberg New York, pp 277–300 Tovar J, Fischer A, Clark CG (1999) The mitosome a novel organelle related to mitochondria in the amitochondrial parasite Entamoeba histolytica. Mol Microbiol 32:1013– 1021 Tovar J, Leon-Avila G, Sanchez LB, Sutak R, Tachezy J, van der Giezen M, Hernandez M, Müller M, Lucocq JM (2003) Mitochondrial remnant organelles of Giardia function in iron-sulphur protein maturation. Nature 426:172–176 Treﬁak WD, Desser SS (1973) Crystalloid inclusions in species of Leucocytozoon, Parahaemoproteus and Plasmodium. J Protozool 20:73–80 Uni S, Iseki M, Maekawa T, Moriya K, Takada S (1987) Ultrastructure of Cryptosporidium muris (strain RN 66) parasitizing the murine stomach. Parasitol Res 74:123–132 van der Giezen M, Cox S, Tovar J, Clark CG (2005) Mitochondrion-derived organelles in protists and fungi. Int Rev Cytol 244:175–225 van Hoek AHAM, Akhmanova AS, Huynen MA, Hackstein JHP (2000) A mitochondrial ancestry of the hydrogenosomes of Nyctotherus ovalis. Mol Biol Evol 17:202–206 Vivier E, Schrevel J (1966) Les ultrastructures cytoplasmiques de Selenidium hollandei, n. sp. gregarine parasite de Sabellaria alveolata L. J Microscopie 5:213–228 Williams BAP, Keeling PJ (2003) Cryptic organelles in parasitic protists and fungi. Adv Parasitol 54:10–68

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Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_114/Published online: 8 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Mitochondrial Remnant in Blastocystis Nigel Yarlett1,2 (u) · Kevin S. W. Tan3 1 Haskins

Laboratories, Pace University, New York, NY 10038, USA [email protected] 2 Department of Chemistry and Physical Science, Pace University, New York, NY 10038, USA 3 Laboratory of Molecular and Cellular Parasitology and Infectious Diseases Programme, Department of Microbiology, Yong Loo Lin School of Medicine, National University of Singapore, 117597 Singapore 1

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Abstract The intestinal protozoan Blastocystis is an anaerobic organism that contains from a few to numerous mitochondria-like organelles (MLOs). The organelles are round to ovoid in shape, are 0.5–2 µm in diameter, and are bounded by an outer bilaminar membrane and an inner bilaminar membrane, from which the cristae arise. Previous studies have reported that key mitochondrial proteins and cytochromes are absent from these structures. However, recent work by the authors have shown that while key enzymes characterizing hydrogenosomes were absent, a partial Krebs cycle exists. We hypothesize that the Blastocystis MLO functions to couple the succinyl CoA formed from citrate to the generation of succinate required for recycling of acetyl CoA to acetate, and formation of ATP. The primary function of the Blastocystis MLOs may therefore be to act as acetyl CoA producers for the synthesis of succinate and fatty acids. The Blastocystis MLOs have also been reported to be involved in apoptosis-like programmed cell death although MLO-associated apoptotic factors have yet to be identiﬁed.

1 Introduction The compartmentalization of energy generation provides a mechanism for the increased efﬁciency of high energy bond transfer to form the ultimate cellular fuel ATP, and has been the driving force behind the evolution of the mitochondrion. This viewpoint is supported by studies of the mitochondrial proteome, which have demonstrated that proteins of eubacterial origin predominantly

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have roles in bioenergetics whereas the eukaryotic-speciﬁc mitochondrial proteins perform transport and regulatory functions (Gray et al. 2001). This is consistent with the proposed α-proteobacterial origin for this organelle and the switch to aerobic respiration (Kurland and Andersson 2000). The identiﬁcation and characterization of organelles that shuttle electrons to protons forming molecular hydrogen (hydrogenosome) under anaerobic conditions has challenged the concept that the mitochondrion is the sole surviving example of the evolution of an energy-generating organelle (Lindmark and Müller 1973; Yarlett et al. 1981, 1986; Akhmanova et al. 1998). However, as molecular techniques improved it became clear that many anaerobes previously considered to be amitochondriate in fact harbor membranes or highly reduced structures that can be considered of mitochondrial origin (Tovar et al. 1999; Marti et al. 2003; Keithly et al. 2005; Burri et al. 2006). Even the most characterized organelle, the hydrogenosome, which is also the most complex of the anaerobic structures, has been demonstrated to have mitochondrial features (Bui et al. 1996; Akhmanova et al. 1998; Tachezy et al. 2001; Embley et al. 2003; Dacks et al. 2006). These ﬁndings have coalesced into essentially three concepts for the origin and evolution of energy generating organelles: 1. These organelles arose multiple times from multiple origins 2. The various structures that are present in all extant organisms are of the same eubacterial origin, which subsequently diverged 3. The variations observed in organelles are the result of multiple invasions, which have evolved to utilize key functions as each wave occurred with the result that distinctions between each ancestor has blurred into the present day collage There is compelling evidence that the hydrogenosomes described from the trichomonads, rumen protists, rumen fungi and protists inhabiting the cockroach hind gut have many mitochondrial features including the recognition of speciﬁc targeting sequences (Dacks et al. 2006), ADP/ATP carriers (Bui et al. 2006), the production of ATP (Embley et al. 2003; van Weelden et al. 2006), and the iron-sulfur cluster assembly (Tachezy et al. 2001). These studies prompted examination of other previously considered amitochondriate organisms, which led to the ﬁnding that many of these organisms possessed mitochondria-like organelles (MLO), e.g., the mitosome of Entamoeba (Tovar et al. 1999), the crytic organelle of Giardia (Marti et al. 2003), the relict organelle in Cryptosporidia (Keithly et al. 2005), and the Microsporidians, Antonospora locustae and Encephalitozoon cuniculi (Burri et al. 2006). Despite little homology to the mitochondrion, in many cases these organelles have many features in common with each other and with the mitochondrion. These studies have reinforced the conclusion that all of these organelles arose from a common origin. The protozoan Blastocystis is an inhabitant of the intestinal tract of mammals and some reptiles. B. hominis is the most frequently reported protozoon

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Fig. 1 Enzymes localized to B. hominis mitochondrial-like organelle. The enzymes are: 1 malic enzyme, 2 pyruvate : NADP oxidoreductase, 3 acetate : succinate CoA transferase, 4 succinate thiokinase, 5 α-ketoglutarate dehydrogenase, 6 isocitrate dehydrogenase, and 7 aconitase

in the human intestinal tract (Lavier 1952; Lee 1991) and has been reported to cause intestinal disease in a number of cases resulting in abdominal cramps, diarrhea, and signiﬁcant discomfort (Weg et al. 1987). In some cases the parasite has been reported to cause acute illness (Russo et al. 1988; Vannetta et al. 1985) and can be life-threatening when present in immunocompromised individuals (Prasad et al. 2000). In common with many lumen-dwelling parasites, the ﬁrst line of treatment is metronidazole (Zierdt et al. 1983) but resistance has been reported (Haresh et al. 1999). Antimicrobials such as paromomycin and ketoconazole have also been tried with mixed success (Dunn and Boreham 1991).

2 Morphology Within vacuolar forms, MLOs are usually situated within the cytoplasmic rim, near the nucleus, and the conﬁguration of a nuclei ﬂanked on each side by one or more MLO is often observed in transmission electron microscopy (TEM) sections (Fig. 2) (Zierdt 1986; Dunn et al. 1989; Stenzel et al. 1994; Stenzel and Boreham 1996). However, there have been reports of cytoplasmic invagination resulting in organelles, including MLOs, being deposited within the

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Fig. 2 Ultrastructure of Blastocystis. Transmission electron micrograph revealing organelle structures. In this cell, MLOs (M) ﬂank the nucleus (Nu) and are characterized by electron-dense matrix and distinct saccate cristae. Cv Central vacuole. Bar = 1 µm

central vacuolar space (Pakandl 1999; Nasirudeen et al. 2001; Tan et al. 2001). The exact role of this phenomenon is currently unclear although this feature, which has been described in Blastocystis undergoing cell death, was postulated to be part of an apoptotic body deposition process (Tan and Nasirudeen 2005). These organelles usually range from two to four per cell but can reach up to hundreds in giant and older cells (Zierdt et al. 1988). Dunn et al. (1989) performed an ultrastructural study of ten different B. hominis stocks and observed that the MLOs of Blastocystis exhibited considerable variation in morphology, both between different isolates and within the same isolate. In fact, morphological heterogeneity could be observed between MLOs of a single cell (Dunn et al. 1989). Ultrastructurally, Blastocystis MLO possess basic features archetypical of eukaryotic mitochondria. The organelles are round to ovoid in shape, are 0.5–2 µm in diameter, and are bounded by an outer bilaminar membrane and an inner bilaminar membrane, from which the cristae arise. The MLO cristae are pleomorphic and have been described as bulbous, tubular, and vesicular. Occasionally, cristae may take on a circular shape and appear as a ring-like structure within the MLO matrix (Dunn et al. 1989). The MLO matrix may exhibit variation in electron density (Silard et al. 1983; Stenzel et al. 1991), although MLOs with electron-dense matrix are most often described (Zierdt 1991; Boreham and Stenzel 1993; Stenzel and Boreham 1996). An early freeze-fracture study (Yoshikawa et al. 1988) revealed that the MLOs of Blastocystis were round and contained tubular cristae. Intramembranous particles were distributed heterogeneously on both the protoplasmic and exoplasmic surfaces of the MLO outer membrane. The MLOs of Blastocystis are selectively stained by a number of dyes employed for staining mitochondria. Early studies reported the use of Janus green (Lavier 1952; Zierdt 1986; Zierdt et al. 1988) and rhodamine 123 (Zierdt 1986; Zierdt et al. 1988) while more recent reports describe speciﬁc staining with MitoTracker Green (Fig. 3) (Nasirudeen and Tan 2004a; Nasirudeen and Tan 2005) and MitoLight (Nasirudeen and Tan 2004b). Rhodamine 123, MitoTracker, and MitoLight accumulate in mitochondria due to the presence of a mitochondrial membrane potential. This indicates that the MLOs

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Fig. 3 Fluorescence microscopy of Blastocystis revealing MitoTracker Green uptake of MLOs with DAPI colocalization. The MLOs are situated very close to the nucleus (large DAPI-stained structures) and are clustered at opposite poles of the cell

of Blastocystis are functional and are able to generate energy to maintain this potential. Interestingly, the ability of the potential-sensitive ﬂuorophore MitoTracker Green to accumulate in MLOs was compromised in Blastocystis cells undergoing programmed cell death (PCD) (Nasirudeen and Tan 2004a, 2005). This is a common feature of PCD in the Metazoa and functions to release proapoptotic factors sequestered within the mitochondria into the cytoplasm. Pre-incubation of cells with cyclosporine A, a mitochondria permeability transition pore blocker, prevented a decrease in MitoTracker Green uptake in PCD-induced cells, with a concomitant decrease in DNA fragmentation, a key marker of programmed cell death (see next section). Interestingly, the MLO also stain positively for 4 ,6-diamidino-2-phenylindole (DAPI), indicating the presence of DNA within the organelle (Fig. 3) (Matsumoto et al. 1987; Nasirudeen and Tan 2004b). There has been a recent report describing a technique for the isolation and puriﬁcation of Blastocystis MLOs (Nasirudeen and Tan 2004b). After density gradient centrifugation, the authors observed, by

Fig. 4 Negative staining of puriﬁed Blastocystis MLOs. Note the distinct ovoid to elongate shapes, with apparent wrinkled surface texture. Bar = 1

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negative staining and TEM, that the organelles were predominantly elongate, and ranged approximately from 1.2 to 3.0 µm in length and approximately 0.7 to 1.0 µm in width. This is in general agreement with previous reports on MLO size (Dunn et al. 1989). The MLO membrane appeared wrinkled and the organelle appeared morphologically intact (Fig. 4). This method will be useful for addressing the major metabolic functions of the organelle as well as characterizing the DNA contained within.

3 Biochemistry of Mitochondria-Like Organelle of Blastocystis The authors have recently demonstrated the presence of key energy coupling reactions in B. hominis and the localization of these enzymes to a MLO (Lantsman et al. 2007, submitted). Biochemically the metabolism of the Blastocystis organelle reﬂects the anaerobic nature of the organism. Key enzymes characterizing hydrogenosomes, hydrogenase and pyruvate : ferredoxin oxidoreductase, were absent in B. hominis. However, an oxygen-sensitive pyruvate : NADP oxidoreductase, which had characteristics in common with the pyruvate : NADP oxidoreductase present in the mitochondria of Euglena gracilis (Inui et al. 1987), was present that was shown to convert pyruvate to acetyl CoA. A signiﬁcant feature of pyruvate metabolism by the B. hominis organelle is the transformation of pyruvate to acetyl CoA and the conservation of the thioester bond to form ATP. In common with trichomonad and Neocallimastix hydrogenosomes (Steinbuchel and Muller 1986; Yarlett et al. 1986) and mitochondria from promastigote Leishmania mexicana, L. infantum, Phytomonas sp., and procyclic Trypanaosoma brucei, as well as the parasitic helminth Fasciola hepatica (van Hellmond et al. 1998), B. hominis utilizes a two-step mechanism that couples acetate with succinate formation to conserve the energy of the thioester bond to drive subcellular ATP formation (Fig. 1). This ﬁnding is consistent with the proposed common origin of hydrogenosomes and mitochondria (Cavalier-Smith 1987). The B. hominis organelle also has remnants of a Krebs cycle (Fig. 1); in contrast to true mitochondria these enzymes were oxygen sensitive. The key feature of the proposed pathway of B. hominis is the channeling of citrate and pyruvate to succinate resulting in the synthesis of two moles of ATP. The absence of detectable activity for succinyl CoA synthetase, succinate dehydrogenase, and fumarate hydratase (fumarase), which have been demonstrated to have a key role in enzyme-to-enzyme channeling of intermediates of the Krebs cycle in many cells (Malaisse et al. 1996), is indicative that the primary function of the MLO from B. hominis is to couple the succinyl CoA formed from citrate to the generation of succinate required for recycling of acetyl CoA to acetate, and formation of ATP. The primary function of the Blastocystis MLOs may therefore be to act as acetyl CoA producers for the synthesis of succi-

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nate and fatty acids. The presence of a partial TCA cycle has been reported in the mitochondria from procyclic Trypanosoma brucei (van Hellemond et al. 2005; van Weelden et al. 2005), which is proposed to function to export acetyl CoA to the cytoplasm for fatty acid biosynthesis, and the degradation of proline and glutamate to succinate. The plasticity of this organelle is therefore evident from the literature and we therefore conclude that despite the lack of cytochromes or other identiﬁable electron transport proteins the organelles from B. hominis (which are characterized as DNA-containing, double membrane-bound structures having an active ATP generating metabolism) are best described as mitochondria-like organelles. Inevitably, all of these organelles function as redox organelles to shuttle protons and electrons derived from pyruvate to produce acetyl CoA for ATP synthesis or fatty acid biosynthesis.

4 Role of MLO in Blastocystis Programmed Cell Death Programmed cell death (PCD) is a cell suicide mechanism that enables multicellular organisms to control cell number and eliminate undesirable cells. It has recently been established that PCD also exists among the unicellular eukaryotes and probably function similarly, when these microbes are viewed from a population-based perspective (DosReis and Barcinski 2001). Blastocystis has also been reported to undergo PCD when exposed to a variety of undesirable stimuli (Tan and Nasirudeen 2005). As in higher eukaryotic mitochondria, the MLOs are also involved in PCD. Mitochondrial outer membrane potential (MOMP), an indication of an actively respiring organelle, was compromised in cells that were exposed to a cytotoxic monoclonal antibody (Nasirudeen and Tan 2005). In mammalian cells, MOMP is directly governed by a subset of proapoptotic proteins, which induce disruptions in the outer mitochondrial membrane and subsequent release of death-promoting proteins like cytochrome c. Cytochrome c triggers a cascade of events that lead to the activation of caspases, enzymes that are intimately involved in dismantling the dying cell. It is currently unknown what proapoptotic factors are released by Blastocystis upon PCD induction, since cytochromes have not been identiﬁed in this organism. PCD involving mitochondria can be either caspase-dependent or caspase-independent. Interestingly, blocking caspases in PCD-induced cells led to a greater compromise in MOMP, suggesting that MLOs trigger a compensatory caspase-independent pathway (Nasirudeen and Tan 2005). Perhaps one of the key roles of Blastocystis MLO, is the regulation of PCD during times of cellular starvation and stress, as has been reported for other cell types (Goyeneche et al. 2006; Yeo et al. 2006). The identiﬁcation of novel MLO-associated PCD mediators in Blastocystis may provide novel strategies for antiparasite therapies.

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5 Future Directions The presence of key energy coupling reactions in B. hominis conﬁrms the role of this organelle in the compartmentalization of redox reactions and energy generation. These biochemical ﬁndings support the naming of this organelle as a MLO and the concept of a common origin for all characterized redox organelles and their remnants from the same α-proteobacterial endosymbiont. The key feature of the proposed pathway of B. hominis is the channeling of citrate and pyruvate to succinate, resulting in the synthesis of two moles of ATP. The absence of detectable activity for succinyl CoA synthetase, succinate dehydrogenase, and fumarate hydratase (fumarase), which have been demonstrated to have a key role in enzyme-to-enzyme channeling of intermediates of the Krebs cycle in many cells (Malaisse et al. 1996), is indicative that the primary function of the MLO from B. hominis is to couple the succinyl CoA formed from citrate to the generation of succinate required for recycling of acetyl CoA to acetate, and formation of ATP. The primary function of the MLOs may therefore be to act as acetyl CoA producers for the synthesis of succinate and fatty acids. The role of fatty acid metabolism in the function of energy metabolism in B. hominis MLOs will be explored in future studies. Acknowledgements This work was supported in part by grants from the National Institutes of Health NCDDG AI40320 (NY), the Biomedical Research Council (KSWT), and the National Medical Research Council (KSWT).

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Stenzel DJ, Boreham PF (1996) Blastocystis hominis revisited. Clin Microbiol Rev 9:563– 584 Stenzel DJ, Boreham PF, McDougall R (1991) Ultrastructure of Blastocystis hominis in human stool samples. Int J Parasitol 21:807–812 Stenzel DJ, Cassidy MF, Boreham PF (1994) Morphology of Blastocystis sp. from domestic birds. Parasitol Res 80:131–137 Tachezy J, Sánchez LB, Müller M (2001) Mitochondrial type iron-sulfur cluster assembly in the amitochondriate eukaryotes Trichomonas vaginalis and Giardia intestinalis, as indicated by the phylogeny of IscS. Mol Biol Evol 18:1919–1928 Tan KS, Howe J, Yap EH, Singh M (2001) Do Blastocystis hominis colony forms undergo programmed cell death? Parasitol Res 87:362–367 Tan KS, Nasirudeen AM (2005) Protozoan programmed cell death – insights from Blastocystis deathstyles. Trends Parasitol 21:547–550 Tovar J, Fischer A, Clark CG (1999) The mitosome, a novel organelle related to mitochondria in the amitochondrial parasite Entamoeba histolytica. Mol Microbiol 32:1013– 1021 van Hellemond JJ, Opperdoes FR, Tielens AG (1998) Trypanosomatidae produce acetate via a mitochondrial acetate:succinate CoA transferase. Proc Natl Acad Sci USA 95:3036–3041 van Hellemond JJ, Opperdoes FR, Tielens AG (2005) The extraordinary mitochondrion and unusual citric acid cycle in Trypanosoma brucei. Biochem Soc Trans 33:967–971 van Weelden SWH, van Hellemond JJ, Opperdoes FR, Tielens AGM (2005) New functions for parts of the Krebs cycle in procyclic Trypanosoma brucei, a cycle not operating as a cycle. J Biol Chem 280:12451–12460 Vannetta JB, Adamson D, Mullican K (1985) Blastocystis hominis infection presenting as recurrent diarrhea. Ann Int Med 102:495–496 Weg AL, Soave R, Jacobson IM (1987) The signiﬁcance of intestinal Blastocystis hominis infection. Gastroent 92:1688 Yarlett N, Hann AC, Lloyd D, Williams AG (1981) Hydrogenosomes in the rumen protozoon Dasytricha ruminantium Schuberg. Biochem J 200:365–372 Yarlett N, Orpin CG, Munn EA, Yarlett NC, Greenwood CA (1986) Hydrogenosomes in the rumen fungus Neocallimastix patriciarum. Biochem J 236:729–739 Yeo JH, Lo JC, Nissom PM, Wong VV (2006) Glutamine or glucose starvation in hybridoma cultures induces death receptor and mitochondrial apoptotic pathways. Biotechnol Lett 28:1445–1452 Yoshikawa H, Yamada M, Yoshida Y (1988) Freeze-fracture study of Blastocystis hominis. J Protozool 35:522–528 Zierdt CH, Swan JC, Hosseini J (1983) In vitro response of Blastocystis hominis to antiprotozoal drugs. J Protozool 30:332–334 Zierdt CH (1986) Cytochrome-free mitochondria of an anaerobic protozoan – Blastocystis hominis. J Protozool 33:67–69 Zierdt CH (1991) Blastocystis hominis – past and future. Clin Microbiol Rev 4:61–79 Zierdt CH, Donnolley CT, Muller J, Constantopoulos G (1988) Biochemical and ultrastructural study of Blastocystis hominis. J Clin Microbiol 26:965–970

Microbiol Monogr (9) J. Tachezy: Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes DOI 10.1007/7171_2007_107/Published online: 7 December 2007 © Springer-Verlag Berlin Heidelberg 2007

Possible Mitochondria-Related Organelles in Poorly-Studied “Amitochondriate” Eukaryotes Vladimir Hampl (u) · Alastair G. B. Simpson (u) Department of Biology, Dalhousie University, Halifax, B3H 2N9, Canada [email protected], [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Pelobionts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Metamonada . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Heterolobosea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Postgaardi mariagerensis . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Andalucia incarcerata . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Breviata anathema . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract The diversity of mitochondria-like organelles is not restricted to the beststudied examples presented in the previous chapters. The recognised diversity of anaerobic and microaerophilic organisms is broader and they are distributed all across the eukaryotic tree. Many of them – pelobionts, most or all metamonads, anaerobic heteroloboseans, Postgaardi, Andalucia incarcerata and Breviata – harbour double membranebounded organelles that do not resemble classical mitochondria, in the sense that they do not have cristae, but are probably homologous to them. For various reasons, these organelles, and the organisms in which they are found, have been studied very little. For example, most of these taxa are not signiﬁcant from a medical or economic perspective, some were recognised only recently and many are difﬁcult to handle in the laboratory. Therefore, the relationship of these organelles to mitochondria and their functional properties are deduced, so far, only from indirect evidence including the presence of a double-membrane envelope, their association with hydrogen consuming methanogens and the discovery of mitochondrion- or hydrogenosome-speciﬁc genes in the organism’s genome. Although studies on the metabolic properties of these organelles will struggle with technical obstacles, they promise interesting insights into the versatility of mitochondrial structure and function, and the evolution of anaerobic/microaerophilic life histories.

1 Introduction In this chapter we attempt to complete the list of known eukaryotic taxa that harbour anaerobic mitochondria-like organelles by introducing those

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that are not covered in previous chapters. At present, the information on the organelles in these taxa is generally very sparse and in most cases is limited to electron microscopy observations. However, the current boom of EST projects on several of these species, followed by cell biological and biochemical investigations, promises imminent improvement in this area. It is commonly assumed that all eukaryotes contain the mitochondrion in some form, but the main reason for retaining the organelle, which in many cases may have lost its function in energy production, is not known. Comparative functional analysis of a broad variety of anaerobic mitochondria-like organelles (MLOs) may reveal their full functional potential and may also provide a window into the process of transition to anaerobiosis itself. As we will deal in the text with organisms that fall into various eukaryotic groups, we feel the need to ﬁrst introduce the reader to the current picture of eukaryotic diversity and evolution. According to the prevailing model, all studied eukaryotes fall into six large “supergroups” plus several small lineages with uncertain placement (reviewed in Simpson and Roger 2004a; Keeling et al. 2005; Adl et al. 2005). A schematic tree of eukaryotes is given in Fig. 1. The six supergroups are as follows: Opisthokonta (animals, fungi and several protist lineages), Amoebozoa (entamoebae, pelobionts, mycetozoan slime molds and the various “lobose amoebae”), Rhizaria (e.g. cercozoans, haplosporidia, foraminiferans and radiolarians), Chromalveolata (apicomplexans, ciliates, dinoﬂagellates, stramenopiles, haptophytes, and cryptophytes plus kathablepharids), Archaeplastida (green algae, land plants, red algae and glaucophytes) and Excavata (jakobids, diplomonads, parabasalids, oxymonads, euglenozoans, heteroloboseans and several other genera of aerobic and anaerobic ﬂagellates). Some of these supergroups are somewhat controversial, especially Excavata and Chromalveolata. The position of the root of the eukaryotic tree is very difﬁcult to infer. Three rooting options are currently considered most promising: (1) Rooting inside Excavata on the branch leading to diplomonads is preferred by molecular phylogenies rooted by prokaryotic outgroups (Bapteste et al. 2002; Arisue et al. 2005; Ciccarelli et al. 2006), but is probably caused by the long branch attraction artefact of tree construction (Felsenstein 1978; Bapteste et al. 2002). (2) The distribution in the tree of a gene fusion character and ﬂagellar apparatus maturation patterns, and the evolution of the myosin protein family is interpreted as evidence for a rooting between unikonts (opisthokonts and amoebozoans) and bikonts (archaeplastids, chromalveolates, rhizarians and excavates) (Stechmann and Cavalier-Smith 2002, 2003; Richards and CavalierSmith 2005). (3) Mitochondrial genomes of jakobid ﬂagellates (e.g. Reclinomonas and Jakoba), a subgroup of excavates, suggest that these taxa may be among the earliest diverging eukaryotes, because they encode the most complete set of protein coding genes of any known mitochondria, including bacterial-like RNA polymerase subunits (Lang et al. 1997).

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Fig. 1 Schematic representation of the current knowledge on relationships among eukaryotic taxa. Eukaryotes fall into six large “supergroups” (Opisthokonta, Amoebozoa, Rhizaria, Archaeplastida, Chromalveolata and Excavata) and several minor lineages with uncertain placement (Incertae sedis). Filled circles indicate lineages with no mitochondria whose species either contain mitochondria-like organelles (MLOs) or no organelle has been observed in them so far, open circles indicate lineages containing both species with mitochondria and species with MLOs

Irrespective of where the eukaryotic root lies, the distribution of anaerobic MLOs – including hydrogenosomes and mitosomes – shows them scattered across the tree (Fig. 1). Assuming that oxidative phosphorylation, where present, is homologous and not transferred laterally, these MLOs have probably evolved many times independently. The list of taxa containing MLOs is given in Table 1. The relatively well-characterised organelles in ciliates, parabasalids, chytrids, microsporidia, Cryptosporidium, Entamoeba, and Blastocystis are discussed in detail in previous chapters of this book. The MLOs of the remaining taxa are reviewed below.

Entamoeba histolytica Pelomyxa palustris Mastigamoeba balamuthi Mastigamoeba punctachora Mastigamoeba simplex Mastigella commutans Neocallimastix

Entamoebidae Pelobionts

Chytridiomycetes

Microsporidia Metamonada

Amoebozoa

Opisthokonta

Excavata

EM, protein localisation EM EM EM EM EM EM, protein localisation, biochemistry EM, biochemistry

Evidence

Trachipleistophora hominis EM, protein localisation Giardia intestinalis EM, protein localisation, biochemistry Spironucleus vortens EM Spironucleus elegans EM Spironucleus barkhanus mitochondrial genes in genome Parabasalia – many species EM, protein localisation, biochemistry Carpediemonas membraEM nifera Dysnectes brevis EM Trimastix marina EM

Piromyces

Taxon bearing MLO

Higher ranks

See Henze, Tachezy; Müller, Hrdý, this volume Simpson and Patterson 1999

–

2E

2C, 3C

Yubuki et al. 2007 Simpson et al. 2000

Sterud and Poynton 2002 Brugerolle et al. 1973 Horner and Embley 2001

– 2F –

– –

–

See Tachezy, this volume Seravin and Goodkov 1987 Chavez et al. 1986 Walker et al. 2001 Walker et al. 2001 Walker et al. 2001 See Tielens, Hellemond, Voncken, Hackstein, this volume See Tielens, Hellemond, Voncken, Hackstein, this volume See Tachezy, this volume See Tachezy, this volume

Refs.

– – 3A,B 2A,B – – –

Figure

Table 1 Taxa containing anaerobic mitochondria-like organelles (MLOs) with the brief summary of the information on MLOs and reference to a ﬁgure or original literature. EM – electron microscopy

268 V. Hampl · A.G.B. Simpson

Jakobida Heterolobosea

Incertae sedis

Stramenopila

Euglenozoa Chromalveolata Apicomplexa Ciliophora

Higher ranks

Table 1 (continued)

EM EM

Chilomastix cuspidata

Saccinobaculus doroaxostylus Andalucia incarcerata Psalteriomonas lanterna

Breviata anathema

Blastocystis hominis

Monopylocystis visvesvarai Sawyeria marylandensis Vahlkamﬁa anaerobica Percolomonas descissus Postgaardi mariagerensis Cryptosporidium Many species

Lyromonas vulgaris

EM, mitochondrial genes among ESTs

Trimastix pyriformis

EM EM, predicted protein targeting EM, localization of hydrogenase EM EM EM EM EM EM, protein localisation EM, sequence of genome, biochemistry EM, membrane potential, presence of genome EM

Evidence

Taxon bearing MLO

2L, 3K,L

–

–

– 2H – 2I 2J, 3I

–

2K, 3J 3G,H

3F

2G, 3E

2D, 3D

Figure

Walker et al. 2006

See Yarlett, Tan, this volume

O’Kelly et al. 2003 O’Kelly et al. 2003 Smirnov and Fenchel 1996 Brugerolle and Simpson 2004 Simpson et al. 1997 See Keithly, this volume See Hackstein, this volume

Broers et al. 1993

Simpson and Patterson 2001 Broers et al. 1990

O’Kelly et al. 1999, Brugerolle and Patterson 1997, Hampl et al., unpublished Weerakoon et al. 1999, this chapter Carpenter et al. unpublished

Refs.

Possible Mitochondria-Related Organelles in “Amitochondriate” Eukaryotes 269

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2 Pelobionts Pelobionts – a group of amoeboid organisms that usually also have a single ﬂagellum (Fig. 2A,B) are classiﬁed within supergroup Amoebozoa based on relatively robust molecular evidence (Milyutina et al. 2001; Bapteste et al. 2002; Cavalier-Smith et al. 2004; Nikolaev et al. 2006). Similarly to their closest relatives – entamoebids – no classical mitochondria have been observed in these species. Electron-dense organelles with two surrounding membranes interpreted as mitochondrial homologues, have been reported from Mastigamoeba balamuthi (basionym Phreatamoeba balamuthi) (organellar dimensions 300 × 500 nm) (Fig. 3A,B), Mastigamoeba punctachora (dimensions 200 × 500 nm), Mastigamoeba simplex (dimensions 200 × 500 nm), Mastigella commutans (round, 200 nm in diameter) and Pelomyxa palustris (round to oval 300–500 nm in the longest dimension) (Chavez et al. 1986; Seravin and Goodkov 1987; Walker et al. 2001; Simpson and Roger 2004b). An EST project on Mastigamoeba balamuthi has revealed genes coding for 23 proteins that have homologues which function in the mitochondria of other eukaryotes (Gill et al. 2007, unpublished). These proteins are involved mainly in the tricarboxylic acid cycle, amino acid metabolism and protein import. Six of these genes contained a predicted mitochondrial targeting peptide and one (cpn 60) has been experimentally proven to localise to the organelles. Genes coding for enzymes typical for anaerobic and hydrogenosomal metabolism (pyruvate:ferredoxin oxidoreductase, [FeFe] hydrogenase and ferredoxin) were also detected, although only ferredoxin possessed a predicted mitochondrial targeting peptide. Judging from the EST data, the organelle of Mastigamoeba is metabolically more active than the mitosome of the relatively closely related parasite Entamoeba histolytica (see Tachezy, this volume). At present, the functions of the MLOs of other pelobionts are not understood. Some inferences have been made from the presence of endobiotic bacteria that were found in the cytoplasm of Pelomyxa palustris, Mastigella commutans and Mastigella sp. (van Bruggen et al. 1985, 1988; Walker et al. 2001). All three species contained bacilliform archaea ascribed to the species Methanobacterium formicicum and a second uncharacterised methanogen was observed in Pelomyxa. Indeed, methane production was detected in cells of Pelomyxa palustris (van Bruggen et al. 1988). The presence of methanogens suggests that the organelles may produce H2 , a substrate for methane synthesis. Nonetheless, in addition to methanogens, both Pelomyxa and Mastigella sp. were found to harbour apparently the same species of thick non-methanogenic Gram-positive rod, which was closely associated with the nucleus (van Bruggen et al. 1985, 1988). Van Bruggen et al. (1985, 1988), who could not detect any microbodies in Pelomyxa, hypothesised that the thick rod might substitute for the function of hydrogenosomes and serve as a connection between catabolism of the amoeba and hydrogen supply for the methanogens.

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Fig. 2 (A–B) Mastigamoeba punctachora, from Bernard et al. (2000); A non-ﬂagellated multinucleated cell; B ﬂagellate cell; C Carpediemonas membranifera, from Bernard et al. (2000); D Trimastix pyriformis, from Bernard et al. (2000); E Trimastix marina, from Bernard et al. (2000); F Spironucleus elegans as an example of the genus, redrawn from Brugerolle et al. (1973); G Chilomastix cuspidata, from Bernard et al. (2000); H Sawyeria marylandensis redrawn from O’Kelly et al. (2003); I Percolomonas descissus, from Bernard et al. (2000); J Postgaardi mariagerensis; K Andalucia incarcerata, from Bernard et al. (2000); L Breviata anathema, redrawn from Walker et al. (2006). Scale bar represents 5 µm (approx.) for (F) and 10 µm (approx.) for all other drawings

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3 Metamonada Diplomonads, parabasalids, retortamonads, oxymonads, Trimastix, Carpediemonas and Dysnectes are ﬂagellates placed in the supergroup Excavata. Trimastix, Carpediemonas, Dysnectes, some diplomonads, a couple of parabasalids and a single retortamonad species are free-living bacterivorous organisms (see Bernard et al. 2000). The remainder are various forms of commensals or parasites. They may constitute a monophyletic group, which is now equated with the taxon name Metamonada (Cavalier-Smith 2003; Hampl et al. 2005), but their monophyly needs further study, because available molecular data are not conclusive (Hampl et al. 2005; Simpson et al. 2006). There is, however, sound evidence that Carpediemonas, Dysnectes and retortamonads are closely related to diplomonads (Silberman et al. 2002; Simpson et al. 2002; Yubuki et al. 2007) and that parabasalids constitute a sister clade to these three groups (Henze et al. 2001; Andersson et al. 2005; Hampl et al. 2005, Simpson et al. 2006). It is also clear that Trimastix is closely related to oxymonads (Dacks et al. 2001; Simpson et al. 2006). One common feature of all these taxa is the absence of aerobic mitochondria. The hydrogenosomes of parabasalids and mitosomes of the diplomonad Giardia intestinalis were discussed in detail earlier in this book (see Müller; Dyall and Doleˇzal; Benchimol; Müller and Hrdý; Henze and Tachezy; Kulda and Hrdý; Tachezy, this volume). Double membrane-bounded electron-dense organelles without cristae have also been observed in Carpediemonas membranifera (Simpson and Patterson 1999), Dysnectes brevis (Yubuki et al. 2007) and Trimastix spp. (Brugerolle and Patterson 1997; O’Kelly et al. 1999; Simpson et al. 2000) that were isolated from marine and freshwater anoxic habitats. The examination of published micrographs suggests that similar organelles may also be present in the parasitic diplomonad taxon Spironucleus (Brugerolle et al. 1973; Sterud and Poynton 2002) and in the free-living retortamonad Chilomastix cuspidata (Weerakoon et al. 1999).

Fig. 3 Electron microscope images of some mitochondria-like organelles (MLOs) of protists discussed in this chapter. (A,B) Mastigamoeba balamuthi, courtesy of Jan Tachezy and Iva Dyková; A MLOs in the cytosol around a nucleus; B detail; C Carpediemonas membranifera; D Trimastix pyriformis, courtesy of Guy Brugerolle; E Chilomastix cuspidata, (original Catherine Bernard and Alastair Simpson); F Saccinobaculus doroaxostylus, courtesy of Patrick Keeling; G,H Psalteriomonas lanterna reprinted from Broers et al. (1990) with permission from Elsevier; G cytosolic organelle surrounded by endoplasmic reticulum; H organelles (dark) and endosymbionts (light rods) in the globule; I Postgaardi mariagerensis, organelle located at the periphery of the cell in association with epibiotic bacteria; J Andalucia incarcerata, courtesy of Laura Hug and Andrew Roger; K,L Breviata anathema, courtesy of Giselle Walker; K position of organelle in the cell, L detail. Scale bars represent 2 µm in (A) and (H), 200 nm in all other ﬁgures

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Carpediemonas membranifera contains narrow but elongated (ca. 450 nm) mitochondria-like organelles that are situated posterior to the nucleus (Simpson and Patterson 1999) (Fig. 3C). Oval organelles (ca. 350 nm in diameter) were observed around the nucleus in Dysnectes brevis (Yubuki et al. 2007). Organelles of Trimastix pyriformis (syn. Trimastix convexa) and Trimastix marina are less conspicuous, smaller, round (200 nm wide, 300 nm long) and number approximately 20 per cell (Fig. 3D). They are dispersed throughout the cell and often associated with endoplasmic reticulum (Brugerolle and Patterson 1997; O’Kelly et al. 1999; Simpson et al. 2000). Their biochemical properties are unknown, but an EST project on Trimastix pyriformis has revealed genes coding for proteins that have homologues that function in the mitochondria of other eukaryotes: the tricarboxylic acid cycle enzyme aconitase, pyridine nucleotide transhydrogenase, lipoyltransferase, the complete glycine cleavage system involved in amino acid metabolism, proteins involved in protein transport and maturation (homologues of TOM40, cnp60 and mitochondrial processing peptidase) and three types of molecular carriers (Hampl et al. 2007, unpublished). Experimental evidence for cell localisation of protein products of all these genes is absent but N-terminal pre-sequences (potential mitochondrial targeting peptides) were found on four of these proteins. At the same time, genes coding for enzymes typical for anaerobic and hydrogenosomal metabolism (pyruvate:ferredoxin oxidoreductase, [FeFe] hydrogenase and [FeFe] hydrogenase maturases) were detected but their cell localisation is unknown (Hampl et al. 2007, unpublished). Spironucleus is a member of the hexamitinid clade of diplomonads that is clearly separated from the giardiinid clade (Kolisko et al. 2005; Keeling and Brugerolle 2006; Jørgensen and Sterud 2007). Sterud and Poynton (2002) reported dark membrane-bounded bodies located close to the nuclei in Spironucleus vortens. The bodies are elongated (up to 400 nm in length) and possibly enveloped in a double membrane. Similar bodies are visible in electron micrographs of Spironucleus elegans (Fig. 7 in Brugerolle et al. 1973). The gene for chaperonin cpn60, which is speciﬁcally associated with mitochondrial organelles was found among ESTs from Spironucleus barkhanus (Horner and Embley 2001). The gene contains a putative mitochondrial targeting sequence and is speciﬁcally related to the (mitosome-targeted) homologue in Giardia intestinalis; its cell localisation, however, has not been directly investigated. An elongated (500 nm long) dense structure is shown, without any comment, in an electron microscopic image of the free-living retortamonad Chilomastix cuspidata (Weerakoon et al. 1999). Re-examination of this organelle suggests that it is bounded by two closely adpressed membranes (Fig. 3E). No mitochondria-like organelles were reported from Retortamonas (the only other genus of retortamonads), but a gene for the enzyme [FeFe] hydrogenase was acquired from Retortamonas sp. (Hampl et al. 2007, unpublished).

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The presence of this gene, however, does not itself imply the existence of hydrogenosomes, because the [FeFe] hydrogenase enzyme could function in the cytoplasm, as in Giardia and Entamoeba (Townson et al. 1996; Rodriguez et al. 1998). Oxymonads are a small group of insect and vertebrate gut endobionts. Bloodgood et al. (1974) reported large dense cytoplasmic bodies in the oxymonad Pyrsonympha. The authors consider them to be neither hydrogenosomes nor other derivates of mitochondria. We are also sceptical of the possibility that these structures could represent MLOs and instead consider them more likely to be endosymbiotic or engulfed bacteria. Carpenter et al. (2007, unpublished) observed membrane-bounded, rounded, electron-dense bodies (0.5–1.5 µm) in Saccinobaculus doroaxostylus (Fig. 3F); some micrographs suggest that this body may be bounded by two membranes. Genes for [FeFe] hydrogenase and pyruvate:ferredoxin oxidoreductase enzymes were found in an EST project carried out on the oxymonad Monocercomonoides (Hampl and Dacks 2007, unpublished), but the cell localization of their products is not known.

4 Heterolobosea The majority of amoebae and amoeboﬂagelates from the group Heterolobosea (e.g. Naegleria, Tetramitus) contain functional aerobic mitochondria. So far ﬁve species have been described that inhabit anaerobic environments and contain organelles interpreted as hydrogenosomes: Psalteriomonas lanterna, Lyromonas vulgaris (basionym Psalteriomonas vulgaris), Vahlkampﬁa anaerobica, Sawyeria marylandensis, and Monopylocystis visvesvarai. 18S rRNA gene phylogenies indicate that the hydrogenosome-bearing heterolobosean species might have evolved from a common ancestor (O’Kelly et al. 2003). Psalteriomonas lanterna, Monopylocystis visvesvarai and Sawyeria marylandensis form a robustly supported clade in phylogenetic trees, while sequences of Vahlkampﬁa anaerobica and Lyromonas are not available yet. The poorly studied heterolobosean ﬂagellate Percolomonas descissus also lives in lowoxygen environments and has mitochondrial organelles that seemingly lack cristae, but available micrographs of the organelles are of mediocre quality (Brugerolle and Simpson 2004; Simpson 2007, unpublished). The amoeboﬂagelate Psalteriomonas lanterna was isolated from the anoxic sediment of a sewage water treatment plant (Broers et al. 1990). In the ﬂagellate stage, disc- and sausage- shaped electron-dense hydrogenosome-like organelles (1–1.5 µm in length) are stacked to form a large “globule” in the centre of the cell (Fig. 3H). The globule is not enfolded by a membrane, and besides the organelles it also contains rod-shaped methanogenic prokaryotic symbionts – probably Methanobacterium formicicum (Broers et al. 1990,

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1992b). The methanogens apparently consume hydrogen produced by the organelle and so maintain a low H2 tension in the globule that theoretically might prevent inhibition of organellar metabolism by accumulation of the end product. An increase of O2 in the culture gas phase results in the loss of the endosymbionts and also triggers the transformation of ﬂagellates to amoebae (Broers et al. 1992a). No endosymbionts were reported from the amoeba stage. Both ﬂagellates and amoebae can grow without endosymbionts, but not in strictly anaerobic conditions (Broers et al. 1992a). Another morphotype of elongated hydrogenosome-like organelles (approx. 1 µm in length) is spread through the cytoplasm of both ﬂagellate and amoeboid stages (Fig. 3G). Like the mitochondria of many Heterolobosea (Page and Blanton 1985), these organelles are often surrounded by endoplasmic reticulum. The cytoplasmic organelles are interpreted either as a distinct type of mitochondria-like organelles (Broers et al. 1990, 1992b) or as immature stages of the hydrogenosomes from the globule (Hackstein et al. 2001). The absence of a fully functional respiration chain was conﬁrmed by negative tests for cytochrome oxidase activity (Brul et al. 1994). A sequence of [2Fe-2S] ferredoxin (Brul et al. 1994) and an incomplete sequence of [FeFe] hydrogenase (Voncken et al. 2002) have been reported. The cellular localization of protein products is unknown, however, the ferredoxin N-terminal sequence is strikingly similar to the hydrogenosomal targeting sequence of the Trichomonas vaginalis ferredoxin and suggests targeting to the organelle (Brul et al. 1994). The ﬂagellate Lyromonas vulgaris (basionym Psalteriomonas vulgaris) was isolated from anoxic sediments. A globule aggregate similar to that of Psalteriomonas lanterna was observed in the cell, situated posterior to the nucleus (Broers et al. 1993). It consisted of elongated organelles (1–3 µm long) and the endosymbionts Methanobacterium formicicum. Lyromonas vulgaris was also able to grow without endosymbionts, but not in strictly anaerobic conditions. Unlike P. lanterna, no cytoplasmic organelles enfolded by endoplasmic reticulum were reported. The presence of hydrogenase in the organelles of the globule was shown by cross-reaction with polyclonal hydrogenosomal hydrogenase antiserum (Broers et al. 1993). Monopylocystis visvesvarai and Sawyeria marylandensis were isolated from salt marsh and iron rich creek sediments respectively (O’Kelly et al. 2003). Flagellate stages are unknown. Both contain hydrogenosome-like organelles that are spherical-to-bacilliform (approx. 800 nm long) in Monopylocystis and bacilliform (approx. 2 µm long) in Sawyeria. The exact number of enveloping membranes was impossible to conﬁrm. The organelles have no detectable cristae and a homogeneous (Monopylocystis) or marbled (Sawyeria) matrix when observed by transmission electron microscopy (O’Kelly et al. 2003). The amoeba Vahlkampﬁa anaerobica was isolated from anaerobic marine sediments and grows under strict anaerobiosis, but could possibly tol-

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erate microaerobic conditions (Smirnov and Fenchel 1996). The cytoplasm contains rounded or elongated microbodies with a maximal dimension of 1.5 µm. They seem to be bounded by double membranes, have no cristae and are not associated with endoplasmic reticulum. They are not associated with bacteria that are present freely in the cytoplasm (Smirnov and Fenchel 1996).

5 Postgaardi mariagerensis Postgaardi mariagerensis is an euglenozoan that has been observed in anoxic regions of saline lakes and fjords (Fenchel et al. 1995; Simpson et al. 1997) and salt marsh organic detritus (Simpson, personal observation). The protists were restricted to the anoxic part of the water column in water rich in sulphide (Fenchel et al. 1995; Simpson et al. 1997). Longitudinally arranged rod-shaped epibiotic bacteria cover virtually the whole surface of the cell. From analogy with the epibionts of certain anaerobic ciliates, it is speculated that the bacteria are sulfate reducers, but this has yet to be conﬁrmed by biochemical analysis. Several mitochondria-like organelles are located under the cell surface of Postgaardi, have tubular proﬁles that are over 2 µm in length and run longitudinally, close and parallel to the epibiotic bacteria (Fig. 3I). The organelles are bounded by double membranes and contain a homogeneous matrix; no cristae, nucleoid or other inclusions were observed. To our knowledge, Postgaardi has not been cultured.

6 Andalucia incarcerata Andalucia incarcerata (basionym Jakoba incarcerata) in an excavate ﬂagellate isolated from marine sediments. Morphologically Andalucia resembles jakobids, but a speciﬁc afﬁnity with jakobids has not been proven by molecular studies published to date (Edgcomb et al. 2001; Simpson et al. 2002; Lara et al. 2006). Andalucia incarcerata grows under low oxygen conditions. Behnke et al. (2006) found ﬁve environmental clones of the 18S rRNA gene restricted to anoxic marine water layers that were closely related to Andalucia incarcerata. The cell contains elongated mitochondria-like organelles (0.5 × 2.5 µm) with two bounding membranes, which lie mostly in the vicinity of the nucleus (Fig. 3J). The organellar matrix is of medium density and contains no cristae (Simpson and Patterson 2001). Interestingly, its only relative in culture at present, Andalucia godoyi, is an aerobe and contains a mitochondrion with tubular cristae (Lara et al. 2006).

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7 Breviata anathema Breviata anathema, formerly studied under the name Mastigamoeba invertens, represents a lineage with an unclear position in the eukaryotic tree. Its former identiﬁcation as a pelobiont was shown to be incorrect on the basis of ultrastructural data (Walker et al. 2006) and gene phylogenies (e.g. Dacks et al. 2002; Edgcomb et al. 2002; Walker et al. 2006). The closest relatives of Breviata in SSU rRNA gene trees are two environmental sequences from anoxic sediments (Dawson and Pace 2002; Walker et al. 2006). Candidate sister taxa of this clade are apusomonads and Excavata, but these possibilities are very poorly supported (see discussion in Walker et al. 2006). The ﬂagellated form of Breviata contains one large (4–5 µm long), branched and double membrane-bounded organelle (Fig. 3K–L). It surrounds the nucleus from the anterior side and is visible by light microscopy. A denser area in an otherwise homogeneous matrix was observed in the most anterior part of the anterior edge, close to the ﬂagellar basal bodies. No cristae were observed. There are no data on the biochemistry of the organelle, but its location near the endocytotically active area of the cell suggests that it might be involved in energy generation (Walker et al. 2006). Taxa reviewed in this chapter promise to contain MLOs with so far undescribed functional properties, which may show that the plasticity of these organelles is even higher than we now appreciate. To understand the functions of these organelles, previous and ongoing EM- and EST-based studies will have to be followed by proteomic and biochemical studies. These will entail methodological challenges as most of these organisms are difﬁcult to handle, and so far impossible to keep in axenic culture. At the same time, further sampling of the biodiversity of anaerobes is required. We expect that a second edition of this book would have separate sections for many of the organisms we have covered, and the analogous chapter to this one would be populated by a very different menagerie of organelles from strange protists that have not yet been discovered. Acknowledgements The authors thank Jan Tachezy, Guy Brugerolle, Laura Hug, Andrew Roger, Giselle Walker and Patrick Keeling for kindly providing electron micrographs and Andrew Roger for helpful comments. The authors are supported by the Canadian Institute for Advanced Research, Programs in Evolutionary Biology and Integrated Microbial Biodiversity.

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Subject Index

2D –, analyses, 173 –, electrophoresis, 166 acetate/succinate CoA transferase (ASCT), 8, 9, 104, 115, 116, 118, 119, 126, 128 acetyl –, CoA, 104, 119, 121 –, CoA hydrolase, 126 –, CoA synthase, 102 adenylate kinase, 119, 127 ADP/ATP carrier, 115, 157, 221 aerobic –, resistance, 185 –, respiration, 256 aerobic eukaryotes, 6 alcohol dehydrogenase E, 151 alpha-ketoglutarate dehydrogenase, 103 alternative oxidase (AOX), 223, 243 amitochondriates, 204, 205 Amoebozoa, 266 Amylovorax, 98 anaerobic –, chytrids, 151 –, eukaryotes, 1, 3, 5–7, 10, 11 –, resistance, 187 Andalucia incarcerata, 277 Antonospora, 32 –, locustae, 44, 211, 219, 256 apicomplexans, 232 apicoplast, 233 apoptotic body, 258 Arabidopsis, 164 archaebacteria, 5, 239 Archaeplastida, 266 Archezoa, 204 arginine, 135 –, biosynthesis, 165 –, deiminase, 136

–, dihydrolase pathway, 136 articulin, 245 ATP:AMP phosphotransferase, 127 ATP –, production, 165 –, synthase, 223 –, synthesis, 2, 8–10, 12 ATP/ADP translocase, 164 autophagy, 88 axostyle, 76, 78, 93 Blastocystis, 256 bongkrekic acid, 115 Bradyrhizobium, 11 Breviata anathema, 278 butyryl-CoA, 104 calcium, 81, 86 Canﬁeld oceans, 3, 7, 9–11, 13, 14 carbamate kinase, 136 cardiolipin, 115, 214 Carpediemonas membranifera, 274 catalase, 129 cellulose, 151 chaperone, 58, 157, 216 chaperonin 60, 205 Chilomastix cuspidata, 274 Chlamydomonas reinhardtii, 9 Chromalveolata, 266 chytridiomycete, 10, 148 ciliates, 76, 78 citrulline, 136 class A ﬂavodiiron protein, 131 cleavage site, 41, 56 CO2 , 115, 118, 128 coenzyme A (CoA), 115, 126, 128 coenzyme Q, 125 complex I, 103, 120, 125, 165 complex II, 165

284 confocal, 240 costa, 76, 78, 93 Cpn10, 43 Cpn60, 42, 43, 232 cristae, 236, 258 cryo-electron tomography, 239 crypton, 203 Cryptosporidia, 256 Cryptosporidium, 51, 54, 57, 58, 204, 213 –, parvum, 32, 121, 124, 174, 205, 207, 219 –, sp., 23 crystalline core, 84 cyanide, 128 Cyclidium porcatum, 80 cyclosporine A, 259 cysteine desulfurase (IscS), 43, 132, 133, 205, 215, 217, 219 cytochalasin, 78, 87 cytochrome, 127, 261 –, P450 reductase, 121, 124 –, c oxidase, 209 Dasytricha, 98, 103 Desulfovibrio vulgaris, 131 Dictyostelium discoideum, 212 dihydrofolate reductase, 135 diplomonads, 174 Diploplastron afﬁne, 103, 107 division, 86, 88, 91, 212 dynamin, 92 electron –, transfer, 164 –, transport, 122, 165, 220, 221 Encephalitozoon cuniculi, 2, 32, 44, 54, 57, 58, 173, 211, 256 endobiotic bacteria, 270, 275 endoplasmic reticulum, 24, 87, 88 endosymbiont, 1, 4, 5, 13, 106 endosymbiotic theory, 4–6 Entamoeba, 2, 10, 32, 35, 42, 54, 57, 132, 220, 256 –, histolytica, 23, 122, 174, 204, 206 Entodinium simplex, 103, 107 epibiotic bacteria, 277 Epidinium, 98, 103 Eremoplastron bovis, 103, 107 ethanol, 9, 151, 191 Eudiplodinium, 98, 103

Subject Index Euglena gracilis, 8, 10, 121, 124, 164, 174, 260 eukaryote evolution, 1, 3, 7, 13 eukaryotic supergroups, 266 Euplotes, 102 evolution, 22, 107, 174, 202 Excavata, 213, 266 Fo F1 -ATP synthase, 103, 120, 127 Fasciola hepatica, 260 fatty acids, 261 [FeFe] hydrogenase, 123, 124 ferredoxin, 31, 35, 43, 118–122, 125, 132, 180, 187, 189, 217, 219, 222 –, genes, 194 ferritin, 223 FeS (see iron-sulfur) ﬂavodiiron protein, 131, 186 ﬂavoprotein, 129 FMN, 125, 131 formate, 9, 152 frataxin, 132, 133, 216, 218, 219 freeze-fracture, 79, 81, 83 FtsZ, 92, 212 Fugu, 164 fumarase, 260, 262 fumarate, 107 –, hydratase, 260, 262 –, reductase, 8 fungi, 76, 213 futile cycling, 185 geological model, 6 Giardia intestinalis, 10, 23, 31, 35, 43, 54, 57, 122, 132, 204, 212, 213, 217, 256 glucose, 101, 103 glutaredoxin, 216, 217 glycerol-3-phosphate dehydrogenase (G3PDH), 57, 222 glycine, 135 –, decarboxylase complex (GDC), 134, 135 glycolysis, 116, 202, 240 H cluster, 123, 124 H protein, 134 heat shock protein 70 (see Hsp70) heme, 133 Heterolobosea, 275 hidden Markov model, 33, 173 Hmp31, 45, 115, 137

Subject Index Hmp35, 46, 115, 137 Hsp70, 43, 54, 70, 205, 232 HydE, 124, 137 HydF, 124 HydG, 124 hydrogen, 76, 115, 153, 166, 256 –, peroxide, 128, 129, 175 hydrogenase, 107, 119, 121, 123, 174, 189, 214 –, maturase, 124, 174 hydrogenosomal –, targeting motif, 137 –, terminal oxidase, 186 hydrogenosome, 23, 30, 35, 44, 76, 181, 203, 206 hydroxyurea, 78 Hymenolepis diminuta, 221 immunolabeling, 93 import, 22 –, machinery, 173 –, motor, 53 inner membrane, 232 inner membrane protease, 26 intestinal tract, 202 invasion, 246 invertebrates, 8, 9, 12 iron storage, 245 iron–sulfur (FeS), 113, 116, 119, 122, 123, 125, 129, 131, 137 –, cluster (ISC), 121, 132, 133, 215, 237 –, cluster assembly, 11, 215 –, protein, 133 IscA, 132 IscS (see cysteine desulfurase) IscU, 132, 215 Isd11, 218 isopycnic centrifugation, 138 Isotricha, 98, 103 Janus green, 258 karyokinesis, 93 2-keto acid oxidoreductase, 116, 121, 193 Krebs cycle, 119, 260, 262 L protein, 134 lactate, 9, 191 –, dehydrogenase (LDH), 128, 191, 243 latency, 125

285 lateral gene transfer, 2 Leishmania, 260 Lyromonas vulgaris, 276 lysosome, 88, 136, 138 malate, 116, 128 malic enzyme, 116, 119, 124, 125, 136, 138, 189 mass spectrometry, 166 Mastigamoeba –, balamuthi, 132, 206, 220, 270 –, punctachora, 270 –, simplex, 270 Mastigella commutans, 270 matrix, 81, 84 membrane, 79, 114, 115, 125 –, potential, 220, 244 Metamonada, 273 Methanobacterium formicicum, 106, 270, 275, 276 methanogenic symbiont, 106 Methanoplanus endosymbiosus, 106 methyl viologen, 119 methylmalonyl-CoA, 8 Metopus, 98 –, contortus, 80, 106 –, striatus, 106 metronidazole, 79, 116, 118, 122, 180, 257 –, anion radicals, 181, 185, 189 –, resistance, 184 microaerophile, 128 microsporidia, 23, 32, 44, 57, 204, 207 microtubule, 93 mitochondrion, 24 mitochondrion-like organelle, 232 MitoLight, 258 mitosis, 93 mitosome, 23, 30, 42–44, 116, 203, 204 MitoTracker Green, 258 Monocercomonas, 114 –, sp., 78 Monocercomonoides, 275 Monopylocystis visvesvarai, 276 Mycoplasma, 4 N 5 ,N 10 -methylene tetrahydrofolate, 135 N-acetyl glucosamine, 83 NAD(P)H nitroreductases, 194 NAD-dependent malic enzyme, 182 NADH

286 –, dehydrogenase, 120, 122, 125, 182 –, oxidase, 128 –, ubiquinone oxidoreductase, 125 NADP-speciﬁc malic enzyme, 116 NADPH oxidase, 128 Neocallimastix, 35, 41, 46, 54, 147, 150, 165, 260 –, frontalis, 78, 86, 91 –, sp., 23, 31 NIF, 215 Nim proteins, 194 nitric oxide, 131 nitro anion radicals, 180 5-nitroimidazole, 179 nitrosative stress, 132 nucleus, 23 Nyctotherus, 54, 57, 58, 98, 104, 175 –, ovalis, 23, 126, 165 ocean geochemistry, 6, 13 Ohr protein, 131 Ophryoscolex caudatus, 103, 107 Opisthokonta, 266 organic hydroperoxide, 131 origin, 164, 173 ornidazole, 180 ornithine, 136 –, decarboxylase, 136 OsmC, 131, 175 Ostracodinium obtusum bilobum, 103, 107 OXA, 210 oxidative –, decarboxylation of malate, 182 –, decarboxylation of pyruvate, 181 –, phosphorylation, 4, 128 oxygen, 3, 5, 6, 127, 128, 202 –, respiration, 4 –, scavenging enzymes, 186 P protein, 134 PAM, 27, 53, 210 Pam18, 43 pelobionts, 270 Pelomyxa palustris, 106, 270 PEP carboxykinase, 116 Percoll, 138 Percolomonas descissus, 275 peripheral vesicles, 81 permeability, 115 peroxide, 130

Subject Index peroxiredoxin, 130, 175 peroxisomes, 4 phosphatidylcholine, 115 phosphatidylethanolamine, 115 phosphatidylserine, 115 phosphoenolpyruvate, 116 phospholipid, 115 photosynthesis, 6 Phytomonas, 260 Piromyces, 9, 147, 150, 175 Plagiopyla, 98 –, nasuta, 106 Plasmodium, 173, 202, 209, 215 –, falciparum, 11, 164, 206 polyamines, 135 Polyplastron, 98 Polytomella, 10 Postgaardi mariagerensis, 277 preprotein, 53 presequence, 29, 34, 40, 42, 210 processing peptidase, 25, 34, 43, 55, 211 programmed cell death, 261 propionyl-CoA, 8 protease, 245 protein –, expression, 190 –, import, 28–30, 164, 238 –, trafﬁcking, 22, 23 Psalteriomonas, 175 –, lanterna, 275 –, vulgaris, 106 putrescine, 136 pyridine –, nucleotide, 115, 119 –, nucleotide transhydrogenase, 205, 221 Pyrsonympha, 275 pyruvate, 115, 116, 128, 260 –, decarboxylase, 192 –, dehydrogenase complex, 101, 121, 222 –, formate lyase, 9, 101, 102, 105, 151, 165 –, kinase, 116 pyruvate:ferredoxin oxidoreductase (PFOR), 10, 101, 102, 104, 118, 119, 121, 129, 153, 165, 174, 180, 189, 214, 260 pyruvate:NADP oxidoreductase (PNO), 121, 124, 240, 260 reactive oxygen species, 129, 137 Reclinomonas americana, 209 reductive evolution, 237

Subject Index relict mitochondrion, 205 respiratory chain, 125, 128, 210 Rhizaria, 266 rhodamine 123, 258 rhodoquinone, 8 Rickettsia, 28 –, prowazekii, 56 Rli1, 239 RNA polymerase II, 205 rotenone, 126, 128 rubredoxin, 131 rubrerythrin, 130, 175 rumen, 148 –, ciliates, 107 Saccharomyces cerevisiae, 164, 215, 219 SAM, 25, 46, 47, 211 Sawyeria marylandensis, 276 Schizosaccharomyces pombe, 164 segmentation, 88, 91 segregation, 212 serine, 135 –, hydroxymethyltransferase (SHMT), 134, 135 signal –, peptidase, 24 –, sequence, 24 small subunit ribosomal RNA, 205 Spironucleus, 132 –, barkhanus, 274 –, elegans, 274 –, salmonicida, 212, 218 –, vortens, 274 subcompartments, 236 substrate-level phosphorylation, 116, 126, 128, 156 succinate, 8, 9, 119, 128, 260 –, CoA ligase, 126 –, dehydrogenase, 260, 262 –, thiokinase (STK, see succinyl-CoA synthetase) succinyl –, CoA, 126 –, CoA synthetase (SCS), 10, 103, 119, 126, 260, 262 succinyl-CoA:acetate CoA transferase, 126 sucrose gradient, 138 SUF, 215 sulfate, 6, 11

287 sulﬁde, 6, 11 –, quinone oxidoreductase, 12 superoxide dismutase (SOD), 129, 186, 222 symbiosis, 4 T protein, 134 targeting –, sequence, 124, 126, 130, 166 –, signal, 34, 116 terminal oxidase, 103 tetrahydrofolate, 135 Tetrahymena, 102 thiol-dependent peroxidase, 130 thioredoxin, 130, 175 –, reductase, 130, 131 TIM, 27, 46, 50–52, 210, 238 tinidazole, 180 Tiny TIMS, 238 TOM, 25, 47, 49, 210, 238 tomography, 232 Toxoplasma gondii, 42, 213 Trachipleistophora hominis, 32, 54, 206 translocase, 25, 34, 174, 211 translocation, 25 transporters, 173 treatment failure, 185 tricarboxylic cycle (TCA), 156, 165 Trichomonadida, 76 trichomonads, 78 Trichomonas, 2, 31, 35, 54, 57, 58, 104, 109, 166 –, gallinae, 115 –, vaginalis, 180, 191 Trimastix –, marina, 274 –, pyriformis, 274 Trimyema, 98, 104 Tritrichomonas –, foetus, 84, 114, 123, 166, 180, 191 –, augusta, 91 Trypanaosoma –, brucei, 41, 202, 212, 216, 260 –, cruzi, 43 ubiquinone, 8, 115, 120 Vahlkampﬁa anaerobica, 276 valyl-tRNA synthetase, 205 vertebrates, 8, 11, 12