Mitochondrial Dynamics and Neurodegeneration

Mitochondrial Dynamics and Neurodegeneration

Bingwei Lu Editor

Mitochondrial Dynamics and Neurodegeneration

Editor Bingwei Lu Department of Pathology Stanford University School of Medicine 825 Allardice Way Stanford, CA 94305 USA [email protected]

ISBN 978-94-007-1290-4 e-ISBN 978-94-007-1291-1 DOI 10.1007/978-94-007-1291-1 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2011929228 © Springer Science+Business Media B.V. 2011 No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

Ever since the ancient bacteria entered a primitive eukaryotic host cell billions of years ago, the proto-mitochondrion and its host cells have entered into an intricate symbiosis relationship that has been constantly evolving. Mitochondria and their host cells depend on each other for maintenance and survival. For example, the majority of the components of the mitochondrial proteome are encoded by host nuclear DNA. On the other hand, mitochondria are essential for such diverse functions as energy metabolism, reactive oxygen species production, calcium buffering, and cell death control. As the name implied (mito and chondrion means threaded and grain, respectively, in Greek), the morphology of mitochondrion is dynamic, constantly undergoing fission and fusion processes in vivo, and these morphological changes are believed to be important for the essential mitochondrial functions mentioned above. Given the critical roles of mitochondria for the normal physiology and life and death decisions of its host cells throughout the body of a multicellular organism, it is not surprising that defects in mitochondrial dynamics are linked to human diseases. What is surprising is the relatively selective vulnerability of neurons to defective mitochondrial dynamics. This has fueled considerable efforts in neurodegenerative disease field to use mitochondrial dynamics as a new experimental paradigm for disease mechanism investigation. Although many reviews and books have been written on the contribution of mitochondrial dysfunction to human diseases, this is the first book that integrates diverse topics such as the genetic control of mitochondrial dynamics, the relationship between mitochondrial dynamics and bioenergetics, the roles of mitochondrial dynamics in apoptosis, axonal transport, mitochondrial quality control, and the contribution of defective mitochondrial dynamics to the pathogenesis of neuro­ degenerative diseases. Focusing on the theme of mitochondrial dynamics and its role in neurodegeneration, this book brings together leading scientists and clinicians from around the world to deliver a comprehensive treatise on all aspects of mitochondrial dynamics. In Chap. 1, Dr. McQuibban and his colleagues present a comprehensive historical account of key discoveries in mitochondria research and those that lead to the identification of key molecular players involved in mitochondrial dynamics. They especially emphasize the role played by model organism genetics in these latter discoveries. In Chap. 2, Dr. Rossignol and colleagues describe the funda­ mental mechanisms by which mitochondrial dynamics and cellular energy status v

vi

Preface

can reciprocally regulate each other. Many thought-provoking hypotheses are proposed. In Chap. 3, Dr. Shirihai and colleagues discuss the regulation and interdependence of mitochondrial dynamics and mitochondrial autophagy (mitophagy) in mitochondrial quality control. This is a very timely topic as mitophagy is currently one of the heavily studied areas in neurodegenerative disease research. In Chap. 4, Dr. Youle and colleague highlights the importance of mitochondrial dynamics in the timely execution of apoptosis, one of the fundamental processes involved in neurodegeneration. In Chap. 5, Dr. Sheng and colleague discuss the molecular processes that lead to the transport and distribution of mitochondria to neuronal axons and synapses, arguably one of the most important aspects of neuronal physiology. Although disease relevance is not the main focus of this chapter, understanding the relationships among mitochondrial dynamics, transport, and neuronal plasticity will shed important insights into the disease conditions. In Chap. 6, Dr. Reynier and colleagues presents recent findings on the clinical diversity of diseases caused by mutations in the mitochondrial fusion machinery, the spectrum of mutations in the mitochondrial fusion genes and the corresponding pathophysiology, and the therapeutic perspectives. In Chap. 7, Dr. Chu and colleague summarize a rapidly growing body of literature focusing on the role of mitochondrial dynamics in the pathogenesis of Parkinson’s disease, and they discuss unresolved controversies in the field in the context of a dynamic network of compensatory responses to mitochondrial stress, dysfunction and injury. In Chap. 8, Dr. Lipton and colleagues highlight the role of aberrant mitochondrial dynamics in the pathogenesis of Alzheimer’s disease, and offer a molecular mechanism connecting disease-causing insults and post-translational modification of the mitochondrial fission machinery component Drp1. In Chap. 9, Dr. Monteiro and colleagues discuss the published literature linking mitochondrial dysfunction in Huntington’s disease, focusing on the role that defects in mitochondrial dynamics might play in disease pathogenesis. These chapters discuss the central problems associated with the nascent but rapidly growing research field of mitochondrial dynamics in neurodegeneration. They should serve as an excellent reference book for students, clinicians, and researchers engaged or interested in mitochondria biology, neurobiology, and mechanisms of neuro­degenerative diseases. I want to thank all of our authors for their hard work and Thijs van Vlijmen and Sara Huisman of Springer for their excellent help in making this book a reality. Associate Professor Stanford University School of Medicine Stanford, CA 94305 USA

Bingwei Lu, Ph.D.

Contents

1 The Genetics of Mitochondrial Fusion and Fission................................ Eliana Y. L. Chan, Jarungjit Rujiviphat, and G. Angus McQuibban 2 Relationships Between Mitochondrial Dynamics and Bioenergetics....................................................................................... Giovanni Benard, Nadège Bellance, Caroline Jose, and Rodrigue Rossignol 3 Mitochondrial Dynamics and Autophagy................................................ Linsey Stiles, Andrew Ferree, and Orian Shirihai

1

47

69

4 Mitochondrial Dynamics and Apoptosis.................................................. 109 Megan M. Cleland and Richard J. Youle 5 Mitochondrial Dynamics and Axonal Transport.................................... 139 Qian Cai and Zu-Hang Sheng 6 Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery.................................................. 169 Guy Lenaers, Dominique Bonneau, Cécile Delettre, Patrizia Amati-Bonneau, Emmanuelle Sarzi, Dan Miléa, Christophe Verny, Vincent Procaccio, Christian Hamel, and Pascal Reynier 7 Mitochondrial Fission-Fusion and Parkinson’s Disease: A Dynamic Question of Compensatory Networks.................................. 197 Charleen T. Chu and Sarah B. Berman 8 Role of the Mitochondrial Fission Protein Drp1 in Synaptic Damage and Neurodegeneration.............................................................. 215 Tomohiro Nakamura, Dong-Hyung Cho, and Stuart A. Lipton

vii

viii

Contents

9 Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate.......................................................................................... 235 Hongmin Wang, Mariusz Karbowski, and Mervyn J. Monteiro Index.................................................................................................................. 259

Contributors

Patrizia Amati-Bonneau UMR CNRS 6214 - Inserm U771, Angers, France and Département de Biochimie et Génétique, Centre Hospitalier Universitaire, Angers, France Nadège Bellance Laboratoire “MRGM” EA4576 (Maladies Rares: Génétique et Métabolisme), Université de Bordeaux, Bordeaux, France Giovanni Benard Laboratoire “MRGM” EA4576 (Maladies Rares: Génétique et Métabolisme), Université de Bordeaux, Bordeaux, France Sarah B. Berman Department of Neurology, University of Pittsburgh School of Medicine, 3501 Fifth Avenue, BST3, Room 7037, Pittsburgh, PA 15213, USA and The Pittsburgh Institute for Neurodegenerative Diseases, University of Pittsburgh School of Medicine, Pittsburgh, PA 15213, USA Dominique Bonneau UMR CNRS 6214 - Inserm U771, Angers, France and Département de Biochimie et Génétique, Centre Hospitalier Universitaire, Angers, France Qian Cai Synaptic Function Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Building 35, Room 2B-215, 35 Convent Drive, Bethesda, MD 20892–3706, USA Eliana Y.L. Chan Department of Biochemistry, University of Toronto, Toronto, ON, Canada

ix

x

Contributors

Dong-Hyung Cho Graduate School of East-West Medical Science, Kyung Hee University, Yongin, Gyeonggi 446–701, Korea Charleen T. Chu Department of Pathology, University of Pittsburgh School of Medicine, 200 Lothrop St., Pittsburgh, PA 15213, USA [email protected] Megan M. Cleland Biochemistry Section, Surgical Neurology Branch, National Institute of Neurological Disorders and Stroke (NINDS), National Institutes of Health (NIH), 35 Convent Drive, MSC 3704, Building 35/Rm, 2C917, Bethesda, MD 20892, USA Cécile Delettre Inserm U1051, Institut des Neurosciences de Montpellier, Universités de Montpellier I et II, Montpellier, France Andrew Ferree Department of Pharmacology, Boston University School of Medicine, Boston, MA, USA Christian Hamel Inserm U1051, Institut des Neurosciences de Montpellier, Universités de Montpellier I et II, Montpellier, France Caroline Jose Laboratoire “MRGM” EA4576 (Maladies Rares: Génétique et Métabolisme), Université de Bordeaux, Bordeaux, France Mariusz Karbowski Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD 21201, USA and Department of Molecular Biology and Biochemistry, University of Maryland School of Medicine, Baltimore, MD 21201, USA Guy Lenaers Inserm U1051, Institut des Neurosciences de Montpellier, Universités de Montpellier I et II, Montpellier, France [email protected] Stuart A. Lipton Center for Neuroscience, Aging, and Stem Cell Research, Sanford-Burnham Medical Research Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA [email protected]

Contributors

G. Angus McQuibban Department of Biochemistry, University of Toronto, Toronto, ON, Canada [email protected] Dan Miléa UMR CNRS 6214 - Inserm U771, Angers, France and Département d’Ophtalmologie, Centre Hospitalier Universitaire, Angers, France Mervyn J. Monteiro Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD 21201, USA and Department of Anatomy and Neurobiology, University of Maryland School of Medicine, Baltimore, MD 21201, USA [email protected] Tomohiro Nakamura Center for Neuroscience, Aging, and Stem Cell Research, Sanford-Burnham Medical Research Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA Vincent Procaccio UMR CNRS 6214 - Inserm U771, Angers, France and Département de Biochimie et Génétique, Centre Hospitalier Universitaire, Angers, France Pascal Reynier UMR CNRS 6214 - Inserm U771, Angers, France and Département de Biochimie et Génétique, Centre Hospitalier Universitaire, Angers, France Rodrigue Rossignol Laboratoire “MRGM” EA4576 (Maladies Rares: Génétique et Métabolisme), Université de Bordeaux, Bordeaux, France [email protected] Jarungjit Rujiviphat Department of Biochemistry, University of Toronto, Toronto, ON, Canada Emmanuelle Sarzi Inserm U1051, Institut des Neurosciences de Montpellier, Universités de Montpellier I et II, Montpellier, France

xi

xii

Contributors

Zu-Hang Sheng Synaptic Function Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Building 35, Room 2B-215, 35 Convent Drive, Bethesda, MD 20892–3706, USA [email protected]. Orian Shirihai Department of Medicine, Section of Molecular Medicine, Boston University School of Medicine, Boston, MA, USA [email protected] Linsey Stiles Department of Medicine, Section of Molecular Medicine, Boston University School of Medicine, Boston, MA, USA and Department of Pharmacology and Experimental Therapeutics, Tufts University School of Medicine, Boston, MA, USA Christophe Verny UMR CNRS 6214 - Inserm U771, Angers, France and Département de Neurologie, Centre Hospitalier Universitaire, Angers, France Hongmin Wang Division of Basic Biomedical Sciences, University of South Dakota, Vermillion, SD 57069, USA Richard J. Youle Biochemistry Section, Surgical Neurology Branch, National Institute of Neurological Disorders and Stroke (NINDS), National Institutes of Health (NIH), 35 Convent Drive, MSC 3704, Building 35/Rm, 2C917, Bethesda, MD 20892, USA [email protected]

Chapter 1

The Genetics of Mitochondrial Fusion and Fission Eliana Y.L. Chan, Jarungjit Rujiviphat, and G. Angus McQuibban*

Abstract  Mitochondrial research has undergone a true renaissance in the last few years. One can hardly pick up one of the top basic research or clinical research journals without seeing a new discovery about mitochondrial function in cellular/human health and disease. This book describes the most important aspects of mitochondrial function in neurodegeneration. This chapter serves as an introduction to the history of mitochondrial research, and also as an introduction to one of the fundamental cell biological nuances of mitochondrial activity, the gymnastics of double membrane fusion and fission. Here, we will discuss the genetic and biochemical characterization of the molecular machinery that orchestrates these complex processes. Further, we will introduce how these membrane dynamics contribute to both organelle and cellular function, and extend these findings to human disease. Keywords  Mitochondrial fusion and fission • Mitochondrial morphology • Genetic screens • Dynamin-related proteins • Mechanoenzymes • Membrane scission and fusion • Proteolytic processing • Membranes and lipid homeostasis • Mitophagy • Neurodegeneration Abbreviations PA-GFP Fzo EM Mfn1/2 Dnm1 Drp1 Ugo1

photoactivatible green fluorescent protein fuzzy onions electron microscopy mitofusins 1 and 2 dynamin 1 dynamin-related protein 1 ugo is Japanese for fusion

* Eliana Y.L. Chan and Jarungjit Rujiviphat contributed equally to this chapter. E.Y.L. Chan, J. Rujiviphat, and G.A. McQuibban (*) Department of Biochemistry, University of Toronto, Toronto, ON, Canada e-mail: [email protected] B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_1, © Springer Science+Business Media B.V. 2011

1

2

Pcp1 Mgm Mdm10/30/33/36 TetO7 Phb1/2 RNAi dsRNA mtDNA siRNA OPA1 cDNA PARL GTPase GTP GED Vps1 OM IMS IM Fis1 NMR TPR co-IP Mdv1 Caf4 Num1 Gag3 Net2 NTE WD-40 Mff Letm1 CaMKIa GDAP1 CMT Mtp18 MTGM BDLP GDP MTS MPP ATP ADOA TEM

E.Y.L. Chan et al.

processing of cytochrome c peroxidase 1 mitochondrial genome maintenance 1 mitochondrial distribution and morphology 10/30/33/36 tetracycline-regulatable promoter with 7 regulatory units prohibitins 1 and 2 ribonucleic acid interference double-stranded ribonucleic acid mitochondrial deoxyribonucleic acid small interfering ribonucleic acid optic atrophy 1 complementary deoxyribonucleic acid presenilin-associated rhomboid-like guanosine triphosphate hydrolase guanosine triphosphate guanosine triphosphate effector domain vacuolar protein sorting 1 outer membrane intermembrane space inner membrane mitochondrial fission 1 nuclear magnetic resonance tetratricopeptide repeat Co-immunoprecipitation mitochondrial division 1 CCR4 Associated Factor nuclear migration 1 glycerol-adapted growth 3 named after the complex mitochondrial network of the yeast mutant N-terminal extension Beta-transducin repeats mitochondrial fission factor eucine zipper-EF-hand containing transmembrane protein 1 Ca2+/calmodulin-dependent protein kinase Ia Ganglioside-induced differentiation protein 1 Charcot-Marie-Tooth Mitochondrial protein 18 kDa mitochondrial targeting GxxxG motif bacterial dynamin-like protein guanosine diphosphate mitochondrial targeting sequence matrix processing peptidase adenosine triphosphate Autosomal Dominant Optic Atrophy transmission electron microscopy

1  The Genetics of Mitochondrial Fusion and Fission

GTPgS SCF TIM23 PAM MitoPLD CL PA Ups1/2 Gep1 MMM1 CRD1 PSD1 MAPL SUMO1 Bax Bak SENP5 MARCH-V MITOL RING DUBs Cdk1 PKA PP2A ROS ETC BCL2 PINK1 PD

3

guanosine 5¢-O-[gamma-thio]triphosphate Skp Cullin, F-box containing complex translocase of the inner mitochondrial membrane 23 Presequence translocase-associated motor mitochondrial phospholipidase D cardiolipin phosphatidic acid unprocessed 1/2 genetic interactor of prohibitin 1 maintenance of mitochondrial morphology cardiolipin synthase 1 phosphatidylserine decarboxylase 1 mitochondrial-anchored protein ligase small ubiquitin-like modifier 1 Bcl-2-associated X protein Bcl-2-associated K protein ? sentrin/SUMO-specific protease 5 membrane-associated RING-CH-V mitochondrial ubiquitin ligase Really interesting new gene deubiquitinase cyclin-dependent kinase 1 protein kinase A phosphatase 2A reactive oxygen species electron transport chain B-cell leukemia/lymphoma 2 PTEN induced putative kinase 1 Parkinson’s disease

1.1 Mitochondrial Dynamics: 100 years in the Making Since the 1850s, cytologists had observed granular structures in the cytoplasm of living cells. It was in 1857 that Swiss anatomist Rudolph Albert von Kölliker described these granular structures in the sarcoplasm of insect muscle cells. He showed in 1888 that these granules swelled in water and possessed a membrane. Although these structures were named “sarcosomes” in 1890, Carl Benda would introduce the term “mitochondrion” in 1898, derived from the Greek words mitos meaning threaded, and chondron meaning grain. Although the term “mitochondrion” became the most widely accepted name for this organelle, the term “sarcosome” is still used in present day to describe mitochondria in muscle cells. See Fig.  1.1 for a summary and overview of the past 100  years of mitochondrial research and discoveries.

2000

1950

1900

1850

Term “mitochondrion” was introduced First mitochondrial stains were introduced.

First successful mitochondrial purification

High-resolution electron micrographs. Mitochondria were revealed to consist of double membrane and that the inner membrane consisted of transversely oriented ridges.

1898

1948

1952

Present

to

2002

1997 2000 2001

1988

1964

1960

Mitochondria are recognized as dynamic organelles. Revealing players in mitochondrial fusion and fission. Understanding the mechanisms and regulations.

Proper mitochondrial fusion and fission were first shown to be crucial for the transmission of parental mitochondria to zygotes.

Term "Cristae" was introduced.

Named “sarcosomes”

1890

1913

First observed and described as granular structures in sarcoplasm of insect muscle cells

1857

Roles in cell cycle and cellular development. Functions in signalling including apoptotic pathway. Importance of mitophagy and its link to apoptosis.

Mitochondria were first shown to play a role in apoptosis.

Purified mitochondria were shown to contain enzymes for TCA cycle, fatty acid oxidation, and the respiratory assembly. The respiratory system was shown to be present in mitochondria from all cell types. Role in maintain ion gradient was first shown.

1948-1951

First discovery that respiration was associated with the granular structure.

More mitochondria-related genes identified in neurodegenerative diseases. Pathophysiological in-

Mutations in genes required for mitochondrial fusion and fission were identified in patients with neurodegenerative diseases.

Diseases associated with mutations in mitochondrial DNA were first identified.

The presence of mitochondrial DNA was confirmed.

Noticed fibers within the mitochondria were sensitive to nucleic acid stains.

4 E.Y.L. Chan et al.

Fig. 1.1  The past 100 years of mitochondria: Mitochondrial studies began over 100 years ago. The discoveries are categorized as findings in mitochondrial structure (beige), mitochondrial function (green) and mitochondrial DNA and related diseases (blue). It has been more than 100  years in attempting and ­refining the visualization of mitochondria. Advances in the successful purification of mitochondria greatly accelerated the discoveries of mitochondrial functions. Around the same time, the presence of mitochondrial DNA was revealed. Subsequently, many studies uncovered mitochondria-related diseases. In the past decade (grey background), tremendous progress has been made in recognizing mitochondria as dynamic organelles, studying its roles in several diverse cellular processes and discovering mitochondria-related genes that are linked to neurodegenerative diseases

1  The Genetics of Mitochondrial Fusion and Fission 5

6

E.Y.L. Chan et al.

Identification of mitochondria in the early days were challenging since there were many granular structures in the cell. The first mitochondrial stains were introduced in 1898 by Benda and Michaelis. Benda introduced crystal violet, and Michaelis introduced Janus green, an oxidation-reduction dye. In 1908, based on their response to different stains, Regaud concluded that mitochondria contain phospholipid and protein, easing the identification of mitochondria in cell types other than muscle. Since the early 1930s, researchers had attempted on many occasions to isolate intact mitochondria. Hogeboom, Schneider and Palade reported the first successful mitochondrial purification in 1948. They used 0.88 M sucrose to disperse rat liver cells, and were able to easily separate nuclei, mitochondria and microsomes by differential centrifugation. More importantly, the isolated mitochondria had a filamentous structure characteristic of those in rat liver cells, indicating that they had purified intact mitochondria (Hogeboom et al. 1947, 1948). Coincidentally, work from other groups had indicated that the oxidation of fatty acids was dependent on certain elements in liver homogenates. In particular, it was imperative that the structure be maintained in this easily damaged element (Munoz and Leloir 1943; Lehninger 1945). The report that intact mitochondria could be purified using 0.88  M sucrose prompted Lehninger and Kennedy to test the ability of isolated mitochondria to oxidize fatty acids. They reported that purified mitochondria could oxidize fatty acids and intermediates of the Krebs cycle at a rate similar to that of intact liver cells. In addition, they also observed that oxidative phosphorylation was concomitant with these oxidations (Kennedy and Lehninger 1948). Over time, work by others confirmed that this respiratory system is present in mitochondria from all cell types examined, regardless of plant or animal origin (Schneider 1948; Lehninger 1949; Kennedy and Lehninger 1949; Potter et  al. 1951). Although Warburg had discovered in 1913 that respiration was associated with granular insoluble structures in the cell, it was not until these breakthroughs approximately 40 years later that confirmed mitochondria were the sites of respiration and oxidative phosphorylation. Soon after the discovery that mitochondria catalyzed oxidative phosphorylation, several groups found that mitochondria also maintained ion gradients. These ions included Na+, K+ and Ca2+ (Macfarlane and Spencer 1953; Bartley and Davies 1954). In order to maintain a gradient, it was clear that mitochondria needed to be membrane-bound. High-resolution electron micrographs provided by Palade and Sjöstrand in 1952 and 1953 confirmed the structure of mitochondria (Palade 1952; Sjostrand 1953). They found that mitochondria had a double membrane, and that the inner membrane consisted of transversely oriented ridges. Palade introduced the term “cristae” to describe the ridges seen in the mitochondrial inner membrane. He noticed that the cristae varied in number and spacing depending on the cell type from which the mitochondria were isolated (Palade 1952). It would later be known that these cristae folds increased the surface area of the mitochondrial inner membrane, enabling more efficient oxidative phosphorylation.

1  The Genetics of Mitochondrial Fusion and Fission

7

The next important mitochondrial discovery was the identification that mitochondria contained DNA. Non-mendelian cytoplasmic inheritance had been discovered in plants by Carl Correns since the early 1900s. This discovery in yeast by Boris Ephrussi in 1950 indicated that factors other than chloroplasts could be inherited by non-mendelian genetics. The Ephrussi laboratory used DNA mutagens and found that these mutations were cytoplasmically inherited although there had been no evidence of cytoplasmic DNA at the time (Roman 1980). Visualizing chick embryo mitochondria by electron microscopy in 1963, Margrit and Sylvan Nass noticed fibres within the mitochondria that were sensitive to nucleic acid stains. They showed in two back-to-back publications that these nucleic acid-containing fibres were DNA. The fibres were not removed when fixed with potassium permanganate indicating they were not RNA, and they were sensitive to DNAse (Nass and Nass 1963a, b). The presence of mitochondrial DNA (mtDNA) was further confirmed by Haslbrunner, Tuppy and Schatz. Using purified yeast mitochondria, biochemical analyses indicated that small, but significant amounts of DNA co-purified with mitochondria (Schatz et  al. 1964). The discovery of mtDNA sparked new interest in mitochondrial research and supported speculations that eukaryotic mitochondria evolved from endosymbiotic bacteria. Due to its bioenergetic capability and the early observations of shape and size of mitochondria, it is believed that mitochondria evolved from symbiotic living of bacteria inside primitive cells. Later, evidence such as the existence of a unique mitochondrial genome, ribosomes, and compositions of the inner and outer membranes further support this theory. With the discovery of mtDNA came the identification of diseases associated with its mutations. Wallace and Holt were the first to identify diseases associated with mutations in mtDNA (Wallace et al. 1988; Holt et al. 1988). Since then, many mitochondrial diseases have been identified and are caused not only by mutations in mtDNA, but also by mutations in nuclear genes encoding mitochondrial proteins. Research conducted in yeast to understand the process of mitochondrial inheritance determined that proper mitochondrial fusion and fission were crucial for the transmission of parental mitochondria to zygotes (Nunnari et al. 1997). Mitochondrial dynamics are now known to be crucial mediators of apoptosis and neurodegeneration. In the last decade much of the research work conducted on mitochondrial diseases has focused on the role of fusion and fission in disease pathogenesis. In 2001, Frank and colleagues discovered that mitochondrial fission was a process that preceded apoptosis. Inhibition of a mitochondrial fission factor prevented cell death (Frank et al. 2001). Furthermore, inhibition of mitochondrial fusion with unopposed fission promoted apoptosis (Sugioka et al. 2004). At approximately the same time that mitochondrial dynamics were discovered to be important in cell death, mutations in genes required for mitochondrial fusion and fission were identified in patients with neurodegenerative diseases (Alexander et  al. 2000; Kijima et  al. 2004; Waterham et  al. 2007). These discoveries are at the heart of the mitochondrial disease field at the moment. Nonetheless, none of these recent advances would have been possible without the 100 years of preceding research and remarkable discoveries made by our predecessors. The basic functions of mitochondria are summarized in Fig. 1.2.

8

E.Y.L. Chan et al.

Fig. 1.2  Mitochondrial structures and functions. (a) Static visualization of mitochondria. Thinsectioned transmission electron micrographs provide static views of mitochondria (arrow heads) as individual bean-shaped organelles. (Figure modified from McQuibban et al. 2003) (b) Dynamic observation of mitochondria. Fluorescence microscopy reveals the network and dynamic nature of mitochondria. These are a series of time-resolved images of yeast expressing mitochondrialtargeted GFP in 3-min intervals. Scale bar: 2 mm. (Figure modified from Shaw and Nunnari 2002). (c) Mitochondrial membrane architecture. Mitochondria are double-membrane-bound organelles consisting of four subcompartments: outer membrane, intermembrane space, matrix and inner membrane that folds into cristae. (d) Mitochondrial functions. Mitochondria are sites for several cellular processes, from left to right: metabolism, cellular respiration via the electron transport chain (ETC), calcium buffering and the apoptotic pathway. Metabolic pathways in mitochondria include the

1  The Genetics of Mitochondrial Fusion and Fission

9

1.1.1 First Description of Mitochondrial Dynamics One of the first in-depth descriptions of mitochondria in cells was achieved by Lewis and Lewis in 1915. Using cultured chick cells, they were able to observe live and fixed mitochondria, and could describe mitochondrial movement, quantity and dynamics. What they referred to as mitochondrial fusion and branching/separation (fission) are now known to be distinct, regulated processes required for proper mitochondrial function and inheritance (Nunnari et al. 1997; Twig et al. 2008; Chen 2005; Dimmer 2002). Figure 1.3 shows more recent fluorescent microscope images of mitochondria rapidly undergoing fusion and fission events. Changes in shapes, dislocation, fusion and fission were also noted and sketched out (Fig. 1.3). Despite the evidence of mitochondrial dynamics in 1915, mitochondria were often visualized and recorded by transmission electron microscopy as bean-shaped double-membrane organelles with four subcompartments: outer membrane, ­intermembrane space, inner membrane and matrix (Palade 1952; Sjostrand 1953) (Fig. 1.2). There were several observations that mitochondria have different morphologies according to cell types and stages in life cycle (Bereiter-Hahn 1990; Yaffe 1999). There were speculations that these different morphologies could be due to the dynamics of the organelle. However, it was not until the 1990s with the development of light microscopy and fluorescent probes that the dynamics of mitochondria in living cells were recorded and widely-accepted (Bereiter-Hahn and Voth 1994). Electron tomographs of three-dimensional mitochondrial structures serve as evidence for mitochondrial plasticity (Perkins et al. 2009). With the development of photoactivatible green fluorescent protein (PA-GFP), tracking the dynamics of a single mitochondrion has now become possible (Patterson and Lippincott-Schwartz 2002; Liu et al. 2009). Figures 1.3 and 1.4 summarize examples of mitochondrial visualization in different model organisms with a variety of microscopic approaches. Figure 1.4 also demonstrates the variety of mitochondrial morphologies that exist in different tissues; yet another example that this is a very dynamic and plastic compartment.

Fig.  1.2  (continued) oxidation of fatty acids and the conversion of pyruvate to acetyl CoA by pyruvate dehydrogenase (PDH) to enter the tricarboxylic acid (TCA) cycle. Transferring electrons and oxygen respiration by the ETC results in ATP production. Reactive oxygen species (ROS) are byproducts of cellular respiration. ROS are inactivated by mitochondrial superoxide dismutase (MnSOD/SOD2). The mitochondrial genome encodes subunits of the respiratory chain complexes. The adenine nucleotide translocator (ANT) and the voltage-dependent anion channel (VDAC) mediate shuttling of ATP and ADP. Mitochondria are essential for maintaining calcium ion homeostasis by the function of VDAC in the outer membrane and the Ca2+/H+ anti-porter in the inner membrane. ANT and VDAC also function with proteins in the Bcl-2 family shown in red in the apoptotic pathway. Pro-apoptotic members, Bak and Bax, ­facilitate the release of apoptosis inducing factor (AIF) and cytochrome c (CytC), respectively, activating caspases to trigger apoptosis. On the other hand, anti-apoptotic members, Bcl-2 and Bcl-X, inhibit the release of CytC and AIF

Fig. 1.3  Visualization of mitochondrial dynamics. (a) The first observation of the presence and the dynamic behavior of mitochondria in living cells by bright field microscopy. These sketches were derived from the observations of chick fibroblast cultures. The changes were described to be (a) bending, (b) shortening and elongating, (c) fusion of granules, (d) changing in shape and fusion to form a network, (e) changing in shape and fusion, and (f) changing in shape and fusion to form a single mitochondrion. The number below each sketch indicates the time observed. Figure taken from Lewis and Lewis, 1914. (b) Visualizing mitochondria in living cells by time-lapse fluorescence microscopy. Mitochondria were labeled with YFP. The blue color highlights mitochondria that were being monitored. Fusion was observed in the first three frames and fission was observed in the latter three frames. (Figure taken from Chan 2006) (c) Monitoring mitochondrial fusion by photoactivatable fluorescence microscopy. Mitochondria were labeled with mtRFP and photoactivatable mtGFP (mtPAGFP). A subset of mitochondria that were photoactivated are seen as yellow mitochondria. These mitochondria can then be monitored for a complete fusion process. Time zero indicates the start point of the fusion process. (Figure modified from Liu et al. 2009)

1  The Genetics of Mitochondrial Fusion and Fission

11

Fig.  1.4  Mitochondrial morphology (I) Normal and aberrant mitochondrial morphologies in model organisms. (A) Knocking-out fusion and fission genes in Saccharomyces cerevisiae results in abnormal mitochondrial morphologies. A tubulated mitochondrial network was observed in wild-type cells (Aa) while net formation and fragmentation were observed in Ddnm1 (Ab) and Dfzo1 (Ac) cells, respectively. (Figure modified from Shaw and Nunnari 2002). (B) Mutation in fzo causes a ‘fuzzy onion’ morphology of mitochondria in Drosophila melanogaster spermatids. (Ba) Wild-type mitochondria form into a giant onion in the onion stage of sperm development. (Bb) Instead of one giant mitochondrial derivative, individual mitochondria in fzo appear to wrap around each other and have failed to undergo proper fusion to form the nebenkern. (Figure modified from Hales and Fuller 1997). (C) Mutation in Drp-1 disrupts mitochondrial morphology in Caenorhabditis elegans muscle cells. Mitochondria in wild-type cells (Ca) have normal tubulated morphology whereas mitochondria in cells containing mutant Drp-1 (Cb) are observed as large blebs. (Figure modified from Labrousse et  al. 1999) (D) Mitochondrial network is aberrant in OPA1 mutated mouse embryonic fibroblast cells. Similar to fragmentation of mitochondria in Fzo1 mutated yeast, the mitochondrial network in mutant OPA1 cells (Db) is fragmented, unlike tubulated mitochondria in wild-type cells (Da).(II) Diverse mitochondrial morphologies and distributions in various tissues. Mitochondrial morphology, distribution and mass vary in different cell types. From (A) to (D), electron microscopy images show mitochondrial morphologies observed in lung, kidney, cardiac muscle and neuron

12

E.Y.L. Chan et al.

Since the identification of mitochondrial dynamics, technological advances together with genetic, structural and biochemical approaches have enabled the field to dissect the molecular mechanisms of mitochondrial fusion and fission. This chapter will describe the discoveries in model organisms that have helped this field progress, and outline the mechanisms of mitochondrial fusion and fission. The functional significance of mitochondrial dynamics will also be discussed. The chapter will conclude with what we feel are the most important recent advances in the last couple of years, and discuss genetic and biochemical approaches to more clearly understand the complex cell-biological activity of this important organelle.

1.1.2 The First Identified Components in Fusion/Fission by Genetic Approaches Although mitochondrial fusion and fission had been observed in many cell types and organisms, it was not until 1997 that Hales and colleagues identified and characterized the first gene involved in mitochondrial dynamics. The Drosophila gene fuzzy onions (Fzo) is required for mitochondrial fusion in Drosophila spermatids. Mutations in Fzo result in abnormal sperm development and consequently male sterility (Hales and Fuller 1997). During the onion stage of spermatogenesis, mitochondria aggregate and fuse, forming two mitochondrial derivatives that wrap around each other, creating a spherical structure known as the Nebenkern. Electron microscope (EM) images of the cross section of WT Nebenkerns resemble onion slices whereas fzo Nebenkerns appear fuzzy and misshapen (see Fig. 1.4). Homologs of Drosophila fzo with similar functions in mitochondrial fusion have since been identified in many organisms including the nematode worm C. elegans (Fzo-1), the budding yeast S. cerevisiae (Fzo1) and in mammals (Mitofusins 1 and 2 [Mfn1 and Mfn2]; (Kanazawa et al. 2008; Hermann et al. 1998; Eura 2003; Chen 2003)). Mutational analyses in S. cerevisiae and C. elegans identified the second ­component of mitochondrial dynamics. The identification of yeast Dnm1 and its worm homolog Drp-1 were identified almost simultaneously to be factors required for mitochondrial fission (Bleazard et  al. 1999; Labrousse et  al. 1999). Previous studies in yeast (Dnm1) and mammals (Drp1) had indicated that Dnm1 and Drp1 are required in the maintenance of mitochondrial morphology and distribution (Otsuga et al. 1998; Smirnova et al. 2001). Two separate groups formally showed that Dnm1/Drp1 regulated mitochondrial morphology and distribution by participating in mitochondrial fission. Knocking-out Dnm1 in yeast resulted in a net-like mitochondrial morphology, a phenotype due to insufficient fission and unopposed fusion. Immunogold labeling indicated that Dnm1 was localized to mitochondrial constriction sites about to undergo fission and on tips of mitochondria that had just completed division. Furthermore, the Ddnm1 mutation rescued the fzo1-1 conditional mutant, implicating it as a fission factor (Bleazard et al. 1999). Consistent

1  The Genetics of Mitochondrial Fusion and Fission

13

with the observations in yeast, over-expression of Drp-1 in worms resulted in fragmented mitochondria instead of the long tubular mitochondrial network seen in WT worms. On the contrary, worms expressing mutant forms of Drp-1 had aggregated mitochondria due to lack of mitochondrial fission (Labrousse et  al. 1999). Surprisingly, the aggregated mitochondria were connected by thin tubules. Further analysis by labeling the matrix and outer membrane (OM) with CFP and YFP respectively determined that those thin connecting tubules were tubules of the OM (see Fig. 1.4). This suggested that although mutant Drp-1 did not allow OM scission, IM scission still occurred (Labrousse et al. 1999). Since the identification of the founding fusion and fission genes Fzo and Dnm1/ Drp1, many other genes have been shown to be involved in maintaining proper mitochondrial morphology. Of particular note are genetic screens conducted in yeast and worms that have helped identify many of these new components.

1.1.3 Yeast Screens Ugo1 and Pcp1 (also known as Rbd1) were identified in a yeast screen as genes that suppressed the Dnm1 fission defect, implicating them as fusion factors. Since fission mutants lose their mtDNA, and the malfunction of either fusion or fission is sufficient for mtDNA loss, the authors screened mutants that retained mtDNA in the double mutant (indicating restored balance of fusion and fission), but lose their mtDNA in the presence of functional Dnm1 (indicating functional fission but lack of fusion). Dugo1 and Dpcp1 mutants had fragmented mitochondria reminiscent of Dfzo1 fusion mutants and rescued the Ddnm1 fission defect, strongly implying that they are required for proper mitochondrial fusion (Sesaki and Jensen 2001). Subsequent analysis of Pcp1 revealed that it is a serine protease required for the cleavage of Mgm1 (Sesaki 2003). Mgm1 had originally been identified as a gene required for mitochondrial genome maintenance (Jones and Fangman 1992). Further studies showed that deletion of Mgm1 not only resulted in the loss of mtDNA, but also compromised respiration and mitochondrial fusion (Jones and Fangman 1992; Wong et al. 2000). Identification of Pcp1-dependent Mgm1 cleavage revealed the regulation of mitochondrial dynamics by proteolytic processing (Sesaki 2003; McQuibban et al. 2003). A systematic genetic screen performed on 4,794 knock-out yeast mutants identified components required for the maintenance of mitochondrial morphology and respiratory growth. Mutants were screened for their inability to grow on glycerol (indicating respiratory incompetence) and aberrant mitochondrial morphology. The genes required to maintain normal mitochondrial distribution and morphology (MDM) were divided into three classes (Dimmer 2002; McConnell et  al. 1990). Class I genes were essential for normal mitochondrial morphology; WT mitochondrial morphology was never observed in Class I mutants. Class II and III mutants had either fragmented or aggregated mitochondria but class II mutants were respiratory competent whereas class III mutants were petites (respiratory incompetent).

14

E.Y.L. Chan et al.

This systematic approach enabled genome-wide screening of yeast mutants, and identified 15 novel genes required for the maintenance of mitochondrial morphology (Dimmer 2002). Further characterization of Mdm30, a gene identified in the screen with previously unknown functions led to the discovery that ubiquitination might regulate the stability of Fzo1, a protein directly involved in mitochondrial fusion. Although previous studies had implicated ubiquitin and the proteasome in maintaining normal mitochondrial morphology, this study likely provided the first evidence of mitochondrial protein-targeted ubiquitination (Fritz 2003). The screens performed previously were unable to identify essential genes involved in mitochondrial dynamics (Dimmer 2002; Sesaki and Jensen 2001; McConnell et  al. 1990; Burgess et  al. 1994). Cells with any one of its essential genes knocked out were simply not viable. To examine whether essential genes were required for mitochondrial morphology, Altmann and colleagues used a collection of yeast strains with essential genes expressed under a regulatable TetO7 promoter. Gene repression was achieved by the addition of doxycycline to repress the TetO7 promoter (Gari et al. 1997; Mnaimneh et al. 2004). Of the 728 yeast genes that were screened, 119 of them were required for normal mitochondrial morphology. Expectedly, some of the highly enriched genes included those of the mitochondrial import machinery. Surprisingly, however, many genes involved in other processes were also highly enriched. These included genes involved in ergosterol biosynthesis, vesicular transport and the ubiquitin/26S proteasome-dependent degradation machinery. The results from this screen indicate that, albeit indirectly, many pathways affect mitochondrial morphology (Altmann 2005), and more research has to be conducted to determine the roles that these pathways play in mitochondrial dynamics (see later sections of chapter). Moreover, yeast multi-copy suppressor screens have been widely conducted to elucidate genes participating in the mitochondrial dynamics pathway. Num1 was found as a hit in the multi-copy suppressor screen of the DNM1-109 mutant (Cerveny et al. 2007). A high-copy suppressor of Mdm12 identified Phb2 belonging to the highly conserved family of prohibitins that function in maintaining IM architecture and processing of the mammalian Mgm1 homolog, OPA1. In addition, the synthetic genetic array (SGA) approach is a recent development to screen for genome-wide genetic interactions in a systematic format. For instance, the genetic interactome of prohibitins provided more insight into the function of prohibitins in the inner membrane and uncovered its role in phospholipid homeostasis. This activity may be linked to the regulation of Mgm1 processing and its role in the maintenance of mitochondrial morphology.

1.1.4 Worm Screen An RNAi screen was conducted in C. elegans to determine which of the known mitochondrial proteins were required for the maintenance of normal morphology. Worms expressing RFP-tagged TOM20 were fed E. coli cells expressing dsRNA

1  The Genetics of Mitochondrial Fusion and Fission

15

specific to the target genes. Mitochondria in the body wall muscles of these worms were examined for their morphology. Of the 719 genes predicted to be mitochondrial, more than 80% were required to maintain normal mitochondrial morphology. RNAi knock-down of 574 (79.8%) genes resulted in mitochondria that were fragmented while knock-down of the remaining 25 (3.5%) genes resulted in elongated mitochondria. Classification of the genes whose knock-down resulted in fragmentation yielded an unexpectedly high number of genes required for basic mitochondrial functions such as metabolism and oxidative phosphorylation. The results from this screen highlight the intricate relationship between proper mitochondrial function and dynamics (Ichishita et al. 2007).

1.1.5 Mammalian Studies The functions of several human orthologs of the fusion and fission molecules (as described above and below) have recently been studied in well-characterized cultured cell lines. For instance, over-expression of Drp1 was shown to alter mitochondrial morphology (Smirnova et  al. 1998; Misaka et  al. 2002). Furthermore, Drp1 and mitochondrial fission were shown to be essential for mitochondrial distribution in neurons (Chen and Chan 2009). In addition to simple over-expression systems in cultured cells, the siRNA knockdown approach has also provided great advances to the study of mitochondrial dynamics. For instance, knockdown of OPA1 leads to fragmentation of mitochondria (Olichon et al. 2003). Beyond experiments in tissue culture cells, genetic and biochemical approaches in mice have been powerful approaches in studying the role of mitochondrial dynamics in both cellular development and the pathophysiology of related diseases. The phenotypes and characterization of various mouse models have lead to a better understanding of the pathophysiology of many ­neurodegenerative diseases and uncovered the functional roles of the players in mitochondrial dynamics. Specifically, Mitofusins 1 and 2 (Mfn1/2) were identified as the mammalian orthologs of fly and yeast Fzo1 from a mouse cDNA library (Chen et al. 2003). Both Mfn1 and Mfn2 knock-outs show embryonic lethality of homozygous mutants. In addition, Mfn-deficient mouse embryonic fibroblast cells have altered mitochondrial morphology and dynamics. Similarly, homozygous Opa1 null mice also show early embryonic lethality. Heterozygous Opa1 mice are viable but their inter-crosses give no live progeny of Opa1−/− (Davies et al. 2007). Heterozygous Opa1 mice displayed fragmented mitochondria, abnormal optic nerve myelination and structure as well as impaired visual functions (Davies et al. 2007). PARL is the mammalian homologue of yeast Pcp1/Rbd1, and when its function is disrupted in the mouse, the animal dies at 6  weeks of age. At a molecular level, PARL knock-out mice reiterate the importance of PARL-dependent OPA1 processing, and reveal the pivotal role of OPA1 in apoptosis by regulating cytochrome c storage and release.

16

E.Y.L. Chan et al.

1.2 Mechanistic Models for the Molecular Machines of Mitochondrial Dynamics The pioneering discoveries of the molecular machineries of mitochondrial fusion and fission were largely derived from genetic studies in yeast, flies and worms as described above. Due to the simplicity of a single cell system and powerful genetic approaches, more is known about the molecular mechanisms of yeast mitochondrial dynamics. The key players have been characterized as mechanoenzymes. These mechanoproteins are the dynamin-related proteins (DRPs) (Hoppins et  al. 2007). DRPs are large GTPases that are involved in several membrane remodeling events (Praefcke and McMahon 2004). They are distinguished from other GTPases by their ability to bind to lipids, self-assemble, and to self-stimulate its GTP hydrolysis activity (Praefcke and McMahon 2004). All DRPs contain three conserved domains; the catalytic GTPase domain serves as the site for GTP binding and hydrolysis; the middle domain and GTP effector domain (GED) are essential for oligomerization and self-assembly. The region between the middle domain and the GED serves as the site for lipid interactions (Praefcke and McMahon 2004). The four well-characterized dynamin-related proteins in yeast are Vps1, Dnm1, Mgm1 and Fzo1. Vps1 mediates Golgi vesicle formation while the other three DRPs function in mitochondrial dynamics. Dnm1 directly mediates the fission of mitochondria while Mgm1 and Fzo1 are the key players for mitochondrial fusion (Hoppins et al. 2007). These DRPs are highly conserved proteins and their higher eukaryotic counterparts (summarized in Table 1.1) are also fission and fusion molecules. Beyond the DRPs, there are several other classes of proteins that orchestrate mitochondrial dynamics. The complexity of these processes can be attributed to the fact that unlike other organelles, mitochondria have a double membrane. Hence, proper mitochondrial fusion and fission requires the maintenance of four distinct compartments: the outer membrane (OM), the intermembrane space (IMS), the inner membrane (IM) and the matrix. The following section will focus on the mechanisms of mitochondrial fusion and fission, summarizing the findings in model organisms and key biochemical techniques that helped elucidate these complex mechanisms. Please see Fig. 1.5 for a pictorial summary of these membrane gymnastics.

1.2.1 Mitochondrial Fission 1.2.1.1 The Components of Mitochondrial Fission Dnm1/Drp1 Although the regulation of mitochondrial fission differs slightly from one organism to another, the core components required remain highly conserved. In general, mitochondrial membrane fission requires Fis1 and a dynamin-related protein

–

–

–

–

–

Num1

Mdm36

Mdm33

–

–

NUM1

MDM36

MDM33

MTP18

SH3GLB1

SUMO1

–

–

Caf4

CAF4

–

–

Mdv1

MDV1

MARCHV

Fis-1/2

Fis1

Drp-1

FIS1

Genes implicated in fission DNM1 Dnm1

–

–

–

–

–

–

–

–

Fis1

Drp1

SUMO1

MARCHV

Endophilin B1

MTP18

–

–

–

–

–

hFis1

DRP1

Dynamin-related GTPase acting as a mechanoenzyme that self-assembles to form ring and constricts mitochondria TRP repeats functioning in targeting of Dnm1 WD40 repeats serving as an adaptor for Fis1 and Dnm1 WD40 repeats serving as an adaptor for Fis1 and Dnm1 Cell-cortex anchored protein functioning in targeting of Dnm1 Fission molecule facilitating Dnm1 and Num1 complex Protein in intermembrane space involved in fission of yeast inner mitochondrial membrane Protein in intermembrane space involving in fission of mammalian inner mitochondrial membrane BAR domain containing protein involving in fission of mammalian outer mitochondrial membrane Ubiquitin ligase in mammalian mitochondrial outer membrane adding ubiquitins onto Drp1 and MFN2 Ubiquitin-like modifying enzyme modifying and regulating Drp1

Table 1.1  List of genes characterized to have implicated roles in mitochondrial dynamics Gene name S.ceverisiae C.elegans Drosophila Mammals Known function

(continued)

Related diseases

1  The Genetics of Mitochondrial Fusion and Fission 17

–

–

–

–

–

Pcp1/Rbd1

Mdm30

Ups1

PCP1

MDM30

UPS1

–

–

Rhomboid-7

–

Opa1-like

Eat-3

Ugo1

Fzo

–

Drosophila

Fzo-1

–

C.elegans

UGO

Genes implicated in fusion Fuzzy Fzo1 onions MGM1 Mgm1

GDAP1

Table 1.1  (continued) Gene name S.ceverisiae

PRELI

–

PARL

–

OPA1

MFN1, MFN2

GDAP1

Mammals

Dynamin-related GTPase possessing heptad repeats for tethering of mitochondria Dynamin-related GTPase mediating fusion and maintaining cristae structure Carrier protein coordinating outer and inner mitochondrial membrane fusion. Rhomboid serine protease involving in Mgm1 processing F-Box E3 ligase involving in maintaining Fzo1 level Protein in intermembrane space involving in regulating Mgm1 processing

Ganglioside-induce differentiationassociated protein involving in fission

Known function

ADOA

CMT2A

CMT4A

Related diseases

18 E.Y.L. Chan et al.

1  The Genetics of Mitochondrial Fusion and Fission

19

Dnm1/Drp1 (Bleazard et al. 1999; Mozdy et al. 2000; Tieu and Nunnari 2000). Dnm1/Drp1 are the DRPs involved in mitochondrial membrane fission. Yeast Dnm1 exists as punctate structures in the cytosol and on the mitochondrial OM (Otsuga et al. 1998; Naylor 2005), while mammalian Drp1 is mostly diffuse cytoplasmic, with some punctae on mitochondria (Smirnova et al. 2001). Despite their differences in localization, Dnm1/Drp1 in yeast, worms and mammals were found to localize to the OM at sites of mitochondrial fission (Sesaki and Jensen 1999; Legesse-Miller 2003) consistent with their roles as proteins that divide mitochondria. Although homologs have been identified in many organisms, it is the detailed biochemical and structural analyses on yeast Dnm1 and mammalian Drp1 that have helped shed light on the mechanism of DRP-mediated mitochondrial fission. Like dynamins, Dnm1/Drp1 assemble into dimers/tetramers (Fukushima et al. 2001; Shin et  al. 1999). In addition, they organize into ring-like structures (Smirnova et al. 2001; Ingerman 2005; Yoon et al. 2001). The spirals formed by Dnm1 were able to constrict liposomes, reducing their diameter to that of mitochondrial constriction sites (Ingerman 2005). Independent studies using truncated forms of yeast Dnm1 and mammalian Drp1 identified the GED as an important region for the formation of higher ordered structures (Fukushima et al. 2001; Zhu 2004). The GED of mammalian Drp1 forms intermolecular interactions with itself and intramolecular interactions with the GTPase domain. K679 in the GED was found to be required for intra- not intermolecular interactions since the K679A mutation resulted in reduced intramolecular interactions but did not affect its intermolecular interactions (Zhu 2004). Other than a function in coordinating intra- and intermolecular interactions, the GED was also shown to be sufficient for mitochondrial targeting of Drp1 (Pitts 2004). One of the characteristic features of dynamins and DRPs is oligomerizationstimulated GTP hydrolysis. The first evidence that the mitochondrial fission DRPs also had this feature came from the analysis of yeast Dnm1. Ingerman, et  al (Ingerman 2005) characterized the kinetics of GTP hydrolysis by Dnm1 and observed a lag in attaining the maximum rate of GTP hydrolysis. This lag was not observed in a Dnm1 middle domain mutant (Dnm1G385D) defective in self-assembly. Although this mutant could dimerize, it failed to form higher order structures that facilitated GTP hydrolysis (Ingerman 2005). A lethal mutation identified in the Drp1 middle domain (Drp1A395D) also compromised its organization into higher order oligomers. While Drp1 monomers still formed dimers/tetramers, they failed to organize into higher order structures (Chang et  al. 2010). Localization of the Dnm1G385D middle domain mutant to mitochondria indicates that the oligomeri­ zation occurred after recruitment of the dimer to the membrane (Bhar 2006). These data strongly suggest that Dnm1/Drp1 self-assemble into higher order oligomers upon recruitment to the mitochondria, and that this self-assembly is the ratelimiting step in GTP hydrolysis (Ingerman 2005; Bhar 2006). The lethality of the reported Drp1A395D mutation points to the importance of this domain and its role in oligomerization and proper Dnm1/Drp1 function.

20

E.Y.L. Chan et al.

Fig. 1.5  Mechanistic models of yeast mitochondrial fusion and fission. The diagram illustrates proposed models of mitochondrial dynamics and mechanisms of actions by the key players. Blue boxes, (a–c), show sequential steps of mitochondrial fusion whereas red boxes, (d–f), present the proposed mitochondrial fission event. Grey boxes, (g–i), show the currently known regulatory pathways in mitochondrial fusion and fission. Proposed model of mitochondrial fusion: (a) The mitochondrial fusion machinery complex consists of Fzo1 and Ugo1, anchored to the outer membrane, l-Mgm1, anchored to the inner membrane, and s-Mgm1, peripherally associated with the inner membrane. Fzo1 tethers mitochondria by forming a complex in trans via its coiled-coil region. Following the tethering step, it is yet unclear how Fzo1 causes fusion of the outer membrane bilayers. Ugo1 is suggested to be crucial for the lipid-mixing step. (b) Both l-Mgm1 and s-Mgm1 are the key players in the inner membrane fusion reaction. l-Mgm1 is proposed to serve as a docking site for s-Mgm1 recruitment. Homo- and hetero-oligomerization of the two isoforms in both cis and trans are important for tethering of the opposing inner membranes. As observed in OPA1, membrane tubulation is a possible action that drives membrane fusion (blue question mark). Tubulation increases the curvature of the membrane, creating high-energy stress that can be relieved upon fusion. (c) The mechanism following the tethering of inner membranes is largely unknown. Besides functioning in mitochondrial fusion, Mgm1 also plays a crucial role in the maintenance of cristae structures (asterisk). Proposed model of mitochondrial fission: (d) Dnm1 acts as a mechanoenzyme to mediate membrane scission. Dnm1 is targeted to, oligomerizes, and assembles onto the cytosolic side of the outer membrane. Inner membrane fission may be mediated separately from outer membrane fission by the protein Mdm33. (e) Dnm1 self-assembles into a high-order helical structure in a GTP-dependent manner. This structure results in mitochondria wrapped by Dnm1 and membrane constriction by Dnm1 self-assembly. (f) Upon GTP hydrolysis, the fission event is completed and Dnm1 is released from the outer membrane. Regulation of mitochondrial dynamics: (g) Proteolytic processing of Mgm1 serves as a mode of regulation since equal molarity of s- and l-Mgm1 is critical for proper fusion. Alternative topogenesis is the

1  The Genetics of Mitochondrial Fusion and Fission

21

Fis1 Fis1 is an integral membrane protein on the mitochondrial OM that recruits Dnm1/ Drp1 to mitochondria (Mozdy et al. 2000; Yoon et al. 2001). In the absence of Fis1, Dnm1/Drp1 localization to mitochondria is severely compromised (Mozdy et  al. 2000; Yoon et al. 2001). The N-terminus of Fis1 faces the cytosol and its C-terminus faces the IMS (Fig.  1.5; (Mozdy et  al. 2000; James 2003)). Most of the protein resides in the cytosol, and is anchored to the OM by its C-terminal TM segment (Stojanovski 2004). Disruption of its short C-terminal tail with or without its TM segment disrupts yeast (Fis1) and human Fis1 (hFis1) localization to mitochondria (Mozdy et al. 2000; Stojanovski 2004; Yoon et al. 2003). Unlike Dnm1/Drp1 that appear as discrete punctae, Fis1 is uniformly distributed throughout the membrane (Mozdy et al. 2000). Solution NMR spectroscopy revealed that the cytosolic portion of yeast Fis1 (Fis1) and human Fis1 (hFis1) organizes into six a-helices (a1–a6) that resemble tetratricopeptide repeat (TPR) folds (Suzuki et al. 2003; Suzuki 2005). TPR motifs mediate protein-protein interactions and are usually a part of multiprotein complexes (reviewed in Blatch and Lassle, 1999 (Blatch and Lassle 1999)). Subsequent crystal structure analysis indicates that a2–a3 and a4–a5 of Fis1/hFis1 form two TPR motifs (TPR1/2) resulting in a concave hydrophobic binding pocket (Suzuki 2005; Dohm et al. 2004). The folding and structure of the six helices are very similar in Fis1 and hFis1. However, the N-terminus of hFis1 is largely unstructured (Suzuki et al. 2003) whereas that of Fis1 is longer and forms an a-helix strikingly similar to that of a1 in hFis1 (Suzuki 2005). Since TPR motifs mediate protein-protein interactions, it would seem intuitive that the TPRs in Fis1/hFis1 serve to recruit Dnm1/Drp1 to mitochondria during fission. Co-immunoprecipitation (co-IP) experiments performed after chemical cross-linking showed that hFis1 and Drp1 physically interact (Yoon et  al. 2003). Subsequently, using truncated forms of Fis1/hFis1, it was determined that Fis1/hFis1 and Dnm1/Drp1 interacted in a TPR-dependent manner. However, contrary to the

Fig. 1.5  (continued) proposed mechanism for the balanced formation of s- and l-Mgm1. In this model, precursor Mgm1 translocates through the inner membrane and stops at the first hydrophobic segment. The N-terminal MTS is cleaved off resulting in l-Mgm1 anchored to the inner membrane. Alternatively, high mitochondrial ATP levels allow precursor Mgm1 to be pulled further into the matrix until the second hydrophobic segment that contains a sequence recognized by Rbd1. Rbd1 cleaves precursor Mgm1 into, s-Mgm1. Other proteins such as Ups1, Phb2 and Gep1 that have implicated roles in phospholipid homeostasis (represented by the purple sphere) may play a role in Mgm1 processing. (h) Fzo1 levels are regulated by ubiquitination and 26S proteasome degradation. Alpha-factor pheromone treatment causes Fzo1 proteasome-dependent degradation. Fzo1 was also shown to be ubiquitinated by Mdm30. However, it is uncertain whether ubiquitinated Fzo1 is targeted to 26S proteasome for degradation. (i) Targeting Dnm1 to outer membrane. Fis1/Mdv1 targeting of Dnm1 to mitochondria is essential for the fission event. The other two pathways, Fis1/Caf4 and Num1, are non-essential but important for Dnm1 targeting to the cell cortex. Mdm36 is involved in Num1 function. See main text for additional details

22

E.Y.L. Chan et al.

co-IPs performed after cross-linking, little to no binding between Dnm1/Drp1 and full-length Fis1/hFis1 was observed (Yu 2005; Wells et al. 2007). These data suggest that adaptor proteins may be required to bridge the interaction between Dnm1/ Drp1 and Fis1/hFis1. Although no adaptor proteins have been identified in mammals, in yeast, Mdv1, Caf4, Num1 and Mdm36 serve as adaptors to recruit Dnm1 to mitochondria (Cerveny et  al. 2007; Tieu and Nunnari 2000; Griffin 2005; Cerveny 2003; Hammermeister et al. 2010). Mdv1/Caf4 Mdv1 and Caf4 are the yeast adaptor proteins required for the recruitment of Dnm1 to the mitochondrial surface. Mdv1 (also known as Gag3 and Net2) is a peripheral protein of the mitochondrial OM. It has an N-terminal extension (NTE), a centre coiled-coil domain and six or seven WD-40 repeats in its C-terminus (Tieu and Nunnari 2000; Fekkes et al. 2000; Cerveny et al. 2001). Mdv1 interacts with Fis1 and Dnm1 through its NTE and WD-40 repeats respectively (Tieu and Nunnari 2000; Naylor 2005). While data convincingly shows that Dnm1 binds efficiently to the Mdv1 WD-40 repeat only in its GTPbound state (Naylor 2005), the Mdv1 binding site on Fis1 is still controversial (Wells et al. 2007; Zhang and Chan 2007). Co-IPs performed using a dimeric form of Dnm1 (Dnm1G385D) showed that Mdv1-dependent recruitment of Dnm1 to mitochondria occurs before Dnm1 higher order assembly (Bhar 2006). Interestingly, Dnm1 localization to mitochondria is maintained in Dmdv1 mutants and mitochondrial fission is only partially compromised (Cerveny et al. 2001; Griffin 2006). These phenotypes are due to the redundant activity of Caf4. Caf4 is a protein that also contains WD-40 repeats and interacts with Fis1 and Mdv1 through its N-terminus. It also interacts with Dnm1 through its C-terminal WD-40 domain (Griffin 2005). Although the deletion of Mdv1 or Caf4 alone had little effect on Dnm1 localization, deleting both genes severely compromised its localization, to an extent similar to that of the Dfis1 mutant (Griffin 2005). This indicates that Mdv1 and Caf4 function as adaptor proteins, albeit with slight redundancy, to recruit Dnm1 to the mitochondrial membrane. Dnm1/Drp1 and Fis1/hFis1 are the core components in the mitochondrial fission machinery. Although adaptor proteins appear to be required for the recruitment of Dnm1/Drp1 to mitochondria, they have only been identified in yeast. Nevertheless, the mechanism of mitochondrial fission mediated by Dnm1/Drp1 and Fis1/hFis1 remains highly conserved. The OM protein Fis1/hFis1 recruits dimeric/tetrameric Dnm1/Drp1 to mitochondria in a TRP-dependent fashion mediated by adaptor proteins. Upon recruitment, the Dnm1/Drp1 dimers/tetramers bind GTP and assemble into ring-like higher order oligomers, constricting the mitochondrial membrane. Oligomerization-stimulated GTP hydrolysis provides the energy required to sever the membrane and subsequent dissociation of the Dnm1/Drp1 oligomer (see Fig. 1.5).

1  The Genetics of Mitochondrial Fusion and Fission

23

1.2.1.2 Additional Fission Proteins Mdm36/Num1 Complex Num1 is a protein that was isolated as a multi-copy supressor of a dominant-negative form of Dnm1 (Cerveny et al. 2007). Num1 had been previously characterized to be responsible for nuclear migration (Farkasovsky and Kuntzel 1995). Here, the authors demonstrated cortical localization of Num1 and frequent co-localization with Dnm1, and an activity of Num1 in mitochondrial division and inheritance. A recent study has further demonstrated the Num1/Dnm1 complex by characterizing the fission promoting protein Mdm36 (Hammermeister et  al. 2010). This study showed that Mdm36 is responsible for maintaining protein complexes at the cell cortex to promote mechanical forces on mitochondrial membranes that can lead to membrane fission. Mff The Drosophila homolog of human Mff (mitochondrial fission factor) was identified in an RNAi screen for disruption in mitochondrial morphology using Drosophila S2R + cells. Immunofluorescence and protease protection experiments determined that Mff is a TM protein of the mitochondria OM. siRNA knock-down of Mff resulted in elongated mitochondrial tubules similar to those of Drp1 knockdown, but more severe than Fis1 knock-down mitochondria. Consistent with a function in mitochondrial fission, knock-down of Mff suppressed Mfn fusion defects and inhibited fragmentation-induced apoptosis. However, over-expression of Mff did not cause mitochondrial fragmentation, a phenotype identical to that of Drp1 over-expression. This suggests that although Mff, like Drp1, is a pro-fission protein, is not responsible for the rate-limiting step in mitochondrial fission (Gandre-Babbe and van der Bliek 2008). Letm1 LETM1 is a gene whose deletion has been associated with seizures in Wolf-Hirschhorn syndrome (Zollino et al. 2003). Characterization of the protein indicates Letm1 is an integral membrane protein of the mitochondrial IM, with the bulk of the protein facing the matrix (Schlickum et al. 2004; Dimmer et al. 2007). Like its yeast homolog Mdm38, Letm1 is required for the maintenance of proper mitochondrial morphology (Dimmer 2002; Dimmer et  al. 2007). Overexpression or knock-down of Letm1 by RNAi resulted in fragmented ring-shaped mitochondria but did not affect mitochondrial function. Interestingly, fragmentation occurred in a Drp1-independent manner, but was rescued by nigericin, a pharmacological agent that restores the K+/H+ antiport. This implied that the fusion/fission balance was not disrupted, but rather reflected mitochondrial

24

E.Y.L. Chan et al.

plasticity. Mitochondria adapted to ion imbalance by changing its morphology in order to maintain function (Dimmer et al. 2007). Coincidentally, an independent study that identified Drp1 phosphorylation by the Ca2+/calmodulin-dependent protein kinase Ia (CaMKIa) observed similar ring-like mitochondria upon treatment with high levels of K+ (see regulating mitochondrial fission; (Han et  al. 2008)). Recently we have characterized Letm1 in the fly (McQuibban et al. 2010) and demonstrated its potential role in neuronal activity and mitophagy, the autophagic clearance of damaged mitochondria (discussed in more detail later). Together, these data further support the notion that mitochondrial morphology is closely related to cellular ion levels, and can regulate the health of the organism in several ways. GDAP1 Ganglioside-induced differentiation protein 1 (GDAP1) is a protein implicated in Charcot-Marie-Tooth (CMT) disease. Mutations in GDAP1 were reported in patients with CMT, although the function of this protein has not been characterized (Baxter et al. 2002; Cuesta et al. 2002; Di Maria et al. 2004). Cell culture and protease protection analyses determined that GDAP1 is an integral membrane protein of the mitochondrial OM. Over-expression of GDAP1 resulted in highly fragmented mitochondria, while RNAi knock-down caused mitochondrial elongation. Although the mechanism by which GDAP1 promotes mitochondrial fission remains to be determined, these data demonstrate that GDAP1 is a newly identified member of the mitochondrial fission machinery (Niemann 2005). 1.2.1.3 Inner Membrane Fission Despite Dnm1/Drp1 and Fis1/hFis1 being the best characterized mitochondrial ­fission proteins, studies in yeast and worms indicate that they may only be responsible for OM fission. Mitochondria of C. elegans expressing mutant forms of Drp-1 were still connected by thin tubules of the OM while IM fission still occurred (Labrousse et al. 1999). Similarly, yeast Ddnm1 mutants did not achieve complete ­mitochondrial fission, but had matrix separation (Jakobs 2003). These data suggest that Dnm1/Drp1 and Fis1 are not required for IM fission, and that other components mediate this process. Yeast Mdm33, mammalian Mtp18 and MTGM were later identified and proposed to be factors mediating mitochondrial IM fission (Messerschmitt 2003; Tondera 2004, 2005; Zhao et  al. 2009). Mdm33 has no known mammalian homologs while Mtp18 has known homologs only in metazoans (Messerschmitt 2003; Tondera 2004, 2005). MTGM, on the other hand, has homologs in eukaryotes from yeast to human. Remarkably, all mammalian orthologs of MTGM have 100% amino acid sequence identity (Zhao et  al. 2009). Reduction of Mdm33,

1  The Genetics of Mitochondrial Fusion and Fission

25

Mtp18 and MTGM protein levels resulted in elongated mitochondria whereas over-expression led to fragmentation. These phenotypes are reminiscent of fission mutants, implicating them in the mitochondrial fission process. Although the topology of Mtp18 remains unclear, it is proposed to reside in the IMS (Tondera 2005). Mdm33 is an integral protein of the inner membrane with its C-terminus in the IMS and predicted N-terminal coiled-coil domains in the matrix. Analysis of its oligomeric state showed that Mdm33 has homotypic interactions, likely mediated by its coiled-coil domains. Over-expression of Mdm33 caused IM fragmentation and loss of cristae, suggesting IM fission defects. Based on these observations, Mdm33 on apposing IMs are proposed to form homotypic interactions through their coiled-coil domains in the matrix, mediating IM fission (Messerschmitt 2003). MTGM was identified as a human nuclear gene whose product is highly enriched in brain tumor cell lines and tumor tissues. Like Mdm33, MTGM is an integral membrane protein of the mitochondrial IM. Rather than coiled-coil domains, MTGM has a highly conserved tetrad of GxxxG motif that also mediates protein-protein interactions. MTGM-induced mitochondrial fragmentation is Drp1-dependent, indicating that it could be a regulator of Drp1. However, its mitochondrial IM localization suggests that MTGM might mediate IM fission (Zhao et  al. 2009). Additional experiments have to be conducted to determine if MTGM is directly involved in IM fission or whether it is simply a regulator of Drp1.

1.2.2 Mitochondrial Fusion Mitochondrial fusion is distinct from other intracellular membrane fusion events because it requires the fusion of two separate membranes coordinately even though these two fusion processes are rather separable events (Hoppins et  al. 2007). In yeast, while Fzo1 mediates outer and Mgm1 mediates inner membrane fusions, Ugo1 may be the key to coordinating these two events and has been recently shown to be involved in the lipid-mixing step. Importantly, these three key players are known to form a stable complex (Hoppins et al. 2007). While the mechanism of how the DRPs and other factors function in membrane fission/scission is largely known (as detailed above), the mechanism of the opposite reaction, membrane fusion, is still unclear due to lack of detailed structural information. There are an increasing number of studies that suggest DRPs can mediate fusion in a similar mechanism as fission by membrane tubulation. This tubulation of the mitochondrial membrane would increase curvature stress and drive the fusion reaction. The crystal structure of bacterial dynamin-like protein BDLP and its liposome tubulation structure have shed some light on our understanding of the potential DRP-mediated fusion reaction mechanism (Low and Lowe 2006; Low et al. 2009). We will now detail what is presently known about mitochondrial membrane fusion (see also Fig. 1.5)

26

E.Y.L. Chan et al.

1.2.2.1 Outer Membrane Fusion Fzo1 Fzo1 is anchored to the outer mitochondrial membrane with both its N- and C-termini facing the cytoplasm. Similar to Dnm1, self-assembly and GTPase activity are crucial for Fzo1 function. Fzo1 possesses predicted coiled-coil regions of the heptad repeat domains for stable cis and trans Fzo1-Fzo1 interactions. Cis interactions of Fzo1 in the same membrane may lead to self-assembly of Fzo1, and trans interactions of Fzo1 is essential for tethering the fusing mitochondria (Hoppins et al. 2007). MFN1 and MFN2 are mammalian orthologs of Fzo1. Even though the two mitofusins share 80% similarity, evidence suggests that they have functional differences. The trans interactions for tethering mitochondria are observed in MFN1 but to a lesser extent in MFN2 (Benard and Karbowski 2009). Apoptosis-related proteins Bax and Bak specifically associate with MFN2 but not MFN1. In addition, mutations in MFN2 but not MFN1 are linked to Charcot-Marie-Tooth ­disease type 2A. Mechanistic Model of Outer Membrane Fusion As mentioned above, the trans Fzo1 complex formation through anti-parallel interactions of coiled-coil regions is likely the key step in outer membrane fusion. However, it is as yet unknown what the subsequent mechanistic steps are. In fact, few details are known due to a lack of in  vitro structural studies on Fzo1. Since BDLP is closely related to Fzo1, the mechanistic function of Fzo1 may be similar to that of BDLP. We know that BDLP binds to, tubulates, and may also induce a compressed bilayer by distorting the lipid tails (Low and Lowe 2006; Low et al. 2009). GDP-bound BDLP has less affinity for the lipid bilayer and may possess a dimeric structure that differs from that of the GTP-bound form. Hence, a model is proposed that GTP binding induces the assembly and tubulation of membranes, and GTP hydrolysis leads to its depolymerization, leaving a highly curved outer leaflet in a high-energy state which will be stabilized upon fusion (Low et al. 2009). 1.2.2.2 Inner Membrane Fusion Mechanistic Model of Inner Membrane Fusion Mgm1/OPA1 Mgm1 was first identified in a screen for genes involved in mitochondrial genome maintenance. Mutations in MGM1 caused temperature-sensitive loss of mtDNA (Jones and Fangman 1992). Mgm1 was later shown to play a role in mitochondrial

1  The Genetics of Mitochondrial Fusion and Fission

27

fusion and subsequently shown to be a key mediator of inner membrane fusion. Deletion and mutations in MGM1 were shown to result in fragmented mitochondria and impaired mitochondrial fusion during mating that could be rescued by a DNM1 deletion (Wong et al. 2000, 2003). By an in vitro mitochondrial fusion assay, Mgm1 was shown to directly mediate inner membrane fusion. Temperature sensitive mutants of Mgm1 are defective in inner but not outer membrane fusion (Meeusen et al. 2004). Mgm1 is a nuclear-encoded protein containing an N-terminal mitochondrial targeting sequence (MTS) that is followed by two hydrophobic segments. It is targeted to the mitochondrial IM via its MTS and is proteolytically processed into two isoforms, l-Mgm1 and s-Mgm1, by the matrix processing peptidase (MPP) and the rhomboid serine protease Rbd1/Pcp1, respectively (McQuibban et al. 2003; Herlan et al. 2003). Maintaining approximately equal molar concentrations of l-Mgm1 and s-Mgm1 is crucial, and the formation of s-Mgm1 was shown to be ATP dependent (Herlan et al. 2004). Both isoforms are necessary for Mgm1 function, and they may serve different roles in the fusion process since l-Mgm1 does not require a functional GTPase domain and localizes more to the IM cristae (Zick et al. 2009). Like the other DRPs, Mgm1 consists of three core dynamin domains: the GTPase domain, a Middle domain and the GED. In addition, our studies suggest that Mgm1 may contain a lipid-binding domain located between middle and GED domain (De Vay et al. 2009; Rujiviphat et al. 2009). Consistent with other DRPs, Mgm1 function also relies on a functional GTPase domain and GED. Mitochondria of yeast strains containing mutations in the GTPase domain and the GED fail to fuse in the in vitro fusion assay (Meeusen et al. 2006). Both cis and trans Mgm1 self-interactions have been observed by immunoprecipitation and immuno-gold labeling thinsectioned EM (Zick et  al. 2009; Meeusen et  al. 2006). GTP hydrolysis, selfoligomerization and lipid interactions have been shown to be essential characteristics of Mgm1 since mutants lacking these activity in vitro have phenotypes similar to that of MGM1 deleted strain (Zick et  al. 2009; Rujiviphat et  al. 2009; Meeusen et al. 2006; Meglei and McQuibban 2009). The mammalian ortholog of Mgm1 is OPA1 whose mutations cause Autosomal Dominant Optic Atrophy (ADOA) (Delettre et  al. 2000). There are eight splice variants of OPA1 and each one undergoes at least one proteolytic processing event resulting in several long- and short-isoforms of OPA1 (Delettre et  al. 2001). In addition to the cleavage of OPA1 by PARL, the ortholog of Rbd1/Pcp1, the precursor of OPA1 is also cleaved by the intermembrane space and matrix AAA-proteases (McQuibban et  al. 2006; Ishihara et  al. 2006; Duvezin-Caubet et  al. 2007). The  exact functions of each isoform and specific OPA1 processing pathways remain to be elucidated. OPA1 has also been shown to hydrolyze GTP and was recently shown to have the ability to tubulate liposomes. In addition, purified bacterially-expressed OPA1 showed GTPase activity (Griparic and van der Bliek 2005). Like s-Mgm1, purified s-OPA1 also binds to phospholipids and possesses lipid-dependent stimulated GTPase activity (Ban et  al. 2010). Most importantly, these lipid-binding activities seem to be crucial for OPA1 function because several OPA1 disease alleles lack these characteristics (Ban et al. 2010).

28

E.Y.L. Chan et al.

How Does the Inner Membrane Really Fuse? Based on previous evidence from the in  vitro fusion assay, Mgm1 might mediate mitochondrial inner membrane fusion via a two-step mechanism: tethering and lipid mixing (Meeusen et al. 2006). The tethering step relies on the self-oligomerization of Mgm1 via the GED since a tight interface of the inner membranes following outer membrane fusion was not observed in Mgm1 with mutations in the GED by thinsectioned TEM images (Meeusen et al. 2006). This tethering step ensures the tight interface of the inner membranes to facilitate fusion. Similar to Fzo1, membrane tubulation may also facilitate the lipid-mixing step by increasing membrane curvature where membrane stress could be relieved upon fusion. Interestingly, liposome tubulation by OPA1 has been recently reported (Ban et al. 2010). Even though it is known that a functional GTPase domain is required for the lipid-mixing step, it is still unclear how the GTPase cycle comes into play in the fusion mechanism. Understanding how nucleotide hydrolysis is required for lipid fusion will be the next big advance in our understanding of how this class of mechanoenzymes works. Of note, GTPgS binding abolishes OPA1 liposome tubulation. Nevertheless, studies show that Mgm1/OPA1 can self-assemble into a crystalline array in the absence of GTP, indicating that GTP is not essential for binding or self-assembly onto the membrane (Rujiviphat et  al. 2009; Ban et  al. 2010). Moreover, tethering can still be observed in temperature-sensitive Mgm1 with a mutation in the GTPase domain (Meeusen et al. 2006). Hence, GTPase activity may cause changes in Mgm1/OPA1 structure and membrane topology during the lipid-mixing step. Further studies are required to understand this important aspect of Mgm1/OPA1 function and will be discussed at the end of this chapter. A possible model for how Mgm1/OPA1 works to fuse membranes is shown in Fig. 1.6. 1.2.2.3 Coordinating Outer and Inner Membrane Fusion Given the fact that mitochondria have four distinct biochemical compartments, and that disruption of this organization can lead to cell death by apoptosis, the coordination and regulation of membrane fusion is critically important. In yeast, there exists a relatively well-characterized protein that could serve a bridging function during this complex process. Ugo1 is a multi-spanning membrane protein that contains a carrier domain. Ugo1 is believed to serve as the bridge to physically tether Mgm1 and Fzo1. Ugo1 binds to both Fzo1 and Mgm1 independently and at different sites. In addition, the Fzo1-Mgm1 interaction is abolished in absence of Ugo1 (Sesaki and Jensen 2004). However, despite the identification of this fusion machinery protein complex, it is still unclear what the exact role of Ugo1 is. It is proposed that Ugo1 may have a structural role and that its interactions in the complex promote structural changes on Fzo1 to mediate fusion (Hoppins and Nunnari 2009). On the other hand, Ugo1 is also proposed to act in the lipid-mixing step (Hoppins and Nunnari 2009). Ugo1 was recently shown to be essential for both outer and inner membrane fusion but required after the tethering step (Hoppins et  al. 2009). To

1  The Genetics of Mitochondrial Fusion and Fission

29

Fig. 1.6  Mechanistic model of Mgm1-mediated membrane fusion: From the top left to the bottom right corner shows an averaged image of s-Mgm1 oligomeric rings to a proposed model of Mgm1mediated mitochondrial fusion. The background pattern shows s-Mgm1 assembled onto liposomes as oligomeric rings. An averaged image of the s-Mgm1 oligomer consists of six monomer densities (a). These six monomers can be interpreted as two stacks of trimers shown in red and yellow (b). These cstacking trimers could be how s-Mgm1 tethers two opposing membranes to mediate membrane fusion (c and d)

date, no human homolog of Ugo1 has been identified. Due to the critical role that Ugo1 plays in fusion, it is proposed that a protein with a similar function to Ugo1 is required for bridging OPA1 and MFN1/2 activities and plays a role in mammalian mitochondrial fusion. The identity and characterization of this key biochemical activity is needed to fully dissect mammalian mitochondrial fusion and represents a key missing piece of the puzzle.

1.2.3 Regulatory Pathways of Membrane Fusion 1.2.3.1 Ubiquitination It is well-known that proper mitochondrial morphology requires the correct maintenance of Fzo1 protein levels and that this is regulated by 26S proteasome degradation (Hoppins and Nunnari 2009). It is also known that treatment with the

30

E.Y.L. Chan et al.

alpha-factor pheromone triggers Fzo1 proteasome-dependent degradation (Hoppins and Nunnari 2009). Further, Fzo1 is known to be ubiquitinated and degraded in an Mdm30-dependent manner (Hoppins and Nunnari 2009). Mdm30 is an F-box containing protein and is a component of the SCF E3 ubiquitin ligase complex that ubiquitinates Fzo1 and targets it for degradation by the 26S proteasome. However, there is conflicting evidence that ubiquitinated Fzo1 degradation is independent of the SCF complex and the 26S proteasome. In addition to its role in Drp1 stability, the E3 ubiquitin ligase MARCH-V has been shown to interact with the pro-fusion protein MFN2 (Nakamura et al. 2006), and MFN1 was recently shown to be its main ubiquitination substrate (Park et al. 2010). Further, a ubiquitinated form of OPA1 has also been observed, but the pathway or significance of this is not yet known. These recent demonstrations are summarized in Fig. 1.7. These data that demonstrate the key role of adding ubiquitins to these fusion molecules begs the question: what is the de-ubiquitylation factor and how must it contribute to this aspect of mitochondrial regulation and dynamics? 1.2.3.2 Proteolytic Processing Alternative topogenesis is the widely-accepted model of Mgm1 processing (Herlan et al. 2004). In this model, the processing of Mgm1 is regulated by ATP levels and the mitochondrial membrane potential (Herlan et al. 2004). Likewise, it has been demonstrated that mitochondrial ATP levels and membrane potential are important factors for OPA1 processing (Guillery et  al. 2008; Baricault et  al. 2007). In an attempt to identify proteins involved in Mgm1 processing, Ups1 was identified. Knocking-out Ups1 led to the abolishment of s-Mgm1 formation due to a defect in Mgm1 sorting (Sesaki et  al. 2006). Instead of having a regulatory role on Rbd1/ Pcp1 activity, Ups1 appears to play a role in cardiolipin homeostasis. This key mitochondrial lipid is required for stabilizing the TIM23-PAM complex that is a key molecular component of the mitochondrial inner membrane import system (Tamura et al. 2009). Yet another family of proteins, the prohibitins are known to impact processing of Mgm1/OPA1. In this case, knocking-out Phb2 in mice leads to a reduction in the OPA1 long isoform (Merkwirth et  al. 2008). Interestingly, recent studies show that Phb2 forms a multimeric complex with Phb1 and together they act as a scaffold in the inner membrane. Taken together, these data underscore the complexity and diversity of regulating the processing of the key machineries of mitochondrial membrane fusion. 1.2.3.3 Lipid Metabolism There has been increasing evidence demonstrating the cross talk between lipid metabolism and mitochondrial membrane dynamics (Furt and Moreau 2009). Knocking-down MitoPLD, the enzyme that converts cardiolipin (CL) to phosphatidic acid (PA), leads to less fusion, and its over-expression leads to aggregation of mitochondria (Choi et al. 2006). Interestingly, these mitochondria are separated by

1  The Genetics of Mitochondrial Fusion and Fission

31

Fig. 1.7  Regulation of the mammalian mitochondrial fusion and fission machinery: The diagrams illustrate elucidated proteins with implicated regulatory roles in mammalian mitochondrial fusion and fission. Arrowhead bold lines represent well-established positive regulations. Arrowhead dash lines represent possible positive regulations. Blunt end bold lines represent negative regulations. (a) The regulation of mitochondrial fusion. MARCH-V interacts and ubiquitinates MFN1, and the degradation of MFN1 is proteasome-dependent. MitoPLD is required for fusion and promotes the tethering of mitochondria in an MFN-dependent manner. PSID was shown to be essential for mitochondrial fusion but the exact function is still unknown. Since PSID converts PS to PE, providing negative membrane curvature, it may play a role in fusion by altering the lipid composition. Proteins in the Bcl-2 family have implicated roles in fusion. Members like Bax and Bak are profusion proteins under normal conditions but pro-fission in apoptotic cells. The Bcl-2 protein may modulate MFN and OPA1 functions in mitochondrial fusion. The function of Bcl-2 proteins is also linked to cytochrome c release that could be due to outer membrane permeabilization or cristae junction disruption. (b) The regulations on mitochondrial fission. hFis1 functions in the localization of DRP1, but the recruitment of Drp1 is not entirely dependent on hFis1. DRP1 and hFis1 are ubiquitinated and degraded in a proteasome-dependent manner. E3 ligases found to mediate this ubiquitination are MARCH-V and Parkin. Since GDAP1 has an implicated role in hFis1 activity and lipid homeostasis, GDAP1 may play a role in fission by modulating hFis1 function or lipid composition. Drp1 is regulated by sumoylation. Sumoylation with SUMO1 by MAPL stabilizes Drp1 and promotes fission. SENP5 is required for DRP1 desumoylation. Phosphorylation is another known regulator of DRP1. DRP1 is phosphorylated by several kinases. MTP18 affects DRP1-dependent fission but its exact function is unknown. There are speculations that MTP18 might mediate inner membrane fission or is involved in fission complex formation. Bcl-2 proteins also have implicated roles in modulating DRP1 function and mitochondrial fission

a distance equivalent to half that of heptad repeat-mediated tethered mitochondria. Therefore, these aggregated mitochondria may be due to an excess of tethering but a complete lack of the lipid mixing suggesting that MitoPLD may function in the lipid-mixing step of outer membrane fusion. Recently, there have been several findings that connect the functional relationship between phospholipid metabolism with Mgm1 processing and mitochondrial

32

E.Y.L. Chan et al.

morphology. Ups1, which has an implicated role in Mgm1 sorting/processing, was shown to act antagonistically with Ups2/Gep1 in regulating cardiolipin levels (Tamura et al. 2009). Phb1 that forms into a multimeric complex with Phb2 was shown to genetically interact with several genes involved in mitochondrial morphology maintenance such as MDM10 and MMM1. PHB1 also has genetic interactions with UPS1, GEP1 as well as genes directly involved in lipid metabolism, CRD1 and PSD1. Altered l- and s-Mgm1 ratios were observed in GEP1, PSD1 and CRD1 knock-out strains (Osman et al. 2009). It is clear that there is still much to discover with respect to the control, orchestration and regulation of mitochondrial membrane fusion. With respect to mitochondrial membrane fission events, the core machinery as described above suggests that cytoplasmic regulators exist for this complex process. In particular, fission factors are almost all cytoplasmic facing, making them very accessible to many post-translational modifying enzymes of the cell. Now, we will discuss the best-characterized activities in regulating the division of mitochondria.

1.2.4 Regulatory Pathways of Membrane Fission 1.2.4.1 Sumoylation Drp1 is covalently conjugated to a small ubiquitin-like modifier SUMO1. Sumoylation of Drp1 by MAPL protects it from degradation, increasing mitochondrial fragmentation (Harder et  al. 2004; Braschi et  al. 2009). Although Drp1 is transiently sumoylated at steady state, induction of apoptosis leads to increased Drp1 sumoylation in a Bax/Bak-dependent fashion. This increase in Drp1 stability on mitochondria also coincides with Bax localization to the mitochondrial membrane (Wasiak et al. 2007). Interestingly, endophilin B1, a fatty acyl transferase, has been shown to regulate Bax localization to mitochondria (Takahashi et al. 2005). Knock-down of endophilin B1 by RNAi caused mitochondrial elongation, implying that it is a pro-fission factor (Karbowski 2004). Together, these data suggest that endophilin B1 might regulate Bax-dependent sumoylation of Drp1, increasing its stability to promote mitochondrial fission. SENP5 is a SUMO protease that catalyzes the removal of Sumo1 from mitochondrial proteins including Drp1. Over-expression of SENP5 led to increased removal of Sumo1 from Drp1 and destabilization of the protein, promoting its degradation. As a result, mitochondria became elongated. In contrast, knockingdown SENP5 by shRNA resulted in increased Drp1 sumoylation and consequent mitochondrial fragmentation (Zunino et al. 2007). Besides steady state desumoylation of Drp1, SENP5 also desumoylates Drp1 during the G2/M transition of the cell cycle. As previously demonstrated, regulating mitochondrial fission during the cell cycle is intimately linked to the regulation of Drp1 (Taguchi et al. 2007). Drp1 is phosphorylated during the G1/S transition to ensure mitochondrial fragmentation in early mitosis and proper mitochondrial inheritance to daughter

1  The Genetics of Mitochondrial Fusion and Fission

33

cells (Taguchi et al. 2007; Boldogh et al. 2001). SENP5 desumoylation of Drp1 during mitosis increases Drp1 oligomerization, facilitating mitochondrial fission (Zunino et  al. 2009). Although desumoylation during the G2/M transition increases fission, steady state desumoylation of Drp1 promoted mitochondrial elongation (Zunino et al. 2007, 2009). These data underscore the importance of this post-translational modification, and it is likely that several other factors undergo sumoylation to regulate mitochondrial division in the different contexts during the life of the cell. 1.2.4.2 Ubiquitination It is becoming increasingly clear that ubiquitination is a key regulator of mitochondrial function. Later in the chapter, we will discuss an emerging theme in mitochondrial research – quality control that is intimately regulated by ubiquitination. With respect to fusion and fission, MARCH-V (also known as MARCH5 and MITOL) is a mitochondrial OM-localized E3 ubiquitin ligase with an N-terminal RING finger motif (Nakamura et  al. 2006; Yonashiro et  al. 2006). It binds to and ubiquitinates hFis1 and Drp1, promoting their degradation (Nakamura et al. 2006; Yonashiro et al. 2006). hFis1 is ubiquinated in cells expressing WT MARCH-V, but not a catalytically inactive form of the protein. Similarly, Drp1 was also ubiquitinated by WT MARCH-V, albeit to a lesser degree than hFis1, possibly owing to its primarily cytosolic localization (Yonashiro et al. 2006). It has also been well established that the pro-fusion proteins Fzo1/MFN are subject to ubiquitination (Hoppins and Nunnari 2009). As more studies are published, the list of ubiquitin-tagged mitochondrial proteins will grow. Interestingly, there have been very few reports that characterize the enzymes that remove ubiquitin (the so-called DUBs) from their substrates. 1.2.4.3 Phosphorylation During mitosis, mitochondria undergo morphological changes to ensure proper inheritance to daughter cells (Boldogh et al. 2001). In HeLa cells, the tubular mitochondrial network becomes fragmented in early mitosis and reforms into an interconnected network after telophase and cytokinesis. This early mitotic fragmentation is Drp1-dependent. Biochemical analyses indicated that Drp1 is phosphorylated by Cdk1/cyclin B during the G1/S transition on S616 within the Drp1 GED. Mutating S616 to an alanine (Drp1S616A) abolished phosphorylation and resulted in a less fragmented mitochondrial network in early mitosis. This demonstrates that Cdk1/ cyclin B phosphorylation of Drp1 during early mitosis promotes mitochondrial fission (Taguchi et al. 2007). Drp1 is also phosphorylated by the cAMP-dependent kinase (PKA) and the Ca2+/ calmodulin-dependent protein kinase Ia (CaMKIa) at S637 (Han et al. 2008; Chang and Blackstone 2007; Cribbs and Strack 2007). PKA phosphorylation negatively regulates Drp1 intramolecular interactions and GTPase activity. Consequently,

34

E.Y.L. Chan et al.

over-expression of phosphomimetic mutants prevented mitochondrial fission, resulting in the accumulation of highly interconnected mitochondria (Chang and Blackstone 2007; Cribbs and Strack 2007). S637 is also the phosphorylation site for CaMKIa. CaMKIa phosphorylation can be selectively stimulated by treatment with high K+. However, unlike phosphorylation by PKA that reduced fission, phosphorylation by CaMKIa resulted in increased recruitment of Drp1 to mitochondria and a consequent increase in mitochondrial fission (Han et  al. 2008; Cribbs and Strack 2007). Due to the importance in maintaining proper mitochondrial dynamics, this discrepancy might reflect the fine-tuned regulation of Drp1-dependent mitochondrial fission. Since the Drp1 sequence around S637 is highly conserved, phosphorylation on S637 might serve as a universal regulatory mechanism for Dnm1/Drp1 across species (Han et  al. 2008; Chang and Blackstone 2007; Cribbs and Strack 2007). Unexpectedly, the phosphomimetic mutant of S616 (S616D) did not show any change in intra- or intermolecular interactions, GTPase activity or mitochondrial morphology (Chang and Blackstone 2007). However, since phosphorylation on S616 is a cell cycle-dependent event, the S616D mutation might not have physiological significance in this study and hence resulted in no phenotype abnormalities. Phosphorylation on S637 is a reversible event mediated by the Ca2+-dependent phosphatase calcineurin (Cribbs and Strack 2007; Cereghetti et al. 2008). Mitochondrial depolarization results in fragmentation and concomitant calcineurin dephosphorylation of Drp1 on S637. Inhibition of calcineurin prevents mitochondrial fragmentation, indicating that dephosphorylation of Drp1 promotes mitochondrial fission (Cereghetti et al. 2008). These results are consistent with the anti-fission effect of Drp1 phosphorylation by PKA (Chang and Blackstone 2007; Cribbs and Strack 2007). Accordingly, the Drp1S637D phosphomimetic mutant was almost exclusively cytosolic while the Drp1S637A dephosphomimetic mutant localized to mitochondria. This demonstrates that dephosphorylation of Drp1 results in its recruitment to mitochondria promoting fission (Cereghetti et al. 2008). Although it is unclear whether calcineurin dephosphorylation has kinase specificity, these data suggest that calcineurin might selectively dephosphorylate PKA-dependent phospho-Drp1. Recently, protein phosphatase 2A (PP2A) and its regulatory subunit Bb2 were identified as regulators of mitochondrial fission. Upon induction of apoptosis, Bb2 recruited PP2A to the mitochondrial surface and promoted mitochondrial fission in a Drp1- and Fis1-dependent manner (Dagda et al. 2008). Although the mechanism of PP2A/Bb2 in the regulation of mitochondrial fission was not examined, a possibility might be the dephosphorylation of Drp1 by PP2A. This would be consistent with calcineurin dephosphorylation of Drp1 promoting mitochondrial fission (Cereghetti et al. 2008). 1.2.4.4 S-Nitrosylation Another post-translational modification of Drp1 is S-nitrosylation. This is not surprising since dynamins have been shown to be S-nitrosylated (Kang-Decker et al. 2007; Wang et al. 2006). Nitric oxide produced in response to the Ab peptide implicated in Alzheimer’s disease resulted in S-nitrosylation of Drp1. A C644A

1  The Genetics of Mitochondrial Fusion and Fission

35

mutation that abrogated Drp1 S-nitrosylation prevented Ab-induced neurotoxicity. S-nitrosylation of Drp1 induces mitochondrial fission by promoting Drp1 dimerization and increased GTPase activity (Cho et al. 2009). It is clear that mitochondrial fusion and fission is a highly complex, and highly regulated process. While researchers can all agree that the mitochondrion is a very dynamic compartment, there is less consensus regarding the role for ­mitochondrial dynamics, in either a single cell like yeast, or in the context of a metazoan such as ourselves.

1.3 Possible Roles of Mitochondrial Dynamics The remaining chapters in this book will discuss in detail the various contexts that the cell utilizes mitochondrial dynamics, and the resulting pathologies that incur when these dynamics are perturbed. However, we will discuss very briefly some of the more well-characterized cell biological roles for mitochondrial dynamics in every cell.

1.3.1 Roles in Cell Cycle and Development In yeast, mitochondrial fission and fusion is important during budding and mating. In the same way, mitochondrial dynamics are crucial for cell development in higher eukaryotes and embryogenesis in mammals. Mitochondrial dynamics are also important for several cellular processes. Mitochondrial dynamics supports intracellular mitochondrial motility. These dynamics are required for the distribution of mitochondria to supply diverse bioenergetic requirements especially in specialized cells. For instance, mitochondrial fusion and fission are required for distributing mitochondria to neuron dendrites and axon terminals for synaptic activity (Kann and Kovacs 2007). Mitochondrial dynamics have a direct role in the apoptosis pathway (see Fig. 1.2). Specifically, cytochrome c that is normally stored in cristae is released to the cytosol and activates caspases that in turn switch on the apoptosis pathways. Up-regulated mitochondrial fission leads to more cytochrome c release and cell death, and up-regulated mitochondrial fusion can suppress the induction of apoptosis (Kann and Kovacs 2007).

1.3.2 Roles in Mitochondrial Genome Maintenance Due to the nature of the mitochondrial nucleiod, ROS production from the electron transport chain (ETC) and mtDNA replication/repair systems, the mitochondrial genome is prone to DNA damages. Mitochondrial dynamics may have evolved to diminish these damages and maintain the integrity of the mitochondrial genome. Since the mitochondrial genome encodes subunits of the complexes of the ETC,

36

E.Y.L. Chan et al.

mtDNA mutations can lead to impaired ETC and ATP production. Possibly as a means of cross-complimentation, fusion of a mitochondrion containing undamaged mitochondrial DNA could functionally replace the damaged component to result in the rapid recovery of ATP production. In addition, mixing of mitochondrial genomes allows transcomplementation of mtDNA and the dilution and tolerance of mtDNA mutations (Schon and Gilkerson 2010; Chen et  al. 2010). On the other hand, fission may lead to selecting out malfunctioning mitochondria (see Fig. 1.1). In support of this idea, studies show that only transient but not complete mitochondrial fusion was observed in OPA1 knock-down or over-expressing cells (Liu et al. 2009), and mitochondria with abnormal shape have a reduced ability to fuse to normal ones (Navratil et al. 2008).

1.3.3 Roles in Quality Control Systems Mitophagy and apoptosis are quality control mechanisms at organellar and cellular levels, respectively. Mitophagy is a mechanism by which severely damaged mitochondria are degraded to prevent them from fusing and sharing toxic material with other mitochondria in the network (discussed in more detail below). Impaired mitophagy can have a deleterious effect on cellular processes and result in the activation of apoptosis. Apoptosis prevents the spreading and accumulation of these defective cells and can rescue the organism from the pathology of necrosis. In this way, the cell has a means to either rescue a cell, or commit it to death if the damage is too great (see Fig. 1.8). It is becoming more and more realized that there is a tight relationship between maintaining mitochondrial dynamics, functions and various quality control systems. Specifically, impaired mitochondrial dynamics causes mitochondrial dysfunction. At the same time, mitochondrial dysfunction can affect OPA1 processing leading to fragmentation and sequestering of damaged mitochondria by mitophagy (Tatsuta and Langer 2008). In apoptotic cells, BCL2-like proteins positively affect DRP1 activity but negatively affect MFN2 and OPA1 activity causing ­mitochondrial fragmentation. This fragmentation facilitates mitophagy and the release of cytochrome c to activate apoptosis (Fig.  1.8) (Tatsuta and Langer 2008). In general, the cell has evolved subtle, refined and elegant ways to preserve function.

1.4 Disruption of Mitochondrial Dynamics Causes Neurodegeneration For many years it was not clear why mitochondrial disruption led to an impairment of neuronal function. However, several key aspects of the neuronal system make it vulnerable to even minor deficiencies in mitochondrial dynamics and activity that

1  The Genetics of Mitochondrial Fusion and Fission

37

Fig. 1.8  Roles of mitochondrial dynamics in genome maintenance and organellar and cellular quality controls: Mitochondria are prone to cellular stresses, DNA damages and reactive oxygen species (ROS) that can cause mutations in mitochondrial DNA (mtDNA). The mitochondrial genome (double-helix circles) encodes for subunits of complexes in the electron transport chain, therefore damage of the mitochondrial genome (colored in black) also leads to impaired oxidative phosphorylation (OXPHOS). These deficiencies can be rescued by mitochondrial fusion (blue arrows) and fission (red arrows) events. On the left side, mitochondrial fusion results in a rapid energy supply and replenishment of impaired OXPHOS by providing intact essential proteins. After several rounds of mtDNA replication and mitochondrial fusion, mtDNA complementation and dilution of mutated mtDNA occur (colored in grey) to maintain the integrity of the mitochondrial genome. On the right side, mitochondrial fission also has a protective role. Mitochondrial fission may selectively (or by chance) segregate out mitochondria that contain damaged genomes. These mitochondria can undergo several rounds of fusion to complement their defects as mentioned above. Alternatively, severely damaged mitochondria are subject to mitophagy where fission/fragmentation facilitates engulfment by the autophagosome. If the damage is too excessive, fission also leads to the release of cytochrome c that activates caspases and subjects the cell to apoptosis, and the game is over

include: (1) the requirement for the distribution of mitochondria to distant axon terminal and the dendrite; (2) the requirement of mitochondrial ATP production and Ca2+ buffering for proper neuronal activity; (3) accumulated mtDNA mutational load is common in most neurodegenerative diseases, likely due to (2); and (4) the

38

E.Y.L. Chan et al.

Fig. 1.9  Disruption in mitochondrial dynamics causes neurodegeneration. (a) Fusion and fission are required for distributing mitochondria to axon terminals and dendrites. In normal cells, mitochondria, shown as white elongated shapes, are distributed throughout the neuron and enriched around terminal buttons to supply ATP for synaptic activity. In fusion and fission defective neurons, mitochondria do not get distributed but get concentrated in the cell body. Fragmentation and elongation due to lack of fusion and fission, respectively, also cause mitochondria to accumulate damages and to malfunction, colored in black. (b) Abnormality in mitochondrial dynamics causes neurodegeneration. As described in (a), mitochondrial dynamics are essential for mitochondrial distribution to synaptic sites. Poor mitochondrial distribution causes insufficient ATP production and Ca2+ buffering that are crucial for synapses. Hence, synaptic dysfunction results in the aftermath of disrupted mitochondrial dynamics, leading to neurodegeneration. In addition, abnormal mitochondrial dynamics, especially fragmentation, causes unregulated apoptosis that leads to neuronal cell death, and once again, it’s game over

quality control system given that unregulated apoptosis induced by disrupted ­mitochondrial dynamics and excessive mitophagy leads to neuronal cell death with no regeneration system in place. All of these aspects can lead to synaptic degeneration and neuronal cell death (see Fig. 1.9). Several chapters in this book will describe these events in detail, and in the context of specific human diseases.

1.5 Future of the Field and Compelling Questions 1.5.1 Compelling Questions in Mitochondrial Fusion and Fission Events This past decade has seen a true resurgence in mitochondrial research that has elaborated on many aspects of mitochondrial dynamics. Genetic studies have identified key players and their regulators while biochemical studies have provided

1  The Genetics of Mitochondrial Fusion and Fission

39

insight into their functions and mechanisms. However, several questions remain to be answered with respect to both regulatory pathways and detailed molecular mechanisms of fusion and fission. Even though we know quite a bit about both the molecular players of mitochondrial fission and how it works at a biophysical level, many genes implicated in the fission event still have unknown functions. For instance, besides its role in lipid metabolism, the exact role of GDAP1 in mitochondrial fission is still unknown. Mutations in GDAP1 cause CMT4A implying that GDAP1 plays a significant role in dynamics. Thus, studies on GDAP1 may provide insight into regulation of the fission event and pathophysiology underlining neurodegenerative diseases. Further studies on the phosphorylation regulation of Drp1 will shed light on the synchronization of mitochondrial dynamics with the cell cycle and with signaling by kinases. In addition, PINK1, which is linked to familial Parkinson’s disease (PD), is one of the kinases that can modulate mitochondrial fission in a Drp1-dependent manner. It is still unclear whether this is due to direct phosphorylation by PINK1 on fission molecules or an indirect involvement of a kinase cascade. More understanding of PINK1 will provide a clearer picture on fission regulation and the functional role of PINK1 in PD. Loss-of-function approaches by a double knock-down of PINK1 with other candidates in the kinase cascade or fusion/fission genes may provide more insight into its pathway in relation to mitochondrial dynamics. With respect to studies on mitochondrial fusion, many questions remain unanswered. It is still unclear how outer and inner membrane fusions occur as separate events but yet coordinated. Specific studies that reveal how Fzo1/MFN and all the isoforms of Mgm1/ OPA1 mediate fusion will clarify the overall picture of mitochondrial fusion and provide more insight into the role of these previously identified genes. For instance, to determine whether the role of Ugo1 in fusion is indeed in the lipid-mixing step of Fzo1 and Mgm1, we need to first understand how Fzo1- and Mgm1-mediated fusion works. In addition, Ugo1 at present appears to be fungal only. Do we have a Ugo1 ortholog, or have we evolved a different mechanism of fusion? Perhaps OPA1 is the answer. Indeed, why are there only two isoforms of Mgm1 as compared to eight for OPA1 with the complicating factor that several proteases have been identified to process OPA1? Synthetic genetic array or multi-copy suppressor screens could be ways to reveal additional genetic interactors that may be important in mitochondrial fusion in yeast. With respect to the challenge of OPA1, it is difficult to truly understand how and why the OPA1 isoforms have come about. One approach could be to determine the expression pattern of all the isoforms and correlate that with all of the putative OPA1 proteases. This could be done in different cell types and stress conditions to determine whether there is a regulation at post-transcriptional or post-translational level. These types of studies are required to truly understand this most complex process. Recent studies have also indicated that lipid metabolism may be another main regulatory pathway in mitochondrial dynamics. Several genes implicated in mitochondrial fusion and fission such as Endophilin B1, GDAP1, mitoPLD, PSD1 and PHB1/2, also have a role in lipid metabolism. Besides the speculation that some phospholipids can create more membrane curvature than others, the exact functional relationship is unclear. Future studies on lipid homeostasis are necessary

40

E.Y.L. Chan et al.

since an important aspect of mitochondrial function is lipid metabolism and the functions of fusion/fission molecules are modulated by their interactions with lipids and overall lipid compositions of the mitochondrial membranes. Identifying genetic and physical interactors will provide greater insight into the pathways interconnecting lipid homeostasis with mitochondrial dynamics.

1.6 The Most Recent and Important Discoveries in Mitochondrial Dynamics While the last 100 years of mitochondrial research has revealed several key discoveries, just in the last couple of years or so mitochondrial research has undergone a true resurgence with respect to understanding both the homeostasis and the pathology of mitochondrial activity within the cell. It appears that since the endosymbiotic event, the host cell has evolved an elegant mechanism to make sure its mitochondrial content remains as healthy as possible. We have already mentioned this quality control mechanism of autophagic removal of damaged mitochondria, now termed mitophagy (Tatsuta and Langer 2008). It is our belief that this discovery will truly revolutionalize and direct mitochondrial research for the next decade (even in the context of what we feel our three key outstanding questions discussed below). It is the simplicity of design in the quality control mechanism that is most appealing. Mitochondria are very harsh environments where proteins, lipids, and nucleic acids are constantly damaged. The cell must maintain a healthy mitochondrial network to thrive. In order to isolate and remove damaged compartments from the network, selective fission occurs. These newly formed mitochondria, due to their damage, can no longer provide energy nor maintain a potential across its inner membrane. This results in the deficient organelle being isolated as it can no longer undergo fusion with the rest of the network. The cell recognizes this damaged entity and flags it with ubiquitin moieties. This serves to recruit the autophagosomal machinery, with the end result of lysosomal destruction and recycling of basic building blocks back into the cytoplasm. Of course it all makes sense, but there are several key and obvious gaps in our understanding of the molecular mechanisms at play. This will be discussed in greater detail in Chap. 3.

1.7 Future Discoveries that will Advance the Field of Mitochondrial Dynamics 1.7.1 How Does Mitochondrial Fusion Work? While we, as authors of this chapter, may be biased towards this question as it is one of our main research pursuits, we feel it is still of fundamental importance to understand the biophysical details of membrane fusion. As more and more components

1  The Genetics of Mitochondrial Fusion and Fission

41

of the molecular machinery are identified, we should be able to formulate testable hypotheses as to how these complexes orchestrate the regulated mixing of membranes. Beyond understanding how a protein like Mgm1 and its two isoforms function to fuse the membranes in terms of torsional forces, figuring out its regulation is key. A mitochondrion is filled with folded invaginations of inner membrane cristae that remain distinct and do not undergo spontaneous fusion. However, when two mitochondria fuse their outer membranes, the respective inner membranes now undergo rapid and efficient fusion. What signal is sent or received to up-regulate the activity of Mgm1-mediated inner membrane fusion? As more detailed studies are conducted on additional components and the regulatory GTP cycle, we should be able to elucidate this basic activity.

1.7.2 How Is the Mitochondrial Genome Compartmentalized and Maintained? As stated in the introduction to this chapter, researchers have known for quite some time that mitochondria house their own genome. Over time the majority of the mitochondrial genome has been incorporated into the host’s nuclear genome. Most species have retained only a handful of genes on their mitochondrial genome, and all are components of the energy production system. Beyond understanding which genes have been retained (which in and of itself is an interesting and important question), figuring out how the genome is compartmentalized (the so-called mitochondrial nucleoid) is extremely important to elucidate mechanisms of inheritance and DNA quality control. It has long been known that as an organism ages it accumulates mutations in its mitochondrial genome largely due to the harsh chemical environment within the organelle. In 2004, a groundbreaking study directly demonstrated that accumulation of mutations in the mitochondrial genome directly resulted in aging in the mouse (Trifunovic et al. 2004; Chan 2006). Together, these data support the notion that maintaining the fidelity of the nucleotide sequence, as well as ensuring faithful transmission during cellular division is of the utmost importance for the well-being of the cell and organism. These functions must occur in a regulated manner, both spatially in the context of the nucleiod, and temporally with respect to the cell cycle. Our knowledge of these processes is currently in its infancy so we predict many more groundbreaking studies with respect to the mitochondrial genome.

1.7.3 How Is the Mitochondrion Integrated into the Rest of the Cellular Networks? We have come a long way from considering the mitochondrion to be an autonomous organelle that drifts amidst the cytoplasm generating the bulk of energy for its host. Perhaps the most convincing notion of mitochondrial integration into the life

42

E.Y.L. Chan et al.

of the cell is its pivotal role in directing apoptosis. While this was a key discovery, most cells are not undergoing apoptosis, as this represents extreme modes of developmental biology and cellular response to stress. One can imagine several homeostatic mechanisms that would rely on active signalling between the mitochondria and the cytoplasm, the cell surface, the nucleus, peroxisomes, etc. As genetic networks are characterized and protein complexes are understood, we should be able to unravel the vast array of mitochondrial directed activities in a normal healthy cell. It is our prediction that the next 100 years of mitochondrial research will be as exciting and informative as the last 100 years. Acknowledgments  Research in the McQuibban lab is supported by grants from the Canadian Institutes of Health Research, the Natural Sciences and Engineering Research Council of Canada, and the Parkinson Society of Canada.

References Alexander, C., Votruba, M., Pesch, U. E., Thiselton, D. L., Mayer, S., Moore, A., Rodriguez, M., Kellner, U., Leo-Kottler, B., Auburger, G., Bhattacharya, S. S., and Wissinger, B. (2000) Nat Genet 26, 211–215 Altmann, K. (2005) Molecular Biology of the Cell 16, 5410–5417 Ban, T., Heymann, J. A., Song, Z., Hinshaw, J. E., and Chan, D. C. (2010) Hum Mol Genet 19, 2113–2122 Baricault, L., Segui, B., Guegand, L., Olichon, A., Valette, A., Larminat, F., and Lenaers, G. (2007) Exp Cell Res 313, 3800–3808 Bartley, W., and Davies, R. E. (1954) Biochem J 57, 37–49 Baxter, R. V., Ben Othmane, K., Rochelle, J. M., Stajich, J. E., Hulette, C., Dew-Knight, S., Hentati, F., Ben Hamida, M., Bel, S., Stenger, J. E., Gilbert, J. R., Pericak-Vance, M. A., and Vance, J. M. (2002) Nat Genet 30, 21–22 Benard, G., and Karbowski, M. (2009) Semin Cell Dev Biol 20, 365–374 Bereiter-Hahn, J. (1990) Int Rev Cytol 122, 1–63 Bereiter-Hahn, J., and Voth, M. (1994) Microsc Res Tech 27, 198–219 Bhar, D. (2006) J Biol Chem 281, 17312–17320 Blatch, G. L., and Lassle, M. (1999) Bioessays 21, 932–939 Bleazard, W., McCaffery, J. M., King, E. J., Bale, S., Mozdy, A., Tieu, Q., Nunnari, J., and Shaw, J. M. (1999) Nat Cell Biol 1, 298–304 Boldogh, I. R., Yang, H. C., and Pon, L. A. (2001) Traffic 2, 368–374 Braschi, E., Zunino, R., and McBride, H. M. (2009) EMBO reports 10, 748–754 Burgess, S. M., Delannoy, M., and Jensen, R. E. (1994) J Cell Biol 126, 1375–1391 Cereghetti, G. M., Stangherlin, A., de Brito, O. M., Chang, C. R., Blackstone, C., Bernardi, P., and Scorrano, L. (2008) Proceedings of the National Academy of Sciences 105, 15803–15808 Cerveny, K. L. (2003) Molecular Biology of the Cell 14, 4126–4139 Cerveny, K. L., McCaffery, J. M., and Jensen, R. E. (2001) Mol Biol Cell 12, 309–321 Cerveny, K. L., Studer, S. L., Jensen, R. E., and Sesaki, H. (2007) Dev Cell 12, 363–375 Chan, D. C. (2006) Cell 125, 1241–1252 Chang, C. R., and Blackstone, C. (2007) J Biol Chem 282, 21583–21587 Chang, C. R., Manlandro, C. M., Arnoult, D., Stadler, J., Posey, A. E., Hill, R. B., and Blackstone, C. (2010) J Biol Chem 285, 32494–32503

1  The Genetics of Mitochondrial Fusion and Fission

43

Chen, H. (2003) J Cell Biol 160, 189–200 Chen, H. (2005) J Biol Chem 280, 26185–26192 Chen, H., and Chan, D. C. (2009) Hum Mol Genet 18, R169-176 Chen, H., Detmer, S. A., Ewald, A. J., Griffin, E. E., Fraser, S. E., and Chan, D. C. (2003) J Cell Biol 160, 189–200 Chen, H., Vermulst, M., Wang, Y. E., Chomyn, A., Prolla, T. A., McCaffery, J. M., and Chan, D. C. (2010) Cell 141, 280–289 Cho, D. H., Nakamura, T., Fang, J., Cieplak, P., Godzik, A., Gu, Z., and Lipton, S. A. (2009) Science 324, 102–105 Choi, S. Y., Huang, P., Jenkins, G. M., Chan, D. C., Schiller, J., and Frohman, M. A. (2006) Nat Cell Biol 8, 1255–1262 Cribbs, J. T., and Strack, S. (2007) EMBO reports 8, 939–944 Cuesta, A., Pedrola, L., Sevilla, T., Garcia-Planells, J., Chumillas, M. J., Mayordomo, F., LeGuern, E., Marin, I., Vilchez, J. J., and Palau, F. (2002) Nat Genet 30, 22–25 Dagda, R. K., Merrill, R. A., Cribbs, J. T., Chen, Y., Hell, J. W., Usachev, Y. M., and Strack, S. (2008) Journal of Biological Chemistry 283, 36241–36248 Davies, V. J., Hollins, A. J., Piechota, M. J., Yip, W., Davies, J. R., White, K. E., Nicols, P. P., Boulton, M. E., and Votruba, M. (2007) Hum Mol Genet 16, 1307–1318 Delettre, C., Lenaers, G., Griffoin, J. M., Gigarel, N., Lorenzo, C., Belenguer, P., Pelloquin, L., Grosgeorge, J., Turc-Carel, C., Perret, E., Astarie-Dequeker, C., Lasquellec, L., Arnaud, B., Ducommun, B., Kaplan, J., and Hamel, C. P. (2000) Nat Genet 26, 207–210 Delettre, C., Griffoin, J. M., Kaplan, J., Dollfus, H., Lorenz, B., Faivre, L., Lenaers, G., Belenguer, P., and Hamel, C. P. (2001) Hum Genet 109, 584–591 DeVay, R. M., Dominguez-Ramirez, L., Lackner, L. L., Hoppins, S., Stahlberg, H., and Nunnari, J. (2009) J Cell Biol 186, 793–803 Di Maria, E., Gulli, R., Balestra, P., Cassandrini, D., Pigullo, S., Doria-Lamba, L., Bado, M., Schenone, A., Ajmar, F., Mandich, P., and Bellone, E. (2004) J Neurol Neurosurg Psychiatry 75, 1495–1498 Dimmer, K. S. (2002) Molecular Biology of the Cell 13, 847–853 Dimmer, K. S., Navoni, F., Casarin, A., Trevisson, E., Endele, S., Winterpacht, A., Salviati, L., and Scorrano, L. (2007) Human Molecular Genetics 17, 201–214 Dohm, J. A., Lee, S. J., Hardwick, J. M., Hill, R. B., and Gittis, A. G. (2004) Proteins 54, 153–156 Duvezin-Caubet, S., Koppen, M., Wagener, J., Zick, M., Israel, L., Bernacchia, A., Jagasia, R., Rugarli, E. I., Imhof, A., Neupert, W., Langer, T., and Reichert, A. S. (2007) Mol Biol Cell 18, 3582–3590 Eura, Y. (2003) J Biochem 134, 333–344 Farkasovsky, M., and Kuntzel, H. (1995) J Cell Biol 131, 1003–1014 Fekkes, P., Shepard, K. A., and Yaffe, M. P. (2000) J Cell Biol 151, 333–340 Frank, S., Gaume, B., Bergmann-Leitner, E. S., Leitner, W. W., Robert, E. G., Catez, F., Smith, C. L., and Youle, R. J. (2001) Dev Cell 1, 515–525 Fritz, S. (2003) Molecular Biology of the Cell 14, 2303–2313 Fukushima, N. H., Brisch, E., Keegan, B. R., Bleazard, W., and Shaw, J. M. (2001) Mol Biol Cell 12, 2756–2766 Furt, F., and Moreau, P. (2009) Int J Biochem Cell Biol 41, 1828–1836 Gandre-Babbe, S., and van der Bliek, A. M. (2008) Molecular Biology of the Cell 19, 2402–2412 Gari, E., Piedrafita, L., Aldea, M., and Herrero, E. (1997) Yeast 13, 837–848 Griffin, E. E. (2005) The Journal of Cell Biology 170, 237–248 Griffin, E. E. (2006) Journal of Biological Chemistry 281, 16599–16606 Griparic, L., and van der Bliek, A. M. (2005) Methods Enzymol 404, 620–631 Guillery, O., Malka, F., Landes, T., Guillou, E., Blackstone, C., Lombes, A., Belenguer, P., Arnoult, D., and Rojo, M. (2008) Biol Cell 100, 315–325

44

E.Y.L. Chan et al.

Hales, K. G., and Fuller, M. T. (1997) Cell 90, 121–129 Hammermeister, M., Schodel, K., and Westermann, B. (2010) Mol Biol Cell 21, 2443–2452 Han, X. J., Lu, Y. F., Li, S. A., Kaitsuka, T., Sato, Y., Tomizawa, K., Nairn, A. C., Takei, K., Matsui, H., and Matsushita, M. (2008) The Journal of Cell Biology 182, 573–585 Harder, Z., Zunino, R., and McBride, H. (2004) Current Biology 14, 340–345 Herlan, M., Vogel, F., Bornhovd, C., Neupert, W., and Reichert, A. S. (2003) J Biol Chem 278, 27781–27788 Herlan, M., Bornhovd, C., Hell, K., Neupert, W., and Reichert, A. S. (2004) J Cell Biol 165, 167–173 Hermann, G. J., Thatcher, J. W., Mills, J. P., Hales, K. G., Fuller, M. T., Nunnari, J., and Shaw, J. M. (1998) J Cell Biol 143, 359–373 Hogeboom, G. H., Schneider, W. C., and Pallade, G. E. (1947) Proc Soc Exp Biol Med 65, 320 Hogeboom, G. H., Schneider, W. C., and Pallade, G. E. (1948) J Biol Chem 172, 619–635 Holt, I. J., Harding, A. E., and Morgan-Hughes, J. A. (1988) Nature 331, 717–719 Hoppins, S., and Nunnari, J. (2009) Biochim Biophys Acta 1793, 20–26 Hoppins, S., Lackner, L., and Nunnari, J. (2007) Annu Rev Biochem 76, 751–780 Hoppins, S., Horner, J., Song, C., McCaffery, J. M., and Nunnari, J. (2009) J Cell Biol 184, 569–581 Ichishita, R., Tanaka, K., Sugiura, Y., Sayano, T., Mihara, K., and Oka, T. (2007) J Biochem 143, 449–454 Ingerman, E. (2005) J Cell Biol 170, 1021–1027 Ishihara, N., Fujita, Y., Oka, T., and Mihara, K. (2006) EMBO J 25, 2966–2977 Jakobs, S. (2003) Journal of Cell Science 116, 2005–2014 James, D. I. (2003) J Biol Chem 278, 36373–36379 Jones, B. A., and Fangman, W. L. (1992) Genes Dev 6, 380–389 Kanazawa, T., Zappaterra, M. D., Hasegawa, A., Wright, A. P., Newman-Smith, E. D., Buttle, K. F., McDonald, K., Mannella, C. A., van der Bliek, A. M., and Lu, B. (2008) PLoS Genetics 4, e1000022 Kang-Decker, N., Cao, S., Chatterjee, S., Yao, J., Egan, L. J., Semela, D., Mukhopadhyay, D., and Shah, V. (2007) J Cell Sci 120, 492–501 Kann, O., and Kovacs, R. (2007) Am J Physiol Cell Physiol 292, C641-657 Karbowski, M. (2004) The Journal of Cell Biology 166, 1027–1039 Kennedy, E. P., and Lehninger, A. L. (1948) J Biol Chem 172, 847–848 Kennedy, E. P., and Lehninger, A. L. (1949) J Biol Chem 179, 957–972 Kijima, K., Numakura, C., Izumino, H., Umetsu, K., Nezu, A., Shiiki, T., Ogawa, M., Ishizaki, Y., Kitamura, T., Shozawa, Y., and Hayasaka, K. (2004) Human Genetics 116, 23–27 Labrousse, A. M., Zappaterra, M. D., Rube, D. A., and van der Bliek, A. M. (1999) Mol Cell 4, 815–826 Legesse-Miller, A. (2003) Molecular Biology of the Cell 14, 1953–1963 Lehninger, A. L. (1945) The Journal of Biological Chemistry 161, 437–451 Lehninger, A. L. (1949) J Biol Chem 178, 625–644 Lewis, M. R., and Lewis, W. H. (1915) Am J Anat 17, 339–401 Liu, X., Weaver, D., Shirihai, O., and Hajnoczky, G. (2009) EMBO J 28, 3074–3089 Low, H. H., and Lowe, J. (2006) Nature 444, 766–769 Low, H. H., Sachse, C., Amos, L. A., and Lowe, J. (2009) Cell 139, 1342–1352 Macfarlane, M. G., and Spencer, A. G. (1953) Biochem J 54, 569–575 McConnell, S. J., Stewart, L. C., Talin, A., and Yaffe, M. P. (1990) J Cell Biol 111, 967–976 McQuibban, G. A., Saurya, S., and Freeman, M. (2003) Nature 423, 537–541 McQuibban, G. A., Lee, J. R., Zheng, L., Juusola, M., and Freeman, M. (2006) Curr Biol 16, 982–989 McQuibban, A. G., Joza, N., Megighian, A., Scorzeto, M., Zanini, D., Reipert, S., Richter, C., Schweyen, R. J., and Nowikovsky, K. (2010) Hum Mol Genet 19, 987–1000 Meeusen, S., McCaffery, J. M., and Nunnari, J. (2004) Science 305, 1747–1752 Meeusen, S., DeVay, R., Block, J., Cassidy-Stone, A., Wayson, S., McCaffery, J. M., and Nunnari, J. (2006) Cell 127, 383–395

1  The Genetics of Mitochondrial Fusion and Fission

45

Meglei, G., and McQuibban, G. A. (2009) Biochemistry 48, 1774–1784 Merkwirth, C., Dargazanli, S., Tatsuta, T., Geimer, S., Lower, B., Wunderlich, F. T., von Kleist-Retzow, J. C., Waisman, A., Westermann, B., and Langer, T. (2008) Genes Dev 22, 476–488 Messerschmitt, M. (2003) The Journal of Cell Biology 160, 553–564 Misaka, T., Miyashita, T., and Kubo, Y. (2002) J Biol Chem 277, 15834–15842 Mnaimneh, S., Davierwala, A. P., Haynes, J., Moffat, J., Peng, W.-T., Zhang, W., Yang, X., Pootoolal, J., Chua, G., and Lopez, A. (2004) Cell 118, 31–44 Mozdy, A. D., McCaffery, J. M., and Shaw, J. M. (2000) J Cell Biol 151, 367–380 Munoz, J. M., and Leloir, L. F. (1943) The Journal of Biological Chemistry 147, 355–362 Nakamura, N., Kimura, Y., Tokuda, M., Honda, S., and Hirose, S. (2006) EMBO Rep 7, 1019–1022 Nass, M. M., and Nass, S. (1963) J Cell Biol 19, 593–611 Nass, S., and Nass, M. M. (1963) J Cell Biol 19, 613–629 Navratil, M., Terman, A., and Arriaga, E. A. (2008) Exp Cell Res 314, 164–172 Naylor, K. (2005) J Biol Chem 281, 2177–2183 Niemann, A. (2005) The Journal of Cell Biology 170, 1067–1078 Nunnari, J., Marshall, W. F., Straight, A., Murray, A., Sedat, J. W., and Walter, P. (1997) Mol Biol Cell 8, 1233–1242 Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P., and Lenaers, G. (2003) J Biol Chem 278, 7743–7746 Osman, C., Haag, M., Potting, C., Rodenfels, J., Dip, P. V., Wieland, F. T., Brugger, B., Westermann, B., and Langer, T. (2009) J Cell Biol 184, 583–596 Otsuga, D., Keegan, B. R., Brisch, E., Thatcher, J. W., Hermann, G. J., Bleazard, W., and Shaw, J. M. (1998) J Cell Biol 143, 333–349 Palade, G. E. (1952) Anat Rec 114, 427–451 Park, Y. Y., Lee, S., Karbowski, M., Neutzner, A., Youle, R. J., and Cho, H. (2010) J Cell Sci 123, 619–626 Patterson, G. H., and Lippincott-Schwartz, J. (2002) Science 297, 1873–1877 Perkins, G. A., Sun, M. G., and Frey, T. G. (2009) Methods Enzymol 456, 29–52 Pitts, K. R. (2004) J Biol Chem 279, 50286–50294 Potter, V. R., Lyle, G. G., and Schneider, W. C. (1951) J Biol Chem 190, 293–301 Praefcke, G. J., and McMahon, H. T. (2004) Nat Rev Mol Cell Biol 5, 133–147 Roman, H. (1980) Annu Rev Genet 14, 447–450 Rujiviphat, J., Meglei, G., Rubinstein, J. L., and McQuibban, G. A. (2009) J Biol Chem 284, 28682–28686 Schatz, G., Haslbrunner, E., and Tuppy, H. (1964) Biochemical and Biophysical Research Communications 15, 127–132 Schlickum, S., Moghekar, A., Simpson, J. C., Steglich, C., O’Brien, R. J., Winterpacht, A., and Endele, S. U. (2004) Genomics 83, 254–261 Schneider, W. C. (1948) The Journal of Biological Chemistry 176, 259–266 Schon, E. A., and Gilkerson, R. W. (2010) Biochim Biophys Acta 1800, 245–249 Sesaki, H. (2003) Biochemical and Biophysical Research Communications 308, 276–283 Sesaki, H., and Jensen, R. E. (1999) J Cell Biol 147, 699–706 Sesaki, H., and Jensen, R. E. (2001) J Cell Biol 152, 1123–1134 Sesaki, H., and Jensen, R. E. (2004) J Biol Chem 279, 28298–28303 Sesaki, H., Dunn, C. D., Iijima, M., Shepard, K. A., Yaffe, M. P., Machamer, C. E., and Jensen, R. E. (2006) J Cell Biol 173, 651–658 Shaw, J. M., and Nunnari, J. (2002) Trends Cell Biol 12, 178–184 Shin, H. W., Takatsu, H., Mukai, H., Munekata, E., Murakami, K., and Nakayama, K. (1999) J Biol Chem 274, 2780–2785 Sjostrand, F. S. (1953) Nature 171, 30–31 Smirnova, E., Shurland, D. L., Ryazantsev, S. N., and van der Bliek, A. M. (1998) J Cell Biol 143, 351–358 Smirnova, E., Griparic, L., Shurland, D. L., and van der Bliek, A. M. (2001) Mol Biol Cell 12, 2245–2256

46

E.Y.L. Chan et al.

Stojanovski, D. (2004) J Cell Sci 117, 1201–1210 Sugioka, R., Shimizu, S., and Tsujimoto, Y. (2004) J Biol Chem 279, 52726–52734 Suzuki, M. (2005) J Biol Chem 280, 21444–21452 Suzuki, M., Jeong, S.-Y., Karbowski, M., Youle, R. J., and Tjandra, N. (2003) Journal of Molecular Biology 334, 445–458 Taguchi, N., Ishihara, N., Jofuku, A., Oka, T., and Mihara, K. (2007) Journal of Biological Chemistry 282, 11521–11529 Takahashi, Y., Karbowski, M., Yamaguchi, H., Kazi, A., Wu, J., Sebti, S. M., Youle, R. J., and Wang, H. G. (2005) Molecular and Cellular Biology 25, 9369–9382 Tamura, Y., Harada, Y., Shiota, T., Yamano, K., Watanabe, K., Yokota, M., Yamamoto, H., Sesaki, H., and Endo, T. (2009) J Cell Biol 184, 129–141 Tatsuta, T., and Langer, T. (2008) EMBO J 27, 306–314 Tieu, Q., and Nunnari, J. (2000) J Cell Biol 151, 353–366 Tondera, D. (2004) Journal of Biological Chemistry 279, 31544–31555 Tondera, D. (2005) Journal of Cell Science 118, 3049–3059 Trifunovic, A., Wredenberg, A., Falkenberg, M., Spelbrink, J. N., Rovio, A. T., Bruder, C. E., Bohlooly, Y. M., Gidlof, S., Oldfors, A., Wibom, R., Tornell, J., Jacobs, H. T., and Larsson, N. G. (2004) Nature 429, 417–423 Twig, G., Elorza, A., Molina, A. J., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E., and Shirihai, O. S. (2008) EMBO J 27, 433–446 Wallace, D. C., Singh, G., Lott, M. T., Hodge, J. A., Schurr, T. G., Lezza, A. M., Elsas, L. J., 2nd, and Nikoskelainen, E. K. (1988) Science 242, 1427–1430 Wang, G., Moniri, N. H., Ozawa, K., Stamler, J. S., and Daaka, Y. (2006) Proc Natl Acad Sci U S A 103, 1295–1300 Wasiak, S., Zunino, R., and McBride, H. M. (2007) The Journal of Cell Biology 177, 439–450 Waterham, H. R., Koster, J., van Roermund, C. W., Mooyer, P. A., Wanders, R. J., and Leonard, J. V. (2007) N Engl J Med 356, 1736–1741 Wells, R. C., Picton, L. K., Williams, S. C. P., Tan, F. J., and Hill, R. B. (2007) J Biol Chem 282, 33769–33775 Wong, E. D., Wagner, J. A., Gorsich, S. W., McCaffery, J. M., Shaw, J. M., and Nunnari, J. (2000) J Cell Biol 151, 341–352 Wong, E. D., Wagner, J. A., Scott, S. V., Okreglak, V., Holewinske, T. J., Cassidy-Stone, A., and Nunnari, J. (2003) J Cell Biol 160, 303–311 Yaffe, M. P. (1999) Nat Cell Biol 1, E149-150 Yonashiro, R., Ishido, S., Kyo, S., Fukuda, T., Goto, E., Matsuki, Y., Ohmura-Hoshino, M., Sada, K., Hotta, H., Yamamura, H., Inatome, R., and Yanagi, S. (2006) EMBO J 25, 3618–3626 Yoon, Y., Pitts, K. R., and McNiven, M. A. (2001) Mol Biol Cell 12, 2894–2905 Yoon, Y., Krueger, E. W., Oswald, B. J., and McNiven, M. A. (2003) Molecular and Cellular Biology 23, 5409–5420 Yu, T. (2005) Journal of Cell Science 118, 4141–4151 Zhang, Y., and Chan, D. C. (2007) Proceedings of the National Academy of Sciences 104, 18526–18530 Zhao, J., Liu, T., Jin, S. B., Tomilin, N., Castro, J., Shupliakov, O., Lendahl, U., and Nister, M. (2009) Journal of Cell Science 122, 2252–2262 Zhu, P. P. (2004) J Biol Chem 279, 35967–35974 Zick, M., Duvezin-Caubet, S., Schafer, A., Vogel, F., Neupert, W., and Reichert, A. S. (2009) FEBS Lett 583, 2237–2243 Zollino, M., Lecce, R., Fischetto, R., Murdolo, M., Faravelli, F., Selicorni, A., Butte, C., Memo, L., Capovilla, G., and Neri, G. (2003) Am J Hum Genet 72, 590–597 Zunino, R., Schauss, A., Rippstein, P., Andrade-Navarro, M., and McBride, H. M. (2007) Journal of Cell Science 120, 1178–1188 Zunino, R., Braschi, E., Xu, L., and McBride, H. M. (2009) Journal of Biological Chemistry 284, 17783–17795

Chapter 2

Relationships Between Mitochondrial Dynamics and Bioenergetics Giovanni Benard, Nadège Bellance, Caroline Jose, and Rodrigue Rossignol

Abstract  In this chapter we describe the fundamental mechanisms by which mammalian cells regulate energy production, and we put emphasis on the importance of mitochondrial dynamics for the regulation of bioenergetics. We discuss both the impact of shape changes of the mitochondrion on organellar energy production, and the existence of reverse mechanisms of regulation of mitochondrial fusion and fission by the cellular energy state. Hence, in complement to pioneering concepts of metabolic control which only considered the key controlling steps of energy fluxes at the level of the respiratory chain, the recent study of mitochondrial dynamics highlights new possibilities for OXPHOS control. The implications of such a regulatory loop between mitochondrial dynamics and bioenergetics impacts several fields of human biology, as diverse as embryonic development, energy storage, cell motility, lipid and membrane biogenesis, intracellular trafficking and cell death. In addition, most neurodegenerative diseases such as Alzheimer’s disease, Parkinson’s disease, Huntington’s disease and Hereditary Spastic Paraplegia are associated with defects in mitochondrial dynamics and bioenergetics. Therefore, to unravel the fundamental mechanisms by which mitochondrial form interacts with mitochondrial function could permit to increase our basic knowledge on the regulation of energy metabolism and to decipher the pathophysiology of a group of rare neuronal diseases. Keywords  Mitochondria • Bioenergetics • Oxidative Phosphorylation • Dynamics • Neurodegenerative diseases Abbreviations ADP adenosine diphosphate ANT adenine nucleotide translocator ATP adenosine triphosphate G. Benard, N. Bellance, C. Jose, and R. Rossignol () Laboratoire “MRGM” EA4576 (Maladies Rares: Génétique et Métabolisme), Université de Bordeaux, Bordeaux, France e-mail: [email protected] B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_2, © Springer Science+Business Media B.V. 2011

47

48

CCCP COX CM CoQ Cyt c DAPI Dy DNP DRP1 EGFP EM ETC FADH2 FCCP FISH FMN 4Pi MICROSCOPE FRET GFP GTP H2O2 IBM ICS IM IMS JO2 MCA mPTP mt-NETWORK NADH OM OPA1 OXPHOS PDH PLD RCR RFP ROS SDH STED MICROSCOPY TMRM

G. Benard et al.

carbonyl cyanide m-chlorophenylhydrazone cytochrome c oxidase cristae membrane coenzyme Q cytochrome c diamidino-4¢,6-phénylindol-2 dichlorhydrate mitochondrial membrane electric potential 2,4-Dinitrophenol dynamin-related protein 1 enhanced GFP electron microscopy electron transfer chain flavin adenine dinucleotide reduced form carbonylcyanide-p-trifluoromethoxyphenylhydrazone fluorescent in situ hybridization flavin mononucleotide confocal microscope with two opposing lenses used for high resolution imaging of fluorescence fluorescence resonance energy transfer green fluorescent protein guanidin triphosphate hydrogen peroxyde inner boundary membrane intra cristae space inner membrane inter-membrane space respiratory rate metabolic control analysis mitochondrial permeability transition pore mitochondrial network nicotinamide adenine dinucleotide reduced form outer membrane gene encoding a dynamin-related mitochondrial protein causing autosomal dominant optic atrophy oxidative phosphorylation pyruvate dehydrogenase complex phospholipase D respiratory control ratio red fluorescent protein reactive oxygen species succinate dehydrogenase stimulated emission depletion microscopy tetramethyl rhodamine methyl ester

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

49

2.1 Introduction Rapid progress in mitochondrial research recently demonstrated a critical role for mitochondrial fusion and fission in the modulation of mitochondrial functions as diverse as apoptosis, energetics, calcium signaling, and ROS generation (Rossignol and Karbowski 2009). Alternative observations also reported changes in mitochondrial form upon activation of apoptosis, cell division, or adaptation to low energy states. Hence, the link between mitochondrial dynamics and energy production can be investigated in two directions. In this chapter we present first the observations in favor of a regulation of mitochondrial dynamics by the mitochondrial energy state, and secondly we summarize the experimental results indicating a possible control of mitochondrial energy production by organellar, shape changes. Prior to develop these two hypotheses we describe briefly the mechanisms of cellular and mitochondrial energy production, as well as their regulation, and we give a basic introduction on mitochondrial dynamics. At the end of this chapter we highlight the importance of the link between mitochondrial dynamics and energetics in human pathology.

2.2 The Biochemistry of Energy Production In most human tissues, mitochondria provide the energy necessary for cell growth, and biological activities. It has been estimated that about 90% of mammalian oxygen consumption is mitochondrial, which primarily serves to synthesize ATP, although in variable levels according to the tissue considered and the organism’s activity status (Benard et al. 2006). Mitochondria intervene in the ultimate phase of cellular catabolism, following the enzymatic reactions of intermediate metabolism that degrade carbohydrates, fats and proteins into smaller molecules such as pyruvate, fatty acids and amino acids, respectively (Fig.  2.1). Mitochondria further transform these energetic elements into NADH and/or FADH2, through ß-oxidation and the Krebs cycle. Those reduced equivalents are then degraded by the mito­ chondrial respiratory chain in a global energy converting process called oxidative phosphorylation (OXPHOS) where the electrons liberated by the oxidation of NADH and FADH2 are passed along a series of carriers regrouped under the name of “respiratory chain” or “electron transport chain” (ETC), and ultimately transferred to molecular oxygen (Fig. 2.2). ETC is located in mitochondrial inner membrane, with an enrichment in the cristae. ETC consists of four enzyme complexes (complexes I – IV), and two mobile electron carriers (coenzyme Q and cytochrome c). These complexes are composed of numerous subunits encoded by both nuclear genes and mitochondrial DNA at the exception of complex II (nuclear only). It was demonstrated that these complexes can assemble into supramolecular assemblies called “supercomplexes” or respirasomes.

50

G. Benard et al.

Fig. 2.1  Main pathways of cellular and mitochondrial energy metabolism. The two main metabolic pathways, i.e glycolysis and oxidative phosphorylation are linked by the enzyme complex pyruvate dehydrogenase. Briefly, glucose is transported inside the cell and oxidized to pyruvate. Under aerobic conditions, the complete oxidation of pyruvate occurs through the TCA cycle to produce NADH, H+ and/or FADH2. These reduced equivalents are oxidized further by the mitochondrial respiratory chain

In presence of energy substrate (NADH or FADH2), the transfer of electrons from complex I (and/or II) to complex IV mediates the extrusion of protons from the matrix to the inter-membrane space, thus generating an electrochemical gradient of protons (DmH+) which is finally used by the F1-Fo ATP synthase (i.e. complex V) to produce adenosine triphosphate (ATP) the energetic currency of the cell. This gradient has two components: an electric potential (DY) and a chemical potential (DmH+) that can also be expressed as a pH gradient (DpH). According to the chemios­motic theory (Mitchell 1961), DmH+ = DY−ZDpH, with Z = −2.303 RT/F. Under physiological conditions, mitochondrial energy production can alternate between two energy steady-states: basically, at state 4, respiration is slow and ATP is not produced (DY is high), while during state 3, respiration is faster and ATP is largely produced (DY is lower). In particular conditions, such as mitochondrial inner membrane permeabilization or the use of a chemical uncoupler, DY can be totally dispersed. As a consequence, respiration is accelerated and ATP production annihilated. The inhibition of respiratory chain complexes also generally decreases DY. Under physiological conditions, it is considered that mitochondria produce ATP in an intermediate state lying between state 3 and state 4. ATP is the only form

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

H+

H+

H+

51

Intermembrane space

Cyt C

Q

e-

NADH +H+

e-

1/202

NAD+

Complex l NADH dehydrogenase 47 Subunits 7 mtDNA/40 nDNA

Matrix

H2O

FAD H2

ADP Pi

FAD

Complex ll Succinate dehydrogenase 4 Subunits 0 mtDNA/4 nDNA

Complex lll Complex lV Ubiquinol cytochrome Cytochrome C oxidase C oxidoreductase 11 Subunits 1 mtDNA /10 nDNA

13 Subunits 3 mtDNA/10 nDNA

H+

ATP

Complex V ATP synthase

17 Subunits 2 mtDNA/15 nDNA

Fig.  2.2  Mitochondrial respiratory chain. For mammals, the respiratory chain consists of four enzyme complexes (complexes I – IV) and two intermediary substrates (coenzyme Q and cytochrome c). The NADH + H+ and FADH2 produced by the intermediate metabolism are oxidized further by the mitochondrial respiratory chain to establish an electrochemical gradient of protons, which is finally used by the F1F0-ATP synthase (complex V) to produce ATP, the only form of energy used by the cell. In this simple representation of the respiratory chain, the supramolecular organization (supercomplexes, dimers) is not shown

of energy used by the cell, and when produced in the mitochondrion it is exported to the cytosol by the adenine nucleotide translocator (ANT) in exchange for cytosolic ADP. Generally, the transport of energy metabolites, nucleotides and cofactors into and out of the mitochondrial matrix is performed by transporters located in the inner membrane (Palmieri et al. 1996). Mitochondria also contain “shuttle” systems that permit the transport of NADH.

2.3 The Regulation of Cellular and Mitochondrial Energy Production In mammalian cells, energy homeostasis requires a constant coordination between cell activity, nutrient availability and the regulation of energy transformation processes. This is obtained via a complex system of signaling linking energy sensing and nutrient sensing to cellular effectors that include kinases and transcription factors. The AMP-activated protein kinases are activated upon alterations in the cellular AMP/ATP ratio, which is dictated by the balance between energy supply (ATP production) and energy demand (ATP consumption). When activated by AMP,

52

G. Benard et al.

the AMPK initiate a cascade of phosphorylation to switch on the catabolic pathways that produce ATP (glycolysis, oxidative phosphorylation via the stimulation of mitochondrial biogenesis), and to switch-off the anabolic pathways that consume ATP (protein synthesis, fatty acid synthesis, cholesterol synthesis) (for review see (Hardie 2007)). More recently, it was discovered that AMPK further regulates energy metabolism through the activation of the sirtuin SIRT1 (Canto and Auwerx 2009; Canto et  al. 2009) and the downstream modulation of SIRT1 targets that include the peroxisome proliferator-activated receptor-y coactivator l alpha (PGC1a), and the forkhead box O1 (FOXO1) and 03 (FOXO3a) transcription factors. In the ­present chapter, we will discuss how mitochondrial dynamics interacts with the regulatory pathway of energy metabolism governed by AMPK and sirtuins, and reciprocally. The regulation of mitochondrial energy production is concerted and multi-site (Benard et al. 2010). The different levels of OXPHOS regulation include (1) the direct modulation of respiratory chain kinetic parameters, (2) modulation of OXPHOS intrinsic efficiency by changes in the basal proton conductance or the induced proton conductance, (3) possible changes in the morphological state of the mitochondrial compartment (as discussed here), (4) modulation of mitochondrial biogenesis and degradation, and (5) in situ regulation of mitochondrial heterogeneity by the cellular and the mitochondrial microenvironment. Most of these regulatory mechanisms (Fig. 2.3) of mitochondrial energy production were discovered at the

Fig.  2.3  Multi-site regulation of mitochondrial oxidative phosphorylation. The modulation of OXPHOS capacity and activity occurs at different levels to adapt mitochondrial energy production to the cellular needs and environmental bioenergetic constraints

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

53

level of the respiratory chain and its surrounding lipid environment. Below, we discuss whether and how changes in mitochondrial fusion and fission impact these conserved mechanisms of bioenergetic regulation.

2.4 Mitochondrial Structure and Dynamics In the past decade, the development of fluorescence microscopy has allowed the gathering of a three dimensional view of the mitochondrion in living human cells (Bereiter-Hahn and Voth 1994; Griparic and van der Bliek 2001; Yaffe 1999). In these studies, the mitochondria looked as an organelle that appeared more like a wide network of long tubules rather than a collection of small individual vesicles. In Fig. 2.4, we see the arborescence of the mitochondrial network in living human cells. Mitochondrial dynamics is performed by fusion proteins and fission proteins, which could reveal new possibilities for the control of mitochondrial energy produc­ tion and cell viability (Benard and Karbowski 2009). In mammals, the proteins involved in mitochondrial fusion include the mitofusins MFN1 and MFN2, and OPA1 (different isoforms of OPA1 are generated by alternative splicing). SLP2 (Stoml2), MarchV, Bax and Bak interact with MFN2 to regulate fusion. In addition, different ATP-dependent or ATP independent proteases regulate fusion via OPA1 processing (Ehses et al. 2009). Low mitochondrial ATP levels, or the dissipation of the mitochondrial electric membrane potential across the inner membrane induce OPA1 cleavage by PARL and the matrix AAA (m-AAA) protease (Duvezin-Caubet et al. 2006). OMA1 mediates OPA1 processing if m-AAA proteases are absent or mitochondrial activities are impaired (Ehses et al. 2009). PISD and mito-PLD are proteins involved in the metabolism of phospholipid and could regulate mitochondrial

Intracristae space

lntermembrane Inner boundary membrane (IBM) space (IMS) One cristae

junction pore

Cristae membrane (CM)

Matrix

IBM

Respiratory chain (6 fold enrichment in CM as compared to IBM)

260 nm

IMS OM Outer membrane (OM)

Fig.  2.4  Mitochondrial structure. (a) Overview of the mitochondrial network in living human HeLa cells. The mitochondrial network was imaged by fluorescence microscopy (bi-photonic), using a matrix-targeted GFP. (b) Internal organization of the mitochondrial network; section of a tubule. This scheme illustrates the “cristae junction model” of mitochondrial interior. In this view, the pores could serve to regulate the release of cytochrome c during apoptosis

54

G. Benard et al.

fusion via changes in membrane composition (Furt and Moreau 2009). The role of MIB is unclear, and might also participate to mitochondrial fusion via MFN2 binding. The proteins involved in mitochondrial fission include FIS1 and DRP1. Mitochondrial Fission Factor (MFF) also plays a role in mitochondrial fission. The regulators of fission include MARCHV, Mff, Bcl-w and different kinases required for the phosphorylation of DRP1. DRP1 can also be nitrosylated (see below). So far, the interaction between mitochondrial dynamics and mitochondrial energetics remain unclear. To investigate the link between mitochondrial energy production and organellar shape changes, appropriate methods are needed to quantify changes both in mitochondrial form and function. Firstly, one can observe the shape of the mitochondrial network by epifluorescence or confocal microscopy in living cells using various fluorescent probes targeted to the mitochondrial matrix (GFPs, mitotrackers, TMRM…). For instance, inhibited fusion leads to a fragmented mitochondrial network while inhibited fission generates long tubules with signs of hyperfusion. The morphometric analysis of the mitochondrial compartment can be performed on microscopy images by using an automated computerized method to assess the length and branching degree of the mitochondrial particles (Koopman et al. 2005b). This method gives a quantitative evaluation based on the measurement of a form factor (combined measure of length and degree of branching), an aspect ratio (measure of length), and the overall content of mitochondria in the cell. In situations of altered fusion and fission (as occurs in neurological diseases caused by mutations in OPA1 and Mfn2, or DRP1, respectively) the viscosity of the mitochondrial matrix can also change (Benard and Rossignol 2008a; Koopman et  al. 2008). Likewise, a test of mitochondrial fusion is available on cells containing different fluorescent proteins targeted to the mitochondrial matrix (matrix red fluorescent protein; mtRFP) or matrix green fluorescent protein (mtGFP). These two types of cells are fused with polyethylene glycol (PEG) to allow cell membrane fusion and the resulting polykaryons are analysed by confocal microscopy on the basis of their level of mtRFP and mtGFP colocalization. The mixing of green and red mitochondrial matrix-targeted proteins due to mitochondrial fusion leads to cells containing “yellow mitochondria” (Legros et al. 2002). Yet, the polyethylene glycol used for cell membrane fusion might also interfere with mitochondrial membrane fusion and perturbate the assay. Another possibility to measure mitochondrial fusion is to use mitochondrially targeted photoconvertible GFP (mito-Dendra) which changes color from green to red once activated by a blue laser. The mixing of the activated GFP with mitochondrial tubules generates yellow regions which can be counted as function of time to evaluate fusion, fission and transport (Koutsopoulos et  al. 2010). Similar assays were also developed to investigate automatically the activity of mitochondrial fusion and fission (Jourdain and Martinou 2010). To further assess the activity of mitochondrial fission, it is also possible to use the so-called “CCCPassay” which inhibits fusion and allows fission to proceed. The time required, or the CCCP amount needed to visualize fission gives a measure of its activity. This test is indirect and considers that OPA1 processivity is unchanged in the different conditions. This test was used in Ehses et al. (2009) and Ishihara et al. (2003, 2006).

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

55

2.5 Control of Mitochondrial Dynamics by Energy Metabolism (Hypothesis 1) The possible control of mitochondrial fusion by energy state was first suggested by studies (Legros et al. 2002; Ishihara et al. 2003) showing that mitochondrial fusion is altered by the collapse of mitochondrial membrane potential (DY). This was later partly explained by a DY-dependent cleavage of OPA1 (Duvezin-Caubet et al. 2006) and the subsequent inhibition of mitochondrial fusion. Fission can also be controlled by energy dependent processes since the phosphorylation of DRP1 by a cAMP-dependent kinase activates fission and promotes cell division. Hypothesis 1 (Fig. 2.5) was also inspired by a wide range of studies showing the alteration of cellular energy productions by the perturbation of mitochondrial fusion and fission (notably in different pathologies), as we reviewed previously in (Benard and Rossignol 2008b) and (Benard et al. 2010). Yet, the signals that mediate changes in energy state to the fusion-fission machinery remain poorly understood, and different possibilities can be proposed. A conceivable hypothesis of a bioenergetic control of mitochondrial fusion and fission by the mitochondrial and/or the cellular energy states arise from the evidence that OPA1, MFN1, MFN2 and DRP1 require GTP to perform their mecha­nical activity. This idea was initially proposed by Petr Jezek in 2009 (Jezek and PlecitaHlavata 2009; Jezek et al. 2009). The GTP formation by the Krebs cycle followed by GTP extrusion by the ANT could directly modulate the extent of mitochondrial fusion (Jezek and Plecita-Hlavata 2009; Jezek et  al. 2009). The conversion of

Exercise training Energy needs (AMPK) MARCHV

selective degradation Uncouplin

g inhibits

Low DY

Functional complementation

fusion

High DY

OPA1 long isoforms

Mfn2

Mitochondrial network and sub-networks

Metabolites diffusion Respirasome activity channeling

Fusion 3 Mitochondrial particles (fragmented network)

GTP DRP1 binding

NDP kinase

Apoptosis Cell division

ATP Fission GTP NDP kinase ATP

Oxidative Phosphorylation

Fig. 2.5  Mutual link between mitochondrial bioenergetics and dynamics (schematic representation of hypotheses 1 and 2 described in the text)

56

G. Benard et al.

ATP to GTP by nucleoside diphosphate kinases (NDP kinases) located either on the surface of the mitochondrion, or in the intermembrane space (Chen and Douglas 1987) might also play an important role in the bioenergetic control of these dynamins. The NDP kinase D (NM23-H4) was shown to be associated with the outer and inner mitochondrial membranes (Lacombe et al. 2000; Milon et al. 2000) and was not found in the soluble fractions. NDP kinase D and porin distribute similarly among the fractions, thus strongly suggesting that NDP kinase D is associated, like porin, with contact sites between the outer and the inner membrane. In addition to NDPkinase D, intra-mitochondrial NDP kinases were found in pigeon. The Nm23-H6 partially colocalizes with mitochondria (Tsuiki et al. 1999). Besides this possibility, the analysis of the different proteins involved in mitochondrial fusion or fission has evidenced post-translational modifications for DRP1, OPA1, and MFN2, which might also intervene in the modulation of mitochondrial dynamics and the subsequent control of OXPHOS capacity in response to cellular energy state and nutrients availability. First, DRP1 can be phosphorylated, ubiquitylated, nitrosylated and sumoylated. So far, the impact of these changes on mitochondrial energy production have not been investigated, and solely the effects on DRP1 recruitment to mitochondrial membrane, DRP1 turnover, fission activity or the sensitivity to apoptosis were analysed (Karbowski et al. 2007; Ishihara et  al. 2003; Figueroa-Romero et  al. 2009; Nakamura et  al. 2006). The lowered expression of DRP1 by RNA interference induces the impairment of OXPHOS with a strong reduction of ATP synthesis capacity (Benard et al. 2007). Regarding OPA1, it is well documented that a loss of mitochondrial membrane potential (i.e. uncoupling by CCCP (Duvezin-Caubet et  al. 2006)) triggers the cleavage of long isoforms of OPA1 to shorter forms, and reduces the ability for mitochondrial fusion (Legros et al. 2002). This cleavage can be performed by different m-AAA proteases which may or may not require ATP and can be controlled by prohibitins; the interaction of both the proteases and their regulators by the cellular energy states also remains to be clarified. How the energy state modifies their expression level and self-proteolytic state remains elusive. Likewise, 8 OPA1 isoforms result from alternate splicing of three exons (Ex4, Ex4b and Ex5b) and the impact of variable energy states on this pattern of OPA splicing is unknown. Alternative splicing is an important mechanism to create protein diversity and to regulate gene expression in a tissue- or developmental-specific manner. Interestingly, the activity of the spliceosome depends on the levels of ATP, since the PRP16 RNA-dependent ATPase is required for the second catalytic step of pre-mRNA splicing (Schwer and Guthrie 1992). The downregulation of OPA1 expression in human cells, or the occurence of pathogenic mutations in this protein are associated with an alteration of OXPHOS functioning (Chen et  al. 2005; Chevrollier et  al. 2008; Olichon et al. 2003). Similar results were obtained with Mfn2 downregulation in human cells or transgenic mice (Chen and Chan 2009). Again, it is still unknown how these changes in mitochondrial fusion and fission alter the mechanisms of energy production. Mfn2 can also be conjugated with MARCHV (Nakamura et  al. 2006; Karbowski et al. 2007) to stimulate its fusion activity. Recent observations also describe a role for

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

57

MFN2 in controlling ATP/ADP exchanges, possibly through protein-protein interactions, which might offer an additional mechanism by which mitochondrial fusion could control mitochondrial energy production (Guillet et al. 2009). Lastly, Mfn2 was recently identified at the interface between the endoplasmic reticulum and the mitochondrion, with a possible role in the regulation of mitochondrial Ca2+ uptake (de Brito and Scorrano 2008). Since matricial calcium can stimulate different mitochondrial dehydrogenases and transporters, which participate in energy production, Mfn2 might also be engaged in the regulation of mitochondrial energetics via calcium signalling. The hypothesis of a role for OPA1, DRP1 and MFN2 in the regulation of mitochondrial and cellular energy production is emphasized by the discovery of diseases caused by mutations in these proteins (see below). As discussed above, energy metabolism is regulated by different pathways involved in nutrient sensing, oxygen sensing or energy needs sensing. How these master regulators of energy metabolism interact with the mitochondrial fusion and fission proteins also remain to be investigated. The activation of Sirt1 by resveratrol stimulates AMPK activity and induces the upregulation of Mfn2 in neurons (Dasgupta and Milbrandt 2007). Likewise, in skeletal muscle, the expression of an active form of AMPK (transgenic mice Tg-AMPKg3225Q) triggers the upregulation of MFN-2 (mitofusin 2), OPA-1 (dynamin-like GTPase-optic atrophy 1) and DRP-1 (dynaminrelated protein 1) (Garcia-Roves et al. 2008).

2.6 Control of Mitochondrial Energetics by Organellar Dynamics (Hypothesis 2) Theoretical studies on the regulation of oxidative phosphorylation in skeletal muscle mitochondria suggest that only a direct activation by “some cytosolic factors” of all oxidative phosphorylation enzymes is able to account for the large increase in VO2 and ATP turnover accompanied by only a very moderate increase in ADP concentration during the rest to work transition (Korzeniewski 2000; Korzeniewski and Mazat 1996). Yet, recent findings in mitochondrial physiology might allow to propose a hypothetic role for mitochondrial dynamics in the rapid regulation of OXPHOS output. Indeed, a recent study by the group of JC. Martinou evidenced a rapid stress-induced mitochondrial hyperfusion (SIMH), which is accompanied by an increase in mitochondrial ATP production (Tondera et  al. 2009). This process involved the mitochondrial inner membrane protein SLP-2 and was dependent on mitochondrial oxidative phosphorylation since it was blocked in RhoO cells that exhibited a normal mitochondrial membrane potential, but were unable to synthesize ATP. The SIMH was observed in situations of stress, as induced by actinomycin D, cycloheximide or UVs, which also induced a large increase in the cellular ATP content (220%, 160% and 120% of the untreated control, respectively). The increased ATP levels associated with SIMH could be attributed to a combined stimulation of OXPHOS and glycolysis, as well as a possible inhibition of the ATP consuming activities. Still, the mechanisms by which this

58

G. Benard et al.

(hyper)fusion event could initiate an increase in mitochondrial and glycolytic ATP synthesis remain unexplained. It may be postulated that mitochondrial fusion could increase the speed of diffusion of energy metabolites (ADP, NADH) within the mitochondrial tubules, or that such conformational changes impact the respirasome organization and activate signalling cascades which remain to be deciphered. Likewise, the overexpression of mitofusin 2 improves the mitochondrial capacity to produce ATP and stimulates glucose oxidation (Bach et  al. 2005). This could be explained by a higher fusion of the mitochondrial network, but also a stimulation of mitochondrial biogenesis, as Mfn2 plays a role in both processes (Zorzano et al. 2009a, b). Furthermore, MFN2 interacts with the proto-oncogene Ras, and Ras activity was associated with the modulation of mitochondrial respiration through the regulation of complex I (Baracca et al. 2010). Hence, the interaction between Mfn2 and Ras could participate to the regulation of mitochondrial ATP production, but this point remains to be evaluated. Lastly, new insights into the critical relationships between mitochondrial dynamics and programmed cell death have demonstrated an interaction between Bax and Bak with Mfn2 (Karbowski et al. 2006) which might suggest that Bcl-2 family members may also regulate apoptosis through organelle morphogenesis machineries. Interestingly, Bcl2 can also regulate directly mitochondrial respiration, through the modulation of cytochrome c oxidase (COX) activity (Chen and Pervaiz 2009, 2007). A closer link between mitochondrial energetics and apoptosis could emerge in the coming years. In addition to Mfn2, the fusion protein OPA1 could also participate in the control of mitochondrial respiration since the removal of the m-AAA protease (which cleaves OPA1 in a Dy-dependent manner) limits the capacity of the oxidative phosphorylation system to supply ATP under conditions of high energy demand (Ehses et al. 2009). Hypothesis 2 (Fig. 2.5) also includes a control of mitochondrial dynamics by the cellular energy state via changes in mitochondrial membrane composition. Mitochondrial function relies fundamentally on organellar membranes properties, since chemiosmosis cannot exist without membrane impermeability to protons (Mitchell 1961), respiratory chain supercomplexes organization is conditioned by the lipid surrounding (Pfeiffer et al. 2003) and changes in mitochondrial membrane fluidity can alter bioenergetics (Benard et al. 2007; Benard and Rossignol 2008b, a; Aleardi et  al. 2005). Thus, modifications of the phospholipid composition of mitochondrial membranes could participate to the control of energy production or apoptosis. For instance, it was shown that changes in organellar membranes fluidity alter both the functioning of OXPHOS and the overall dynamics of the mitochondrial network, in a mutual way (Benard et al. 2007). The control of the mitochondrial apoptotic pathway is also conditioned by changes in organellar membrane integrity (MOMP) and composition (Sandra et al. 2005; Newmeyer and FergusonMiller 2003; Wei et al. 2001; Kuwana et al. 2002). It remains to be shown whether changes in mitochondrial membrane composition induced by difference in energy state could contribute to the observed changes in mitochondrial network morphology. A study revealed that an ancestral member of the phospholipase D (PLD) superfamily of lipid-modifying enzymes is required for mitochondrial fusion by hydrolysing cardiolipin to generate phosphatidic acid (Choi et  al. 2006). In this

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

59

study, the silencing of MitoPLD significantly reduced mitochondrial membrane potential and presumably associated mitochondrial energy production. To conclude on hypothesis 2, only few observations are available regarding the stimulation of OXPHOS by an increase in mitochondrial fusion, while numerous reports indicate an impairment of mitochondrial respiration in situations of altered fusion or fission (Benard and Rossignol 2008b). Therefore, more evidences are needed to prove that mitochondrial fusion and fission proteins actively participate in the regulation of mitochondrial respiration.

2.7 Mitochondrial Fusion and Functional Complementation (Consequence of Hypothesis 2) When two mitochondrial particles fuse, they can exchange materials that could permit compensation for a defective mitochondrial energy production at the cellular level. The study of Nakada et al. (2001) was the first to show the existence of mitochondrial genetic and functional complementation in vivo by analyzing the skeletal muscle of different transgenic mice containing various degrees of heteroplasmy (percentage of mutant mitochondrial DNA per cell). These authors never observed the coexistence of respiratory chain complex IV active and inactive mitochondria by histochemical staining within single muscle fibers despite the presence of 89% heteroplasmy. This might indicate that genetic complementation was attained in the muscle fibers, possibly via the inter-mitochondrial particles fusion and mixing of their content. During aging, the different proteins located in the mitochondrial network (as well as mitochondrial DNA molecules) can be progressively damaged by the high local concentration of reactive oxygen species (ROS), thus creating a heterogeneous situation where active DNAs and protein complexes could coexist with damaged ones along the tubules (heteroplasmy). In this case, the fusion of mitochondrial particles could allow functional complementation (Benard and Rossignol 2008b). Depending on the morphological state of the mt-network, the internal complementation can be variable. In the fragmented state, no exchange can occur between the different particles, creating an heterogeneous situation inside the cells. Therefore, reactions of fusion and fission could modulate the network organization and impact the capacity for compensating OXPHOS defects. Furthermore, we showed that such compensation exists until a threshold is reached (Rossignol et al. 1999, 2000, 2003) (upon strong inhibition of mitochondrial respiratory chain), and we also showed that mitochondrial fragmentation occurs once this threshold is reached (Benard et al. 2007). Accordingly, forced fusion of the mitochondrial network might allow a higher resistance (higher threshold) to a pathogenic respiratory chain impairment induced by a specific inhibitor of complex I. The mitochondrial threshold effect shows that large defects in the activity of respiratory chain complexes (induced by aging or pathogenic mutations) can occur in muscle or brain without triggering a reduction of the mitochondrial ATP production. We postulate that mitochondrial fusion and the exchange of materials such as energy metabolites and mitochondrial subunits could permit functional complementation.

60

G. Benard et al.

The role of mitochondrial fusion in promoting intramitochondrial diffusion of metabolites could be important since studies indicate the existence of segmentation along the tubules that sequester the matrix into successive domains within the same tubule. This segmentation was observed in different analyses of mitochondrial interior physical continuity, using either JC1 or photoactivable GFP, as well as FRAP experiments (Collins et al. 2002; Partikian et al. 1998; Twig et al. 2006, 2008a). The use of rosGFP1, or GFPpH also reveals zones of variable intensity in the mtnetwork of a same cell. In this case, the lipid nature of the intratubular barriers remains to be clarified, as well as their protein composition. The existence of internal barriers inside the tubule has important repercussions for the modalities of energy production, as they could abrogate the diffusion of substrates and metabolites required for oxidative phosphorylation. Different pre-fragmented domains could exist next to each other along the tubules, with differences in energy states, determined by the composition of OXPHOS complexes, the local concentration of substrates and activators of the energetic machinery, the variable volume of the different compartments (ICS and IMS), and the surrounding cytoplasm ATP needs. Thus, the idea of mitochondrial tubules operating as long uninterrupted electric cables might need to be revisited. Furthermore, it is also possible that the internal heterogeneity of mitochondrial tubule, in regard to OXPHOS activity, could be dictated by the local needs for ATP and the demand for oxygen (Mironov 2007; Mironov and Symonchuk 2006), but more studies are needed to validate this view. Mitochondrial networking could also play a role in genetic inter-mitochondrial particles complementation (Nakada et al. 2001), but this requires complete fusion and physical continuity. An alternative could be the existence of partial and transient sub-networks that will define larger functional domain. Twig et al. validated this view by using a matrix targeted photoactivable GFP (Twig et al. 2006). They showed that different subnetworks are coexisting and can undergo internal fission or fusion, without loss of their global architecture. Each network is defined by luminal continuity, equipotentiality, and boundaries which cannot be predicted without consideration of these parameters. This idea of subnetwork was previously proposed by the group of Dimitri Zorov that described the existence of Clusters Formed by Chains of Mitochondria that were called Streptio mitochondriale (Bakeeva et al. 1978). Taken together, these different observations permit the definition of four hierarchical levels of organization of the mitochondrion: the segment, the tubule, the sub-network and the global network. Each of these structures could play different roles in the synthesis and delivery of the vital ATP to various cellular areas, as well as to coordinate the different functions of the mitochondrion.

2.8 Mitochondrial Dynamics, Cell Cycle, and Organogenesis The overall networked and ramified architecture of the mitochondrion could be conserved throughout species. In some eukaryotic microorganisms such as Trypanosoma the mitochondrion is typically observed as a single highly ramified organelle.

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

61

Likewise, the protozoan Toxoplasma present only one ramified mitochondrion. Yet, the comparison of the mitochondrial network in different cell types (Kuznetsov et al. 2009) as well as the study of its internal organization in muscle, heart, liver, kidney and brain taken from rats shows a large diversity in mitochondrial shape and organization which could be partly explained by differences in energy supply and demand, as well as a tissue-specific regulation of fusion and fission. A link between the cell cycle and mitochondrial dynamics was evidenced in Hela cells, as the mitochondrial network becomes fragmented in early mitotic phase, and fused again in the daughter cells (Margineantu et al. 2002; Chang and Blackstone 2007). This process depends on Drp1 phosphorylation by Cdk1/cyclin. Mitochondrial dynamics might also play a determinant role during cell development as Campello S et al. showed that mitochondria specifically concentrate at the uropod during lymphocyte migration, by a process involving rearrangements of their shape (Campello et al. 2006). In this study, mitochondrial fission facilitated relocation of the organelles and promoted lymphocyte chemotaxis, whereas mitochondrial fusion inhibited both processes. In Drosophila, the polar granules are tightly associated with mitochondria in early embryos, suggesting that mitochondria could contribute to pole cell formation (Amikura et al. 2001a, b). It was even reported that mitochondrial large and small rRNAs are transported from mitochondria to polar granules prior to pole cell formation. Fujiwara S. and Satoh N. showed that during ooplasmic segregation, mitochondria are mainly distributed to the myoplasmic region in embryos of the ascidian HaloCynthia roretzi (Fujiwara et  al. 1993). Recently, a transgenic mice model showed that brain-specific Drp1 ablation caused developmental defects of the cerebellum (Wakabayashi et  al. 2009). Likewise, a primary culture of NS-Drp1(−/−) mouse forebrain showed a decreased number of neurites and defective synapse formation (Ishihara et al. 2009). Lastly, the removal of Mfn2 from the cerebellum in a transgenic mice model (Chen et al. 2007), triggered neurodegeneration caused by loss of mitochondrial fusion. In this model, during development and after maturity, Purkinje cells required Mfn2 (but not Mfn1) for dendritic outgrowth, spine formation, and cell survival.

2.9 Mitochondrial Dynamics, Energetics and Diseases The recent discoveries of pathogenic mutations in genes essentials for fusion and fission of the mitochondrial network have complicated the physiopathological mechanisms of these diseases (Detmer and Chan 2007), and led to questions about the similarities between enzymatic defects in respiratory chain complexes versus the perturbation of mt-network dynamics. Why do they lead to similar clinical consequences, how do they interact, and what are the molecular and signaling bases that connect each other? Recent hypotheses in the field of mitochondrial physiopathology even consider that defects of the mitochondrial network could be the final cause of the disease, consequently to mutations affecting primarily the respiratory chain complexes. For instance, the work of Duvezin et  al. (2007) analyzed the

62

G. Benard et al.

fusion protein OPA1 in different cells models with a primary genetic defect in the respiratory chain complexes. These included cybrids created from a patient with myoclonus epilepsy and ragged-red fibers (MERRF) syndrome, mouse embryonic fibroblasts harboring an error-prone mitochondrial mtDNA polymerase gamma, heart tissue derived from heart-specific TFAM knock-out mice suffering from mitochondrial cardiomyopathy, and skeletal muscles from patients suffering from mitochondrial myopathies such as myopathy encephalopathy lactic acidosis and stroke-like episodes. They observed that dissipation of the mitochondrial membrane potential led to fast induction of proteolytic processing of OPA1 and concomitant fragmentation of mitochondria. Moreover, recovery of mitochondrial fusion depended on protein synthesis and was accompanied by resynthesis of large isoforms of OPA1. This could designate a novel mechanism by which the proteolytic processing of OPA1 could induce the fragmentation of energetically compromised mitochondrial segments. This could prevent the fusion of dysfunctional mitochondrial particle with the functional mitochondrial network. As discussed above, the relationships between mitochondrial energy production and mt-network organization are mutual (Benard et al. 2007). Hence, pathological mutations affecting either the respiratory chain, or the dynamic machinery, can both lead to OXPHOS deficiency and abnormal shaping of the mitochondrion. The observations of mt-network architecture in cells from patients with a genetic defect in respiratory chain show inconsistent abnormalities of the mt-network. The patterns of these changes can vary form one patient to another, as the clinical signs typically do. Most studies looking at these aspects concerned the complex I deficiencies, in human skin fibroblasts, since it is the most common cause of mitochondrial diseases. The group of Robinson B. analyzed the mitochondrial structure and motion dynamics in living cells with energy metabolism defects by real time microscope imaging (Pham et al. 2004). They concluded that skin fibroblasts from patients with mitochondrial complex I deficiency and normal fibroblasts treated with rotenone, or antimycin A, contained higher proportions of mitochondria in the swollen filamentous forms, nodal filaments, and ovoid forms rather than the slender filamentous forms found in normal cells. They also reported a decreased motility with more ovoid mitochondrial forms compared to the filamentous forms. Likewise, it was shown that when complex I activity was chronically reduced by 80% in human skin fibroblasts, using rotenone treatment, the percentage of moving mitochondria and their velocity decreased by 30% (Koopman et al. 2007). It was proposed that ROS generated by the ETC in pathological situations could be responsible for such observed changes in mt-network organization. To test this hypothesis, Werner Koopman and Sjoerd Verkaart developed a fine and reliable method for the simultaneous quantification of oxidant levels and cell spreading (Koopman et al. 2005a, b, 2006a, b), based the monitoring of CM-H2DCF conversion into DCF upon intracellular oxidation, by video-rate confocal microscopy. After an extensive validation of their protocol, these authors looked at the ROS steady-state levels in fibroblasts taken from patients with a complex I deficiency and demonstrated a 2.5-fold higher oxidative stress in these cells. In another study, the same group demonstrated that superoxide production is increased in complex I deficient cell lines, in proportion

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

63

to the enzymatic activity decrease. Interestingly, the redox state of these cells remained unaffected, even though oxidative stress was higher (Koopman et  al. 2005b). Lastly, they analyzed the consequence of complex I deficiency on the mitochondrial network organization and observed a higher branching and elongation of the tubules, qualified as “mitochondrial outgrowth” (Koopman et al. 2005a). The questions arise about the importance of mt-network adaptations for the diagnosis of mitochondrial diseases. This could allow a first check as an easy test on blood cells, with a rapid mitochondrial staining and direct visualization on a fluorescent microscope. All OXPHOS defects will not be identified this way (Guillery et al. 2008; Benard et al. 2007), but large deficiencies could be observed at the level of the mt-network, owing that the cell type under consideration expresses sufficiently the metabolic defect. It was reported that an heterozygous, dominant-negative mutation in the dynamin-like protein 1 gene (DLP1) could cause a pathological conditions associated with microcephaly, abnormal brain development, optic atrophy and hypoplasia, persistent lactic acidemia, and a mildly elevated plasma concentration of very-long-chain fatty acids. The DLP1 protein is involved in the fission of mt-network tubules, and we showed that its absence can impair the synthesis of ATP by the mitochondrion (Benard et al. 2007). Thus, the mitochondrial network organization is a new parameter that must be taken into account for physiopathological analyses of mitochondrial diseases. The possible interaction between mitochondrial dynamics and energy production could offer an opportunity for the discovery of drugs that target mitochondrial fusion or fission with a stimulatory effect on energy metabolism, to the benefit of patients suffering from obesity, diabetes and mitochondrial/neuromuscular diseases (Zorzano et al. 2009a, b). Defective fusion or fission are the cause of rare human neurological disorders such as Charcot-Marie Tooth type 2a, Autosomal Dominant Optic Atrophy, Lethal Defect of Mitochondrial and Peroxisomal Fission, Spastic Paraplegia or a dominant form of spino-cerebellar ataxia, SCA28 (defective proteins MFN2, OPA1/3, DRP1, Paraplegin or AFG3l2 respectively), the pathophysiological molecular mechanisms of which still remain unknown. Recent evidence also suggest that mitochondrial dynamics is impaired in Parkinson’s disease, and that Parkin could be involved in the selective degradation of altered mitochondrial particles (Narendra et al. 2008, 2009). Twig and colleagues proposed a life cycle of mitochondrial particles where regions of the mitochondrial network with low membrane potential could be selectively targeted and brought to degradation (Twig et al. 2008a, b), presumably through a Parkin-mediated mechanism. In Alzheimer’s disease, elevated levels of S-nitrosylated Drp1 were found in brain samples from AD patients and AD mouse models, along with extensive mitochondrial fragmentation (Cho et al. 2009). For all of the above listed diseases, the impairment of mitochondrial energy production as observed in fibroblasts taken from patients with OPA1 or MFN2 mutations still remain unexplained at the molecular level. Acknowledgments  We thank the French National Institute for Scientific and Medical Research (INSERM), Université Victor Segalen Bordeaux 2, Région Aquitaine, Ammi, and Cancéropôle Grand Sud-Ouest for financial support. N. Bellance was supported by a Grant from INSERM/ Région Aquitaine and G. Benard by a grant from ANR.

64

G. Benard et al.

References Aleardi, A. M., Benard, G., Augereau, O., Malgat, M., Talbot, J. C., Mazat, J. P., Letellier, T., Dachary-Prigent, J., Solaini, G. C. & Rossignol, R. (2005) Gradual Alteration of Mitochondrial Structure and Function by beta-Amyloids: Importance of Membrane Viscosity Changes, Energy Deprivation, Reactive Oxygen Species Production, and Cytochrome c Release. J Bioenerg Biomembr, 37, 207–225. Amikura, R., Hanyu, K., Kashikawa, M. & Kobayashi, S. (2001a) Tudor protein is essential for the localization of mitochondrial RNAs in polar granules of Drosophila embryos. Mech Dev, 107, 97–104. Amikura, R., Kashikawa, M., Nakamura, A. & Kobayashi, S. (2001b) Presence of mitochondriatype ribosomes outside mitochondria in germ plasm of Drosophila embryos. Proc Natl Acad Sci U S A, 98, 9133–8. Bach, D., Naon, D., Pich, S., Soriano, F. X., Vega, N., Rieusset, J., Laville, M., Guillet, C., Boirie, Y., Wallberg-Henriksson, H., Manco, M., Calvani, M., Castagneto, M., Palacin, M., Mingrone, G., Zierath, J. R., Vidal, H. & Zorzano, A. (2005) Expression of Mfn2, the Charcot-MarieTooth neuropathy type 2A gene, in human skeletal muscle: effects of type 2 diabetes, obesity, weight loss, and the regulatory role of tumor necrosis factor alpha and interleukin-6. Diabetes, 54, 2685–93. Bakeeva, L. E., Chentsov YU, S. & Skulachev, V. P. (1978) Mitochondrial framework (reticulum mitochondriale) in rat diaphragm muscle. Biochim Biophys Acta, 501, 349–69. Baracca, A., Chiaradonna, F., Sgarbi, G., Solaini, G., Alberghina, L. & Lenaz, G. (2010) Mitochondrial Complex I decrease is responsible for bioenergetic dysfunction in K-ras transformed cells. Biochim Biophys Acta, 1797, 314–23. Benard, G. & Karbowski, M. (2009) Mitochondrial fusion and division: Regulation and role in cell viability. Semin Cell Dev Biol, 20, 365–74. Benard, G. & Rossignol, R. (2008a) Mitochondrial fluidity matters. Focus on “Inherited complex I deficiency is associated with faster protein diffusion in the matrix of moving mitochondria”. Am J Physiol Cell Physiol, 294, C1123. Benard, G. & Rossignol, R. (2008b) Ultrastructure of the mitochondrion and its bearing on function and bioenergetics. Antioxid Redox Signal, 10, 1313–42. Benard, G., Faustin, B., Passerieux, E., Galinier, A., Rocher, C., Bellance, N., Delage, J. P., Casteilla, L., Letellier, T. & Rossignol, R. (2006) Physiological diversity of mitochondrial oxidative phosphorylation. Am J Physiol Cell Physiol. Benard, G., Bellance, N., James, D., Parrone, P., Fernandez, H., Letellier, T. & Rossignol, R. (2007) Mitochondrial bioenergetics and structural network organization. J Cell Sci, 120, 838–48. Benard, G., Bellance, N., Jose, C., Melser, S., Nouette-Gaulain, K. & Rossignol, R. (2010) Multi-site control of cellular and mitochondrial energy production. BBA Bioenergetics Jun-Jul;1797(6–7):698–709. Bereiter-Hahn, J. & Voth, M. (1994) Dynamics of mitochondria in living cells: shape changes, dislocations, fusion, and fission of mitochondria. Microsc Res Tech, 27, 198–219. Campello, S., Lacalle, R. A., Bettella, M., Manes, S., Scorrano, L. & Viola, A. (2006) Orchestration of lymphocyte chemotaxis by mitochondrial dynamics. J Exp Med, 203, 2879–86. Canto, C. & Auwerx, J. (2009) PGC-1alpha, SIRT1 and AMPK, an energy sensing network that controls energy expenditure. Curr Opin Lipidol, 20, 98–105. Canto, C., Gerhart-Hines, Z., Feige, J. N., Lagouge, M., Noriega, L., Milne, J. C., Elliott, P. J., Puigserver, P. & Auwerx, J. (2009) AMPK regulates energy expenditure by modulating NAD(+) metabolism and SIRT1 activity. Nature. Chang, C. R. & Blackstone, C. (2007) Drp1 phosphorylation and mitochondrial regulation. EMBO Rep, 8, 1088–9; author reply 1089–90. Chen, H. & Chan, D. C. (2009) Mitochondrial dynamics – fusion, fission, movement, and mitophagy – in neurodegenerative diseases. Hum Mol Genet, 18, R169-76.

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

65

Chen, W. J. & Douglas, M. G. (1987) Phosphodiester bond cleavage outside mitochondria is required for the completion of protein import into the mitochondrial matrix. Cell, 49, 651–8. Chen, Z. X. & Pervaiz, S. (2007) Bcl-2 induces pro-oxidant state by engaging mitochondrial respiration in tumor cells. Cell Death Differ, 14, 1617–27. Chen, Z. X. & Pervaiz, S. (2009) Involvement of cytochrome c oxidase subunits Va and Vb in the regulation of cancer cell metabolism by Bcl-2. Cell Death Differ. Chen, H., Chomyn, A. & Chan, D. C. (2005) Disruption of fusion results in mitochondrial heterogeneity and dysfunction. J Biol Chem, 280, 26185–92. Chen, H., Mccaffery, J. M. & Chan, D. C. (2007) Mitochondrial fusion protects against neurodegeneration in the cerebellum. Cell, 130, 548–62. Chevrollier, A., Guillet, V., Loiseau, D., Gueguen, N., De Crescenzo, M. A., Verny, C., Ferre, M., Dollfus, H., Odent, S., Milea, D., Goizet, C., Amati-Bonneau, P., Procaccio, V., Bonneau, D. & Reynier, P. (2008) Hereditary optic neuropathies share a common mitochondrial coupling defect. Ann Neurol, 63, 794–8. Cho, D. H., Nakamura, T., Fang, J., Cieplak, P., Godzik, A., Gu, Z. & Lipton, S. A. (2009) S-nitrosylation of Drp1 mediates beta-amyloid-related mitochondrial fission and neuronal injury. Science, 324, 102–5. Choi, S. Y., Huang, P., Jenkins, G. M., Chan, D. C., Schiller, J. & Frohman, M. A. (2006) A common lipid links Mfn-mediated mitochondrial fusion and SNARE-regulated exocytosis. Nat Cell Biol. Collins, T. J., Berridge, M. J., Lipp, P. & Bootman, M. D. (2002) Mitochondria are morphologically and functionally heterogeneous within cells. Embo J, 21, 1616–27. Dasgupta, B. & Milbrandt, J. (2007) Resveratrol stimulates AMP kinase activity in neurons. Proc Natl Acad Sci U S A, 104, 7217–22. De Brito, O. M. & Scorrano, L. (2008) Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature, 456, 605–10. Detmer, S. A. & Chan, D. C. (2007) Functions and dysfunctions of mitochondrial dynamics. Nat Rev Mol Cell Biol, 8, 870–9. Duvezin-Caubet, S., Jagasia, R., Wagener, J., Hofmann, S., Trifunovic, A., Hansson, A., Chomyn, A., Bauer, M. F., Attardi, G., Larsson, N. G., Neupert, W. & Reichert, A. S. (2006) Proteolytic processing of OPA1 links mitochondrial dysfunction to alterations in mitochondrial morphology. J Biol Chem, 281, 37972–9. Duvezin-Caubet, S., Koppen, M., Wagener, J., Zick, M., Israel, L., Bernacchia, A., Jagasia, R., Rugarli, E. I., Imhof, A., Neupert, W., Langer, T. & Reichert, A. S. (2007) OPA1 processing reconstituted in yeast depends on the subunit composition of the m-AAA protease in mitochondria. Mol Biol Cell, 18, 3582–90. Ehses, S., Raschke, I., Mancuso, G., Bernacchia, A., Geimer, S., Tondera, D., Martinou, J. C., Westermann, B., Rugarli, E. I. & Langer, T. (2009) Regulation of OPA1 processing and mitochondrial fusion by m-AAA protease isoenzymes and OMA1. J Cell Biol, 187, 1023–36. Figueroa-Romero, C., Iniguez-Lluhi, J. A., Stadler, J., Chang, C. R., Arnoult, D., Keller, P. J., Hong, Y., Blackstone, C. & Feldman, E. L. (2009) SUMOylation of the mitochondrial fission protein Drp1 occurs at multiple nonconsensus sites within the B domain and is linked to its activity cycle. Faseb J, 23, 3917–27. Fujiwara, S., Kawahara, H., Makabe, K. W. & Satoh, N. (1993) A complementary DNA for an ascidian embryonic nuclear antigen Hgv2 encodes a protein closely related to the amphibian histone-binding protein N1. J Biochem, 113, 189–95. Furt, F. & Moreau, P. (2009) Importance of lipid metabolism for intracellular and mitochondrial membrane fusion/fission processes. Int J Biochem Cell Biol, 41, 1828–36. Garcia-Roves, P. M., Osler, M. E., Holmstrom, M. H. & Zierath, J. R. (2008) Gain-of-function R225Q mutation in AMP-activated protein kinase gamma3 subunit increases mitochondrial biogenesis in glycolytic skeletal muscle. J Biol Chem, 283, 35724–34. Griparic, L. & Van Der Bliek, A. M. (2001) The many shapes of mitochondrial membranes. Traffic, 2, 235–44.

66

G. Benard et al.

Guillery, O., Malka, F., Frachon, P., Milea, D., Rojo, M. & Lombes, A. (2008) Modulation of mitochondrial morphology by bioenergetics defects in primary human fibroblasts. Neuromuscul Disord, 18, 319–30. Guillet, V., Gueguen, N., Verny, C., Ferre, M., Homedan, C., Loiseau, D., Procaccio, V., AmatiBonneau, P., Bonneau, D., Reynier, P. & Chevrollier, A. (2009) Adenine nucleotide translocase is involved in a mitochondrial coupling defect in MFN2-related Charcot-Marie-Tooth type 2A disease. Neurogenetics. Hardie, D. G. (2007) AMP-activated/SNF1 protein kinases: conserved guardians of cellular energy. Nat Rev Mol Cell Biol, 8, 774–85. Ishihara, N., Jofuku, A., Eura, Y. & Mihara, K. (2003) Regulation of mitochondrial morphology by membrane potential, and DRP1-dependent division and FZO1-dependent fusion reaction in mammalian cells. Biochem Biophys Res Commun, 301, 891–8. Ishihara, N., Fujita, Y., Oka, T. & Mihara, K. (2006) Regulation of mitochondrial morphology through proteolytic cleavage of OPA1. Embo J, 25, 2966–77. Ishihara, N., Nomura, M., Jofuku, A., Kato, H., Suzuki, S. O., Masuda, K., Otera, H., Nakanishi, Y., Nonaka, I., Goto, Y., Taguchi, N., Morinaga, H., Maeda, M., Takayanagi, R., Yokota, S. & Mihara, K. (2009) Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nat Cell Biol, 11, 958–66. Jezek, P. & Plecita-Hlavata, L. (2009) Mitochondrial reticulum network dynamics in relation to oxidative stress, redox regulation, and hypoxia. Int J Biochem Cell Biol, 41, 1790–804. Jezek, P., Plecita-Hlavata, L., Smolkova, K. & Rossignol, R. (2009) Distinctions and similarities of cell bioenergetics and the role of mitochondria in hypoxia, cancer, and embryonic development. Int J Biochem Cell Biol. Jourdain, A. & Martinou, J. C. (2010) Mitochondrial dynamics: quantifying mitochondrial fusion in vitro. BMC Biol, 8, 99. Karbowski, M., Norris, K., MM., C., Jeong, S. & Youle, R. J. (2006) Role of BAX and BAK in mitochondrial morphogenesis. Nature, Online publication doi:10.1038/nature05111. Karbowski, M., Neutzner, A. & Youle, R. J. (2007) The mitochondrial E3 ubiquitin ligase MARCH5 is required for Drp1 dependent mitochondrial division. J Cell Biol, 178, 71–84. Koopman, W. J., Verkaart, S., Visch, H. J., Van Der Westhuizen, F. H., Murphy, M. P., Van Den Heuvel, L. W., Smeitink, J. A. & Willems, P. H. (2005a) Inhibition of complex I of the electron transport chain causes O2-. -mediated mitochondrial outgrowth. Am J Physiol Cell Physiol, 288, C1440-50. Koopman, W. J., Visch, H. J., Verkaart, S., Van Den Heuvel, L. W., Smeitink, J. A. & Willems, P. H. (2005b) Mitochondrial network complexity and pathological decrease in complex I activity are tightly correlated in isolated human complex I deficiency. Am J Physiol Cell Physiol, 289, C881-90. Koopman, W. J., Verkaart, S., Van Emst-De Vries, S. E., Grefte, S., Smeitink, J. A. & Willems, P. H. (2006a) Simultaneous quantification of oxidative stress and cell spreading using 5-(and-6)chloromethyl-2¢,7¢-dichlorofluorescein. Cytometry A, 69, 1184–92. Koopman, W. J., Visch, H. J., Smeitink, J. A. & Willems, P. H. (2006b) Simultaneous quantitative measurement and automated analysis of mitochondrial morphology, mass, potential, and motility in living human skin fibroblasts. Cytometry A, 69, 1–12. Koopman, W. J., Hink, M. A., Verkaart, S., Visch, H. J., Smeitink, J. A. & Willems, P. H. (2007) Partial complex I inhibition decreases mitochondrial motility and increases matrix protein diffusion as revealed by fluorescence correlation spectroscopy. Biochim Biophys Acta, 1767, 940–7. Koopman, W. J., Distelmaier, F., Hink, M. A., Verkaart, S., Wijers, M., Fransen, J., Smeitink, J. A. & Willems, P. H. (2008) Inherited complex I deficiency is associated with faster protein diffusion in the matrix of moving mitochondria. Am J Physiol Cell Physiol, 294, C1124–32. Korzeniewski, B. (2000) Regulation of ATP supply in mammalian skeletal muscle during resting state – intensive work transition. Biophys Chem, 83, 19–34. Korzeniewski, B. & Mazat, J. (1996) Theoretical studies on control of oxidative phosphorylation in muscle mitochondria at different energy demands and oxygen concentrations. Acta Biotheoretica, 44, 263–269.

2  Relationships Between Mitochondrial Dynamics and Bioenergetics

67

Koutsopoulos, O. S., Laine, D., Osellame, L., Chudakov, D. M., Parton, R. G., Frazier, A. E. & Ryan, M. T. (2010) Human Miltons associate with mitochondria and induce microtubule-dependent remodeling of mitochondrial networks. Biochim Biophys Acta, 1803, 564–74. Kuwana, T., Mackey, M. R., Perkins, G., Ellisman, M. H., Latterich, M., Schneiter, R., Green, D. R. & Newmeyer, D. D. (2002) Bid, Bax, and lipids cooperate to form supramolecular openings in the outer mitochondrial membrane. Cell, 111, 331–42. Kuznetsov, A. V., Hermann, M., Saks, V., Hengster, P. & Margreiter, R. (2009) The cell-type specificity of mitochondrial dynamics. Int J Biochem Cell Biol, 41, 1928–39. Lacombe, M. L., Milon, L., Munier, A., Mehus, J. G. & Lambeth, D. O. (2000) The human Nm23/ nucleoside diphosphate kinases. J Bioenerg Biomembr, 32, 247–58. Legros, F., Lombes, A., Frachon, P. & Rojo, M. (2002) Mitochondrial fusion in human cells is efficient, requires the inner membrane potential, and is mediated by mitofusins. Mol Biol Cell, 13, 4343–54. Margineantu, D., Cox, W., Sundell, L., Sherwood, S., Beechen, J. & Capaldi, R. (2002) Cell cycle dependent morphology changes and associated mitochondrial DNA redistribution in mitochondria of human cell lines. Mitochondrion, 1, 397–478. Milon, L., Meyer, P., Chiadmi, M., Munier, A., Johansson, M., Karlsson, A., Lascu, I., Capeau, J., Janin, J. & Lacombe, M. L. (2000) The human nm23-H4 gene product is a mitochondrial nucleoside diphosphate kinase. J Biol Chem, 275, 14264–72. Mironov, S. L. (2007) ADP regulates movements of mitochondria in neurons. Biophys J, 92, 2944–52. Mironov, S. L. & Symonchuk, N. (2006) ER vesicles and mitochondria move and communicate at synapses. J Cell Sci, 119, 4926–34. Mitchell, P. (1961) Coupling of phosphorylation to electron and hydrogen transfer by a chemiosmotic type of mechanism. Nature, 191, 144–148. Nakada, K., Inoue, K., Ono, T., Isobe, K., Ogura, A., Goto, Y. I., Nonaka, I. & Hayashi, J. I. (2001) Inter-mitochondrial complementation: Mitochondria-specific system preventing mice from expression of disease phenotypes by mutant mtDNA. Nat Med, 7, 934–40. Nakamura, N., Kimura, Y., Tokuda, M., Honda, S. & Hirose, S. (2006) MARCH-V is a novel mitofusin 2- and Drp1-binding protein able to change mitochondrial morphology. EMBO Rep, 7, 1019–22. Narendra, D., Tanaka, A., Suen, D. F. & Youle, R. J. (2008) Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. J Cell Biol, 183, 795–803. Narendra, D., Tanaka, A., Suen, D. F. & Youle, R. J. (2009) Parkin-induced mitophagy in the pathogenesis of Parkinson disease. Autophagy, 5, 706–8. Newmeyer, D. D. & Ferguson-Miller, S. (2003) Mitochondria: releasing power for life and unleashing the machineries of death. Cell, 112, 481–90. Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P. & Lenaers, G. (2003) Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J Biol Chem, 278, 7743–6. Palmieri, F., Bisaccia, F., Capobianco, L., Dolce, V., Fiermonte, G., Iacobazzi, V., Indiveri, C. & Palmieri, L. (1996) Mitochondrial metabolite transporters. Biochim Biophys Acta, 1275, 127–32. Partikian, A., Olveczky, B., Swaminathan, R., Li, Y. & Verkman, A. S. (1998) Rapid diffusion of green fluorescent protein in the mitochondrial matrix. J Cell Biol, 140, 821–9. Pfeiffer, K., Gohil, V., Stuart, R. A., Hunte, C., Brandt, U., Greenberg, M. L. & Schagger, H. (2003) Cardiolipin stabilizes respiratory chain supercomplexes. J Biol Chem, 278, 52873–80. Pham, N. A., Richardson, T., Cameron, J., Chue, B. & Robinson, B. H. (2004) Altered mitochondrial structure and motion dynamics in living cells with energy metabolism defects revealed by real time microscope imaging. Microsc Microanal, 10, 247–60. Rossignol, R. & Karbowski, M. (2009) Editorial of the directed issue on mitochondrial dynamics in biology and medicine. Int J Biochem Cell Biol, 41, 1748–9. Rossignol, R., Malgat, M., Mazat, J.-P. & Letellier, T. (1999) Threshold Effect and Tissue Specificity. J of Biological Chemistry, 274, 33426–33432.

68

G. Benard et al.

Rossignol, R., Letellier, T., Malgat, M., Rocher, C. & Mazat, J. P. (2000) Tissular variation in the control of oxidative phosphorylations, implication for mitochondrial diseases. Biochemical Journal, 347, 45–53. Rossignol, R., Faustin, B., Rocher, C., Malgat, M., Mazat, J. P. & Letellier, T. (2003) Mitochondrial threshold effects. Biochem J, 370, 751–62. Sandra, F., Degli Esposti, M., Ndebele, K., Gona, P., Knight, D., Rosenquist, M. & Khosravi-Far, R. (2005) Tumor necrosis factor-related apoptosis-inducing ligand alters mitochondrial membrane lipids. Cancer Res, 65, 8286–97. Schwer, B. & Guthrie, C. (1992) A conformational rearrangement in the spliceosome is dependent on PRP16 and ATP hydrolysis. Embo J, 11, 5033–9. Tondera, D., Grandemange, S., Jourdain, A., Karbowski, M., Mattenberger, Y., Herzig, S., Da Cruz, S., Clerc, P., Raschke, I., Merkwirth, C., Ehses, S., Krause, F., Chan, D. C., Alexander, C., Bauer, C., Youle, R., Langer, T. & Martinou, J. C. (2009) SLP-2 is required for stress-induced mitochondrial hyperfusion. Embo J, 28, 1589–600. Tsuiki, H., Nitta, M., Furuya, A., Hanai, N., Fujiwara, T., Inagaki, M., Kochi, M., Ushio, Y., Saya, H. & Nakamura, H. (1999) A novel human nucleoside diphosphate (NDP) kinase, Nm23-H6, localizes in mitochondria and affects cytokinesis. J Cell Biochem, 76, 254–69. Twig, G., Graf, S. A., Wikstrom, J. D., Mohamed, H., Haigh, S. E., Elorza, A., Deutsch, M., Zurgil, N., Reynolds, N. & Shirihai, O. S. (2006) Tagging and tracking individual networks within a complex mitochondrial web with photoactivatable GFP. Am J Physiol Cell Physiol, 291, C176-84. Twig, G., Elorza, A., Molina, A. J., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E. & Shirihai, O. S. (2008a) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. Embo J, 27, 433–46. Twig, G., Hyde, B. & Shirihai, O. S. (2008b) Mitochondrial fusion, fission and autophagy as a quality control axis: the bioenergetic view. Biochim Biophys Acta, 1777, 1092–7. Wakabayashi, J., Zhang, Z., Wakabayashi, N., Tamura, Y., Fukaya, M., Kensler, T. W., Iijima, M. & Sesaki, H. (2009) The dynamin-related GTPase Drp1 is required for embryonic and brain development in mice. J Cell Biol, 186, 805–16. Wei, M. C., Zong, W. X., Cheng, E. H., Lindsten, T., Panoutsakopoulou, V., Ross, A. J., Roth, K. A., Macgregor, G. R., Thompson, C. B. & Korsmeyer, S. J. (2001) Proapoptotic BAX and BAK: a requisite gateway to mitochondrial dysfunction and death. Science, 292, 727–30. Yaffe, M. P. (1999) Dynamic mitochondria. Nat Cell Biol, 1, E149-50. Zorzano, A., Liesa, M. & Palacin, M. (2009a) Role of mitochondrial dynamics proteins in the pathophysiology of obesity and type 2 diabetes. Int J Biochem Cell Biol, 41, 1846–54. Zorzano, A., Sebastian, D., Segales, J. & Palacin, M. (2009b) The molecular machinery of mitochondrial fusion and fission: An opportunity for drug discovery? Curr Opin Drug Discov Devel, 12, 597–606.

Chapter 3

Mitochondrial Dynamics and Autophagy Linsey Stiles, Andrew Ferree, and Orian Shirihai

Abstract  Efficient mitochondrial quality control is critical for maintenance of a healthy mitochondrial population. Both mitochondrial dynamics and selective mitochondrial autophagy, termed mitophagy, contribute to mitochondrial turnover and quality control. Mitochondrial fusion and fission allow for complementation of mitochondrial solutes, proteins, and DNA but also for generation of unequal daughter organelles. Selective fusion is utilized for incorporation of polarized mitochondria back into the network, while a depolarized mitochondrion will not fuse, but instead will be targeted for elimination by mitophagy. Mitophagy is dependent on mitochondrial dysfunction, such as depolarization, and a number of proteins are required for core autophagic machinery, signaling, and mitochondrial segregation and targeting. The relationship between mitochondrial dynamics and autophagy and how they may contribute to both mitochondrial and cellular quality control is beginning to be elucidated. Even with the questions that remain in regards to the regulation and interdependence of mitochondrial dynamics and mitophagy, it is clear that alterations in these processes lead to mitochondrial dysfunction and pathological states such as neurodegeneration. Keywords  Mitochondrial dynamics • Mitochondrial quality control • Mitophagy • Aging • Neurodegeneration

L. Stiles Department of Medicine, Section of Molecular Medicine, Boston University School of Medicine, Boston, MA, USA and Department of Pharmacology and Experimental Therapeutics, Tufts University School of Medicine, Boston, MA, USA A. Ferree Department of Pharmacology, Boston University School of Medicine, Boston, MA, USA O. Shirihai () Department of Medicine, Section of Molecular Medicine, Boston University School of Medicine, Boston, MA, USA e-mail: [email protected] B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_3, © Springer Science+Business Media B.V. 2011

69

70

L. Stiles et al.

Abbreviations A9-DA AD ALS AMPK ATG CA CCCP CMA CMT2A Cvt DA DLB DN DNM1L Drp1 ER ERK1/2 ETC Fzo1p GAP HIF LAMP-2A LC3; (MAP)LC3 MAO mdivi-1 Mfn1/2 mPTP MPTP mtDNA mTOR mtPA-GFP NGF OPA1 PBMC PD PE PI PINK1 RGC Rheb ROS TMRE

A9-subtype dopaminergic neurons of the substantia nigra pars compacta Alzheimer’s disease amyotrophic lateral sclerosis 5¢adenosine-monophosphate activated protein kinase autophagy-related genes constitutively active carbonyl cyanide m-chlorophenylhydrazone chaperone-mediated autophagy Charcot-Marie-Tooth neuropathy type 2A cytoplasmic to vacuole targeting dopaminergic dementia with Lewy Bodies dominant negative dynamin 1-like dynamin related protein 1 endoplasmic reticulum extracellular signal-regulated protein kinase 1/2 electron transport chain fuzzy onion protein GTPase-activating protein hypoxia-inducing factor lysosomal-associated membrane protein 2A microtubule-associated protein light chain 3 monoamine oxidase mitochondrial division inhibitor mitofusin 1 and 2 mitochondrial permeability transition pore 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine mitochondrial DNA mammalian target of Rapamycin mitochondrial photoactivatable green fluorescent protein nerve growth factor optic atrophy protein 1 peripheral blood mononuclear cells Parkinson’s disease phosphatidylethanolamine phosphatidylinositol PTEN-induced kinase 1 retinal ganglion cells Ras homolog enriched in brain reactive oxygen species tetramethylrhodamine ethyl ester

3  Mitochondrial Dynamics and Autophagy

TOR TORC1/2 Ulk1/2 VDAC1 Vps

71

target of Rapamycin TOR complex 1 and 2 Unc-51-like kinase 1 and 2 voltage-dependent anion channel 1 vacuolar protein-sorting.

3.1 A Brief Overview of Mitochondrial Dynamics Mitochondria function as heterogeneous networks that undergo frequent fusion and fission events, constituting mitochondrial dynamics, which regulate their morpho­ logy, number, and function (Bereiter-Hahn and Voth 1994; Chen and Chan 2005). Mitochondrial dynamics have been demonstrated to contribute to mitochondrial function in a number of systems including budding yeast, pancreatic b-cells, muscle, and neurons (Liesa et al. 2009). It has been established that mitochondrial dynamics can influence almost every aspect of the mitochondrion including biogenesis, bioenergetics, heterogeneity and elimination (Wikstrom et al. 2009; Hyde et al. 2010). Proteins that mediate mitochondrial fusion and fission have been identified. In mammals, fusion is regulated by at least three mitochondrial localized GTPases: mitofusin 1 (Mfn1), mitofusin 2 (Mfn2), and optic atrophy protein 1 (OPA1) (Santel and Fuller 2001; Misaka et  al. 2002). The yeast homologs of these proteins are Fzo1p (Mfn1/2) and Mgm1p (OPA1) (Merz et al. 2007). Mfn1 and Mfn2 are loca­ lized to the outer mitochondrial membrane, while OPA1 is an inner mitochondrial membrane protein. Mitochondrial fusion is a two step process where fusion of the inner and outer membranes occurs as separate events. Fission is mediated by the transmembrane protein Fis1 and cytosolic GTPase dynamin related protein 1 (Drp1/ DNM1L); with Fis1 as the rate limiting factor of fission in some models. The yeast homologs to the fission proteins are Dnm1p (Drp1) and Fis1p (James et al. 2003; Praefcke and McMahon 2004). Drp1 translocates from the cytosol to scission sites on the outer mitochondrial membrane to initiate fission events. It should be noted that many of these proteins have roles outside of mitochondrial fusion and fission. For instance, it has been shown that Mfn2 is located on endoplasmic reticulum (ER), where it can regulate ER morphology and ER to mitochondria tethering; a process required for efficient Ca2+ homeostasis (de Brito and Scorrano 2008). Fis1 and Drp1 are localized to peroxisomes and have been shown to mediate peroxisomal fission in a similar manner to that of mitochondrial fission (Koch et al. 2003, 2005). Mitochondrial dynamics are essential to maintain a metabolically efficient mitochondrial population and disruption of either fusion or fission alters mitochondrial morphology and functionality. Fusion contributes to maintenance of oxidative phosphorylation and mitochondrial membrane potential, with loss of fusion gene­ rally resulting in mitochondrial fragmentation, decreased mitochondrial membrane potential and oxygen consumption, and often increased reactive oxygen species (ROS) production and susceptibility to apoptosis (Chen and Chan 2010; Zorzano et al. 2010). Fusion allows for the generation of continuous membranes and matrix

72

L. Stiles et al.

lumen, which subsequently allows for complementation of solutes, metabolites, and proteins. Complementation is thought to be a key mechanism by which mitochondria can rescue a damaged unit within the network. Mitochondrial fission is also a modulator of mitochondrial function. Fission regulates inheritance of mitochondria by daughter cells within dividing cells, cellular differentiation (neuronal, cardiac, and muscle cells), and progression of apoptosis. Loss of fission results in increased mitochondrial connectivity, loss of mtDNA, bioenergetic deficiency and changes in apoptosis (Liesa et al. 2009; Karbowski 2010). Alterations to mitochondrial fusion and fission have been demonstrated in a number of conditions including neurodegeneration, obesity, and type II diabetes. Additionally, mutations in mitochondrial dynamics proteins have been identified in human pathologies. Mutations in OPA1 lead to autosomal dominant optic atrophy, which is the most common form of hereditary optic neuropathy (Alexander et al. 2000). Charcot-Marie-Tooth (CMT) disease is a group of genetically heterogeneous diseases of the peripheral nervous system, characterized by distal muscle atrophy. Mutations in Mfn2 have been identified that result in CMT Type 2A (Zuchner et al. 2004). These data place mitochondrial dynamics and function at the crossroads of human pathologies, in particular neurodegenerative diseases.

3.2 Autophagy Christian de Duve coined the term ‘autophagy’ meaning ‘self-eating’ in 1963 (Klionsky 2007). In 1967, he published work showing that mitochondria are located within lysosomes when rat liver was perfused with glucagon (Deter et  al. 1967; Deter and De 1967). Recently, the field of autophagy has rapidly expanded with research demonstrating that autophagy contributes to survival in the face of starvation (nutrient-deprivation) and to turnover of damaged organelles and proteins with long half-lives. Autophagy is a key regulator of human health and aging, as it has been implicated in a number of diseases including type II diabetes, cardiomyopathies, and neurodegeneration (Essick and Sam 2010; Fujitani et al. 2010). There are three broad classifications of autophagy: chaperone-mediated autophagy, microautophagy, and macroautophagy. All three forms of autophagy promote proteolytic degradation of cytosolic components within the lysosome. In chaperonemediated autophagy (CMA), individual proteins with a consensus binding sequence (KFERQ) are recognized and delivered to the lysosome by a complex of chaperone proteins, such as heat shock cognate protein of 70 kDa (hsc70). The CMA targeted proteins then cross the lysosomal membrane via a translocation complex that includes lysosomal-associated membrane protein 2A, LAMP-2A (Orenstein and Cuervo 2010). During microautophagy, cytosolic components are directly taken up via invagination of the lysosomal membrane. In contrast, macroautophagy is the process by which macromolecular cytosolic components are degraded via sequestration in a double-membrane structure, which fuses with the lysosome to deliver the enclosed material for degradation (Glick et al. 2010; Yang and Klionsky 2010b;

3  Mitochondrial Dynamics and Autophagy

73

Goldman et  al. 2010). Macroautophagy, herein referred to as autophagy, is the process by which mitochondria amongst other organelles (such as ER, ribosomes, and peroxisomes) are degraded for recycling and elimination.

3.2.1 Autophagic Machinery The process of autophagy (Fig.  3.1) begins with an isolation membrane derived from a lipid bilayer contributed by ER and/or trans-Golgi, endosomes, and possibly mitochondria (Hailey et al. 2010); although the exact origin of the isolation membrane, also known as the phagophore, is controversial (Eskelinen 2008). The membrane expands to engulf its cargo, which is then contained within a double-membraned structure known as the autophagosome. The autophagosome fuses with the lysosome to form an autolysosome, also called an autophagolysosome, which degrades its contents via lysosomal acidic hydrolases. The resulting breakdown products are delivered back to the cytosol by lysosomal permeases and transporters where they can be reused (Yang and Klionsky 2010a).

Phagophore initiation/ formation

Elongation

Closure

Maturation/ Fusion

Autophagosome

Bafilomycin chloroquine

3MA wortmannin

Lysosome Ulk1/2 complex

mAtg9

Atg16L complex

LC3-II

Class III PtdIns3K

Permease

Acid Hydrolase

Pepstatin A + E64D

Degradation

Fig. 3.1  Overview of the mammalian autophagic machinery. Autophagy is a multi-step process that is initiated by the formation of the phagophore. This step can be blocked by using the chemical PI3-kinase inhibitors 3-methyladenine (3MA), LY294002, or, wortmannin. Phagophore formation is followed by elongation, and closure of the phagophore to form the autophagosome. Maturation of the autophagosome occurs upon fusion with a lysosome, generating the autophagolysosome. The maturation step can be inhibited by bafilomycin, an inhibitor of vacuolar H + ATPase (V-ATPase), and the lysosomotropic agent, chloroquine. Acid hydrolases degrade the cargo within the autophagolysosome, which can be blocked by the lysosomal protease inhibitors, pepstatin A and E64D. Breakdown products can be transported out of the autophagolysosome through permeases for recycling. Adapted from (Yang and Klionsky 2010a)

74

L. Stiles et al.

3.2.2 Molecular Mechanism of Autophagy in Yeast Major advancements in understanding molecular mechanisms of autophagy came from the discovery of several autophagy-related genes (ATG) in Saccharomyces cerevisiae yeast. Many mammalian homologs exist and a subset of these Atg proteins are considered the core components required for autophagosome formation (Yang and Klionsky 2010b). In yeast, Atg1 exists in a complex, with Atg13 and Atg17, which is required for initial phagophore formation. Atg13 is phosphorylated in a Target of Rapamycin (TOR) kinase dependent manner, which prevents its binding to Atg1 and Atg17. Inactivation of TOR kinase complex 1 (TORC1) leads to dephosphorylation of Atg13, thereby promoting Atg1-Atg-13-Atg17 complex formation, generation of the phagophore, and autophagy. There are two complexes that incorporate Vps34, the only PI 3-kinase in yeast. Complex I consists of the proteins Vps34, Vps15, Atg6, and Atg14; this complex is necessary for the initiation of autophagy. Vps34 uses phosphatidylinositol (PI) as a substrate to generate phosphatidylinositol triphosphate, which is required for elongation of the phagophore as well as recruitment of other Atg proteins to the phagophore assembly site. The function of Vps34 is dependent on Vps15, a serine/ threonine kinase, that modulates membrane recruitment and stimulation of the PI 3-kinase activity of Vps34 (Stack et  al. 1995). Complex II, consisting of Vps34, Vps15, Atg6, and Vps38, is involved in vacuolar sorting. Two ubiquitin-like systems are essential for autophagy in yeast: Atg5-Atg12 conjugation and Atg8 processing. These two conjugation systems are evolutionarily conserved from yeast to mammals and the nomenclature is essentially the same, except for Atg8. A number of mammalian homologs to Atg8 have been identified, the most prominent being microtubule associated protein light chain 3A (LC3). Additional Atg8 homologs include: GATE16, GABARAP, and Atg8L, all of which are processed in the same manner as Atg8 is in yeast (Tanida et al. 2006). See the mammalian molecular mechanisms of autophagy section below for more detailed information on the Atg5-Atg12 and LC3 conjugation systems.

3.2.3 Mammalian Homologs and Molecular Mechanism of Autophagy The Unc-51-like kinases,Ulk1 and Ulk2, are homologous to yeast Atg1 and participate in complex formation with mAtg13 and FIP200 (the mammalian homolog of yeast Atg17). Unlike the yeast complex, mAtg13 interacts with Ulk1, Ulk2, and FIP200 independently of its phosphorylation state. A great deal about how the phosphorylation requirements relate to autophagic activity is beginning to be elucidated. For example, it is currently known that mAtg13, Ulk1, Ulk2, FIP200, and mTOR are all phosphorylated/dephosphorylated within the complex and that

3  Mitochondrial Dynamics and Autophagy

75

this is regulated by nutritional state and modulates autophagic activity. Interestingly, one study demonstrated a role for Ulk1 in neurotrophic signaling (Zhou et  al. 2007). Activation of TrkA receptors by nerve growth factor (NGF) leads to ubiquitination of Ulk1, subsequent association with p62, and binding to the NGF/TrkA complex. Through interactions with synaptic mediators of endocytosis, Ulk1 facilitated trafficking of the ligand-receptor complex into endosomes. This novel role of Ulk1 represents an important potential mechanism for axonal membrane homeostasis via autophagic and endosomal crosstalk (Komatsu et al. 2007). The role of the class III PI-3 kinase Vps34 and its binding partner Beclin-1 (homolog of yeast Atg6) has been studied extensively. The complex of Vps34, Beclin-1, and other regulatory proteins, such as p150 (homolog of Vps15), is required for the induction of autophagy. The role of Vps34 in complex I is conserved in yeast and mammals. UVRAG, BIF-1, Atg14L, Ambra, Rubicon, and Bcl-2 are additional regulatory proteins that complex with Vps34 and Beclin-1. UVRAG, BIF-1, Atg14L, and Ambra promote autophagy while Rubicon and Bcl-2 inhibit autophagy. The two ubiquitin-like conjugation systems that are essential for autophagy in mammals are Atg5-Atg12 conjugation and LC3 processing. In the Atg5-Atg12 system, Atg7 acts as an E1-like ubiquitin activating enzyme, which activates Atg12 by binding to its carboxyterminal glycine residue in an ATP-dependent manner. Atg12 is transferred to the E2-like carrier protein Atg10, which increases the efficiency of covalent binding of Atg12 to Atg5 (on lysine 130). The conjugated Atg5Atg12 interacts noncovalently with Atg16L, which oligomerizes to form the Atg16L complex: a large multimeric complex that associates with the extending phagophore. This association is thought to induce curvature in the expanding phagophore through recruitment of processed Atg8/LC3. The Atg16L complex dissociates from the phagophore membrane once the autophagosome is formed. Microtubule-associated protein light chain 3 (LC3) is encoded for by the mammalian homolog of yeast Atg8. LC3 is expressed as a full-length cytosolic protein, which is proteolytically cleaved to LC3-I by Atg4, a cysteine protease, upon induction of autophagy. This cleavage exposes a glycine, which is subsequently activated by Atg7, an E1-like enzyme, in an ATP-dependent manner. Activated LC3-I is transferred to Atg3, an E2-like carrier protein, before phosphatidylethanolamine (PE) is conjugated to the glycine to generate LC3-II. This lipidated form of LC3 is integrated into both internal and external faces of the phagophore membrane, where it is involved in fusion of membranes. LC3-II may also act as a ‘receptor’ on the phagophore surface that can interact with target material via adaptor molecules, such as ubiquitinated proteins and p62, to induce selective cargo uptake. The two conjugation systems (Atg5-Atg12 and LC3) are closely connected with Atg16L, which acts as an E3-like enzyme and determines the sites of LC3 recruitment and lipidation. In addition, it has been proposed that formation of the Atg16L complex may be dependent on Atg8/LC3 machinery given that mice lacking the ability to process LC3 (via knockout of Atg3) have reduced Atg5-Atg12 conjugation (Sou et al. 2008). Sou et al. have also demonstrated that autophagosomes from Atg3-deficient animals are smaller and ‘open-ended’, providing evidence that the

76

L. Stiles et al.

conjugation systems are involved in elongation and closure of the growing phagophore. This is further supported by experiments showing that autophagosomes are not closed in cells where LC3 lipidation is inhibited by overexpression of an inactive mutant of Atg4 (Fujita et al. 2008). The autophagosome is generated when the expanding ends of the phagophore fuse; although before the autophagosome is ready to fuse with the lysosome, it must first go through additional maturation steps by undergoing fusion events with early and late endosomes. It is thought that this connects endocytic and autophagic pathways to deliver both cargo and membrane fusion machinery to the autophagosome and to lower the pH of the vesicle. The lower pH of the autophagic vesicle makes it more suitable to house the lysosomal proteases that will breakdown the engulfed cargo. The source of lipid used for autophagosome formation and the process by which this lipid is moved to the site of assembly remains largely unknown. mAtg9 is a transmembrane protein that is required for mammalian autophagy and is hypothesized to act as a ‘membrane carrier’. Located in the trans-Golgi network and late endosomes, mAtg9 overlaps with LC3-positive autophagosomes upon starvation or rapamycin induction of autophagy. During starvation, the cycling of mAtg9 has been shown to require Ulk1 and hVps34 (Reggiori et al. 2004; Young et al. 2006). It has been proposed that mAtg9 may contribute to the formation of the autophagosome by delivering membrane components. This hypothesis is based on its function in yeast, yet this still remains to be demonstrated in mammalian cells. Nishida et al. have shown that there is the possibility of an alternative macroautophagy pathway (Nishida et al. 2009). They found that mouse cells lacking Atg5 or Atg7 can still form autophagosomes and autolysosomes and perform autophagymediated protein and mitochondrial degradation. Interestingly, lipidation of LC3 did not occur in the Atg5/Atg7-independent process of ‘macroautophagy’. The alternative autophagy pathway still utilized and was regulated by known autophagic proteins, such as Ulk1 and Beclin1. This implies that there may be more than one pathway in which mammalian macroautophagy can occur, including an Atg5/Atg7-dependent conventional pathway and an Atg5/Atg7-independent alternative pathway.

3.2.4 Signal Transduction Regulation of Autophagy Autophagy occurs both at a basal level to maintain cellular homeostasis and at a stimulated level when induced by cellular stressors, such as nutrient starvation, oxidative stress, and hypoxia. A number of signaling pathways upstream of the ‘Atg’ machinery, or the mammalian equivalents (Yang and Klionsky 2010a), modulate autophagy in response to intracellular and extracellular stresses as shown in Fig. 3.2. In yeast there are two functionally distinct protein complexes for the protein target of rapamycin (TOR): TOR complex 1 and 2 (TORC1 and TORC2). TORC1 has the primary role in regulating autophagy in response to nutrient availability.

3  Mitochondrial Dynamics and Autophagy

77

Fig. 3.2  Signal Regulation of Autophagy in Mammalian Cells. Autophagy is highly regulated by signaling networks, with mTOR as a common downstream target of many of the signal transduction regulators of autophagy. When mTOR is activated in conditions such as nutrient availability or Akt signaling, autophagy is inhibited. Activation of AMPK upon energy depletion causes inhibition of mTOR and stimulation of autophagy. ERK1/2 signaling and PI3K III/Beclin 1 complex formation induce autophagy. The anti-apoptotic protein Bcl-2 prevents the association of Beclin 1 and PI3K III

The mammalian homolog of TOR (mTOR) is a highly conserved serine/threonine protein kinase which acts as a key regulator of autophagy by sensing nutrient availability, growth factors, and cellular energy. mTOR activation inhibits autophagy under conditions where nutrients are readily available and is a downstream signal of PI 3-kinase, insulin signaling, growth factor receptor signaling, and the Akt pathways (Young et al. 2009). For instance, Akt signaling leads to phosphorylation and inhibition of the heterodimer Tsc1/Tsc2, which is a GTPase-activating protein (GAP) for the GTPase Ras homolog enriched in brain (Rheb). Rheb is required for mTOR activity, with the GDP-bound form inhibiting mTOR, while the GTP-bound form of Rheb stimulates mTOR. Akt inhibition of the Tsc1/Tsc2 complex activates mTOR signaling and thus has an inhibitory effect on autophagy. When mTOR is inhibited by stress conditions, such as hypoxia and starvation, autophagy is induced. Upstream signals of mTOR in this scenario include 5¢adenosine-monophosphate activated protein kinase (AMPK), which is activated by depleted ATP levels. Activation of AMPK promotes the activity of the Tsc1/Tsc2 complex, which favors the GDP-bound form of Rheb, leading to inhibition of mTOR and induction of autophagy. In response to hypoxia, autophagy is induced in part by mTOR inhibition and in part by hypoxia-inducing factor (HIF), which acts independently of mTOR.

78

L. Stiles et al.

HIF induces autophagy by targeting BNIP3 and BNIP3L, which are members of the Bcl-2 family of cell death regulators (Mazure and Pouyssegur 2009; Glick et al. 2010). In both the neonatal and adult brain, hypoxia induces a rapid and extensive autophagic induction (Adhami et  al. 2006). It appears this response may not be entirely beneficial as mice lacking Atg7 are almost completely protected from ischemia-induced neurodegeneration. While much remains to be elucidated, this striking result suggests that modulation or temporary inhibition of neuronal autophagy may be a potential treatment strategy for ischemic stroke victims (Yue et al. 2009). One important effector of autophagy with connections to neurodegeneration is extracellular signal-regulated protein kinase 1/2 (ERK1/2). Brains from Parkinson’s disease (PD) patients exhibit phosphorylated ERK2 granules that colocalize with autophagocytosed mitochondria (Zhu et al. 2003). In a neuroblastoma model, expression of either constitutively active (CA) or wildtype ERK2 was sufficient to promote activation of autophagy with CA-ERK2 inducing mitophagy to a greater extent than wildtype (Dagda et al. 2008). Another protein family that plays a prominent role in regulating both autophagy and mitophagy is the Bcl-2 proteins. Inhibition of autophagy occurs via the antiapoptotic members such as Bcl-2, Bcl-XL, and Bcl-w, while proapoptotic BH3-only proteins, such as BNIP3, Bad, Bix, and BimEL can induce autophagy. These proteins seem to play a more prominent role in regulating mitophagy.

3.3 Mitophagy Mitochondria are essential for a number of critical cellular processes. They are also the main source and target of reactive oxygen species (ROS); therefore, it is essential that the pool of mitochondria within a cell remain functional and energetically efficient. Mitochondrial autophagy, or mitophagy, refers to the selective removal of mitochondria by the autophagic machinery and is another method utilized for quality control by promoting turnover (autophagy of dysfunctional mitochondria in conjunction with biogenesis). Mitophagy serves to degrade mitochondria, effectively removing them from the dynamic network. Several questions remain regarding exactly how defective mitochondria are specifically targeted for degradation via autophagy. This section will overview mitophagy and what is known about whether and how mitochondria are selectively targeted for autophagy. It has been known since 1957 that mammalian mitochondria are degraded by autophagy with Clark’s discovery that mitochondria are located in autophagosomes in the kidney of newborn mice (Clark 1957). Since then mitophagy has been documented in a number of different tissues including brain, heart, liver, reticulocytes, and pancreatic b-cells. Mitophagy has been shown to be involved in cellular quality control, differentiation, and disease pathogenesis. Initially, it was thought that autophagy was a nonselective process, arbitrarily engulfing cytosolic components, but this notion is changing with evidence that specific, defective cargo is targeted for degradation. Certain proteins that are involved with targeting mitochondria for sequestration into autophagosomes have been discovered in yeast. While this

3  Mitochondrial Dynamics and Autophagy

79

process remains largely unclear in mammalian cells, potential tags are beginning to be identified, including mitochondrial depolarization and opening of the mitochondrial permeability transition pore (mPTP).

3.3.1 Mitophagy in Yeast Takeshige et  al. were the first to show in yeast that an induction of autophagy occurs when yeast are subjected to a stressful environment, such as starvation. This was also the first work to demonstrate that autophagy in yeast was similar to mammalian autophagy. They proposed a mechanism by which autophagy could remove cytosolic components, such as mitochondria, although they believed this to be an unselective process (Takeshige et al. 1992; Goldman et al. 2010). Evidence for selective mitochondrial autophagy was first provided by Campbell and Thorsness using an experimental design that was able to distinguish mitochondria in an acidic compartment. This technique was used to demonstrate increased mitochondrial degradation under conditions such as loss of mitochondrial membrane potential and mtDNA (Campbell and Thorsness 1998). Their work provided a connection between mitochondrial damage and increased degradation, although did not provide direct evidence for mitochondrial quality control via autophagy. Early experimental evidence for this came from Kissova et al. when they demonstrated that knockout of the gene UTH1, which encodes for a yeast outer mitochondrial membrane protein, resulted in a loss of mitophagy, while other kinds of macroautophagy remained intact (Kissova et al. 2004). In addition, autophagy deficient yeast display defects in mitochondrial biogenesis, decreased oxygen consumption and mitochondrial membrane potential, and increased ROS (Zhang et al. 2007). This work highlights the role of mitophagy as a quality control mechanism that is required for maintenance of mitochondrial integrity. Further evidence for this role in quality control is provided by studies showing that mitochondrial dysfunction resulting from loss of specific proteins or pharmacologic treatments also increase mitophagy of damaged mitochondria. Removal of functionally damaged mitochondria helps maintain the mitochondrial network by keeping the system energetically efficient and removing both the major source and target of cellular ROS and, therefore, protects mtDNA. Recently, a large amount of work has gone into identifying ‘mitophagy receptors’ in both yeast and mammalian systems. Since mitophagy is a subset of macroautophagy, it utilizes the basic core autophagic machinery common with autophagy described above. On the other hand, mitophagy does utilize some additional ATG genes that are required for other pathways. Atg11, Atg 20, and Atg24, which are proteins involved in the cytoplasmic to vacuole targeting (Cvt) pathway have been shown to be required for mitochondrial autophagy (Nice et  al. 2002; Kanki and Klionsky 2010). This may demonstrate that these proteins are a general requirement for selective autophagy. ATG32 and ATG33 were identified as mitophagy-related genes in yeast by using genomic screens for mutants defective in selective mitochondrial autophagy

80

L. Stiles et al.

(Kanki et  al. 2009a; Kanki and Klionsky 2010). Atg32 is an outer mitochondrial membrane protein that has emerged as a mitophagy receptor by interacting with Atg proteins that are essential for autophagosome formation. Atg32 binds to Atg11 during mitophagy and uses it as an adaptor protein to recruit mitochondria to the phagophore assembly site (Okamoto et al. 2009b; Kanki et al. 2009b). In addition, Atg32 has a binding motif (WXXI/L/V) for binding to Atg8/LC3 family members; this binding motif is also present in Atg19 and the mammalian protein p62 (Okamoto et  al. 2009a). Okamoto et  al. showed through studies using mutations in the WXXI/L/V binding motif of Atg32 that the binding of Atg32 to Atg8 is required for sequestration of mitochondria by the phagophore. Atg33 is also localized to the outer mitochondrial membrane and may detect or present aged mitochondria for selective mitochondrial autophagy when cells have reached the stationary phase (Kanki and Klionsky 2010). While Atg32 and Uth1 have been identified as the main proteins involved with regulating mitophagy in yeast; other proteins that play a role have been established. Nowikovsky et  al. demonstrate that knockout of DMN1, the yeast homolog of Drp1, was found to have an inhibitory effect on mitophagy in a strain (mdm38 conditional knockout) known to induce mitophagy. This study also demonstrated that osmotic swelling of mitochondria induces selective mitochondrial autophagy in yeast (Nowikovsky et  al. 2007). Another protein that was demonstrated to be required for mitophagy in yeast is Aup1p. This mitochondrial intermembrane space protein is necessary for efficient targeted degradation of mitochondria via autophagy during the stationary phase of yeast (Tal et al. 2007). Interestingly, mitochondrial membrane potential depolarization induced by the oxidative phosphorylation uncoupler, carbonyl cyanide m-chlorophenylhydrazone (CCCP), does not stimulate mitochondrial autophagy in yeast (Kissova et  al. 2004; Kanki et  al. 2009a), yet mitochondrial depolarization is a necessary, although not sufficient, requirement for mitophagy in mammals.

3.3.2 Mitophagy in Mammalian Cells Mitophagy has been shown to play a key physiological role in a number of different mammalian tissues. While no mammalian homologs of ATG32 and UTH1 genes have been identified, selective removal of mitochondria does occur in mammalian cells and is mediated by mitochondrial membrane potential as well as a number of required proteins. It was first demonstrated in hepatocytes that disturbances in mitochondrial membrane potential by mPTP stimulates degradation of mitochondria via autophagy (Lemasters et al. 1998; Elmore et al. 2001). This work contributed to the knowledge that mitophagy functions to maintain cellular homeostasis by removing damaged mitochondria from healthy cells. It also acts as a mechanism to remove increased levels of ROS generated by dysfunctional mitochondria and keep cells energetically efficient (Goldman et al. 2010). Certain cell-type specific proteins that are required for mitochondrial targeting to the autophagosome have been identified. For instance, the protein Ulk1 has been

3  Mitochondrial Dynamics and Autophagy

81

shown to be necessary for autophagy during erythropoiesis. Reticulocyte maturation is an example of a specialized system that demonstrates the role of autophagic degradation of mitochondria in cellular maintenance and physiology. During the final step of erythrocyte differentiation, mitophagy is utilized to remove reticulocyte mitochondria and other organelles to create a mature red blood cell. It has been shown that Unc51-like kinase (Ulk1) is not necessary for general autophagic activity outside of the reticulocyte, but is required to selectively remove ribosomes and mitochondria during red blood cell differentiation (Kundu et al. 2008). This is in contrast with what was already reported about Ulk1 in previous sections. The discrepancy may be due to the fact that the work showing Ulk1 is a nonessential mechanistic component of mammalian autophagy was done in a Ulk1 conditional knockout where Ulk2 compensation is a possible confounding factor. Interestingly, in the Ulk1 knockout mice, not only are mitochondria not removed by autophagy during reticulocyte maturation but they also maintain their membrane potential (Zhang et al. 2009). Nix is another protein that may be required for red blood cell mitophagy, which is upstream of Ulk1. It is located on the outer mitochondrial membrane and is a member of the Bcl-2 family of proteins, specifically the BH3only proapoptotic subfamily. Nix deficient blood contains a large number of erythrocytes that retain their mitochondria, although in this system ribosomes are cleared, providing evidence that it is a mitophagy specific protein (Zhang and Ney 2008). Nix is required for mobilization of autophagic machinery and mitochondrial depolarization following CCCP treatment (Ding et  al. 2010). The mechanism behind how Nix is involved in mitophagy is still controversial, but it may play a role in mitochondrial membrane potential destabilization, which is a trigger for mitophagy. Although, some studies have reported that Nix has a membrane potential independent role in mitophagy. These findings suggest there are multiple pathways that lead to mitophagy in mammals. Nix, also known as Bnip3L, is not the only member of the Bcl-2 family of proteins that has been shown to play a role in mitophagy. Bnip3 is also a member of this family that is hypothesized to trigger autophagy by causing mitochondrial depolarization. Bnip3 has been shown to be activated following ischemia-reperfusion in cardiac myocytes, where it induces autophagy as a protective response to apoptotic signaling (Gottlieb and Carreira 2010). It is important to note that both Nix and Bnip3 play a dual role in autophagy and apoptosis. Autophagy in this system can be both protective or associated with autophagic cell death depending on the conditions (Zhang and Ney 2009). A number of mechanisms have been proposed for how Bnip3 induces autophagy including mitochondria permeability transition, mitochondrial depolarization, and/or inhibition of mitochondrial dynamics machinery.

3.3.3 Mitophagy in Neurons Under normal conditions a very small amount of autophagosomes can be detected in neurons by electron or light microscopy. However inhibition of autophagic

82

L. Stiles et al.

clearance by pharmacologic or genetic tools results in a dramatic accumulation of autophagosomes, many of which contain mitochondria. These findings indicate that not only does mitophagy occur in neurons but also the normal flux from autophagosome formation to degradation is very rapid and efficient. In fact, proper autophagic processing is absolutely crucial for neuronal survival. The dependence on autophagy for clearance of damaged macromolecules is less drastic in dividing cells as cellular debris can be diluted in the daughter cells. This protective mechanism is not an option for long-lived post-mitotic cells. Thus neurons must have functional autophagic clearance mechanisms to avoid the accumulation of hazardous waste material such as damaged mitochondria and protein aggregates. It is not surprising therefore that strong evidence exists linking defects in autophagy with neurodegeneration. To date the clearest evidence demonstrating a causal role of autophagic impairment leading to neuronal loss comes from knockout animal models. Mice with neuronal-specific Atg5 or Atg7 deficiency display massive accumulation of polyubiquitinated protein aggregates and severe neurodegeneration (Hara et  al. 2006; Komatsu et al. 2006). It is noteworthy that the neuronal loss and protein aggregates occur without additional stresses used to model neurodegeneration, such as exposure to neurotoxins or protein overexpression (Cuervo 2006). The phenotypes of these mice emphasize the essential role of basal autophagy in maintaining quality control for neuronal maintenance. The importance of autophagy in the human brain is underscored by autophagic abnormalities seen in several neurodegenerative diseases, including Alzheimer’s disease (AD) (Nixon et al. 2005; Nixon 2006) frontotemporal dementia (Lee and Gao 2008a; Ju and Weihl 2010) Parkinson’s disease (Anglade et al. 1997; Dehay et  al. 2010), and Dementia with Lewy Bodies (Zhu et  al. 2003). The autopsied brains from patients with these chronic diseases exhibit increased numbers of autophagic vesicles, protein aggregates, and dysfunctional mitochondria. In isolation the finding of increased autophagosome number has different viable potential explanations. First, the observed vesicles could result from an increase in activation and thus increased formation of autophagosomes. Alternatively the increased autophagic vesicles could represent a loss of clearance capacity through the lysosome leading to an accumulation of autophagosomes. A third possibility is a combination of increased activation in the presence of impaired degradation. At the time of autopsy the latter possibility seems most probable, as patients tend to be at late stages of a chronic disease progression spanning years to decades. It is likely to be the case in many age-related neurodegenerative diseases that impaired lysosomal degradation capacity leads to the accumulation of intracellular debris, such as damaged mitochondria and protein aggregates. This accumulation in turn signals the activation of autophagy and further increase in autophagosome number. While it is far from clear what comes first in this feed forward scenario, future therapeutics targeting enhancement of autophagic function should consider all stages of autophagosome maturation and seek to improve complete clearance of autophagosomal cargo.

3  Mitochondrial Dynamics and Autophagy

83

3.4 Mitochondrial Dynamics and Mitophagy 3.4.1 The PINK1/Parkin Pathway in Mitophagy and Mitochondrial Dynamics Recently, a large effort has been dedicated to understanding how the PINK1/parkin pathway contributes to the maintenance of cellular and mitochondrial homeostasis. There is evidence showing that this pathway is involved with the regulation of selective mitophagy of damaged mitochondria, as well as mitochondrial function and dynamics. Moreover, improper regulation of the PINK1/parkin pathway, mitophagy, and mitochondrial fusion and fission have been implicated in disease pathology, specifically in Parkinson’s disease. PTEN-induced kinase 1 (PINK1) is a Ser/Thr protein kinase with an N-terminal mitochondrial targeting signal, a putative transmembrane domain, and a C-terminal regulatory domain that governs kinase activity and substrate selectivity (Mills et al. 2008; Chu 2010). More than 50 mutations have been mapped throughout the kinase and C-terminal domain of PINK1, which have differing effects on the activity and stability of the protein (Dagda and Chu 2009). Parkin is a cytosolic E3 ubiquitin ligase. Mutations in parkin have been identified with disease causing mutations often disrupting parkin ligase activity (Lee et al. 2010). Loss-of-function mutations in PINK1 and parkin are the main cause of early-onset autosomal recessive forms of PD (Gasser 2009), which supports a role for these proteins in neuroprotection and cellular quality control. Interestingly, mice that have PINK1 and parkin knocked out display a mild phenotype and do not have the dopaminergic (DA) neuron loss in the substantia nigra observed in humans (Whitworth and Pallanck 2009). These single gene knockout mice display similar functional defects in the dopaminergic system, yet the morphology and numbers of DA neurons are normal (Goldberg et al. 2003; Kitada et al. 2007). PINK1 deficient mice display mitochondrial dysfunction prior to the onset of any neurodegeneration (Gispert et al. 2009). A triple knockout of PINK1, parkin, and DJ-1 also display normal morphology and cell number in the substantia nigra and is insufficient to cause significant nigral degeneration in the mice (Kitada et al. 2009). Again, as yet unknown, compensatory mechanisms may be held responsible. Both PINK1 and parkin play a role in mitophagy, although the mechanism is still not fully understood. It is known that upon selective recruitment of cytosolic parkin to depolarized mitochondria, the dysfunctional units are eliminated via the autophagic machinery (Narendra et  al. 2010). PINK1 expression and function is required for this process (Vives-Bauza et al. 2010; Geisler et al. 2010; Vives-Bauza and Przedborski 2010). Overexpression of wild-type PINK1 is sufficient to induce recruitment of parkin to mitochondria, even if mitochondrial membrane potential remains intact. In addition, mutations in either PINK1 or parkin inhibit parkin recruitment to mitochondria and subsequent mitophagy (Vives-Bauza and Przedborski 2010). PINK1 or parkin mutations alone or PINK1/parkin double

84

L. Stiles et al.

mutants display a similar phenotype, indicating that they operate in the same pathway. Interestingly, overexpression of parkin can compensate for loss of PINK1, while the reverse does not occur, suggesting that parkin acts downstream of PINK1 (Clark et al. 2006; Whitworth and Pallanck 2009). Loss of PINK1 leads to alterations in mitochondrial function and morphology in a number of mammalian systems. There seems to be a universal decrease in mitochondrial membrane potential and ATP synthesis in the systems where this has been studied (Exner et al. 2007; Liu et al. 2009a; Chu 2010). The same phenotype is observed with expression of mutated forms of PINK1 (Grunewald et al. 2009; Marongiu et al. 2009). Loss of PINK1 also seems to increase ROS in mammalian cells. An increase in superoxide has been demonstrated in mouse cortical neurons, human dopaminergic neurons (Wood-Kaczmar et al. 2008; Gandhi et al. 2009), and human SH-SY5Y neuroblastoma cells (Dagda et al. 2009), as well as a decrease in glutathione levels in some of these cell lines. There is also evidence for induction of mitophagy or mitochondrial association with lysosomes in a number of these studies. Interestingly, there are also changes in mitochondrial morphology with loss of PINK1, in mammalian systems this generally correlates with an increase in fragmentation (Sandebring et al. 2009). Unraveling the connection between autophagy, the PINK1/parkin pathway, and mitochondrial morphology is critical to further understanding the role of mitochondrial quality control. Although, the mitochondrial morphology phenotype of PINK1/parkin deficiency remains controversial between studies in Drosophila and higher eukaryotes, one thing is clear: loss of PINK1/parkin does lead to alterations in mitochondrial morphology. The PINK1/parkin pathway interacts with components of the mitochondrial dynamics machinery. Much of the work dissecting the connection between PINK1/ parkin and mitochondrial fusion and fission machinery has been done in Drosophila. The expression of mitofusin is inversely correlated with the activity of PINK1 and parkin in Drosophila. On the other hand, loss of PINK1 or parkin does not change the steady-state expression of Drp1 or Opa1, or the subcellular distribution of Drp1 (Poole et al. 2010). Ziviani et al. demonstrated that Mfn is ubiquitinated by parkin when it is recruited to dysfunctional mitochondria (Ziviani et al. 2010). They also show that loss of PINK1 or parkin leads to an increase in the expression of Mfn and an elongated mitochondrial morphology. Additionally, knockdown of Mfn or OPA1 or overexpression of Drp1, rescues the mitochondrial abnormalities in PINK1 or parkin mutants (Poole et  al. 2008; Park et  al. 2009). There is also evidence that the PINK1/parkin pathway negatively regulates Mfn and OPA1 function and positively regulates Drp1 (Deng et al. 2008). Yang et al. propose that Fis1 acts between PINK1 and Drp1 promoting mitochondrial fission (Yang et  al. 2008). All of this experimental evidence suggests that the PINK1/parkin pathway regulates mitochondrial morphology by either promoting mitochondrial fission or inhibiting fusion in Drosophila. This fits with the hypothesized model that the higher the frequency of mitochondrial fission, the higher the probability that dysfunctional units will be segregated and eliminated, as will be discussed in the following sections. These data suggest a scheme where PINK1/parkin promote mitochondrial fission and elimination via mitophagy, therefore providing a connection between this pathway and both mitochondrial dynamics and autophagy.

3  Mitochondrial Dynamics and Autophagy

85

Ubiquitination of Mfn by parkin is hypothesized to promote autophagy of damaged mitochondria. It is thought that the mechanism by which this occurs is that ubiquitination of the outer mitochondrial membrane acts as a recruitment signal for p62/SQSTM1, which is required for PINK1/parkin-mediated mitophagy (Geisler et  al. 2010). Although, this could also be mediated by parkin-mediated poly-ubiquitination of the voltage-dependent anion channel 1 (VDAC1), which has been shown to be necessary for PINK1/parkin-directed mitophagy. Ziviani and Whitworth provide additional potential mechanisms for the role of Mfn ubiquitination in mitophagy that focus on the function of Mfn in mitochondrial fusion. For instance, poly-ubiquitinated Mfn may be degraded consequently inhibiting fusion, with the possibility that this mechanism is specific to damaged mitochondria. If Mfn is mono-ubiquitinated, it is unlikely that it will be degraded, but it could still provide a useful regulatory mechanism. An example of this is that ubiquitination may prevent dimerization and therefore tethering of Mfn and in this manner prevents mitochondrial fusion (Ziviani and Whitworth 2010). Interestingly, contradictory mitochondrial morphology findings have been reported in human SH-SY5Y neuroblastoma cells and other mammalian systems. Dagda et al. demonstrate that stable PINK1 knockdown in SH-SY5Y cells results in a fragmented mitochondrial morphology, increased oxidative stress, and an increase in mitophagy that correlates with a 50% reduction in mitochondrial mass, which was assessed by western blotting for mitochondrial proteins (Dagda et  al. 2009). Conversely, overexpression of PINK1 lead to an increase in mitochondrial connectivity and an elongated morphology with some abnormally enlarged mitochondria, yet there was no autophagic response to these irregular mitochondria; in fact PINK1 overexpression inhibited 6-hydroxydopamine (6-ODHA)-induced mitophagy. Interestingly, dominant negative Drp1 expression rescued the morphology and autophagy effect in the PINK1 knockdown, leading to more elongated mitochondria and decreased mitophagy (Dagda et al. 2009). This study also shows that autophagy machinery is necessary for mitochondrial fragmentation, which was demonstrated by utilizing siRNA targeting Atg7 and Atg8/LC3B. Knockdown of these proteins inhibited autophagy and reversed the mitochondrial fragmentation observed in PINK1 knockdown cells to levels similar to that in control cells. Decreased PINK1 expression in SH-SY5Y cells causes reduced autophagic flux that corresponds with decreased ATP synthesis (Gegg et  al. 2010),which can be rescued by overexpression of parkin. Inducing mitochondrial dysfunction using the electron transport chain uncoupler, CCCP, demonstrated that PINK1 and parkin are required for mitophagy. Gegg et  al. show that ubiquitination of Mfn1 and Mfn2 occurs within 3 h of CCCP treatment and that this process may identify mitochondria for degradation via mitophagy. The ubiquitination of these mitochondrial dynamics proteins is inhibited with knockdown of PINK1 or parkin expression, implicating that PINK1 and parkin are essential for mitophagy, specifically for removal of damaged mitochondria. These data point to interdependency between mitochondrial dynamics and autophagy, demonstrating that mitochondrial fusion/ fission machinery is required for PINK1 mediated mitochondrial degradation and suggests that autophagy participates in mitochondrial remodeling.

86

L. Stiles et al.

Loss of PINK1 or parkin in SH-SY5Y cells causes a fragmented mitochondrial morphology (Lutz et al. 2009). Overexpression of Mfn2 and OPA1 or a dominant negative mutation of Drp1 (Drp1-DN) were able to rescue the effects loss of PINK1 or parkin had on mitochondrial morphology and function. Conversely, expression of parkin or PINK1 was able to suppress mitochondrial fragmentation induced by Drp1 expression. Furthermore, Drp1-DN expressing cells that were also knockdown for PINK1 or parkin did not display the expected fragmented mitochondrial morphology phenotype or a decrease in ATP. This implies that mitochondrial alterations in parkin- or PINK1-deficient cells are associated with an increase in mitochondrial fission and relies on mitochondrial dynamics proteins and machinery. In agreement with this research, expression of human mutant PINK1 in a rat dopaminergic cell line leads to fragmented mitochondria via an increased ratio of fission to fusion proteins thereby promoting mitochondrial fission. Mitochondrial division inhibitor (mdivi-1), a small molecule inhibitor of Drp1, rescued both the morphological and functional mitochondrial defects caused by the expression of mutant PINK1 (Cui et al. 2010). Lutz et  al. suggest that possible explanations for the difference observed in Drosophila and mammalian cells could be the time points that analysis of PINK1/ parkin deficiency occurred and/or that there could be fundamental differences in the regulation of mitochondrial fusion and fission and mitophagy in mammalian systems (Lutz et al. 2009). Mitochondrial dynamics and mitophagy could also be regulated in a tissue-specific manner. Regardless of the controversial conclusions obtained when examining PINK1/parkin, it is clear that this pathway provides an interesting mechanism that may connect mitochondrial dynamics with mitophagy. The mechanism behind how loss of mitochondrial membrane potential triggers PINK1 signaling, PINK1 triggers parkin recruitment, and parkin is localized to mitochondria remain unknown at this time. Yet, what is known is that PINK1 and parkin affect mitochondrial morphology. It has also been demonstrated that mitochondrial depolarization is critical to the PINK1/parkin pathway and is necessary for mitophagy. The connection between mitochondrial fusion and fission, membrane potential, and mitophagy has been studied extensively in mammalian cells in the context of the lifecycle of a mitochondrion. Currently, interdependence between mitochondrial dynamics, function, and depolarization has been shown to influence selective degradation of damaged mitochondria through autophagy.

3.4.2 The Lifecycle of a Mitochondrion As previously discussed, mitochondria go through continuous cycles of fusion and fission, which is thought to play a role in mitochondrial complementation, allowing for mixing of matrix solutes and metabolites, mtDNA, and mitochondrial membrane components (these occur at different rates with membrane components having the slowest rate). This sharing of proteins and solutes between mitochondria through fusion occurs as a first line of defense in an attempt to recover function in damaged

3  Mitochondrial Dynamics and Autophagy

87

mitochondria. This complementary rescue ensures that dysfunctional mitochondria do not become an energetic strain on the rest of the mitochondrial population. Mitochondrial turnover, which is a balance between mitochondrial degradation and biogenesis, is also required for a healthy mitochondrial network. This allows dysfunctional mitochondrial to be removed by mitophagy while maintaining the population and allowing propagation of the most apt mitochondria. The interplay between mitochondrial dynamics and mitophagy is a key determinant of mitochondrial homeostasis, which can be exemplified by the mitochondrial life cycle. The life cycle of a mitochondrion has been studied by Twig et  al. in INS-1 cells, a rat insulinoma cell line, where mitochondrial fission, selective fusion, and autophagy are thought to contribute to mitochondrial quality control, as shown schematically in Fig. 3.3 (Twig et al. 2008a; Mouli et al. 2009). It involves cycles of fusion periods, each followed by fission. Following the fission event, mitochondria enter a solitary state and are available for subsequent fusion events. This solitary state is ~20-fold longer than the fused period and if a mitochondrion maintains a polarized membrane potential during this period, it can re-enter the mitochondrial web by means of a subsequent fusion event with another mitochondrion.

Mitophagy 5 opa1 mfn1 PINK1 Parkin

Fission 2

Fusion

pre-autophagic pool

3

4

Sustained depolarization 1

Transient depolarization opa1

1 1

2

3

4

mfn1

fis1 drp1

PINK1

Parkin

5 Polarized mitochondria

Depolarized mitochondria

Fig.  3.3  The lifecycle of a mitochondrion; the role of fission, fusion, and autophagy in the segregation of dysfunction mitochondria. The mitochondrion shifts between a networked postfusion state (1) and a solitary post-fission state (2). Following a fission event, the daughter mitochondrion may either maintain membrane potential (red mitochondrion), or depolarize (green mitochondrion). If depolarization occurs, the mitochondrion is unlikely to undergo further fusion events for the entire depolarization interval (3). If mitochondrial depolarization is transient and membrane potential is restored, fusion capacity is also restored. However, if mitochondrial membrane potential depolarization is sustained (4), reduction in OPA1 levels and increases in PINK1 activity and parkin translocation follows and elimination by autophagy occurs (5) (Figure courtesy of Twig and Shirihai 2010)

88

L. Stiles et al.

On the other hand, if the daughter mitochondrion loses membrane potential and becomes depolarized, it will remain segregated and it can either be targeted for autophagy or it may repolarize and be recovered by the network. This suggests that mitochondrial fusion is selective for functional mitochondrial units. It is reasonable to consider that fission may merely decrease mitochondrial size making it more accessible for degradation. Although upon further reflection the creation of two dissimilar daughter units argues against this availability theory, given that size will be similar but it is function that becomes divergent. In addition, it has been shown that mitochondrial fusion depends on mitochondrial membrane potential and cannot occur in depolarized mitochondria regardless of size or the close proximity of other polarized, fusion-ready mitochondria (Twig and Shirihai 2010). These metabolically dysfunctional, depolarized mitochondria cannot fuse with other mitochondria and are preferentially targeted for degradation by the autophagosome, although the exact mechanism behind this remains unclear. However, it suggests that fusion itself may be partially responsible by unevenly redistributing components between mitochondria, which with the successive fission event results in two functionally divergent mitochondria. This ability to reorganize mitochondrial components may allow the mitochondrial population to separate and remove damaged units at a faster rate making them less energetically wasteful. The role of depolarization and how it prevents further fusion is in question when examining the life cycle of a mitochondrion. While all of the mitochondria that are targeted for autophagy are depolarized, the timeline of depolarization to engulfment provides further information regarding how autophagy is induced in these cells and whether depolarization is a cause or effect of autophagy. Twig et al. found that mitochondrial depolarization occurs at least 1 h before engulfment, indicating that removal by autophagy is not the explanation for why depolarized mitochondria cannot re-fuse with the network (Twig et al. 2008a). This time gap between depolarization and mitophagy demonstrates that mitochondrial fission is a principal route by which depolarized mitochondria are generated and fusion is selective for polarized units. It is important to note that while mitochondrial depolarization is a necessary prerequisite for mitophagy, depolarization alone is not sufficient to cause mitochondrial autophagy. With the knowledge that a mitochondrion spends a substantial amount of time as a non-fusing, segregated mitochondrion before it is degraded, the question is raised as to what the mechanistic explanation is for the observation that a depolarized mitochondrion cannot go through a subsequent fusion event. It was shown that the underlying mechanism for the membrane potential dependency of fusion is actually an impaired fusion capacity within these mitochondria. Indeed, co-staining mitochondria with mitochondrial matrix targeted photo-activatable green fluorescent protein (mtPA-GFP), to identify non-fusing mitochondria, and TMRE, a mitochondrial membrane potential dependent dye, or an anti-OPA1 antibody provided some mechanistic insight into the membrane potential and fusion state of these mitochondria. Twig et al. show that non-fusing mitochondria are depolarized and display an increase in proteolytic cleavage, or degradation, of the fusion protein, OPA1, with approximately 50% less OPA1 than in wild-type fusing mitochondria (Twig et al. 2008a).

3  Mitochondrial Dynamics and Autophagy

89

Cleavage of OPA1 has been shown to be triggered by reduction in mitochondrial ATP and by mitochondrial depolarization (Griparic et  al. 2004; Duvezin-Caubet et  al. 2006; Song et al. 2007). This implicates a targeted fusion deficiency as the explanation for a lack of mitochondrial fusion in depolarized mitochondria. The interdependence of mitochondrial dynamics and mitophagy are also revealed by utilizing tools to either knockdown or overexpress mitochondrial dynamics proteins prior to studying autophagy (Table 3.1). Retinal ganglion cells (RGCs) from mice heterozygous for a nonsense mutation in OPA1 display an increase in autophagosomes (White et  al. 2009). The authors suggest that this induction of autophagy is due to accumulation of abnormal mitochondria in the RGC layer. Induction of fusion by overexpression of OPA1 using adenovirus in INS1 cells resulted in a decrease in mitophagy by 64%. This was not due to increased mitochondrial size, but to an increase in fusion capacity. Mitochondrial fission is also essential for mitophagy. Stimulation of fission decreases mitochondrial mass in HeLa cells (Frieden et al. 2004; Gomes and Scorrano 2008) and INS1 cells (Park et al. 2008), consistent with the concept of an increase in mitophagy. Overexpression of Fis1 causes cells to accumulate fragmented mitochondria and autophagosomes, with the observation that there is selective autophagy of damaged mitochondria. Using Fis1 mutants, it was demonstrated that induction of punctate mitochondria and autophagic vesicles correlated with mitochondrial dysfunction not fragmentation, indicating that it is function not morphology that determines whether a cell will trigger mitophagy (Gomes and Scorrano 2008). Promoting fission by overexpression of Drp1 stimulates mitochondrial elimination under proapoptotic stimuli (Arnoult et al. 2005). Furthermore, inhibiting fission using either Fis1 RNAi or Drp1-DN results in a reduction in mitochondrial-specific autophagy (Barsoum et al. 2006; Twig et al. 2008a; Gottlieb and Carreira 2010). This reduction in autophagy is mitophagy specific because other forms of autophagy, such as ER autophagy (reticulophagy) remain intact, indicated by no alterations in the levels of ER mass (Twig et  al. 2008a). Moreover, pharmacological inhibition of autophagy affects mitochondrial dynamics and metabolic function. Treatment of rat myoblasts with 3MA to inhibit autophagy leads to a decrease in mitochondrial membrane potential, inhibition of mitochondrial fusion (attributed to a decrease in OPA1), and accumulation of ‘giant’ mitochondria (Navratil et  al. 2008). These works imply that both fusion and fission play a critical role in mitochondrial turnover and quality control by influencing mitophagy.

3.4.3 Lessons from Simulation Experiments Further support for the potential role of mitochondrial dynamics in mitochondrial quality control comes from simulation experiments from Mouli et  al. They measured parameters of fusion and fission that may influence maintenance of mitochondrial function. By using a programmed simulation based on experimental findings they were able to investigate the effects of fusion, fission and autophagy

Fragmentation and perinuclear clustering Fragmentation

Retinal ganglion cells INS1 b -cells

INS1 b -cells

MEFs

Opa1 heterozygous mutated mice Mfn1 overexpression

Mfn1 loss-of function (overexpression of Mfn1 DN) Mfn2 knockout

DJ-1 knockdown

PINK1 overexpression

Increased interconnectivity and elongation Abnormally enlarged mitochondria Fragmentation

Fragmentation

INS1 b -cells

Opa1 overexpression

Human neuronal SH-SY5Y cells Human neuronal SH-SY5Y cells M17 human dopaminergic cell line

Reduction in mitochondrial size with intact fusion ability Increased percentage of opaque mitochondria Mitochondrial aggregation

HeLa cells HeLa cells

PINK1 knockdown

Fragmented, swollen mitochondria Tubular, elongated mitochondria Fragmentation

INS1 b -cells

Drp1K38A (dominant negative Drp1) Drp1 RNAi Drp1 overexpression

Increased general autophagy

Increased autophagic sequestration of depolarized mitochondria Decreased in mitochondrial volume by 50% Autophagic vacuole number remained low

Inhibition of autophagy

Decrease in mitochondrial volume by 40% Unchanged mitochondrial mass

Reduction in autophagosomes containing mitochondria by 75% Decrease in mitophagy (qualitative) Decrease in mitochondrial mass by 70% Reduction in autophagosomes containing mitochondria by 63% An increase in general autophagy

Table 3.1  The effect of manipulations in mitochondrial protein expression on mitochondrial morphology and mitophagy Manipulation Cell type Effect on morphology Effect on mitophagy Fis1 RNAi Elongation Reduction in autophagosomes containing INS1 b-cells mitochondria by 70% Fis1 overexpression HeLa cells INS1 Fragmentation Enhanced autophagosome b -cells formation by 50% Reduce total mitochondrial volume by 50%

Thomas et al. (2010)

Dagda et al. (2009)

Dagda et al. (2009)

Hailey et al. (2010)

Park et al. (2008)

Park et al. (2008)

White et al. (2009)

Twig et al. (2008a)

Parone et al. (2008) Arnoult et al. (2005)

Frieden et al. (2004), Gomes and Scorrano (2008), Park et al. (2008) Twig et al. (2008a)

Reference Twig et al. (2008a)

90 L. Stiles et al.

3  Mitochondrial Dynamics and Autophagy

91

over an extended time period in a large population of mitochondria (Mouli et al. 2009). A model was developed where there is an optimal frequency of fusion and fission events that maintains mitochondrial function, such as respiration, even with a persistent damaging algorithm that should compromise mitochondrial function by inactivating mitochondrial components. In this model, where the extent of fusion and fission contribute to mitochondrial activity, it was determined that autophagy alone was the process by which damaged, “low-activity” mitochondria were removed, working to preserve function, even when this process is nonselective. Indeed, even in the absence of repair mechanisms, mitochondrial dynamics and autophagy may be adequate to restore mitochondrial function, given that fusion allows for redistribution of damaged components that can be removed quickly and preferentially by mitophagy. When mitochondrial activity under selective fusion is considered, especially with high levels of damage, there is a considerable affect on mitochondrial function. Selectivity allows for removal of damaged units by promoting increased fusion frequency of functional mitochondrial. This is accomplished without compromising autophagy of damaged mitochondria. This gives selective fusion a two-fold purpose in maintaining mitochondrial function, first as a means of intramitochondrial complementation and second as a way to segregate damaged mitochondria prior to autophagy (Nakada et al. 2001; Mouli et al. 2009). It also suggests that along with the key element of selective fusion, it is the rate of fission (not just the balance between fusion and fission) that determines the efficacy of quality control mechanisms (fusion, fission, and autophagy). A higher frequency of fission results in an increased probability of dysfunctional mitochondria being isolated and eliminated by mitophagy. This work further explores the interconnected nature of fusion, ­fission, and autophagy and how the occurrence and frequency of these processes maintains mitochondrial function.

3.4.4 Mitochondrial Motility and Dynamics Mitochondrial movement regulates mitochondrial morphology by controlling the spatial distribution of mitochondria. Mitochondrial fusion requires and in some cases can promote (i.e. transient fusion) mitochondrial motility. In H9c2 cells, there is a similar prevalence of moving and stationary mitochondria within the general mitochondrial population, yet fusion events involve moving mitochondria approximately 90% of the time (Twig et al. 2010). This work demonstrates that mitochondrial motility facilitates fusion. The extent of mitochondrial fusion depends on the location of mitochondria along microtubules, whether they are located on the same or separate microtubules, and the direction in which they are moving (Liu et al. 2009a). Mitochondria move along microtubules via kinesin motors with the assistance of the Miro/ Milton complex. Miro1 and Miro2 are atypical RhoGTPases located on the outer mitochondrial membrane that allow for the kinesin-1 motor to interact with

92

L. Stiles et al.

mitochondria. Milton is an adaptor protein, which connects Miro and kinesin to facilitate mitochondrial motility and trafficking. Miro has two Ca2+–binding EF-hands, which controls calcium-sensitivity (Chen and Chan 2009). Mitochondria are immobilized by high calcium concentrations, which is thought to assist in calcium buffering by mitochondria and localized ATP production, specifically in neurons, which have high energy demands for neurotransmitter release (Liu and Hajnoczky 2009; Stephenson 2010). The connection between mitochondrial motility and mitophagy has not yet been determined, although it has been demonstrated that PINK1 interacts with the Miro/ Milton complex (Weihofen et  al. 2009). The interdependence of mitochondrial motility and morphology is beginning to be elucidated. Mfn1 and Mfn2 have been shown to interact with the Miro/Milton complex (Misko et al. 2010). In addition, Mfn2 is necessary for axonal transport of mitochondria in dorsal root ganglia neuronal cultures, which is independent of its role in mitochondrial fusion. Loss of OPA1 has no effect on motility indicating that inner membrane fusion is not required for mitochondrial axonal transport along the microtubule. Overexpression of Miro and Milton causes mitochondrial elongation, clustering and aggregation and inhibits the mitochondrial fragmentation caused by loss of PINK1 (Glater et al. 2006; Saotome et al. 2008; Weihofen et al. 2009). Loss of Miro function leads to suppression of mitochondrial motility and Drp1 mediated mitochondrial fragmentation (Fransson et al. 2006; Saotome et al. 2008; Liu and Hajnoczky 2009). These data provide evidence that mitochondrial motility and dynamics occur in a common pathway and suggest that motility may also influence mitophagy. Figure 3.4 overviews

Fig. 3.4  The role of PINK1 in mitochondrial motility, morphology, and mitophagy. PINK1 has been shown to interact with the mitochondrial motility complex Miro/Milton, yet the exact role PINK1 plays in this complex has yet to be determined. However, it is hypothesized that PINK1 may pay a role in mitochondrial trafficking. A connection between mitochondrial motility and morphology exists, where fusion requires and can promote motility. It has been well established that changes in PINK1 expression affects mitochondrial morphology; in mammalian cells PINK1 promotes fusion and an elongated mitochondrial morphology. PINK1 also promotes mitophagy upon parkin recruitment to the mitochondria. A relationship between mitochondrial dynamics and mitophagy exists where fission followed by selective fusion allows for segregation of dysfunctional mitochondria that can be targeted for mitophagy. Mitochondrial motility, morphology, and autophagy are thought to exert mitochondrial quality control, which determines mitochondrial function. Thus, as PINK1 has been shown to regulate all these processes, it is a solid candidate as a master regulator of mitochondrial quality control

3  Mitochondrial Dynamics and Autophagy

93

the role of PINK1 in mitochondrial motility, morphology, and autophagy, presenting the possibility that PINK1 may play a role in mitochondrial quality control (Whitworth and Pallanck 2009).

3.4.5 Mitochondrial Motility, Dynamics, and Mitophagy as a Quality Control Axis Mitochondrial motility along microtubules is required for mitochondrial fusion (Liu et al. 2009b). The observation that mitochondrial fusion is a membrane potentialdependent process guarantees that dysfunctional organelles will avoid fusion, while intact mitochondria will benefit from complementation. Segregation of dysfunctional mitochondria from the fusing population results in the generation of small, depolarized mitochondria that can be targeted for degradation by mitophagy. This sets up a possible hierarchy of mitochondrial quality control mechanisms with mitochondrial movement at the top, followed by fusion, which utilizes a complementary “rescue” mechanism, followed by fission which provides dissimilar daughter cells and ending with mitophagy as the recipient of the depolarized units. Importantly, the idea of this quality control axis advocates that it is not just the balance between fusion and fission that can contribute to maintenance of the mitochondrial population, but also the absolute rate of events. Specifically, it is thought that the higher the fission frequency, the higher the probability that dysfunctional units will be segregated and eliminated (Twig et al. 2008b).

3.5 When Quality Control Breaks Down: Implications in Aging and Neurodegeneration Mitochondrial quality control is critical for maintenance of a healthy mitochondrial population. Both mitochondrial dynamics and selective mitochondrial autophagy are thought to play key roles in maintaining mitochondrial function and promoting turnover. Therefore, it is not surprising that these processes have been implicated in a number of diseases.

3.5.1 Role of Mitochondrial Dynamics and Mitophagy in Aging Mitochondria have been proposed to play a prominent role in the aging process, namely dysfunctional mitochondrial quality control leads to an accumulation of damaged mitochondria, which could lead to further cellular impairment. ‘Giant’ or swollen mitochondria are often observed in aged cells. These mitochondria are

94

L. Stiles et al.

enlarged with accumulated mtDNA and oxidized proteins, which are indicative of a lack of mitochondrial turnover (Yen and Klionsky 2008). This implicates mitochondrial fusion and fission, as well as mitophagy, as mechanisms of aging. Indeed, mitochondrial dynamics, selective mitochondrial autophagy, ROS, and mtDNA maintenance are all hypothesized to play a role in the aging process (Weber and Reichert 2010). There are a number of other aging theories that have been proposed and even additional mitochondrial-related pathways that are thought to be involved with this process, such as caloric restriction, the insulin signaling pathway/insulin sensitivity, and sirtuins, but these do not strictly fit into the topic of mitochondrial dynamics and autophagy and are beyond the scope of this chapter. Oxidative stress is an interesting starting point when examining the role mitochondria play in the aging process. Mitochondria are both the main source and, due to their close proximity and less efficient DNA proofreading enzymes, target of ROS. The ‘free radical theory of aging’ proposed by Harman in the 1950s is based on the fact that ROS induce mitochondrial dysfunction and mitochondrial dysfunction generates more ROS, leading to a vicious cycle of mitochondrial damage (Harman 1956). It is important to note that not all ROS is deleterious, it also acts as a physiological cellular signal. For instance, at low levels ROS can be a signal to stimulate ROS scavenging pathways, mitochondrial biogenesis, and mPTP, which may lead to mitophagy. As ROS levels and oxidative damage increase, instead of promoting survival of mitochondria they may instead trigger release of factors that initiate apoptosis. Increased ROS levels may also elevate mitochondrial DNA mutations. Mitochondria have increased incidences for DNA mutation compared to genomic DNA due to a number of reasons including being in close proximity to higher levels of ROS and decreased repair mechanisms. Increased accumulation of mtDNA mutations has also been proposed as an aging mechanism and is a cause for heterogeneity in the mitochondrial population. Accumulation of mutated mtDNA is an indication that mitochondrial quality control mechanisms are dysfunctional. It has been hypothesized that certain mtDNA mutations that decrease mitochondrial respiration and therefore ROS production (a seemingly beneficial result in circumstances of mitochondrial damage and high ROS production) may also decrease mitophagy (Lemasters 2005). This provides another mechanism by which impaired mitochondrial dynamics and mitophagy is implicated in the aging process. Mitophagy is known to be upregulated in response to dysfunctional mitochondria and has been proposed to slow the aging process. Specifically, it has been shown that there is an age-related decline in lysosomal systems. Accumulation of damaged mitochondria due to impaired removal/degradation of these organelles is thought to result in decreased cell viability. Therefore, functional selective autophagy of damaged mitochondria is crucial for the preservation of a healthy mitochondrial population and maintenance of cellular integrity. All of the aging pathways described thus far have a common feature in that they converge together to lead to mitochondrial damage and impaired quality control. Mitochondrial dynamics may also play a role in the aging process. Yeast deficient in Dmn1 (homolog of Drp1) have an increased lifespan. It has been demonstrated in P. anserine that mitochondrial fragmentation occurs progressively with age, which

3  Mitochondrial Dynamics and Autophagy

95

was reversed by Dnm1 knockout (Scheckhuber et al. 2007), although this does not occur in mammalian systems where Drp1 knockout is embryonically lethal. Although, this does not rule out the possibility that a small shift in the balance of mitochondrial dynamics could be beneficial to the cellular environment. Additionally, changes in mitochondrial morphology are observed in disease states that are concurrent with aging. It is often difficult to determine whether changes to mitochondrial dynamics are the cause or an effect of pathology, likely it is probably not that simple with a more cyclical mechanism expected. We do know, however, that effective mitochondrial dynamics, including complementation and selective fusion, influence mitophagy and are key determinants of quality control, which helps to maintain a healthy mitochondrial and therefore cellular population.

3.5.2 Mitochondrial-Lysosomal Axis Theory of Aging While essential, the task of degrading mitochondria via autophagy is no easy task. With lipid-rich membranes and several iron containing enzyme complexes, mitochondria represent a challenging substrate for the lysosome. Moreover damaged mitochondria are a serious hazard through their capacity to produce large quantities of ROS. This delicate and dangerous dynamic is especially relevant for neurons and has been elegantly described in the mitochondrial-lysosomal axis theory of aging proposed by Terman and Brunk (Terman et al. 2010). In their theory, high levels of intralysosomal iron combined with hydrogen peroxide leads to peroxidation of cargo in autophagosomes and lysosomes (Kurz et al. 2008b). This oxidative modification renders the cargo resistant to degradation by lysosomal hydrolytic enzymes. A non-degradable polymer of cross-linked protein and lipid residues, known as lipofuscin or ceroid, accumulates in the lysosomes and decreases auto­ phagic clea­rance. Impaired mitophagic clearance, in turn exacerbates the situation by allowing more time for reactive oxidative species to escape from damaged mitochondria. Under extreme oxidative stress, damage to the membrane of the lysosome leads to subsequent permeabilization and the pro-apoptotic leakage of lysosomal enzymes into the cytoplasm (Terman et al. 2006; Kurz et al. 2008a). Mechanisms of aging and neurodegeneration in humans are inherently difficult to prove yet there is a rapidly growing body of evidence in support of the mitochondrial-lysosomal axis theory. Examples specific to neurodegeneration come from studies of autopsied brains of patients with Alzheimer’s disease and Parkinson’s disease. Several researchers have documented evidence of aberrant mitophagy in AD (Moreira et  al. 2007b; Chen and Chan 2009; Santos et  al. 2010) including increased quantities of mitochondrial lipoic acid associated with lipofuscin (Moreira et  al. 2007a, b). Nixon and collaborators have documented numerous lysosomal-related abnormalities in sporadic AD suggestive of impaired clearance, including a striking accumulation of autophagosomes and autophagolysosomes within degenerating neurites (Nixon et al. 2008). Further mechanistic support for their interpretations come from studies of presenilin-1, the most common cause of

96

L. Stiles et al.

early-onset familial Alzheimer’s Disease (Lee et al. 2010). Tissues from patients and transgenic animals with presenilin-1 mutations display several lines of evidence indicating impaired autophagic clearance. In relation to PD, Dehay et  al. report lysosomal depletion following administration of the neurotoxin MPTP (1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine) which targets mitochondrial complex I (Vila et al. 2010; Dehay et al. 2010). The loss of lysosomes was preceded by a permeabilization of lysosomal membranes due to oxidative damage resulting from increased mitophagy. Supporting their data in toxin models, Dehay et al., also found evidence of lysosomal breakdown and autophagosome accumulation in brain tissue from PD patients. Enhancement of autophagic-lysosomal function through genetic and pharmacologic modifications was protective against the toxicity of MPTP and, most importantly, attenuated the PD-related dopaminergic neurodegeneration. These studies illustrate the lysosomal hazards of mitophagy and point to the great potential for treatments of neurodegenerative conditions through modulation of the mitochondrial-lysosomal axis.

3.5.3 Mitochondrial Dynamics, Mitophagy, and Neurodegeneration All of the factors that have been implicated in aging also play a role in neurodegeneration. It is, therefore, not surprising that neurodegeneration is often a late-onset, age-associated disease. It has even been suggested that aging is a ‘benign’ form of neurodegeneration. Excessive and unbalanced mitochondrial dynamics have been shown to have deleterious effects in a variety of tissues and cell types including, but not limited to, pancreatic b-cells, skeletal and cardiac myocytes, and neurons. Mitochondrial dynamics and mitophagy are required for quality control in neurons, and it has been shown that both are essential for normal neuronal function and disruptions in these processes can lead to disease states. In particular, dysfunctional mitophagy has been implicated and extensively studied in the pathogenesis of Parkinson’s disease, although it has also been associated with Alzheimer’s disease, amyotrophic lateral sclerosis (ALS), and Huntington’s disease. Parkinson’s disease is the most common neurodegenerative movement disorder and mutations in PINK1 and PARKIN genes are the most frequent cause of autosomal recessive familial Parkinson’s disease. PD is characterized by degeneration of dopaminergic neurons in the substantia nigra and often by the formation of protein aggregates called Lewy bodies. The primary symptom of PD is a progressive dysfunction in movement, including impairment in motor skills and speech. Interestingly, chemical compounds that inhibit complex I of the electron transport chain (ETC), such as rotenone or MPTP, also cause PD-like symptoms in humans and rodents. Mitochondrial dysfunction has been observed in both human PD and murine animal models of the disease. Deletions and mutations in mtDNA have been found in these models as well as increased oxidative stress. A decrease in mitochondrial respiratory complexes is present in PD models and patients (including sporadic PD

3  Mitochondrial Dynamics and Autophagy

97

patients), specifically a deficiency in complex I expression and activity has been observed. An accumulation of a-synuclein, the major component of Lewy bodies, in mitochondria of dopaminergic neurons may be responsible, at least in part, for the ETC complex deficiency and also increased ROS production. PINK1 deficient mice and Drosophila display age-related decreases in complex I activity and mitochondrial depolarization. Of the 13 ETC subunits encoded for by mtDNA, eight encode for complex I, again implying that mutations or deletions in mtDNA in PD may be responsible for the loss of complex I activity. There is experimental evidence that mitochondrial fusion and fission are altered in animal models of PD. Neurotoxin exposure (such as MPTP) leads to mitochondrial fragmentation and swelling along with increased levels of ROS, mitochondrial dysfunction, and mitophagy. There is also data showing morphological changes in various tissues from PD patients. In brain biopsies from PD patients, heterogeneity in mitochondrial size was observed, as well as mitochondrial swelling. Increased ERK1/2 phosphorylation localized to mitochondria in autophagomes has also been observed in brains of PD patients, which is indicative of an induction of mitophagy and/or impaired clearance of autophagosomes (Zhu et al. 2003; Dagda et al. 2008). Studies of cytoplasmic hybrid (cybrid) cell lines, which contain mtDNA from PD subjects, have shown alterations in mitochondrial respiration and dramatic restructuring of mitochondrial architecture (Esteves et al. 2010), including mitochondrial swelling, changes to cristae structure and the outer mitochondrial membrane. Autophagy and mitophagy dysfunction in PD have been a focus of research in this field. It has been reported that in human PD both increased autophagy and mitophagy are present (Zhu and Chu 2010). Activation of the autophagic response is observed in peripheral blood mononuclear cells (PBMCs) from PD patients, which is hypothesized to be a protective mechanism against abnormal protein accumulation and help prevent or slow a-synuclein aggregation (Prigione et  al. 2010). This finding from peripheral tissue may also represent an impairment of flux, with increased signaling for activation of autophagy concurrent with decreased clearance of autophagosomes. Dementia with Lewy Bodies (DLB) patients display elevated levels of neuronal mTOR and reduced Atg7 expression, although this was not observed in brains of PD patients (Crews et al. 2010). The DLB patients also showed increased expression of lysosomal markers in neurons, with the presence of enlarged lysosomes and abundant, abnormal autophagosomes. In addition, alterations in autophagy and/or mitophagy have been reported in other models of PD, such as MPTP and PINK1 or parkin deficiency. A loss in the ability of neurons to clear damaged mitochondria or proteins may be a main mechanism in the pathogenesis of PD. It is evident that PD is associated with impairment in the ability of mitochondria to regulate repair, recovery, and/or recycling of damaged units. Yet these changes to mitochondrial function do not reveal whether the alterations in mitochondrial function and the quality control axis are a cause or consequence of PD. Although, familial PD caused by mutations in PINK1 and parkin suggests that mitochondrial quality control plays a mechanistic role in the early pathogenesis of PD. Additionally, alterations in mitochondrial fusion and fission proteins are known to

98

L. Stiles et al.

be associated with human neurodegenerative disorders. Mitochondrial dysfunction has been implicated in a number of diseases and determining the mechanisms by which these pathways impact disease progression may provide pharmaceutical targets for treatment.

3.5.4 The A9-Dopaminergic Neuron: A Mitochondrial Perfect Storm Selective vulnerability is a common characteristic of many neurodegenerative diseases such as Parkinson’s disease, Alzheimer’s disease, Huntington’s disease and amyotrophic lateral sclerosis. These diseases are marked by a patterned pathology in specific neuronal sub-populations that likely provides a clue to the disease mechanism. Of all the cells in the body, neurons are among the most sensitive to interruptions in mitochondria dynamics, motility, and autophagy. This vulnerability results from extreme energy demands and complex, polarized cell structures, exemplified by neurons that degenerate in forms of parkinsonism, the A9-subtype dopaminergic neurons of the substantia nigra pars compacta (A9-DA). The A9-DA neurons have multiple physical and physiological characteristics that likely render them highly susceptible to loss of function in the mitochondrial network (Fig. 3.5). Nigral A9-DA neurons are examples of extreme physical polarization with somata accounting for less than 1% of cell volume due to massive axonal and dendritic branching (Sulzer 2007). These projection neurons have long, thin, poorlymyelinated axons that terminate in the striatum (Braak et al. 2004; Matsuda et al. 2009). Each of these morphological aspects increases metabolic requirements and risk of oxidative stress. Long axons raises ATP demand and travel time needed for transporting cargo back and forth from soma to synapse. Longer transport times for damaged mitochondria autophagocytosed at synapses increases risk and extent of oxidative damage en route to lysosomes located in the soma (Yue 2007; Terman et al. 2010). Like lanes on a highway, thin axons have reduced transport capacity. This spatial limitation impairs motility for both mitochondria and autophagosomes, and also has the potential to lengthen transport time for mitophagic vesicles. Increased surface to volume ratios cause thin caliber axons to have elevated Na+/K+ ATPase activity, this in turn increases energy demands and has been shown to directly impair mitochondrial motility by inhibiting transport by miro (Zhang et al. 2010). Myelination has a more obvious impact on energy demands, as unmyelinated axons have a less efficient action potential conduction mechanism. Moreover, well-myelinated axons benefit by concentrating their mitochondria at nodes of Ranvier the unmyelinated gaps at the sites of highest Na+/K+ ATPase activity. The thin, poorly-myelinated axons of A9-DA neurons require high Na+/K+ ATPase activity throughout the length of the axon and thus are likely denied the benefits of  mitochondrial clustering. Each A9-DA neuron is estimated to have as many as  ~370,000 synapses in their axonal field (Arbuthnott and Wickens 2007;

3  Mitochondrial Dynamics and Autophagy

99

The A9 Dopaminergic Neuron of the Substantia Nigra pars compacta The perfect storm for interruptions in mitochondrial dynamics, motility, and autophagy

Long

Thin

Massive Poor Myelination Arborization

Energy Demands Energy Demands + + Na+ / K + ATPase Na / K ATPase Decreased mitochondrial clustering Motility Consequences Motility Consequences lncreases transit time to lysosome Limit microtubule capacity Energy Demands Na+ / K + ATPase Dynein/Kinesin

Energy Demands Na+ / K + ATPase Synapses Motility Consequences Highly dispersed Mito-network

Na+ / K + ATPase

O O2 2O 2 O2

Fig.  3.5  The A9-dopaminergic neuron has axonal characteristics that simultaneously increase metabolic demand while impeding mitochondrial dynamics, transport, and mitophagy. Long axons have higher energy demands to accommodate transport machinery and longer autophagosomal transit times from synapse to soma. Thin caliber axons have higher metabolic demands from increased Na+ / K + ATPase activity and reduced transport capacity. Lack of nodes of Ranvier also elevates Na+ / K + ATPase activity and necessitates a more distributed source of energy, thus denying the benefits of nodal mitochondrial clustering seen in well-myelinated axons. Massive axonal branching requires a highly dispersed mitochondrial network and increases metabolic demands at energy-intensive synapses

Surmeier  et  al. 2010). Synapses are inherently energy demanding and sites of voltage-gated calcium influx and thus are packed with mitochondria. The tremendous axonal branching of the A9-DA neuron establishes a logistical challenge for its mitochondrial network in terms of dynamics and transport. Synapses are also points of vulnerability and considered the earliest sites of pathology in many forms of neurodegeneration, underscoring the need for effective autophagic clearance. In addition to the extreme energy demands resulting from physical characteristics, unique physiological characteristics of A9-DA neurons places strain on the mitochondrial and lysosomal networks. Surmeier and colleagues have convincingly implicated L-type calcium channels that drive spontaneous activity in A9-DA neurons as key components of their selective vulnerability (Surmeier et al. 2010). Without protective binding proteins such as calbindin, A9-DA neurons rely heavily on mitochondria to constantly supply energy for calcium pumps and to buffer high levels of cytosolic calcium directly. In addition to stress on the mitochondrial

100

L. Stiles et al.

The A9 Dopaminergic Neuron of the Substantia Nigra pars compacta The perfect storm for interruptions in mitochondrial dynamics, motility, and autophagy Soma Dendrites Massive arborization Extreme polarization−Soma 1% of cell volume Pacemaking through L-type Ca2+ channels Lack of protective Ca2+ buffering proteins Exterme Na+ / K+ATPase activity Lipofuscin /neuromelanin-laden lysosomes

2+

Ca 2+ Ca Ca2+

MOA

Nucleus

O2− O2− O2−

Cytosolic Dopamine

Axon Long and thin poor myelinatation Massive ardorization Dopamine release

Mitochondrial Network Bidirectional Transport and Fusion /Fission Dynamics

Microtubule Network

Autophagosomes Lysosomes

Retrograde Transport Endosomal and lysosomal fusion

Healthy and lipofuscin/Neuromelanin-ladden

Fig.  3.6  Morphological and physiological characteristics of the dentrites, soma, and axon of the A9-dopaminergic neuron increase susceptibility to dysfunction in mitochondrial dynamics, motility, and mitophagy. The mitochondrial network is highly dispersed over an exceptionally polarized cell structure yet must accommodate extremely high Na+ / K + ATPase activity. Mitochondria also serve as a protective barrier against toxicities associated with cytosolic dopamine and calcium. Finally, lysosomes laden with lipofuscin and neuromelanin indicate compromised lysosomal capacity for autophagosome clearance

network by calcium overloading, high cytosolic calcium limits mitochondrial transport by binding to Miro and causing detachment from microtubules (Saotome et  al. 2008; Wang and Schwarz 2009; Macaskill et  al. 2009). Another crucial protective role for mitochondria in A9-DA neurons results from the oxidizing properties of dopamine. The outer mitochondrial membrane contains monoamine oxidase (MAO), which is an enzyme essential for defending the A9-DA neurons from toxicity associated with non-vesicular cytosolic dopamine. This protection comes with a tradeoff as increased MAO activity inhibits mitochondrial respiration (Gluck and Zeevalk 2004). Finally, as a consequence of high levels of ROS from mitochondria and dopamine itself, the cell bodies of A9-DA neurons contain lysosomes with high levels of lipofuscin and neuromelanin that likely reduce degradation capacity (Fig. 3.6). Collectively, the morphological and physiological characteristics of the A9-DA neuron represent a perfect storm for heightened sensitivity to disruptions in mitochondrial dynamics, motility, and mitophagy. Acknowledgements  We are grateful to Drs. Marc Liesa, Gilad Twig, and Dani Dagan for insightful comments during the writing of this chapter.

3  Mitochondrial Dynamics and Autophagy

101

References Adhami F, Liao G, Morozov YM, Schloemer A, Schmithorst VJ, Lorenz JN, Dunn RS, Vorhees CV, Wills-Karp M, Degen JL, Davis RJ, Mizushima N, Rakic P, Dardzinski BJ, Holland SK, Sharp FR, and Kuan CY (2006) Cerebral ischemia-hypoxia induces intravascular coagulation and autophagy. Am J Pathol, 169, 566–583. Alexander C, Votruba M, Pesch UE, Thiselton DL, Mayer S, Moore A, Rodriguez M, Kellner U, Leo-Kottler B, Auburger G, Bhattacharya SS, and Wissinger B (2000) OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat Genet, 26, 211–215. Anglade P, Vyas S, Javoy-Agid F, Herrero MT, Michel PP, Marquez J, Mouatt-Prigent A, Ruberg M, Hirsch EC, and Agid Y (1997) Apoptosis and autophagy in nigral neurons of patients with Parkinson’s disease. Histol Histopathol, 12, 25–31. Arbuthnott GW and Wickens J (2007) Space, time and dopamine. Trends Neurosci, 30, 62–69. Arnoult D, Rismanchi N, Grodet A, Roberts RG, Seeburg DP, Estaquier J, Sheng M, and Blackstone C (2005) Bax/Bak-dependent release of DDP/TIMM8a promotes Drp1-mediated mitochondrial fission and mitoptosis during programmed cell death. Curr Biol, 15, 2112–2118. Barsoum MJ, Yuan H, Gerencser AA, Liot G, Kushnareva Y, Graber S, Kovacs I, Lee WD, Waggoner J, Cui J, White AD, Bossy B, Martinou JC, Youle RJ, Lipton SA, Ellisman MH, Perkins GA, and Bossy-Wetzel E (2006) Nitric oxide-induced mitochondrial fission is regulated by dynamin-related GTPases in neurons. EMBO J, 25, 3900–3911. Bereiter-Hahn J and Voth M (1994) Dynamics of mitochondria in living cells: shape changes, dislocations, fusion, and fission of mitochondria. Microsc Res Tech, 27, 198–219. Braak H, Ghebremedhin E, Rub U, Bratzke H, and Del TK (2004) Stages in the development of Parkinson’s disease-related pathology. Cell Tissue Res, 318, 121–134. Campbell CL and Thorsness PE (1998) Escape of mitochondrial DNA to the nucleus in yme1 yeast is mediated by vacuolar-dependent turnover of abnormal mitochondrial compartments. J Cell Sci, 111 ( Pt 16), 2455–2464. Chen H and Chan DC (2005) Emerging functions of mammalian mitochondrial fusion and fission. Hum Mol Genet, 14 Spec No. 2, R283-R289. Chen H and Chan DC (2009) Mitochondrial dynamics--fusion, fission, movement, and mitophagy-in neurodegenerative diseases. Hum Mol Genet, 18, R169-R176. Chen H and Chan DC (2010) Physiological functions of mitochondrial fusion. Ann N Y Acad Sci, 1201, 21–25. Chu CT (2010) Tickled PINK1: mitochondrial homeostasis and autophagy in recessive Parkinsonism. Biochim Biophys Acta, 1802, 20–28. Clark IE, Dodson MW, Jiang C, Cao JH, Huh JR, Seol JH, Yoo SJ, Hay BA, and Guo M (2006) Drosophila pink1 is required for mitochondrial function and interacts genetically with parkin. Nature, 441, 1162–1166. Clark SL Jr (1957) Cellular differentiation in the kidneys of newborn mice studies with the electron microscope. J Biophys Biochem Cytol, 3, 349–362. Crews L, Spencer B, Desplats P, Patrick C, Paulino A, Rockenstein E, Hansen L, Adame A, Galasko D, and Masliah E (2010) Selective molecular alterations in the autophagy pathway in patients with Lewy body disease and in models of alpha-synucleinopathy. PLoS One, 5, e9313. Cuervo AM (2006) Autophagy in neurons: it is not all about food. Trends Mol Med, 12, 461–464. Cui M, Tang X, Christian WV, Yoon Y, and Tieu K (2010) Perturbations in mitochondrial dynamics induced by human mutant PINK1 can be rescued by the mitochondrial division inhibitor mdivi-1. J Biol Chem, 285, 11740–11752. Dagda RK, Cherra SJ, III, Kulich SM, Tandon A, Park D, and Chu CT (2009) Loss of PINK1 function promotes mitophagy through effects on oxidative stress and mitochondrial fission. J Biol Chem, 284, 13843–13855.

102

L. Stiles et al.

Dagda RK and Chu CT (2009) Mitochondrial quality control: insights on how Parkinson’s disease related genes PINK1, parkin, and Omi/HtrA2 interact to maintain mitochondrial homeostasis. J Bioenerg Biomembr, 41, 473–479. Dagda RK, Zhu J, Kulich SM, and Chu CT (2008) Mitochondrially localized ERK2 regulates mitophagy and autophagic cell stress: implications for Parkinson’s disease. Autophagy, 4, 770–782. de Brito OM and Scorrano L (2008) Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature, 456, 605–610. Dehay B, Bove J, Rodriguez-Muela N, Perier C, Recasens A, Boya P, and Vila M (2010) Pathogenic lysosomal depletion in Parkinson’s disease. J Neurosci, 30, 12535–12544. Deng H, Dodson MW, Huang H, and Guo M (2008) The Parkinson’s disease genes pink1 and parkin promote mitochondrial fission and/or inhibit fusion in Drosophila. Proc Natl Acad Sci USA, 105, 14503–14508. Deter RL, Baudhuin P, and De DC (1967) Participation of lysosomes in cellular autophagy induced in rat liver by glucagon. J Cell Biol, 35, C11-C16. Deter RL and De DC (1967) Influence of glucagon, an inducer of cellular autophagy, on some physical properties of rat liver lysosomes. J Cell Biol, 33, 437–449. Ding WX, Ni HM, Li M, Liao Y, Chen X, Stolz DB, Dorn Ii GW, and Yin XM (2010) Nix is critical to two distinct phases of mitophagy: reactive oxygen species (ROS)-mediated autophagy induction and Parkin-ubiqutin-p62-mediated mitochondria priming. J Biol Chem Duvezin-Caubet S, Jagasia R, Wagener J, Hofmann S, Trifunovic A, Hansson A, Chomyn A, Bauer MF, Attardi G, Larsson NG, Neupert W, and Reichert AS (2006) Proteolytic processing of OPA1 links mitochondrial dysfunction to alterations in mitochondrial morphology. J Biol Chem, 281, 37972–37979. Elmore SP, Qian T, Grissom SF, and Lemasters JJ (2001) The mitochondrial permeability transition initiates autophagy in rat hepatocytes. FASEB J, 15, 2286–2287. Eskelinen EL (2008) New insights into the mechanisms of macroautophagy in mammalian cells. Int Rev Cell Mol Biol, 266, 207–247. Essick EE and Sam F (2010) Oxidative stress and autophagy in cardiac disease, neurological disorders, aging and cancer. Oxid Med Cell Longev, 3, 168–177. Esteves AR, Lu J, Rodova M, Onyango I, Lezi E, Dubinsky R, Lyons KE, Pahwa R, Burns JM, Cardoso SM, and Swerdlow RH (2010) Mitochondrial respiration and respiration-associated proteins in cell lines created through Parkinson’s subject mitochondrial transfer. J Neurochem, 113, 674–682. Exner N, Treske B, Paquet D, Holmstrom K, Schiesling C, Gispert S, Carballo-Carbajal I, Berg D, Hoepken HH, Gasser T, Kruger R, Winklhofer KF, Vogel F, Reichert AS, Auburger G, Kahle PJ, Schmid B, and Haass C (2007) Loss-of-function of human PINK1 results in mitochondrial pathology and can be rescued by parkin. J Neurosci, 27, 12413–12418. Fransson S, Ruusala A, and Aspenstrom P (2006) The atypical Rho GTPases Miro-1 and Miro-2 have essential roles in mitochondrial trafficking. Biochem Biophys Res Commun, 344, 500–510. Frieden M, James D, Castelbou C, Danckaert A, Martinou JC, and Demaurex N (2004) Ca(2+) homeostasis during mitochondrial fragmentation and perinuclear clustering induced by hFis1. J Biol Chem, 279, 22704–22714. Fujita N, Hayashi-Nishino M, Fukumoto H, Omori H, Yamamoto A, Noda T, and Yoshimori T (2008) An Atg4B mutant hampers the lipidation of LC3 paralogues and causes defects in autophagosome closure. Mol Biol Cell, 19, 4651–4659. Fujitani Y, Ueno T, and Watada H (2010) Autophagy in health and disease. 4. The role of pancreatic beta-cell autophagy in health and diabetes. Am J Physiol Cell Physiol, 299, C1-C6. Gandhi S, Wood-Kaczmar A, Yao Z, Plun-Favreau H, Deas E, Klupsch K, Downward J, Latchman DS, Tabrizi SJ, Wood NW, Duchen MR, and Abramov AY (2009) PINK1-associated Parkinson’s disease is caused by neuronal vulnerability to calcium-induced cell death. Mol Cell, 33, 627–638. Gasser T (2009) Molecular pathogenesis of Parkinson disease: insights from genetic studies. Expert Rev Mol Med, 11, e22.

3  Mitochondrial Dynamics and Autophagy

103

Gegg ME, Cooper JM, Chau KY, Rojo M, Schapira AH, and Taanman JW (2010) Mitofusin 1 and mitofusin 2 are ubiquitinated in a PINK1/parkin-dependent manner upon induction of mitophagy. Hum Mol Genet, 19, 4861–4870. Geisler S, Holmstrom KM, Skujat D, Fiesel FC, Rothfuss OC, Kahle PJ, and Springer W (2010) PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nat Cell Biol, 12, 119–131. Gispert S, Ricciardi F, Kurz A, Azizov M, Hoepken HH, Becker D, Voos W, Leuner K, Muller WE, Kudin AP, Kunz WS, Zimmermann A, Roeper J, Wenzel D, Jendrach M, Garcia-Arencibia M, Fernandez-Ruiz J, Huber L, Rohrer H, Barrera M, Reichert AS, Rub U, Chen A, Nussbaum RL, and Auburger G (2009) Parkinson phenotype in aged PINK1-deficient mice is accompanied by progressive mitochondrial dysfunction in absence of neurodegeneration. PLoS One, 4, e5777. Glater EE, Megeath LJ, Stowers RS, and Schwarz TL (2006) Axonal transport of mitochondria requires milton to recruit kinesin heavy chain and is light chain independent. J Cell Biol, 173, 545–557. Glick D, Barth S, and Macleod KF (2010) Autophagy: cellular and molecular mechanisms. J Pathol, 221, 3–12. Gluck MR and Zeevalk GD (2004) Inhibition of brain mitochondrial respiration by dopamine and its metabolites: implications for Parkinson’s disease and catecholamine-associated diseases. J Neurochem, 91, 788–795. Goldberg MS, Fleming SM, Palacino JJ, Cepeda C, Lam HA, Bhatnagar A, Meloni EG, Wu N, Ackerson LC, Klapstein GJ, Gajendiran M, Roth BL, Chesselet MF, Maidment NT, Levine MS, and Shen J (2003) Parkin-deficient mice exhibit nigrostriatal deficits but not loss of dopaminergic neurons. J Biol Chem, 278, 43628–43635. Goldman SJ, Taylor R, Zhang Y, and Jin S (2010) Autophagy and the degradation of mitochondria. Mitochondrion, 10, 309–315. Gomes LC and Scorrano L (2008) High levels of Fis1, a pro-fission mitochondrial protein, trigger autophagy. Biochim Biophys Acta, 1777, 860–866. Gottlieb RA and Carreira RS (2010) Autophagy in health and disease. 5. Mitophagy as a way of life. Am J Physiol Cell Physiol, 299, C203-C210. Griparic L, van der Wel NN, Orozco IJ, Peters PJ, and van der Bliek AM (2004) Loss of the intermembrane space protein Mgm1/OPA1 induces swelling and localized constrictions along the lengths of mitochondria. J Biol Chem, 279, 18792–18798. Grunewald A, Gegg ME, Taanman JW, King RH, Kock N, Klein C, and Schapira AH (2009) Differential effects of PINK1 nonsense and missense mutations on mitochondrial function and morphology. Exp Neurol, 219, 266–273. Hailey DW, Rambold AS, Satpute-Krishnan P, Mitra K, Sougrat R, Kim PK, and LippincottSchwartz J (2010) Mitochondria supply membranes for autophagosome biogenesis during starvation. Cell, 141, 656–667. Hara T, Nakamura K, Matsui M, Yamamoto A, Nakahara Y, Suzuki-Migishima R, Yokoyama M, Mishima K, Saito I, Okano H, and Mizushima N (2006) Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature, 441, 885–889. Harman D (1956) Aging: a theory based on free radical and radiation chemistry. J Gerontol, 11, 298–300. Hyde BB, Twig G, and Shirihai OS (2010) Organellar vs cellular control of mitochondrial dynamics. Semin Cell Dev Biol, 21, 575–581. James DI, Parone PA, Mattenberger Y, and Martinou JC (2003) hFis1, a novel component of the mammalian mitochondrial fission machinery. J Biol Chem, 278, 36373–36379. Ju JS and Weihl CC (2010) Inclusion body myopathy, Paget’s disease of the bone and frontotemporal dementia: a disorder of autophagy. Hum Mol Genet, 19, R38-R45. Kanki T and Klionsky DJ (2010) The molecular mechanism of mitochondria autophagy in yeast. Mol Microbiol, 75, 795–800. Kanki T, Wang K, Baba M, Bartholomew CR, Lynch-Day MA, Du Z, Geng J, Mao K, Yang Z, Yen WL, and Klionsky DJ (2009a) A genomic screen for yeast mutants defective in selective mitochondria autophagy. Mol Biol Cell, 20, 4730–4738.

104

L. Stiles et al.

Kanki T, Wang K, Cao Y, Baba M, and Klionsky DJ (2009b) Atg32 is a mitochondrial protein that confers selectivity during mitophagy. Dev Cell, 17, 98–109. Karbowski M (2010) Mitochondria on guard: role of mitochondrial fusion and fission in the regulation of apoptosis. Adv Exp Med Biol, 687, 131–142. Kissova I, Deffieu M, Manon S, and Camougrand N (2004) Uth1p is involved in the autophagic degradation of mitochondria. J Biol Chem, 279, 39068–39074. Kitada T, Pisani A, Porter DR, Yamaguchi H, Tscherter A, Martella G, Bonsi P, Zhang C, Pothos EN, and Shen J (2007) Impaired dopamine release and synaptic plasticity in the striatum of PINK1-deficient mice. Proc Natl Acad Sci USA, 104, 11441–11446. Kitada T, Tong Y, Gautier CA, and Shen J (2009) Absence of nigral degeneration in aged parkin/ DJ-1/PINK1 triple knockout mice. J Neurochem, 111, 696–702. Klionsky DJ (2007) Autophagy: from phenomenology to molecular understanding in less than a decade. Nat Rev Mol Cell Biol, 8, 931–937. Koch A, Thiemann M, Grabenbauer M, Yoon Y, McNiven MA, and Schrader M (2003) Dynaminlike protein 1 is involved in peroxisomal fission. J Biol Chem, 278, 8597–8605. Koch A, Yoon Y, Bonekamp NA, McNiven MA, and Schrader M (2005) A role for Fis1 in both mitochondrial and peroxisomal fission in mammalian cells. Mol Biol Cell, 16, 5077–5086. Komatsu M, Waguri S, Chiba T, Murata S, Iwata J, Tanida I, Ueno T, Koike M, Uchiyama Y, Kominami E, and Tanaka K (2006) Loss of autophagy in the central nervous system causes neurodegeneration in mice. Nature, 441, 880–884. Komatsu M, Wang QJ, Holstein GR, Friedrich VL, Jr., Iwata J, Kominami E, Chait BT, Tanaka K, and Yue Z (2007) Essential role for autophagy protein Atg7 in the maintenance of axonal homeostasis and the prevention of axonal degeneration. Proc Natl Acad Sci USA, 104, 14489–14494. Kundu M, Lindsten T, Yang CY, Wu J, Zhao F, Zhang J, Selak MA, Ney PA, and Thompson CB (2008) Ulk1 plays a critical role in the autophagic clearance of mitochondria and ribosomes during reticulocyte maturation. Blood, 112, 1493–1502. Kurz T, Terman A, Gustafsson B, Brunk UT (2008a) Lysosomes and oxidative stress in ageing and apoptosis. Biochim Biophys Acta, 1780, 1291–1303. Kurz T, Terman A, Gustafsson B, Brunk UT (2008b) Lysosomes in iron metabolism, ageing, and apoptosis. Histochem Cell Biol, 129, 389–406. Lee JA and Gao FB (2008a) ESCRT, autophagy, and frontotemporal dementia. BMB Rep, 41, 827–832. Lee JH, Yu WH, Kumar A, Lee S, Mohan PS, Peterhoff CM, Wolfe DM, Martinez-Vicente M, Massey AC, Sovak G, Uchiyama Y, Westaway D, Cuervo AM, Nixon RA (2010) Lysosomal proteolysis and autophagy require presenilin 1 and are disrupted by Alzheimer-related PS1 mutations. Cell, 141, 1146–1158. Lee JY, Nagano Y, Taylor JP, Lim KL, and Yao TP (2010) Disease-causing mutations in parkin impair mitochondrial ubiquitination, aggregation, and HDAC6-dependent mitophagy. J Cell Biol, 189, 671–679. Lemasters JJ (2005) Selective mitochondrial autophagy, or mitophagy, as a targeted defense against oxidative stress, mitochondrial dysfunction, and aging. Rejuvenation Res, 8, 3–5. Lemasters JJ, Nieminen AL, Qian T, Trost LC, Elmore SP, Nishimura Y, Crowe RA, Cascio WE, Bradham CA, Brenner DA, and Herman B (1998) The mitochondrial permeability transition in cell death: a common mechanism in necrosis, apoptosis and autophagy. Biochim Biophys Acta, 1366, 177–196. Liesa M, Palacin M, and Zorzano A (2009) Mitochondrial dynamics in mammalian health and disease. Physiol Rev, 89, 799–845. Liu W, Vives-Bauza C, Acin P, Yamamoto A, Tan Y, Li Y, Magrane J, Stavarache MA, Shaffer S, Chang S, Kaplitt MG, Huang XY, Beal MF, Manfredi G, and Li C (2009a) PINK1 defect causes mitochondrial dysfunction, proteasomal deficit and alpha-synuclein aggregation in cell culture models of Parkinson’s disease. PLoS One, 4, e4597. Liu X and Hajnoczky G (2009) Ca 2+−dependent regulation of mitochondrial dynamics by the Miro-Milton complex. Int J Biochem Cell Biol, 41, 1972–1976. Liu X, Weaver D, Shirihai O, and Hajnoczky G (2009b) Mitochondrial ‘kiss-and-run’: interplay between mitochondrial motility and fusion-fission dynamics. EMBO J, 28, 3074–3089.

3  Mitochondrial Dynamics and Autophagy

105

Lutz AK, Exner N, Fett ME, Schlehe JS, Kloos K, Lammermann K, Brunner B, Kurz-Drexler A, Vogel F, Reichert AS, Bouman L, Vogt-Weisenhorn D, Wurst W, Tatzelt J, Haass C, and Winklhofer KF (2009) Loss of parkin or PINK1 function increases Drp1-dependent mitochondrial fragmentation. J Biol Chem, 284, 22938–22951. Macaskill AF, Rinholm JE, Twelvetrees AE, Rancibia-Carcamo IL, Muir J, Fransson A, Aspenstrom P, Attwell D, and Kittler JT (2009) Miro1 is a calcium sensor for glutamate receptor-dependent localization of mitochondria at synapses. Neuron, 61, 541–555. Marongiu R, Spencer B, Crews L, Adame A, Patrick C, Trejo M, Dallapiccola B, Valente EM, and Masliah E (2009) Mutant Pink1 induces mitochondrial dysfunction in a neuronal cell model of Parkinson’s disease by disturbing calcium flux. J Neurochem, 108, 1561–1574. Matsuda W, Furuta T, Nakamura KC, Hioki H, Fujiyama F, Arai R, and Kaneko T (2009) Single nigrostriatal dopaminergic neurons form widely spread and highly dense axonal arborizations in the neostriatum. J Neurosci, 29, 444–453. Mazure NM and Pouyssegur J (2009) Atypical BH3-domains of BNIP3 and BNIP3L lead to autophagy in hypoxia. Autophagy, 5, 868–869. Merz S, Hammermeister M, Altmann K, Durr M, and Westermann B (2007) Molecular machinery of mitochondrial dynamics in yeast. Biol Chem, 388, 917–926. Mills RD, Sim CH, Mok SS, Mulhern TD, Culvenor JG, and Cheng HC (2008) Biochemical aspects of the neuroprotective mechanism of PTEN-induced kinase-1 (PINK1). J Neurochem, 105, 18–33. Misaka T, Miyashita T, and Kubo Y (2002) Primary structure of a dynamin-related mouse mitochondrial GTPase and its distribution in brain, subcellular localization, and effect on mitochondrial morphology. J Biol Chem, 277, 15834–15842. Misko A, Jiang S, Wegorzewska I, Milbrandt J, and Baloh RH (2010) Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex. J Neurosci, 30, 4232–4240. Moreira PI, Siedlak SL, Wang X, Santos MS, Oliveira CR, Tabaton M, Nunomura A, Szweda LI, Aliev G, Smith MA, Zhu X, and Perry G (2007a) Autophagocytosis of mitochondria is prominent in Alzheimer disease. J Neuropathol Exp Neurol, 66, 525–532. Moreira PI, Siedlak SL, Wang X, Santos MS, Oliveira CR, Tabaton M, Nunomura A, Szweda LI, Aliev G, Smith MA, Zhu X, and Perry G (2007b) Increased autophagic degradation of mitochondria in Alzheimer disease. Autophagy, 3, 614–615. Mouli PK, Twig G, and Shirihai OS (2009) Frequency and selectivity of mitochondrial fusion are key to its quality maintenance function. Biophys J, 96, 3509–3518. Nakada K, Inoue K, Ono T, Isobe K, Ogura A, Goto YI, Nonaka I, and Hayashi JI (2001) Intermitochondrial complementation: Mitochondria-specific system preventing mice from expression of disease phenotypes by mutant mtDNA. Nat Med, 7, 934–940. Narendra DP, Jin SM, Tanaka A, Suen DF, Gautier CA, Shen J, Cookson MR, and Youle RJ (2010) PINK1 is selectively stabilized on impaired mitochondria to activate Parkin. PLoS Biol, 8, e1000298. Navratil M, Terman A, and Arriaga EA (2008) Giant mitochondria do not fuse and exchange their contents with normal mitochondria. Exp Cell Res, 314, 164–172. Nice DC, Sato TK, Stromhaug PE, Emr SD, and Klionsky DJ (2002) Cooperative binding of the cytoplasm to vacuole targeting pathway proteins, Cvt13 and Cvt20, to phosphatidylinositol 3-phosphate at the pre-autophagosomal structure is required for selective autophagy. J Biol Chem, 277, 30198–30207. Nishida Y, Arakawa S, Fujitani K, Yamaguchi H, Mizuta T, Kanaseki T, Komatsu M, Otsu K, Tsujimoto Y, and Shimizu S (2009) Discovery of Atg5/Atg7-independent alternative macroautophagy. Nature, 461, 654–658. Nixon RA (2006) Autophagy in neurodegenerative disease: friend, foe or turncoat? Trends Neurosci, 29, 528–535. Nixon RA, Wegiel J, Kumar A, Yu WH, Peterhoff C, Cataldo A, and Cuervo AM (2005) Extensive involvement of autophagy in Alzheimer disease: an immuno-electron microscopy study. J Neuropathol Exp Neurol, 64, 113–122.

106

L. Stiles et al.

Nixon RA, Yang DS, Lee JH (2008) Neurodegenerative lysosomal disorders: a continuum from development to late age. Autophagy, 4, 590–599. Nowikovsky K, Reipert S, Devenish RJ, and Schweyen RJ (2007) Mdm38 protein depletion causes loss of mitochondrial K+/H + exchange activity, osmotic swelling and mitophagy. Cell Death Differ, 14, 1647–1656. Okamoto K, Kondo-Okamoto N, and Ohsumi Y (2009a) A landmark protein essential for mitophagy: Atg32 recruits the autophagic machinery to mitochondria. Autophagy, 5, 1203–1205. Okamoto K, Kondo-Okamoto N, and Ohsumi Y (2009b) Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev Cell, 17, 87–97. Orenstein SJ and Cuervo AM (2010) Chaperone-mediated autophagy: Molecular mechanisms and physiological relevance. Semin Cell Dev Biol. Park J, Lee G, and Chung J (2009) The PINK1-Parkin pathway is involved in the regulation of mitochondrial remodeling process. Biochem Biophys Res Commun, 378, 518–523. Park KS, Wiederkehr A, Kirkpatrick C, Mattenberger Y, Martinou JC, Marchetti P, Demaurex N, and Wollheim CB (2008) Selective actions of mitochondrial fission/fusion genes on metabolism-secretion coupling in insulin-releasing cells. J Biol Chem, 283, 33347–33356. Parone PA, Da CS, Tondera D, Mattenberger Y, James DI, Maechler P, Barja F, and Martinou JC (2008) Preventing mitochondrial fission impairs mitochondrial function and leads to loss of mitochondrial DNA. PLoS One, 3, e3257. Poole AC, Thomas RE, Andrews LA, McBride HM, Whitworth AJ, and Pallanck LJ (2008) The PINK1/Parkin pathway regulates mitochondrial morphology. Proc Natl Acad Sci USA, 105, 1638–1643. Poole AC, Thomas RE, Yu S, Vincow ES, and Pallanck L (2010) The mitochondrial fusion-promoting factor mitofusin is a substrate of the PINK1/parkin pathway. PLoS One, 5, e10054. Praefcke GJ and McMahon HT (2004) The dynamin superfamily: universal membrane tubulation and fission molecules? Nat Rev Mol Cell Biol, 5, 133–147. Prigione A, Piazza F, Brighina L, Begni B, Galbussera A, Difrancesco JC, Andreoni S, Piolti R, and Ferrarese C (2010) Alpha-synuclein nitration and autophagy response are induced in peripheral blood cells from patients with Parkinson disease. Neurosci Lett, 477, 6–10. Reggiori F, Tucker KA, Stromhaug PE, and Klionsky DJ (2004) The Atg1-Atg13 complex regulates Atg9 and Atg23 retrieval transport from the pre-autophagosomal structure. Dev Cell, 6, 79–90. Sandebring A, Thomas KJ, Beilina A, Vander BM, Cleland MM, Ahmad R, Miller DW, Zambrano I, Cowburn RF, Behbahani H, Cedazo-Minguez A, and Cookson MR (2009) Mitochondrial alterations in PINK1 deficient cells are influenced by calcineurin-dependent dephosphorylation of dynamin-related protein 1. PLoS One, 4, e5701. Santel A and Fuller MT (2001) Control of mitochondrial morphology by a human mitofusin. J Cell Sci, 114, 867–874. Santos RX, Correia SC, Wang X, Perry G, Smith MA, Moreira PI, and Zhu X (2010) A synergistic dysfunction of mitochondrial fission/fusion dynamics and mitophagy in Alzheimer’s disease. J Alzheimers Dis, 20 Suppl 2, S401-S412. Saotome M, Safiulina D, Szabadkai G, Das S, Fransson A, Aspenstrom P, Rizzuto R, and Hajnoczky G (2008) Bidirectional Ca2+−dependent control of mitochondrial dynamics by the Miro GTPase. Proc Natl Acad Sci USA, 105, 20728–20733. Scheckhuber CQ, Erjavec N, Tinazli A, Hamann A, Nystrom T, and Osiewacz HD (2007) Reducing mitochondrial fission results in increased life span and fitness of two fungal ageing models. Nat Cell Biol, 9, 99–105. Song Z, Chen H, Fiket M, Alexander C, and Chan DC (2007) OPA1 processing controls mitochondrial fusion and is regulated by mRNA splicing, membrane potential, and Yme1L. J Cell Biol, 178, 749–755. Sou YS, Waguri S, Iwata J, Ueno T, Fujimura T, Hara T, Sawada N, Yamada A, Mizushima N, Uchiyama Y, Kominami E, Tanaka K, and Komatsu M (2008) The Atg8 conjugation system is indispensable for proper development of autophagic isolation membranes in mice. Mol Biol Cell, 19, 4762–4775.

3  Mitochondrial Dynamics and Autophagy

107

Stack JH, DeWald DB, Takegawa K, and Emr SD (1995) Vesicle-mediated protein transport: regulatory interactions between the Vps15 protein kinase and the Vps34 PtdIns 3-kinase essential for protein sorting to the vacuole in yeast. J Cell Biol, 129, 321–334. Stephenson FA (2010) Activity-dependent immobilization of mitochondria: the role of miro. Front Mol Neurosci, 3, 9. Sulzer D (2007) Multiple hit hypotheses for dopamine neuron loss in Parkinson’s disease. Trends Neurosci, 30, 244–250. Surmeier DJ, Guzman JN, Sanchez-Padilla J, and Goldberg JA (2010) What causes the death of dopaminergic neurons in Parkinson’s disease? Prog Brain Res, 183, 59–77. Takeshige K, Baba M, Tsuboi S, Noda T, and Ohsumi Y (1992) Autophagy in yeast demonstrated with proteinase-deficient mutants and conditions for its induction. J Cell Biol, 119, 301–311. Tal R, Winter G, Ecker N, Klionsky DJ, and Abeliovich H (2007) Aup1p, a yeast mitochondrial protein phosphatase homolog, is required for efficient stationary phase mitophagy and cell survival. J Biol Chem, 282, 5617–5624. Tanida I, Sou YS, Minematsu-Ikeguchi N, Ueno T, and Kominami E (2006) Atg8L/Apg8L is the fourth mammalian modifier of mammalian Atg8 conjugation mediated by human Atg4B, Atg7 and Atg3. FEBS J, 273, 2553–2562. Terman A, Kurz T, Gustafsson B, Brunk UT (2006) Lysosomal Labilization. IUBMB Life, 58, 531–539. Terman A, Kurz T, Navratil M, Arriaga EA, and Brunk UT (2010) Mitochondrial turnover and aging of long-lived postmitotic cells: the mitochondrial-lysosomal axis theory of aging. Antioxid Redox Signal, 12, 503–535. Thomas KJ, McCoy MK, Blackinton J, Beilina A, van der BM, Sandebring A, Miller D, Maric D, Cedazo-Minguez A, and Cookson MR (2010) DJ-1 acts in parallel to the PINK1/parkin pathway to control mitochondrial function and autophagy. Hum Mol Genet. Twig G, Elorza A, Molina AJ, Mohamed H, Wikstrom JD, Walzer G, Stiles L, Haigh SE, Katz S, Las G, Alroy J, Wu M, Py BF, Yuan J, Deeney JT, Corkey BE, and Shirihai OS (2008a) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J, 27, 433–446. Twig G, Hyde B, and Shirihai OS (2008b) Mitochondrial fusion, fission and autophagy as a quality control axis: the bioenergetic view. Biochim Biophys Acta, 1777, 1092–1097. Twig G, Liu X, Liesa M, Wikstrom JD, Molina AJ, Las G, Yaniv G, Hajnoczky G, and Shirihai OS (2010) Biophysical properties of mitochondrial fusion events in pancreatic beta-cells and cardiac cells unravel potential control mechanisms of its selectivity. Am J Physiol Cell Physiol, 299, C477-C487. Twig G and Shirihai OS (2010) The interplay between mitochondrial dynamics and mitophagy. Antioxid Redox Signal. Vila M, Bove J, Dehay B, Rodriguez-Muela N, and Boya P (2010) Lysosomal membrane permeabilization in Parkinson disease. Autophagy, 7, 98–100. Vives-Bauza C and Przedborski S (2010) PINK1 points Parkin to mitochondria. Autophagy, 6. Vives-Bauza C, Zhou C, Huang Y, Cui M, de Vries RL, Kim J, May J, Tocilescu MA, Liu W, Ko HS, Magrane J, Moore DJ, Dawson VL, Grailhe R, Dawson TM, Li C, Tieu K, and Przedborski S (2010) PINK1-dependent recruitment of Parkin to mitochondria in mitophagy. Proc Natl Acad Sci USA, 107, 378–383. Wang X and Schwarz TL (2009) The mechanism of Ca2+ −dependent regulation of kinesinmediated mitochondrial motility. Cell, 136, 163–174. Weber TA and Reichert AS (2010) Impaired quality control of mitochondria: aging from a new perspective. Exp Gerontol, 45, 503–511. Weihofen A, Thomas KJ, Ostaszewski BL, Cookson MR, and Selkoe DJ (2009) Pink1 forms a multiprotein complex with Miro and Milton, linking Pink1 function to mitochondrial trafficking. Biochemistry, 48, 2045–2052. White KE, Davies VJ, Hogan VE, Piechota MJ, Nichols PP, Turnbull DM, and Votruba M (2009) OPA1 deficiency associated with increased autophagy in retinal ganglion cells in a murine model of dominant optic atrophy. Invest Ophthalmol Vis Sci, 50, 2567–2571.

108

L. Stiles et al.

Whitworth AJ and Pallanck LJ (2009) The PINK1/Parkin pathway: a mitochondrial quality control system? J Bioenerg Biomembr, 41, 499–503. Wikstrom JD, Twig G, and Shirihai OS (2009) What can mitochondrial heterogeneity tell us about mitochondrial dynamics and autophagy? Int J Biochem Cell Biol, 41, 1914–1927. Wood-Kaczmar A, Gandhi S, Yao Z, Abramov AY, Miljan EA, Keen G, Stanyer L, Hargreaves I, Klupsch K, Deas E, Downward J, Mansfield L, Jat P, Taylor J, Heales S, Duchen MR, Latchman D, Tabrizi SJ, and Wood NW (2008) PINK1 is necessary for long term survival and mitochondrial function in human dopaminergic neurons. PLoS One, 3, e2455. Yang Y, Ouyang Y, Yang L, Beal MF, McQuibban A, Vogel H, and Lu B (2008) Pink1 regulates mitochondrial dynamics through interaction with the fission/fusion machinery. Proc Natl Acad Sci USA, 105, 7070–7075. Yang Z and Klionsky DJ (2010b) Eaten alive: a history of macroautophagy. Nat Cell Biol, 12, 814–822. Yang Z and Klionsky DJ (2010b) Mammalian autophagy: core molecular machinery and signaling regulation. Curr Opin Cell Biol, 22, 124–131. Yen WL and Klionsky DJ (2008) How to live long and prosper: autophagy, mitochondria, and aging. Physiology (Bethesda ), 23, 248–262. Young AR, Chan EY, Hu XW, Kochl R, Crawshaw SG, High S, Hailey DW, Lippincott-Schwartz J, and Tooze SA (2006) Starvation and ULK1-dependent cycling of mammalian Atg9 between the TGN and endosomes. J Cell Sci, 119, 3888–3900. Young JE, Martinez RA, and La Spada AR (2009) Nutrient deprivation induces neuronal autophagy and implicates reduced insulin signaling in neuroprotective autophagy activation. J Biol Chem, 284, 2363–2373. Yue Z (2007) Regulation of neuronal autophagy in axon: implication of autophagy in axonal function and dysfunction/degeneration. Autophagy, 3, 139–141. Yue Z, Friedman L, Komatsu M, and Tanaka K (2009) The cellular pathways of neuronal autophagy and their implication in neurodegenerative diseases. Biochim Biophys Acta, 1793, 1496–1507. Zhang CL, Ho PL, Kintner DB, Sun D, and Chiu SY (2010) Activity-dependent regulation of mitochondrial motility by calcium and Na/K-ATPase at nodes of Ranvier of myelinated nerves. J Neurosci, 30, 3555–3566. Zhang J, Kundu M, and Ney PA (2009) Mitophagy in mammalian cells: the reticulocyte model. Methods Enzymol, 452, 227–245. Zhang J and Ney PA (2008) NIX induces mitochondrial autophagy in reticulocytes. Autophagy, 4, 354–356. Zhang J and Ney PA (2009) Role of BNIP3 and NIX in cell death, autophagy, and mitophagy. Cell Death Differ, 16, 939–946. Zhang Y, Qi H, Taylor R, Xu W, Liu LF, and Jin S (2007) The role of autophagy in mitochondria maintenance: characterization of mitochondrial functions in autophagy-deficient S cerevisiae strains. Autophagy, 3, 337–346. Zhou X, Babu JR, Da SS, Shu Q, Graef IA, Oliver T, Tomoda T, Tani T, Wooten MW, and Wang F (2007) Unc-51-like kinase 1/2-mediated endocytic processes regulate filopodia extension and branching of sensory axons. Proc Natl Acad Sci USA, 104, 5842–5847. Zhu J and Chu CT (2010) Mitochondrial dysfunction in Parkinson’s disease. J Alzheimers Dis, 20 Suppl 2, S325-S334. Zhu JH, Guo F, Shelburne J, Watkins S, and Chu CT (2003) Localization of phosphorylated ERK/MAP kinases to mitochondria and autophagosomes in Lewy body diseases. Brain Pathol, 13, 473–481. Ziviani E, Tao RN, and Whitworth AJ (2010) Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc Natl Acad Sci USA, 107, 5018–5023. Ziviani E and Whitworth AJ (2010) How could Parkin-mediated ubiquitination of mitofusin promote mitophagy? Autophagy, 6. Zorzano A, Liesa M, Sebastian D, Segales J, and Palacin M (2010) Mitochondrial fusion proteins: dual regulators of morphology and metabolism. Semin Cell Dev Biol, 21, 566–574. Zuchner S, Mersiyanova IV, Muglia M, Bissar-Tadmouri N, Rochelle J, Dadali EL, Zappia M, Nelis E, Patitucci A, Senderek J, Parman Y, Evgrafov O, Jonghe PD, Takahashi Y, Tsuji S, Pericak-Vance MA, Quattrone A, Battaloglu E, Polyakov AV, Timmerman V, Schroder JM, and Vance JM (2004) Mutations in the mitochondrial GTPase mitofusin 2 cause CharcotMarie-Tooth neuropathy type 2A. Nat Genet, 36, 449–451.

Chapter 4

Mitochondrial Dynamics and Apoptosis Megan M. Cleland and Richard J. Youle

Abstract  The dynamic nature of mitochondria is not only important for maintaining normal healthy cells, but is also very important in the timely execution of apoptosis. The machinery involved in mediating mitochondrial fission and fusion, namely the large GTPases Drp1, OPA1, Mfn1 and Mfn2, can be modulated to either intensify or reduce apoptosis. During the intrinsic/mitochondrial apoptotic pathway, Drp1 promotes mitochondrial fragmentation, while proteolytic cleavage and release of OPA1 from the mitochondria enhances this fragmentation and results in a swollen/ altered cristae morphology. Conditions that enhance mitochondrial fusion, such as overexpression of OPA1, Mfn1, Mfn2 or inhibition of Drp1, generally result in a delay of apoptosis while enhancing mitochondrial fission, through overexpression of Drp1 or down regulating OPA1, Mfn1 or Mfn2, augments apoptosis. Keywords  Mitochondrial dynamics • Apoptosis • Mitofusin • OPA1 • Drp1 Abbreviations BH CCCP CMT2A GED IMM IMS mdivi-1 MEFs

Bcl-2 homology Carbonyl cyanide 3-chlorophenylhydrazone Charcot-Marie Tooth Type 2A GTPase effector domain Inner mitochondrial membrane Inner membrane space mitochondrial division inhibitor Mouse embryonic fibroblasts

M.M. Cleland and R.J. Youle (*) Biochemistry Section, Surgical Neurology Branch, National Institute of Neurological Disorders and Stroke (NINDS), National Institutes of Health (NIH), 35 Convent Drive, MSC 3704, Building 35/Rm, 2C917, Bethesda, MD 20892, USA e-mail: [email protected] B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_4, © Springer Science+Business Media B.V. 2011

109

110

MOMP mtPTP OMM OMMAD SIMH TM

M.M. Cleland and R.J. Youle

mitochondrial outer membrane permeabilization Mitochondrial permeability transition pore Outer mitochondrial membrane Outer mitochondrial membrane associated degradation Stress induced mitochondrial hyperfusion Transmembrane

4.1 Introduction 4.1.1 Mitochondria Are Key to Eukaryotic Cell Vitality The maintenance of mitochondrial integrity and function is vital for the health of the eukaryotic cell. Mitochondria perform key roles in a variety of cellular functions including: metabolism; biosynthesis of steroids, heme and iron-sulfur clusters; and maintenance of calcium concentration through calcium buffering. In addition to these prominent cellular functions required for eukaryotic cell viability, mitochondria are central participants in promotion of programmed cell death or apoptosis.

4.1.2 Bcl-2 Family Members Regulate the Intrinsic Mitochondrial Death Pathway Apoptosis (Greek: apo – from/off/without, ptosis – falling) is an effective means for dismantling and cleanly disposing of a cell that has been infected, damaged or genetically programmed to die. Although the first usage of the term apoptosis was used by Hippocrates of Cos in the fourth century BC and the first description of apoptotic cell death attributed to Rudolph Virchow in 1860, it was not until 1972 and the work by Kerr, Wyllie and Currie in defining apoptosis by EM that triggered the ensuing monumental discoveries toward identifying the genes that controlled cell death (Diamantis et  al. 2008). One of those genes, Bcl-2, was discovered in B-cell follicular lymphomas to be excessively transcribed due to a chromosome translocation that placed the gene under the control of the immunoglobin heavy chain promoter and enhancer (Bakhshi et al. 1985; Tsujimoto et al. 1985; Cleary et al. 1986). Overexpression of Bcl-2 did not promote cell proliferation in a manner similar to other oncogenes, but rather inhibited cell death (Vaux et al. 1988). There are two pathways that mediate apoptosis: the extrinsic pathway mediated by FAS and TNFR1 and the intrinsic or mitochondrial pathway that is mediated by the Bcl-2 family proteins. This family that consists of a variety of pro- (Bax and Bak) and anti- (Bcl-2, Bcl-xL and Mcl-1) apoptotic proteins containing 2 - 4 Bcl-2 homology (BH) domains, are primarily alpha-helical in structure and typically contain a c-terminal membrane anchor (Youle and Strasser 2008). The anti-apoptotic

4  Mitochondrial Dynamics and Apoptosis

111

members normally act by restraining the pro-apoptotic members, until a signal releases the restraint thereby allowing activation. When Bax and Bak become activated, they oligomerize on the mitochondria and induce mitochondrial outer membrane permeabilization (MOMP), which in turn releases cytochrome c and initiates the caspase cascade to trigger the down stream effects of apoptosis. The so-called BH3 only proteins (Bid, Bim, Bad, BMF, NOXA, HRK/DP5, PUMA, BIK and MULE) contain an essential BH3 domain and promote apoptosis via two possible models: (1) binding to and inhibiting the anti-apoptotic Bcl-2 family members which lifts the inhibition of Bax and Bak (Willis et al. 2007) or (2) direct activation of Bax and Bak (Letai et al. 2002).

4.1.3 Mitochondria Are Dynamic Organelles As central components of the intrinsic apoptosis pathway, mitochondria have long been known to exist in many distinct lengths and shapes, but only in the last two decades have many of the key mediators of mitochondrial morphology been elucidated allowing us to begin to understand how mitochondria fission and fusion events maintain steady state mitochondrial morphology. While a variety of proteins have been shown to affect the rates of fission and fusion, four large GTPase proteins (Mfn1, Mfn2, OPA1 and Drp1) comprise the core machinery. As double membrane organelles, mitochondria must unite two separate and mechanistically distinct membranes to complete an organelle fusion event (Meeusen et al. 2004). Mfn1 and Mfn2 first mediate fusion of the outer mitochondrial membrane (OMM) and then OPA1 mediates fusion of the inner mitochondrial membrane (IMM) (Song et  al. 2009). Drp1 on the other hand mediates mitochondrial fission (Smirnova et al. 2001).

4.1.4 Mitochondrial Dynamics Are Linked to Diverse Processes Different cell types and processes display distinct mitochondrial morphologies. For example, during developmental differentiation mitochondrial elongation is required for myogenesis (De Palma et al. 2010), while during adipocyte differentiation mitochondria change from elongated to fragmented (Kita et  al. 2009). Indicative of their importance in metabolism, mitochondria are sensitive to treatment with chemical inhibitors of electron transport, the ATP synthase complex or mtPTP, which all result in reversible fragmentation of mitochondria (De Vos et al. 2005). Recently, an RNAi screen in C. elegans demonstrated how intricately mitochondrial dynamics are linked to a variety of functions in the cell. This study showed that knockdown of 80% of the 719 predicted mitochondrial proteins caused defects in mitochondria morphology, thus indicating that proteins with diverse functions such as metabolism, oxidative phosphorylation, protein folding, protein synthesis and transport can all affect mitochondrial shape and size when

112

M.M. Cleland and R.J. Youle

mutated (Ichishita et  al. 2008). Mitochondria also fragment during autophagy (Twig et al. 2008), the cell cycle (Gorsich and Shaw 2004; Taguchi et al. 2007) and apoptosis (Frank et al. 2001).

4.1.5 Mitochondrial Fragmentation Is an Early Event During Apoptosis For a substantial portion of the last century it was known that mitochondria change morphology during cell death (Glucksmann 1951), with more recent reports describing the changes as smaller size and proliferation in number of mitochondria (Mancini et al. 1997) or a transition to a condensed conformation (Desagher and Martinou 2000). However, work reported in 2001 definitively demonstrated that mitochondria undergo fragmentation during cell death, transitioning from a spaghetti-like network to distinct round fragmented mitochondria (Frank et al. 2001). Treatment with an apoptotic stimulus triggers Bax to translocate from evenly distributed in the cytosol to form distinct foci on the mitochondria (Nechushtan et al. 2001). These Bax foci, which colocalize with Mfn1, Mfn2 and Drp1, eventually become scission sites and result in the apoptotic fragmentation of mitochondria (Karbowski et al. 2002; Cleland et al. 2011). This excessive apoptotic mitochondrial fragmentation can be explained by the following observations (1) Drp1 increases its localization to mitochondria thereby promoting fragmentation (Frank et  al. 2001) and (2) mitochondrial fusion halts, which occurs in a similar time frame as Bax translocation and MOMP (Karbowski et al. 2004). Interestingly, anti-apoptotic Bcl-2 does not block mitochondrial fragmentation during cell death nor does treatment with the caspase inhibitor zVAD, indicating mitochondrial fragmentation is an early event in apoptosis (Sugioka et al. 2004).

4.1.6 Mitochondrial Remodeling and Cytochrome C Release During Apoptosis Although cytochrome c was long known as an important component of electron transport and oxidative phosphorylation, in 1996 it was discovered to be involved in the intrinsic pathway of apoptosis (Liu et al. 1996). Following or concurrent to mitochondrial fragmentation (Frank et  al. 2001) cytochrome c is released from mitochondria (Yang et al. 1997; Kluck et al. 1997) where it binds to APAF1 and initiates caspase activation (Li et al. 1997). Several other proteins have also been shown to be released from the mitochondria following MOMP during apoptosis including Smac/DIABLO, adenylate kinase, Omi/HtrA2, DDP, EndoG and cleaved OPA1 (Wang and Youle 2009).

4  Mitochondrial Dynamics and Apoptosis

113

During cell death the cristae of the mitochondria become highly disorganized. Cytochrome c normally resides within cristae folds. Although it was initially thought that cytochrome c becomes more accessible to release subsequent to MOMP though widening of cristae junctions (Scorrano et al. 2002; Ott et al. 2002) subsequent work correlating light microscopy with EM microscopy found that cristae remodeling is not required for efficient cytochrome c release and that swelling of the mitochondria occurs late in apoptosis after cytochrome c release and membrane potential loss (Sun et al. 2007). It was also demonstrated that MOMP and cristae junction opening can be pharmacologically separated; blocking MOMP did not block OPA1 processing (see Sect. 4.3.4) or cristae junction opening, but did block cytochrome c release and Bak oligomerization (Yamaguchi et al. 2008). Thus, OPA1 processing seems important for MOMP independent of cristae remodeling. With the dramatic changes to the mitochondria that occur during apoptosis and the key apoptotic transduction components that are unleashed from the mitochondria, regulation of these changes is imperative to maintaining the health of the cell or to instigate its destruction in a timely manner. The mitochondrial fusion/fission machinery and their intersection with the intrinsic mitochondrial apoptotic pathway provide a key means for regulation. The remainder of this chapter will define the core mitochondrial dynamic machinery, describe how they function in healthy cells and how each of these proteins can be modulated to exacerbate or prevent apoptosis.

4.2 Drp1 4.2.1 Drp1 Mediates Mitochondrial Fission Dynamin-related protein 1 (Drp1), a large GTPase located both in the cytosol and on the mitochondria, mediates fission. Initially, it was proposed that Drp1 had an impact on mitochondrial distribution as a dominant negative GTPase mutant (Drp1K38A) induced aggregation of the mitochondria (Smirnova et  al. 1998). However, experiments using nocodazole to disrupt microtubules demonstrated that the aggregated mitochondria actually consisted of a highly interconnected network of mitochondria thus demonstrating Drp1 is required for mitochondrial fission (Smirnova et al. 2001). Furthermore, Drp1 RNAi leads to dramatic interconnected nets of mitochondria (Lee et al. 2004). Drp1 homologs also mediate mitochondria fission in C. elegans (Drp1) (Labrousse et al. 1999), Arabidopsis (ADL2b) (Arimura and Tsutsumi 2002), Drosophila (DRP) (Rikhy et al. 2007) and S. cerevisiae (Dnm1) (Otsuga et al. 1998). Interestingly, Drp1 not only mediates normal mitochondrial fission, but it also mediates peroxisome fission (Koch et al. 2003) and has an important role in promoting mitochondrial fragmentation during apoptosis (Frank et al. 2001) (Fig. 4.1). The role of Drp1 in mitochondrial fragmentation and programmed cell death is conserved in C. elegans (Jagasia et al. 2005; Breckenridge et al. 2008), Drosophila (Goyal et al. 2007; Abdelwahid et al. 2007) and also in models of S. cerevisiae death (Fannjiang et al. 2004).

114

M.M. Cleland and R.J. Youle

Fig.  4.1  Drp1 translocation to mitochondria initiates mitochondrial fragmentation. To initiate mitochondrial fragmentation, both in healthy and apoptotic conditions, Drp1 translocates from the cytosol to mitochondria where it binds to Mff and forms large oligomeric structures on the mitochondria, which ultimately instigate fission via GTP hydrolysis. This translocation is sensitive to phosphorylation, mdivi-1, SUMOylation or S-nitrosylation. In healthy cells, mitochondrial fragmentation is reversible and is not coupled to Bax/Bak activation. In apoptotic cells, however, there is excess mitochondrial fission and the large oligomeric Drp1 structures colocalize with oligomeric Bax/Bak, which then initiate MOMP and the downstream effects of apoptosis

4.2.2 Cellular Effects of Drp1 Down-Regulation, Knock-Out and Chemical Inhibition The excess fusion of mitochondria resulting from down regulation of Drp1 can lead to loss of mtDNA, decreased respiration, increase in cellular ROS and diminished ATP levels, ultimately halting cell proliferation (Parone et al. 2008). Low levels of

4  Mitochondrial Dynamics and Apoptosis

115

Drp1 in human cells heteroplasmic for pathological A3243G mtDNA mutation resulted in an increase in mutant mtDNA relative to WT (Malena et  al. 2009). A lethal patient mutation (A395D) caused microcephaly, abnormal brain development, optic atrophy and lactic acidemia (Waterham et al. 2007), likely due to the impaired oligomerization of Drp1 (Chang et  al. 2010). Recently, two research groups demonstrated that the complete deletion of Drp1 in mice causes embryonic lethality, while brain tissue specific deletion leads to cerebellar and forebrain developmental defects and ultimate death due to brain hypoplasia with apoptosis (Ishihara et al. 2009; Wakabayashi et al. 2009). Cells from the knock-out mice had normal ATP levels despite extensively fused mitochondria and peroxisomes (Wakabayashi et al. 2009). The down regulation of Drp1 or expression of the dominant negative GTPase domain mutant, Drp1K38A, both of which inhibit mitochondrial fission, lead to a substantial protection against apoptotic mitochondrial fragmentation and the ensuing cytochrome c release (Frank et al. 2001; Breckenridge et al. 2003; Lee et al. 2004) (Fig. 4.2). In spite of this inhibition of cytochrome c release, Drp1RNAi or Drp1K38A do not block Bax translocation (Karbowski et al. 2002; Lee et al. 2004) nor do they alter Smac/DIABLO, Omi/HtrA2, adenylate kinase 2 or deafness dystonia

Fig.  4.2  Modulation of mitochondrial dynamics alters apoptosis. Enhancing fragmentation augments apoptosis while promoting fusion impedes apoptosis. The central image is a healthy HeLa cell; to the left are the fragmented Mfn2 KO MEF and HeLa cell overexpressing Bax; to the right are the elongated Drp1 RNAi HeLa cell and an Mfn2 KO MEF treated with low level of Actinomycin D thus demonstrating SIMH (Image contributed by Dr. Jean Claude Martinuo)

116

M.M. Cleland and R.J. Youle

peptide/TIMM8a release from the mitochondria thus demonstrating that blocking fission slows down, but does not completely inhibit protein release from the mitochondria followed by cell death (Parone et al. 2006; Estaquier and Arnoult 2007). A recent report has helped shed light on the interesting dilemma of normal Smac, Omi, etc. release but impaired cytochrome c release in Drp1 RNAi cells. Martinou and colleagues demonstrate that Drp1, in a GTPase independent manner, stimulates Bax oligomerization, likely via the promotion of cardiolipin containing hemi-membrane fission intermediates (Montessuit et al. 2010). Although Bax can translocate to mitochondria independently of Drp1, it appears that this Bax is not as efficient at releasing cytochrome c in the absence of Drp1 induced hemi-fission intermediates. A chemical screen for inhibitors of mitochondrial fission using yeast identified mdivi-1 (mitochondrial division inhibitor-1), which was shown to attenuate mitochondrial fission in both mammals and yeast by inhibiting Drp1 and Dnm1 GTPase activity, respectively (Cassidy-Stone et al. 2008). Mdivi-1 also slows down apoptosis by inhibiting MOMP, preventing Drp1 accumulation on the mitochondria and ultimately blocking mitochondrial fragmentation (Fig.  4.1). Furthermore, mdivi-1 was able to block cytochrome c release in vitro (Cassidy-Stone et al. 2008) and in vivo therapeutic applications of this compound are beginning to unfold (Lackner and Nunnari 2010). For example, blocking mitochondrial fission, through expression of Drp1K38A or treatment with mdivi-1, was able to confer protection against cell death that occurs from mPTP opening during ischemia/reperfusion injury (Ong et al. 2010).

4.2.3 Drp1 Localization In healthy cells Drp1 has a predominantly cytosolic distribution with a fraction localized to the mitochondria in distinct foci that often mark future mitochondrial fission events (Pitts et al. 1999; Smirnova et al. 2001). These foci often align with microtubules (Varadi et  al. 2004). Drp1 homologs in Arabidopsis (Arimura and Tsutsumi 2002), S. cerevisiae (Otsuga et al. 1998; Bleazard et al. 1999) and C. elegans (Labrousse et al. 1999) are also found in foci along mitochondria, specifically at mitochondrial tips and constriction points. Purified human Drp1 self assembles in ring-like structures in solution (Smirnova et al. 2001) and yeast Dnm1 forms spirals in vitro as well as on mitochondria and liposomes (Ingerman et al. 2005). Therefore, the current model is that Drp1 foci on mitochondria represent spirals then undergo conformational changes to constrict the membrane and induce fission (Fig. 4.1). In yeast, Fis1 and/or the adaptor proteins Mdv1 and Caf4 are required for Dnm1 mitochondrial localization and thus are important regulators of mitochondrial fission (Tieu and Nunnari 2000; Mozdy et  al. 2000; Tieu et  al. 2002; Griffin et al. 2005; Schauss et al. 2006). However, in mammals Fis1 is not required for Drp1 localization (Lee et al. 2004) nor has an Mdv1 or Caf4 homolog protein

4  Mitochondrial Dynamics and Apoptosis

117

been found. This would indicate that yeast and mammalian mitochondrial fission machinery may operate differently and the mammalian Drp1 mitochondrial receptor has yet to be elucidated. In 2008 a new player in the mammalian fission machinery, Mff (Mitochondrial Fission Factor) was identified by an RNAi screen in Drosophila cells. Downregulation of Mff drastically inhibits mitochondrial fission resulting in completely fused mitochondria that are quite resistant to uncoupler induced fragmentation (Gandre-Babbe and van der Bliek 2008). Recently, Mihara and colleagues demonstrated that in the absence of Mff, Drp1 does not localize to mitochondria, while overexpression of Mff leads to enhanced Drp1 mitochondrial localization and enhanced fission, thus indicating that Mff is a key component in mitochondrial recruitment of Drp1 (Otera et  al. 2010) (Fig.  4.1). Furthermore, these authors demonstrated through various RNAi and genetic ablation strategies that Fis1 is dispensable for mitochondrial fission; showing that Fis1 RNAi did not produce completely interconnected networks as Drp1 and Mff RNAi did and that Fis1 KO HCT116 cells had similar mitochondrial morphology to their WT counterparts (Otera et al. 2010). Therefore, in mammalian cells Mff, rather than Fis1, has a prominent role in Drp1 recruitment and induction of mitochondrial fission. It is possible that Fis1’s role in mitochondrial fission is masked by an unknown functionally redundant protein and/or that Fis1 mediated fission is downstream of Mff recruitment of Drp1. During apoptosis Drp1 (Frank et  al. 2001) and Bax translocate (Wolter et  al. 1997) from the cytosol to the mitochondria, where they ultimately coalesce at future mitochondrial scission sites (Karbowski et al. 2002) and likely cooperate to induce cytochrome c release (Montessuit et al. 2010) (Fig. 4.1). Drp1 foci, which become stably associated with mitochondria after Bax translocation but before MOMP (Wasiak et al. 2007), form prior to fragmentation and membrane potential loss (Germain et  al. 2005). Interestingly, expression of anti-apoptotic Bcl-2 does not block Drp1 recruitment, but prevents cell death indicating that Drp1 activation occurs independently of Bcl-2 (Sugioka et al. 2004). On the other hand Bax translocation is upstream of Drp1 translocation (Frank et al. 2001). Blocking the ability of Drp1 to localize to mitochondria by downregulating Mff inhibits apoptotic mitochondrial fragmentation and cytochrome c release (Otera et al. 2010). Therefore, inhibiting the translocation of Drp1 to mitochondria during cell death offers an efficient means of regulating cell death.

4.2.4 Drp1 Post-Translational Modifications Drp1 has been shown to be regulated by multiple pathways of post-translational modification including phosphorylation (Taguchi et al. 2007; Chang and Blackstone 2007; Cribbs and Strack 2007), SUMOylation (Zunino et  al. 2007; Wasiak et  al. 2007), S-nitrosylation (Cho et al. 2009) and ubiquitination (Yonashiro et al. 2006; Nakamura et al. 2006; Karbowski et al. 2007). Regulation of the mitochondrial fission activity of Drp1 typically involves either stimulating or inhibiting the GTPase

118

M.M. Cleland and R.J. Youle

activity. While the specific details of each of the mediators of these post-translational modifications are still being debated, it is clear that Drp1 is a highly regulated protein due to its important function in mediating fission of mitochondria. These modifications are particularly important during apoptosis. Dephosphorylation activates Drp1 and triggers mitochondrial fragmentation as evidenced by the phosphomimetic mutants displaying reduced activity in fission, elongated mitochondria and resistance to apoptosis (Cribbs and Strack 2007) whereas a phosphorylation deficient Drp1 mutant appears constitutively active with consequent mitochondrial fragmentation and enhanced apoptosis (Cribbs and Strack 2007). During apoptosis, Drp1 interacts with the E2 enzyme Ubc9 and SUMO-1 (Harder et al. 2004) and becomes SUMOylated in a Bax/Bak-dependent manner (Wasiak et al. 2007). S-nitrosylation regulates Drp1 dimerization, GTPase activity and mitochondrial fragmentation and mutants devoid of S-nitrosylation are inactive and therefore do not promote cell death (Cho et al. 2009). Therefore, phosphorylation prevents activation of Drp1, while S-nitrosylation and SUMOylation increase Drp1 activity (Fig. 4.1).

4.3 OPA1 Whereas mitochondrial fission requires one large GTPase, Drp1, mitochondrial fusion requires three large GTPases, Mfn1, Mfn2 and OPA1 to mediate OMM and IMM fusion. The first reports of the large mitochondrial GTPase OPA1 regarded the yeast homolog, Mgm1. Initially, Mgm1 was thought to be important for mitochondrial genome maintenance (Jones and Fangman 1992; Guan et  al. 1993; Shepard and Yaffe 1999), however, it was later determined that Mgm1 was primarily necessary for mitochondrial fusion and its genome maintenance role was via this process (Wong et al. 2000, 2003).

4.3.1 OPA1 Mutations Result in Human Disease The human optic atrophy 1 (OPA1) was discovered to be mutated in dominant optic atrophy (DOA) (Alexander et  al. 2000; Delettre et  al. 2000), the most prevalent hereditary optic neuropathy (Lenaers et al. 2009) before its mitochondrial roles were established. Although OPA1 is broadly expressed, it is most abundant in the retina (Alexander et al. 2000). OPA1 contains a mitochondrial targeting sequence (MTS), two hydrophobic regions, a GTPase domain, a middle domain and a GED domain (Hoppins et al. 2007). The majority of DOA patient mutations lie within the GTPase domain of OPA1 (Votruba 2004), with additional mutations occurring in the mitochondrial targeting region, middle and GED domain (Ferré et al. 2009).

4  Mitochondrial Dynamics and Apoptosis

119

4.3.2 Cellular Effects of OPA1 Deletion Cells lacking OPA1 have excessively fragmented mitochondria (Cipolat et al. 2004; Griparic et  al. 2004) that have dissipated or heterogeneous membrane potential (Olichon et  al. 2003) as well as deficient oxygen consumption and cell growth (Chen et  al. 2005). These defects are likely due to the fact that knocking down OPA1 also severely impairs the cristae morphology resulting in swollen/altered cristae (Olichon et al. 2003; Griparic et al. 2004). Mutation or deletion of the S. cerevisiae (Mgm1) or C. elegans (Eat-3) homologs of OPA1 lead to impaired mitochondrial inheritance (S. cerevisiae), mitochondrial fragmentation, loss of mtDNA and swollen cristae (Shepard and Yaffe 1999; Amutha et al. 2004; Kanazawa et al. 2008). Interestingly, OMM fusion can still occur in OPA1 null mouse cells (Song et al. 2009) or in yeast harboring mutant Mgm1 (Meeusen et al. 2006), but IMM fusion is defective thus indicating that the mitofusins first mediate OMM fusion followed by OPA1-mediated fusion of the IMM. The first solid link between mitochondrial fusion machinery and apoptosis was the report that loss of OPA1 via siRNA treatment committed cells to apoptosis, which can be prevented by overexpressing anti-apoptotic Bcl-2 (Olichon et  al. 2003). Loss of OPA1 can sensitize or accelerate the rate of apoptosis and even leads to spontaneous cell death from cytochrome c release and severe disruption of the cristae morphology (Lee et  al. 2004; Frezza et  al. 2006; Arnoult et  al. 2005) (Fig. 4.2). Overexpression of WT OPA1 protects cells from apoptosis induced by intrinsic stimuli by maintaining normal cristae morphology, delaying the release of cytochrome c and preventing the loss of membrane potential (Frezza et al. 2006) (Fig.  4.2). Curiously, the C. elegans homolog, Eat-3, mutants do not show an increase in cell death (Kanazawa et al. 2008).

4.3.3 OPA1 Mediates Inner Mitochondrial Membrane Fusion OPA1 is found in the mitochondrial intermembrane space (IMS) as well as bound to the surface of the IMM (Griparic et al. 2004) where it promotes fusion of the IMM. Overexpression of OPA1 can promote elongated mitochondria (Cipolat et al. 2004), or fragmented mitochondria that tend to cluster around the nucleus (Misaka et al. 2002; Griparic et al. 2004). This fragmentation is likely due to an overexpression artifact or the delicate nature of IMM morphology. In in vitro mitochondrial fusion assays, yeast mitochondria require a functional Mgm1 on both mitochondria (Meeusen et al. 2006), consistent with intermolecular complex formation mediating IMM fusion and cristae maintenance (Amutha et al. 2004; Olichon et al. 2003). The addition of GTP to OPA1 promotes in vitro complex assembly, while GDP blocks complex formation (Yamaguchi et  al. 2008). Therefore, OPA1 complex formation is key in maintaining IMM morphology.

120

M.M. Cleland and R.J. Youle

4.3.4 OPA1 Splice Variants and Proteolytic Processing OPA1 has eight splice variants due to alternative splicing of exon 4, 4b and 5b (Delettre et al. 2001) that appear to have differential roles in mitochondrial dynamics and susceptibility to apoptosis (Olichon et  al. 2007). Interestingly, Exon 4, which is conserved through evolution, is involved in the mitochondrial membrane potential maintenance as well as fusion of the mitochondrial network, while the vertebrate specific exon 4b and 5b appear to define OPA1 function in cytochrome c release (Olichon et al. 2007). This could indicate that OPA1 evolved this additional function to become an important component of the intrinsic apoptosis pathway and could also explain why ablation/mutation of the C. elegans homolog Eat-3 does not show an increase in cell death. OPA1 not only has eight splice variants, but also undergoes both constitutive and mitochondrial depolarization induced proteolytic processing. Each splice variant contains a MPP (mitochondrial processing peptidase) cleavage site just after the MTS, in addition to cleavage sites within exons 5 and 5b (Song et al. 2007). Once inserted into the IMM, cleavage by MPP generates the mature long isoform of OPA1 (L-OPA1), while cleavage in the peptide encoded by exons 5 and 5b generate the short isoforms (S-OPA1) (Song et al. 2007). Initial reports suggested that only the L-OPA1 was fusion competent (Ishihara et  al. 2006), however other reports indicate that only long isoforms that can be constitutively processed to generate one or more shorter isoforms are able to support fusion activity (Song et al. 2007). The yeast homolog Mgm1, which is also processed by proteases, requires the presence of both long and short isoforms for maintenance of mitochondrial morphology and mtDNA (Herlan et al. 2003). Until recently, it was unclear what role the constitutively cleaved form of OPA1 played. Purified S-OPA1 (Ban et al. 2010; DeVay et al. 2009) and yeast homolog S-Mgm1 (Meglei and McQuibban 2009) were shown to have a low basal GTP hydrolysis rate that was stimulated upon binding negative phospholipids like cardiolipin; specifically, binding to lipids with a similar composition to the IMM resulted in a 50-fold increase in GTPase activity of Mgm1 (Rujiviphat et al. 2009). Upon association with lipids (specifically cardiolipin) S-OPA1 assembles into higher order oligomers and tubulates membranes (Ban et al. 2010). Since L-OPA1 and S-OPA1 both appear to be required for fusion (Song et  al. 2007), one could speculate that L-OPA1/S-OPA1 complex that forms during fusion would expose S-OPA1 to the IMM and therefore increase the GTP hydrolysis rate and complete membrane fusion. While it is clear that low levels of constitutive processing of OPA1 are important for maintenance of mitochondrial fusion, excessive processing of OPA1 from mitochondrial depolarization-induced proteolytic processing due to chemical depolarization or apoptosis induced membrane potential loss clearly results in mitochondrial fragmentation (Fig. 4.3). Depletion of mitochondrial membrane potential by CCCP treatment results in high levels of rapid proteolytic processing of OPA1, occurring within 10–15  min. This fragmentation can be reversed by removing CCCP but

4  Mitochondrial Dynamics and Apoptosis

121

Fig.  4.3  OPA1 induced proteolysis is mediated by OMA1. OPA1 proteolytic processing by OMA1 during uncoupling of the IMM results in fragmented mitochondria and an accumulation of S-OPA1 in the IMS. During apoptosis, however, OPA1 proteolytic processing by OMA1 is coupled with Bax/Bak promoted MOMP thereby resulting in an increase in S-OPA1 concurrent with release of S-OPA1 and cytochrome c from the IMS

requires de  novo protein synthesis (Duvezin-Caubet et  al. 2006; Ishihara et  al. 2006; Griparic et al. 2007). Interestingly, OPA1 processing is enhanced in a variety of cell types modeling disease conditions that display heightened mitochondrial fragmentation (Duvezin-Caubet et al. 2006). Recent analysis of the various types of fusion events that occur revealed that OPA1 is important for complete mitochondrial fusion, as decreasing the amount of OPA1 results in lack of complete fusion, while transient fusions events could still occur (Liu et al. 2009). Non-fusing mitochondria, which tend to undergo autophagy, have reduced levels of OPA1 as well as reduced membrane potential (Twig et  al. 2008). Ultimately, OPA1 proteolytic cleavage allows the cell to select healthy from unhealthy mitochondria, subjecting the unhealthy mitochondria to elimination by autophagy.

122

M.M. Cleland and R.J. Youle

During apoptosis the long isoforms of OPA1 disappear due to excessive proteolytic processing (Ishihara et al. 2006) and the resulting short isoforms are released into the cytosol along with cytochrome c and Smac/DIABLO (Arnoult et  al. 2005) (Fig. 4.3), a process which, in neurons, occurs simultaneously with mitochondrial fragmentation and cytochrome c release (Loucks et al. 2009). Although release of the short Opa1 isoforms occurs simultaneous with MOMP, BH3 only proteins Bid and Bim can induce OPA1 complex disassembly even when MOMP is blocked by Drp1 RNAi, thus indicating that OPA1 processing precedes MOMP (Yamaguchi et al. 2008). Curiously OPA1 is processed to a greater extent in Drp1 RNAi cells (Estaquier and Arnoult 2007). Expression of a mutant of OPA1 (Q297V) that mimics its GTP bound form doesn’t undergo proteolytic processing, blocks cytochrome c, Omi/Htra2 and Smac/DIABLO release, but not Bax activation (Yamaguchi et al. 2008). Therefore, interfering with apoptosis-induced proteolytic cleavage of OPA1 offers an appealing strategy to prevent apoptosis. Early efforts to identify the protease mediating OPA1 cleavage following membrane potential loss focused on the rhomboid protease PARL (Presenilin-associated rhomboid-like) due to the clear indication that yeast Mgm1 was cleaved by the yeast homolog of PARL, Pcp1 (Herlan et al. 2003; Sesaki et al. 2003; McQuibban et al. 2003). Although initial reports indicated that PARL−/− cells displayed reduced levels of soluble IMS short OPA1, but have unchanged levels of long OPA1 (Cipolat et al. 2006), subsequent studies revealed that OPA1 cleavage pattern, both constitutive and uncoupler-induced, is normal in PARL−/− and PARL siRNA cells (Guillery et al. 2007; Griparic et al. 2007). Moreover, OPA1 expressed in yeast cells is not cleaved by Pcp1 (Duvezin-Caubet et al. 2007). Initial experiments demonstrated that overexpression of Paraplegin fragmented the mitochondria and decreased L-OPA1 while Paraplegin RNAi modestly compromised OPA1 processing (Ishihara et al. 2006), however it was later demonstrated that OPA1 processing, both constitutive and excessive CCCP-induced, occurs normally in Paraplegin deficient Spg7−/− cells (Duvezin-Caubet et al. 2007). Another candidate for CCCP-induced OPA1 cleavage was the i-AAA protease Yme1. Knockdown of Yme1 was initially reported to have no affect on OPA1 processing (Ishihara et al. 2006), however it was later demonstrated to prevent the constitutive cleavage of a subset of OPA1 splice variants (Griparic et al. 2007). Furthermore, Yme1 either has a minor role or does not mediate the OPA1 cleavage induced by CCCP or apoptosis (Griparic et  al. 2007; Guillery et  al. 2007). Thus the current consensus is that Yme1 mediates constitutive processing of a subset of OPA1 splice variants, while PARL and Paraplegin have minor if any roles in OPA1 processing. Recent work from two groups identified that OPA1 inducible cleavage is mediated by OMA1, which is a mitochondrial zinc metalloprotease with multiple membrane spanning segments and a zinc-binding motif (Head et al. 2009; Ehses et al. 2009) (Fig. 4.3). OMA1 is also constitutively cleaved, but is stabilized upon CCCP treatment allowing the full length OMA1 to initiate rapid OPA1 cleavage (Head et  al. 2009). Interestingly, subunits of m-AAA protease AFG3L1/2 may control OMA1 cleavage since deleting AFG3L1/2 results in excessive processing of OPA1

4  Mitochondrial Dynamics and Apoptosis

123

by OMA1 (Ehses et al. 2009). Depletion of OMA1 inhibits OPA1 inducible cleavage (Head et al. 2009) and prevents the excessive OPA1 processing in cells devoid of AFG3L1 and AFG3L2 (Ehses et al. 2009). Prohibitin-1 and Prohibitin-2 are also involved in the proteolytic processing of OPA1 by acting as scaffolds on the IMM to assist in defining the spatial compartments for OPA1 processing and membrane fusion (Merkwirth et al. 2008). Due to the potent processing of OPA1 by OMA1, it is plausible that OMA1 may require restraints beyond proteolytic destabilization, such as residing in a different spatial compartment than OPA1 due to partitioning by the Prohibitins (McBride and Soubannier 2010). Although initially it appeared that the metalloprotease-mediated OPA1 processing detailed above did not appear to be required for the apoptotic release of cytochrome c and OPA1 from the IMS (Guillery et al. 2007), recent work suggests that metalloprotease-mediated OPA1 processing may be important given that silencing OMA1, which delays the excessive processing of OPA1, leads to a delay in apoptosis (Head et al. 2009). Interestingly, OPA1 oligomers disappear faster in PARL−/− mitochondria (Frezza et al. 2006), which have faster apoptotic cristae remodeling and cytochrome c release (Cipolat et  al. 2006). Moreover, WT OPA1 protects WT MEFs, but not PARL−/− MEFs from apoptosis and targeting OPA1 to the IMS protects PARL−/− MEFs from apoptosis. This suggests that OPA1 must undergo constitutive proteolytic processing for its anti-apoptotic function and that there is some balance required between its long and short forms to achieve this protection (Cipolat et  al. 2006). Furthermore, this could indicate that perhaps OMA1 is stabilized in PARL−/− mitochondria thus causing excessive processing of OPA1. Ultimately, low level constitutive processing appears to be important for maintaining OPA1’s anti-apoptotic function through maintaining OPA1 oligomers, cristae morphology and promoting IMM fusion, while induced excessive proteolytic processing and release of OPA1 from the mitochondria results in the disruption of cristae morphology and leads to a timely execution of apoptosis.

4.4 Mfn1 and Mfn2 The first member of the mitofusin family was discovered in Drosophila by Hales and Fuller in 1997, when the fuzzy onion gene (fzo; named for the smaller mitochondria that wrap around each other in the onion stage of nebekern formation) was shown to be a mediator of mitochondrial fusion (Hales and Fuller 1997). The importance of the mitofusins in mitochondrial fusion was further confirmed when homologs in S. cerevisiae (Fzo1) (Hermann et al. 1998), mammals (which express two distinct homologs: Mfn1 and Mfn2 (Santel and Fuller 2001)) and C. elegans (Fzo-1) (Ichishita et  al. 2008) were discovered and also shown to mediate mitochondrial fusion. Later studies in Drosophila revealed a second mitofusin gene, Marf, which has more broad tissue expression than fzo (Hwa et al. 2002).

124

M.M. Cleland and R.J. Youle

4.4.1 Mfn2 Mutation Results in CMT2A Mfn1 and Mfn2, in the same large GTPase family as Drp1 and OPA1, are OMM proteins which contain beyond the GTPase domain, two coiled-coil domains and two membrane-spanning motifs (Santel and Fuller 2001), which orient the GTPase domain and coiled-coil domains to face the cytosol (Rojo et al. 2002). Mutations in the GTPase domain, N-terminus and coiled-coil domain in Mfn2 have been found to cause CharcotMarie-Tooth type 2A (CMT2A), a peripheral neuropathy (Züchner et  al. 2004). Curiously, Mfn1, but not Mfn2, can complement the fusion deficient CMT2A disease allele mutants and restore mitochondrial fusion. This revealed the importance of Mfn1 levels in tissues affected by CMT2A disease (Detmer and Chan 2007). Work using transgenic mice expressing these mutants has recently begun with a transgenic mouse expressing R94Q Mfn2, a prominent CMT2A disease allele, in the neuron specific enolase locus. This mouse had locomotion and gait defects, which resembled CMT2A disease phenotypes and had increased ­numbers of mitochondria in the distal end of the sciatic nerve, reminiscent of a mitochondrial fusion defect (Cartoni et al. 2010). Other neuropathies, such as hereditary motor and sensory neuropathy type VI are also associated with Mfn2 mutations (Züchner et al. 2006). Ultimately, it is unclear why OPA1 mutations result in DOA while Mfn2 mutations result in CMT2A, since both OPA1 and Mfn2 are important for mitochondrial fusion. Perhaps the resolution to this lies in ­functions besides mitochondrial fusion such as cristae maintenance (OPA1), apoptosis regulation, ER-tethering (Mfn2) or in differential tissue expression.

4.4.2 Mitofusin Deletion Results in Fragmented Mitochondria Deletion of FZO1 in yeast leads to loss of mtDNA, mitochondrial fragmentation and a block in fusion following yeast mating (Rapaport et al. 1998; Hermann et al. 1998). In mammals, removal of either mitofusin is lethal with Mfn1 KO and Mfn2 KO mice both dying in mid-gestation (Chen et al. 2003). Mitochondria in MEFs derived from Mfn1 KO and Mfn2 KO mice are more fragmented than mitochondria in WT MEFs and display much slower rates of fusion (Chen et  al. 2005). Intriguingly, Mfn1 had a more profound effect on fusion than Mfn2 as evidenced by Mfn1’s complete restoration of fusion in both Mfn1 and Mfn2 KO MEFs, while Mfn2 completely restored fusion in Mfn2 KO MEFs but only partially restored fusion in Mfn1KO MEFs (Chen et  al. 2003). Depletion of both Mfn1 and Mfn2 leads to enhanced mitochondrial fragmentation and a complete block in fusion (Eura et  al. 2003; Chen et  al. 2005). Mfn1/2 DKO MEFs grow slowly and their mitochondria display heterogeneous membrane potential and decreased cellular respiration (Chen et al. 2005). Furthermore, conditional deletion of Mfn1 or Mfn2 revealed their importance in Purkinje cell development (Chen et al. 2007) and skeletal muscle morphology (Chen et al. 2010). Mfn2 is expressed more highly in heart and skeletal muscle than in brain, kidney and liver and is induced during myogenesis (Bach et  al. 2003). Mfn1 and Mfn2

4  Mitochondrial Dynamics and Apoptosis

125

levels are increased 24  h post-exercise in a PGC-1alpha driven process (Cartoni et al. 2005). Mfn2 knockdown fragments the mitochondria of myotubes and impairs glucose oxidation, membrane potential and oxygen consumption (Bach et al. 2003). Mfn2 levels are lower in the skeletal muscle from obese Zucker rats (Bach et al. 2003) and in skeletal muscles from Early-Onset Type 2 Diabetic subjects, where during chronic exercise the Mfn2 level in skeletal muscles increases in control subjects, but fails to increase in diabetic subjects (Hernández-Alvarez et al. 2010). These studies taken together highlight the importance of the mitofusins in maintaining mitochondrial health in a variety of cell types. Although, how mitochondrial fusion maintains mitochondrial fidelity is not clear, one possibility is that repeated rounds of fission and fusion allow segregation of damaged materials and autophagic removal of impaired mitochondria (Twig et al. 2008).

4.4.3 Fusion Mechanism: GTPase Function and Oligomeric Interactions Mfn1 and Mfn2 interact and form both cis (same mitochondria) and trans (adjacent mitochondria) hetero and homotypic complexes (Chen et al. 2003; Eura et al. 2003) which are required on adjacent mitochondria for fusion to occur (Koshiba et  al. 2004). Intriguingly, in vitro mitochondrial fusion assays demonstrated that Mfn1/ Mfn2 heterotypic trans complexes are more efficient at inducing fusion than homotypic complexes (Hoppins et  al. 2011). Experiments in yeast revealed that trans interactions require one interacting partner to have functional coiled-coil domain while the other must have a functional GTPase domain thereby indicating that the GTP domain interactions with coiled-coil domains are essential for fusion (Griffin and Chan 2006). Mutations within the GTPase domain hinder the mitofusin mitochondrial fusion activity (Hermann et al. 1998; Santel and Fuller 2001). Mfn2K109A or Mfn1K88T, which are homologous mutations within the GTPase domains of these proteins, have lost the ability to restore fusion in Mfn1 and Mfn2 KO cells (Chen et al. 2003). Mfn2K109T, which acts as a dominant negative inhibitor of fusion, induces significant mitochondrial fragmentation and clumping in WT cells (Santel and Fuller 2001). This mutant appears to malfunction at an intermediate stage in mitochondrial fusion since it accumulates at contact points between mitochondria, or on the tips of mitochondria (Eura et  al. 2003). Mfn1K88T, on the other hand, is not as effective as Mfn2K109T at blocking fusion in WT cells (Eura et al. 2003). Thus distinct differences are apparent between Mfn1 and Mfn2 in GTPase domain characteristics and adjacent mitochondria tethering efficiencies (Ishihara et al. 2004). Endogenous Mfn2 forms foci on tips and constriction points of mitochondria, and overexpressing Mfn2-YFP results in a more distinct punctate localization (Cleland et al. 2011). Interestingly, this punctate localization of Mfn2 appears to be linked to the nucleotide bound state and GTPase activity since the dominant-negative Mfn2K109T forms larger and more prominent foci, while the dominant active

126

M.M. Cleland and R.J. Youle

Mfn2G12V does not form such distinct puncta of WT Mfn2 (Neuspiel et al. 2005; Karbowski et al. 2006). Furthermore, expression of Mfn1 and Mfn2 mutants that lack the c-terminal coiled-coil domain, and thus are devoid of tethering and fusion functions, are not able to form foci (Karbowski et al. 2006). The mitofusins are also regulated by ubiquitination. For example, cell cycle arrest in yeast leads to mitochondrial fragmentation and proteasome-mediated degradation of Fzo1 in an Mdm30 (an F-Box protein that is found in subunits of Skp1Cdc53-F-box protein ubiquitin ligases) independent manner (Neutzner and Youle 2005; Escobar-Henriques et al. 2006). On the other hand, at steady state Mdm30 mediates Fzo1 ubiquitination and degradation (Cohen et al. 2008). Interestingly, a CMT2A mutation (human I213T; yeast V327T) engineered into Fzo1 had diminished GTP hydrolysis, membrane fusion activity and reduced Mdm30-mediated ubiquitination and degradation thus linking turnover of Fzo1 with fusion activity (Amiott et al. 2009). Recently, Mfn1 and Mfn2 as well as Drosophila Marf were shown to be ubiquitinated and degraded by the proteasome during mitochondrial membrane potential loss in a PINK1, Parkin and p97 dependent manner (Ziviani et  al. 2010; Poole et  al. 2010; Gegg et  al. 2010; Tanaka et  al. 2010). Thus outer mitochondrial membrane associated degradation (OMMAD) offers an efficient means to modulate mitochondrial fusion by regulating the level of mitofusins. However, it remains to be determined if the mitofusins must be ubiquitinated and degraded during cycles of mitochondrial fusion.

4.4.4 Excess Mitochondrial Fusion Delays Cell Death Consistent with OPA1-mediated fusion, overexpression of rat Mfn1/2 diminishes or delays mitochondrial fragmentation, Bax activation, cytochrome c release and cell death induced by etoposide treatment, while silencing Mfn1/2 causes a mild increase in apoptosis (Sugioka et al. 2004) (Fig. 4.2). Furthermore, expression of the dominant active Mfn2G12V protects against STS-induced cell death better than WT Mfn2 in Cos-7 cells (Neuspiel et al. 2005). In cerebellar neurons, Mfn2 knockdown induces cell death, while Mfn2 overexpression induces mitochondrial fusion and prevents apoptosis induced by DNA damage, oxidative stress or potassium deprivation (Jahani-Asl et  al. 2007). Similar to Cos-7 cells, Mfn2G12V protects neurons from apoptosis better than WT Mfn2, specifically attenuating cytochrome c release (Jahani-Asl et al. 2007). Curiously, oxidative stress induces Mfn2 expression and apoptosis in rat neonatal cardiomyocytes thus indicating that under these conditions in cardiomyocytes Mfn2 is no longer protective against apoptosis (Shen et al. 2007). Conversely, in the HL-1 cardiac cell line mitochondria fragment during ischemia/reperfusion injury and overexpression of Mfn1 and Mfn2 blocks cell death that occurs from this insult (Ong et al. 2010). Prior to the fragmentation induced by apoptosis, mitochondria become excessively long and interconnected during a process termed stress-induced mitochondrial hyperfusion (SIMH). This can occur during certain stresses such as UV

4  Mitochondrial Dynamics and Apoptosis

127

irradiation or actinomycin D treatment and requires L-Opa, Mfn1 and SLP-2 (Stomatin-like protein 2, (Hájek et al. 2007)), but not Mfn2, Bax/Bak or the prohibitins. SIMH correlates with an increase in ATP production by the mitochondria and ultimately protects against apoptosis (Tondera et al. 2009) (Fig. 4.2). Certain cancer cells (LS174 colon carcinoma) evade apoptosis during hypoxia conditions through the induction of mitochondrial fusion during which Mfn1 is slightly upregulated (Chiche et al. 2010). Viruses also modulate mitochondrial dynamics to evade apoptosis. Interestingly, the cytomegalovirus protein vMIA (viral mitochondrial inhibitor of apoptosis) induces substantial mitochondrial fragmentation without the induction of apoptosis (McCormick et  al. 2003; Karbowski et  al. 2006). This fragmentation without ensuing apoptosis is a result of the vMIA-mediated inactivation and sequestration of Bax (Arnoult et al. 2004; Poncet et al. 2004) and Bak (Karbowski et al. 2006) on the mitochondria which prevents them from oligomerizing and releasing cytochrome c. This fragmentation might also be a strategy of the virus to evade the host antiviral response. Recent reports show that activation of retinoic acid-inducible gene 1-like receptor, which responds to viral RNA, inducing mitochondrial elongation through the activation of MAVS (mitochondrial antiviral signaling), which then prompts production of type 1 interferons and proinflammatory cytokines to combat the viral invasion (Castanier et al. 2010). MAVS downstream signaling is increased upon excessive fusion (Drp1 shRNA) and decreased upon excess fission due to OPA1 shRNA, Mfn1 shRNA or expression of vMIA (Castanier et al. 2010), perhaps indicating that inducing fission may be an efficient way to evade the viral immune response, while inducing fusion heightens the antiviral response.

4.5 Mitochondrial Dynamics and the Bcl-2 Family An interesting new connection between the machinery controlling mitochondrial morphology and that controlling apoptosis has emerged in the last 5 years. In addition to their role in regulating apoptosis, members of the Bcl-2 family in C. elegans, Drosophila and mammals regulate mitochondrial fission and fusion even in healthy cells. In HeLa cells, overexpressed CED-9 (the C. elegans Bcl-2 family member) or Bcl-xL promotes mitochondrial fusion and clumping which is likely mediated by their interaction with Mfn2 (Delivani et  al. 2006). In neurons, overexpressed Bcl-xL binds to and stimulates the GTPase activity of Drp1 resulting in an increased density of synapses and synaptic vesicles as well as an increased number of mitochondria localized in synapses likely due to enhanced mitochondrial fission (Li et al. 2008). Similarly, anti-apoptotic Bcl-w promotes mitochondrial fragmentation in Purkinje cell dendrites (Liu and Shio 2008). Through a careful examination of fission and fusion events, Berman et  al. found that Bcl-xL overexpression in neurons increases both the fusion and fission rates in addition to increasing mitochondrial mass (Berman et  al. 2009). In C. elegans increased CED-9 expression leads to longer mitochondria in muscle cells in an FZO-1 and EAT-3 (OPA1)

128

M.M. Cleland and R.J. Youle

dependent manner, though CED-9 mutants display normal mitochondrial morphology (Tan et  al. 2008; Rolland et  al. 2009; Breckenridge et  al. 2009). CED-9 and FZO-1 have been shown to interact, likely via the coiled-coil domain located between the GTPase domain and the TM domain of FZO-1 (Rolland et al. 2009). Although the downstream mechanisms of this interaction have not been fully explored, it is likely to be important given the role of these two proteins in mitochondrial morphology maintenance. Recently it was shown that the dynamics of mitochondria during germline cell death in Drosophila are dependent on the Bcl-2 family proteins Buffy and Debcl (Tanner et al. 2010) indicating that the role of the Bcl-2 family in mitochondrial morphology is distinct from their roles in apoptosis. Bax and Bak also seem to be important for mitochondrial fusion as elimination or inhibition of Bax and Bak through genetic manipulation, RNAi or inactivation by a viral protein results in fragmented mitochondria that show a distinct defect in mitochondrial fusion (Karbowski et al. 2006; Cleland et al. 2011). Using a cell free mitochondrial fusion assay, it was further demonstrated that recombinant Bax stimulates mitochondrial fusion in an Mfn2 dependent manner more than 50% (Hoppins et al. 2011). This is in agreement with earlier studies demonstrating that the defect in fusion from the absence of Bax/Bak was attributed to the loss of focal localization of Mfn2 and thus loss of fusogenic function of Mfn2 (Karbowski et al. 2006). Interestingly, not only does Bax promote the foci formation of Mfn2 in healthy cells, it also forms apoptotic mitochondrial localized foci that colocalize with Mfn1, Mfn2 and Drp1 (Karbowski et al. 2002; Cleland et al. 2011). The exact role of these foci remains to be determined. There have been several reports of the mitofusins interacting with Bcl-2 family proteins (Karbowski et al. 2006; Delivani et al. 2006; Brooks et al. 2007; Rolland et al. 2009; Cleland et al. 2011). One study focused on whether Bax and Bak interacted with Mfn1 and Mfn2 during apoptosis. Accordingly, Brooks and colleagues found that Bax interacts with Mfn2 similarly in the absence and presence of apoptosis, but interacts more with Mfn1 during apoptosis. Bak, on the other hand, switches from interacting with Mfn2 to Mfn1 during apoptosis. The inactive BH3 domain mutant, Bak L75E, no longer promoted mitochondrial fragmentation during apoptosis nor was it able to dissociate from Mfn2 during apoptosis (Brooks et al. 2007). In accordance with these results, IL-6 treatment, which increases Bcl-2 expression and thereby decreases hyperoxic lung injury and cell death, ultimately prevents the dissociation of Bak from Mfn2 in addition to preventing the interaction between Mfn1 and Bak (Waxman and Kolliputi 2009). Further indicating the delicate and dynamic nature of the mitofusin/Bax interaction, a chimeric protein between Bcl-xL and Bax containing the majority of Bcl-xL with helix 5 of Bax inserted, had substantially increased binding to the Mfn1/2 relative to WT Bax and WT Bcl-xL and induced substantial mitochondrial fragmentation (Cleland et  al. 2011). Interestingly, this chimera also had an enhanced colocalization with the mitofusins and appeared to increase the size of endogenous Mfn2 foci (Cleland et al. 2011). Therefore, altering the efficacy of the mitofusin/Bax interaction offers yet another strategy to alter mitochondrial morphology.

4  Mitochondrial Dynamics and Apoptosis

129

4.6 Conclusion In normal healthy cells the large GTPase machineries of the OMM and IMM membrane, Drp1, Mfn1, Mfn2 and OPA1, maintain mitochondrial morphology. These proteins are regulated primarily through modulation of their GTPase functions by phosphorylation, SUMOylation, S-nitrosylation, ubiquitination and protease cleavage that ultimately affects localization, oligomeric activity and protein level. During cell death, Drp1 translocates from the cytosol to colocalize with Bax, Mfn1 and Mfn2 at future mitochondrial scission sites. Drp1 then forms large oligomeric structures, which likely constrict the mitochondria to ultimately undergo massive fragmentation. While the fission proceeds, mitochondrial fusion is inhibited, OPA1 oligomers begin to disassemble and the cristae morphology becomes swollen and disordered. Close in time with the fragmentation and cristae remodeling, MOMP occurs and various small proteins such as cytochrome c and SMAC are released from the IMS and proceed to activate the downstream effectors of the intrinsic apoptotic pathway. Thus, the processes of mitochondrial fission appear intimately linked to normal progression through the intrinsic apoptotic pathway. Acknowledgements  We would like to thank Dr. Lesley Kane for her thoughtful reading and comments on the chapter and Dr. Jean Claude Martinou for the SIMH image. Work in the Youle Lab is supported in part by the Intramural Research Program, NINDS, NIH.

References Abdelwahid, E., Yokokura, T., Krieser, R. J., Balasundaram, S., Fowle, W. H. & White, K. (2007) Mitochondrial disruption in Drosophila apoptosis. Dev Cell, 12, 793–806. Alexander, C., Votruba, M., Pesch, U. E., Thiselton, D. L., Mayer, S., Moore, A., Rodriguez, M., Kellner, U., Leo-Kottler, B., Auburger, G., Bhattacharya, S. S. & Wissinger, B. (2000) OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat Genet, 26, 211–5. Amiott, E., Cohen, M., Saint-Georges, Y., Weissman, A. & Shaw, J. (2009) A Mutation Associated with CMT2A Neuropathy Causes Defects in Fzo1 GTP Hydrolysis, Ubiquitylation, and Protein Turnover. Mol Biol Cell. Amutha, B., Gordon, D. M., Gu, Y. & Pain, D. (2004) A novel role of Mgm1p, a dynamin-related GTPase, in ATP synthase assembly and cristae formation/maintenance. Biochem J, 381, 19–23. Arimura, S.-I. & Tsutsumi, N. (2002) A dynamin-like protein (ADL2b), rather than FtsZ, is involved in Arabidopsis mitochondrial division. Proc Natl Acad Sci USA, 99, 5727–31. Arnoult, D., Bartle, L. M., Skaletskaya, A., Poncet, D., Zamzami, N., Park, P. U., Sharpe, J., Youle, R. J. & Goldmacher, V. S. (2004) Cytomegalovirus cell death suppressor vMIA blocks Bax- but not Bak-mediated apoptosis by binding and sequestering Bax at mitochondria. Proc Natl Acad Sci USA, 101, 7988–93. Arnoult, D., Grodet, A., Lee, Y.-J., Estaquier, J. & Blackstone, C. (2005) Release of OPA1 during apoptosis participates in the rapid and complete release of cytochrome c and subsequent mitochondrial fragmentation. J Biol Chem, 280, 35742–50. Bach, D., Pich, S., Soriano, F. X., Vega, N., Baumgartner, B., Oriola, J., Daugaard, J. R., Lloberas, J., Camps, M., Zierath, J. R., Rabasa-Lhoret, R., Wallberg-Henriksson, H., Laville, M., Palacín, M., Vidal, H., Rivera, F., Brand, M. & Zorzano, A. (2003) Mitofusin-2 determines

130

M.M. Cleland and R.J. Youle

mitochondrial network architecture and mitochondrial metabolism. A novel regulatory mechanism altered in obesity. J Biol Chem, 278, 17190–7. Bakhshi, A., Jensen, J. P., Goldman, P., Wright, J. J., Mcbride, O. W., Epstein, A. L. & Korsmeyer, S. J. (1985) Cloning the chromosomal breakpoint of t(14;18) human lymphomas: clustering around JH on chromosome 14 and near a transcriptional unit on 18. Cell, 41, 899–906. Ban, T., Heymann, J. A. W., Song, Z., Hinshaw, J. E. & Chan, D. C. (2010) OPA1 disease alleles causing dominant optic atrophy have defects in cardiolipin-stimulated GTP hydrolysis and membrane tubulation. Hum Mol Genet. Berman, S. B., Chen, Y.-B., Qi, B., Mccaffery, J. M., Rucker, E. B., Goebbels, S., Nave, K.-A., Arnold, B. A., Jonas, E. A., Pineda, F. J. & Hardwick, J. M. (2009) Bcl-x L increases mitochondrial fission, fusion, and biomass in neurons. J Cell Biol, 184, 707–19. Bleazard, W., Mccaffery, J. M., King, E. J., Bale, S., Mozdy, A., Tieu, Q., Nunnari, J. & Shaw, J. M. (1999) The dynamin-related GTPase Dnm1 regulates mitochondrial fission in yeast. Nat Cell Biol, 1, 298–304. Breckenridge, D. G., Kang, B.-H., Kokel, D., Mitani, S., Staehelin, L. A. & Xue, D. (2008) Caenorhabditis elegans drp-1 and fis-2 regulate distinct cell-death execution pathways downstream of ced-3 and independent of ced-9. Mol Cell, 31, 586–97. Breckenridge, D. G., Kang, B.-H. & Xue, D. (2009) Bcl-2 proteins EGL-1 and CED-9 do not regulate mitochondrial fission or fusion in Caenorhabditis elegans. Curr Biol, 19, 768–73. Breckenridge, D. G., Stojanovic, M., Marcellus, R. C. & Shore, G. C. (2003) Caspase cleavage product of BAP31 induces mitochondrial fission through endoplasmic reticulum calcium signals, enhancing cytochrome c release to the cytosol. J Cell Biol, 160, 1115–27. Brooks, C., Wei, Q., Feng, L., Dong, G., Tao, Y., Mei, L., Xie, Z.-J. & DONG, Z. (2007) Bak regulates mitochondrial morphology and pathology during apoptosis by interacting with mitofusins. Proc Natl Acad Sci USA, 104, 11649–54. Cartoni, R., Arnaud, E., Médard, J.-J., Poirot, O., Courvoisier, D. S., Chrast, R. & Martinou, J.-C. (2010) Expression of mitofusin 2(R94Q) in a transgenic mouse leads to Charcot-Marie-Tooth neuropathy type 2A. Brain, 133, 1460–9. Cartoni, R., Leger, B., Hock, M. B., Praz, M., Crettenand, A., Pich, S., Ziltener, J.-L., Luthi, F., Dériaz, O., Zorzano, A., Gobelet, C., Kralli, A. & Russell, A. P. (2005) Mitofusins 1/2 and ERRalpha expression are increased in human skeletal muscle after physical exercise. J Physiol (Lond), 567, 349–58. Cassidy-Stone, A., Chipuk, J. E., Ingerman, E., Song, C., Yoo, C., Kuwana, T., Kurth, M. J., Shaw, J. T., Hinshaw, J. E., Green, D. R. & Nunnari, J. (2008) Chemical inhibition of the mitochondrial division dynamin reveals its role in Bax/Bak-dependent mitochondrial outer membrane permeabilization. Dev Cell, 14, 193–204. Castanier, C., Garcin, D., Vazquez, A. & Arnoult, D. (2010) Mitochondrial dynamics regulate the RIG-I-like receptor antiviral pathway. EMBO Rep, 11, 133–8. Chang, C.-R. & Blackstone, C. (2007) Cyclic AMP-dependent protein kinase phosphorylation of  Drp1 regulates its GTPase activity and mitochondrial morphology. J Biol Chem, 282, 21583–7. Chang, C. R., Manlandro, C. M., Arnoult, D., Stadler, J., Posey, A. E., Hill, R. B. & Blackstone, C. (2010) A lethal de novo mutation in the middle domain of the dynaminrelated GTPase Drp1 impairs higher order assembly and mitochondrial division. J Biol Chem, 285, 32494–503. Chen, H., Chomyn, A. & Chan, D. C. (2005) Disruption of fusion results in mitochondrial heterogeneity and dysfunction. J Biol Chem, 280, 26185–92. Chen, H., Detmer, S. A., Ewald, A. J., Griffin, E. E., Fraser, S. E. & Chan, D. C. (2003) Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J Cell Biol, 160, 189–200. Chen, H., Mccaffery, J. M. & Chan, D. C. (2007) Mitochondrial fusion protects against neurodegeneration in the cerebellum. Cell, 130, 548–62. Chen, H., Vermulst, M., Wang, Y. E., Chomyn, A., Prolla, T. A., Mccaffery, J. M. & Chan, D. C. (2010) Mitochondrial fusion is required for mtDNA stability in skeletal muscle and tolerance of mtDNA mutations. Cell, 141, 280–9.

4  Mitochondrial Dynamics and Apoptosis

131

Chiche, J., Rouleau, M., Gounon, P., Brahimi-Horn, M. C., Pouysségur, J. & Mazure, N. M. (2010) Hypoxic enlarged mitochondria protect cancer cells from apoptotic stimuli. J Cell Physiol, 222, 648–57. Cho, D.-H., Nakamura, T., Fang, J., Cieplak, P., Godzik, A., Gu, Z. & Lipton, S. A. (2009) S-nitrosylation of Drp1 mediates beta-amyloid-related mitochondrial fission and neuronal injury. Science, 324, 102–5. Cipolat, S., Martins De Brito, O., Dal Zilio, B. & Scorrano, L. (2004) OPA1 requires mitofusin 1 to promote mitochondrial fusion. Proc Natl Acad Sci USA, 101, 15927–32. Cipolat, S., Rudka, T., Hartmann, D., Costa, V., Serneels, L., Craessaerts, K., Metzger, K., Frezza, C., Annaert, W., D’adamio, L., Derks, C., Dejaegere, T., Pellegrini, L., D’hooge, R., Scorrano, L. & De Strooper, B. (2006) Mitochondrial rhomboid PARL regulates cytochrome c release during apoptosis via OPA1-dependent cristae remodeling. Cell, 126, 163–75. Cleary, M. L., Smith, S. D. & Sklar, J. (1986) Cloning and structural analysis of cDNAs for bcl-2 and a hybrid bcl-2/immunoglobulin transcript resulting from the t(14;18) translocation. Cell, 47, 19–28. Cleland, M. M., Norris, K. L., Karbowski, M., Wang, C., Suen, D. F., Jiao, S., George, N. M., Luo, X., Li, Z. & Youle, R. J. (2011) Bcl-2 family interaction with the mitochondrial morphogenesis machinery. Cell Death Differ, 18, 235–247. Cohen, M. M. J., Leboucher, G. P., Livnat-Levanon, N., Glickman, M. H. & Weissman, A. M. (2008) Ubiquitin-proteasome-dependent degradation of a mitofusin, a critical regulator of mitochondrial fusion. Mol Biol Cell, 19, 2457–64. Cribbs, J. T. & Strack, S. (2007) Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein kinase and calcineurin regulates mitochondrial fission and cell death. EMBO Rep, 8, 939–44. De Palma, C., Falcone, S., Pisoni, S., Cipolat, S., Panzeri, C., Pambianco, S., Pisconti, A., Allevi, R., Bassi, M. T., Cossu, G., Pozzan, T., Moncada, S., Scorrano, L., Brunelli, S. & Clementi, E. (2010) Nitric oxide inhibition of Drp1-mediated mitochondrial fission is critical for myogenic differentiation. Cell Death Differ. De Vos, K. J., Allan, V. J., Grierson, A. J. & Sheetz, M. P. (2005) Mitochondrial function and actin regulate dynamin-related protein 1-dependent mitochondrial fission. Curr Biol, 15, 678–83. Delettre, C., Griffoin, J. M., Kaplan, J., Dollfus, H., Lorenz, B., Faivre, L., Lenaers, G., Belenguer, P. & Hamel, C. P. (2001) Mutation spectrum and splicing variants in the OPA1 gene. Hum Genet, 109, 584–91. Delettre, C., Lenaers, G., Griffoin, J. M., Gigarel, N., Lorenzo, C., Belenguer, P., Pelloquin, L., Grosgeorge, J., Turc-Carel, C., Perret, E., Astarie-Dequeker, C., Lasquellec, L., Arnaud, B., Ducommun, B., Kaplan, J. & Hamel, C. P. (2000) Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat Genet, 26, 207–10. Delivani, P., Adrain, C., Taylor, R. C., Duriez, P. J. & Martin, S. J. (2006) Role for CED-9 and Egl-1 as regulators of mitochondrial fission and fusion dynamics. Mol Cell, 21, 761–73. Desagher, S. & Martinou, J. C. (2000) Mitochondria as the central control point of apoptosis. Trends Cell Biol, 10, 369–77. Detmer, S. A. & Chan, D. C. (2007) Complementation between mouse Mfn1 and Mfn2 protects mitochondrial fusion defects caused by CMT2A disease mutations. J Cell Biol, 176, 405–14. Devay, R. M., Dominguez-Ramirez, L., Lackner, L. L., Hoppins, S., Stahlberg, H. & Nunnari, J. (2009) Coassembly of Mgm1 isoforms requires cardiolipin and mediates mitochondrial inner membrane fusion. J Cell Biol, 186, 793–803. Diamantis, A., Magiorkinis, E., Sakorafas, G. H. & Androutsos, G. (2008) A brief history of apoptosis: from ancient to modern times. Onkologie, 31, 702–6. Duvezin-Caubet, S., Jagasia, R., Wagener, J., Hofmann, S., Trifunovic, A., Hansson, A., Chomyn, A., Bauer, M. F., Attardi, G., Larsson, N.-G., Neupert, W. & Reichert, A. S. (2006) Proteolytic processing of OPA1 links mitochondrial dysfunction to alterations in mitochondrial morphology. J Biol Chem, 281, 37972–9. Duvezin-Caubet, S., Koppen, M., Wagener, J., Zick, M., Israel, L., Bernacchia, A., Jagasia, R., Rugarli, E. I., Imhof, A., Neupert, W., Langer, T. & Reichert, A. S. (2007) OPA1 processing reconstituted in yeast depends on the subunit composition of the m-AAA protease in mitochondria. Mol Biol Cell, 18, 3582–90.

132

M.M. Cleland and R.J. Youle

Ehses, S., Raschke, I., Mancuso, G., Bernacchia, A., Geimer, S., Tondera, D., Martinou, J.-C., Westermann, B., Rugarli, E. I. & Langer, T. (2009) Regulation of OPA1 processing and mitochondrial fusion by m-AAA protease isoenzymes and OMA1. J Cell Biol, 187, 1023–36. Escobar-Henriques, M., Westermann, B. & Langer, T. (2006) Regulation of mitochondrial fusion by the F-box protein Mdm30 involves proteasome-independent turnover of Fzo1. J Cell Biol, 173, 645–50. Estaquier, J. & Arnoult, D. (2007) Inhibiting Drp1-mediated mitochondrial fission selectively prevents the release of cytochrome c during apoptosis. Cell Death Differ, 14, 1086–94. Eura, Y., Ishihara, N., Yokota, S. & Mihara, K. (2003) Two mitofusin proteins, mammalian homologues of FZO, with distinct functions are both required for mitochondrial fusion. J Biochem, 134, 333–44. Fannjiang, Y., Cheng, W.-C., Lee, S. J., Qi, B., Pevsner, J., Mccaffery, J. M., Hill, R. B., Basañez, G. & Hardwick, J. M. (2004) Mitochondrial fission proteins regulate programmed cell death in yeast. Genes Dev, 18, 2785–97. Ferré, M., Bonneau, D., Milea, D., Chevrollier, A., Verny, C., Dollfus, H., Ayuso, C., Defoort, S., Vignal, C., Zanlonghi, X., Charlin, J.-F., Kaplan, J., Odent, S., Hamel, C. P., Procaccio, V., Reynier, P. & Amati-Bonneau, P. (2009) Molecular screening of 980 cases of suspected hereditary optic neuropathy with a report on 77 novel OPA1 mutations. Hum Mutat, 30, E692–705. Frank, S., Gaume, B., Bergmann-Leitner, E. S., Leitner, W. W., Robert, E. G., Catez, F., Smith, C. L. & Youle, R. J. (2001) The role of dynamin-related protein 1, a mediator of mitochondrial fission, in apoptosis. Dev Cell, 1, 515–25. Frezza, C., Cipolat, S., Martins De Brito, O., Micaroni, M., Beznoussenko, G. V., Rudka, T., Bartoli, D., Polishuck, R. S., Danial, N. N., De Strooper, B. & Scorrano, L. (2006) OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion. Cell, 126, 177–89. Gandre-Babbe, S. & Van Der Bliek, A. M. (2008) The novel tail-anchored membrane protein Mff controls mitochondrial and peroxisomal fission in mammalian cells. Mol Biol Cell, 19, 2402–12. Gegg, M. E., Cooper, J. M., Chau, K. Y., Rojo, M., Schapira, A. H. & Taanman, J. W. (2010) Mitofusin 1 and mitofusin 2 are ubiquitinated in a PINK1/parkin-dependent manner upon induction of mitophagy. Hum Mol Genet, 19, 4861–70. Germain, M., Mathai, J. P., Mcbride, H. M. & Shore, G. C. (2005) Endoplasmic reticulum BIK initiates DRP1-regulated remodelling of mitochondrial cristae during apoptosis. EMBO J, 24, 1546–56. Glucksmann, A. (1951) Cell Deaths in Normal Vertebrate Ontogeny. Biol Rev Camb Philos Soc, 26, 59–86 Gorsich, S. W. & Shaw, J. M. (2004) Importance of mitochondrial dynamics during meiosis and sporulation. Mol Biol Cell, 15, 4369–81. Goyal, G., Fell, B., Sarin, A., Youle, R. J. & Sriram, V. (2007) Role of mitochondrial remodeling in programmed cell death in Drosophila melanogaster. Dev Cell, 12, 807–16. Griffin, E. E. & Chan, D. C. (2006) Domain interactions within Fzo1 oligomers are essential for mitochondrial fusion. J Biol Chem, 281, 16599–606. Griffin, E. E., Graumann, J. & Chan, D. C. (2005) The WD40 protein Caf4p is a component of the mitochondrial fission machinery and recruits Dnm1p to mitochondria. J Cell Biol, 170, 237–48. Griparic, L., Kanazawa, T. & Van Der Bliek, A. M. (2007) Regulation of the mitochondrial dynamin-like protein Opa1 by proteolytic cleavage. J Cell Biol, 178, 757–64. Griparic, L., Van Der Wel, N. N., Orozco, I. J., Peters, P. J. & Van Der Bliek, A. M. (2004) Loss of the intermembrane space protein Mgm1/OPA1 induces swelling and localized constrictions along the lengths of mitochondria. J Biol Chem, 279, 18792–8. Guan, K., Farh, L., Marshall, T. K. & Deschenes, R. J. (1993) Normal mitochondrial structure and genome maintenance in yeast requires the dynamin-like product of the MGM1 gene. Curr Genet, 24, 141–8. Guillery, O., Malka, F., Landes, T., Guillou, E., Blackstone, C., Lombès, A., Belenguer, P., Arnoult, D. & Rojo, M. (2007) Metalloprotease-mediated OPA1 processing is modulated by the mitochondrial membrane potential. Biol Cell.

4  Mitochondrial Dynamics and Apoptosis

133

Hájek, P., Chomyn, A. & Attardi, G. (2007) Identification of a novel mitochondrial complex containing mitofusin 2 and stomatin-like protein 2. J Biol Chem, 282, 5670–81. Hales, K. G. & Fuller, M. T. (1997) Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase. Cell, 90, 121–9. Harder, Z., Zunino, R. & Mcbride, H. (2004) Sumo1 conjugates mitochondrial substrates and participates in mitochondrial fission. Curr Biol, 14, 340–5. Head, B., Griparic, L., Amiri, M., Gandre-Babbe, S. & Van Der Bliek, A. M. (2009) Inducible proteolytic inactivation of OPA1 mediated by the OMA1 protease in mammalian cells. J Cell Biol, 187, 959–66. Herlan, M., Vogel, F., Bornhovd, C., Neupert, W. & Reichert, A. S. (2003) Processing of Mgm1 by the rhomboid-type protease Pcp1 is required for maintenance of mitochondrial morphology and of mitochondrial DNA. J Biol Chem, 278, 27781–8. Hermann, G. J., Thatcher, J. W., Mills, J. P., Hales, K. G., Fuller, M. T., Nunnari, J. & Shaw, J. M. (1998) Mitochondrial fusion in yeast requires the transmembrane GTPase Fzo1p. J Cell Biol, 143, 359–73. Hernández-Alvarez, M. I., Thabit, H., Burns, N., Shah, S., Brema, I., Hatunic, M., Finucane, F., Liesa, M., Chiellini, C., Naon, D., Zorzano, A. & Nolan, J. J. (2010) Subjects with early-onset type 2 diabetes show defective activation of the skeletal muscle PGC-1{alpha}/Mitofusin-2 regulatory pathway in response to physical activity. Diabetes Care, 33, 645–51. Hoppins, S., Edlich, F., Cleland, M. M., Banerjee, S., Mccaffery, J. M., Youle, R. J. & Nunnari, J. (2011) The soluble form of Bax regulates mitochondrial fusion via MFN2 homotypic complexes. Mol Cell, 41, 150–60. Hoppins, S., Lackner, L. & Nunnari, J. (2007) The machines that divide and fuse mitochondria. Annu Rev Biochem, 76, 751–80. Hwa, J. J., Hiller, M. A., Fuller, M. T. & Santel, A. (2002) Differential expression of the Drosophila mitofusin genes fuzzy onions (fzo) and dmfn. Mech Dev, 116, 213–6. Ichishita, R., Tanaka, K., Sugiura, Y., Sayano, T., Mihara, K. & Oka, T. (2008) An RNAi screen for mitochondrial proteins required to maintain the morphology of the organelle in Caenorhabditis elegans. J Biochem, 143, 449–54. Ingerman, E., Perkins, E. M., Marino, M., Mears, J. A., Mccaffery, J. M., Hinshaw, J. E. & Nunnari, J. (2005) Dnm1 forms spirals that are structurally tailored to fit mitochondria. J Cell Biol, 170, 1021–7. Ishihara, N., Eura, Y. & Mihara, K. (2004) Mitofusin 1 and 2 play distinct roles in mitochondrial fusion reactions via GTPase activity. J Cell Sci, 117, 6535–46. Ishihara, N., Fujita, Y., Oka, T. & Mihara, K. (2006) Regulation of mitochondrial morphology through proteolytic cleavage of OPA1. EMBO J, 25, 2966–77. Ishihara, N., Nomura, M., Jofuku, A., Kato, H., Suzuki, S. O., Masuda, K., Otera, H., Nakanishi, Y., Nonaka, I., Goto, Y.-I., Taguchi, N., Morinaga, H., Maeda, M., Takayanagi, R., Yokota, S. & Mihara, K. (2009) Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nat Cell Biol, 11, 958–66. Jagasia, R., Grote, P., Westermann, B. & Conradt, B. (2005) DRP-1-mediated mitochondrial fragmentation during EGL-1-induced cell death in C. elegans. Nature, 433, 754–60. Jahani-Asl, Cheung, Neuspiel, Maclaurin, Fortin, Park, Mcbride & Slack (2007) Mitofusin 2 protects cerebellar granule neurons against injury induced cell death. J Biol Chem. Jones, B. A. & Fangman, W. L. (1992) Mitochondrial DNA maintenance in yeast requires a protein containing a region related to the GTP-binding domain of dynamin. Genes Dev, 6, 380–9. Kanazawa, T., Zappaterra, M. D., Hasegawa, A., Wright, A. P., Newman-Smith, E. D., Buttle, K. F., Mcdonald, K., Mannella, C. A. & Van Der Bliek, A. M. (2008) The C. elegans Opa1 homologue EAT-3 is essential for resistance to free radicals. PLoS Genet, 4, e1000022. Karbowski, M., Arnoult, D., Chen, H., Chan, D. C., Smith, C. L. & Youle, R. J. (2004) Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J Cell Biol, 164, 493–9. Karbowski, M., Lee, Y.-J., Gaume, B., Jeong, S.-Y., Frank, S., Nechushtan, A., Santel, A., Fuller, M., Smith, C. L. & Youle, R. J. (2002) Spatial and temporal association of Bax with mitochondrial fission sites, Drp1, and Mfn2 during apoptosis. J Cell Biol, 159, 931–8.

134

M.M. Cleland and R.J. Youle

Karbowski, M., Neutzner, A. & Youle, R. J. (2007) The mitochondrial E3 ubiquitin ligase MARCH5 is required for Drp1 dependent mitochondrial division. J Cell Biol, 178, 71–84. Karbowski, M., Norris, K. L., Cleland, M. M., Jeong, S.-Y. & Youle, R. J. (2006) Role of Bax and Bak in mitochondrial morphogenesis. Nature, 443, 658–62. Kita, T., Nishida, H., Shibata, H., Niimi, S., Higuti, T. & Arakaki, N. (2009) Possible role of mitochondrial remodelling on cellular triacylglycerol accumulation. J Biochem, 146, 787–96. Kluck, R. M., Bossy-Wetzel, E., Green, D. R. & Newmeyer, D. D. (1997) The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science, 275, 1132–6. Koch, A., Thiemann, M., Grabenbauer, M., Yoon, Y., Mcniven, M. A. & Schrader, M. (2003) Dynamin-like protein 1 is involved in peroxisomal fission. J Biol Chem, 278, 8597–605. Koshiba, T., Detmer, S. A., Kaiser, J. T., Chen, H., Mccaffery, J. M. & Chan, D. C. (2004) Structural basis of mitochondrial tethering by mitofusin complexes. Science, 305, 858–62. Labrousse, A. M., Zappaterra, M. D., Rube, D. A. & Van Der Bliek, A. M. (1999) C. elegans dynamin-related protein DRP-1 controls severing of the mitochondrial outer membrane. Mol Cell, 4, 815–26. Lackner, L. L. & Nunnari, J. (2010) Small molecule inhibitors of mitochondrial division: tools that translate basic biological research into medicine. Chem Biol, 17, 578–83. Lee, Y.-J., Jeong, S.-Y., Karbowski, M., Smith, C. L. & Youle, R. J. (2004) Roles of the mammalian mitochondrial fission and fusion mediators Fis1, Drp1, and Opa1 in apoptosis. Mol Biol Cell, 15, 5001–11. Lenaers, G., Reynier, P., Elachouri, G., Soukkarieh, C., Olichon, A., Belenguer, P., Baricault, L., Ducommun, B., Hamel, C. & Delettre, C. (2009) OPA1 functions in mitochondria and dysfunctions in optic nerve. Int J Biochem Cell Biol, 41, 1866–74. Letai, A., Bassik, M. C., Walensky, L. D., Sorcinelli, M. D., Weiler, S. & Korsmeyer, S. J. (2002) Distinct BH3 domains either sensitize or activate mitochondrial apoptosis, serving as prototype cancer therapeutics. Cancer Cell, 2, 183–92. Li, H., Chen, Y., Jones, A. F., Sanger, R. H., Collis, L. P., Flannery, R., Mcnay, E. C., Yu, T., Schwarzenbacher, R., Bossy, B., Bossy-Wetzel, E., Bennett, M. V. L., Pypaert, M., Hickman, J. A., Smith, P. J. S., Hardwick, J. M. & Jonas, E. A. (2008) Bcl-xL induces Drp1-dependent synapse formation in cultured hippocampal neurons. Proc Natl Acad Sci USA, 105, 2169–74. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., Alnemri, E. S. & Wang, X. (1997) Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell, 91, 479–89. Liu, Q. A. & Shio, H. (2008) Mitochondrial morphogenesis, dendrite development, and synapse formation in cerebellum require both Bcl-w and the glutamate receptor delta2. PLoS Genet, 4, e1000097. Liu, X., Kim, C. N., Yang, J., Jemmerson, R. & Wang, X. (1996) Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell, 86, 147–57. Liu, X., Weaver, D., Shirihai, O. & Hajnóczky, G. (2009) Mitochondrial ‘kiss-and-run’: interplay between mitochondrial motility and fusion-fission dynamics. EMBO J, 28, 3074–89. Loucks, F. A., Schroeder, E. K., Zommer, A. E., Hilger, S., Kelsey, N. A., Bouchard, R. J., Blackstone, C., Brewster, J. L. & Linseman, D. A. (2009) Caspases indirectly regulate cleavage of the mitochondrial fusion GTPase OPA1 in neurons undergoing apoptosis. Brain Res, 1250, 63–74. Malena, A., Loro, E., Di Re, M., Holt, I. J. & Vergani, L. (2009) Inhibition of mitochondrial fission favours mutant over wild-type mitochondrial DNA. Hum Mol Genet, 18, 3407–16. Mancini, M., Anderson, B. O., Caldwell, E., Sedghinasab, M., Paty, P. B. & Hockenbery, D. M. (1997) Mitochondrial proliferation and paradoxical membrane depolarization during terminal differentiation and apoptosis in a human colon carcinoma cell line. J Cell Biol, 138, 449–69. Mcbride, H. & Soubannier, V. (2010) Mitochondrial function: OMA1 and OPA1, the grandmasters of mitochondrial health. Curr Biol, 20, R274–6. Mccormick, A. L., Smith, V. L., Chow, D. & Mocarski, E. S. (2003) Disruption of mitochondrial networks by the human cytomegalovirus UL37 gene product viral mitochondrion-localized inhibitor of apoptosis. J Virol, 77, 631–41.

4  Mitochondrial Dynamics and Apoptosis

135

Mcquibban, G. A., Saurya, S. & Freeman, M. (2003) Mitochondrial membrane remodelling regulated by a conserved rhomboid protease. Nature, 423, 537–41. Meeusen, S., Devay, R., Block, J., Cassidy-Stone, A., Wayson, S., Mccaffery, J. M. & Nunnari, J. (2006) Mitochondrial inner-membrane fusion and crista maintenance requires the dynaminrelated GTPase Mgm1. Cell, 127, 383–95. Meeusen, S., Mccaffery, J. M. & Nunnari, J. (2004) Mitochondrial fusion intermediates revealed in vitro. Science, 305, 1747–52. Meglei, G. & Mcquibban, G. A. (2009) The dynamin-related protein Mgm1p assembles into oligomers and hydrolyzes GTP to function in mitochondrial membrane fusion. Biochemistry, 48, 1774–84. Merkwirth, C., Dargazanli, S., Tatsuta, T., Geimer, S., Löwer, B., Wunderlich, F. T., Von KleistRetzow, J.-C., Waisman, A., Westermann, B. & Langer, T. (2008) Prohibitins control cell proliferation and apoptosis by regulating OPA1-dependent cristae morphogenesis in mitochondria. Genes Dev, 22, 476–88. Misaka, T., Miyashita, T. & Kubo, Y. (2002) Primary structure of a dynamin-related mouse mitochondrial GTPase and its distribution in brain, subcellular localization, and effect on mitochondrial morphology. J Biol Chem, 277, 15834–42. Montessuit, S., Somasekharan, S. P., Terrones, O., Lucken-Ardjomande, S., Herzig, S., Schwarzenbacher, R., Manstein, D. J., Bossy-Wetzel, E., Basanez, G., Meda, P. & Martinou, J. C. (2010) Membrane remodeling induced by the dynamin-related protein Drp1 stimulates Bax oligomerization. Cell, 142, 889–901. Mozdy, A. D., Mccaffery, J. M. & Shaw, J. M. (2000) Dnm1p GTPase-mediated mitochondrial fission is a multi-step process requiring the novel integral membrane component Fis1p. J Cell Biol, 151, 367–80. Nakamura, N., Kimura, Y., Tokuda, M., Honda, S. & Hirose, S. (2006) MARCH-V is a novel mitofusin 2- and Drp1-binding protein able to change mitochondrial morphology. EMBO Rep, 7, 1019–22. Nechushtan, A., Smith, C. L., Lamensdorf, I., Yoon, S. H. & Youle, R. J. (2001) Bax and Bak coalesce into novel mitochondria-associated clusters during apoptosis. J Cell Biol, 153, 1265–76. Neuspiel, M., Zunino, R., Gangaraju, S., Rippstein, P. & Mcbride, H. (2005) Activated mitofusin 2 signals mitochondrial fusion, interferes with Bax activation, and reduces susceptibility to radical induced depolarization. J Biol Chem, 280, 25060–70. Neutzner, A. & Youle, R. J. (2005) Instability of the mitofusin Fzo1 regulates mitochondrial morphology during the mating response of the yeast Saccharomyces cerevisiae. J Biol Chem, 280, 18598–603. Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P. & Lenaers, G. (2003) Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J Biol Chem, 278, 7743–6. Olichon, A., Elachouri, G., Baricault, L., Delettre, C., Belenguer, P. & Lenaers, G. (2007) OPA1 alternate splicing uncouples an evolutionary conserved function in mitochondrial fusion from a vertebrate restricted function in apoptosis. Cell Death Differ, 14, 682–92. Ong, S.-B., Subrayan, S., Lim, S. Y., Yellon, D. M., Davidson, S. M. & Hausenloy, D. J. (2010) Inhibiting Mitochondrial Fission Protects the Heart Against Ischemia/Reperfusion Injury. Circulation. Otera, H., Wang, C., Cleland, M. M., Setoguchi, K., Yokota, S., Youle, R. J. & Mihara, K. (2010) Mff is an essential factor for mitochondrial recruitment of Drp1 during mitochondrial fission in mammalian cells. J Cell Biol, 191, 1141–58. Otsuga, D., Keegan, B. R., Brisch, E., Thatcher, J. W., Hermann, G. J., Bleazard, W. & Shaw, J. M. (1998) The dynamin-related GTPase, Dnm1p, controls mitochondrial morphology in yeast. J Cell Biol, 143, 333–49. Ott, M., Robertson, J. D., Gogvadze, V., Zhivotovsky, B. & Orrenius, S. (2002) Cytochrome c release from mitochondria proceeds by a two-step process. Proc Natl Acad Sci USA, 99, 1259–63. Parone, P. A., Da Cruz, S., Tondera, D., Mattenberger, Y., James, D. I., Maechler, P., Barja, F. & Martinou, J.-C. (2008) Preventing mitochondrial fission impairs mitochondrial function and leads to loss of mitochondrial DNA. PLoS ONE, 3, e3257.

136

M.M. Cleland and R.J. Youle

Parone, P. A., James, D. I., Da Cruz, S., Mattenberger, Y., Donzé, O., Barja, F. & Martinou, J.-C. (2006) Inhibiting the mitochondrial fission machinery does not prevent Bax/Bak-dependent apoptosis. Mol Cell Biol, 26, 7397–408. Pitts, K. R., Yoon, Y., Krueger, E. W. & Mcniven, M. A. (1999) The dynamin-like protein DLP1 is essential for normal distribution and morphology of the endoplasmic reticulum and mitochondria in mammalian cells. Mol Biol Cell, 10, 4403–17. Poncet, D., Larochette, N., Pauleau, A. L., Boya, P., Jalil, A. A., Cartron, P. F., Vallette, F., Schnebelen, C., Bartle, L. M., Skaletskaya, A., Boutolleau, D., Martinou, J. C., Goldmacher, V. S., Kroemer, G. & Zamzami, N. (2004) An anti-apoptotic viral protein that recruits Bax to mitochondria. J Biol Chem, 279, 22605–14. Poole, A. C., Thomas, R. E., YU, S., Vincow, E. S. & Pallanck, L. (2010) The mitochondrial fusion-promoting factor mitofusin is a substrate of the PINK1/parkin pathway. PLoS ONE, 5, e10054. Rapaport, D., Brunner, M., Neupert, W. & Westermann, B. (1998) Fzo1p is a mitochondrial outer membrane protein essential for the biogenesis of functional mitochondria in Saccharomyces cerevisiae. J Biol Chem, 273, 20150–5. Rikhy, R., Kamat, S., Ramagiri, S., Sriram, V. & Krishnan, K. S. (2007) Mutations in dynaminrelated protein result in gross changes in mitochondrial morphology and affect synaptic vesicle recycling at the Drosophila neuromuscular junction. Genes Brain Behav, 6, 42–53. Rojo, M., Legros, F., Chateau, D. & Lombès, A. (2002) Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo. J Cell Sci, 115, 1663–74. Rolland, S. G., Lu, Y., David, C. N. & Conradt, B. (2009) The BCL-2-like protein CED-9 of C. elegans promotes FZO-1/Mfn1,2- and EAT-3/Opa1-dependent mitochondrial fusion. J Cell Biol, 186, 525–40. Rujiviphat, J., Meglei, G., Rubinstein, J. L. & Mcquibban, G. A. (2009) Phospholipid association is essential for dynamin-related protein Mgm1 to function in mitochondrial membrane fusion. J Biol Chem, 284, 28682–6. Santel, A. & Fuller, M. T. (2001) Control of mitochondrial morphology by a human mitofusin. J Cell Sci, 114, 867–74. Schauss, A. C., Bewersdorf, J. & Jakobs, S. (2006) Fis1p and Caf4p, but not Mdv1p, determine the polar localization of Dnm1p clusters on the mitochondrial surface. J Cell Sci, 119, 3098–106. Scorrano, L., Ashiya, M., Buttle, K., Weiler, S., Oakes, S. A., Mannella, C. A. & Korsmeyer, S. J. (2002) A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis. Dev Cell, 2, 55–67. Sesaki, H., Southard, S. M., Hobbs, A. E. A. & Jensen, R. E. (2003) Cells lacking Pcp1p/Ugo2p, a rhomboid-like protease required for Mgm1p processing, lose mtDNA and mitochondrial structure in a Dnm1p-dependent manner, but remain competent for mitochondrial fusion. Biochem Biophys Res Commun, 308, 276–83. Shen, T., Zheng, M., Cao, C., Chen, C., Tang, J., Zhang, W., Cheng, H., Chen, K.-H. & Xiao, R.-P. (2007) Mitofusin-2 is a major determinant of oxidative stress-mediated heart muscle cell apoptosis. J Biol Chem, 282, 23354–61. Shepard, K. A. & Yaffe, M. P. (1999) The yeast dynamin-like protein, Mgm1p, functions on the mitochondrial outer membrane to mediate mitochondrial inheritance. J Cell Biol, 144, 711–20. Smirnova, E., Griparic, L., Shurland, D. L. & Van Der Bliek, A. M. (2001) Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells. Mol Biol Cell, 12, 2245–56. Smirnova, E., Shurland, D. L., Ryazantsev, S. N. & Van Der Bliek, A. M. (1998) A human dynamin-related protein controls the distribution of mitochondria. J Cell Biol, 143, 351–8. Song, Z., Chen, H., Fiket, M., Alexander, C. & Chan, D. C. (2007) OPA1 processing controls mitochondrial fusion and is regulated by mRNA splicing, membrane potential, and Yme1L. J Cell Biol, 178, 749–55. Song, Z., Ghochani, M., Mccaffery, J. M., Frey, T. G. & Chan, D. C. (2009) Mitofusins and OPA1 mediate sequential steps in mitochondrial membrane fusion. Mol Biol Cell, 20, 3525–32.

4  Mitochondrial Dynamics and Apoptosis

137

Sugioka, R., Shimizu, S. & Tsujimoto, Y. (2004) Fzo1, a protein involved in mitochondrial fusion, inhibits apoptosis. J Biol Chem, 279, 52726–34. Sun, M. G., Williams, J., Munoz-Pinedo, C., Perkins, G. A., Brown, J. M., Ellisman, M. H., Green, D. R. & Frey, T. G. (2007) Correlated three-dimensional light and electron microscopy reveals transformation of mitochondria during apoptosis. Nat Cell Biol, 9, 1057–65. Taguchi, N., Ishihara, N., Jofuku, A., Oka, T. & Mihara, K. (2007) Mitotic phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial fission. J Biol Chem, 282, 11521–9. Tan, F. J., Husain, M., Manlandro, C. M., Koppenol, M., Fire, A. Z. & Hill, R. B. (2008) CED-9 and mitochondrial homeostasis in C. elegans muscle. J Cell Sci, 121, 3373–82. Tanaka, A., Cleland, M. M., Xu, S., Narendra, D. P., Suen, D. F., Karbowski, M. & Youle, R. J. (2010) Proteasome and p97 mediate mitophagy and degradation of mitofusins induced by Parkin. J Cell Biol, 191, 1367-80. Tanner, E. A., Blute, T. A., Brachmann, C. B. & Mccall, K. (2010) Bcl-2 proteins and autophagy regulate mitochondrial dynamics during programmed cell death in the Drosophila ovary. Development, 138, 327–38. Tieu, Q. & Nunnari, J. (2000) Mdv1p is a WD repeat protein that interacts with the dynaminrelated GTPase, Dnm1p, to trigger mitochondrial division. J Cell Biol, 151, 353–66. Tieu, Q., Okreglak, V., Naylor, K. & Nunnari, J. (2002) The WD repeat protein, Mdv1p, functions as a molecular adaptor by interacting with Dnm1p and Fis1p during mitochondrial fission. J Cell Biol, 158, 445–52. Tondera, D., Grandemange, S., Jourdain, A., Karbowski, M., Mattenberger, Y., Herzig, S., Da Cruz, S., Clerc, P., Raschke, I., Merkwirth, C., Ehses, S., Krause, F., Chan, D. C., Alexander, C., Bauer, C., Youle, R., Langer, T. & Martinou, J.-C. (2009) SLP-2 is required for stress-induced mitochondrial hyperfusion. EMBO J, 28, 1589–600. Tsujimoto, Y., Cossman, J., Jaffe, E. & Croce, C. M. (1985) Involvement of the bcl-2 gene in human follicular lymphoma. Science, 228, 1440–3. Twig, G., Elorza, A., Molina, A. J. A., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E. & Shirihai, O. S. (2008) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J, 27, 433–46. Varadi, A., Johnson-Cadwell, L. I., Cirulli, V., Yoon, Y., Allan, V. J. & Rutter, G. A. (2004) Cytoplasmic dynein regulates the subcellular distribution of mitochondria by controlling the recruitment of the fission factor dynamin-related protein-1. J Cell Sci, 117, 4389–400. Vaux, D. L., Cory, S. & Adams, J. M. (1988) Bcl-2 gene promotes haemopoietic cell survival and cooperates with c-myc to immortalize pre-B cells. Nature, 335, 440–2. VOTRUBA, M. (2004) Molecular genetic basis of primary inherited optic neuropathies. Eye (Lond), 18, 1126–32. Wakabayashi, J., Zhang, Z., Wakabayashi, N., Tamura, Y., Fukaya, M., Kensler, T. W., Iijima, M. & Sesaki, H. (2009) The dynamin-related GTPase Drp1 is required for embryonic and brain development in mice. J Cell Biol, 186, 805–16. Wang, C. & Youle, R. (2009) The Role of Mitochondria in Apoptosis. Annu Rev Genet. Wasiak, S., Zunino, R. & Mcbride, H. M. (2007) Bax/Bak promote sumoylation of DRP1 and its stable association with mitochondria during apoptotic cell death. J Cell Biol, 177, 439–50. Waterham, H. R., Koster, J., Van Roermund, C. W., Mooyer, P. A., Wanders, R. J. & Leonard, J. V. (2007) A lethal defect of mitochondrial and peroxisomal fission. N Engl J Med, 356, 1736–41. Waxman, A. & Kolliputi, N. (2009) IL-6 Protects Against Hyperoxia Induced Mitochondrial Damage via BCL-2 induced BAK Interactions with Mitofusions. Am J Respir Cell Mol Biol. Willis, S. N., Fletcher, J. I., Kaufmann, T., Van Delft, M. F., Chen, L., Czabotar, P. E., Ierino, H., Lee, E. F., Fairlie, W. D., Bouillet, P., Strasser, A., Kluck, R. M., Adams, J. M. & Huang, D. C. (2007) Apoptosis initiated when BH3 ligands engage multiple Bcl-2 homologs, not Bax or Bak. Science, 315, 856–9. Wolter, K. G., Hsu, Y. T., Smith, C. L., Nechushtan, A., Xi, X. G. & Youle, R. J. (1997) Movement of Bax from the cytosol to mitochondria during apoptosis. J Cell Biol, 139, 1281–92.

138

M.M. Cleland and R.J. Youle

Wong, E. D., Wagner, J. A., Gorsich, S. W., Mccaffery, J. M., Shaw, J. M. & Nunnari, J. (2000) The dynamin-related GTPase, Mgm1p, is an intermembrane space protein required for maintenance of fusion competent mitochondria. J Cell Biol, 151, 341–52. Wong, E. D., Wagner, J. A., Scott, S. V., Okreglak, V., Holewinske, T. J., Cassidy-Stone, A. & Nunnari, J. (2003) The intramitochondrial dynamin-related GTPase, Mgm1p, is a component of a protein complex that mediates mitochondrial fusion. J Cell Biol, 160, 303–11. Yamaguchi, R., Lartigue, L., Perkins, G., Scott, R. T., Dixit, A., Kushnareva, Y., Kuwana, T., Ellisman, M. H. & Newmeyer, D. D. (2008) Opa1-mediated cristae opening is Bax/Bak and BH3 dependent, required for apoptosis, and independent of Bak oligomerization. Mol Cell, 31, 557–69. Yang, J., Liu, X., Bhalla, K., Kim, C. N., Ibrado, A. M., Cai, J., Peng, T. I., Jones, D. P. & Wang, X. (1997) Prevention of apoptosis by Bcl-2: release of cytochrome c from mitochondria blocked. Science, 275, 1129–32. Yonashiro, R., Ishido, S., Kyo, S., Fukuda, T., Goto, E., Matsuki, Y., Ohmura-Hoshino, M., Sada, K., Hotta, H., Yamamura, H., Inatome, R. & Yanagi, S. (2006) A novel mitochondrial ubiquitin ligase plays a critical role in mitochondrial dynamics. EMBO J, 25, 3618–26. Youle, R. J. & Strasser, A. (2008) The BCL-2 protein family: opposing activities that mediate cell death. Nat Rev Mol Cell Biol, 9, 47–59. Ziviani, E., Tao, R. N. & Whitworth, A. J. (2010) Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc Natl Acad Sci U S A, 107, 5018–23. Züchner, S., De Jonghe, P., Jordanova, A., Claeys, K. G., Guergueltcheva, V., Cherninkova, S., Hamilton, S. R., Van Stavern, G., Krajewski, K. M., Stajich, J., Tournev, I., Verhoeven, K., Langerhorst, C. T., De Visser, M., Baas, F., Bird, T., Timmerman, V., Shy, M. & Vance, J. M. (2006) Axonal neuropathy with optic atrophy is caused by mutations in mitofusin 2. Ann Neurol, 59, 276–81. Züchner, S., Mersiyanova, I. V., Muglia, M., Bissar-Tadmouri, N., Rochelle, J., Dadali, E. L., Zappia, M., Nelis, E., Patitucci, A., Senderek, J., Parman, Y., Evgrafov, O., Jonghe, P. D., Takahashi, Y., Tsuji, S., Pericak-Vance, M. A., Quattrone, A., Battaloglu, E., Polyakov, A. V., Timmerman, V., Schröder, J. M., Vance, J. M. & Battologlu, E. (2004) Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat Genet, 36, 449–51. Zunino, R., Schauss, A., Rippstein, P., Andrade-Navarro, M. & Mcbride, H. M. (2007) The SUMO protease SENP5 is required to maintain mitochondrial morphology and function. J Cell Sci, 120, 1178–88.

Chapter 5

Mitochondrial Dynamics and Axonal Transport Qian Cai and Zu-Hang Sheng

Abstract  Mitochondria are essential organelles for neuronal survival and function through ATP generation, calcium buffering, and apoptotic signaling. Due to their extreme polarity, neurons utilize specialized mechanisms to regulate mitochondrial transport along axons to areas in which energy production and calcium homeostasis are in high demand, namely, active growth cones, nodes of Ranvier, and synaptic terminals. Axonal mitochondria display complex mobility patterns and undergo saltatory and bidirectional movement. While one-third of axonal mitochondria are mobile, the rest remains stationary. Docked mitochondria serve as local power plants and maintain local Ca2+ homeostasis. The balance between mobile and stationary mitochondria is influenced by the diverse physiological states of axons and synapses. The coordination of mitochondrial mobility by axonal physiology is crucial for neuronal development and synaptic function. Defects in mitochondrial transport have been implicated in the pathologic processes of several major neurodegenerative diseases. Thus, understanding the regulation of axonal mitochondrial transport will increase our knowledge of fundamental processes that may affect human neurological disorders. In this chapter, we introduce recent advances in our understanding of motor-adaptor complexes and docking machinery that mediate mitochondrial transport and axonal distribution. We will also discuss the molecular mechanisms underlying the complex mobility patterns of axonal mitochondria and how mitochondrial mobility impacts the physiology and function of synapses. Keywords  Mitochondria • Axonal transport • Docking • Synaptic plasticity • Kinesin • Motor adaptor • Anterograde transport • Retrograde transport • Stationary mitochondria • Mitochondrial mobility

Q. Cai and Z.-H. Sheng () Synaptic Function Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Building 35, Room 2B-215, 35 Convent Drive, Bethesda, MD 20892–3706, USA e-mail: [email protected]. B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_5, © Springer Science+Business Media B.V. 2011

139

140

Q. Cai and Z.-H. Sheng

Abbreviations (MT) (NF) (KIFs) (KHC) (KLC) (DHC) (DIC) (DLIC) (DLC) (dMiro) (TRAK1) (FEZ1) (JIP1) (SNPH) (FCCP) (DRG) (NO) (Mfn2) (EPSCs)

Microtubule Neurofilament kinesin superfamily proteins kinesin heavy chains kinesin light chains dynein heavy chains dynein intermediate chains dynein light intermediate chains dynein light chains drosophila mitochondrial Rho-GTPase trafficking protein kinesin-binding 1 fasciculation and elongation protein zeta-1 c-Jun N-terminal kinase (JNK)-interacting protein syntaphilin cyanide-p-trifluoromethoxyphenyhydrazone dorsal root ganglion nitric oxide mitofusin 2 excitatory postsynaptic currents

5.1 Introduction Mitochondria are thought to produce more than 90% of the cellular ATP in neurons and support many essential functions including the mobilization of synaptic vesicles during intensive neuronal activity (Verstreken et  al. 2005), assembly of the actin cytoskeleton within synaptic boutons (Lee and Peng 2008), and reversal of the ion influxes across their electrochemical gradients for the generation and propagation of action potential, and the maintenance of synaptic potential. In addition to aerobic ATP production, mitochondria have been implicated in certain forms of short-term synaptic plasticity via their ability to buffer Ca2+ at nerve terminals (Tang and Zucker 1997; Billups and Forsythe 2002; Levy et al. 2003; Yang et al. 2003; Kang et al. 2008). The loss of mitochondria from axon terminals results in impaired synaptic transmission (Stowers et al. 2002; Guo et al. 2005; Verstreken et  al. 2005; Ma et  al. 2009). The dysfunction and defective transport of axonal mitochondria have been implicated in the pathogenesis of neurodegenerative diseases such as Alzheimer’s, Huntington’s, and amyotrophic lateral sclerosis (see reviews by Chan 2006; Stokin and Goldstein 2006. Please also refer to other chapters of this book). Neurons are highly polarized and consist of three distinct functional and subcellular domains: a cell body (or soma), a long axon with a uniform diameter, and

5  Mitochondrial Dynamics and Axonal Transport

141

thick dendrites with many branches. The majority of mitochondria are generated within the soma and delivered to their final destination via long neuronal processes. Mitochondria are transported down axons and stop at areas where metabolic demand is high, including active growth cones, axonal branches, synapses, nodes of Ranvier, myelination boundaries and regions of demyelination (Povlishock 1976; Gotow et al. 1991; Bogan and Cabot 1991; Fabricius et al. 1993; Morris and Hollenbeck 1993; Mutsaers and Carroll 1998; Bristow et  al. 2002; Ruthel and Hollenbeck 2003; Kang et al. 2008; Zhang et al. 2010). Axonal distribution of mitochondria is achieved by dynamic and bidirectional mitochondrial movements along microtubule (MT)-based tracks, characterized by frequent stops, changes in direction, and persistent docking in certain regions. Individual mitochondria exhibit frequent pauses and reversals. The complex mobility patterns suggest that axonal mitochondria are coupled to two opposing molecular motors- kinesins and dyneins- along with docking/anchoring machinery. While kinesin motors are responsible for anterograde transport, cytoplasmic dynein motors are the driving force behind retrograde movement (Tanaka et  al. 1998; Martin et al. 1999; Górska-Andrzejak et al. 2003; Pilling et al. 2006). Syntaphilin targets axonal mitochondria and mediates their stationary docking by interacting with MTs (Kang et al. 2008; Chen et al. 2009). Such a mechanism enables neurons to maintain proper mitochondrial density along axons and near synapses. While one-third of axonal mitochondria are mobile in mature neurons, a large fraction (~65–80%) remains stationary. The net movement of individual mitochondria is significantly influenced by their time spent between stationary or mobile states. The average mitochondrial velocity along axons falls between fast-moving vesicles and slow-moving cytoskeletal proteins. Saltatory and bidirectional movements through a combination of dynamic events result in net velocities of 0.3–2.0 mm s−1 (Morris and Hollenbeck 1993; Ligon and Steward 2000). Patterns of neuronal activity within the brain are constantly changing. Both the structure and function of axons and synapses are highly plastic and undergo spontaneous and activity-dependent remodeling, thereby changing mitochondrial mobility. In addition, mitochondrial trafficking and anchoring are tightly regulated during development for rapid redistribution into areas with increased energy or calcium buffering requirements. Thus, the mechanisms controlling the balance between the mobile and stationary phases of axonal mitochondria are possible targets of regulation. The coordination of mitochondrial mobility by axonal physiology is crucial for proper neuronal development and synaptic function. Mobile mitochondria are recruited to the stationary pool near synapses in response to elevated cytosolic Ca2+ and synaptic activity (Rintoul et al. 2003; Yi et al. 2004). Recent identification of KIF5-Milton (TRAK1/2), a mitochondrial motor-adaptor complex, and Miro, a calcium sensor that controls mitochondrial movement, provide molecular targets for such regulation (Glater et al. 2006; Saotome et al. 2008; Macaskill et al. 2009; Wang and Schwarz 2009; Cai and Sheng 2009). Understanding the mechanisms underlying the regulation of axonal mitochondrial mobility will advance our knowledge of fundamental processes that may be essential for neurons to maintain axonal and synaptic homeostasis.

142

Q. Cai and Z.-H. Sheng

5.2 Cytoskeleton Required for Axonal Mitochondrial Transport Cytoskeletal elements not only maintain the highly specialized structures of neurons but also support organelle transport and stationary docking events. The axonal cytoskeleton consists of three major components: MTs, actin, and intermediate neurofilaments (NFs). Only MTs and actin filaments play critical roles in mitochondrial transport. Long-range axonal transport of mitochondria occurs mainly along MTs. While dendritic MTs exhibit mixed polarity, axonal MTs are organized into a polar array whose ‘plus’ end is directed distally and whose ‘minus’ end is directed toward the soma. MT polarity and organization in axons are critical for the targeted transport of mitochondria from the soma to distal regions of axons and synaptic terminals. MT-associated motor proteins, including members of the kinesin superfamily and cytoplasmic dynein, are the primary motors that drive long-distance axonal transport. While most kinesin motors are plus-end directed, dynein travels toward the minus end of MTs. Therefore, kinesin motors generally mediate anterograde axonal transport, and dynein drives retrograde axonal transport of mitochondria (Fig. 5.1). Myosin-driven transport along the actin-based cytoskeleton contributes to local and short-range transport of mitochondria at nerve terminals, growth cones, and subcortical plasma membrane regions (Langford 2002). Actin filaments are enriched in several ­cellular compartments, including presynaptic terminals and dendritic spines. Actin monomers assemble into helical polymers with two distinct ends: fast- and slow-­ growing. In neurons, actin filaments bundle into a network. Although the organization of these structures in nerve terminals has not been fully elucidated, the orientation of the filaments can directly affect the activity of actin-based myosin motors (Bridgman 2004). The physical interaction between MT- and actin-based motors suggests that mitochondrial transport can be coordinated along both of these cytoskeletal systems (Bridgman 2004). A coupled “dual transport” model was proposed in which MT-based motors ensure long-range axonal transport, while actin-based myosin motors allow for local and short-range movement. It was reported that axonal mitochondria are anchored to MTs, actins, or NFs directly or via docking adaptors such as syntaphilin (Kang et al. 2008). Association of mitochondria with NFs has been suggested by ultrastructural observations (Hirokawa 1982) and in  vitro binding experiments (Wagner et  al. 2003). Ultrastructural studies have also shown structural links between mitochondria and MTs (Lindén et al. 1989a, b; Leterrier et al. 1994). Thus, the cytoskeleton serves not only as the tracks for transportation, but also as docking platforms.

5.3 Motor Proteins Required for Axonal Mitochondrial Transport Long-distance fast transport of axonal mitochondria depends on MT-based kinesin and dynein motor protein families, which use the energy of ATP hydrolysis to ­generate movement (Hollenbeck 1996). While cytoplasmic dynein motors are the

5  Mitochondrial Dynamics and Axonal Transport

143

Fig. 5.1  Axonal mitochondrial transport. In axons, microtubules are uniformly organized with the plus (+) ends facing toward the axonal terminals and the minus (−) ends toward the cell body. However, the organization of microtubules in dendrites shows mixed orientation. Polarity and organization of microtubules in axons are critical for the targeted transport of synaptic cargoes and organelles by microtubule-associated motor proteins. While kinesin motors are mostly plus-end directed, dynein travels toward the minus ends of microtubules. Therefore, kinesin motors generally mediate anterograde axonal transport of mitochondria and dynein drives retrograde axonal transport of mitochondria. At nerve terminals, mitochondria undergo short-range movement driven by actin-based myosin motors

driving force behind retrograde movement toward the soma (Pilling et  al. 2006), kinesin motors are responsible for anterograde transport of mitochondria to distal regions of axons and synaptic terminals (Fig. 5.1) (Hurd and Saxton 1996; Tanaka et al. 1998; Stowers et al. 2002; Cai et al. 2005; Glater et al. 2006).

5.3.1 Kinesin Motors and Anterograde Mitochondrial Transport Kinesin superfamiliy proteins (KIFs) comprise a large family with at least 45 different genes in humans and mice (Hirokawa and Takemura 2004). Kinesin-1 was the first motor identified to drive plus end-directed transport in  vitro (Vale et  al. 1985; Hirokawa et al. 1991). Kinesin-1 contains two heavy chains (KHCs) and two light chains (KLCs) (Fig. 5.2a). The heavy chains, known as KIF5s, form homo- or heterodimers through the coiled-coil regions in their stalk domains and can function

144

Q. Cai and Z.-H. Sheng

Fig.  5.2  Structure of motor proteins. (a) KIF5 motors form homodimers among themselves through the coiled-coil region in the stalk domains. While KIF5 itself possesses motor function, it also binds to the light chain of kinesin-1 (KLC) through its domain located at the stalk and tail regions. The specific association of KIF5 with cargoes or organelles can be mediated either directly through the cargo-binding region in its tail domain, or indirectly via the COOH-terminal domains of KLC, indicating the existence of two forms of KIF5 motor-cargo coupling. (b) Cytoplasmic dynein. Cytoplasmic dyneins consist of heavy chains (DHC), light intermediate chains (DLIH), intermediate chains (DIC) and light chains (DLC). To transport cargos, cytoplasmic dynein needs to bind with the dynactin complex (not shown). (c) Myosin motor. Myosin V consists of two heavy chains

as motors. Drosophila have only one KIF5 motor isoform, while mammals have three: KIF5A, -B and -C. While KIF5B is present in almost all cell types, KIF5A and C are neuron-specific motors (Kanai et al. 2000, also see review by Hirokawa and Takemura 2005). Each KIF5 heavy chain contains an amino terminal motor domain that binds directly to MTs, whereas its carboxyl terminal domain, named

5  Mitochondrial Dynamics and Axonal Transport

145

the cargo-binding domain, mediates the association of KIF5 with KLC or directly interacts with mitochondrial adaptor proteins. Thus, there are two distinct forms of motor-mitochondria coupling: kinesin-1 can either attach directly to mitochondrial adaptor proteins through its cargo-binding domain or the coupling can occur indirectly via KLC. Recent study, however, indicates that KIF5 motors drive mitochondrial transport independent of its KLC binding (Glater et al. 2006). KIF5 has a prominent role in the anterograde transport of axonal mitochondria (Hurd and Saxton 1996; Tanaka et al. 1998; Stowers et al. 2002; Cai et al. 2005; Pilling et al. 2006; Glater et al. 2006). Targeted disruption of kif5b results in perinuclear clustering of mitochondria instead of their being transported throughout the cytoplasm and toward the cell periphery in undifferentiated extra-embryonic cells (Tanaka et al. 1998). Drosophila kif5 mutants exhibit impaired mitochondrial transport and reduced mitochondrial distribution in larval motor axons (Pilling et al. 2006). Biochemical and imaging analyses show that KIF5 motors are associated with brain mitochondria (Hirokawa et al. 1991; Cai et al. 2005; Pilling et al. 2006; Macaskill et  al. 2009; Wang and Schwarz 2009). Hippocampal neurons expressing the KIF5 cargo-binding domain exhibit impaired axonal mitochondrial transport and reduced mitochondrial density in distal axons (Cai et al. 2005), further highlighting the role of KIF5 motors in driving anterograde transport of axonal mitochondria. KIF1Ba, a member of the kinesin–3 family enriched in the brain, is also reported to interact directly with mitochondria (Nangaku et al. 1994). KIF1Ba can transport purified mitochondria along MTs in  vitro at a velocity of 0.5  mm/s. Although the mutation of kif1B leads to peripheral neuropathies in mice, a role for KIF1Ba in anterograde mitochondrial transport is still unclear.

5.3.2 Dynein Motors and Retrograde Mitochondrial Transport Cytoplasmic dynein is the principal motor driving retrograde transport in axons and is a multisubunit complex composed of two heavy chains (DHC) and several intermediate (DIC), light intermediate (DLIC) and light chains (DLC). These polypeptides are thought to mediate the association of the dynein motor with cargoes or to regulate motility (Fig. 5.2b). Cargoes attach to the dynein motor with a degree of specificity in neurons. Relative to the Kinesin superfamily, few dynein heavy chains have been identified. Thus, the selectivity of specific cargoes likely depends on a large number of accessory proteins, including DLCs and dynactin (Susalka and Pfister 2000). Dynactin is a large complex composed of 11 different subunits and can bind directly to cytoplasmic dynein and MTs through its p150Glued component. The dynactin complex is not essential but can enhance the processivity of the dynein motor, mediate interactions with some cargoes (Waterman-Storer et  al. 1995; Karki and Holzbaur 1999; King and Schroer 2000), and coordinate bidirectional axonal transport (Haghnia et  al. 2007). Studies on single dynein–dynactin motor complex reveal that the complex can move bidirectionally, although there is

146

Q. Cai and Z.-H. Sheng

a directional bias towards the microtubule minus end (Mallik et al. 2005). This ability may enable the dynein motor to bypass obstacles during transport within the cell. The function of the dynein–dynactin complex is extremely important in motor neurons, as this neuronal population is the most vulnerable to defects in dynein function (Hafezparast et al. 2003; Puls et al. 2003; Levy et al. 2006). In the Drosophila nervous system, dynein and dynactin components associate with purified mitochondria, and cytoplasmic dynein is essential for mediating mitochondrial retrograde transport (Pilling et al. 2006). Mutations of the dynein heavy chain and dynactin p150Glued disrupt fast organelle transport in both directions, resulting in axonal swellings composed of retrograde and anterograde cargoes including mitochondria, a phenotype similar to those caused by kinesin mutations (Martin et al. 1999). In addition, expressing a mutant p150Glued causes mitochondrial accumulation within the cell body (Levy et al. 2006). Given that axonal mitochondria display bidirectional movement and that dynein has been observed co-localized with both anterogradely and retrogradely moving axonal mitochondria (Hirokawa et  al. 1990), it is likely that two opposing kinesin and dynein motors coordinate mitochondrial transport in axons. Emerging lines of evidence indicate that motor adaptors may mediate functional coupling of two opposing motors on individual mitochondria. APLIP1, a Drosophila homologue of mammalian kinesin-interacting JIP-1/JNK Scaffold Protein, is implicated in retrograde mitochondrial transport (Horiuchi et al. 2005). A recent work also demonstrates that a Drosophila mitochondria adaptor protein, dMiro, plays a role in regulating both anterograde and retrograde mitochondrial transport in axons (Russo et al. 2009). Thus, it will be interesting to determine if there is regulatory crosstalk between the two opposing motors or their adaptors and how these motoradaptor complexes help coordinate the overall movement and distribution of axonal mitochondria in response to changes in synaptic activity and neuronal physiology.

5.3.3 Myosin Motors and Local Short-Range Mitochondrial Movement While kinesins and dyneins quickly transport cargoes and organelles along lengthy axons, myosins drive short-distance trafficking along actin filaments within presynaptic terminals and growth cones. Although there is no direct evidence of a role for myosins in neuronal actin-based mitochondrial transport, actin-dependent mitochondrial mobility is well characterized in plant cells and Aspergillus, a filamentous fungus. Myosin V-associated cargoes and mitochondria moving along axons exhibit similar transport velocities (Morris and Hollenbeck 1995; Bridgman 2004; Brown et al. 2004), which are slower than those of other axonal vesicles and organelles (Cheney et al. 1993; Wolenski et al. 1995). The movement of mitochondria along actin filaments supports the idea that myosin motors may also drive shortrange mitochondrial transport within certain regions of axons, particularly at nerve terminals where actin filaments are relatively enriched (Fig. 5.1).

5  Mitochondrial Dynamics and Axonal Transport

147

A recent study from cultured Drosophila neurons showed that myosin V depletion increases mitochondrial mean velocity in both anterograde and retrograde directions. Myosin VI loss-of-function selectively increases retrograde transport, while knocking down myosin II does not affect mitochondrial transport in axons. These findings provide evidence that myosin V and VI, but not myosin II, modulate axonal mitochondrial transport (Pathak et al. 2010). Myosin V can interact directly with kinesin motors to form a hetero-motor complex or with the 8-kDa dynein light chain (Naisbitt et  al. 2000), which raises the possibility that these dual-motor ­complexes may coordinate mitochondrial long-range transport along MTs and short-range movement on actin filaments. Alternatively, myosins may facilitate mitochondrial offloading from MT tracks or enhance their docking onto actin by competing with MT-based motors. In this scenario, myosins could increase the frequency of short pauses during long-range bidirectional transport. It is unknown how myosin V and VI attach to mitochondria, how they regulate MT-based motor activity, and whether both of these myosin motors are required for synapse-directed mitochondria transport.

5.4 Motor Adaptors Essential for Mitochondrial Transport 5.4.1 Kinesin Motor Adaptors Anterograde axonal transport is mediated by kinesin motors that associate with their cargoes via adaptor proteins (or receptors). To preserve cargo identity and enable targeted trafficking, cargo vesicles must attach to their transport motors with a degree of specificity. KIF5s, key motors for driving anterograde mitochondrial movement, may attach directly to the mitochondrial membrane or indirectly via motor adaptors that either bind lipids or have transmembrane domains. Identification of mitochondrial receptors or motor–adaptor complexes is critical to understand the mechanisms regulating mitochondrial transport and distribution in neurons. 5.4.1.1 Milton and Miro The Drosophila protein Milton is a well-characterized adaptor for mitochondrial transport in neurons. Milton attaches indirectly to mitochondria via an interaction with Miro, a mitochondrial outer membrane protein. The milton mutation in Drosophila results in the depletion of mitochondria at synaptic terminals and axons (Stowers et al. 2002). Miro is a member of the mitochondrial Rho-GTPase family, has two EF hand Ca2+-binding domains (Fransson et al. 2003), and is required for the association of Milton and KIF5 with mitochondria (Fig.  5.3a) (Glater et  al. 2006). Mutation of the dmiro gene in Drosophila disrupts anterograde mitochondrial transport and prevents mitochondria from entering the axon and synapses,

148

Q. Cai and Z.-H. Sheng

Fig. 5.3  KIF5 motor adaptors for mitochondrial transport. (a) Milton-Miro adaptor complex. Miro is a member of the mitochondrial Rho-GTPase family and a mitochondrial outer membrane protein. Milton attaches indirectly to mitochondria via an interaction with Miro and recruits KIF5 to mitochondria independently of KLC. Two mammalian Milton orthologues, TRAK1 and TRAK2 can form complexes with the two mammalian orthologues of dMiro (Miro1 and Miro2). (b) Syntabulin is a KIF5 adaptor and targets to mitochondria via its carboxyl-terminal transmembrane domain, thus linking KIF5 to mitochondria and mediating mitochondrial anterograde transport. In addition, FEZ1, RanBP2, and caytaxin were reported to serve as candidate kinesin motor adaptors contributing to mitochondrial transport

which impairs neurotransmitter release and Ca2+ buffering during prolonged stimulation (Guo et  al. 2005). Furthermore, biochemical and genetic evidence demonstrates that the recruitment of KIF5 to mitochondria occurs independently of KLC, which instead inhibit KIF5 binding to Milton (Glater et al. 2006). Two mammalian Milton orthologues, TRAK1 and TRAK2 (also known as OIP106 and OIP98/Grif-1), can form complexes with the two mammalian orthologues of dMiro (Miro1 and Miro2) (Fransson et al. 2003; Macaskill et al. 2009). There are several functional similarities and differences between Milton and the two TRAKs. In Drosophila, Milton acts as a mitochondrial adaptor and links Miro to KIF5 (Fig.  5.3a) (Glater et  al. 2006; Wang and Schwarz 2009). In contrast, the TRAK proteins, which share 33.5% and 35.3% homology with Milton, respectively, are involved in the intracellular transport of other cargoes and organelles, in addition to mitochondria (Grishin et al. 2006; Kirk et al. 2006; Webber et al. 2008). Although TRAKs are important for mitochondrial trafficking (Smith et  al. 2006; Macaskill et al. 2009), it is unclear if they functions as a bridge linking Miro to KIF5 or, alternatively, act as regulatory components for motor activity (Macaskill et al. 2009). 5.4.1.2 Syntabulin Syntabulin, as an adaptor linking syntaxin-containing vesicles to KIF5 (Su et al. 2004), plays an essential role in the axonal transport of the presynaptic active

5  Mitochondrial Dynamics and Axonal Transport

149

zone components and is necessary for presynaptic assembly and activity-dependent plasticity in developing neurons (Cai et  al. 2007). In addition, syntabulin targets to mitochondria via its carboxyl-terminal transmembrane domain, thus also linking KIF5 to mitochondria and playing a critical role in the anterograde transport of mitochondria (Cai et al. 2005). Knocking down syntabulin or blocking the syntabulin–KIF5 interaction with binding-domain transgenes dramatically changes mitochondrial distribution in cultured hippocampal neurons. In this experiment, mitochondria were predominantly clustered in the soma but distributed sparsely in the processes. This distribution profile was consistent with observations from mobility analysis in live neurons: syntabulin loss-of-function reduces anterograde, but not retrograde, transport of mitochondria along axonal processes. This loss-of-function phenotype suggests that syntabulin serves as an adaptor linking KIF5 to mitochondria, thus mediating mitochondrial anterograde transport (Fig. 5.3b).

5.4.1.3 Other Kinesin Adaptors In addition to Milton/Miro and syntabulin, several other molecules have been proposed as candidate adaptors for kinesin-1 required for mitochondrial transport (Fig.  5.3b). Fasciculation and elongation protein zeta-1 (FEZ1), a mammalian orthologue of the Caenorhabditis elegans UNC-76 protein, is a brain-specific coiled-coil protein involved in axonal outgrowth (Kuroda et al. 1999). In Drosophila, the FEZ1 orthologue UNC-76 associates with the kinesin motor enabling axonal transport of vesicles and organelles (Gindhart et al. 2003). It was recently reported that the association of FEZ1 with the c-Jun N-terminal kinase (JNK)-interacting protein (JIP1) efficiently activates kinesin motor activity (Blasius et  al. 2007). Moreover, FEZ1 was shown to be essential for the anterograde transport of mitochondria along hippocampal axons (Ikuta et al. 2007), possibly because it stabilizes motor-MT binding and/or regulates motor activity. The hereditary ataxia protein caytaxin was reported to associate with kinesin-1 through a direct interaction with KLC (Aoyama et al. 2009). It localizes to both axons and dendrites, where it partially co-localized with mitochondria, and is required for the proper distribution of mitochondria in distal neurites. Caytaxin may serve as a scaffold for signaling proteins in the phosphatidylinositol-calcium signaling pathways and perform similar functions to those of JIPs, which bind to KLC and facilitate the attachment of various cargoes to KLC for transport by kinesin-1. RanBP2 is a large mosaic protein abundant in the ellipsoid compartment of photoreceptors. It co-localizes and interacts directly with KIF5B and KIF5C, but not KIF5A (Mavlyutov et al. 2002). The association of the KIF5-binding domain of RanBP2 to KIF5B and KIF5C regulates mitochondrial transport in non-neuronal cells (Cho et al. 2007). Little is known, however, about the potential role for this protein in the regulation of axonal mitochondrial transport.

150

Q. Cai and Z.-H. Sheng

5.4.2 Dynein Motor Adaptors Relative to the kinesin-adaptor transport machinery that controls anterograde mitochondrial transport, the underlying mechanisms which link the dynein motor to mitochondria are less characterized. Although dynactin is thought to link dyneins to cargo membranes (Karki and Holzbaur 1999; Schroer 2004), a recent study highlights a potential role for dynactin in coordinating the activity of opposing motors (Haghnia et al. 2007). An alternative model suggests that DLCs and DICs may link dyneins to membrane-associated proteins independent of dynactin (Tai et al. 1999; Tynan et al. 2000). Tctex 1, a dynein light chain, associates with the mitochondrial outer membrane, probably via an interaction with the mitochondrial voltage-dependent anion-selective channel 1 (Schwarzer et  al. 2002). This finding raises a possibility for its role in attaching dynein motors to mitochondria. Although APLIP1 is a kinesin-binding protein, APLIP1 mutation selectively reduces retrograde mitochondrial transport (Horiuchi et al. 2005). A recent study indicates a mechanism by which an activated JNK pathway regulates axonal transport by disrupting APLIP1-kinesin-1 coupling (Horiuchi et al. 2007). Thus, it is necessary to determine whether vertebrate APLIP orthologues, JIP1-3s, play a role in the retrograde transport of mitochondria in axons. Interestingly, Miro, a KIF5 adaptor for mitochondria, is required for both kinesin-driven anterograde and dynein-mediated retrograde transport of mitochondria in Drosophila axons (Russo et al. 2009). These observations suggest that Miro may serve as a general motor adaptor for both KIF5 and dynein, although the association of Miro with dynein has not been demonstrated.

5.5 Docking Mechanisms Controlling Mitochondrial Mobility In mature axons, approximately two-thirds of the total mitochondrial population remains stationary at any given time (Hollenbeck 1996). Stationary mitochondria may reflect disassociation of motor proteins from the MT track. A growing body of evidence suggests that static anchors serve as effective cytoskeletal attachment points for mitochondria. Docking receptors anchor mitochondria to MTs, actin and neurofilaments at specific locations on axons where energy production and calcium buffering capacity are in high demand. Mitochondrial docking mechanisms may enable neurons to maintain proper densities of stationary mitochondria within axons and near synapses. Considering the limited diffusion of ATP in an intracellular environment (Belles et al. 1987; Hubley et al. 1996), docked stationary mitochondria likely serve as local energy stations and provide the ATP needed to maintain the high activity of Na+–K+ ATPase and fast spike propagation.

5  Mitochondrial Dynamics and Axonal Transport

151

To achieve the desired balance between mobile and stationary mitochondria in axons, neurons utilize special mechanisms to recruit mobile mitochondria to the stationary pool in response to changes in neuronal activity and synaptic modification. Thus, identifying proteins that can mediate mitochondrial docking and facilitate their retention within axons and at synapses will provide new molecular targets by which to regulate axonal mitochondrial distribution.

5.5.1 Syntaphilin-Mediated Docking of Axonal Mitochondria Using genetic mouse models combined with time-lapse imaging analysis, a recent study has identified syntaphilin (SNPH) as a “static anchor” for axonal mitochondria (Kang et al. 2008). SNPH is a neuron-specific axonal mitochondria-targeted protein (Fig.  5.4a). SNPH associates with mitochondria via its carboxyl-terminal tail. Intriguingly, endogenous SNPH associates with 65 ± 14% of axonal mitochondria, which strongly correlates to the proportion of stationary axonal mitochondria (62 ± 15%). SNPH-mediated immobilization of axonal mitochondria occurs when it anchors to the MT-based cytoskeleton. Deletion of the snph gene in mice dramatically increases mitochondrial mobility and decreases the density of mitochondria within axons (Fig. 5.4b). The reduced mitochondrial density also reflects a diminished interbouton mitochondrial density in the snph (−/−) neurons relative to wild-type controls. This study identifies SNPH as an anchoring receptor for axonal mitochondrial docking (Fig. 5.4c). Such a mechanism enables neurons to maintain proper densities of stationary mitochondria within axons and near synapses. A subsequent study further suggests that the mechanism underlying SNPH-mediated docking of axonal mitochondria can be further regulated by the dynein light chain LC8 (Chen et al. 2009). LC8 regulates axonal mitochondrial mobility by binding to SNPH, thus enhancing the SNPH-MT docking interaction. These findings provide new mechanistic insights into the control of mitochondrial mobility through the dynamic interaction between mitochondrial docking receptors and the axonal cytoskeleton. Identification of SNPH as a docking protein provides a molecular target for further investigation into how mobile mitochondria are recruited to the stationary pool in response to changes in axonal activity and synaptic modification. The snph mutant mice also provide a unique model for studying the physiological impact of mitochondrial docking and retention on synaptic transmission, axonal homeostasis, and axonal degeneration. It is expected that defective mitochondrial docking/ anchoring could affect neuronal function, particularly in motor neurons due to their long axonal processes. There is mounting evidence that defective mitochondrial trafficking is involved in the pathology of several neurodegenerative diseases. Future studies which cross snph mice with these disease mouse models may augment the turnover of dysfunctional mitochondria by increasing their transport to the soma where they may be repaired or degraded.

152

Q. Cai and Z.-H. Sheng

Fig.  5.4  SNPH acts as a receptor for docking/anchoring axonal mitochondria. (a) Axonal mitochondrial targeting of SNPH in cultured hippocampal neurons. Hippocampal neurons at DIV14 were co-immunostained for SNPH and mitochondrial marker Cytochrome c. Arrows point to SNPH-associated mitochondria and arrowheads indicate SNPH-negative mitochondria within an axonal process. Scale bars, 10 mm. (b) SNPH immobilizes axonal mitochondria and deletion of snph gene in mice robustly increases mobility of axonal mitochondria. Mobility of axonal mitochondria was observed in live neurons one week after transfection. Motion data are presented in kymograph, in which vertical lines represent stationary mitochondria and slant lines or curves indicate motile ones. Left panel: wild-type neurons co-transfected at DIV6 with DsRed-mito (red) and GFP-SNPH (green); middle panel: wild-type neurons transfected with DsRed-Mito alone; right panel: snph (−/−) neurons transfected with DsRed-Mito alone. (c) Schematic diagram for the proposed role of SNPH. SNPH acts as a receptor for docking/anchoring mitochondria in axons

5  Mitochondrial Dynamics and Axonal Transport

153

5.5.2 Actin and Neurofilament-Based Mitochondrial Docking In addition to MTs, actin and neurofilaments have been shown to be responsible for anchoring axonal and dendritic mitochondria. Actin is relatively enriched in neuronal subdomains where ATP production is in high demand such as growth cones, presynaptic terminals, and dendritic spines. Additionally, neuronal activity has been shown to induce mitochondrial retention within dendritic spines in an actin-dependent manner (Li et al. 2004; Sung et al. 2008; Macaskill et al. 2009). In axons, disrupting the actin cytoskeleton and actin-based myosin motors results in increased mitochondrial movement (Ligon and Steward 2000; Morris and Hollenbeck 1995; Pathak et  al. 2010). NGF-dependent local recruitment of mitochondria in axons also requires actin, probably via the actin effector WAVE1 (Chada and Hollenbeck 2003, 2004; Sung et al. 2008). Lysophosphatidic acid (LPA), a growth factor present in serum, inhibits fast transport of mitochondria along the actin cytoskeleton by activating RhoA, a small GTPase (Minin et al. 2006). Therefore, actin-dependent mitochondrial tethering may be coupled to signals that recruit and retain mitochondria along actin filaments or enhance their association with actin-based transport machinery. In non-neuronal cells or in vitro systems, mitochondria are tethered to neurofilaments and neuron-specific intermediate filaments (Toh et al. 1980; Winter et al. 2008). Interestingly, the ability of mitochondria to anchor to neurofilaments depends upon the mitochondrial membrane potential. Mitochondria with high membrane potentials are more likely to bind to neurofilaments, whereas inducing mitochondrial depolarization by FCCP treatment releases mitochondria from neurofilaments (Wagner et al. 2003). These observations highlight an attractive model in which healthy mitochondria are anchored stably along neurofilaments, while dysfuntional ones are transported to the soma for recycling.

5.6 Regulation of Mitochondrial Transport 5.6.1 Activity-Dependent Regulation of Mitochondrial Transport Mitochondria produce ATP and serve as neuronal Ca2+ reserves. Neuronal mitochondria display distinct saltatory and bidirectional movement. The relative levels of mobile and stationary mitochondria are determined by fluctuations in metabolic

Fig. 5.4  (continued) and is required for maintaining a large number of axonal mitochondria in a stationary state by interacting with the microtubule-based cytoskeleton. (Images in (a) are adapted with permission from Jian-Sheng Kang, Jin-Hua Tian, Philip Zald, Ping-Yue Pan, Cuiling Li, Chuxia Deng, and Zu-Hang Sheng. Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation. Cell 132, 137–148, 2008. Images in (b) and schematic diagram in (c) are adapted with permission from Qian Cai, Zu-Hang Sheng. Mitochondrial transport and docking in axons. Experimental Neurology 218, 257–267, 2009)

154

Q. Cai and Z.-H. Sheng

demand. The depletion of local ATP, achieved by the application of glutamate, results in decreased mitochondrial velocity near synapses. Additionally, the consumption of synaptic ATP facilitates the targeting of mitochondria to synapses by increasing local ADP levels (Mironov 2007, 2009). As mitochondria often reside in subcellular regions with high metabolic demands, this mechanism may facilitate the recruitment of mitochondria to those subdomains requiring additional ATP due to high levels of synaptic activity. In addition to ATP levels, mitochondrial mobility may also be modulated in response to altered intracellular Ca2+ or neurotransmitter release (Yi et  al. 2004). In cultured neurons, mitochondrial movement increases after treatment with tetrodotoxin, which blocks action potentials, but decreases after KCl depolarization (Li et al. 2004) and Ca2+ influx through NMDA receptors (Rintoul et al. 2003). The inhibition of synaptic activity, via N-type Ca2+ and Na2+ channel blocker application, increases mitochondrial transport. The duration of intracellular Ca2+ elevation correlates with reduced mitochondrial movements (Mironov 2006). Ca2+ influx occurs at presynaptic terminals and postsynaptic dendritic spines, where mitochondria are often retained to maintain Ca2+ homeostasis by producing ATP to pump Ca2+ across the plasma membrane or by the sequestration of Ca2+ into mitochondria. Therefore, during sustained synaptic activity, neurons must utilize special mechanisms to achieve activity-dependent retention of mitochondria at synaptic terminals. How are mobile mitochondria recruited to the stationary pool in response to elevated cytosolic Ca2+ and sustained synaptic activity? Recent advances in our understanding of the KIF5–Milton–Miro complex have further elucidated the molecular mechanisms underlying the regulation of mitochondrial mobility by intracellular Ca2+ levels (Saotome et al. 2008; Wang and Schwarz 2009; Macaskill et al. 2009). Miro acts as a Ca2+ sensor via its EF hands’s ability to bind the ion. The binding of Miro to Ca2+ results in the arrest of mitochondrial mobility. The activation of glutamate receptors or increasing action potential firing rates by application of an electric field recruits mitochondria to activated synapses through a Miro-mediated Ca2+-sensing pathway. In neurons expressing a mutant Miro EF hand defective in Ca2+ binding, the Ca2+-induced cessation of mitochondrial mobility was effectively blocked. Two alternative mechanisms have been proposed to explain how Ca2+-induced suppression of mitochondrial mobility is achieved by regulating motor-adaptor coupling, thereby changing the engagement of mitochondria with microtubules (Fig. 5.5) (Cai and Sheng 2009). Wang and Schwarz (2009) proposed a motor–adaptor switch model. They show that the C-terminal tail of KIF5, in the absence of Ca2+, binds to mitochondria through an interaction with the Milton–Miro complex, allowing its N-terminal motor domain to engage microtubules and facilitate anterograde transport. Ca2+ binding to the EF hands induces a conformational change, resulting in a direct interaction between the motor domain and Miro and prevents KIF5–microtubule interaction. Thus, mitochondrial mobility is tightly regulated by synaptic activity through a Ca2+-binding switch mechanism that turns motor–microtubule engagement “on” or “off” (Fig.  5.5a). In contrast, Macaskill et  al. (2009) propose that Miro mediates mitochondrial transport by linking mitochondria to KIF5. Ca2+

5  Mitochondrial Dynamics and Axonal Transport

155

Fig. 5.5  Schematic diagram for two proposed models of Miro as a Ca2+ sensor in regulating mitochondrial mobility. Miro contains two GTPase domains and calcium-binding EF hand motifs, thus regulates axonal mitochondrial motility by either GTP hydrolysis or calcium binding in response to calcium signals and synaptic activity. (a) Model 1: Ca2+-binding turns “off” KIF5 engagement with MTs. The tail of KIF5 is linked to Miro via milton in a Ca2+-independent manner, thus leaving its motor domain to engage with MTs. Ca2+-binding to the EF hands triggers the direct interaction of the motor domain with Miro, thus preventing the motor from engaging with MTs (Wang and Schwarz 2009). (b) Model 2: Ca2+-binding detaches KIF5 from mitochondria. Mitochondrial transport is mediated by linking Miro to KIF5. Ca2+-binding to the EF hands dissociates Miro from KIF5 while KIF5-binding protein GRIF-1/TRAK2 (a mammalian homologue of Milton) remains bound to Miro1 (Macaskill et al. 2009). (The model diagram is adapted with permission from Qian Cai and Zu-Hang Sheng. Moving or stopping mitochondria: Miro as a ­traffic cop by sensing calcium. Neuron 61, 493–496. 2009)

binding to the EF hands releases KIF5 from Miro-attached mitochondria. Thus, activity-triggered Ca2+ influx could recruit mobile mitochondria to the activated synapses (Fig. 5.5b). While these models are attractive, several questions must still be answered. Miro-mediated and Ca2+-dependent suppression of mitochondrial mobility affects both anterograde and retrograde transport. Is Miro linked mechanistically to the

156

Q. Cai and Z.-H. Sheng

dynein motor complex? Complex mitochondrial mobility patterns suggest that mitochondria are linked to opposing KIF5 and dynein motors, allowing them to move bidirectionally. When kinesin-driven movement is disturbed, dynein does not simply take over. Thus, motor adaptors are likely candidates for the coordination of the activity of opposing motors bound to a single organelle. Recent identification of syntaphilin as a “static anchor” for axonal mitochondria provides another promising mechanism (Fig.  5.4c) (Kang et  al. 2008). Do the motor-adaptor complex and docking receptor share a single system of regulation? And could this regulation involve the physical displacement of the motor-adaptor complex by docking interactions (or vice versa)? What are the neuronal signals that regulate the interplay between motor-docking machineries and maintain the proper balance between the mobile and stationary pools in response to changes in neuronal activity? Future studies using genetic mouse models will provide insight into the molecular mechanisms underlying the complex regulation of mitochondrial bidirectional movement and their temporal-spatial localization in axons and near synapses.

5.6.2 Neuronal Signaling Pathways for Regulating Mitochondrial Movement Axonal mitochondria undergo anterograde transport toward the distal portions of axons and retrograde movement in the opposite direction. Tight regulation of these events allows for efficient response to changes in metabolic demands and local energy states. Mitochondria are efficiently transported into nascent axons and recruited into the stationary pools within active growth cones, branches of developing neurons, nodes of Ranvier, myelination boundaries, sites of axonal protein synthesis, and synaptic terminals (Hollenbeck and Saxton 2005). In developing neurons, mitochondria exhibit greater anterograde movement and increased density along elongating axons (Pilling et  al. 2006; O’Toole et  al. 2008). An increase in retrograde mitochondrial movement is probably responsible for the significantly reduced mitochondrial density observed during the cessation of axonal growth (Morris and Hollenbeck 1993; Ruthel and Hollenbeck 2003). The direction of mitochondrial transport is closely linked to mitochondrial membrane potential, which is heterogeneous and varies over time and space in response to changes in metabolic demand (Overly et al. 1996). Mitochondria moving anterogradely toward the axon terminal exhibit high membrane potentials. Conversely, those moving towards the cell body have low membrane potentials, suggesting that aged or damaged mitochondria are transported to the soma for repair (Miller and Sheetz 2004). These findings support the notion that mitochondrial fission within the soma generates healthy mitochondria that are subsequently transported along axons toward distal regions or synapses, while dysfunctional mitochondria are delivered to the soma where they undergo fusion with healthy mitochondria or are degraded through mitophagy (Chang and Reynolds 2006). A recent study, however, found no difference in membrane potential between stationary, anterogradely and

5  Mitochondrial Dynamics and Axonal Transport

157

retrogradely moving mitochondria (Verburg and Hollenbeck 2008). NGF and semaphorin signaling can effectively elicit local receptor-mediated increases in mitochondrial membrane potential within growth cones of chick sensory neurons. These data suggest that the membrane potential of axonal mitochondria can, like their distribution, be regulated locally, highlighting an important role for cellular signaling in the regulation of neuronal mitochondrial membrane potential. NGF serves as a docking signal that immobilizes axonal mitochondria in dorsal root ganglion (DRG) neuron cultures. This docking process probably occurs on F-actin filaments and depends upon the PI3K phosphorylation pathway (Chada and Hollenbeck 2004; Reynolds and Rintoul 2004). In addition, serotonin (5-HT) promotes axonal mitochondrial transport by activating the 5-HT1A receptor and Akt (Chen et  al. 2007). In contrast, dopamine inhibits mitochondrial mobility in hippocampal neurons through the same signaling cascade (Chen et al. 2008). As Akt activity is regulated by PI3K, NGF and serotonin/dopamine signaling pathways likely alter mitochondrial transport via a conserved regulatory mechanism under the control of PI3K. Nitric oxide (NO) has several physiological and pathophysiological effects in the nervous system and can modulate mitochondrial mobility. Elevated NO levels are associated with reduced mitochondrial membrane potentials, which result in rapid termination of mitochondrial movement (Rintoul et al. 2006). Hypoxia is also found to impair mitochondrial movement via NO-mediated signaling pathways in cortical neurons (Zanelli et al. 2006). Mitochondria have been identified as targets of the neurotoxic actions of zinc. Elevated intraneuronal zinc impairs mitochondrial trafficking in neurons, suggesting that mitochondrial transport may be closely coupled to neuronal viability (Malaiyandi et al. 2005). Mitochondrial dynamics involve movement at two levels: organelle transport and membrane fusion/fission. Thus, regulation of mitochondrial transport can also be achieved by altering mitochondrial membrane fusion/fission events. The mitochondrial fusion process requires the apposition of two adjacent mitochondria, presumably via transport machinery. Moreover, organelle size affects mitochondrial mobility. Thus, the coordination of mitochondrial transport and membrane fusion/fission events in neurons likely involves similar mechanisms. The mutation of the mitochondrial protein Drp1, a dynamin-like GTPase, impairs axonal transport and synaptic targeting of mitochondria by interfering with the fission process (Verstreken et al. 2005). A recent study shows that neurons with deleted mitofusin 2 (Mfn2), an outer mitochondrial membrane protein involved in mitochondrial fusion, exhibit impaired axonal mitochondrial transport (Misko et al. 2010). Mfn2 disruption alters mitochondrial transport in both anterograde and retrograde directions. The mutations of Mfn2 have been identified as a common cause of Chaecot-Marie-Tooth type 2, a dominantly inherited disease characterized by the degeneration of peripheral sensory and motor axons. Expressing mutated Mfn2 in neurons significantly impairs mitochondrial transport in axons (Baloh et al. 2007). Therefore, the crosstalk between membrane fusion proteins and motor transport machinery is likely through the interaction between Mfn2 and the Miro/Milton complex (Misko et al. 2010).

158

Q. Cai and Z.-H. Sheng

Miro contains two atypical GTPase domains flanking its Ca2+-binding EF hand motifs (Fransson et al. 2003), Ca2+-regulated transport is likely coordinated by GTP hydrolysis-mediated fission. Miro enhances mitochondrial fragmentation (or fission) by its Ca2+-sensing and GTPase activity, thus controlling mitochondrial mobility and density in dendrites (Saotome et  al. 2008). That Miro has duel functions is quite interesting. Fission defects result in long, tubular mitochondria that are less mobile than short tubular or vesicular structures.

5.7 The Impact of Axonal Mitochondria Mobility on Synaptic Function and Axonal Degeneration 5.7.1 Role of Kinesin Motor Adaptors in Synaptic Plasticity It is well documented that proper distribution of mitochondria, particularly at synapses, is required to maintain normal neurotransmission by buffering calcium levels and producing ATP during intense stimulation. Presynaptic mitochondria maintain neurotransmission by accelerating recovery from synaptic depression after a period of moderate activity at mammalian central synapses (Billups and Forsythe 2002). Mitochondria maintain calcium homeostasis at some synapses by sequestering excess intracellular Ca2+ during tetanic stimulation and by releasing calcium after stimulation (Jonas 2006). Drosophila dMiro, a mutant that causes defects in axonal mitochondrial transport, results in the loss of mitochondria from neuromuscular junctions and contributes to the observed reduction of Ca2+ buffering and impaired neurotransmitter release during prolonged stimulation (Guo et  al. 2005). Mitochondrial ATP production is also necessary to mobilize synaptic vesicles from the reserve pool during intense synaptic activity. In drp1 mutant Drosophila, the loss of mitochondria from neuromuscular junctions results in faster synaptic vesicle depression during prolonged pulse train stimulation (Verstreken et  al. 2005). Photoreceptors expressing mutant Milton show aberrant synaptic transmission due to a reduced distribution of mitochondria at synapses (Stowers et al. 2002). Using cultured superior cervical ganglion neurons combined with gene manipulation, a recent study shows that syntabulin-mediated and KIF5-driven mitochondrial transport has critical roles in the maintenance of presynaptic function (Ma et  al. 2009). Syntabulin loss-of-function delays the initiation of synaptic activity in developing neurons and impairs synaptic transmission in mature neurons, as demonstrated by reduced basal activity, accelerated synaptic depression during high-frequency firing, slowed recovery rates after synaptic vesicle depletion, and impaired presynaptic short-term plasticity. These defects are associated with reduced mitochondrial density along neuronal processes and could be rescued by the application of ATP within presynaptic neurons. These results suggest that syntabulin assists in the axonal transport of mitochondria and concomitant ATP production within presynaptic terminals. ATP supply from locally stationed ­mitochondria is necessary for the efficient mobilization of synaptic vesicles into

5  Mitochondrial Dynamics and Axonal Transport

159

the readily releasable pool. These findings emphasize the critical role of axonal mitochondrial transport in the maintenance of presynaptic function and regulation of synaptic plasticity.

5.7.2 Role of Docking Protein Syntaphilin in Short-Term Presynaptic Plasticity A fundamental question is whether changes in axonal mitochondrial mobility or docking state have an impact on synaptic transmission. Identification of syntaphilin (SNPH) as an axonal mitochondrial docking protein provides a genetic model to address this issue. Taking advantage of the unique phenotypes of snph (−/−) neurons, Kang et al. (2008) examined synaptic physiology using dual whole-cell patchclamp recordings of paired hippocampal neurons in culture. The increased mobility and reduced density of axonal mitochondria induced by deleting the snph gene in neurons does not significantly impact basal synaptic transmission, as measured by the frequency and amplitude of mini AMPA events and averaged amplitude and kinetics of EPSCs. During the application of short stimulus trains (20 Hz, 1 s) to the presynaptic neuron at 10-second intervals, however, persistent enhanced facilitation was reproduced in the snph (−/−), not in (+/+), neurons under repetitive pulse train stimulation (Fig.  5.6a). The observed phenotype could be fully rescued by re-introducing the snph gene into mutant neurons, confirming that the enhanced short-term facilitation was due to SNPH deficiency. The deletion of the snph gene results in enhanced short-term facilitation but does not significantly affect basal synaptic transmission. One explanation for these observations is that a rapid buildup of intracellular Ca2+ at presynaptic terminals may occur during intense stimulation. Testing this hypothesis, Kang et al. (2008) further demonstrate that disruption of the docking mechanism changes global Ca2+ dynamics at presynaptic boutons during intense and prolonged stimulation probably through a reduction in mitochondrial Ca2+-buffering capacity or an impaired ability of the terminals to pump Ca2+ out of the cell due to insufficient ATP supply (Fig.  5.6b). Altogether, these electrophysiological and calcium imaging studies, combining genetic and cell biological approaches, provide the first direct evidence that manipulating axonal mitochondrial docking impacts short-term presynaptic plasticity by affecting calcium dynamics at nerve terminals.

5.7.3 Defects of Mitochondrial Transport and Neuronal Degeneration Mitochondria are essential organelles for neuronal survival and function by ­producing ATP and buffering Ca2+. Due to their highly polarized morphology, neurons require specialized mechanisms to regulate mitochondrial transport along axons as well as retain them in active growth cones, nodes of Ranvier, and synaptic

160

Q. Cai and Z.-H. Sheng

Fig.  5.6   Role of docking protein syntaphilin in short-term presynaptic plasticity. (a) Deletion of the snph gene results in sustained synaptic short-term facilitation. 20  Hz, 1  s stimulus train was delivered repetitively six times at 10-second intervals (upper panel). Normalized EPSC amplitudes were plotted against stimulus number (lower panel). Note that persistent facilitation in synaptic responses was shown only in the snph (−/−) neurons (red circles). Reintroducing the snph gene into the mutant presynaptic neuron (purple circles) eliminates the short-term facilitation and fully rescues the (+/+) phenotype (blue triangles). (b) Presynaptic calcium dynamics in the snph (+/+) and snph (−/−) neurons. Upper panel: representative calcium images at presynaptic boutons of the snph (+/+) and (−/−) neurons before stimulation (1), at the beginning (2) and the end (3), and after stimulation (4). Calcium transients within presynaptic boutons labeled with DsRed-monomer-synaptophysin were imaged using Fluo-4NW at 50-ms intervals upon stimulation at

5  Mitochondrial Dynamics and Axonal Transport

161

terminals, where energy production and calcium homeostasis are critical. In some motor neurons, mitochondria are required to travel up to one meter from the cell body to their destinations. Thus, defects in mitochondrial transport could cause local energy depletion and toxic changes in Ca2+ buffering, thus triggering synaptic dysfunction and loss. In addition, mitochondrial morphology and membrane dynamics, required for maintaining mitochondrial function, depend on proper mitochondrial mobility in neurons (Chen and Chan 2009). Altered transport and aberrant distribution of mitochondria along axons has been implicated in the pathologic processes of neurodegenerative diseases such as Alzheimer’s, Huntington’s and Amyotrophic lateral sclerosis (ALS) (Stamer et al. 2002; Pigino et al. 2003; Trushina et al. 2004; Chan 2006; Chang and Reynolds 2006; ChevalierLarsen and Holzbaur 2006; Orr et  al. 2008; Magrané and Manfredi 2009). However, it has not been established whether altered mitochondrial mobility and distribution plays a critical role in axonal degeneration or is merely a side effect of general transport defects or a downstream response to primary pathogenic changes. Further characterization of mitochondrial transport in mature neurons from aged neurodegenerative disease mouse models is an important step in understanding the cellular mechanisms of neurodegeneration. ALS is a late-onset neurodegenerative disease that causes motor neuron loss (Cleveland and Rothstein 2001). Mutations in Cu/Zn superoxide dismutase (SOD) genes have been found to cause familial ALS (Rosen et al. 1993). Transgenic mice expressing mutant human SOD1 are clinically and pathologically similar to human ALS patients (Gurney et al. 1994), becoming paralyzed in one or more limbs due to a loss of motor neurons in the spinal cord. Mutant human SOD1 accumulates in mitochondria of the brain, spinal cord, and motor neurons. Several laboratories have reported altered transport of axonal mitochondria in ALS patients and mutant SOD1 mice, as well as neurons and motor neuron NSC34 cell lines expressing SOD1G93A (Sasaki and Iwata 1996; De Vos et al. 2007; Magrané et al. 2009). The presence of metabolically dysfunctional mitochondria at distal axons could be a consequence of impaired recycling or degradation of abnormal mitochondria due to altered axonal transport. To slow the rate of axonal degeneration, it is proposed that increased mitochondrial transport might aid the efficient delivery of healthy mitochondria to axons and/or the removal of damaged mitochondria from distal

Fig. 5.6  (continued) 10 Hz for 10 s. The images are pseudocolored, with blue representing low [Ca2+] concentration and red representing high [Ca2+] concentration. Scale bars: 1  mm. Lower panel: time course of changes in presynaptic fluorescent intensity over baseline (∆F/F0) from the snph (+/+) (black, n=32) and (−/−) (red, n=33) neurons. Inset: peak values of intracellular [Ca2+] levels within boutons were averaged from last 10 stimuli (20 frames of calcium imaging), expressed as % increase of fluorescence intensity over baseline (∆F/F0), and are significantly different (p<0.001, t test) between the snph (+/+) (24%±2%) and (−/−) (33% ± 2%) neurons. Error bars: s.e.m (Adapted with permission from Jian-Sheng Kang, Jin-Hua Tian, Philip Zald, Ping-Yue Pan, Cuiling Li, Chuxia Deng, and Zu-Hang Sheng. Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation. Cell 132, 137–148, 2008)

162

Q. Cai and Z.-H. Sheng

synapses for degradation via mitophagy or repair via fusion/fission. Therefore, it is important to assess whether enhanced (rescued) axonal mitochondrial mobility in disease models has a beneficial impact on the pathogenesis of motor neuron degeneration.

5.8 Conclusions Both the structure and function of axons and synapses are highly plastic and undergo spontaneous and activity-dependent remodeling, thereby changing mitochondrial mobility. The mechanisms that determine the balance between the mobile and stationary phases of axonal mitochondria are likely regulated by intracellular signaling and synaptic activity. How are mobile mitochondria recruited to the stationary pool (or vice versa) in response to changes in neuronal activity and synaptic physiology? Mitochondrial dysfunction, fragmentation and impaired transport in axons are implicated in the pathogenesis of some neurodegenerative diseases. Whether impaired mitochondrial transport plays a critical role in axonal degeneration or is a side effect of more general defective axonal transport has been a subject of debate. Does mitochondrial mobility impact the turnover of dysfunctional mitochondria through membrane fusion/fission or mitophagy processes? In particular, it will be interesting to determine whether the enhanced mobility of axonal mitochondria has a positive impact on disease pathogenesis by contributing to the efficient removal of dysfunctional mitochondria from synapses and distal axons. The recent identification of motor–adaptor transport complexes and docking machinery provide new molecular and genetic tools to address these issues. Future studies using genetic mouse models, along with live cell tracking of mitochondrial mobility in vivo, will help elucidate the molecular and cellular mechanisms regulating mitochondrial mobility, distribution and factors impacting their function and turnover in axons and near synapses. These studies will shed light on fundamental neuronal processes that may advance our understanding of human neurodegenerative disorders. Acknowledgements  We thank the members of the Sheng lab for helpful discussions and M. Davis for editing of the manuscript. The authors are supported by the Intramural Research Program of NINDS, NIH.

References Aoyama, T, Hata, S, Nakao, T, Tanigawa, Y, Oka, C, Kawaichi, M, 2009, Cayman ataxia protein caytaxin is transported by kinesin along neurites through binding to kinesin light chains, J Cell Sci 122:4177–85 Baloh, RH, Schmidt, RE, Pestronk, A, Milbrandt, J, 2007, Altered axonal mitochondrial transport in the pathogenesis of Charcot-Marie-Tooth disease from mitofusin 2 mutations, J Neurosci 27:422–30

5  Mitochondrial Dynamics and Axonal Transport

163

Belles, B., Hescheler, J, Trube, G, 1987, Changes of membrane currents in cardiac cells induced by long whole-cell recordings and tolbutamide, Pflugers Arch 409:582–8 Billups, B, Forsythe, ID, 2002, Presynaptic mitochondrial calcium sequestration influences transmission at mammalian central synapses, J Neurosci 22:5840–47 Blasius, TL, Cai, D, Jih, GT, Toret, CP, Verhey, KJ, 2007, Two binding partners cooperate to activate the molecular motor Kinesin-1, J Cell Biol 176:11–7 Bogan, N, Cabot, JB, 1991, Light and electron microscopic analyses of intraspinal axon collaterals of sympathetic preganglionic neurons, Brain Res 41:241–51 Bridgman, PC, 2004, Myosin-dependent transport in neurons, J Neurobiol 58:164–74 Bristow, EA, Griffiths, PG, Andrews, RM, Johnson, MA, Turnbull, DM, 2002, The distribution of mitochondrial activity in relation to optic nerve structure, Arch Ophthalmol 120: 791–6 Brown, JR, Stafford, P, Langford, GM, 2004, Short-range axonal/dendritic transport by myosin-V: A model for vesicle delivery to the synapse, J Neurobiol 58:175–88 Cai, Q, Gerwin, C, Sheng, ZH, 2005, Syntabulin-mediated anterograde transport of mitochondria along neuronal processes, J Cell Biol 170:959–69 Cai, Q, Pan, PY, Sheng, ZH, 2007, Syntabulin-kinesin-1 family member 5B-mediated axonal transport contributes to activity-dependent presynaptic assembly, J Neurosci 27:7284–96 Cai, Q, Sheng, ZH, 2009, Moving or stopping mitochondria: Miro as a traffic cop by sensing calcium, Neuron, 61:493–6 Chada, SR, Hollenbeck, PJ, 2003, Mitochondrial movement and positioning in axons: the role of growth factor signaling, J Exp Biol 206:1985–92 Chada, SR, Hollenbeck, PJ, 2004, Nerve growth factor signaling regulates motility and docking of axonal mitochondria, Curr Biol 14:1272–76 Chan, DC., 2006, Mitochondria: dynamic organelles in disease, aging, and development, Cell, 125:1241–52 Chang, DT, Reynolds, IJ, 2006, Mitochondrial trafficking and morphology in healthy and injured neurons, Prog Neurobiol 80:241–68 Chen, H, Chan, DC, 2009, Mitochondrial dynamics – fusion, fission, movement, and mitophagy – in neurodegenerative diseases. Hum Mol Genet 18:169–76 Chen, S, Owens, GC, Crossin, KL, Edelman, DB, 2007, Serotonin stimulates mitochondrial transport in hippocampal neurons, Mol Cell Neurosci 36:472–83 Chen, S, Owens, GC, Edelman, DB, 2008, Dopamine inhibits mitochondrial motility in hippocampal neurons, PLoS ONE, 30:e2804 Chen, YM, Gerwin, C, Sheng, ZH, 2009, Dynein light chain LC8 regulates syntaphilin-mediated mitochondrial docking in axons, J Neurosci 29:9429–38 Cheney, RE, O’Shea, MK, Heuser, JE, Coelho, MV, Wolenski, JS, Espreafico, EM, Forscher, P, Larson, RE, Mooseker, MS, 1993, Brain myosin-V is a two-headed unconventional myosin with motor activity. Cell, 75:13–23 Chevalier-Larsen, E, Holzbaur, EL, 2006, Axonal transport and neurodegenerative disease, Biochim. Biophys Acta 1762:1094–108 Cho, KI, Cai, Y, Yi, H, Yeh, A, Aslanukov, A, Ferreira, PA, 2007, Association of the kinesinbinding domain of RanBP2 to KIF5B and KIF5C determines mitochondria localization and function, Traffic, 8:1722–35 Cleveland, DW, Rothstein, JD, 2001, From Charcot to Lou Gehrig: deciphering selective motor neuron death in ALS. Nat Rev Neurosci 2:806–19 De Vos, KJ, Chapman, AL, Tennant, ME, Manser, C, Tudor, EL, Lau, KF, Brownlees, J, Ackerley, S, Shaw, PJ, McLoughlin, DM, Shaw, CE, Leigh, PN, Miller, CC, Grierson, AJ, 2007, Familial amyotrophic lateral sclerosis-linked SOD1 mutants perturb fast axonal transport to reduce axonal mitochondria content. Hum Mol Genet 16:2720–28 Fabricius, C, Berthold, C,H, Rydmark, M, 1993, Axoplasmic organelles at nodes of Ranvier. II. Occurrence and distribution in large myelinated spinal cord axons of the adult cat, J Neurocytol 22:941–54

164

Q. Cai and Z.-H. Sheng

Fransson, A, Ruusala, A, Aspenström, P, 2003, Atypical Rho GTPases have roles in mitochondrial homeostasis and apoptosis, J Biol Chem 278:6495–502 Gindhart, JG, Chen, J, Faulkner, M, Gandhi, R, Doerner, K, Wisniewski, T, Nandlestadt, A, 2003, The kinesin-associated protein UNC-76 is required for axonal transport in the Drosophila nervous system, Mol Biol Cell, 14:3356–65 Glater, EE, Megeath, LJ, Stowers, RS, Schwarz, TL, 2006, Axonal transport of mitochondria requires milton to recruit kinesin heavy chain and is light chain independent, J Cell Biol 173:545–57 Górska-Andrzejak, J, Stowers, RS, Borycz, J, Kostyleva, R, Schwarz TL, Meinertzhagen, IA, 2003, Mitochondria are redistributed in Drosophila photoreceptors lacking milton, a kinesinassociated protein, J Comp Neurol 463:372–88 Gotow, T, Miyaguchi, K, Hashimoto, PH, 1991, Cytoplasmic architecture of the axon terminal: filamentous strands specifically associated with synaptic vesicles, Neuroscience, 40:587–98 Grishin, A, Li, H, Levitan, ES, Zaks-Makhina, E, 2006, Identification of gamma-aminobutyric acid receptor-interacting factor 1 (TRAK2) as a trafficking factor for the K + channel Kir2.1, J Biol Chem 281:30104–11 Guo, X, Macleod, GT, Wellington, A, Hu, F, Panchumarthi, S, Schoenfield, M, Marin, L, Charlton, MP, Atwood HL, Zinsmaier, KE, 2005, The GTPase dMiro is required for axonal transport of mitochondria to Drosophila synapses, Neuron, 47:379–93 Hafezparast, M et  al., 2003, Mutations in dynein link motor neuron degeneration to defects in retrograde transport, Science, 300:808–12 Gurney, ME, Pu, H, Chiu, AY, Dal Canto, MC, Polchow, CY, Alexander, DD, Caliendo, J, Hentati, A, Kwon, YW, Deng, HX, et  al. 1994, Motor neuron degeneration in mice that express a human Cu,Zn superoxide dismutase mutation. Science 264:1772–5 Haghnia, M, Cavalli, V, Shah, SB, Schimmelpfeng, K, Brusch, R, Yang, G, Herrera, C, Pilling, A, Goldstein, LS, 2007, Dynactin is required for coordinated bidirectional motility, but not for dynein membrane attachment, Mol Biol Cell, 18:2081–89 Hirokawa, N, 1982, Cross-linker system between neurofilaments, microtubules, and membranous organelles in frog axons revealed by the quick-freeze, deep-etching method, J Cell Biol 94:129–42 Hirokawa, N, Sato-Yoshitake, R, Yoshida, T, Kawashima, T, 1990, Brain dynein (MAP1C) localizes on both anterogradely and retrogradely transported membranous organelles in vivo, J Cell Biol 111:1027–37 Hirokawa, N, Sato-Yoshitake, R, Kobayashi, N, Pfister, KK, Bloom, GS, Brady, ST, 1991, Kinesin associates with anterogradely transported membranous organelles in  vivo, J Cell Biol 114:295–302 Hirokawa, N, Takemura, R, 2004, Kinesin superfamily proteins and their various functions and dynamics, Exp Cell Res 301:50–9 Hirokawa, N, Takemura, R, 2005, Molecular motors and mechanisms of directional transport in neurons, Nat Rev Neurosci 6:201–14 Hollenbeck, PJ, 1996, The pattern and mechanism of mitochondrial transport in axons, Front Biosci 1:d91-102 Hollenbeck, PJ, Saxton, WM, 2005, The axonal transport of mitochondria, J Cell Sci 118:5411–19 Horiuchi, D, Barkus, RV, Pilling, AD, Gassman, A, Saxton, WM, 2005, APLIP1, a kinesin binding JIP-1/JNK scaffold protein, influences the axonal transport of both vesicles and mitochondria in Drosophila, Curr Biol 15:2137–41 Horiuchi, D, Collins, CA, Bhat, P, Barkus, RV, Diantonio, A, Saxton, WM, 2007, Control of a kinesin-cargo linkage mechanism by JNK pathway kinases, Curr Biol 17:1313–7 Hubley, MJ, Locke, BR, Moerland, TS, 1996, The effects of temperature, pH, and magnesium on the diffusion coefficient of ATP in solutions of physiological ionic strength, Biochim Biophys Acta 1291:115–21 Hurd, DD, Saxton, WM, 1996, Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila, Genetics, 144:1075–85

5  Mitochondrial Dynamics and Axonal Transport

165

Ikuta, J, Maturana, A, Fujita, T, Okajima, T, Tatematsu, K, Tanizawa, K, Kuroda, S. Fasciculation and elongation protein zeta-1 (FEZ1) participates in the polarization of hippocampal neuron by controlling the mitochondrial motility, Biochem Biophys Res Commun 353:127–32 Jonas, E, 2006, BCL-xL regulates synaptic plasticity, Mol Interv 6:208–22 Kanai, Y, Okada, Y, Tanaka, Y, Harada, A, Terada, S, Hirokawa, N, 2000, KIF5C, a novel neuronal kinesin enriched in motor neurons, J Neurosci 20:6374–84 Kang, JS, Tian, JH, Pan, PY, Zald, P, Li, C, Deng, C, Sheng, ZH, 2008, Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation, Cell, 132:137–48 Karki, S, Holzbaur, EL, 1999, cytoplasmic dynein and dynactin in cell division and intracellular transport, Curr Opin Cell Biol 11:45–53 King, SJ, Schroer, TA, 2000, Dynactin increases the processivity of the cytoplasmic dynein motor, Nat Cell Biol 2:20–4 Kirk, E, Chin, LS, Li, L, 2006, GRIF1 binds Hrs and is a new regulator of endosomal trafficking, J Cell Sci 119:4689–701 Kuroda, S, Nakagawa, N, Tokunaga, C, Tatematsu, K, Tanizawa, K, 1999, Mammalian homologue of the Caenorhabditis elegans UNC-76 protein involved in axonal outgrowth is a protein kinase C zeta-interacting protein, J Cell Biol 144:403–11 Langford, GM, 2002, Myosin-V, a versatile motor for short-range vesicle transport, Traffic, 3:859–65 Lee, CW, Peng, HB, 2008, The function of mitochondria in presynaptic development at the neuromuscular junction, Mol Biol Cell, 19:150–8 Leterrier, JF, Rusakov, DA, Nelson, BD, Linden, M, 1994, Interactions between brain mitochondria and cytoskeleton: evidence for specialized outer membrane domains involved in the association of cytoskeleton-associated proteins to mitochondria in situ and in vitro, Microsc Res Tech 27:233–61 Levy, M, Faas, GC, Saggau, P, Craigen WJ, Sweatt, JD, 2003, Mitochondrial regulation of synaptic plasticity in the hippocampus, J Biol Chem 278:17727–34 Levy, JR, Sumner, CJ, Caviston, JP, Tokito, MK, Ranganathan, S, Ligon, LA, Wallace, KE, LaMonte, BH, Harmison, GG, Puls, I, Fischbeck, KH, Holzbaur, EL, 2006, A motor neuron disease-associated mutation in p150Glued perturbs dynactin function and induces protein aggregation, J Cell Biol 172:733–45 Li, Z, Okamoto, K, Hayashi, Y, Sheng, M, 2004, The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses, Cell, 119:873–87 Ligon, LA, Steward, O, 2000, Movement of mitochondria in the axons and dendrites of cultured hippocampal neurons, J Comp Neurol 427:340–50 Lindén, M, Nelson, BD, Loncar, D, Leterrier, JF, 1989a, Studies on the interaction between mitochondria and the cytoskeleton, J Bioenerg. Biomembr 21:507–18 Lindén, M, Nelson, BD, Leterrier, JF, 1989b, The specific binding of the microtubule-associated protein 2 (MAP2) to the outer membrane of rat brain mitochondria, Biochem J 261:167–73 Ma, H, Cai, Q, Lu, W, Sheng, ZH, Mochida, S, 2009, KIF5B motor adaptor syntabulin maintains synaptic transmission in sympathetic neurons, J Neurosci 29:13019–29 Macaskill, AF, Rinholm, JE, Twelvetrees, AE, Arancibia-Carcamo, IL, Muir, J, Fransson, A, Aspenstrom, P, Attwell, D, Kittler, JT. 2009, Miro1 is a calcium sensor for glutamate receptordependent localization of mitochondria at synapses, Neuron, 61:541–55 Magrané, J, Hervias, I, Henning, MS, Damiano, M, Kawamata, H, Manfredi, G, 2009, Mutant SOD1 in neuronal mitochondria causes toxicity and mitochondrial dynamics abnormalities. Hum Mol Genet 18:4552–64 Magrané, J, Manfredi, G, 2009, Mitochondrial function, morphology, and axonal transport in amyotrophic lateral sclerosis, Antioxid Redox Signal, 11:1615–26 Malaiyandi, LM, Vergun, O, Dineley, KE, Reynolds, IJ, 2005, Direct visualization of mitochondrial zinc accumulation reveals uniporter-dependent and -independent transport mechanisms, J Neurochem 93:1242–50 Mallik, R, Petrov, D, Lex, SA, King, SJ, Gross, SP. 2005, Building complexity: an in vitro study of cytoplasmic dynein with in vivo implications, Curr Biol 15:2075–85

166

Q. Cai and Z.-H. Sheng

Martin, M, Iyadurai, SJ, Gassman, A, Gindhart, JG, Jr Hays, TS, Saxton, WM, 1999, Cytoplasmic dynein, the dynactin complex, and kinesin are interdependent and essential for fast axonal transport, Mol Biol Cell, 10:3717–28 Mavlyutov, TA, Cai, Y, Ferreira, PA, 2002, Identification of RanBP2- and kinesin-mediated transport pathways with restricted neuronal and subcellular localization, Traffic, 3:630–40 Miller, KE, Sheetz, MP, 2004, Axonal mitochondrial transport and potential are correlated, J Cell Sci 117:2791–804 Minin, AA, Kulik, AV, Gyoeva, FK, Li, Y, Goshima, G, Gelfand, VI, 2006, Regulation of mitochondria distribution by RhoA and formins, J Cell Sci 119:659–70 Misko, A, Jiang, S, Wegorzewska, I, Milbrandt, J, Baloh, RH, 2010, Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex, J Neurosci 30:4232–40 Mironov, SL, 2006, Spontaneous and evoked neuronal activities regulate movements of single neuronal mitochondria, Synapse, 59:403–11 Mironov, SL, 2007, ADP regulates movements of mitochondria in neurons, Biophys J 92:2944–52 Mironov, SL, 2009, Complexity of mitochondrial dynamics in neurons and its control by ADP produced during synaptic activity, Int J Biochem Cell Biol 41:2005–14 Morris, RL, Hollenbeck, PJ, 1993, The regulation of bidirectional mitochondrial transport is coordinated with axonal outgrowth, J Cell Sci 104:917–27 Morris, RL, Hollenbeck, PJ, 1995, Axonal transport of mitochondria along microtubules and F-actin in living vertebrate neurons, J Cell Biol 131:1315–26 Mutsaers, SE, Carroll, WM, 1998, Focal accumulation of intra-axonal mitochondria in demyelination of the cat optic nerve, Acta Neuropathol 96:139–43 Nangaku, M, Sato-Yoshitake, R, Okada, Y, Noda, Y, Takemura, R, Yamazaki, H, Hirokawa, N, 1994, KIF1B, a novel microtubule plus end-directed monomeric motor protein for transport of mitochondria, Cell, 79:1209–20 Naisbitt, S, Valtschanoff, J, Allison, DW, Sala, C, Kim, E, Craig, AM, Weinberg, RJ, Sheng, M, 2000, Interaction of the postsynaptic density-95/guanylate kinase domain-associated protein complex with a light chain of myosin-V and dynein, J Neurosci 20:4524–34 Orr, AL, Li, S, Wang, CE, Li, H, Wang, J, Rong, J, Xu, X, Mastroberardino, PG, Greenamyre, JT, Li, XJ, 2008, N-terminal mutant huntingtin associates with mitochondria and impairs mitochondrial trafficking, J Neurosci 28:2783–92 O’Toole, M, Latham, R, Baqri, RM, Miller, KE, 2008, Modeling mitochondrial dynamics during in vivo axonal elongation, J Theor Biol 255:369–77 Overly, CC, Rieff, HI, Hollenbeck, PJ, 1996, Organelle motility and metabolism in axons vs dendrites of cultured hippocampal neurons, J Cell Sci 109:971–80 Pathak, D, Sepp, KJ, Hollenbeck, PJ, 2010, Evidence that myosin activity opposes microtubulebased axonal transport of mitochondria, J Neurosci 30:8984–92 Pigino, G, Morfini, G, Pelsman, A, Mattson, MP, Brady, ST, Busciglio, J, 2003, Alzheimer’s presenilin 1 mutations impair kinesin-based axonal transport, J Neurosci 23:4499–508 Pilling, AD, Horiuchi, D, Lively, CM, Saxton, WM, 2006, Kinesin-1 and Dynein are the primary motors for fast transport of mitochondria in Drosophila motor axons, Mol Biol Cell, 17:2057–68 Povlishock, JT, 197 The fine structure of the axons and growth cones of the human fetal cerebral cortex, Brain Res 114:379-9i Puls, I, Jonnakuty, C, LaMonte, BH, Holzbaur, EL, Tokito, M, Mann, E, Floeter, MK, Bidus, K, Drayna, D, Oh, SJ, Brown, RH, Jr, Ludlow CL, Fischbeck, KH, 2003, Mutant dynactin in motor neuron disease, Nat Genet 33:455–6 Reynolds, IJ, Rintoul, GL, 2004, Mitochondrial stop and go: signals that regulate organelle movement, Sci STKE 21:PE46 Rintoul, GL, Filiano, AJ, Brocard, JB, Kress, GJ, Reynolds, IJ, 2003, Glutamate decreases mitochondrial size and movement in primary forebrain neurons, J Neurosci 23:7881–88 Rintoul, GL, Bennett, VJ, Papaconstandinou, NA, Reynolds, IJ, 2006, Nitric oxide inhibits mitochondrial movement in forebrain neurons associated with disruption of mitochondrial membrane potential, J Neurochem 97:800–6

5  Mitochondrial Dynamics and Axonal Transport

167

Rosen, DR, Siddique, T, Patterson, D, Figlewicz, DA, Sapp, P, Hentati, A, Donaldson, D, Goto, J, O’Regan, JP, Deng, HX, et  al. 1993, Mutations in Cu/Zn superoxide dismutase gene are associated with familial amyotrophic lateral sclerosis. Nature 362:59–62 Russo, GJ, Louie, K, Wellington, A, Macleod, GT, Hu, F, Panchumarthi, S, Zinsmaier, KE, 2009, Drosophila Miro is required for both anterograde and retrograde axonal mitochondrial transport, J Neurosci 29:5443–55 Ruthel, G, Hollenbeck, PJ, 2003, Response of mitochondrial traffic to axon determination and differential branch growth, J Neurosci 23:8618–24 Saotome, M, Safiulina, D, Szabadkai, G, Das S, Fransson, A, Aspenstrom, P, Rizzuto, R, Hajnóczky, G, 2008, Bidirectional Ca2+−dependent control of mitochondrial dynamics by the Miro GTPase, Proc Natl Acad Sci U S A 105:20728–33 Sasaki, S, Iwata, M, 1996, Impairment of fast axonal transport in the proximal axons of anterior horn neurons in amyotrophic lateral sclerosis. Neurology 47:535–40 Schroer, TA, 2004, Dynactin, Annu Rev Cell Dev Biol 20:759–79 Schwarzer, C, Barnikol-Watanabe, S, Thinnes, FP, Hilschmann, N, 2002, Voltage-dependent anion-selective channel (VDAC) interacts with the dynein light chain Tctex1 and the heatshock protein PBP74, Int J Biochem Cell Biol 34:1059–70 Smith, MJ, Pozo, K, Brickley, K, Stephenson, FA, 2006, Mapping the GRIF-1 binding domain of the kinesin, KIF5C, substantiates a role for GRIF-1 as an adaptor protein in the anterograde trafficking of cargoes, J Biol Chem 281:27216–28 Stamer, K, Vogel, R, Thies, E, Mandelkow, E, Mandelkow, EM, 2002, Tau blocks traffic of organelles, neurofilaments, and APP vesicles in neurons and enhances oxidative stress, J Cell Biol 156:1051–63 Stokin, GB, Goldstein, LS, 2006, Axonal transport and Alzheimer’s disease, Annu Rev Biochem 75:607–27 Stowers, RS, Megeath, LJ, Górska-Andrzejak, J, Meinertzhagen IA, Schwarz, TL, 2002, Axonal transport of mitochondria to synapses depends on milton, a novel Drosophila protein, Neuron, 36:1063–77 Su, Q, Cai, Q, Gerwin, C, Smith, CL, Sheng, ZH, 2004, Syntabulin is a microtubule-associated protein implicated in syntaxin transport in neurons, Nat Cell Biol 6:941–53 Sung, JY, Engmann, O, Teylan, MA, Nairn, AC, Greengard, P, Kim, Y. 2008, WAVE1 controls neuronal activity-induced mitochondrial distribution in dendritic spines, Proc Natl Acad Sci USA 105:3112–6 Susalka, SJ, Pfister, KK, 2000, Cytoplasmic dynein subunit heterogeneity: implications for axonal transport, J Neurocytol 29:819–29 Tai, AW, Chuang, JZ, Bode, C, Wolfrum, U, Sung, CH, 1999, Rhodopsin’s carboxy-terminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1, Cell, 97:877–87 Tanaka, Y, Kanai, Y, Okada, Y, Nonaka, S, Takeda, S, Harada, A, Hirokawa, N, 1998, Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria, Cell, 93:1147–58 Tang, Y, Zucker, RS, 1997, Mitochondrial involvement in post-tetanic potentiation of synaptic transmission, Neuron, 18:483–91 Toh, BH, Lolait, SJ, Mathy, JP, Baum, R. 1980, Association of mitochondria with intermediate filaments and of polyribosomes with cytoplasmic actin, Cell Tissue Res 211:163–9 Trushina, E, Dyer, RB, Badger, JD 2nd, Ure D, Eide L, Tran, DD, Vrieze, BT, LegendreGuillemin, V, McPherson, PS, Mandavilli, BS, Van Houten, B, Zeitlin, S, McNiven, M, Aebersold, R, Hayden, M, Parisi, JE, Seeberg, E, Dragatsis, I, Doyle, K, Bender, A, Chacko, C, McMurray, CT, 2004, Mutant huntingtin impairs axonal trafficking in mammalian neurons in vivo and in vitro, Mol Cell Biol 24:8195–209 Tynan, SH, Purohit, A, Doxsey, SJ, Vallee, RB, 2000, Light intermediate chain 1 defines a functional subfraction of cytoplasmic dynein which binds to pericentrin, J Biol Chem 275:32763–8 Vale, RD, Reese, TS, Sheetz, MP, 1985, Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility, Cell, 42:39–50

168

Q. Cai and Z.-H. Sheng

Verburg, J, Hollenbeck, PJ, 2008, Mitochondrial membrane potential in axons increases with local nerve growth factor or semaphorin signaling, J Neurosci 28:8306–15 Verstreken, P, Ly, CV, Venken, KJ, Koh, TW, Zhou, Y, Bellen, HJ, 2005, Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions, Neuron 47:365–78 Wagner, OI, Lifshitz, J, Janmey, PA, Linden, M, McIntosh, TK, Leterrier, JF, 2003, Mechanisms of mitochondria-neurofilament interactions, J Neurosci 23:9046–58 Wang, X, Schwarz, TL, 2009, The mechanism of Ca2+ −dependent regulation of kinesin-mediated mitochondrial motility, Cell 2009 136:163–74 Waterman-Storer, CM, Karki, S, Holzbaur, EL, 1995, The p150Glued component of the dynactin complex binds to both microtubules and the actin-related protein centractin (Arp-1), Proc Natl Acad Sci USA 92:1634–38 Webber, E, Li, L, Chin, LS, 2008, Hypertonia-associated protein Trak1 is a novel regulator of endosome-to-lysosome trafficking, J Mol Biol 382:638–51 Winter, L, Abrahamsberg, C, Wiche, G, 2008, Plectin isoform 1b mediates mitochondrionintermediate filament network linkage and controls organelle shape, J Cell Biol 181:903–11 Wolenski, JS, Cheney, RE, Mooseker, MS, Forscher, P, 1995, In vitro motility of immunoadsorbed brain myosin-V using a Limulus acrosomal process and optical tweezer-based assay, J Cell Sci 108:1489–96 Yang, F, He, XP, Russell, J, Lu, B, 2003, Ca2+ influx-independent synaptic potentiation mediated by mitochondrial Na(+)-Ca2+ exchanger and protein kinase C, J Cell Biol 163:511–23 Yi, M, Weaver, D, Hajnóczky, G, 2004, Control of mitochondrial motility and distribution by the calcium signal: a homeostatic circuit, J Cell Biol 167:661–72 Zanelli, SA, Trimmer, PA, Solenski, NJ, 2006, Nitric oxide impairs mitochondrial movement in cortical neurons during hypoxia, J Neurochem 97:724–36 Zhang, CL, Ho, PL, Kintner, DB, Sun, D, Chiu, SY, 2010, Activity-dependent regulation of mitochondrial motility by calcium and Na/K-ATPase at nodes of Ranvier of myelinated nerves, J Neurosci 30:3555–66

Chapter 6

Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery Guy Lenaers, Dominique Bonneau, Cécile Delettre, Patrizia Amati-Bonneau, Emmanuelle Sarzi, Dan Miléa, Christophe Verny, Vincent Procaccio, Christian Hamel, and Pascal Reynier Abstract  Dominant optic atrophy (DOA) and Charcot-Marie-Tooth type 2A disease (CMT2A) are inherited neurological disorders primarily affecting retinal ganglion cells and peripheral nerves, and secondarily, other neuronal cells with high energetic requirements. The discovery of OPA1 mutations in DOA patients and MFN2 ­mutations in CMT2A patients, has defined a new subset of mitochondrial diseases related to mitochondrial dynamics. OPA1 and MFN2 are both dynamin GTPases that promote the fusion of the mitochondrial network. Since diseases related to OPA1 and MFN2 mutations show some clinical similarities, their pathophysiological mechanisms are believed to share some common mitochondrial dysfunctions. These mechanisms are closely ­interdependent and involve mitochondrial fusion, motility along the axons, energy ­production and metabolism, maintenance of the organelle and its genome, as well as ­susceptibility to apoptosis. This chapter presents recent findings on: (1) the clinical diversity of OPA1- and MFN2-related diseases; (2) the spectrum of mutations in the OPA1 and MFN2 genes and their consequences at the protein level; and (3) the pathophysiology of DOA and CMT2A as characterized in cellular and mouse models. Finally, we discuss the therapeutic perspectives opened up by the current knowledge of these diseases. G. Lenaers (), C. Delettre, E. Sarzi, and C. Hamel Inserm U1051, Institut des Neurosciences de Montpellier, Universités de Montpellier I et II, Montpellier, France e-mail: [email protected] D. Bonneau, P. Amati-Bonneau, V. Procaccio, and P. Reynier UMR CNRS 6214 - Inserm U771, Angers, France and Département de Biochimie et Génétique, Centre Hospitalier Universitaire, Angers, France D. Miléa UMR CNRS 6214 - Inserm U771, Angers, France and Département d’Ophtalmologie, Centre Hospitalier Universitaire, Angers, France C. Verny UMR CNRS 6214 - Inserm U771, Angers, France and Département de Neurologie, Centre Hospitalier Universitaire, Angers, France B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_6, © Springer Science+Business Media B.V. 2011

169

170

G. Lenaers et al.

Keywords  Dominant optic atrophy • Charcot-Marie-Tooth disease • Mitochondria • MFN2 • OPA1 Abbreviations aa ADP ANT ATP CMT disease CPEO DOA DRG ENU GTP GTPase HR IMM LHON MEF MFN2 mtDNA OMA1 OMM OPA1 OXPHOS Phb1/2 RGC TM VEP YME1L

amino acid adenosine-5’-diphosphate adenine nucleotide translocator adenosine-5’-triphosphate Charcot-Marie-Tooth disease chronic progressive external ophthalmoplegia dominant optic atrophy dorsal root ganglion N-ethyl-N-nitrosurea guanosine triphosphate guanosine triphosphate phosphorylase heptad repeat domain inner mitochondrial membrane. Leber’s hereditary optic neuropathy mouse embryonic fibroblasts Mitofusin 2 mitochondrial DNA overlapping activity with M-AAA protease 1 outer mitochondrial membrane optic atrophy 1 oxidative phosphorylation prohibitins 1 and 2 retinal ganglion cell transmembrane domain visual evoked potential YME1-like protein 1

6.1 OPA1-Associated Diseases 6.1.1 Historical Description of Dominant Optic Atrophy (DOA) In 1959, the Danish ophthalmologist Poul Kjer described an infantile optic atrophy with a dominant mode of inheritance identified in 19 families (Kjer 1959). Dominant optic atrophy (DOA; MIM #165500), also called Kjer disease, is classically described as a progressive, isolated optic neuropathy, developing insidiously

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 171

over the first two decades of life. Rarely, the clinical presentation of the disease is atypical including acute, severe visual loss, reversible visual loss, or late occurrence in the life course. Bilateral visual loss is generally moderate, but, in some cases, leading to legal blindness (Votruba et al. 1998). The loss of the central visual field, due to centrocecal, central or paracentral scotoma, is commonly bilateral and sometimes progressive, associated with difficulty in discriminating colors in the blue-yellow axis, corresponding to dyschromatopsia of the tritanopic type. Ophthalmoscopic examination discloses temporal or diffuse pallor of the optic nerve heads, sometimes associated with optic disc excavation. The rare histopathological postmortem studies published several years ago described a selective loss of retinal ganglion cells (RGC) that form the optic nerve (Johnston et al. 1979; Kjer et  al. 1983). More recently, in  vivo optical coherence tomography confirmed the ­thinning of the optic fiber layer around the optic disc in patients, whereas the other retinal layers remain unaltered (Milea et al. 2010). From a genetic point of view, DOA has incomplete penetrance with highly heterogeneous clinical expressivity even within families (Hoyt 1980; Votruba et al. 1998). The classical clinical presentation of DOA is distinct from that of Leber’s hereditary optic neuropathy (LHON), which is characterized by an acute and sequential onset of visual loss in both eyes occurring at an older age, typically 18–35 years (Carelli et al. 2004). LHON, which is exclusively transmitted maternally through mitochondrial DNA (mtDNA) mutations, predominantly affects males, whereas DOA affects males and females equally.

6.1.2 The Discovery of OPA1 The locus for the main DOA-causing gene, optic atrophy 1 (OPA1, MIM *605290), was mapped to chromosome 3q28-29 (Eiberg et  al. 1994; Lunkes et  al. 1995). Subsequently, DOA was found to be genetically heterogeneous as two further loci were identified, i.e. OPA4 (MIM %605293) and OPA5 (MIM %610708), mapping to chromosomes 18q12.2-q12.3 (Kerrison et al. 1998) and 22q12.1-q13.1 (Barbet et al. 2005) respectively. In 2000, studies on mitochondrial dynamics in yeast, led to the identification of the human ortholog gene of the MGM1/Msp1 genes of Saccharomyces cerevisiae and Schizosaccharomyces pombe. This gene, encoding a mitochondrial GTPase of the dynamin family was localized on chromosome 3 at the morbid locus linked to DOA. Gene sequencing in six families suffering from DOA unraveled pathogenic mutations, indicating that the ortholog of the MGM1/ Msp1 genes was indeed OPA1, the gene responsible for DOA (Delettre et al. 2000). Meanwhile, another team from Germany and the United Kingdom reached the same conclusion using positional cloning (Alexander et  al. 2000). These two articles published in Nature Genetics in 2000, marked the beginning of the explo­ ration of a new subtype of mitochondrial disorders involving mitochondrial network dynamics.

172

G. Lenaers et al.

6.1.3 Revisiting Clinical Phenotypes of DOA on the Basis of Current Knowledge of the OPA1 Gene Since 2000, the systematic sequencing of OPA1 in patients with different forms of optic atrophy has led to a better understanding of the pathophysiology of DOA and to refining the clinical spectrum of the disease, which has turned out to be larger than expected. Indeed, OPA1 mutations are responsible for many different phenotypes ranging from mild disorders affecting only the RGCs, as in DOA, to severe and complex phenotypes, named “DOA+”, which are multi-systemic diseases rather similar to mitochondrial diseases due to oxidative phosphorylation (OXPHOS) defects. The routine screening of patients with isolated optic nerve atrophies, which could not be classified as DOA on the basis of genetic or clinical parameters, has led to the discovery of novel OPA1 mutations and phenotype-genotype correlations. Moreover, the identification of multi-systemic forms of DOA has revealed that RGCs, although highly sensitive to these diseases, are not the only cells affected by OPA1 dysfunction. The molecular screening of 980 patients with sporadic and familial optic neuropathies showed that 295 patients (30.1%) carried an OPA1 mutation, 131 patients (13.4%) a LHON-related mtDNA mutation and 14 patients (1.4%) an OPA3 mutation responsible for a rare form of DOA associated with cataract (Ferre et  al. 2009; Reynier et al. 2004). In addition, this study showed the high prevalence of pathogenic OPA1 mutations in 157 of the 392 (40%) patients with apparent sporadic optic atrophy, demonstrating the frequent absence of the dominant inheritance considered to be characteristic of this disease. DOA is classically associated with a progressive, irreversible loss of vision. We have reported several cases of DOA presenting with a large clinical heterogeneity, including cases of spontaneous recovery of vision (Cornille et al. 2008), or with late-onset, LHON-like acute optic atrophy in a 62-year-old woman carrying an OPA1 mutation (Nochez et al. 2009). Several LHON patients referred to our laboratory for mtDNA-mutation screening were actually found to harbor an OPA1 mutation, suggesting a degree of clinical overlap between the two disorders. A recent study of 104 patients, from 45 unrelated families, sums up the new clinical forms of DOA described over the past 10 years (Yu-Wai-Man et al. 2010a). Nearly 20% of patients with DOA have extra-ocular impairment associated with optic neuropathy; these clinical forms being referred to as “DOA+” (Table 6.1). The commonest form of this impairment is bilateral, progressive sensorineural deafness beginning in adolescence (Amati-Bonneau et  al. 2005). The association of DOA with deafness was initially reported in patients carrying the same OPA1 p.R445H missense mutation (Amati-bonneau et al. 2003, 2005; Li et al. 2005; Payne et al. 2004; Shimizu et al. 2003) and other OPA1 mutations were later reported (Ke et al. 2006; Puomila et al. 2005; Yu-Wai-Man et al. 2010b). Another “DOA+” clinical phenotype associates ataxia, myopathy, peripheral neuropathy, and chronic progressive external ophthalmoplegia (Amati-Bonneau et al. 2008; Hudson et al. 2008). Finally, more rarely, other disorders mimicking

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 173 Table  6.1  Clinical and pathophysiological similarities and differences between OPA1- and MFN2-related diseases. CPEO: Chronic progressive external ophthalmoplegia DOA/OPA1 Clinical phenotypes CMT2A/MFN2 + Optic neuropathy +/− +/− Peripheral neuropathy + +/− Sensorineural deafness +/− +/− Encephalopathy +/− +/− Myopathy and CPEO − +/− Multiple sclerosis-like − +/− Tremor +/− + + ? + + + +/− +/−

Pathophysiological conditions Mitochondrial morphological anomalies Energy production defect Axonal transport alteration Alteration of neuron connectivity Increased susceptibility to apoptosis Mitochondrial DNA instability Haploinsufficiency Dominant negative effect

+ + + ? − − − +

hereditary spastic paraparesis (Yu-Wai-Man et  al. 2010b), peripheral neuropathy (Spinazzi et al. 2008), or multiple sclerosis have been reported (Verny et al. 2008). Interestingly, in a few cases, the clinical presentations of OPA1 mutations excluded optic nerve involvement (Milone et al. 2009; Yu-Wai-Man et al. 2010b), showing that some OPA1-associated diseases tend towards phenotypes far removed from the initial description of DOA. The “DOA+” multi-systemic syndrome is described as beginning with optic ­atrophy during childhood and then extending to chronic progressive external ­ophthalmoplegia (CPEO), ataxia, sensorineural deafness, sensory-motor neuropathy and myopathy in adult life (Amati-Bonneau et al. 2008; Hudson et al. 2008; Stewart et al. 2008). The muscle abnormalities observed in patients with “DOA+” were similar to those found in mitochondrial myopathies, with the presence of cytochrome c oxidase-negative fibers and ragged red fibers. In fact, these patients harbored multiple mtDNA deletions in skeletal muscle, suggesting that mitochondrial fusion and the OPA1 protein, in particular, play a key role in mtDNA maintenance. OPA1 mutations are by far the commonest genetic causes of DOA. However, the rate of detection of OPA1 mutations in DOA families is highly variable, ranging from 32% to 89% (Cohn et al. 2007; Delettre et  al. 2001; Marchbank et  al. 2002; Pesch et al. 2001; Thiselton et al. 2002; Toomes et al. 2001). OPA1-related DOA has incomplete penetrance, and several carriers of OPA1 mutations remain asymptomatic. Initially, the penetrance of the disease was estimated to be as high as 98% (Kivlin et  al. 1983), but the identification of OPA1 mutations led to a ­re-evaluation of this figure, with results ranging from 43% to 90% in more recent studies (Cohn et al. 2007; Puomila et al. 2005; Toomes et al. 2001).

174

G. Lenaers et al.

6.1.4 Epidemiology of DOA DOA and LHON are the two commonest forms of hereditary optic atrophy. The prevalence of DOA is estimated at about 1/50,000 in the general population (Lyle 1990), but the actual prevalence was found to be much higher in Denmark (1/12,000) (Kjer et  al. 1996; Thiselton et  al. 2001). The discovery of OPA1 also helped to show that the higher prevalence of DOA observed in the Danish population was due to a founder effect involving the c.2826delT mutation, hypothesized to have been introduced at least 69 generations ago, much before the Viking era (Thiselton et al. 2001). Recently, the prevalence of DOA due to OPA1 mutations has been re-evaluated to over 1/35,000 in a large cohort of patients of the north of England (Yu-Wai-Man et al. 2010a).

6.2 MFN2-Associated Charcot-Marie-Tooth Diseases 6.2.1 Clinical Description and Classification of Charcot-MarieTooth Diseases Charcot-Marie-Tooth disease (CMT) owes its name to three neurologists who first described the disorder in 1886 (Charcot and Marie 1886; Tooth 1886). CMT is now known to include a heterogeneous group of hereditary chronic neuropathies, affecting sensory and motor neurons (Skre 1974), characterized by motor weakness and muscle atrophy affecting the limbs, reduction or abolition of tendon reflexes, and distal sensory loss, often associated with orthopedic deformities, such as hollow feet. The disease progresses slowly, gradually leading to impaired mobility, and often to wheelchair dependency. The CMT group of diseases is clinically heterogeneous, even within the same family. Several forms of CMT have been described on the basis of electrophysiological criteria (nerve conduction velocities), modes of inheritance, and the molecular genetic classification. The electrophysiology helps to distinguish the axonal forms from the demyelinating forms of CMT since the latter are specifically associated with motor nerve conduction velocities lower than 38  m/s. The axonal forms ­correspond to predominantly axonal lesions that do not affect the integrity of the myelin sheath. The modes of inheritance may be autosomal dominant, X-linked or autosomal recessive. The most common forms of CMT are the autosomal dominant demyelinating form (CMT1), the axonal form (CMT2), and the X-linked forms (CMTX). Autosomal recessive forms of CMT are rarer but more severe, and may be either axonal (AR-CMT2) or demyelinating (CMT4). So far, more than 30 genes have been linked to the different forms of CMT. Among these genes, four encode mitochondrial proteins, two of which are involved in mitochondrial dynamics: mitofusin 2 (MFN2) and the ganglioside-induced differentiation-associated protein 1 (GDAP1).

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 175

6.2.2 The Discovery of MFN2 In 2004, the MFN2 gene was linked with axonal, autosomal dominant Charcot-Marie-Tooth disease (CMT type 2A; MIM #118210), (Zuchner et al. 2006). Subsequently, it was found that MFN2 mutations occurred in 9–33% of CMT2 patients (Calvo et al. 2009; Kijima et al. 2005; Verhoeven et al. 2006). On the basis of clinical criteria, the disease penetrance was estimated at 74–100% (Lawson et al. 2005).

6.2.3 “CMT2A+” Phenotypes Like DOA linked to OPA1 mutations, CMT due to MFN2 mutations may be associated with other neurological disorders, as in the case of hereditary motor and sensory neuropathy type VI, which includes peripheral and optic neuropathies (Zuchner et  al. 2006). Patients with this disorder suffer from severe peripheral neuropathy, often with a subacute onset of optic atrophy followed by slow recovery of visual acuity. This observation, illustrating the similarities between the peripheral neuropathy found in some patients carrying OPA1 mutations and the optic neuropathy reported in some patients with MFN2 mutations, suggests the operation of common pathophysiological mechanisms. Hearing loss, found in patients carrying OPA1 mutations, was also detected in some patients carrying MFN2 mutations (Calvo et  al. 2009; Verhoeven et  al. 2006). A more complex form of CMT, associating peripheral neuropathy with cognitive impairment and defective vision was also reported in a family carrying a missense MFN2 mutation (Del Bo et al. 2008), with one member of the family additionally presenting spastic paraplegia. This observation, as well as the presence of pyramidal signs in patients, (Calvo et al. 2009; Zhu et al. 2005), demonstrates the possible involvement of the central nervous system in the pathophysiology of MFN2-related CMT. Recently the clinical spectrum of MFN2 mutations was further extended to the CMT1 or demyelinating forms of the disease, as well as to intermediate forms of the disease (Braathen et al. 2010).

6.3 Pathophysiology of OPA1- and MFN2-Associated Diseases The diversity and variable severity of the phenotypes associated with OPA1 and MFN2 mutations indicate the operation of complex pathogenic mechanisms, linking neuronal and mitochondrial pathophysiology. Considering the energy requirement of RGCs and the complexity of axonal transport, the specific structure of the RGCs and the length of the peripheral nerve are likely to be critical pathophysiological parameters. Moreover, mitochondrial involvement is highly complex since it concerns several essential, interdependent functions, such as energy production,

176

G. Lenaers et al.

apoptosis, oxidative stress, maintenance and expression of mtDNA, mitophagy, and cell signaling.

6.3.1 Mutation Spectrum of the OPA1 Gene The OPA1 gene, located on chromosome 3q28-29, spans approximately 100 kb and is composed of 31 exons, including three that undergo alternative splicing (4, 4b and 5b) corresponding to a total of eight protein isoforms (Delettre et al. 2001). The main isoform, excluding exon 4b et 5b, encoded by a 2,880 nucleotide-long open reading frame, is a 960 amino acid protein ubiquitously expressed and located in the inter-membrane space, associated with the inner mitochondrial membrane (IMM). The main protein domains are the N-terminal mitochondrial targeting sequence (aa 1–88), a transmembrane domain (aa 97–115), the GTPase domain (aa 260–567), the dynamin central region (aa 568–901), the coiled-coil domain (aa 209–260), and the GTPase putative effector domain (aa 902–960) (Fig. 6.1). We have developed a locus specific database called eOPA1 (http://lbbma.univangers.fr/eOPA1/) as the main depository of OPA1 sequence variations (Ferre et al. 2005, 2009). As of October 2010, 224 pathogenic mutations have been published in this database. These mutations span all the exons, with the exception of exons 4, 4b and 5, in which no mutations have so far been reported. The exons encoding the GTPase domain and the C-terminal coiled-coil domain are more frequently mutated. Heterozygous mutations are mostly family specific, but the mutation c.2708_2711delTTAG in exon 27, is present in about 17% of cases. A total of 41% of the mutations results in premature termination of translation, 25% are missense mutations, 25% are in frame splice variants, and 6% are micro-deletions or duplications. In addition, large genomic rearrangements of OPA1 gene have been revealed by fluorescence in situ hybridization (FISH) or multiplex ligation-dependent probe amplification (MLPA); (Fuhrmann et  al. 2009, 2010; Toomes et  al. 2001; Yu-Wai-Man et al. 2010b). OPA1 mutations leading to splicing defects and premature stop codons induce non-sense mRNA decay, and the reduction of mRNA levels (Schimpf et al. 2008). These observations concord with the low mRNA levels found in fibroblasts from OPA1 patients and mouse models, with a decrease of up to 50% in OPA1 transcripts; in one patient the entire OPA1 gene was found to be deleted, suggesting that haplo-insufficiency is the main pathophysiological mechanism in “pure” DOA (Alavi et al. 2007; Davies et al. 2008; Olichon et al. 2007b). Consequently, phenotype-genotype correlations have been difficult to establish for most OPA1 mutations leading to DOA, suggesting that other gene modifiers, such as those involved in mRNA decay, may modulate the severity of the disease. Nevertheless, in syndromic forms of DOA, missense mutations have been systematically identified downstream to, or within, the GTPase domain, with one exception situated in the GED C-terminal domain (Amati-Bonneau et al. 2008; Hudson et al. 2008). In these cases, the OPA1 protein levels were always normal,

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 177

Fig. 6.1  (continued)

178

G. Lenaers et al.

suggesting a mutant dominant negative effect. Interestingly, the R445H mutation affecting a highly conserved amino acid in the GTPase domain is responsible for the clinical presentation of most of these “DOA+” patients, ranging from DOA and deafness to severe multi-systemic disorders.

6.3.2 Mutation Spectrum of the MFN2 Gene The MFN2 gene, located on chromosome 1p36.2, comprises 19 exons (Zuchner et  al. 2004). The main transcript, which is 4,685 nucleotides long, including an open reading frame containing 2,271 nucleotides, encodes a protein composed of 757 amino acids with a Ras domain (aa 84–99), a GTPase domain (aa 100–260), two transmembrane domains (aa 628–647), two hydrophobic heptad repeat domains (HR1 and HR2, aa 405–425 and 702–742) allowing coiled-coil interaction and fusion between adjacent mitochondria. CMT2A is inherited as a dominant trait caused by heterozygous mutations in the MFN2 gene; however, in some rare cases, compound heterozygous or homozygous mutations have been found (Calvo et  al. 2009; Nicholson et  al. 2008). To date, more than 70 mutations have been identified (see http://www.molgen.ua.ac.be/ CMTMutations/Mutations/Mutations.cfm) (Calvo et al. 2009), the large majority of these (>90%) being missense mutations, whereas only a few truncating mutations have been reported; these later mutations target either the last 10 amino acids of the C-terminal domain, the last 340 amino acids including the two TM and the HR2 domains, or result from splicing defects. So far, no duplication or deletion in the MFN2 gene has been reported. The distribution of mutations in the MFN2 protein is not homogeneous, indeed most of the mutations (>85%) affect the half of the N-terminal domain of the protein including the GTPase domain, and the two ­surrounding domains (aa 1–99 and 260–405), the structure and function of which remain unknown. Conversely, from the first coiled-coil HR1 domain to the carboxy-end of the protein, only a few mutations have been identified, none of which affect the transmembrane TM1 and TM2 domains (Cartoni and Martinou 2009). This suggests that the dysfunctional element of the mutated MFN2 protein is strictly located on the external side of the outer mitochondrial membrane (OMM), i.e. towards the cytoplasm.

Fig. 6.1  (continued) Mutation spectrum in OPA1 and MFN2 proteins and clinical presentations of syndromic DOA and CMT2A. OPA1 (left) and MFN2 (right) proteins are schematically represented. MTS: mitochondrial targeting sequence; TM: transmembrane domain; CC/HR: coiled coil domains/ heptad repeats; RAS: Ras binding domain; GTPase: guanosine tri-phosphate hydrolase domain; and GED: GTPase effector domain. In addition, for OPA1, the domains corresponding to the alternate spliced exons 4b and 5b are shown. The domains mutated in OPA1 and MFN2 in “pure” DOA (left) and CMT2A (right) patients are indicated by dotted lines. Mutations responsible for peculiar syndromic phenotypes are represented by arrowheads of different colors. DOA: Dominant optic atrophy; CMT2A: Charcot Marie Tooth type 2A; +MS: plus multiple ­sclerosis; +SP: plus spastic paraplegia; ROA: recessive optic atrophy; PEO: progressive external ophthalmoplegia (without DOA); +ND: plus neurosensorial deafness; +Tr: plus tremor; and + Enc: plus encephalopathy

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 179

On the basis of the accumulated clinical and genetic data collected over the past few years, it now appears possible to evaluate genotype-phenotype correlations, although this should be done with caution since the severity of the disease remains highly variable between individuals sharing the same molecular defect. Thus, patients presenting a CMT2A syndrome associated with optic atrophy carry ­mutations in the domain (aa 260–405) located between the GTPase and the HR1 domains. Similarly, CMT2A patients with neurosensorial hearing loss carry mutations in the GTPase domain, while the two most common mutations at position R94Q (Ras domain) and R104W (GTPase domain) are responsible for CMT2A plus optic atrophy and hearing loss. In addition, CMT2A patients with tremor carry MFN2 mutations in the amino-terminal domain upstream of the GTPase domain. Finally, one of the two splicing mutations in MFN2, which induce erroneous ­splicing of intron 13 leading to truncated forms of the MFN2 protein, is associated with a subacute and severe fatal encephalopathy that follows shortly the onset of the polyneuropathy (Boaretto et al. 2010) (Fig. 6.1). Taken together, these findings have three important consequences on the pathophysiological hypothesis concerning the relationship between CMT2A and the mutant MFN2 protein. Firstly, the different amino-terminal domains of MFN2 have distinct functions. Secondly, MFN2 must be anchored to the outer mitochondrial membrane by the two TM domains and exerts its pathological effect by interacting with partners thanks to the integrity of the HR domains. Thirdly, CMT2A is caused by a dominant negative pathophysiological mechanism depending on the mutant MFN2 protein. This idea is supported by the fact that the amount of MFN2 remains unchanged in fibroblasts from CMT2A patients (Amiott et  al. 2008). Moreover, the heterozygous MFN2+/− mouse does not present any symptom of the disease (Chen et al. 2003), thus ruling out the hypothesis that haplo-insufficiency might be the main causative mechanism of CMT2A.

6.3.3 Consequences on Mitochondrial Network Morphology The dynamin OPA1, and the mitofusins MFN1 and MFN2, are the only actors known to be involved in the fusion of the mitochondrial network (Song et al. 2009; Chen and Chan 2010). The mitofusins are responsible for tethering adjacent mitochondria and promoting the fusion of the OMMs in which they are embedded (Rojo et al. 2002). OPA1, which is closely associated with the IMM in the intermembrane space and cristae (Olichon et al. 2002), is directly involved in the fusion of the IMM (Song et al. 2009). We may therefore expect OPA1 and MFN2 mutations to lead to various defects in mitochondrial network structure and function in DOA and CMT2A. Primary cultures of skin fibroblasts from DOA and CMT2A patients have been studied to investigate the pathophysiology of these diseases. Simultaneously, mutated alleles of OPA1 and MFN2 were expressed in common cell lines, such as HeLa and MEF cells, to study the consequences of the mutations in a homogeneous genetic background; RNA-interference experiments targeting the OPA1 and

180

G. Lenaers et al.

MFN2 genes were also carried out. Similar gene overexpression and silencing experiments were performed in other cell types, such as purified neurons from the retina, the cerebellum, or the spinal cord. Finally, murine models have recently allowed the investigation of the consequences of OPA1 and MFN2 mutations on animal physiology. The convergence of the results obtained with these different models has helped to unravel the pathophysiology of DOA and CMT2A. In fibroblasts from patients ­carrying OPA1 mutations leading to haplo-insufficiency, there were practically no detectable defects in mitochondrial morphology under normal growth conditions (Olichon et al. 2007b). Nevertheless, when galactose was substituted for glucose in the cell culture medium to force the cells to use mitochondrial OXPHOS, OPA1 fibroblasts showed a fragmented mitochondrial network compared to controls (Zanna et al. 2008). This phenotype with highly punctuated mitochondria was also prominent in fibroblasts with OPA1 dominant negative mutations found in DOA+, independently of the media used, suggesting that missense OPA1 alleles have a more severe effect on fusion than the 50% reduction in OPA1 abundance. (Amati-Bonneau et al. 2005). These findings were confirmed in HeLa cells in which the expression of different missense OPA1 mutations induced severe fragmentation, whereas that of truncative OPA1 mutations had no effect on the mitochondrial network. In cells carrying truncative OPA1 mutations, the level of OPA1 expression was markedly reduced, suggesting that the truncated protein is unstable, thus reinforcing the hypothesis of haplo-insufficiency in “pure” DOA (Olichon et al. 2007b). In addition, some elegant recent research has demonstrated that OPA1 associates with cardiolipins, which are negatively charged phospholipids enriched in the IMM, to stimulate GTP hydrolysis and membrane tubulation. Interestingly, different missense OPA1 mutations induce specific defects in membrane scaffolding, thus revealing for the first time the possible consequences of these mutations on the disorganization of the IMM (Ban et al. 2010). This is of further relevance to the apoptotic process since the release of cytochrome c during programmed cell death depends on major ­structural changes of the IMM, associated with the processing and release of OPA1 in the cytoplasm. These findings are concordant with the results of silencing specific OPA1 isoforms, which induce independent effects on mitochondrial dynamics and apoptosis, suggesting that these processes operate independently in mitochondria (Olichon et  al. 2003, 2007a). Similar experiments on mouse RGCs suggest that OPA1 is required for the mitochondrial fusion and necessary to protect these cells from apoptotic insults (Kamei et al. 2005) and for the distribution of mitochondria in the somas and dendrites of the RGCs (Williams et al. 2010). The investigation of the dysfunction of MFN2 in fibroblasts from CMT2A patients revealed limited alteration of the mitochondrial network, suggesting that the MFN2 defect in fibroblast cells either has mild consequences on the equilibrium between fusion and fission, or is readily compensated by other processes. It is thus possible that the role of MFN2 in fibroblasts is secondary to that of MFN1, and that the eightfold higher GTPase activity of MFN1 compared to that of MFN2 is responsible for most of the membrane tethering and fusion activity in fibroblasts

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 181

(Amiott et  al. 2008; Ishihara et  al. 2004). More importantly, the absence of any major effect on the structure of the mitochondrial network may be related to the lack of susceptibility to apoptosis found in MFN2 fibroblasts (Guillet et  al. 2010). Similarly, since MFN2 is responsible for tethering the endoplasmic reticulum to the mitochondrial network, the absence of network modification suggests that the physical relationship between the ER and the mitochondrial network, as well as the calcium uptake by mitochondria, remain unaffected in MFN2 mutated cells (de Brito and Scorrano 2008). Conversely, the overexpression of MFN2 alleles either with truncated TM or HR1/2 domains, or mutated in the GTPase domain, lead to a dominant negative effect on fusion, leaving fragmented mitochondria that remain merely apposed but unfused (Eura et al. 2003; Honda et al. 2005). Interestingly, the overexpression of wild-type MFN2 induces mitochondrial fragmentation and ­dysfunction with concomitant cell death, illustrating the need for a balance between mitochondrial fusion and fission (Huang et al. 2007). Similarly, the expression of MFN2-mutated proteins in neurons induces the clustering of fragmented mitochondria in the soma, by slowing down the physiological anterograde and retrograde mitochondrial transport in axons, without ­affecting the transport of other cargoes (Baloh et al. 2007; Misko et al. 2010). These phenomena clearly play a role in the peculiar cell structure of RGCs and on the axonal length of neurons in patients affected by CMT2A.

6.3.4 Consequences on Mitochondrial Metabolism Cells carrying OPA1 mutations present a significant deficit of mitochondrial respiration. The study of fibroblasts from DOA patients reveals a coupling defect associated with the reduction of membrane potential and respiratory complex activities (Chevrollier et al. 2008). The reduction of the quantity of OPA1 protein in these fibroblasts can alter the structure of the mitochondrial cristae and reduce the impermeability of the inner membrane, allowing a mild dissipation of the proton gradient (Olichon et  al. 2003, 2007a). This observation is in agreement with the finding of a partial uncoupling of the respiration in OPA1-mutated fibroblasts, correlated with the severity of the disease, (Chevrollier et al. 2008). Moreover, the calcium clearance was found to be impaired in RGCs silenced for OPA1, underscoring the importance of membrane potential in importing cytoplasmic calcium into the matrix (Dayanithi et al. 2010). Similarly, mild alteration of the structure of the IMM was found to disrupt the organization between the respiratory complexes, explaining the reduction of complex IV activity observed in DOA fibroblasts, that is compensated in basic growth condition to allowing normal ATP production (Chevrollier et  al. 2008). However, when galactose was used instead of glucose, forcing cells to rely on OXPHOS alone, OPA1-mutated fibroblasts presented impaired complex I-driven production of ATP compared to control cells (Zanna et al. 2008). A similar alteration of ATP production was reported in DOA patients, with an in vivo study of the calf muscle by phosphorus magnetic resonance spectroscopy showing that the de novo synthesis of phosphocreatine was delayed after exercise

182

G. Lenaers et al.

(Lodi et al. 2004, 2010). Taken together, these observations support the hypothesis of an energetic defect as one of the principal causes of DOA. Hence, a reduction in respiration coupling may progressively impair neuronal functions, particularly in the transmission of action potentials and the transport of cargoes along the axons. Although the MFN2 mutations identified in CMT2A affect the cytoplasmic domains of the protein, it is now well established that they are responsible for severe defects within mitochondria, and not only in the process of oxidative respiration. MFN2 mutations reduce mitochondrial membrane potential and induce partial uncoupling of respiration, reducing ATP production. This process appears to be compensated by a higher respiratory rate, although in normal conditions of cell culture, i.e. in the presence of glucose, respiratory complex activities were found to be normal in fibroblasts from CMT2A patients (Loiseau et  al. 2007). The mitochondrial phenotype and energetic metabolism of such fibroblasts cultured in a galactose medium have not yet been investigated, but MFN2 mutations may induce subtle defects under restrictive oxidative metabolism. The effect of silencing MFN2, studied in L6E9 myotubes, revealed increased complex II activity and a shift of mitochondrial substrate consumption with a reduced input of pyruvate, glucose, and fatty acids into mitochondrial fuelling. The amounts of the respiratory complexes present, except for complex IV, appear to be linked to the integrity of the first 600 amino acids of MFN2, independently of the fusion activity or attachment of the protein to the OMM (Pich et al. 2005). In addition, the level of expression of ANT3, an adenine nucleotide translocase embedded in the IMM, depends on the integrity of MFN2, suggesting that in parallel to its fusion activity, MFN2 could modulate the mitochondrial ATP/ADP exchanges (Guillet et al. 2010). These findings suggest that MFN2 has an indirect regulatory function in modulating gene expression, uncoupled from its fusion activity in the mitochondrial network. This suggests that MFN2 contributes to a regulatory cross-talk pathway linking mitochondrial dynamics to nuclear gene expression. Studies on dorsal root ganglion cells (DRG), expressing the mutated forms of MFN2, have revealed normal ATP levels and oxidative respiration, although the mitochondrial network appeared to be affected with fragmented and aggregated mitochondria surrounding the nucleus (Baloh et al. 2007). These results further illustrate the dichotomy of MFN2 functions, making it critical to choose appropriate cellular models for studying the pathophysiological mechanisms of MFN2 mutations.

6.3.5 Consequences on the Integrity of MtDNA It has been recently demonstrated that mitochondrial fusion plays an important role in the maintenance of the integrity of the mitochondrial genome. Indeed, studies of muscle biopsies from patients with syndromic DOA revealed the presence of mtDNA deletions; however, such deletions were not found in fibroblasts from these

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 183

patients (Amati-Bonneau et al. 2008; Hudson et al. 2008). The characterization of the deletion boundaries showed that the deletions were similar to those observed in other disorders associated with mtDNA deletions, suggesting a common mechanism underlying the instability of mtDNA. Another study found similar mtDNA deletions in patients with non-syndromic DOA, suggesting that there may be no correlation between the clinical presentation of DOA and the presence of mtDNA deletions (Yu-Wai-Man et al. 2010c). One study indicated that lymphocytes from OPA1 patients contained smaller amounts of mtDNA than controls (Kim et al. 2005). However, this finding was not confirmed in any other studies involving fibroblasts from OPA1 patients or OPA1 mouse models. In contrast, a recent study revealed that the amount of mtDNA increased two to fourfold in skeletal fibers with even higher levels of amplification in “DOA+” samples (Yu-Wai-Man et al. 2010c). More importantly, in addition to the increased mtDNA copy number, the ratio of wild-type mitochondrial genome versus mutant genome decreased, supporting the hypothesis that the inhibition of mitochondrial dynamics may somehow favor mutant over wild-type mtDNA proliferation (Malena et al. 2009). Few studies have addressed the consequences of MFN2 mutations on mtDNA integrity. To date, two reports on fibroblasts from CMT2A patients have suggested that mtDNA content and integrity are unaffected by pathological MFN2 alleles (Amiott et al. 2008; Guillet et al. 2010); however, as in the case of fibroblasts from OPA1 patients, the fibroblasts from CMT2A patients may not offer the best approach for the investigation of mtDNA integrity. Since MFN2 is not uniformly expressed in all tissues, we cannot rule out the contribution of neuronal mtDNA deletions to the pathophysiological mechanism of CMT2A, particularly in syndromic cases. Indeed, the deletion of both mitofusins in mouse muscle has drastic effects on the quantitative and qualitative integrity of mtDNA, highlighting the importance of mitofusins in mtDNA maintenance (Chen et al. 2010).

6.3.6 Mouse Models of DOA and CMT2A The development and investigation of animal models of DOA and CMT2A are of critical importance to gain deep insights in the pathophysiological mechanisms of these diseases. Indeed, in vitro primary cultures of RGCs or peripheral neurons fail to reproduce several of the main constraints imposed on these cells under physiological conditions. For example, in the eye the RGC axons are unmyelinated and are constantly solicited for the transduction of visual information. Moreover, these axons are subjected to the assaults of the intra-ocular pressure and exposure to daylight, which generate oxidative stress. Similarly, the peripheral neurons have the longest axons in the body. This means that axonal transport, including that of mitochondria, as well as the energy distribution must be maintained at a high level of efficiency to avoid jeopardizing cell survival. Thus, only animal models of DOA and CMT2A can satisfactorily reproduce these neuronal specificities, allowing the chronological

184

G. Lenaers et al.

follow-up of the impact of the disease on mitochondrial pathophysiology, cell survival and neuronal functionality. Two OPA1 mouse models have been created from an N-ethyl-N-nitrosurea (ENU) mutagenized library, each encoding a protein with a truncation in the GTPase domain. Molecular analysis has revealed that the mutated allele is counter-selected so that haplo-insufficiency is the princeps mechanism driven by these mutations (Alavi et al. 2007; Davies et al. 2007). Visual assessment tests were performed in order to characterize the clinical phenotype of these animals, but no significant alteration of the electroretinogram was observed, whatever the age of the animals. In contrast, a reduction of amplitude of the scotopic, but not the photopic, visual evoked potential (VEP) was detected in the older animals, reflecting an altered transduction of visual information from the retina to the brain (Heiduschka et al. 2010). This reduction of the VEP followed a decrease in visual acuity detected in 12-month-old mice by means of an optokinetic drum. The absence of any delay in the VEP suggests that RGC dysfunction runs parallel to the phenomenon of apoptosis, which progresses from the soma to the synapse. Histological examination of the retina revealed a decrease in the dendritic length of the RGC-ON subpopulation, but not that of the RGC-OFF subpopulation (Williams et al. 2010). Since the RGC-ON neurons contact bipolar cells in sublamina b of the inferior parietal lobule, the decreased dendritic length may reflect retarded dendritic outgrowth due to defective mitochondrial transport. Moreover, autophagosomes have been found in the RGC layer of DOA mouse models, attesting the autophagy of RGCs and the clearance of apoptotic by-products (White et al. 2009). In the optic nerve of older DOA mouse models, some abnormal myelination was observed beyond the lamina cribrosa, with an increase in the number of microglial cells together with the loss of the largest axons. In parallel, alterations of the IMM structure were observed in the OPA1(+/−) animals with greater numbers of cristae, forming vesicule-like structures. However, no abnormalities were reported in the mitochondrial network or abundance of the organelles, and no mtDNA deletions were found, but whether the mitochondrial activity is altered in these models has yet to be determined. Surprisingly, some secondary clinical, neurological and neuromuscular symptoms were observed, although in a very mild presentation. Thus, locomotor activity was reduced and some tremor was observed in the older mouse models, but no hearing loss was detected. Similarly, although food intake was normal during life, lower weight and less body fat were found in old OPA1 mice, possibly reflecting the lasting uncoupling of mitochondrial respiratory activity (Alavi et al. 2009). No MFN2 knock-in animal models have yet been developed. However, two transgenic mouse strains, expressing a pathological MFN2 allele only in neurons, have been developed using neuron-specific promoters. One of these mouse strains has a mutation in the GTPase domain (T105 M) (Detmer et al. 2008), and the other has a mutation upstream of the GTPase domain (R94Q) (Cartoni et al. 2010), corresponding to two commonest mutations found in CMT2A patients. Both mouse models present locomotor impairment and hindleg gait defects, mimicking the major syndrome of the disease. Nevertheless, the phenotype and its severity result from a subtle but significant difference in the level of expression of the pathological

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 185

alleles. Indeed, the expression of the T105M allele induces a dose-dependent gradient of mitochondria, with several aggregates around the nucleus and clusters in the proximal axonal region, but few in the distal region. Conversely, the expression of R94Q leads to an increase in the number of mitochondria, particularly in the distal part of axons of small diameter, thus reproducing some of the findings in CMT2A patients and suggesting that the underlying pathophysiological mechanisms may be similar. In parallel, the T105M expression induced a reduction of the number of motoneuron axons, whereas the number of small axons increased in the R94Q mouse model (Cartoni et al. 2010; Detmer et al. 2008). Thus, both models mimic the clinical presentation of CMT2A but possibly with different pathophysiological mechanisms.

6.4 Therapeutic Strategies for DOA and CMT2A DOA and CMT2A remain orphan pathologies with no currently available treatment. As in the case of several other genetic diseases, the two therapeutic approaches that may be envisaged are based on gene therapy and pharmacological therapy. The direct involvement of OPA1 in the function and regulation of mitochondrial networks in patients with DOA, and that of MFN2 in patients with CMT2A, suggests some promising therapeutic strategies.

6.4.1 Gene Therapy The theory of gene therapy is fairly straightforward. It states that if a pathological condition is due to the presence of reduced quantities of a certain protein then the normal protein content may be restored by using an appropriate vector carrying the transgene encoding the protein. Could this theory have a practical application in diseases such as DOA and CMT2A? In most cases of DOA, the disease is caused by OPA1 haplo-insufficiency. Moreover, RGC somas, which form the innermost layer of the retina facing the vitreous humor, are readily accessible to micro-injections into the vitreous humor of the eye globe. Thus, microinjections of a vector with an affinity for RGCs and encoding the OPA1 gene should enhance the protein content and restore its full activity in the mitochondrial network of the target cells. However, this attractive strategy remains conditioned by the possibility of controlling the ectopic production of OPA1 since overexpression of this protein might be more deleterious than beneficial. In addition, it would be of capital importance to choose the appropriate OPA1 isoform to be expressed, since all eight isoforms of OPA1 are expressed in the RGCs. The tools required to address these problems have now been assembled and the first pre-clinical trials on mouse models should soon be under way.

186

G. Lenaers et al.

In contrast to the therapeutic strategy outlined for DOA, CMT2A offers a much greater challenge. Since the disease is caused by the dominant negative effect of the mutated MFN2 protein on the wild-type protein, it is far from obvious that the pathological phenotype would be rescued by the ectopic expression of MFN2. Indeed, fibroblast studies have shown that the overexpression of MFN2 does not in itself complement the mutated MFN2, whereas that of MFN1 does so by forming hetero-oligomeric complexes with the mutated MFN2 protein (Detmer and Chan 2007). The specific effects of the overexpression of MFN1 and MFN2 in peripheral nerves expressing mutated MFN2 have yet to be investigated. Another important aspect that needs to be considered is the delivery of the therapeutic vectors to the soma of the peripheral neurons affected in CMT2A. It would obviously be difficult to design a practical strategy that would allow the exhaustive transduction of a ­vector to neurons located all along the spinal cord and in the dorsal root ganglia. Furthermore, the current mouse models of CMT2A were obtained by transgenesis of a mutated allele, rather than by knocking-in the MFN2 gene. For all these ­reasons, we are still far from the point of testing the benefits of gene therapy for CMT2A patients.

6.4.2 Pharmacological Therapy Pharmacological therapy offers an interesting alternative to gene therapy. Although no pharmacological therapy could be expected to correct gene mutations, it may prove effective in dealing with various mechanisms affected by these genetic ­diseases. In particular, it would be important to elucidate the factors involved in the production and function of OPA1 in DOA, and MFN2 in CMT2A, so as to determine how their activities could be appropriately modulated by drugs. The pathways regulating the production and function of MFN2 (Fig.  6.2) and OPA1 (Fig.  6.3) suggest the eventuality of novel therapeutic approaches to the treatment of CMT2A and DOA. Whereas little is known about the regulation of the expression of the OPA1 gene, three pathways have been identified in skeletal muscle for the regulation of the MFN2 gene. They involve PGC1a and PGC1b, which are PPARg co-activators, the estrogen-related receptor a, which acts in synergy with PPARg and the antitumoral protein p53; all of these induce the transcription of MFN2 by binding to its promoter (Wang et al. 2010; Zorzano 2009). It may therefore be hypothesized that drugs activating these pathways, such as glitazones, resveratrol and estrogen mimetics, could increase the abundance of MFN2 in peripheral neurons. However, it is not clear whether the increase of mutated as well as wild-type MFN2 proteins would exert a positive or negative effect on the basic functions of the protein. It further remains to be demonstrated that these pathways are active in peripheral neurons, and that they can be stimulated through oral treatment. A second pathway regulates the turnover of OPA1 and MFN2 according to their rate of degradation. The degradation may target a particular protein, or globally

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 187

Fig. 6.2  MFN2 regulatory pathways. The processes shown in rectangles indicate the different pathways regulating the expression, the stability and the activities of MFN2. The processes shown in ovals indicate mechanisms that depend upon the MFN2 activities affected in CMT2A. Full arrows represent direct functional involvements of MFN2; dotted arrows represent links between processes that are indirectly regulated by MFN2; and small curved arrows indicate the contribution of specific partners to the control of the activities and abundance of MFN2

target the mitochondria. The term “mitophagy” refers specifically to the degradation of impaired mitochondria. Both forms of degradation require Pink1 and Parkin proteins, which are mutated in hereditary forms of Parkinson’s disease (Matsuda and Tanaka 2010), for the ubiquitylation of specific mitochondrial proteins. Whether OPA1 is a direct target of Pinkl and Parkin proteins in vertebrate cells is unknown, but the abundance of MFN2 is clearly regulated by this pathway, since the loss of function of either Pink1 or Parkin stabilizes MFN2 and induces elongation of the mitochondrial network (Detmer and Chan 2007). In addition, some pathogenic mutations in MFN2 are known to stabilize the abundance of the protein because of reduced ubiquitylation (Amiott et al. 2009). Thus, it is still difficult to say whether MFN2 ubiquitylation should be stimulated to reduce the amount of the mutated protein, or whether it should be inhibited to increase the ratio of the wild-type to mutated protein. These possibilities could be readily tested in fibroblasts from

188

G. Lenaers et al.

Fig.  6.3  OPA1 regulatory pathways. The processes shown in rectangles indicate the different pathways regulating the expression, the stability and the activities of OPA1. The processes shown in ovals indicate mechanisms that depend upon the OPA1 activities affected in DOA. Full arrows represent direct functional involvements of OPA1; dotted arrows represent links between processes that are indirectly regulated by OPA1; and small curved arrows indicate the contribution of specific partners to the control of the activities and abundance of OPA1

CMT2A patients by means of drugs or siRNA targeting the Parkin E3-ubiquitin ligase, and evaluating MFN2 turnover, the mitochondrial network structure and respiration parameters. Mitophagy follows a quality-control process, called “kiss-and-run”, which evaluates the functionality of mitochondria by assessing their membrane potential (Twig et  al. 2008). OPA1 is required for this process, and its downregulation in DOA could affect its sensor activity, leading to the progressive accumulation of impaired mitochondria (Liu et al. 2009). Thus, stabilizing the amount of OPA1 or increasing its expression could favor the maintenance of efficient mitochondria and sustain the perennity of their distribution. A third pathway controlling the activities of OPA1 and MFN2 corresponds to the post-translational modifications of these proteins. It is not yet clear whether MFN2 undergoes other modifications than the ubiquitinylation that controls its turn-over. But the involvement of OPA1 in mitochondrial fusion is clearly regulated by an amino-terminal cleavage that in physiological conditions affects about 50% of all isoforms, except for those, including the alternate-spliced exon 4b, which are 100%

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 189

cleaved (Song et al. 2007). Under conditions of stress, all OPA1 isoforms undergo cleavage (Baricault et  al. 2007; Guillery et  al. 2008) leading to a markedly fragmented mitochondrial network and increasing susceptibility to apoptosis (Griparic et al. 2007). Thus, the reduction of OPA1 cleavage, through the modulation of the inner membrane proteases, YME1L (Song et al. 2007) and OMA1 (Ehses et al. 2009; Head et  al. 2009), involved in this proteolytic process, should offer an interesting approach towards increasing the resistance to apopotic stimuli. This could further be promoted by stimulating the activity of the Phb1 and Phb2 prohibitins, which control the proteases involved in OPA1 processing (Merkwirth et al. 2008). The three pathways described above sustain OPA1 and MFN2 activity by increasing their active amounts in mitochondria. However, the drugs capable of acting on these pathways are far from being adequately characterized. Other strategies for the treatment of DOA and CMT2A diseases may rely on products acting on the downstream mitochondrial activities affected by OPA1 and MFN2 mutations. Thus, patients may expect to benefit from strategies, compatible with chronic treatment, that would stimulate mitochondrial fusion, foster mitochondrial transport, restore membrane potential, boost mitochondrial respiration and ATP production, promote calcium homeostasis, and inhibit the apoptotic process. Drugs targeting these processes remain to be characterized, but the methodology that would enable their identification is now available. High-throughput drug screening on fibroblasts from DOA and CMT2A patients would help to determine the most potent drugs for these diseases. Further testing would allow identification of the precise targets and pathways stimulated or inhibited by these drugs. Furthermore, the drugs identified as potentially active against DOA and CMT2A would probably also prove useful for the treatment of other mitochondrial diseases. To conclude this section on therapeutic strategies, it should be mentioned that prosthetic therapy using a cochlear implant has been successfully used to treat neurosensorial deafness in syndromic DOA patients with the OPA1-R445H mutation, leading to marked auditive improvement (Huang et al. 2009). This prosthetic approach, unfortunately not applicable in the optic and peripheral neuropathies in cases of DOA and CMT2A, could nevertheless be used to treat the neurosensorial deafness of syndromic patients with either disease. Acknowledgements  We are grateful to Kanaya Malkani for critical reading and comments on the manuscript. Our work is supported by INSERM, the University Hospital of Angers (PHRC 04–12), the University of Angers, France; and by grants from the following patients’ associations: “Association contre les Maladies Mitochondriales (AMMi)”, “Ouvrir les Yeux (OLY)”, “Retina France” and “Union Nationale des Aveugles et Déficients Visuels (UNADEV)”.

References Alavi, M. V., Bette, S., Schimpf, S., Schuettauf, F., Schraermeyer, U., Wehrl, H. F., Ruttiger, L., Beck, S. C., Tonagel, F., Pichler, B. J., Knipper, M., Peters, T., Laufs, J. & Wissinger, B. (2007) A splice site mutation in the murine Opa1 gene features pathology of autosomal dominant optic atrophy. Brain, 130, 1029–42.

190

G. Lenaers et al.

Alavi, M. V., Fuhrmann, N., Nguyen, H. P., Yu-Wai-man, P., Heiduschka, P., Chinnery, P. F. & Wissinger, B. (2009) Subtle neurological and metabolic abnormalities in an Opa1 mouse model of autosomal dominant optic atrophy. Exp Neurol, 220, 404–9. Alexander, C., Votruba, M., Pesch, U. E., Thiselton, D. L., Mayer, S., Moore, A., Rodriguez, M., Kellner, U., Leo-Kottler, B., Auburger, G., Bhattacharya, S. S. & Wissinger, B. (2000) OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat Genet, 26, 211–5. Amati-bonneau, P., Guichet, A., Olichon, A., Chevrollier, A., Viala, F., Miot, S., Ayuso, C., Odent, S., Arrouet, C., Verny, C., Calmels, M. N., Simard, G., Belenguer, P., Wang, J., Puel, J. L., Hamel, C., Malthiery, Y., Bonneau, D., Lenaers, G. & Reynier, P. (2005) OPA1 R445H mutation in optic atrophy associated with sensorineural deafness. Ann Neurol, 58, 958–63. Amati-bonneau, P., Odent, S., Derrien, C., Pasquier, L., Malthiery, Y., Reynier, P. & Bonneau, D. (2003) The association of autosomal dominant optic atrophy and moderate deafness may be due to the R445H mutation in the OPA1 gene. Am J Ophthalmol, 136, 1170–1. Amati-bonneau, P., Valentino, M. L., Reynier, P., Gallardo, M. E., Bornstein, B., Boissiere, A., Campos, Y., Rivera, H., De la Aleja, J. G., Carroccia, R., Iommarini, L., Labauge, P., Figarellabranger, D., Marcorelles, P., Furby, A., Beauvais, K., Letournel, F., Liguori, R., La Morgia, C., Montagna, P., Liguori, M., Zanna, C., Rugolo, M., Cossarizza, A., Wissinger, B., Verny, C., Schwarzenbacher, R., Martin, M. A., Arenas, J., Ayuso, C., Garesse, R., Lenaers, G., Bonneau, D. & Carelli, V. (2008) OPA1 mutations induce mitochondrial DNA instability and optic atrophy ‘plus’ phenotypes. Brain, 131, 338–51. Amiott, E. A., Cohen, M. M., Saint-georges, Y., Weissman, A. M. & Shaw, J. M. (2009) A mutation associated with CMT2A neuropathy causes defects in Fzo1 GTP hydrolysis, ubiquitylation, and protein turnover. Mol Biol Cell, 20, 5026–35. Amiott, E. A., Lott, P., Soto, J., Kang, P. B., Mccaffery, J. M., Dimauro, S., Abel, E. D., Flanigan, K. M., Lawson, V. H. & Shaw, J. M. (2008) Mitochondrial fusion and function in CharcotMarie-Tooth type 2A patient fibroblasts with mitofusin 2 mutations. Exp Neurol, 211, 115–27. Baloh, R. H., Schmidt, R. E., Pestronk, A. & Milbrandt, J. (2007) Altered axonal mitochondrial transport in the pathogenesis of Charcot-Marie-Tooth disease from mitofusin 2 mutations. J Neurosci, 27, 422–30. Ban, T., Heymann, J. A., Song, Z., Hinshaw, J. E. & Chan, D. C. (2010) OPA1 disease alleles causing dominant optic atrophy have defects in cardiolipin-stimulated GTP hydrolysis and membrane tubulation. Hum Mol Genet, 19, 2113–22. Barbet, F., Hakiki, S., Orssaud, C., Gerber, S., Perrault, I., Hanein, S., Ducroq, D., Dufier, J. L., Munnich, A., Kaplan, J. & Rozet, J. M. (2005) A third locus for dominant optic atrophy on chromosome 22q. J Med Genet, 42, e1. Baricault, L., Segui, B., Guegand, L., Olichon, A., Valette, A., Larminat, F. & Lenaers, G. (2007) OPA1 cleavage depends on decreased mitochondrial ATP level and bivalent metals. Exp Cell Res, 313, 3800–8. Boaretto, F., Vettori, A., Casarin, A., Vazza, G., Muglia, M., Rossetto, M. G., Cavallaro, T., Rizzuto, N., Carelli, V., Salviati, L., Mostacciuolo, M. L. & Martinuzzi, A. (2010) Severe CMT type 2 with fatal encephalopathy associated with a novel MFN2 splicing mutation. Neurology, 74, 1919–21. Braathen, G. J., Sand, J. C., Lobato, A., Hoyer, H. & Russell, M. B. (2010) MFN2 point mutations occur in 3.4% of Charcot-Marie-Tooth families. An investigation of 232 Norwegian CMT families. BMC Med Genet, 11, 48. Calvo, J., Funalot, B., Ouvrier, R. A., Lazaro, L., Toutain, A., de Mas, P., Bouche, P., Gilbertdussardier, B., Arne-bes, M. C., Carriere, J. P., Journel, H., Minot-myhie, M. C., Guillou, C., Ghorab, K., Magy, L., Sturtz, F., Vallat, J. M. & Magdelaine, C. (2009) Genotype-phenotype correlations in Charcot-Marie-Tooth disease type 2 caused by mitofusin 2 mutations. Arch Neurol, 66, 1511–6. Carelli, V., Ross-cisneros, F. N. & Sadun, A. A. (2004) Mitochondrial dysfunction as a cause of optic neuropathies. Prog Retin Eye Res, 23, 53–89.

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 191 Cartoni, R., Arnaud, E., Medard, J. J., Poirot, O., Courvoisier, D. S., Chrast, R. & Martinou, J. C. (2010) Expression of mitofusin 2(R94Q) in a transgenic mouse leads to Charcot-Marie-Tooth neuropathy type 2A. Brain, 133, 1460–9. Cartoni, R. & Martinou, J. C. (2009) Role of mitofusin 2 mutations in the physiopathology of Charcot-Marie-Tooth disease type 2A. Exp Neurol, 218, 268–73. Charcot, J. & Marie, P. (1886) Sur une forme particulière d’atrophie musculaire progressive souvent familiale débutant par les pieds et les jambes et atteignant plus tard les mains. Revue de Médecine, 6, 97–138. Chen, H. & Chan, D. C. (2010) Physiological functions of mitochondrial fusion. Ann N Y Acad Sci, 1201, 21–5. Chen, H., Detmer, S. A., Ewald, A. J., Griffin, E. E., Fraser, S. E. & Chan, D. C. (2003) Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J Cell Biol, 160, 189–200. Chen, H., Vermulst, M., Wang, Y. E., Chomyn, A., Prolla, T. A., Mccaffery, J. M. & Chan, D. C. (2010) Mitochondrial fusion is required for mtDNA stability in skeletal muscle and tolerance of mtDNA mutations. Cell, 141, 280–9. Chevrollier, A., Guillet, V., Loiseau, D., Gueguen, N., de Crescenzo, M. A., Verny, C., Eng, M. F., Dollfus, H., Odent, S., Milea, D., Goizet, C., Amati-bonneau, P., Procaccio, V., Bonneau, D. & Reynier, P. (2008) Hereditary optic neuropathies share a common mitochondrial coupling defect. Ann Neurol, 63, 794–8. Cohn, A. C., Toomes, C., Potter, C., Towns, K. V., Hewitt, A. W., Inglehearn, C. F., Craig, J. E. & Mackey, D. A. (2007) Autosomal dominant optic atrophy: penetrance and expressivity in patients with OPA1 mutations. Am J Ophthalmol, 143, 656–62. Cornille, K., Milea, D., Amati-bonneau, P., Procaccio, V., Zazoun, L., Guillet, V., El Achouri, G., Delettre, C., Gueguen, N., Loiseau, D., Muller, A., Ferre, M., Chevrollier, A., Wallace, D. C., Bonneau, D., Hamel, C., Reynier, P. & Lenaers, G. (2008) Reversible optic neuropathy with OPA1 exon 5b mutation. Ann Neurol, 63, 667–71. Davies, V. J., Hollins, A. J., Piechota, M. J., Yip, W., Davies, J. R., White, K. E., Nicols, P. P., Boulton, M. E. & Votruba, M. (2007) Opa1 deficiency in a mouse model of autosomal dominant optic atrophy impairs mitochondrial morphology, optic nerve structure and visual function. Hum Mol Genet, 16, 1307–18. Davies, V. J., Powell, K. A., White, K. E., Yip, W., Hogan, V., Hollins, A. J., Davies, J. R., Piechota, M., Brownstein, D. G., Moat, S. J., Nichols, P. P., Wride, M. A., Boulton, M. E. & Votruba, M. (2008) A missense mutation in the murine Opa3 gene models human Costeff syndrome. Brain, 131, 368–80. Dayanithi, G., Chen-kuo-chang, M., Viero, C., Hamel, C., Muller, A. & Lenaers, G. (2010) Characterization of Ca2+ signalling in postnatal mouse retinal ganglion cells: involvement of OPA1 in Ca2+ clearance. Ophthalmic Genet, 31, 53–65. de Brito, O. M. & Scorrano, L. (2008) Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature, 456, 605–10. Del Bo, R., Moggio, M., Rango, M., Bonato, S., D’angelo, M. G., Ghezzi, S., Airoldi, G., Bassi, M. T., Guglieri, M., Napoli, L., Lamperti, C., Corti, S., Federico, A., Bresolin, N. & Comi, G. P. (2008) Mutated mitofusin 2 presents with intrafamilial variability and brain mitochondrial dysfunction. Neurology, 71, 1959–66. Delettre, C., Griffoin, J. M., Kaplan, J., Dollfus, H., Lorenz, B., Faivre, L., Lenaers, G., Belenguer, P. & Hamel, C. P. (2001) Mutation spectrum and splicing variants in the OPA1 gene. Hum Genet, 109, 584–91. Delettre, C., Lenaers, G., Griffoin, J. M., Gigarel, N., Lorenzo, C., Belenguer, P., Pelloquin, L., Grosgeorge, J., Turc-carel, C., Perret, E., Astarie-dequeker, C., Lasquellec, L., Arnaud, B., Ducommun, B., Kaplan, J. & Hamel, C. P. (2000) Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat Genet, 26, 207–10. Detmer, S. A. & Chan, D. C. (2007) Complementation between mouse Mfn1 and Mfn2 protects mitochondrial fusion defects caused by CMT2A disease mutations. J Cell Biol, 176, 405–14.

192

G. Lenaers et al.

Detmer, S. A., Vande Velde, C., Cleveland, D. W. & Chan, D. C. (2008) Hindlimb gait defects due to motor axon loss and reduced distal muscles in a transgenic mouse model of Charcot-Marie-Tooth type 2A. Hum Mol Genet, 17, 367–75. Ehses, S., Raschke, I., Mancuso, G., Bernacchia, A., Geimer, S., Tondera, D., Martinou, J. C., Westermann, B., Rugarli, E. I. & Langer, T. (2009) Regulation of OPA1 processing and mitochondrial fusion by m-AAA protease isoenzymes and OMA1. J Cell Biol, 187, 1023–36. Eiberg, H., Kjer, B., Kjer, P. & Rosenberg, T. (1994) Dominant optic atrophy (OPA1) mapped to chromosome 3q region. I. Linkage analysis. Hum Mol Genet, 3, 977–80. Eura, Y., Ishihara, N., Yokota, S. & Mihara, K. (2003) Two mitofusin proteins, mammalian homologues of FZO, with distinct functions are both required for mitochondrial fusion. J Biochem, 134, 333–44. Ferre, M., Amati-bonneau, P., Tourmen, Y., Malthiery, Y. & Reynier, P. (2005) eOPA1: an online database for OPA1 mutations. Hum Mutat, 25, 423–8. Ferre, M., Bonneau, D., Milea, D., Chevrollier, A., Verny, C., Dollfus, H., Ayuso, C., Defoort, S., Vignal, C., Zanlonghi, X., Charlin, J. F., Kaplan, J., Odent, S., Hamel, C. P., Procaccio, V., Reynier, P. & Amati-bonneau, P. (2009) Molecular screening of 980 cases of suspected hereditary optic neuropathy with a report on 77 Novel OPA1 mutations. Hum Mutat, 30, E692–E705. Fuhrmann, N., Alavi, M. V., Bitoun, P., Woernle, S., Auburger, G., Leo-kottler, B., Yu-wai-man, P., Chinnery, P. & Wissinger, B. (2009) Genomic rearrangements in OPA1 are frequent in patients with autosomal dominant optic atrophy. J Med Genet, 46, 136–44. Fuhrmann, N., Schimpf, S., Kamenisch, Y., Leo-kottler, B., Alexander, C., Auburger, G., Zrenner, E., Wissinger, B. & Alavi, M. V. (2010) Solving a 50 year mystery of a missing OPA1 mutation: more insights from the first family diagnosed with autosomal dominant optic atrophy. Mol Neurodegener, 5, 25. Griparic, L., Kanazawa, T. & Van der Bliek, A. M. (2007) Regulation of the mitochondrial dynamin-like protein Opa1 by proteolytic cleavage. J Cell Biol, 178, 757–64. Guillery, O., Malka, F., Landes, T., Guillou, E., Blackstone, C., Lombes, A., Belenguer, P., Arnoult, D. & Rojo, M. (2008) Metalloprotease-mediated OPA1 processing is modulated by the mitochondrial membrane potential. Biol Cell, 100, 315–25. Guillet, V., Gueguen, N., Verny, C., Ferre, M., Homedan, C., Loiseau, D., Procaccio, V., Amatibonneau, P., Bonneau, D., Reynier, P. & Chevrollier, A. (2010) Adenine nucleotide translocase is involved in a mitochondrial coupling defect in MFN2-related Charcot-Marie-Tooth type 2A disease. Neurogenetics, 11, 127–33. Head, B., Griparic, L., Amiri, M., Gandre-babbe, S. & Van der Bliek, A. M. (2009) Inducible proteolytic inactivation of OPA1 mediated by the OMA1 protease in mammalian cells. J Cell Biol, 187, 959–66. Heiduschka, P., Schnichels, S., Fuhrmann, N., Hofmeister, S., Schraermeyer, U., Wissinger, B. & Alavi, M. V. (2010) Electrophysiological and histologic assessment of retinal ganglion cell fate in a mouse model for OPA1-associated autosomal dominant optic atrophy. Invest Ophthalmol Vis Sci, 51, 1424–31. Honda, S., Aihara, T., Hontani, M., Okubo, K. & Hirose, S. (2005) Mutational analysis of action of mitochondrial fusion factor mitofusin-2. J Cell Sci, 118, 3153–61. Hoyt, C. S. (1980) Autosomal dominant optic atrophy. A spectrum of disability. Ophthalmology, 87, 245–51. Huang, P., Yu, T. & Yoon, Y. (2007) Mitochondrial clustering induced by overexpression of the mitochondrial fusion protein Mfn2 causes mitochondrial dysfunction and cell death. Eur J Cell Biol, 86, 289–302. Huang, T., Santarelli, R. & Starr, A. (2009) Mutation of OPA1 gene causes deafness by affecting function of auditory nerve terminals. Brain Res, 1300, 97–104. Hudson, G., Amati-bonneau, P., Blakely, E. L., Stewart, J. D., He, L., Schaefer, A. M., Griffiths, P. G., Ahlqvist, K., Suomalainen, A., Reynier, P., Mcfarland, R., Turnbull, D. M., Chinnery, P. F. & Taylor, R. W. (2008) Mutation of OPA1 causes dominant optic atrophy with external ophthalmoplegia, ataxia, deafness and multiple mitochondrial DNA deletions: a novel disorder of mtDNA maintenance. Brain, 131, 329–37.

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 193 Ishihara, N., Eura, Y. & Mihara, K. (2004) Mitofusin 1 and 2 play distinct roles in mitochondrial fusion reactions via GTPase activity. J Cell Sci, 117, 6535–46. Johnston, P. B., Gaster, R. N., Smith, V. C. & Tripathi, R. C. (1979) A clinicopathologic study of autosomal dominant optic atrophy. Am J Ophthalmol, 88, 868–75. Kamei, S., Chen-kuo-chang, M., Cazevieille, C., Lenaers, G., Olichon, A., Belenguer, P., Roussignol, G., Renard, N., Eybalin, M., Michelin, A., Delettre, C., Brabet, P. & Hamel, C. P. (2005) Expression of the Opa1 mitochondrial protein in retinal ganglion cells: its downregulation causes aggregation of the mitochondrial network. Invest Ophthalmol Vis Sci, 46, 4288–94. Ke, T., Nie, S. W., Yang, Q. B., Liu, J. P., Zhou, L. N., Ren, X., Liu, J. Y., Wang, Q. & Liu, M. G. (2006) The G401D mutation of OPA1 causes autosomal dominant optic atrophy and hearing loss in a Chinese family. Zhonghua Yi Xue Yi Chuan Xue Za Zhi, 23, 481–5. Kerrison, J. B., Koenekoop, R. K., Arnould, V. J., Zee, D. & Maumenee, I. H. (1998) Clinical features of autosomal dominant congenital nystagmus linked to chromosome 6p12. Am J Ophthalmol, 125, 64–70. Kijima, K., Numakura, C., Izumino, H., Umetsu, K., Nezu, A., Shiiki, T., Ogawa, M., Ishizaki, Y., Kitamura, T., Shozawa, Y. & Hayasaka, K. (2005) Mitochondrial GTPase mitofusin 2 mutation in Charcot-Marie-Tooth neuropathy type 2A. Hum Genet, 116, 23–7. Kim, J. Y., Hwang, J. M., Ko, H. S., Seong, M. W., Park, B. J. & Park, S. S. (2005) Mitochondrial DNA content is decreased in autosomal dominant optic atrophy. Neurology, 64, 966–72. Kivlin, J. D., Lovrien, E. W., Bishop, D. T. & Maumenee, I. H. (1983) Linkage analysis in dominant optic atrophy. Am J Hum Genet, 35, 1190–5. Kjer, B., Eiberg, H., Kjer, P. & Rosenberg, T. (1996) Dominant optic atrophy mapped to chromosome 3q region. II. Clinical and epidemiological aspects. Acta Ophthalmol Scand, 74, 3–7. Kjer, P. (1959) Infantile optic atrophy with dominant mode of inheritance: a clinical and genetic study of 19 Danish families. Acta Ophthalmol Suppl, 164, 1–147. Kjer, P., Jensen, O. A. & Klinken, L. (1983) Histopathology of eye, optic nerve and brain in a case of dominant optic atrophy. Acta Ophthalmol (Copenh), 61, 300–12. Lawson, V. H., Graham, B. V. & Flanigan, K. M. (2005) Clinical and electrophysiologic features of CMT2A with mutations in the mitofusin 2 gene. Neurology, 65, 197–204. Li, C., Kosmorsky, G., Zhang, K., Katz, B. J., Ge, J. & Traboulsi, E. I. (2005) Optic atrophy and sensorineural hearing loss in a family caused by an R445H OPA1 mutation. Am J Med Genet A, 138A, 208–11. Liu, X., Weaver, D., Shirihai, O. & Hajnoczky, G. (2009) Mitochondrial ‘kiss-and-run’: interplay between mitochondrial motility and fusion-fission dynamics. Embo J, 28, 3074–89. Lodi, R., Tonon, C., Valentino, M. L., Iotti, S., Clementi, V., Malucelli, E., Barboni, P., Longanesi, L., Schimpf, S., Wissinger, B., Baruzzi, A., Barbiroli, B. & Carelli, V. (2004) Deficit of in vivo mitochondrial ATP production in OPA1-related dominant optic atrophy. Ann Neurol, 56, 719–23. Lodi, R., Tonon, C., Valentino, M. L., Manners, D., Testa, C., Malucelli, E., La Morgia, C., Barboni, P., Carbonelli, M., Schimpf, S., Wissinger, B., Zeviani, M., Baruzzi, A., Liguori, R., Barbiroli, B. & Carelli, V. (2010) Defective Mitochondrial Adenosine Triphosphate Production in Skeletal Muscle From Patients With Dominant Optic Atrophy Due to OPA1 Mutations. Arch Neurol. Loiseau, D., Chevrollier, A., Verny, C., Guillet, V., Gueguen, N., Pou de Crescenzo, M. A., Ferre, M., Malinge, M. C., Guichet, A., Nicolas, G., Amati-bonneau, P., Malthiery, Y., Bonneau, D. & Reynier, P. (2007) Mitochondrial coupling defect in Charcot-Marie-Tooth type 2A disease. Ann Neurol, 61, 315–23. Lunkes, A., Hartung, U., Magarino, C., Rodriguez, M., Palmero, A., Rodriguez, L., Heredero, L., Weissenbach, J., Weber, J. & Auburger, G. (1995) Refinement of the OPA1 gene locus on chromosome 3q28-q29 to a region of 2-8 cM, in one Cuban pedigree with autosomal dominant optic atrophy type Kjer. Am J Hum Genet, 57, 968–70. Lyle, W. (1990) Genetic Risk. University of Waterloo Press. Malena, A., Loro, E., Di Re, M., Holt, I. J. & Vergani, L. (2009) Inhibition of mitochondrial fission favours mutant over wild-type mitochondrial DNA. Hum Mol Genet, 18, 3407–16. Marchbank, N. J., Craig, J. E., Leek, J. P., Toohey, M., Churchill, A. J., Markham, A. F., Mackey, D. A., Toomes, C. & Inglehearn, C. F. (2002) Deletion of the OPA1 gene in a

194

G. Lenaers et al.

dominant optic atrophy family: evidence that haploinsufficiency is the cause of disease. J Med Genet, 39, e47. Matsuda, N. & Tanaka, K. (2010) Uncovering the roles of PINK1 and parkin in mitophagy. Autophagy, 6, 952–4. Merkwirth, C., Dargazanli, S., Tatsuta, T., Geimer, S., Lower, B., Wunderlich, F. T., Von Kleistretzow, J. C., Waisman, A., Westermann, B. & Langer, T. (2008) Prohibitins control cell proliferation and apoptosis by regulating OPA1-dependent cristae morphogenesis in mitochondria. Genes Dev, 22, 476–88. Milea, D., Sander, B., Wegener, M., Jensen, H., Kjer, B., Jorgensen, T. M., Lund-andersen, H. & Larsen, M. (2010) Axonal loss occurs early in dominant optic atrophy. Acta Ophthalmol, 88, 342–6. Milone, M., Younge, B. R., Wang, J., Zhang, S. & Wong, L. J. (2009) Mitochondrial disorder with OPA1 mutation lacking optic atrophy. Mitochondrion, 9, 279–81. Misko, A., Jiang, S., Wegorzewska, I., Milbrandt, J. & Baloh, R. H. (2010) Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex. J Neurosci, 30, 4232–40. Nicholson, G. A., Magdelaine, C., Zhu, D., Grew, S., Ryan, M. M., Sturtz, F., Vallat, J. M. & Ouvrier, R. A. (2008) Severe early-onset axonal neuropathy with homozygous and compound heterozygous MFN2 mutations. Neurology, 70, 1678–81. Nochez, Y., Arsene, S., Gueguen, N., Chevrollier, A., Ferre, M., Guillet, V., Desquiret, V., Toutain, A., Bonneau, D., Procaccio, V., Amati-bonneau, P., Pisella, P. J. & Reynier, P. (2009) Acute and late-onset optic atrophy due to a novel OPA1 mutation leading to a mitochondrial coupling defect. Mol Vis, 15, 598–608. Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P. & Lenaers, G. (2003) Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J Biol Chem, 278, 7743–6. Olichon, A., Elachouri, G., Baricault, L., Delettre, C., Belenguer, P. & Lenaers, G. (2007a) OPA1 alternate splicing uncouples an evolutionary conserved function in mitochondrial fusion from a vertebrate restricted function in apoptosis. Cell Death Differ, 14, 682–92. Olichon, A., Emorine, L. J., Descoins, E., Pelloquin, L., Brichese, L., Gas, N., Guillou, E., Delettre, C., Valette, A., Hamel, C. P., Ducommun, B., Lenaers, G. & Belenguer, P. (2002) The human dynamin-related protein OPA1 is anchored to the mitochondrial inner membrane facing the inter-membrane space. FEBS Lett, 523, 171–6. Olichon, A., Landes, T., Arnaune-pelloquin, L., Emorine, L. J., Mils, V., Guichet, A., Delettre, C., Hamel, C., Amati-bonneau, P., Bonneau, D., Reynier, P., Lenaers, G. & Belenguer, P. (2007b) Effects of OPA1 mutations on mitochondrial morphology and apoptosis: relevance to ADOA pathogenesis. J Cell Physiol, 211, 423–30. Payne, M., Yang, Z., Katz, B. J., Warner, J. E., Weight, C. J., Zhao, Y., Pearson, E. D., Treft, R. L., Hillman, T., Kennedy, R. J., Meire, F. M. & Zhang, K. (2004) Dominant optic atrophy, sensorineural hearing loss, ptosis, and ophthalmoplegia: a syndrome caused by a missense mutation in OPA1. Am J Ophthalmol, 138, 749–55. Pesch, U. E., Leo-kottler, B., Mayer, S., Jurklies, B., Kellner, U., Apfelstedt-sylla, E., Zrenner, E., Alexander, C. & Wissinger, B. (2001) OPA1 mutations in patients with autosomal dominant optic atrophy and evidence for semi-dominant inheritance. Hum Mol Genet, 10, 1359–68. Pich, S., Bach, D., Briones, P., Liesa, M., Camps, M., Testar, X., Palacin, M. & Zorzano, A. (2005) The Charcot-Marie-Tooth type 2A gene product, Mfn2, up-regulates fuel oxidation through expression of OXPHOS system. Hum Mol Genet, 14, 1405–15. Puomila, A., Huoponen, K., Mantyjarvi, M., Hamalainen, P., Paananen, R., Sankila, E. M., Savontaus, M. L., Somer, M. & Nikoskelainen, E. (2005) Dominant optic atrophy: correlation between clinical and molecular genetic studies. Acta Ophthalmol Scand, 83, 337–46. Reynier, P., Amati-bonneau, P., Verny, C., Olichon, A., Simard, G., Guichet, A., Bonnemains, C., Malecaze, F., Malinge, M. C., Pelletier, J. B., Calvas, P., Dollfus, H., Belenguer, P., Malthiery, Y., Lenaers, G. & Bonneau, D. (2004) OPA3 gene mutations responsible for autosomal dominant optic atrophy and cataract. J Med Genet, 41, e110.

6  Neurological Diseases Associated with Mutations in the Mitochondrial Fusion Machinery 195 Rojo, M., Legros, F., Chateau, D. & Lombes, A. (2002) Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo. J Cell Sci, 115, 1663–74. Schimpf, S., Fuhrmann, N., Schaich, S. & Wissinger, B. (2008) Comprehensive cDNA study and quantitative transcript analysis of mutant OPA1 transcripts containing premature termination codons. Hum Mutat, 29, 106–12. Shimizu, S., Mori, N., Kishi, M., Sugata, H., Tsuda, A. & Kubota, N. (2003) A novel mutation in the OPA1 gene in a Japanese patient with optic atrophy. Am J Ophthalmol, 135, 256–7. Skre, H. (1974) Genetic and clinical aspects of Charcot-Marie-Tooth’s disease. Clin Genet, 6, 98–118. Song, Z., Chen, H., Fiket, M., Alexander, C. & Chan, D. C. (2007) OPA1 processing controls mitochondrial fusion and is regulated by mRNA splicing, membrane potential, and Yme1L. J Cell Biol, 178, 749–55. Song, Z., Ghochani, M., Mccaffery, J. M., Frey, T. G. & Chan, D. C. (2009) Mitofusins and OPA1 mediate sequential steps in mitochondrial membrane fusion. Mol Biol Cell, 20, 3525–32. Spinazzi, M., Cazzola, S., Bortolozzi, M., Baracca, A., Loro, E., Casarin, A., Solaini, G., Sgarbi, G., Casalena, G., Cenacchi, G., Malena, A., Frezza, C., Carrara, F., Angelini, C., Scorrano, L., Salviati, L. & Vergani, L. (2008) A NOVEL DELETION IN THE GTPase DOMAIN OF OPA1 CAUSES DEFECTS IN MITOCHONDRIAL MORPHOLOGY AND DISTRIBUTION, BUT NOT IN FUNCTION. Hum Mol Genet. Stewart, J. D., Hudson, G., Yu-wai-man, P., Blakeley, E. L., He, L., Horvath, R., Maddison, P., Wright, A., Griffiths, P. G., Turnbull, D. M., Taylor, R. W. & Chinnery, P. F. (2008) OPA1 in multiple mitochondrial DNA deletion disorders. Neurology, 71, 1829–31. Thiselton, D. L., Alexander, C., Morris, A., Brooks, S., Rosenberg, T., Eiberg, H., Kjer, B., Kjer, P., Bhattacharya, S. S. & Votruba, M. (2001) A frameshift mutation in exon 28 of the OPA1 gene explains the high prevalence of dominant optic atrophy in the Danish population: evidence for a founder effect. Hum Genet, 109, 498–502. Thiselton, D. L., Alexander, C., Taanman, J. W., Brooks, S., Rosenberg, T., Eiberg, H., Andreasson, S., van Regemorter, N., Munier, F. L., Moore, A. T., Bhattacharya, S. S. & Votruba, M. (2002) A comprehensive survey of mutations in the OPA1 gene in patients with autosomal dominant optic atrophy. Invest Ophthalmol Vis Sci, 43, 1715–24. Toomes, C., Marchbank, N. J., Mackey, D. A., Craig, J. E., Newbury-ecob, R. A., Bennett, C. P., Vize, C. J., Desai, S. P., Black, G. C., Patel, N., Teimory, M., Markham, A. F., Inglehearn, C. F. & Churchill, A. J. (2001) Spectrum, frequency and penetrance of OPA1 mutations in dominant optic atrophy. Hum Mol Genet, 10, 1369–78. Tooth, H. (1886) The peroneal type of progressive muscular atrophy. HK Lewis, London. Twig, G., Elorza, A., Molina, A. J., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E. & Shirihai, O. S. (2008) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. Embo J, 27, 433–46. Verhoeven, K., Claeys, K. G., Zuchner, S., Schroder, J. M., Weis, J., Ceuterick, C., Jordanova, A., Nelis, E., de Vriendt, E., van Hul, M., Seeman, P., Mazanec, R., Saifi, G. M., Szigeti, K., Mancias, P., Butler, I. J., Kochanski, A., Ryniewicz, B., de Bleecker, J., Van Den Bergh, P., Verellen, C., Van Coster, R., Goemans, N., Auer-grumbach, M., Robberecht, W., Milic Rasic, V., Nevo, Y., Tournev, I., Guergueltcheva, V., Roelens, F., Vieregge, P., Vinci, P., Moreno, M. T., Christen, H. J., Shy, M. E., Lupski, J. R., Vance, J. M., De Jonghe, P. & Timmerman, V. (2006) MFN2 mutation distribution and genotype/phenotype correlation in Charcot-MarieTooth type 2. Brain, 129, 2093–102. Verny, C., Loiseau, D., Scherer, C., Lejeune, P., Chevrollier, A., Gueguen, N., Guillet, V., Dubas, F., Reynier, P., Amati-Bonneau, P. & Bonneau, D. (2008) Multiple sclerosis-like disorder in OPA1-related autosomal dominant optic atrophy. Neurology, 70, 1152–3. Votruba, M., Moore, A. T. & Bhattacharya, S. S. (1998) Clinical features, molecular genetics, and pathophysiology of dominant optic atrophy. J Med Genet, 35, 793–800. Wang, W., Cheng, X., Lu, J., Wei, J., Fu, G., Zhu, F., Jia, C., Zhou, L., Xie, H. & Zheng, S. (2010) Mitofusin-2 is a novel direct target of p53. Biochem Biophys Res Commun.

196

G. Lenaers et al.

White, K. E., Davies, V., Hogan, V., Piechota, M., Nichols, P., Turnbull, D. M. & Votruba, M. (2009) OPA1 deficiency is associated with increased autophagy in retinal ganglion cells in a murine model of dominant optic atrophy. Invest Ophthalmol Vis Sci. Williams, P. A., Morgan, J. E. & Votruba, M. (2010) Opa1 deficiency in a mouse model of dominant optic atrophy leads to retinal ganglion cell dendropathy. Brain. Yu-wai-man, P., Griffiths, P. G., Burke, A., Sellar, P. W., Clarke, M. P., Gnanaraj, L., Ah-kine, D., Hudson, G., Czermin, B., Taylor, R. W., Horvath, R. & Chinnery, P. F. (2010a) The prevalence and natural history of dominant optic atrophy due to OPA1 mutations. Ophthalmology, 117, 1538–46, 1546 e1. Yu-wai-man, P., Griffiths, P. G., Gorman, G. S., Lourenco, C. M., Wright, A. F., Auer-grumbach, M., Toscano, A., Musumeci, O., Valentino, M. L., Caporali, L., Lamperti, C., Tallaksen, C. M., Duffey, P., Miller, J., Whittaker, R. G., Baker, M. R., Jackson, M. J., Clarke, M. P., Dhillon, B., Czermin, B., Stewart, J. D., Hudson, G., Reynier, P., Bonneau, D., Marques, W., Jr., Lenaers, G., Mcfarland, R., Taylor, R. W., Turnbull, D. M., Votruba, M., Zeviani, M., Carelli, V., Bindoff, L. A., Horvath, R., Amati-bonneau, P. & Chinnery, P. F. (2010b) Multi-system neurological disease is common in patients with OPA1 mutations. Brain, 133, 771–86. Yu-wai-man, P., Sitarz, K. S., Samuels, D. C., Griffiths, P. G., Reeve, A. K., Bindoff, L. A., Horvath, R. & Chinnery, P. F. (2010c) OPA1 mutations cause cytochrome c oxidase deficiency due to loss of wild-type mtDNA molecules. Hum Mol Genet, 19, 3043–52. Zanna, C., Ghelli, A., Porcelli, A. M., Karbowski, M., Youle, R. J., Schimpf, S., Wissinger, B., Pinti, M., Cossarizza, A., Vidoni, S., Valentino, M. L., Rugolo, M. & Carelli, V. (2008) OPA1 mutations associated with dominant optic atrophy impair oxidative phosphorylation and mitochondrial fusion. Brain, 131, 352–67. Zhu, D., Kennerson, M. L., Walizada, G., Zuchner, S., Vance, J. M. & Nicholson, G. A. (2005) Charcot-Marie-Tooth with pyramidal signs is genetically heterogeneous: families with and without MFN2 mutations. Neurology, 65, 496–7. Zorzano, A. (2009) Regulation of mitofusin-2 expression in skeletal muscle. Appl Physiol Nutr Metab, 34, 433–9. Zuchner, S., de Jonghe, P., Jordanova, A., Claeys, K. G., Guergueltcheva, V., Cherninkova, S., Hamilton, S. R., Van Stavern, G., Krajewski, K. M., Stajich, J., Tournev, I., Verhoeven, K., Langerhorst, C. T., De Visser, M., Baas, F., Bird, T., Timmerman, V., Shy, M. & Vance, J. M. (2006) Axonal neuropathy with optic atrophy is caused by mutations in mitofusin 2. Ann Neurol, 59, 276–81. Zuchner, S., Mersiyanova, I. V., Muglia, M., Bissar-tadmouri, N., Rochelle, J., Dadali, E. L., Zappia, M., Nelis, E., Patitucci, A., Senderek, J., Parman, Y., Evgrafov, O., Jonghe, P. D., Takahashi, Y., Tsuji, S., Pericak-vance, M. A., Quattrone, A., Battaloglu, E., Polyakov, A. V., Timmerman, V., Schroder, J. M. & Vance, J. M. (2004) Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat Genet, 36, 449–51.

Chapter 7

Mitochondrial Fission-Fusion and Parkinson’s Disease: A Dynamic Question of Compensatory Networks Charleen T. Chu and Sarah B. Berman

Abstract  Mitochondrial pathobiology has long been linked to the pathogenesis of neurodegenerative diseases, in part because neurons are highly dependent upon mitochondrial metabolism. In particular, mitochondrial dysfunction represents a prominent feature of Parkinson’s disease (PD) and related familial forms of ­parkinsonism. Mitochondria are able to divide by fission, move around the cell and fuse with other mitochondria, and these dynamic changes are essential for regulating not only mitochondrial functions, but also its lifecycle within a cell. A rapidly growing body of recent literature adds a new twist to the saga, putting the focus on mitochondrial dynamics in the pathogenesis of PD. Here we review the literature on toxin (rotenone, 1-methyl-4-phenylpyridinium: MPP+, 6-hydroxydopamine) and genetic (parkin, PTEN-induced kinase 1: PINK1, DJ-1, a-synuclein) models of PD pathogenesis with respect to fission, fusion and mitochondrial autophagy (mitophagy). Strengths and weaknesses of various approaches to studying mitochondrial fission and fusion are highlighted, and unresolved controversies in the field are discussed in the context of a dynamic network of compensatory responses to mitochondrial stress, dysfunction and injury. Keywords  Mitochondrial complex I inhibitors • Mitochondrial dynamics • Parkin • Parkinson Disease • PINK1

C.T. Chu () Department of Pathology, University of Pittsburgh School of Medicine, 200 Lothrop St., Pittsburgh, PA 15213, USA e-mail: [email protected] S.B. Berman Department of Neurology, University of Pittsburgh School of Medicine, 3501 Fifth Avenue, BST3, Room 7037, Pittsburgh, PA 15213, USA and The Pittsburgh Institute for Neurodegenerative Diseases, University of Pittsburgh School of Medicine, Pittsburgh, PA 15213, USA B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_7, © Springer Science+Business Media B.V. 2011

197

198

C.T. Chu and S.B. Berman

Abbreviations 6-OHDA ERK1/2 FRAP MPP+ MPTP PD PINK1

6-hydroxydopamine extracellular signal-regulated protein kinase 1/2 fluorescence recovery after photobleaching 1-methyl-4-phenylpyridinium 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine Parkinson’s disease PTEN-induced kinase 1

7.1 Introduction Parkinson’s disease (PD), dementia with Lewy bodies and related familial parkinsonian syndromes are characterized by degeneration of the dopaminergic midbrain neurons of the nigrostriatal projection, with varying degrees of involvement by other brainstem, autonomic and cortical regions. Early efforts to model parkinsonian neurodegeneration involved administration of toxins such as 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) and 6-hydroxydopamine (6-OHDA), both of which cause relatively selective damage to dopaminergic neurons due to dopamine transporter-mediated uptake. The active metabolite of MPTP functions as a mitochondrial complex I inhibitor, and human tissue studies reveal reduced complex I activity in PD (Schapira et al. 1990). Interestingly, PD patients also exhibit peripheral decreases in complex I activity (Haas et al. 1995), and it was later discovered that systemic administration of rotenone causes selective nigrostriatal degeneration in rats (Betarbet et al. 2000). These observations implicate enhanced vulnerability of dopaminergic neurons to altered mitochondrial function. Mitochondrial function is regulated by a continuous process of remodeling involving the mitochondrial fission and fusion machinery. In general, a fused mitochondrial reticulum enhances respiration, mtDNA stability (Chen et  al. 2010a) and calcium buffering capacity (Frieden et  al. 2004). Mitochondrial fission is necessary for synaptogenesis and cell division (Li et al. 2008; Li et al. 2004), and in replicative cells, senescence is associated with fused mitochondria (Park et al. 2010). Many forms of cellular injury stimulate mitochondrial fission, which in some cases is obligatory for subsequent apoptotic cell death (Youle and Karbowski 2005). On the other hand, fission permits the selective degradation of damaged mitochondrial segments by macroautophagy (Twig et al. 2008), playing a neuroprotective role in a recessive PD model (Dagda et  al. 2009). These apparently disparate fission-dependent processes may be reconciled by conceptualizing fission as a temporary injury response (Chu 2010b), isolating dysfunctional portions of the mitochondrial network. The potential outcomes include local repair and ability to rejoin the mitochondrial reticulum through fusion, permanent sequestration within autophagosomes, and if both these measures fail, leakage of apoptotic mediators from the intermembrane space (Chu 2010b). Under homeostatic conditions,

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

199

mitochondria appear as a mixture of “threads” and “grains”, reflecting fused and fissioned segments; however, dynamic changes to mitochondrial morphology with stress or injury reflects a balance of not just fission and fusion, but also of mitochondrial autophagy, biogenesis and transport (Fig. 7.1). The recent discoveries that several PD-linked gene products exhibit regulated translocation to mitochondria have stimulated a renaissance of interest in mitochondrial dysfunction as a primary mechanism in parkinsonian neurodegeneration (Reviewed in (Winklhofer and Haass 2010)). In particular, the protein products of three mutated genes in familial recessive PD, parkin (PARK2), PTEN-induced kinase 1 (PINK1; PARK6) and DJ-1 (PARK7) have been implicated in key roles regulating mitochondrial quality control (Whitworth and Pallanck 2009; Chu 2010a; Hao et al. 2010). Loss-of-function in each of these proteins exhibit striking effects on mitochondrial morphology in mammalian cells and Drosophila, although their roles in regulating fission or fusion remain controversial. In subsequent sections, we review key methodologies used to study or infer mitochondrial dynamics with respect to their use in different PD model systems. Specific roles of PD-implicated proteins in the regulation of mitochondrial function, oxidative stress, fission-fusion or autophagic mitochondrial recycling remain to be fully elucidated, but undoubtedly these dynamic functional considerations will impact the steady state complement of fused, aggregated or fissioned mitochondria observed in PD and its model systems.

7.2 The Study of Mitochondrial Dynamics in Neurons and Non-Neuronal Model Systems Mitochondrial dynamics encompasses mitochondrial fission, fusion, transport, biogenesis, and degradation. Here, we will focus on methods employed to analyze mitochondrial fission and fusion in non-neuronal and neuronal cells, particularly with reference to Parkinson’s disease. To assess mitochondrial fission and fusion in whole cell systems, mitochondrial morphology is often utilized, analyzed by imaging fluorescently labeled mitochondria (Fig. 7.1, insets) or through electron microscopy. In some cases, cells are subjectively scored as containing fragmented or interconnected mitochondria, e.g. (Exner et al. 2007). However, this assumes that the process being studied will affect all mitochondria in a cell, and is less suited for cells such as neurons that natively express relatively discrete, highly motile mitochondria. In other cases, the distribution of mitochondrial sizes or lengths is used to derive a quantitative index of mitochondrial morphology reflective of the balance of fission to fusion, e.g. (Dagda et al. 2009; Ziviani et al. 2010). These methods provide rapid static measurements of mitochondrial morphology under various conditions. Hypothesized changes in mitochondrial fission or fusion are then confirmed using molecular genetic methods to alter the expression of known fission or fusion factors, or pharmacologic inhibition of fission, to rescue the abnormal phenotype, e.g. (Cui et al. 2010; Deng et al. 2008; Dagda et al. 2009).

200

C.T. Chu and S.B. Berman

Fig. 7.1  Dynamic changes in mitochondrial morphology and distribution reflect functional and pathologic demands. Mitochondria undergo multiple intersecting cycles of biogenesis and degradation, fission and fusion, and bidirectional transport in neurons. While fission and fusion are both necessary for proper distribution and function of mitochondria in somatic, dendritic, axonal and synaptic compartments, fission is a common response to multiple forms of cellular and mitochondrially targeted injuries. If localized repair is successful, fissioned mitochondria can rejoin the mitochondrial reticulum through fusion. Fragmentation and removal of damaged segments of mitochondria are coordinated during the process of mitophagy, which is believed to prevent release of pro-apoptotic mediators. Other injury responses commonly observed in neurons include perinuclear clustering, which may facilitate autophagic clearance. a-Synuclein mutants, neurotoxins and deficiency in recessive PD genes have been implicated in promoting mitochondrial dysfunction. While DJ-1, PINK1 and Parkin are each implicated in promoting a healthy mitochondrial complement, it is less clear the relative contributions of upstream injury prevention versus effective repair/removal of damaged mitochondria. Insets show examples of altered steady state appearances of mitochondria. (a) Mitochondrial interconnectivity and elongation are increased in SH-SY5Y cells stably overexpressing wild type PINK1 and stained with mitotracker green. (b) In contrast, COX8-GFP expressing SH-SY5Y cells stably defective in PINK1 exhibit fragmented mitochondria. (c) In primary midbrain neuron cultures treated with 5 mM MPP+, the unaffected nondopaminergic neuron (asterisk) exhibits mitochondria (green, AIF) dispersed throughout soma and neuritic extensions. In contrast, the fluorescent mitochondrial signal forms an enlarged perinuclear structure (yellow, arrow) in the injured dopaminergic neuron (red, tyrosine hydroxylase). While changes in morphology are often interpreted as consequences of altered fission or fusion, multiple other factors can affect the appearance of somatic mitochondria in neurons. These include changes in growth, clustering, transport and/or degradation that affect subsets of shorter or longer mitochondria. Panels (a) and (b) are unpublished images of SH-SY5Y clones #9 and D14, respectively, from a previously analyzed dataset (Dagda et al. 2009), and panel (c) represents an alternative image from a previously described experiment (Chu et al. 2005). Scale bars in each panel: 5 microns

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

201

However, potential interpretive caveats exist. For example, apparently larger or longer fluorescently-labeled mitochondria, usually interpreted as an increase in mitochondrial fusion, could potentially represent clusters of small mitochondria (Huang et  al. 2007). While this can be addressed through electron microscopy, quantifying mitochondrial length in electron micrographs is suboptimal unless 3-D tomography is employed, as the apparent mitochondrial segment length is influenced by the plane of section. In addition, in neurons, there is evidence that altered rates of mitochondrial fission and/or fusion can lead to minimal changes in overall mitochondrial morphology (Arnold et al. 2011), presumably due to compensatory changes in biogenesis or degradation. For example, an increase in fission could be masked by selective autophagic degradation of smaller (damaged) mitochondria (Chu 2010b). Live cell imaging techniques are also used to directly evaluate mitochondrial fission and fusion events in cells. Connectivity of mitochondria can be studied using fluorescence recovery after photobleaching (FRAP). An area of fluorescently labeled mitochondria is photobleached, and recovery of fluorescence depends on fusion and connectivity of neighboring mitochondria, with resulting redistribution of non-bleached fluorescent molecules to the pre-bleached region, e.g. (Abassi et al. 2004; Karbowski et al. 2006; Sandebring et al. 2009). This is particularly useful in non-neuronal cells with minimal mitochondrial movement across the cell, and assumes uniform rates of fission-fusion across the cell, as only a limited area is photobleached. Alternatively, double fluorescent studies employing mitochondrial red fluorescence with photoactivatable mitochondrial green fluorescence have been utilized to quantify mitochondrial fusion rates by mixing of fluorescent molecules, in both non-neuronal and neuronal cells (Berman et  al. 2009; Karbowski et  al. 2004). This has the advantage of directly measuring rates of fusion and fission, but is time-intensive making it difficult to perform large-scale analyses. One difficulty with analyzing mitochondrial dynamics is that it is becoming clear that mitochondrial fission/fusion, mitophagy, transport, and biogenesis are interrelated and may be coordinately regulated (Twig et al. 2008; Arnold et al. 2011; Dagda et al. 2009; Kim and Lemasters 2010). For example, while inhibiting fission reduces mitophagy, inhibiting autophagy reduces mitochondrial fragmentation, indicating a cooperative process involving both machineries (Dagda et al. 2009). Thus, development of methods to integrate multiple aspects of mitochondrial dynamics, particularly in neurons, where less is known about these interactions, are needed to better elucidate the role of altered mitochondrial dynamics in PD.

7.3 Mitochondrial Dynamics and PD Toxins Given that neuronal cell loss in the dopaminergic nigrostriatal system gives rise to many of the major PD symptoms, several toxins have been used to produce relatively selective lesions in this system. Two of the major PD toxins function as inhibitors of mitochondrial complex I activity, the pesticide rotenone and 1-methyl-4-phenylpyridinium,

202

C.T. Chu and S.B. Berman

the active metabolite of the heroin contaminant MPTP (Przedborski et  al. 2004; Panov et al. 2005). ROS production, while present to a degree, is not typically robust (Benard et  al. 2007), and cellular effects are frequently attributed to respiratory impairment (Plecita-Hlavata et al. 2008). In contrast, the toxin 6-hydroxydopamine operates directly through oxidative mechanisms (Henze et al. 2005). Each of these stresses is associated with fragmentation of the mitochondrial network, suggesting an alteration in the fission-fusion balance. In non-neuronal cells, either rotenone or a mitochondrial uncoupling ionophore, causes mitochondrial fission and disintegration of the mitochondrial network that cannot be prevented by the MitoQ antioxidant (Plecita-Hlavata et al. 2008). Fission is also induced by rotenone or nitric oxide in cortical neurons, and cell death is reduced by either mitofusin 1 or dominant negative Drp1 (Barsoum et al. 2006), consistent with a pro-apoptotic role for fission in these models. In primary neurons exposed to chronic, sublethal rotenone, early increases in mitochondrial fusion shifts to favor fission over time, and inhibition of mitochondrial fission prevented neurite loss in differentiated dopaminergic cells (Arnold et al. 2011). This suggests that changes in mitochondrial dynamics may play a role early in the pathology of this PD-related model. MPP + elicites caspase-independent cell death in SH-SY5Y cells and primary dopaminergic midbrain neurons (Chu et al. 2005). While mitochondrial autophagy is increased in both transformed and primary neurons (Zhu et al. 2007), mitochondria appear fragmented in SH-SY5Y cells, while perinuclear clustering is observed in a subset of dopaminergic midbrain neurons (Fig. 7.1c). The catecholamine neurotoxin 6-OHDA is capable of generating reactive oxygen species through redox cycling. Although redox cycling of 6-OHDA can occur rapidly in both extracellular or pH neutral intracellular compartments (Kulich et al. 2007), its toxicity in  vivo is not reduced by extracellular superoxide dismutase (Callio et al. 2005). Instead, neuronal cell death is dependent upon mitochondriallyderived reactive oxygen species (Callio et al. 2005), a delayed cellular response to injury (Kulich et al. 2007). This is accompanied by Drp1-dependent mitochondrial fission (Gomez-Lazaro et  al. 2008) and mitochondrial autophagy (Dagda et  al. 2008). As observed in other toxin models, silencing of Drp1 conferred significant protection from cell death (Gomez-Lazaro et al. 2008).

7.4 Mitochondrial Dynamics and DJ-1 DJ-1 is mutated in recessive familial parkinsonism, and has a peroxiredoxin-like activity (Andres-Mateos et al. 2007) as well as a protease-like activity (Chen et al. 2010b). A cysteine at position 106 acts as a redox switch regulating its translocation to mitochondria (Canet-Aviles et  al. 2004), but also rendering it susceptible to oxidative inactivation with aging in Drosophila (Meulener et al. 2006). In fibroblasts derived from DJ-1 knockout mice or from patients with homozygous mutation in the DJ-1 gene, there is decreased mitochondrial membrane potential and

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

203

increased oxidative stress, accompanied by mitochondrial fragmentation and simplification of branch points (Krebiehl et al. 2010). Similarly, shortened mitochondrial lengths were observed in DJ-1 knockout cortical neurons, and DJ-1 fibroblasts or knockdown neuroblastoma cells show reduced rates of mitochondrial fusion, assessed via spreading of a photoactivatable matrix signal (Irrcher et al. 2010; Thomas et al. 2010). Strikingly, dominant negative Drp1 reduced the susceptibility of DJ-1 knockout neurons to MPP + (Irrcher et  al. 2010), suggesting that fission plays an active role in cell death in this context. Mitochondrial dysfunction in DJ-1 null cells is sometimes accompanied by a baseline decrease in the activated (lipidated) form of the autophagosome marker LC3. However, whether this reflects increased or decreased autophagy remains controversial. DJ-1 null fibroblasts show reductions in expression of rapamycin-induced autophagosome markers in one study, consistent with decreased autophagic flux (Krebiehl et  al. 2010). Based on analysis of LC3-II levels in the presence and absence of the lysosomal fusion inhibitor bafilomycin, and observation of decreased steady state levels of p62, an autophagy substrate and cargo adaptor, another study concluded increased autophagic flux (Irrcher et  al. 2010). In other studies, DJ-1 shRNA increases markers of autophagosomes (Thomas et al. 2010). Flux analysis of autophagy or mitophagy can be technically challenging, requiring titration to detect rate differences that may be masked by saturation. Thus, it is still unclear how autophagy is perturbed in these cells, and whether these changes reflect a direct consequence of DJ-1 loss or a more likely a compensatory response. Activity of extracellular signal regulated protein kinase ½ (ERK1/2) is necessary for autophagy/mitophagy induced by either 6-OHDA (Dagda et  al. 2008) or the complex I inhibitor MPP+ (Zhu et al. 2007). Interestingly, reduced mitophagy as reported in one study of DJ-1 knockout fibroblasts is associated with a selective decrease in phosphorylated ERK1/2 specifically in the mitochondrial compartment (Krebiehl et al. 2010). Localized activation of ERK2 at the mitochondria is sufficient to induce mitophagy (Dagda et  al. 2008), and phosphorylated ERK2 is observed in association with mitochondria in post-mortem dopaminergic neurons of patients with PD and other Lewy body diseases (Zhu et al. 2003). These data suggest the possibility of a shared mechanism of mitophagy regulation under genetic, toxic and sporadic contexts.

7.5 Mitochondrial Dynamics and PINK1 The PTEN-induced kinase 1 (PINK1) is a mitochondrially-targeted kinase whose deficiency gives rise to perturbations in mitochondrial respiration, calcium buffering, electron leak, morphology and turnover. The effects upon mitochondrial dynamics have been extensively reviewed elsewhere (Van Laar and Berman 2009; Whitworth and Pallanck 2009; Yang and Lu 2009; Thomas and Cookson 2009; Chu 2010b), and are characterized by significant differences depending upon the model system. Regardless of these differences, and in contrast to the toxin and

204

C.T. Chu and S.B. Berman

DJ-1 models discussed above, suppressing fission exacerbates cellular pathology, implicating fission in a compensatory role. Mitochondrial morphology is dramatically altered in Drosophila and mammalian culture models of PINK1 deficiency, while mouse models show much more subtle morphologic effects (Gispert et al. 2009; Gautier et al. 2008). PINK1 deficiency in Drosophila is associated with condensed or clustered mitochondrial morphology, which is rescued by enhanced Drp1 or decreased mitofusin activity (Deng et  al. 2008; Poole et al. 2008; Yang et al. 2008). In contrast, either acute or chronic PINK1 knockdown in cultured mammalian neuronal cells results in Drp1-dependent fragmentation of the mitochondrial network (Fig. 7.1b) (Dagda et al. 2009; Lutz et al. 2009; Sandebring et al. 2009). More recent work further indicates that PINK1 deficiency has opposite effects on mitofusin expression between these model systems (Ziviani et al. 2010; Cui et al. 2010), perturbing the balance of fission to fusion proteins in directions consistent with the morphologic changes observed in each model. Interestingly, both Drosophila and SH-SY5Y studies implicate mitochondrial fission as a protective response, despite differences in the appearance of mitochondria. Drosophila studies suggest that PINK1 promotes fission, and loss of this protective effect is observed in PINK1 null animals (Poole et al. 2010). SH-SY5Y studies implicate PINK1 overexpression as promoting fusion (Fig.  7.1a), with PINK1 knockdown cells exhibiting fission as a compensatory response, downstream of ROS-generation and calcium dysregulation (Dagda et al. 2009; Sandebring et al. 2009). Cultured cortical neurons from PINK1 −/− mice show an increase in aggregated, but not in elongated, mitochondria within neurites, but only upon added proteasomal stress (Gispert et  al. 2009). These findings support the concept that some of the morphologic changes induced by PINK1 loss of function result indirectly from mitochondrial injury, with fission as a temporary injury response that can resolve in different directions depending upon context (Fig. 7.1). Determining whether or not effects of PINK1 on fission-fusion represent direct or indirect mechanisms in the different model systems would be important. The fate of fissioned mitochondria may differ between Drosophila and vertebrate systems, given differences in regulation of macroautophagy between these species. Both generalized bulk macroautophagy and selective mitophagy are dependent upon covalent attachment of the autophagy protein LC3 to the growing autophagosome membrane (Geng and Klionsky 2008; Gottlieb and Carreira 2010). While autophagy generally plays an adaptive, homeostatic role, we have proposed that high or sustained levels of autophagy or mitophagy may interfere with neuronal function, mediating neurite retraction or even cell death due to metabolic dependence upon mitochondria (Cherra et al. 2010a; Zhu et al. 2007). Indeed, several braking mechanisms to limit the extent of autophagy have been described. These include the ability of Bcl2 to limit the amount of Beclin 1 available for autophagy induction (Pattingre et al. 2005) and activation of a mammalian phosphatidylinositol 3-phosphate phosphatase (Vergne et al. 2009). LC3 itself is directly phosphorylated by protein kinase A, with downregulating effects on autophagy induced by rapamycin, MPP + or the G2019S mutation in leucine rich repeat kinase 2 (Cherra et  al. 2010b). Interestingly, this phosphorylation site is conserved in vertebrate species, but is absent in invertebrates.

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

205

Thus, differences in steady state appearance of mitochondria within Drosophila and mammalian cells deficient in PINK1 may reflect differences in the efficiency of clearance of fissioned mitochondria.

7.6 Mitochondrial Dynamics and Parkin Another PD-related gene, parkin, whose protein product is an E3 ubiquitin ligase and whose recessive mutations cause familial PD (Kitada et  al. 1998), has been found to interact downstream from PINK1 in the pathways involving mitochondrial dynamics, including fission and fusion and, more recently, mitophagy. In Drosophila, it was found that the mitochondrial morphologic changes induced by the loss of PINK1, as described above, are completely rescued by the overexpression of Parkin, while the reverse is not true (i.e. PINK1 overexpression does not rescue the mitochondrial morphology abnormalities caused by loss of Parkin) (Clark et  al. 2006; Park et al. 2006; Yang et al. 2006). Although as noted above, the mitochondrial morphology changes due to loss of PINK1 in mammalian cells differs from that seen in Drosophila, Parkin overexpression was similarly able to rescue the morphologic changes in human cells (Exner et al. 2007). These results suggest that Parkin functions downstream of PINK1 to regulate mitochondrial morphology. Like PINK1, then, Parkin is also suggested to promote fission in flies (Yang et al. 2008; Poole et al. 2008; Park et al. 2009; Deng et al. 2008), whereas it has been suggested to promote fusion in mammalian cells (Lutz et al. 2009). More recently, mitochondrial homeostasis through mitophagy has been identified as an important function of these PD-related proteins. Narendra et al. first found that Parkin translocates to depolarized mitochondria and promotes their mitophagic degradation (Narendra et al. 2008). This function of parkin in response to FCCP/CCCP has been found to be dependent on the presence of PINK1 (Vives-Bauza et al. 2010; Narendra et al. 2010a; Matsuda et al. 2010), which becomes stabilized on mitochondria when membrane potential is reduced (Lin and Kang 2008). Parkin-induced mitophagy involves PINK1 kinase activity (Matsuda et  al. 2010; Narendra et  al. 2010a) and Parkin’s ubiquitin ligase function (Geisler et al. 2010, 2373; Okatsu et al. 2010), though it remains controversial if there is a direct interaction (Xiong et al. 2009; Um et al. 2009; Vives-Bauza et al. 2010; Shiba et al. 2009). In addition, this pathway is dependent on the voltage-dependent anion channel VDAC1, mitophagyassociated protein Nix (Ding et  al. 2010), and perhaps p62 (Geisler et  al. 2010; Okatsu et  al. 2010), though this is not consistently found (Okatsu et  al. 2010; Narendra et al. 2010b). It has been hypothesized that the loss of Parkin’s ability to regulate the degradation of dysfunctional mitochondria could be an underlying etiology of Parkinrelated PD neurodegeneration. This, in turn, suggests that the vulnerable neurons in PD may be more dependent on this mitochondrial homeostatic function. However, the impact of Parkin-accelerated mitophagy on cell survival and death has not been extensively characterized, aside from apparent acceleration of cell death in HEK293

206

C.T. Chu and S.B. Berman

cells (Morrison et al. 2010) and HeLa cells cultured in galactose following 72 h of CCCP (Narendra et  al. 2008). Morever, Parkin-associated mitophagy has largely been studied in non-neuronal cells, which show different metabolic features from neurons (Gusdon and Chu 2011). Recent evidence indicates that bioenergetic dependence on mitochondria inhibits this phenomenon in primary neurons (Van Laar et  al. 2011). Thus, further understanding of mechanisms pertinent to neuronal mitochondrial quality control will help to delineate the role of Parkin in neuronal mitochondrial degradation. In addition, it is likely that the fission/fusion and mitophagy regulatory roles for Parkin are not disparate, unrelated functions, as interactions between mitochondrial degradation, mitochondrial fission and fusion, and mitochondrial transport are being elucidated in this system. Recently, Parkin was reported to ubiquitinate the outer mitochondrial membrane fusion protein mitofusin (Ziviani et  al. 2010; Poole et al. 2010), which provides a potential link between mitochondrial fission/ fusion activity and degradation of dysfunctional mitochondria. This is particularly interesting, given the proposed function of mitochondrial fission and fusion in targeting ­damaged mitochondria for degradation (Twig et  al. 2008), although there is also evidence for reciprocal regulation between the fission-fusion and autophagic machineries (Dagda et  al. 2009), which together provide possible mechanistic interactions between the impacts on mitochondrial fission/fusion and mitophagy. In addition, microtubule-dependent mitochondrial transport to perinuclear regions may be important in the PINK1/Parkin mitophagy pathway (Vives-Bauza et al. 2010; Okatsu et al. 2010), though this is not yet clearly defined. Thus, it is likely that many aspects of mitochondrial dynamics interact to regulate mitochondrial health and homeostasis, and these may be particularly relevant to PD neurodegeneration.

7.7 Mitochondrial Dynamics and a-Synuclein Although perhaps less clearly delineated than for the other PD-related proteins, a-synuclein has also been implicated with relation to mitochondrial dynamics. a-Synuclein (PARK1) gene mutations and duplications/triplications cause autosomal dominant PD (Polymeropoulos et al. 1997; Kruger et al. 1998; Zarranz et al. 2004; Singleton et al. 2003; Bradbury 2003), and a-synuclein is one of the major components of Lewy bodies in PD (Spillantini et al. 1997). Many mechanisms have been proposed for a-synuclein-related neurodegeneration, including proteasomal dysfunction and oxidative stress (reviewed in (Dawson et al. 2010)). Interestingly, alterations in a-synuclein can elicit mitochondrial membrane depolarization and respiratory dysfunction, including inhibition of Complex I activity (Banerjee et al. 2010; Loeb et al. 2010). A proportion of a-synuclein has been localized on or in mitochondria (Martin et al. 2006; Li et al. 2007; Zhang et al. 2008; Shavali et al. 2008; Devi et  al. 2008; Nakamura et  al. 2008), and a-synuclein overexpression sensitizes cells to mitochondrial toxins (Song et al. 2004; Orth et al. 2003).

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

207

a-Synuclein binds lipid membranes and was shown to affect the lipid packing of vesicular membranes (Kamp and Beyer 2006). This lipid binding activity may also contribute to formation of Lewy bodies, as peroxidized cardiolipin-a-synuclein-cytochrome c complexes sequester released cytochrome c to prevent apoptosis (Bayir et  al. 2009). Recently, Kamp et  al. (2010) reported that a-synuclein inhibits mitochondrial fusion in  vitro and causes mitochondrial fragmentation in neurons and muscle of C. elegans (Kamp et al. 2010). Most interestingly, this mitochondrial fragmentation was rescued by overexpression of PINK1, parkin, and DJ-1, but not their disease-causing mutant forms. This suggests the possibility that all of these proteins function in pathways that directly or indirectly modulate mitochondrial dynamics.

7.8 Future Perspectives: A Dynamic Question of Compensatory Networks? In PD, data from both genetic and toxin studies are converging on changes in both mitochondrial function and mitochondrial quality control. It is becoming clear that the consequences of changes in mitochondrial dynamics are more than the simple balance of fission and fusion activities, with excess fission resulting in cell death. Mitochondrial fission, while possibly involved in apoptosis, is also clearly necessary for synaptic function and mitochondrial distribution. In addition, as noted, fission/fusion changes are likely a component of mitophagy regulation (Twig et al. 2008; Dagda et al. 2009; Ziviani et al. 2010) and biogenesis (Berman et al. 2009). Maintenance of mitochondrial function and homeostasis, thus, most likely reflects a dynamic interaction tying fission/fusion to clearance of dysfunctional mitochondria, transport of mitochondria within neurons, and biogenesis of new mitochondria (Fig. 7.1). It is also possible that the mitochondrial homeostatic regulation may differ depending on the cellular environment and may change over time. For example, in cells less dependent on mitochondria, the PINK1/Parkin pathway might regulate mitochondrial quality control by promoting increased fission and mitochondrial clearance by autophagy, while in neuronal cells dependent on mitochondria, the pathway could promote mitochondrial redistribution and repair through fusion. The increasingly compelling evidence that dysfunction in mitochondrial quality control may be a common final pathway in PD-related neurodegeneration suggests a dynamic network of compensatory measures in chronic toxicity that over time can protect mitochondrial integrity through disparate processes. These may include fusion and re-mixing of mitochondrial contents, isolation and selective degradation of dysfunctional mitochondria, and/or perhaps increases in mitochondrial biogenesis. When dysregulation of this dynamic network occurs, however, it could then lead to dysfunctional mitochondrial homeostasis and subsequent neurodegeneration, which could occur through both loss-of-function(s) or imbalanced activation of a particular response. Neurons are extremely dependent on mitochondria for

208

C.T. Chu and S.B. Berman

energy production, calcium regulation, and synaptic function. Interestingly, the neurons vulnerable to PD-related degeneration, share some common features, including long axons that are poorly myelinated (Braak et al. 2003). These features are likely to make PD-vulnerable neurons especially sensitive to changes in the regulation of mitochondrial dynamics. Thus, dysregulation of mitochondrial dynamics could represent a common pathway for neurodegeneration in both sporadic and genetic forms of PD. Acknowledgments  CTC is supported in part by the National Institutes of Health (AG026389, NS065789), and is recipient of an AFAR/Ellison Medical Foundation Julie Martin Mid-Career Award in Aging Research. SB is supported in part by the National Institutes of Health K08NS059576.

References Abassi, Z. A., Binah, O. & Youdim, M. B. 2004. Cardiovascular activity of rasagiline, a selective and potent inhibitor of mitochondrial monoamine oxidase B: comparison with selegiline. Br J Pharmacol, 143, 371–8. Andres-mateos, E., Perier, C., Zhang, L., Blanchard-fillion, B., Greco, T. M., Thomas, B., Ko, H. S., Sasaki, M., Ischiropoulos, H., Przedborski, S., Dawson, T. M. & Dawson, V. L. 2007. DJ-1 gene deletion reveals that DJ-1 is an atypical peroxiredoxin-like peroxidase. Proc Natl Acad Sci USA, 104, 14807–12. Arnold, B., Cassady, S. J., Van Laar, V. S. & Berman, S. B. 2011. Integrating multiple aspects of mitochondrial dynamics in neurons: age-related differences and dynamic changes in a chronic rotenone model. Neurobiol Dis, 41, 189–200. Banerjee, K., Sinha, M., Pham cle, L., Jana, S., Chanda, D., Cappai, R. & Chakrabarti, S. 2010. Alpha-synuclein induced membrane depolarization and loss of phosphorylation capacity of isolated rat brain mitochondria: implications in Parkinson’s disease. FEBS Lett, 584, 1571–6. Barsoum, M. J., Yuan, H., Gerencser, A. A., Liot, G., Kushnareva, Y., Graber, S., Kovacs, I., Lee, W. D., Waggoner, J., Cui, J., White, A. D., Bossy, B., Martinou, J. C., Youle, R. J., Lipton, S. A., Ellisman, M. H., Perkins, G. A. & Bossy-wetzel, E. 2006. Nitric oxide-induced mitochondrial fission is regulated by dynamin-related GTPases in neurons. Embo J, 25, 3900–11. Bayir, H., Kapralov, A. A., Jiang, J., Huang, Z., Tyurina, Y. Y., Tyurin, V. A., Zhao, Q., Belikova, N. A., Vlasova, II, Maeda, A., Zhu, J., Na, H. M., Mastroberardino, P. G., Sparvero, L. J., Amoscato, A. A., Chu, C. T., Greenamyre, J. T. & Kagan, V. E. 2009. Peroxidase mechanism of lipid-dependent cross-linking of synuclein with cytochrome C: protection against apoptosis versus delayed oxidative stress in Parkinson disease. J Biol Chem, 284, 15951–69. Benard, G., Bellance, N., James, D., Parrone, P., Fernandez, H., Letellier, T. & Rossignol, R. 2007. Mitochondrial bioenergetics and structural network organization. J Cell Sci, 120, 838–48. Berman, S. B., Chen, Y., Mccaffery, J. M., Rucker, E. B., Goebbels, S., Nave, K.-A., Arnold, B. A., Jonas, E. A., Pineda, F. J. & Hardwick, J. M. 2009. Bcl-xL increases mitochondrial fission, fusion, and biomass in neurons. Journal of Cell Biology, 184, 707–719. Betarbet, R., Sherer, T. B., Mackenzie, G., Garcia-osuna, M., Panov, A. V. & Greenamyre, J. T. 2000. Chronic systemic pesticide exposure reproduces features of Parkinson’s disease. Nat Neurosci, 3, 1301–6. Braak, H., Del Tredici, K., Rub, U., De Vos, R. A., Jansen Ste Ur, E. N. & Braak, E. 2003. Staging of brain pathology related to sporadic Parkinson’s disease. Neurobiol Aging, 24, 197–211 Bradbury, J. 2003. Alpha-synuclein gene triplication discovered in Parkinson’s disease. Lancet Neurol, 2, 715.

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

209

Callio, J., Oury, T. D. & Chu, C. T. 2005. Manganese superoxide dismutase protects against 6-hydroxydopamine injury in mouse brains. J. Biol. Chem., 280, 18536–18542. Canet-aviles, R. M., Wilson, M. A., Miller, D. W., Ahmad, R., Mclendon, C., Bandyopadhyay, S., Baptista, M. J., Ringe, D., Petsko, G. A. & Cookson, M. R. 2004. The Parkinson’s disease protein DJ-1 is neuroprotective due to cysteine-sulfinic acid-driven mitochondrial localization. Proc Natl Acad Sci USA, 101, 9103–8. Chen, H., Vermulst, M., Wang, Y. E., Chomyn, A., Prolla, T. A., Mccaffery, J. M. & Chan, D. C. 2010a. Mitochondrial fusion is required for mtDNA stability in skeletal muscle and tolerance of mtDNA mutations. Cell, 141, 280–9. Chen, J., Li, L. & Chin, L. S. 2010b. Parkinson disease protein DJ-1 converts from a zymogen to a protease by carboxyl-terminal cleavage. Hum Mol Genet, 19, 2395–408. Cherra, S. J., 3 rd, Dagda, R. & Chu, C. T. 2010a. Autophagy and Neurodegeneration: Survival at a cost? Neuropathol Applied Neurol, 36, 125–132. Cherra, S. J., 3 rd, Kulich, S. M., Uechi, G., Balasubramani, M., Mountzouris, J., Day, B. W. & Chu, C. T. 2010b. Regulation of the autophagy protein LC3 by phosphorylation. J Cell Biol, 190, 533–9. Chu, C. T. 2010a. A pivotal role for PINK1 and autophagy in mitochondrial quality control: implications for Parkinson disease. Hum Mol Genet, 19, R28-37. Chu, C. T. 2010b. Tickled PINK1: Mitochondrial homeostasis and autophagy in recessive Parkinsonism. Biochim Biophys Acta, 1802, 20–8. Chu, C. T., Zhu, J. H., Cao, G., Signore, A., Wang, S. & Chen, J. 2005. Apoptosis inducing factor mediates caspase-independent 1-methyl-4-phenylpyridinium toxicity in dopaminergic cells. J Neurochem, 94, 1685–95. Clark, I. E., Dodson, M. W., Jiang, C., Cao, J. H., Huh, J. R., Seol, J. H., Yoo, S. J., Hay, B. A. & Guo, M. 2006. Drosophila pink1 is required for mitochondrial function and interacts genetically with parkin. Nature, 441, 1162–6. Cui, M., Tang, X., Christian, W. V., Yoon, Y. & Tieu, K. 2010. Perturbations in mitochondrial dynamics induced by human mutant PINK1 can be rescued by the mitochondrial division inhibitor mdivi-1. J Biol Chem, 285, 11740–52. Dagda, R. K., Cherra, S. J., 3 rd, Kulich, S. M., Tandon, A., Park, D. & Chu, C. T. 2009. Loss of pink1 function promotes mitophagy through effects on oxidative stress and mitochondrial fission. J Biol Chem, 284, 13843–55. Dagda, R. K., Zhu, J., Kulich, S. M. & Chu, C. T. 2008. Mitochondrially localized ERK2 regulates mitophagy and autophagic cell stress: implications for Parkinson’s disease. Autophagy, 4, 770–82. Dawson, T. M., Ko, H. S. & Dawson, V. L. 2010. Genetic animal models of Parkinson’s disease. Neuron, 66, 646–61. Deng, H., Dodson, M. W., Huang, H. & Guo, M. 2008. The Parkinson’s disease genes pink1 and parkin promote mitochondrial fission and/or inhibit fusion in Drosophila. Proc Natl Acad Sci USA, 105, 14503–8. Devi, L., Raghavendran, V., Prabhu, B. M., Avadhani, N. G. & Anandatheerthavarada, H. K. 2008. Mitochondrial import and accumulation of alpha-synuclein impair complex I in human dopaminergic neuronal cultures and Parkinson disease brain. J Biol Chem, 283, 9089–100. Ding, W. X., Ni, H. M., Li, M., Liao, Y., Chen, X., Stolz, D. B., Dorn, G. W., 2nd & Yin, X. M. 2010. Nix is critical to two distinct phases of mitophagy, reactive oxygen species-mediated autophagy induction and Parkin-ubiquitin-p62-mediated mitochondrial priming. J Biol Chem, 285, 27879–90. Exner, N., Treske, B., Paquet, D., Holmstrom, K., Schiesling, C., Gispert, S., Carballo-carbajal, I., Berg, D., Hoepken, H. H., Gasser, T., Kruger, R., Winklhofer, K. F., Vogel, F., Reichert, A. S., Auburger, G., Kahle, P. J., Schmid, B. & Haass, C. 2007. Loss-of-function of human PINK1 results in mitochondrial pathology and can be rescued by parkin. J Neurosci, 27, 12413–8. Frieden, M., James, D., Castelbou, C., Danckaert, A., Martinou, J. C. & Demaurex, N. 2004. Ca(2+) homeostasis during mitochondrial fragmentation and perinuclear clustering induced by hFis1. J Biol Chem, 279, 22704–14.

210

C.T. Chu and S.B. Berman

Gautier, C. A., Kitada, T. & Shen, J. 2008. Loss of PINK1 causes mitochondrial functional defects and increased sensitivity to oxidative stress. Proc Natl Acad Sci USA, 105, 11364–9. Geisler, S., Holmstrom, K. M., Skujat, D., Fiesel, F. C., Rothfuss, O. C., Kahle, P. J. & Springer, W. 2010. PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nat Cell Biol, 12, 119–31. Geng, J. & Klionsky, D. J. 2008. The Atg8 and Atg12 ubiquitin-like conjugation systems in macroautophagy. ‘Protein modifications: beyond the usual suspects’ review series. EMBO Rep, 9, 859–64. Gispert, S., Ricciardi, F., Kurz, A., Azizov, M., Hoepken, H. H., Becker, D., Voos, W., Leuner, K., Muller, W. E., Kudin, A. P., Kunz, W. S., Zimmermann, A., Roeper, J., Wenzel, D., Jendrach, M., Garcia-arencibia, M., Fernandez-ruiz, J., Huber, L., Rohrer, H., Barrera, M., Reichert, A. S., Rub, U., Chen, A., Nussbaum, R. L. & Auburger, G. 2009. Parkinson phenotype in aged PINK1-deficient mice is accompanied by progressive mitochondrial dysfunction in absence of neurodegeneration. PLoS One, 4, e5777. Gomez-lazaro, M., Bonekamp, N. A., Galindo, M. F., Jordan, J. & Schrader, M. 2008. 6-Hydroxydopamine (6-OHDA) induces Drp1-dependent mitochondrial fragmentation in SH-SY5Y cells. Free Radic Biol Med, 44, 1960–9. Gottlieb, R. A. & Carreira, R. S. 2010. Autophagy in health and disease. 5. Mitophagy as a way of life. Am J Physiol Cell Physiol, 299, C203-10. Gusdon, A. M. & Chu, C. T. 2011. To Eat or Not to Eat: Neuronal Metabolism, Mitophagy and Parkinson’s Disease. Antioxidants & Redox Signaling, In press. Haas, R. H., Nasirian, F., Nakano, K., Ward, D., Pay, M., Hill, R. & Shults, C. W. 1995. Low platelet mitochondrial complex I and complex II/III activity in early untreated Parkinson’s disease. Ann Neurol, 37, 714–22. Hao, L. Y., Giasson, B. I. & Bonini, N. M. 2010. DJ-1 is critical for mitochondrial function and rescues PINK1 loss of function. Proc Natl Acad Sci USA, 107, 9747–52. Henze, C., Earl, C., Sautter, J., Schmidt, N., Themann, C., Hartmann, A. & Oertel, W. H. 2005. Reactive oxidative and nitrogen species in the nigrostriatal system following striatal 6-hydroxydopamine lesion in rats. Brain Res, 1052, 97–104. Huang, P., Yu, T. & Yoon, Y. 2007. Mitochondrial clustering induced by overexpression of the mitochondrial fusion protein Mfn2 causes mitochondrial dysfunction and cell death. Eur J Cell Biol, 86, 289–302. Irrcher, I., Aleyasin, H., Seifert, E. L., Hewitt, S. J., Chhabra, S., Phillips, M., Lutz, A. K., Rousseaux, M. W., Bevilacqua, L., Jahani-Asl, A., Callaghan, S., Maclaurin, J. G., Winklhofer, K. F., Rizzu, P., Rippstein, P., Kim, R. H., Chen, C. X., Fon, E. A., Slack, R. S., Harper, M. E., Mcbride, H. M., Mak, T. W. & Park, D. S. 2010. Loss of the Parkinson’s disease-linked gene DJ-1 perturbs mitochondrial dynamics. Hum Mol Genet, 19, 3734–3746. Kamp, F. & Beyer, K. 2006. Binding of alpha-synuclein affects the lipid packing in bilayers of small vesicles. J Biol Chem, 281, 9251–9. Kamp, F., Exner, N., Lutz, A. K., Wender, N., Hegermann, J., Brunner, B., Nuscher, B., Bartels, T., Giese, A., Beyer, K., Eimer, S., Winklhofer, K. F. & Haass, C. 2010. Inhibition of mitochondrial fusion by alpha-synuclein is rescued by PINK1, Parkin and DJ-1. EMBO J. Karbowski, M., Arnoult, D., Chen, H., Chan, D. C., Smith, C. L. & Youle, R. J. 2004. Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J Cell Biol, 164, 493–9. Karbowski, M., Norris, K. L., Cleland, M. M., Jeong, S. Y. & Youle, R. J. 2006. Role of Bax and Bak in mitochondrial morphogenesis. Nature, 443, 658–62. Kim, I. & Lemasters, J. J. 2010. Mitochondrial Degradation by Autophagy (Mitophagy) in GFP-LC3 Transgenic Hepatocytes during Nutrient Deprivation. Am J Physiol Cell Physiol. Kitada, T., Asakawa, S., Hattori, N., Matsumine, H., Yamamura, Y., Minoshima, S., Yokochi, M., Mizuno, Y. & Shimizu, N. 1998. Mutations in the parkin gene cause autosomal recessive juvenile parkinsonism. Nature, 392, 605–8. Krebiehl, G., Ruckerbauer, S., Burbulla, L. F., Kieper, N., Maurer, B., Waak, J., Wolburg, H., Gizatullina, Z., Gellerich, F. N., Woitalla, D., Riess, O., Kahle, P. J., Proikas-cezanne, T. & Kruger, R. 2010. Reduced basal autophagy and impaired mitochondrial dynamics due to loss of Parkinson’s disease-associated protein DJ-1. PLoS One, 5, e9367.

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

211

Kruger, R., Kuhn, W., Muller, T., Woitalla, D., Graeber, M., Kosel, S., Przuntek, H., Epplen, J. T., Schols, L. & Riess, O. 1998. Ala30Pro mutation in the gene encoding alpha-synuclein in Parkinson’s disease. Nat Genet, 18, 106–8. Kulich, S., Horbinski, C., Patel, M. & Chu, C. 2007. 6-Hydroxydopamine induces mitochondrial ERK activation. Free Rad Biol Med, 43, 372–383. Li, H., Chen, Y., Jones, A. F., Sanger, R. H., Collis, L. P., Flannery, R., Mcnay, E. C., Yu, T., Schwarzenbacher, R., Bossy, B., Bossy-wetzel, E., Bennett, M. V., Pypaert, M., Hickman, J. A., Smith, P. J., Hardwick, J. M. & Jonas, E. A. 2008. Bcl-xL induces Drp1-dependent synapse formation in cultured hippocampal neurons. Proc Natl Acad Sci USA, 105, 2169–74. Li, W. W., Yang, R., Guo, J. C., Ren, H. M., Zha, X. L., Cheng, J. S. & Cai, D. F. 2007. Localization of alpha-synuclein to mitochondria within midbrain of mice. Neuroreport, 18, 1543–6. Li, Z., Okamoto, K., Hayashi, Y. & Sheng, M. 2004. The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses. Cell, 119, 873–87. Lin, W. & Kang, U. J. 2008. Characterization of PINK1 processing, stability, and subcellular localization. J Neurochem, 106, 464–74. Loeb, V., Yakunin, E., Saada, A. & Sharon, R. 2010. The transgenic overexpression of alpha-synuclein and not its related pathology associates with complex I inhibition. J Biol Chem, 285, 7334–43. Lutz, A. K., Exner, N., Fett, M. E., Schlehe, J. S., Kloos, K., Lammermann, K., Brunner, B., Kurz-drexler, A., Vogel, F., Reichert, A. S., Bouman, L., Vogt-weisenhorn, D., Wurst, W., Tatzelt, J., Haass, C. & Winklhofer, K. F. 2009. Loss of Parkin or PINK1 Function Increases Drp1-dependent Mitochondrial Fragmentation. J Biol Chem, 284, 22938–51. Martin, L. J., Pan, Y., Price, A. C., Sterling, W., Copeland, N. G., Jenkins, N. A., Price, D. L. & Lee, M. K. 2006. Parkinson’s disease alpha-synuclein transgenic mice develop neuronal mitochondrial degeneration and cell death. J Neurosci, 26, 41–50. Matsuda, N., Sato, S., Shiba, K., Okatsu, K., Saisho, K., Gautier, C. A., Sou, Y. S., Saiki, S., Kawajiri, S., Sato, F., Kimura, M., Komatsu, M., Hattori, N. & Tanaka, K. 2010. PINK1 stabilized by mitochondrial depolarization recruits Parkin to damaged mitochondria and activates latent Parkin for mitophagy. J Cell Biol, 189, 211–21. Meulener, M. C., Xu, K., Thomson, L., Ischiropoulos, H. & Bonini, N. M. 2006. Mutational analysis of DJ-1 in Drosophila implicates functional inactivation by oxidative damage and aging. Proc Natl Acad Sci USA, 103, 12517–22. Morrison, E., Thompson, J., Williamson, S. J., Cheetham, M. E. & Robinson, P. A. 2010. A simple cell based assay to measure Parkin activity. J Neurochem. Nakamura, K., Nemani, V. M., Wallender, E. K., Kaehlcke, K., Ott, M. & Edwards, R. H. 2008. Optical reporters for the conformation of alpha-synuclein reveal a specific interaction with mitochondria. J Neurosci, 28, 12305–17. Narendra, D., Tanaka, A., Suen, D. F. & Youle, R. J. 2008. Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. J Cell Biol, 183, 795–803. Narendra, D. P., Jin, S. M., Tanaka, A., Suen, D. F., Gautier, C. A., Shen, J., Cookson, M. R. & Youle, R. J. 2010a. PINK1 is selectively stabilized on impaired mitochondria to activate Parkin. PLoS Biol, 8, e1000298. Narendra, D. P., Kane, L. A., Hauser, D. N., Fearnley, I. M. & Youle, R. J. 2010b. p62/SQSTM1 is required for Parkin-induced mitochondrial clustering but not mitophagy; VDAC1 is dispensable for both. Autophagy, 6. Okatsu, K., Saisho, K., Shimanuki, M., Nakada, K., Shitara, H., Sou, Y. S., Kimura, M., Sato, S., Hattori, N., Komatsu, M., Tanaka, K. & Matsuda, N. 2010. p62/SQSTM1 cooperates with Parkin for perinuclear clustering of depolarized mitochondria. Genes Cells, 15, 887–900. Orth, M., Tabrizi, S. J., Schapira, A. H. & Cooper, J. M. 2003. Alpha-synuclein expression in HEK293 cells enhances the mitochondrial sensitivity to rotenone. Neurosci Lett, 351, 29–32. Panov, A., Dikalov, S., Shalbuyeva, N., Taylor, G., Sherer, T. & Greenamyre, J. T. 2005. Rotenone model of Parkinson disease: multiple brain mitochondria dysfunctions after short term systemic rotenone intoxication. J Biol Chem, 280, 42026–35. Park, J., Lee, G. & Chung, J. 2009. The PINK1-Parkin pathway is involved in the regulation of mitochondrial remodeling process. Biochem Biophys Res Commun, 378, 518–23.

212

C.T. Chu and S.B. Berman

Park, J., Lee, S. B., Lee, S., Kim, Y., Song, S., Kim, S., Bae, E., Kim, J., Shong, M., Kim, J. M. & Chung, J. 2006. Mitochondrial dysfunction in Drosophila PINK1 mutants is complemented by parkin. Nature, 441, 1157–61. Park, Y. Y., Lee, S., Karbowski, M., Neutzner, A., Youle, R. J. & Cho, H. 2010. Loss of MARCH5 mitochondrial E3 ubiquitin ligase induces cellular senescence through dynamin-related protein 1 and mitofusin 1. J Cell Sci, 123, 619–26. Pattingre, S., Tassa, A., Qu, X., Garuti, R., Liang, X. H., Mizushima, N., Packer, M., Schneider, M. D. & Levine, B. 2005. Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell, 122, 927–39. Plecita-hlavata, L., Lessard, M., Santorova, J., Bewersdorf, J. & Jezek, P. 2008. Mitochondrial oxidative phosphorylation and energetic status are reflected by morphology of mitochondrial network in INS-1E and HEP-G2 cells viewed by 4Pi microscopy. Biochim Biophys Acta, 1777, 834–46. Polymeropoulos, M. H., Lavedan, C., Leroy, E., Ide, S. E., Dehejia, A., Dutra, A., Pike, B., Root, H., Rubenstein, J., Boyer, R., Stenroos, E. S., Chandrasekharappa, S., Athanassiadou, A., Papapetropoulos, T., Johnson, W. G., Lazzarini, A. M., Duvoisin, R. C., Di Iorio, G., Golbe, L. I. & Nussbaum, R. L. 1997. Mutation in the alpha-synuclein gene identified in families with Parkinson’s disease. Science, 276, 2045–7. Poole, A. C., Thomas, R. E., Andrews, L. A., Mcbride, H. M., Whitworth, A. J. & Pallanck, L. J. 2008. The PINK1/Parkin pathway regulates mitochondrial morphology. Proc Natl Acad Sci USA, 105, 1638–43. Poole, A. C., Thomas, R. E., Yu, S., Vincow, E. S. & Pallanck, L. 2010. The mitochondrial fusionpromoting factor mitofusin is a substrate of the PINK1/parkin pathway. PLoS One, 5, e10054. Przedborski, S., Tieu, K., Perier, C. & Vila, M. 2004. MPTP as a mitochondrial neurotoxic model of Parkinson’s disease. J Bioenerg Biomembr, 36, 375–9. Sandebring, A., Thomas, K. J., Beilina, A., Van der Brug, M., Cleland, M. M., Ahmad, R., Miller, D. W., Zambrano, I., Cowburn, R. F., Behbahani, H., Cedazo-minguez, A. & Cookson, M. R. 2009. Mitochondrial alterations in PINK1 deficient cells are influenced by calcineurin-dependent dephosphorylation of dynamin-related protein 1. PLoS One, 4, e5701. Schapira, A. H., Cooper, J. M., Dexter, D., Clark, J. B., Jenner, P. & Marsden, C. D. 1990. Mitochondrial complex I deficiency in Parkinson’s disease. J Neurochem, 54, 823–7. Shavali, S., Brown-borg, H. M., Ebadi, M. & Porter, J. 2008. Mitochondrial localization of alphasynuclein protein in alpha-synuclein overexpressing cells. Neurosci Lett, 439, 125–8. Shiba, K., Arai, T., Sato, S., Kubo, S. I., Ohba, Y., Mizuno, Y. & Hattori, N. 2009. Parkin stabilizes PINK1 through direct interaction. Biochem Biophys Res Commun. Singleton, A. B., Farrer, M., Johnson, J., Singleton, A., Hague, S., Kachergus, J., Hulihan, M., Peuralinna, T., Dutra, A., Nussbaum, R., Lincoln, S., Crawley, A., Hanson, M., Maraganore, D., Adler, C., Cookson, M. R., Muenter, M., Baptista, M., Miller, D., Blancato, J., Hardy, J. & Gwinn-hardy, K. 2003. alpha-Synuclein locus triplication causes Parkinson’s disease. Science, 302, 841. Song, D. D., Shults, C. W., Sisk, A., Rockenstein, E. & Masliah, E. 2004. Enhanced substantia nigra mitochondrial pathology in human alpha-synuclein transgenic mice after treatment with MPTP. Exp Neurol, 186, 158–72. Spillantini, M. G., Schmidt, M. L., Lee, V. M., Trojanowski, J. Q., Jakes, R. & Goedert, M. 1997. Alpha-synuclein in Lewy bodies. Nature, 388, 839–40. Thomas, K. J. & Cookson, M. R. 2009. The role of PTEN-induced kinase 1 in mitochondrial dysfunction and dynamics. Int J Biochem Cell Biol, 41, 2025–35. Thomas, K. J., Mccoy, M. K., Blackinton, J., Beilina, A., Van der Brug, M., Sandebring, A., Miller, D., Maric, D., Cedazo-minguez, A. & Cookson, M. R. 2010. DJ-1 acts in parallel to the PINK1/parkin pathway to control mitochondrial function and autophagy. Hum Mol Genet. Twig, G., Elorza, A., Molina, A. J., Mohamed, H., Wikstrom, J. D., Walzer, G., Stiles, L., Haigh, S. E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B. F., Yuan, J., Deeney, J. T., Corkey, B. E. &

7  Mitochondrial Fission-Fusion and Parkinson’s Disease

213

Shirihai, O. S. 2008. Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. Embo J, 27, 433–46. Um, J. W., Stichel-gunkel, C., Lubbert, H., Lee, G. & Chung, K. C. 2009. Molecular interaction between parkin and PINK1 in mammalian neuronal cells. Mol Cell Neurosci, 40, 421–32. Van Laar, V., Arnold, B., Cassady, S., Chu, C., Burton, E. & Berman, S. 2011. Bioenergetics of neurons inhibit the translocation response of Parkin following rapid mitochondrial depolarization. Human Molecular Genetics, 20, 927– 40. Van Laar, V. S. & Berman, S. B. 2009. Mitochondrial dynamics in Parkinson’s disease. Exp Neurol, 218, 247–56. Vergne, I., Roberts, E., Elmaoued, R. A., Tosch, V., Delgado, M. A., Proikas-cezanne, T., Laporte, J. & Deretic, V. 2009. Control of autophagy initiation by phosphoinositide 3-phosphatase jumpy. Embo J, 28, 2244–2258. Vives-bauza, C., Zhou, C., Huang, Y., Cui, M., De Vries, R. L., Kim, J., May, J., Tocilescu, M. A., Liu, W., Ko, H. S., Magrane, J., Moore, D. J., Dawson, V. L., Grailhe, R., Dawson, T. M., Li, C., Tieu, K. & Przedborski, S. 2010. PINK1-dependent recruitment of Parkin to mitochondria in mitophagy. Proc Natl Acad Sci USA, 107, 378–83. Whitworth, A. J. & Pallanck, L. J. 2009. The PINK1/Parkin pathway: a mitochondrial quality control system? J Bioenerg Biomembr, 41, 499–503. Winklhofer, K. F. & Haass, C. 2010. Mitochondrial dysfunction in Parkinson’s disease. Biochim Biophys Acta, 1802, 29–44. Xiong, H., Wang, D., Chen, L., Choo, Y. S., Ma, H., Tang, C., Xia, K., Jiang, W., Ronai, Z., Zhuang, X. & Zhang, Z. 2009. Parkin, PINK1, and DJ-1 form a ubiquitin E3 ligase complex promoting unfolded protein degradation. J Clin Invest, 119, 650–60. Yang, Y., Gehrke, S., Imai, Y., Huang, Z., Ouyang, Y., Wang, J. W., Yang, L., Beal, M. F., Vogel, H. & Lu, B. 2006. Mitochondrial pathology and muscle and dopaminergic neuron degeneration caused by inactivation of Drosophila Pink1 is rescued by Parkin. Proc Natl Acad Sci USA, 103, 10793–8. Yang, Y. & Lu, B. 2009. Mitochondrial morphogenesis, distribution, and Parkinson disease: insights from PINK1. J Neuropathol Exp Neurol, 68, 953–63. Yang, Y., Ouyang, Y., Yang, L., Beal, M. F., Mcquibban, A., Vogel, H. & Lu, B. 2008. Pink1 regulates mitochondrial dynamics through interaction with the fission/fusion machinery. Proc Natl Acad Sci USA, 105, 7070–5. Youle, R. J. & Karbowski, M. 2005. Mitochondrial fission in apoptosis. Nat Rev Mol Cell Biol, 6, 657–63. Zarranz, J. J., Alegre, J., Gomez-esteban, J. C., Lezcano, E., Ros, R., Ampuero, I., Vidal, L., Hoenicka, J., Rodriguez, O., Atares, B., Llorens, V., Gomez Tortosa, E., Del Ser, T., Munoz, D. G. & De Yebenes, J. G. 2004. The new mutation, E46K, of alpha-synuclein causes Parkinson and Lewy body dementia. Ann Neurol, 55, 164–73. Zhang, L., Zhang, C., Zhu, Y., Cai, Q., Chan, P., Ueda, K., Yu, S. & Yang, H. 2008. Semiquantitative analysis of alpha-synuclein in subcellular pools of rat brain neurons: an immunogold electron microscopic study using a C-terminal specific monoclonal antibody. Brain Res, 1244, 40–52. Zhu, J.-H., Guo, F., Shelburne, J., Watkins, S. & Chu, C. T. 2003. Localization of phosphorylated ERK/MAP kinases to mitochondria and autophagosomes in Lewy body diseases. Brain Pathol, 13, 473–481. Zhu, J. H., Horbinski, C., Guo, F., Watkins, S., Uchiyama, Y. & Chu, C. T. 2007. Regulation of autophagy by extracellular signal-regulated protein kinases during 1-methyl-4-phenylpyridiniuminduced cell death. Am J Pathol, 170, 75–86. Ziviani, E., Tao, R. N. & Whitworth, A. J. 2010. Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc Natl Acad Sci USA, 107, 5018–23.

Chapter 8

Role of the Mitochondrial Fission Protein Drp1 in Synaptic Damage and Neurodegeneration Tomohiro Nakamura, Dong-Hyung Cho, and Stuart A. Lipton

Abstract  Under neurodegenerative conditions, abnormal mitochondrial morphology is often associated with synaptic injury and neuronal damage. Mitochondrial fission and fusion are dynamic processes that represent a key feature controlling mitochondrial structure and morphology. Members of the dynamin superfamily of GTPases and associated proteins are known to function in both mitochondrial fission and fusion. For example, mitofusin and OPA1 are essential for mitochondrial fusion, and dynamin-related protein 1 (Drp1), along with critical partners, mediates mitochondrial fission. In this review, we primarily focus on the mitochondrial fission protein Drp1, and discuss recent insights that may potentially link aberrant S-nitrosylation Drp1 as well as other posttranslational modifications (phosphorylation, sumoylation, and ubiquitination) to synaptic damage, neuronal cell injury, and cell death. We propose the hypothesis that dysfunctional Drp1 activity can contribute to the pathogenesis of neurodegenerative diseases including Alzheimer’s and Parkinson’s diseases. Keywords  Mitochondrial dynamics • Neurodegenerative diseases • Dynaminrelated protein 1 • Synaptic damage • Neuronal cell death Abbreviations Ab b-amyloid protein ABAD Ab-binding alcohol dehydrogenase AD Alzheimer’s disease

T. Nakamura and S.A. Lipton () Center for Neuroscience, Aging, and Stem Cell Research, Sanford-Burnham Medical Research Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA e-mail: [email protected] D.-H. Cho Graduate School of East-West Medical Science, Kyung Hee University, Yongin, Gyeonggi 446–701, Korea B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_8, © Springer Science+Business Media B.V. 2011

215

216

ADDL ADOA APP CDK5 CMT2A Drp1 GED HR IMM MAPL Mfn MLS MOMP NO OMM Opa1 PD PTM SENP SUMO

T. Nakamura et al.

Ab-driven diffusible ligand autosomal dominant optic atrophy amyloid precursor protein cyclin-dependent kinase 5 Charcot-Marie-Tooth neuropathy type2A dynamin-related protein 1 GTPase effector domain heptad repeat inner mitochondrial membrane mitochondrial-anchored protein ligase mitofusin mitochondrial localization sequence mitochondrial outer membrane permeabilization nitric oxide outer mitochondrial membrane optic atrophy protein 1 Parkinson’s disease posttranslational modification Sentrin/SUMO-specific protease small ubiquitin-related modifier 1.

8.1 Introduction The mitochondrial respiration system produces ATP from oxygen, glucose, and fatty acids, making the mitochondrion a critical energy source for virtually all eukaryotic cells. In addition to respiration activity, mitochondria are important for Ca2+ storage as well as regulation of apoptotic cell death. The mitochondrion is composed of inner and outer membranes. Fission or fusion of these membranes results in mitochondrial division or unification, respectively, thus regulating processes associated with mitochondrial morphology such as biogenesis, subcellular localization, and distribution of mitochondria, all of which can ultimately affect mitochondrial bioenergetics (Chan 2006a, b; Liesa et  al. 2009). Several important regulators of mitochondrial fission and fusion have been discovered (Alexander et  al. 2000; Delettre et  al. 2000; Hales and Fuller 1997; Santel and Fuller 2001; Shin et  al. 1997; Smirnova et  al. 1998). For instance, when mitochondria undergo fission, the large GTPase, dynamin-related protein 1 (Drp1), translocates from the cytosol to the outer mitochondrial membrane (OMM) to initiate fission or fragmentation (Praefcke and McMahon 2004; Smirnova et al. 2001). RNAi-mediated knockdown of Drp1 or overexpression of a dominant-negative Drp1 mutant (K38A, with defective GTPase activity) inhibits fission and delays cytochrome c release, caspase activation, and thus impedes cell death, suggesting that Drp1-mediated mitochondrial fission can contribute

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

217

to apoptosis, at least under certain conditions (Frank et al. 2001; Smirnova et al. 2001; Yoon et  al. 2001). On the other hand, three large GTPase dynamin like proteins, mitofusin1 (Mfn1), mitofusin2 (Mfn2), and optic atrophy protein 1 (Opa1), mediate mitochondrial fusion (Chan 2006b; Hoppins and Nunnari 2009). Specifically, Mfn1/2 mediates OMM fusion, and Opa1 activity is associated with inner mitochondrial membrane (IMM) fusion. In contrast to mitochondrial fission, fusion functions in part as a cell protective mechanism. Silencing Mfn1/2 results in mitochondrial fragmentation and increases mitochondrial susceptibility to apoptotic stimuli. Similarly, loss of Opa1 induces disruption of mitochondrial cristae as well as spontaneous apoptosis (Olichon et  al. 2003; Suen et al. 2008). Recently, emerging evidence has suggested that an imbalance between mitochondrial fission and fusion (collectively designated ‘mitochondrial dynamics’) contributes to the pathophysiology of neurodegenerative and metabolic disorders (Su et al. 2010; Zorzano et al. 2009). Dysfunction of mitochondrial dynamics and resulting disease can emanate not only from mutations of the genes encoding fission- or fusion-related proteins but also, as our group has recently shown, from posttranslational protein modifications that regulate stability and activity. Evidence of genetic mutations in Mfn1/2 and Opa1 is found in patients with Charcot-Marie-Tooth neuropathy type2A (CMT2A) (Cartoni and Martinou 2009; Lawson et al. 2005; Zuchner et al. 2004) and autosomal dominant optic atrophy (ADOA), respectively (Alexander et al. 2000; Delettre et al. 2000; Olichon et al. 2006). These mutations are thought to underlie the disease process. Furthermore, a heterozygous missense mutation of Drp1 (A395D) has been identified in a patient manifesting both ADOA- and CMT2A-like symptoms (Waterham et  al. 2007). Taken together, these findings suggest that neurons are particularly susceptible to dysfunction of mitochondrial dynamics because these fission- and fusion-related proteins are widely expressed throughout the body yet disease is often manifest in neurons. Posttranslational modifications (PTMs) also regulate the activity of mitochondrial fission/fusion proteins. Among these are phosphorylation, S-nitrosylation, sumoylation, ubiquitination, and proteolytic processing of Drp1 as well as Opa1 (Chang and Blackstone 2007; Cho et  al. 2009; Hajek et al. 2007; Han et al. 2008; Harder et al. 2004; Ishihara et al. 2006; Nakamura et al. 2006; Taguchi et al. 2007; Wasiak et al. 2007; Zunino et al. 2007). These PTMs can also affect intracellular localization and stability, as discussed in detail in a later section. Intriguingly, in experimental models some of the Drp1 PTMs can recapitulate key pathological features seen in patients with genetic mutations, potentially linking the PTMs of fission/fusion proteins to neurodegeneration in sporadic cases. Taken together, dysfunction of mitochondrial dynamics may serve as a hallmark of many common neurodegenerative diseases as well as a contributing factor to their pathophysiology. In this review, we focus on the well-characterized mitochondrial fission protein, Drp1, and discuss mitochondrial dynamics from the perspective of cell death and neurodegeneration.

218

T. Nakamura et al.

8.2 Mitochondrial Fission and Fusion Machinery Despite extensive data currently available regarding the machinery of mitochondrial fission and fusion, the precise molecular mechanisms relevant to mitochondrial dynamics are not entirely understood. However, several key players in mitochondrial dynamics have been identified. Drp1 is a member of the conserved dynamin GTPase superfamily, which includes a broad range of membrane fission proteins (Praefcke and McMahon 2004; Smirnova et al. 2001). Drp1 exhibits specific fission activity on mitochondrial and peroxisomal membranes (Koch et al. 2003; Pitts et al. 2004) (Fig. 8.1). The mechanism exploited in the recruitment of Drp1 to the mitochondrial membrane for fission remains largely unclear. Fis1, localized on the OMM, functions as a Drp1 receptor. Fis1 overexpression accelerates mitochondrial fission, whereas Fis1 silencing by RNAi inhibits mitochondrial fragmentation (James et  al. 2003; Yoon et al. 2003). Although Fis1 acts downstream of Drp1, the role of Fis1 in Drp1 recruitment in mammalian cells is still unclear because Drp1 still localizes to the OMM after RNAi knockdown of Fis1 (James et  al. 2003; Lee et  al. 2004; Yoon et al. 2003). Therefore, further studies will be required to understand the detailed mechanism for the interaction of Fis1 and Drp1. Mitochondrial fusion is a well-coordinated process in which the inner and outer membranes fuse separately. The key components of the machinery of mitochondrial fusion are Mfn1/2 and Opa1. Mfn1/2 is localized to the OMM with both its N- and

Fig. 8.1  Drp1 mediates mitochondrial fission. Drp1 oligomers form a ring-like structure at the site of mitochondrial fission. Constriction of the Drp1 ring structure is thought to produce mitochondrial fission, resulting in two mitochondria

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

219

C-termini facing into the cytosol (Rojo et  al. 2002). Structurally, the mitofusins harbor a GTPase domain in the N-terminus, a bipartite transmembrane domain, and two heptad repeat (HR) coiled-coil domains in the middle and C-terminal regions (Santel 2006). Mfn1/2 function is dependent on GTPase activity. Loss of function mutations in the GTPase domain (K88T, T109A mutants) disrupt Mfn fusion activity, whereas the gain-of-function mutant [Mfn2(G12V)] exhibits increased Mfn fusion activity (Karbowski et al. 2006; Neuspiel et al. 2005; Santel et al. 2003). The HR2 domain initiates mitochondrial fusion via oligomerization of Mfn, which leads to tethering of two adjacent mitochondria. Thus, mutations in the HR region also inhibit the fusion activity of Mfn (Koshiba et al. 2004). Knockout of Mfn1 or Mfn2 results in the formation of small fragmented mitochondria, suggesting that both Mfn1 and Mfn2 mediate mitochondrial fusion (Chen et  al. 2003, 2007). Mfn2 is predominantly expressed in heart and skeletal muscle, in contrast to the more general expression of Mfn1. Mfn1 depletion results in a more pronounced phenotype than Mfn2 (Bach et  al. 2003; Chen et  al. 2005; Neuspiel et  al. 2005). Moreover, Opa1 functions in an Mfn1 but not Mfn2-dependent manner, in the mitochondrial fusion pathway (Cipolat et al. 2004). Therefore, the various isoforms may perform specific roles with distinctive activities. Opa1 is another key factor implicated in mitochondrial fusion and cristae remodeling. The mitochondrial localization sequence (MLS) in the N-terminal region of Opa1 is responsible for importing the protein into the IMM. In addition to the MLS, a transmembrane domain anchors Opa1 in the IMM. In the middle region, Opa1 also contains a GTPase domain that is crucial for activity. Mutations in this GTPase domain are associated with the generation of fragmented mitochondria (Olichon et al. 2007). The middle domain, which is located next to the GTPase domain, is possibly involved in tetramerization and higher order assembly of Opa1 (Ramachandran et al. 2007). The C-terminus harbors a coiled-coil region thought to present in the GTPase effector domain (GED), which is also involved in oligomerization and activation of Opa1 (Praefcke and McMahon 2004). Similar to the effect on mitofusins, RNAi against Opa1 leads to fragmentation of mitochondria (Chen et al. 2005; Olichon et al. 2003). This may be attributable to an increased mitochondrial division rate or to a reduction in the fusion capacity of the mitochondria. It has been reported that overexpression of Opa1 leads to complicated responses, such as mitochondrial elongation or fragmentation, depending on the overexpression system (Chen et al. 2005; Cipolat et al. 2004; Griparic et al. 2004; Olichon et  al. 2003). However, the mechanism of this paradoxical effect remains to be elucidated. Mice lacking Mfn1/2 or Opa1 genes manifest embryonic lethality (Chen et al. 2003; Alavi et al. 2007; Davies et al. 2007), suggesting that these fusion proteins are required for embryonic development in mice. Similarly to ADOA patients, OPA1 heterozygous mice show degeneration in the optic nerve system. Taken together, these findings establish an important role for these dynamin-like proteins in mitochondrial dynamics, and suggest that impaired balance between fission and fusion is associated with neurophathological features.

220

T. Nakamura et al.

8.3 Structure and Function of Mitochondrial Fission Protein Drp1 The human Drp1 gene is located on chromosome 12 and encodes at least four different splice variants (Howng et al. 2004; Smirnova et al. 1998). Drp1 contains several functional domains, including an N-terminal GTPase domain, which is thought to provide mechanical force, a dynamin-like middle domain, an insert B domain, and a GTPase effector domain (GED), which is located in the C-terminal region (Bossy-Wetzel et al. 2003; Liesa et al. 2009) (Fig. 8.2). Intra- and intermolecular interactions between the GTPase domain and GED or intermolecular interaction of GEDs are required for the assembly and promotion of higher order structures (dimers or tetramers) (Zhang and Hinshaw 2001; Zhu et al. 2004). Selfassembly and assembly stimulated GTP hydrolysis are functionally essential features of the dynamin superfamily of proteins (Damke et al. 2001; Ingerman et al. 2005; Lackner et al. 2009; Ramachandran et al. 2007). The GTP-binding defective mutant (K38A) inactivates Drp1 by sequestration of endogenous Drp1, thus inhibiting its mitochondrial localization; therefore, it acts as a dominant negative. GED is also involved in the regulation of GTPase activity and its assembly. Mutations in the GED domain also significantly change Drp1 activity (Zhu et al. 2004). Mice lacking the Drp1 gene manifest embryonic lethality due to developmental abnormalities in heart, liver, forebrain and placenta (Ishihara et al. 2009; Wakabayashi et al. 2009), indicating that Drp1 is required for embryonic development. In embryonic stem cells and fibroblasts obtained from Drp1−/− mice, fluorescence immunocytochemistry and electron microscopy studies revealed the presence of elongated mitochondria, which could be rescued by the expression of Drp1. Furthermore, knockout of Drp1 prevented mitochondrial fragmentation due to CCCP (carbonyl cyanide m-chlorophenylhydrazone, a mitochondrial membrane disrupting agent).

Fig. 8.2  Schematic structure of the domains of Drp1, and posttranslational modification of Drp1. The length and topology of amino acids for each region are shown. Drp1 has a GTPase domain responsible for hydrolysis of GTP. Drp1 also has a middle domain, an insert B domain, and a GTPase effector domain (GED). The locations of posttranslational modifications are indicated by arrows (for phosphorylation and S-nitrosylation) or brackets ‘}’ (for sumoylation at multiple lysine residues in the insert B domain). The effect of posttranslational modifications on mitochondrial fission activity is indicated as increased (⇑) or (decreased (⇓)

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

221

This result is consistent with the notion that Drp1 can mediate mitochondrial fragmentation. Interestingly, Drp1 knockout did not significantly affect mitochondrial respiration, ATP levels, or mitochondrial DNA content. However, it is still not known whether aberrant Drp1 activity can contribute to impaired mitochondrial bioenergetics at a specific subcellular location (e.g., neuronal synapses). Moreover, Drp1-deficient cells grew slightly slower than wild-type cells, suggesting that altered cell proliferation might have caused the embryonic lethality in Drp1 knockout mice. Recently, a human case study reported a dominant-negative, lethal mutation (A395D) in the middle domain of Drp1 (Waterham et al. 2007). The newborn infant with this mutation suffered from microcephaly, aberrant brain development, and optic atrophy, and died suddenly at the age of 37 days. Despite the elevated lactate levels in plasma and cerebrospinal fluid, activity of the mitochondrial respiratory chain remained normal in both cultured skin fibroblast and skeletal-muscle biopsy specimens. Interestingly, fibroblasts derived from this patient displayed elongated mitochondrial structure, while the muscle-biopsy sample showed no morphological abnormalities in mitochondria. Biochemically, the A395D Drp1 mutant retained the ability to form tetramers in vitro; however, it showed defects in forming higher order assembly and subsequent stimulation of GTPase activity. Moreover, A395D mutation impaired the ability of Drp1 to localize to mitochondria (Chang et  al. 2010). These findings were used to argue that lethality occurred because of disrupted Drp1 activity and a resultant decrease in mitochondrial fission in certain cell types (i.e., fibroblasts and presumably neurons).

8.4 Posttranslational Modification of Drp1 8.4.1 Phosphorylation Posttranslational modifications have been implicated as regulatory mechanisms that mediate a variety of Drp1 activities during mitochondrial fission. The first of these modifications is phosphorylation (Fig.  8.2). For example, during mitosis, Drp1 is activated by the CDK1/Cyclin B-mediated phosphorylation of serine 616 on the GED of the longest isoform of human Drp1 (serine 585 in rat) (Taguchi et al. 2007). This phosphorylation event facilitates the proper distribution of mitochondria within daughter nascent cells. Another serine residue (serine 637) within the GED is also phosphorylated by cyclic AMP-dependent protein kinase (PKA) (Chang and Blackstone 2007; Cribbs and Strack 2007). Unlike CDK1, PKA-induced phosphorylation results in reduced activity by inhibiting the intramolecular interaction of Drp1. Overexpression of Drp1(S637D), a phospho-mimetic mutant, significantly elongates mitochondria. Additionally, calcineurin is involved in dephosphorylation and regulates translocation of Drp1 to mitochondria (Cereghetti et  al. 2008). However, a recent study demonstrated that calcium/calmodulin-dependent protein kinase Ia (CaMKIa) phosphorylates Drp1 at the same serine residue (Ser637) and that this phosphorylation is associated with increased translocation of Drp1 to

222

T. Nakamura et al.

mitochondria, resulting in increased affinity for Fis1 (Han et  al. 2008). Finally, cyclin-dependent kinase 5 (CDK5) has been identified as another relevant mitochondrial fission-regulatory kinase, but whether or not CDK5 directly regulates Drp1 remains to be determined (Meuer et al. 2007). These reports thus suggest that phosphorylation of Drp1 has both positive and negative effects on its fission activity. These insights raise the possibility of phosphorylation-dependent regulation of Drp1 in neurodegeneration; additional experiments will be needed to test this hypothesis.

8.4.2 S-Nitrosylation In addition to phosphorylation, our group and subsequently others have recently identified Drp1 as a substrate for reaction with nitric oxide (NO) species, leading to S-nitrosylation, a redox reaction involving transfer of NO to a critical cysteine thiol, which regulates protein function (Cho et  al. 2009; Wang et  al. 2009) (Fig.  8.2). S-Nitrosylation of Drp1 (forming SNO-Drp1), enhances GTPase activity and oligomer formation, leading to excessive mitochondrial fission or fragmentation in neurons. By mutational analysis, we surmised that NO reacts with cysteine residue 644 of Drp1 within the GED. Mutation of this target cysteine prevented excessive mitochondrial fragmentation and ameliorated synaptic damage and neurotoxicity induced by NO or b-amyloid protein (Ab, which indirectly increases NO) (Cho et al. 2009). Cyclic AMP and calcium, as well as NO, are key second messengers in the control of cellular metabolism and homeostasis. Thus, current findings suggest that intracellular signals control and alter mitochondrial morphology by regulating Drp1 activity.

8.4.3 Sumoylation Sumoylation also affects Drp1 activity. Small ubiquitin-related modifier 1 (SUMO1) is bound to the insert B domain of Drp1 and functions as a Drp1stabilizing protein via sumoylation, thereby resulting in mitochondrial fission (Figueroa-Romero et al. 2009; Harder et al. 2004) (Fig. 8.2). In contrast, Sentrin/ SUMO-specific protease (SENP5) works as a desumoylation enzyme on Drp1 (Zunino et al. 2007). Overexpression of SUMO1 evokes mitochondrial fragmentation, while SENP5 rescues SUMO-induced mitochondrial fragmentation (Harder et al. 2004; Zunino et al. 2007).

8.4.4 Ubiquitination Beside sumoylation, ubiquitination of Drp1 also regulates mitochondrial dynamics. Mitochondrial-anchored protein ligase (MAPL) and mitochondrial E3 ligase protein (MARCH5/MITOL) function as ubiquitin E3 ligases for Drp1 (Braschi et al. 2009;

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

223

Karbowski et al. 2007; Nakamura et al. 2006; Yonashiro et al. 2006). However, the effect of MARCH5 on mitochondrial dynamics has not been clearly identified. MARCH5-deficient cells (produced by small interfering RNA knockdown) or cells with a MARCH5 mutant lacking ubiquitin ligase activity manifested aberrant mitochondrial morphology, including fragmentation, in previously reported studies (Nakamura et al. 2006; Yonashiro et al. 2006). However, Karbowski et al. reported that MARCH5 RING mutants and knockdown of MARCH5 by RNAi resulted in abnormal elongation and interconnections of mitochondria (Karbowski et al. 2007). Therefore, Drp1 sumoylation is believed to protect against degradation, but the function of ubiqutination of Drp1 warrants further study.

8.5 Drp1, Mitochondrial Dynamics, and Synaptic Morphology One of the key morphological features that represents a functional, mature neuron is the presence of synapses, where neurons signal to one another. Synaptic loss occurs early in the neurodegenerative process in disorders such as Alzheimer’s disease (AD), Parkinson’s Disease (PD), and Huntington’s Disease (Scheff and Price 1998; Zhan et al. 1993). Currently, it is generally accepted that such synaptic degeneration contributes to the cognitive decline seen in these patients. Proper distribution of mitochondria is critical for synaptic integrity, e.g., the maintenance of dendritic spines (representing the postsynaptic protruberances), because of the role of mitochondria in Ca2+ homeostasis and ATP production (Li et al. 2004). Early investigations by Sheng and colleagues demonstrated that expression of dominant-negative Drp1 caused mitochondrial depletion from dendritic spines, which was associated with a decline in spine density (Li et al. 2004). Similar synaptic aberration was observed in neurons overexpressing OPA1. In contrast, overexpression of Drp1 increases mitochondrial mass at postsynaptic structures. Mitochondria are also present at the synaptic terminal (presynaptically), controlling neurotransmitter release. For instance, in Drosophila neuromuscular junctions, mutations in the drp1 gene resulted in a massive depletion of mitochondria from the presynaptic terminal. Intriguingly, the absence of mitochondria caused defects in the mobilization of the reserve pool of vesicles and failed to maintain normal neurotransmission when synapses were stimulated at high frequency (Verstreken et al. 2005). Addition of ATP partially reversed the observed defects, suggesting that ATP production and bioenergetics were impaired in the Drp1 mutant flies. Along these lines, neuronal cultures prepared from the neuron-specific Drp1 knockout mouse displayed significantly less neurites and dendritic branches, which were associated with diminished expression of synaptic markers, suggesting that normal Drp1 activity is indispensable for synapse formation as well as the maintenance of synaptic structure. Consistent with these findings, enlarged mitochondria were encountered in Drp1-deficient neurons and failed to localize to synapses (Ishihara et al. 2009). Thus, neuron-specific Drp1 knockout mice exhibit enlarged mitochondria, with reduction of forebrain size, increased neuronal cell death in deep cortical layers, and periventricular leukomalacia (Ishihara et al. 2009).

224

T. Nakamura et al.

In living neurons, Bcl-w and Bcl-xL appear to maintain proper spine or synaptic structures via regulation of mitochondrial morphology and biomass (Berman et al. 2009; Li et al. 2008; Liu and Shio 2008). Interestingly, Bcl-xL forms a complex with Drp1 and stimulates its GTPase activity at presynaptic sites, facilitating synaptic formation. However, the mechanism whereby Bcl-2 family proteins regulate Drp1 activity remains to be elucidated (Autret and Martin 2009). Taken together, these observations clearly support the notion that Drp1-regulated mitochondrial dynamics play an important role in synaptic function. Moreover, under neurodegenerative conditions, as detailed in Sects. 8.4.2 and 8.7, S-nitrosylation can excessively activate synaptic Drp1 and contribute to neurodegeneration. Additional molecular and functional analyses of synaptic Drp1 activity will provide further insight into the physiological and pathophysiological roles of Drp1 at synapses.

8.6 Drp1, Mitochondrial Dynamics, and Cell Death Mitochondria play a pivotal role in cell death, mediating both intrinsic and extrinsic signaling pathways. Mitochondrial outer membrane permeabilization (MOMP) leads to increased ROS generation, reduced ATP levels, and the release of several pro-apoptotic factors, including cytochrome C, Smac, and apoptosis-inducing factor (Ow et al. 2008; Zamzami and Kroemer 2001). Mitochondrial membrane permeability is tightly regulated by Bcl-2 family proteins, and is determined by the ratio of pro- and anti-Bcl-2 protein complexes (Chipuk and Green 2008). Mitochondria undergo extensive fragmentation during apoptosis, and Drp1 inhibition suppresses apoptosis, suggesting that mitochondrial fission may be actively involved in apoptosis. Mitochondrial fragmentation occurs before caspase activation. Overexpression of a dominant negative form of Drp1 inhibits not only mitochondrial fission but also loss of mitochondrial membrane potential and cytochrome C release (Frank et  al. 2001). In line with these findings, chemical inhibition of Drp1 (mdivi-1) significantly blocks Bax/Bak-induced cytochrome C and Smac release from mitochondria (Cassidy-Stone et al. 2008). Moreover, inhibition of Fis1 also suppresses apoptosis, whereas overexpression of Fis1 induces both mitochondrial fission and cytochrome C release (James et  al. 2003; Lee et  al. 2004). Conversely, Mfn1/2 overexpression inhibits apoptosis (Sugioka et  al. 2004). Knockdown of Opa1 not only induces fission, but also induces spontaneous cytochrome C release and subsequent cell death (Olichon et al. 2003). Opa1 mutants defective in GTPase activity are incapable of protecting cells from apoptotic stimuli, indicating that the pro-survival and mitochondrial fusion effects of Opa1 might be linked (Frezza et al. 2006). Bcl-2 family proteins, which regulate mitochondrial permeabilization, functionally associate with mitochondrial fission and fusion proteins. For example, during apoptosis, Bax and Bak associate on the surface of the OMM, where they colocalize with Drp1 and Mfn2 (Karbowski et al. 2002). Drp1 can participate in apoptosis signaling via stimulating Bax oligomerization and thus increasing MOMP

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

225

(Montessuit et al. 2010). Furthermore, mitochondrial fragmentation is attenuated in Bak-deficient cells, and Bax activation is inhibited by a Mfn2 dominant-active mutant, suggesting that Bax and Bak also control mitochondrial fusion through Mfn2 (Karbowski et al. 2006; Neuspiel et al. 2005). The above investigations notwithstanding, whether mitochondrial fission plays an active role in MOMP, cytochrome C release, and apoptosis remains unclear. For example, a series of recent studies suggested that Bcl-2 family proteins might influence mitochondrial dynamics independent of anti-apoptotic activity (Sheridan et al. 2008; Delivani et al. 2006). In these reports, Bax induces release of cytochrome C and apoptosis as well as mitochondrial fragmentation; however, overexpression of Bcl-xL or Bcl-2 protein only inhibits Bax-induced release of cytochorome C but not Bax-mediated mitochondrial fragmentation. Additionally, Fis1 stimulates mitochondrial fragmentation without apoptosis, and inhibition of the mitochondrial fission machinery does not prevent Bax/Bak-mediated apoptosis (Alirol et al. 2006; Parone et al. 2006). Moreover, in Mfn1/2 deficient cells, mitochondria are extensively fragmented, but apoptosis is not enhanced (Chen et al. 2003, 2005). Additional experiments with Drp1 knockout cells produced somewhat puzzling results. In immortalized fibroblasts and embryonic stem cells, deletion of the Drp1 gene delays cytochrome C release, caspase activation, and subsequent cell death, but translocation of Bax is not affected. In contrast, the Drp1 knockout (1) increases caspase activation in neuronal progenitor cells, but (2) does not affect caspase activation in non-immortalized fibroblasts. These observations suggest that the effects of Drp1 activity and mitochondrial fission on apoptotic cell death may depend on the cell type or redox environment, as discussed in Sect. 4.2. In summary, current reports suggest that the mitochondrial fission machinery may contribute to or regulate apoptotic events, but is not invariably linked to apoptosis. The exact nature between mitochondrial dynamics and apoptosis needs to be studied further.

8.7 Drp1 and Mitochondrial Dynamics in Alzheimer’s and Parkinson’s Diseases Mitochondria obviously play a critical role in energy generation. This makes mitochondria particularly important in neurons, owing to their high demands for energy because of their specialized functions, complex morphology, and synaptic activity. Mitochondrial function is directly linked to mitochondrial dynamics, and vice versa. Recent studies have linked mitochondrial dysfunction to neurodegenerative diseases (Knott et  al. 2008), suggesting that an imbalance in mitochondrial dynamics may contribute to both familial and sporadic neurodegenerative disorders (Su et al. 2010). In this section, we will discuss the effects of mitochondrial dynamics in neurodegenerative diseases, such as Alzheimer’s disease and Parkinson’s disease.

226

T. Nakamura et al.

AD is the most frequent age-related neurodegenerative disorder and currently has a prevalence approaching 40% among people 80 years of age or older. A typical clinical symptom of AD involves the deterioration of selective cognitive domains, particularly those related to learning and memory (LaFerla et al. 2007). Accumulation of amyloid plaques, representing aggregates of Ab protein, and neurofibrillary tangles, composed of hyperphosphorylated tau, is thought to contribute to the disruption of neuronal synapses, connectivity and plasticity, with eventual neuronal demise (Brunden et al. 2009; Yankner and Lu 2009). b- and g-secretase prototypically process amyloid precursor protein (APP) in the transmembrane region to generate Ab (Haass and Selkoe 2007; LaFerla et  al. 2007). Oligomeric Ab may contribute to pathology by disrupting calcium homeostasis and synaptic function, possibly via increases in reactive oxygen/nitrogen species and inflammatory responses (Selkoe 2000; Wyss-Coray 2006). Several metabolic defects occur in AD (Blass 2000). Not only are mitochondria damaged, but also resulting oxidative stress has been shown to be present in AD. Moreover, several key molecules in oxidative metabolism have been shown to be defective in AD, consistent with the notion that mitochondrial dysfunction plays a prominent role in disease pathogenesis (Wang et al. 2009). One school of thought holds that TOM (translocase of the outer mitochondria membrane) may transport Ab into mitochondria, directly inducing mitochondrial dysfunction (Chen and Yan 2007; Hansson Petersen et  al. 2008). Ab may disrupt mitochondrial function via inhibition of key enzymes in respiratory metabolism, such as pyruvate dehydrogenase, cytochrome oxidase, and ABAD (Ab-binding alcohol dehydrogenase). As discussed above, mitochondrial dynamics influence mitochondrial function; emerging studies indicate that abnormal mitochondrial dynamics may play a crucial role in the pathogenesis of AD. Indeed, as our group and others have shown, overexpression of APP or exposure to Ab induces mitochondrial fragmentation and abnormal mitochondrial distribution, resulting in mitochondrial and neuronal dysfunction (Casley et al. 2002; Crouch et al. 2005; Lustbader et al. 2004; Barsoum et al. 2006). We recently reported that NO can lead to mitochondrial fragmentation via S-nitrosylation of Drp1 at cysteine residue 644, which is located within the GED domain that regulates GTPase activity and oligomerization (Cho et al. 2009). NO is a signaling molecule involved in several important physiological processes including neurotransmitter release and plasticity. However, when NO is excessively produced, it can contribute to neurotoxicity by inducing nitrosative stress, which contributes to the pathogenesis of neurodegenerative diseases. We found that exposure of neurons to oligomerized Ab leads to S-nitrosylation of Drp1 (forming SNO-Drip1), thus hyperactivating this fission protein and causing mitochondrial fragmentation. This fragmentation of mitochondria contributes to synaptic damage and subsequent neuronal cell death (Fig. 8.3). Importantly, substitution of an alanine for cysteine residue 644 in Drp1 suppresses Ab-induced synaptic damage and cell death, suggesting that SNO-Drp1 is a critical step in the Ab-mediated pathogenesis of AD. Subsequently, it was reported that the expression of levels of Drp1, Opa1, and Mfn1/2 are significantly decreased in hippocampal neurons of human AD brains, whereas Fis1 levels are increased (Wang et al. 2009). Moreover, the expression level of Drp1 is reduced in

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

227

Fig.  8.3  Proposed model of SNO-Drp1-induced mitochondrial fission at the synapse. (Left) Under physiological conditions, basal levels of NO, stimulated by normal synaptic activity of N-methyl-d-aspartate-type glutamate receptors (NMDAR), do not impair mitochondrial function. (Right) Under pathophysiological conditions, overactivation of extrasynaptic or perisynaptic NMDA receptors results in increased production of NO. Ab oligomers may also increase neuronal NO production via both NMDA receptor-dependent and –independent (e.g., iNOS) mechanisms. Excessive NO produces S-nitrosylation of Drp1 (forming SNO-Drp1), which contributes to neuronal synaptic injury by leading to excessive mitochondrial fission and bioenergetic impairment

AD brains compared to controls. Nonetheless, the level of mitochondrial Drp1 is similar in neurons from AD samples or even increased after exposure to oligomerized Ab protein (or Ab-driven diffusible ligands (ADDLs)). Interestingly, phosphorylated or S-nitrosylated Drp1, which possesses increased Drp1 activity, is upregulated in AD tissues and ADDL-exposed neurons (Cho et al. 2009; Taguchi et  al. 2007; Wang et  al. 2009). In summary, emerging evidence suggests that impaired mitochondrial dynamics are involved in the pathogenesis of AD. PD is the second most common neurodegenerative disease following AD, and the most common movement disorder, affecting approximately 1% of people aged 60 years or older. It is characterized by the progressively diminished ability to initiate voluntary movements owing to the loss of dopaminergic neurons in the substantia nigra (Abou-Sleiman et al. 2006). Mitochondrial dysfunction has long been implicated in the pathogenesis of PD. Mitochondrial respiratory electron transport chain activity, predominantly NADH dehydrogenase (Complex I), is reduced in the substantia nigra of PD patients, and complex I inhibitors, such as rotenone, MPP+ and pesticides, result in neuropathological changes similar to PD (Schapira et al. 2006). Not only are levels of multiple mitochondrial proteins altered in postmortem samples of PD brains, but also PD-linked genetic mutations in PINK1, Parkin and DJ-1 have been identified to regulate mitochondrial morphology, suggesting that mitochondrial function may be altered in PD (Abou-Sleiman et al. 2006; Jin et al. 2007). Recent evidence suggests that abnormal mitochondrial dynamics may contribute to neuronal injury and death in animal models of PD. For example, both rotenone and 6-hydroxydopamine have been shown to induce Drp1-dependent mitochondrial fragmentation as well as oxidative stress (Barsoum et al. 2006; Gomez-Lazaro et al. 2008). Additionally, loss of function of PINK1 or Parkin leads to mitochondrial fragmentation, which is associated with enhanced mitophagy (Dagda et  al. 2009; Exner et al. 2007; Lutz et al. 2009). Further evidence comes from studies of PINK1

228

T. Nakamura et al.

mutations underlying some forms of hereditary PD. Mammalian fibroblasts carrying PINK1 mutations from PD patients (e.g., Q456X nonsense or V170G missense mutants) also exhibit more fragmented mitochondrial networks (Grunewald et  al. 2009). However, the effects of PINK1 and Parkin on mitochondrial dynamics have been contentious. In Drosophila the PINK1/Parkin pathway appears to promote mitochondrial fission or inhibits mitochondrial fusion (Deng et al. 2008; Poole et al. 2008; Yang et al. 2008). PINK1 and Parkin not only affect mitochondrial morphology but also regulate mitochondrial degradation by mitophagy (Geisler et al. 2010; Matsuda et al. 2010; Narendra et al. 2010; Ziviani et al. 2010). After recruitment by PINK1, mitochondrial Parkin ubiqutinates VDAC1 and Mfn, which may promote autophagic clearance of damaged mitochondria. Although further studies will be needed to understand the significance of these findings for the pathogenesis of PD, it is clear that PD-associated genes exert and influence on mitochondrial dynamics.

8.8 Conclusions Mitochondria are highly dynamic organelles, which continuously divide and fuse. Over the past decade, a significant number of studies regarding mitochondrial dynamics have accumulated. Fusion requires the activity of large GTPase proteins such as Mfn1/2 and Opa1. In contrast, another large GTPase, Drp1 represents a principal component of the mitochondrial fission machinery. The discovery of genetic mutations in Drp1, OPA1 and Mfn that underlie several human neurodegenerative diseases strongly implies that the balance between mitochondrial fission and fusion is linked to the pathogenesis of these diseases. Additionally, oligomerized Ab can induce mitochondrial fragmentation through regulation of Drp1. Moreover, dysfunction in Parkinson’s disease-related genes also induces abnormal mitochondrial morphology. Thus, abnormal mitochondrial dynamics may represent a final common pathway leading to neuronal dysfunction. This hypothesis suggests that the mitochondrial fission and fusion machinery may become a therapeutic target for neurodegenerative diseases, including AD and PD. Acknowledgments  This study was supported in part by NIH grants P01 HD29587, P01 ES01673, P30 NS057096, R01 EY05477, and R01 EY09024 (to S.A.L.). Additional support was provided by a grant from the Korea Healthcare technology R&D Project, Ministry for Health, Welfare and Family Affairs, Republic of Korea (A090013 and A092042) (to D.-H.C.).

References Abou-Sleiman, Pm, Muqit, Mm, Wood, Nw, 2006, Expanding insights of mitochondrial dysfunction in Parkinson’s disease, Nat Rev Neurosci, 7: 207–19. Alavi, Mv, Bette, S, Schimpf, S, Schuettauf, F, Schraermeyer, U, Wehrl, Hf, Ruttiger, L, Beck, Sc, Tonagel, F, Pichler, Bj, Knipper, M, Peters, T, Laufs, J, Wissinger, B, 2007, A splice site mutation

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

229

in the murine Opa1 gene features pathology of autosomal dominant optic atrophy, Brain, 130: 1029–42. Alexander, C, Votruba, M, Pesch, Ue, Thiselton, Dl, Mayer, S, Moore, A, Rodriguez, M, Kellner, U, Leo-Kottler, B, Auburger, G, Bhattacharya, Ss, Wissinger, B, 2000, OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28, Nat Genet, 26: 211–5. Alirol, E, James, D, Huber, D, Marchetto, A, Vergani, L, Martinou, Jc, Scorrano, L, 2006, The mitochondrial fission protein hFis1 requires the endoplasmic reticulum gateway to induce apoptosis, Mol Biol Cell, 17: 4593–605. Autret, A, Martin, Sj, 2009, Emerging role for members of the Bcl-2 family in mitochondrial morphogenesis, Mol Cell, 36: 355–63. Bach, D, Pich, S, Soriano, Fx, Vega, N, Baumgartner, B, Oriola, J, Daugaard, Jr, Lloberas, J, Camps, M, Zierath, Jr, Rabasa-Lhoret, R, Wallberg-Henriksson, H, Laville, M, Palacin, M, Vidal, H, Rivera, F, Brand, M, Zorzano, A, 2003, Mitofusin-2 determines mitochondrial network architecture and mitochondrial metabolism. A novel regulatory mechanism altered in obesity, J Biol Chem, 278: 17190–7. Barsoum, Mj, Yuan, H, Gerencser, Aa, Liot, G, Kushnareva, Y, Graber, S, Kovacs, I, Lee, Wd, Waggoner, J, Cui, J, White, Ad, Bossy, B, Martinou, Jc, Youle, Rj, Lipton, Sa, Ellisman, Mh, Perkins, Ga, Bossy-Wetzel, E, 2006, Nitric oxide-induced mitochondrial fission is regulated by dynamin-related GTPases in neurons, EMBO J, 25: 3900–11. Berman, Sb, Chen, Yb, Qi, B, Mccaffery, Jm, Rucker, Eb, 3rd, Goebbels, S, Nave, Ka, Arnold, Ba, Jonas, Ea, Pineda, Fj, Hardwick, Jm, 2009, Bcl-x L increases mitochondrial fission, fusion, and biomass in neurons, J Cell Biol, 184: 707–19. Blass, Jp, 2000, The mitochondrial spiral. An adequate cause of dementia in the Alzheimer’s syndrome, Ann NY Acad Sci, 924: 170–83. Bossy-Wetzel, E, Barsoum, Mj, Godzik, A, Schwarzenbacher, R, Lipton, Sa, 2003, Mitochondrial fission in apoptosis, neurodegeneration and aging, Curr Opin Cell Biol, 15: 706–16. Braschi, E, Zunino, R, Mcbride, Hm, 2009, MAPL is a new mitochondrial SUMO E3 ligase that regulates mitochondrial fission, EMBO Rep, 10: 748–54. Brunden, Kr, Trojanowski, Jq, Lee, Vm, 2009, Advances in tau-focused drug discovery for Alzheimer’s disease and related tauopathies, Nat Rev Drug Discov, 8: 783–93. Cartoni, R, Martinou, Jc, 2009, Role of mitofusin 2 mutations in the physiopathology of CharcotMarie-Tooth disease type 2A, Exp Neurol, 218: 268–73. Casley, Cs, Canevari, L, Land, Jm, Clark, Jb, Sharpe, Ma, 2002, Beta-amyloid inhibits integrated mitochondrial respiration and key enzyme activities, J Neurochem, 80: 91–100. Cassidy-Stone, A, Chipuk, Je, Ingerman, E, Song, C, Yoo, C, Kuwana, T, Kurth, Mj, Shaw, Jt, Hinshaw, Je, Green, Dr, Nunnari, J, 2008, Chemical inhibition of the mitochondrial division dynamin reveals its role in Bax/Bak-dependent mitochondrial outer membrane permeabilization, Dev Cell, 14: 193–204. Cereghetti, Gm, Stangherlin, A, Martins De Brito, O, Chang, Cr, Blackstone, C, Bernardi, P, Scorrano, L, 2008, Dephosphorylation by calcineurin regulates translocation of Drp1 to mitochondria, Proc Natl Acad Sci USA, 105: 15803–8. Chan, Dc, 2006a, Mitochondria: dynamic organelles in disease, aging, and development, Cell, 125: 1241–52. Chan, Dc, 2006b, Mitochondrial fusion and fission in mammals, Annu Rev Cell Dev Biol, 22: 79–99. Chang, Cr, Blackstone, C, 2007, Cyclic AMP-dependent protein kinase phosphorylation of Drp1 regulates its GTPase activity and mitochondrial morphology, J Biol Chem, 282: 21583–7. lethal de  novo mutation in the middle domain of the dynamin-related GTPase Drp1 impairs higher order assembly and mitochondrial division, J Biol Chem, 285: 32494–503. Chen, H, Chomyn, A, Chan, Dc, 2005, Disruption of fusion results in mitochondrial heterogeneity and dysfunction, J Biol Chem, 280: 26185–92. Chen, H, Detmer, Sa, Ewald, Aj, Griffin, Ee, Fraser, Se, Chan, Dc, 2003, Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development, J Cell Biol, 160: 189–200.

230

T. Nakamura et al.

Chen, H, Mccaffery, Jm, Chan, Dc, 2007, Mitochondrial fusion protects against neurodegeneration in the cerebellum, Cell, 130: 548–62. Chen, Jx, Yan, Sd, 2007, Amyloid-beta-induced mitochondrial dysfunction, J Alzheimers Dis, 12: 177–84. Chipuk, Je, Green, Dr, 2008, How do BCL-2 proteins induce mitochondrial outer membrane permeabilization?, Trends Cell Biol, 18: 157–64. Cho, Dh, Nakamura, T, Fang, J, Cieplak, P, Godzik, A, Gu, Z, Lipton, Sa, 2009, S-Nitrosylation of Drp1 mediates beta-amyloid-related mitochondrial fission and neuronal injury, Science, 324: 102–5. Cipolat, S, Martins De Brito, O, Dal Zilio, B, Scorrano, L, 2004, OPA1 requires mitofusin 1 to promote mitochondrial fusion, Proc Natl Acad Sci USA, 101: 15927–32. Cribbs, Jt, Strack, S, 2007, Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein kinase and calcineurin regulates mitochondrial fission and cell death, EMBO Rep, 8: 939–44. Crouch, Pj, Blake, R, Duce, Ja, Ciccotosto, Gd, Li, Qx, Barnham, Kj, Curtain, Cc, Cherny, Ra, Cappai, R, Dyrks, T, Masters, Cl, Trounce, Ia, 2005, Copper-dependent inhibition of human cytochrome c oxidase by a dimeric conformer of amyloid-beta1-42, J Neurosci, 25: 672–9. Dagda, Rk, Cherra, Sj, 3rd, Kulich, Sm, Tandon, A, Park, D, Chu, Ct, 2009, Loss of PINK1 function promotes mitophagy through effects on oxidative stress and mitochondrial fission, J Biol Chem, 284: 13843–55. Damke, H, Binns, Dd, Ueda, H, Schmid, Sl, Baba, T, 2001, Dynamin GTPase domain mutants block endocytic vesicle formation at morphologically distinct stages, Mol Biol Cell, 12: 2578–89. Davies, Vj, Hollins, Aj, Piechota, Mj, Yip, W, Davies, Jr, White, Ke, Nicols, Pp, Boulton, Me, Votruba, M, 2007, Opa1 deficiency in a mouse model of autosomal dominant optic atrophy impairs mitochondrial morphology, optic nerve structure and visual function, Hum Mol Genet, 16: 1307–18. Delettre, C, Lenaers, G, Griffoin, Jm, Gigarel, N, Lorenzo, C, Belenguer, P, Pelloquin, L, Grosgeorge, J, Turc-Carel, C, Perret, E, Astarie-Dequeker, C, Lasquellec, L, Arnaud, B, Ducommun, B, Kaplan, J, Hamel, Cp, 2000, Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy, Nat Genet, 26: 207–10. Delivani, P, Adrain, C, Taylor, Rc, Duriez, Pj, Martin, Sj, 2006, Role for CED-9 and Egl-1 as regulators of mitochondrial fission and fusion dynamics, Mol Cell, 21: 761–73. Deng, H, Dodson, Mw, Huang, H, Guo, M, 2008, The Parkinson’s disease genes pink1 and parkin promote mitochondrial fission and/or inhibit fusion in Drosophila, Proc Natl Acad Sci USA, 105: 14503–8. Exner, N, Treske, B, Paquet, D, Holmstrom, K, Schiesling, C, Gispert, S, Carballo-Carbajal, I, Berg, D, Hoepken, Hh, Gasser, T, Kruger, R, Winklhofer, Kf, Vogel, F, Reichert, As, Auburger, G, Kahle, Pj, Schmid, B, Haass, C, 2007, Loss-of-function of human PINK1 results in mitochondrial pathology and can be rescued by parkin, J Neurosci, 27: 12413–8. Figueroa-Romero, C, Iniguez-Lluhi, Ja, Stadler, J, Chang, Cr, Arnoult, D, Keller, Pj, Hong, Y, Blackstone, C, Feldman, El, 2009, SUMOylation of the mitochondrial fission protein Drp1 occurs at multiple nonconsensus sites within the B domain and is linked to its activity cycle, FASEB J, 23: 3917–27. Frank, S, Gaume, B, Bergmann-Leitner, Es, Leitner, Ww, Robert, Eg, Catez, F, Smith, Cl, Youle, Rj, 2001, The role of dynamin-related protein 1, a mediator of mitochondrial fission, in apoptosis, Dev Cell, 1: 515–25. Frezza, C, Cipolat, S, Martins De Brito, O, Micaroni, M, Beznoussenko, Gv, Rudka, T, Bartoli, D, Polishuck, Rs, Danial, Nn, De Strooper, B, Scorrano, L, 2006, OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion, Cell, 126: 177–89. Geisler, S, Holmstrom, Km, Skujat, D, Fiesel, Fc, Rothfuss, Oc, Kahle, Pj, Springer, W, 2010, PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1, Nat Cell Biol, 12: 119–31. Gomez-Lazaro, M, Bonekamp, Na, Galindo, Mf, Jordan, J, Schrader, M, 2008, 6-Hydroxydopamine (6-OHDA) induces Drp1-dependent mitochondrial fragmentation in SH-SY5Y cells, Free Radic Biol Med, 44: 1960–9.

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

231

Griparic, L, Van Der Wel, Nn, Orozco, Ij, Peters, Pj, Van Der Bliek, Am, 2004, Loss of the intermembrane space protein Mgm1/OPA1 induces swelling and localized constrictions along the lengths of mitochondria, J Biol Chem, 279: 18792–8. Grunewald, A, Gegg, Me, Taanman, Jw, King, Rh, Kock, N, Klein, C, Schapira, Ah, 2009, Differential effects of PINK1 nonsense and missense mutations on mitochondrial function and morphology, Exp Neurol, 219: 266–73. Haass, C, Selkoe, Dj, 2007, Soluble protein oligomers in neurodegeneration: lessons from the Alzheimer’s amyloid beta-peptide, Nat Rev Mol Cell Biol, 8: 101–12. Hajek, P, Chomyn, A, Attardi, G, 2007, Identification of a novel mitochondrial complex containing mitofusin 2 and stomatin-like protein 2, J Biol Chem, 282: 5670–81. Hales, Kg, Fuller, Mt, 1997, Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase, Cell, 90: 121–9. Han, Xj, Lu, Yf, Li, Sa, Kaitsuka, T, Sato, Y, Tomizawa, K, Nairn, Ac, Takei, K, Matsui, H, Matsushita, M, 2008, CaM kinase I alpha-induced phosphorylation of Drp1 regulates mitochondrial morphology, J Cell Biol, 182: 573–85. Hansson Petersen, Ca, Alikhani, N, Behbahani, H, Wiehager, B, Pavlov, Pf, Alafuzoff, I, Leinonen, V, Ito, A, Winblad, B, Glaser, E, Ankarcrona, M, 2008, The amyloid beta-peptide is imported into mitochondria via the TOM import machinery and localized to mitochondrial cristae, Proc Natl Acad Sci USA, 105: 13145–50. Harder, Z, Zunino, R, Mcbride, H, 2004, Sumo1 conjugates mitochondrial substrates and participates in mitochondrial fission, Curr Biol, 14: 340–5. Hoppins, S, Nunnari, J, 2009, The molecular mechanism of mitochondrial fusion, Biochim Biophys Acta, 1793: 20–6. Howng, Sl, Sy, Wd, Cheng, Ts, Lieu, As, Wang, C, Tzou, Ws, Cho, Cl, Hong, Yr, 2004, Genomic organization, alternative splicing, and promoter analysis of human dynamin-like protein gene, Biochem Biophys Res Commun, 314: 766–72. Ingerman, E, Perkins, Em, Marino, M, Mears, Ja, Mccaffery, Jm, Hinshaw, Je, Nunnari, J, 2005, Dnm1 forms spirals that are structurally tailored to fit mitochondria, J Cell Biol, 170: 1021–7. Ishihara, N, Fujita, Y, Oka, T, Mihara, K, 2006, Regulation of mitochondrial morphology through proteolytic cleavage of OPA1, EMBO J, 25: 2966–77. Ishihara, N, Nomura, M, Jofuku, A, Kato, H, Suzuki, So, Masuda, K, Otera, H, Nakanishi, Y, Nonaka, I, Goto, Y, Taguchi, N, Morinaga, H, Maeda, M, Takayanagi, R, Yokota, S, Mihara, K, 2009, Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice, Nat Cell Biol, 11: 958–66. James, Di, Parone, Pa, Mattenberger, Y, Martinou, Jc, 2003, hFis1, a novel component of the mammalian mitochondrial fission machinery, J Biol Chem, 278: 36373–9. Jin, J, Li, Gj, Davis, J, Zhu, D, Wang, Y, Pan, C, Zhang, J, 2007, Identification of novel proteins associated with both alpha-synuclein and DJ-1, Mol Cell Proteomics, 6: 845–59. Karbowski, M, Lee, Yj, Gaume, B, Jeong, Sy, Frank, S, Nechushtan, A, Santel, A, Fuller, M, Smith, Cl, Youle, Rj, 2002, Spatial and temporal association of Bax with mitochondrial fission sites, Drp1, and Mfn2 during apoptosis, J Cell Biol, 159: 931–8. Karbowski, M, Neutzner, A, Youle, Rj, 2007, The mitochondrial E3 ubiquitin ligase MARCH5 is required for Drp1 dependent mitochondrial division, J Cell Biol, 178: 71–84. Karbowski, M, Norris, Kl, Cleland, Mm, Jeong, Sy, Youle, Rj, 2006, Role of Bax and Bak in mitochondrial morphogenesis, Nature, 443: 658–62. Knott, Ab, Perkins, G, Schwarzenbacher, R, Bossy-Wetzel, E, 2008, Mitochondrial fragmentation in neurodegeneration, Nat Rev Neurosci, 9: 505–18. Koch, A, Thiemann, M, Grabenbauer, M, Yoon, Y, Mcniven, Ma, Schrader, M, 2003, Dynaminlike protein 1 is involved in peroxisomal fission, J Biol Chem, 278: 8597–605. Koshiba, T, Detmer, Sa, Kaiser, Jt, Chen, H, Mccaffery, Jm, Chan, Dc, 2004, Structural basis of mitochondrial tethering by mitofusin complexes, Science, 305: 858–62. Lackner, Ll, Horner, Js, Nunnari, J, 2009, Mechanistic analysis of a dynamin effector, Science, 325: 874–7.

232

T. Nakamura et al.

Laferla, Fm, Green, Kn, Oddo, S, 2007, Intracellular amyloid-beta in Alzheimer’s disease, Nat Rev Neurosci, 8: 499–509. Lawson, Vh, Graham, Bv, Flanigan, Km, 2005, Clinical and electrophysiologic features of CMT2A with mutations in the mitofusin 2 gene, Neurology, 65: 197–204. Lee, Yj, Jeong, Sy, Karbowski, M, Smith, Cl, Youle, Rj, 2004, Roles of the mammalian mitochondrial fission and fusion mediators Fis1, Drp1, and Opa1 in apoptosis, Mol Biol Cell, 15: 5001–11. Li, H, Chen, Y, Jones, Af, Sanger, Rh, Collis, Lp, Flannery, R, Mcnay, Ec, Yu, T, Schwarzenbacher, R, Bossy, B, Bossy-Wetzel, E, Bennett, Mv, Pypaert, M, Hickman, Ja, Smith, Pj, Hardwick, Jm, Jonas, Ea, 2008, Bcl-xL induces Drp1-dependent synapse formation in cultured hippocampal neurons, Proc Natl Acad Sci USA, 105: 2169–74. Li, Z, Okamoto, K, Hayashi, Y, Sheng, M, 2004, The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses, Cell, 119: 873–87. Liesa, M, Palacin, M, Zorzano, A, 2009, Mitochondrial dynamics in mammalian health and disease, Physiol Rev, 89: 799–845. Liu, Qa, Shio, H, 2008, Mitochondrial morphogenesis, dendrite development, and synapse formation in cerebellum require both Bcl-w and the glutamate receptor delta2, PLoS Genet, 4: e1000097. Lustbader, Jw, Cirilli, M, Lin, C, Xu, Hw, Takuma, K, Wang, N, Caspersen, C, Chen, X, Pollak, S, Chaney, M, Trinchese, F, Liu, S, Gunn-Moore, F, Lue, Lf, Walker, Dg, Kuppusamy, P, Zewier, Zl, Arancio, O, Stern, D, Yan, Ss, Wu, H, 2004, ABAD directly links Abeta to mitochondrial toxicity in Alzheimer’s disease, Science, 304: 448–52. Lutz, Ak, Exner, N, Fett, Me, Schlehe, Js, Kloos, K, Laemmermann, K, Brunner, B, Kurz-Drexler, A, Vogel, F, Reichert, As, Bouman, L, Vogt-Weisenhorn, D, Wurst, W, Tatzelt, J, Haass, C, Winklhofer, Kf, 2009, Loss of parkin or PINK1 function increases DRP1-dependent mitochondrial fragmentation, J Biol Chem, 284: 22938–51. Matsuda, N, Sato, S, Shiba, K, Okatsu, K, Saisho, K, Gautier, Ca, Sou, Ys, Saiki, S, Kawajiri, S, Sato, F, Kimura, M, Komatsu, M, Hattori, N, Tanaka, K, 2010, PINK1 stabilized by mitochondrial depolarization recruits Parkin to damaged mitochondria and activates latent Parkin for mitophagy, J Cell Biol, 189: 211–21. Meuer, K, Suppanz, Ie, Lingor, P, Planchamp, V, Goricke, B, Fichtner, L, Braus, Gh, Dietz, Gp, Jakobs, S, Bahr, M, Weishaupt, Jh, 2007, Cyclin-dependent kinase 5 is an upstream regulator of mitochondrial fission during neuronal apoptosis, Cell Death Differ, 14: 651–61. Montessuit, S, Somasekharan, Sp, Terrones, O, Lucken-Ardjomande, S, Herzig, S, Schwarzenbacher, R, Manstein, Dj, Bossy-Wetzel, E, Basanez, G, Meda, P, Martinou, Jc, 2010, Membrane remodeling induced by the dynamin-related protein Drp1 stimulates Bax oligomerization, Cell, 142: 889–901. Nakamura, N, Kimura, Y, Tokuda, M, Honda, S, Hirose, S, 2006, MARCH-V is a novel mitofusin 2- and Drp1-binding protein able to change mitochondrial morphology, EMBO Rep, 7: 1019–22. Narendra, Dp, Jin, Sm, Tanaka, A, Suen, Df, Gautier, Ca, Shen, J, Cookson, Mr, Youle, Rj, 2010, PINK1 is selectively stabilized on impaired mitochondria to activate Parkin, PLoS Biol, 8: e1000298. Neuspiel, M, Zunino, R, Gangaraju, S, Rippstein, P, Mcbride, H, 2005, Activated mitofusin 2 signals mitochondrial fusion, interferes with Bax activation, and reduces susceptibility to radical induced depolarization, J Biol Chem, 280: 25060–70. Olichon, A, Baricault, L, Gas, N, Guillou, E, Valette, A, Belenguer, P, Lenaers, G, 2003, Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis, J Biol Chem, 278: 7743–6. Olichon, A, Guillou, E, Delettre, C, Landes, T, Arnaune-Pelloquin, L, Emorine, Lj, Mils, V, Daloyau, M, Hamel, C, Amati-Bonneau, P, Bonneau, D, Reynier, P, Lenaers, G, Belenguer, P, 2006, Mitochondrial dynamics and disease, OPA1, Biochim Biophys Acta, 1763: 500–9. Olichon, A, Landes, T, Arnaune-Pelloquin, L, Emorine, Lj, Mils, V, Guichet, A, Delettre, C, Hamel, C, Amati-Bonneau, P, Bonneau, D, Reynier, P, Lenaers, G, Belenguer, P, 2007, Effects

8  Role of the Mitochondrial Fission Protein Drp1 in Synaptic

233

of OPA1 mutations on mitochondrial morphology and apoptosis: relevance to ADOA pathogenesis, J Cell Physiol, 211: 423–30. Ow, Yp, Green, Dr, Hao, Z, Mak, Tw, 2008, Cytochrome c: functions beyond respiration, Nat Rev Mol Cell Biol, 9: 532–42. Parone, Pa, James, Di, Da Cruz, S, Mattenberger, Y, Donze, O, Barja, F, Martinou, Jc, 2006, Inhibiting the mitochondrial fission machinery does not prevent Bax/Bak-dependent apoptosis, Mol Cell Biol, 26: 7397–408. Pitts, Kr, Mcniven, Ma, Yoon, Y, 2004, Mitochondria-specific function of the dynamin family protein DLP1 is mediated by its C-terminal domains, J Biol Chem, 279: 50286–94. Poole, Ac, Thomas, Re, Andrews, La, Mcbride, Hm, Whitworth, Aj, Pallanck, Lj, 2008, The PINK1/ Parkin pathway regulates mitochondrial morphology, Proc Natl Acad Sci USA, 105: 1638–43. Praefcke, Gj, Mcmahon, Ht, 2004, The dynamin superfamily: universal membrane tubulation and fission molecules?, Nat Rev Mol Cell Biol, 5: 133–47. Ramachandran, R, Surka, M, Chappie, Js, Fowler, Dm, Foss, Tr, Song, Bd, Schmid, Sl, 2007, The dynamin middle domain is critical for tetramerization and higher-order self-assembly, EMBO J, 26: 559–66. Rojo, M, Legros, F, Chateau, D, Lombes, A, 2002, Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo, J Cell Sci, 115: 1663–74. Santel, A, 2006, Get the balance right: mitofusins roles in health and disease, Biochim Biophys Acta, 1763: 490–9. Santel, A, Frank, S, Gaume, B, Herrler, M, Youle, Rj, Fuller, Mt, 2003, Mitofusin-1 protein is a generally expressed mediator of mitochondrial fusion in mammalian cells, J Cell Sci, 116: 2763–74. Santel, A, Fuller, Mt, 2001, Control of mitochondrial morphology by a human mitofusin, J Cell Sci, 114: 867–74. Schapira, Ah, Bezard, E, Brotchie, J, Calon, F, Collingridge, Gl, Ferger, B, Hengerer, B, Hirsch, E, Jenner, P, Le Novere, N, Obeso, Ja, Schwarzschild, Ma, Spampinato, U, Davidai, G, 2006, Novel pharmacological targets for the treatment of Parkinson’s disease, Nat Rev Drug Discov, 5: 845–54. Scheff, Sw, Price, Da, 1998, Synaptic density in the inner molecular layer of the hippocampal dentate gyrus in Alzheimer disease, J Neuropathol Exp Neurol, 57: 1146–53. Selkoe, Dj, 2000, Toward a comprehensive theory for Alzheimer’s disease. Hypothesis: Alzheimer’s disease is caused by the cerebral accumulation and cytotoxicity of amyloid betaprotein, Ann NY Acad Sci, 924: 17–25. Sheridan, C, Delivani, P, Cullen, Sp, Martin, Sj, 2008, Bax- or Bak-induced mitochondrial fission can be uncoupled from cytochrome C release, Mol Cell, 31: 570–85. Shin, Hw, Shinotsuka, C, Torii, S, Murakami, K, Nakayama, K, 1997, Identification and subcellular localization of a novel mammalian dynamin-related protein homologous to yeast Vps1p and Dnm1p, J Biochem, 122: 525–30. Smirnova, E, Griparic, L, Shurland, Dl, Van Der Bliek, Am, 2001, Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells, Mol Biol Cell, 12: 2245–56. Smirnova, E, Shurland, Dl, Ryazantsev, Sn, Van Der Bliek, Am, 1998, A human dynamin-related protein controls the distribution of mitochondria, J Cell Biol, 143: 351–8. Su, B, Wang, X, Zheng, L, Perry, G, Smith, Ma, Zhu, X, 2010, Abnormal mitochondrial dynamics and neurodegenerative diseases, Biochim Biophys Acta, 1802: 135–42. Suen, Df, Norris, Kl, Youle, Rj, 2008, Mitochondrial dynamics and apoptosis, Genes Dev, 22: 1577–90. Sugioka, R, Shimizu, S, Tsujimoto, Y, 2004, Fzo1, a protein involved in mitochondrial fusion, inhibits apoptosis, J Biol Chem, 279: 52726–34. Taguchi, N, Ishihara, N, Jofuku, A, Oka, T, Mihara, K, 2007, Mitotic phosphorylation of dynaminrelated GTPase Drp1 participates in mitochondrial fission, J Biol Chem, 282: 11521–9. Verstreken, P, Ly, Cv, Venken, Kj, Koh, Tw, Zhou, Y, Bellen, Hj, 2005, Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions, Neuron, 47: 365–78.

234

T. Nakamura et al.

Wakabayashi, J, Zhang, Z, Wakabayashi, N, Tamura, Y, Fukaya, M, Kensler, Tw, Iijima, M, Sesaki, H, 2009, The dynamin-related GTPase Drp1 is required for embryonic and brain development in mice, J Cell Biol, 186: 805–16. Wang, X, Su, B, Lee, Hg, Li, X, Perry, G, Smith, Ma, Zhu, X, 2009, Impaired balance of mitochondrial fission and fusion in Alzheimer’s disease, J Neurosci, 29: 9090–103. Wasiak, S, Zunino, R, Mcbride, Hm, 2007, Bax/Bak promote sumoylation of DRP1 and its stable association with mitochondria during apoptotic cell death, J Cell Biol, 177: 439–50. Waterham, Hr, Koster, J, Van Roermund, Cw, Mooyer, Pa, Wanders, Rj, Leonard, Jv, 2007, A lethal defect of mitochondrial and peroxisomal fission, N Engl J Med, 356: 1736–41. Wyss-Coray, T, 2006, Inflammation in Alzheimer disease: driving force, bystander or beneficial response?, Nat Med, 12: 1005–15. Yang, Y, Ouyang, Y, Yang, L, Beal, Mf, Mcquibban, A, Vogel, H, Lu, B, 2008, Pink1 regulates mitochondrial dynamics through interaction with the fission/fusion machinery, Proc Natl Acad Sci USA, 105: 7070–5. Yankner, Ba, Lu, T, 2009, Amyloid beta-protein toxicity and the pathogenesis of Alzheimer disease, J Biol Chem, 284: 4755–9. Yonashiro, R, Ishido, S, Kyo, S, Fukuda, T, Goto, E, Matsuki, Y, Ohmura-Hoshino, M, Sada, K, Hotta, H, Yamamura, H, Inatome, R, Yanagi, S, 2006, A novel mitochondrial ubiquitin ligase plays a critical role in mitochondrial dynamics, EMBO J, 25: 3618–26. Yoon, Y, Krueger, Ew, Oswald, Bj, Mcniven, Ma, 2003, The mitochondrial protein hFis1 regulates mitochondrial fission in mammalian cells through an interaction with the dynamin-like protein DLP1, Mol Cell Biol, 23: 5409–20. Yoon, Y, Pitts, Kr, Mcniven, Ma, 2001, Mammalian dynamin-like protein DLP1 tubulates membranes, Mol Biol Cell, 12: 2894–905. Zamzami, N, Kroemer, G, 2001, The mitochondrion in apoptosis: how Pandora’s box opens, Nat Rev Mol Cell Biol, 2: 67–71. Zhan, Ss, Beyreuther, K, Schmitt, Hp, 1993, Quantitative assessment of the synaptophysin immuno-reactivity of the cortical neuropil in various neurodegenerative disorders with dementia, Dementia, 4: 66–74. Zhang, P, Hinshaw, Je, 2001, Three-dimensional reconstruction of dynamin in the constricted state, Nat Cell Biol, 3: 922–6. Zhu, Pp, Patterson, A, Stadler, J, Seeburg, Dp, Sheng, M, Blackstone, C, 2004, Intra- and intermolecular domain interactions of the C-terminal GTPase effector domain of the multimeric dynamin-like GTPase Drp1, J Biol Chem, 279: 35967–74. Ziviani, E, Tao, Rn, Whitworth, Aj, 2010, Drosophila parkin requires PINK1 for mitochondrial translocation and ubiquitinates mitofusin, Proc Natl Acad Sci USA, 107: 5018–23. Zorzano, A, Liesa, M, Palacin, M, 2009, Mitochondrial dynamics as a bridge between mitochondrial dysfunction and insulin resistance, Arch Physiol Biochem, 115: 1–12. Zuchner, S, Mersiyanova, Iv, Muglia, M, Bissar-Tadmouri, N, Rochelle, J, Dadali, El, Zappia, M, Nelis, E, Patitucci, A, Senderek, J, Parman, Y, Evgrafov, O, Jonghe, Pd, Takahashi, Y, Tsuji, S, Pericak-Vance, Ma, Quattrone, A, Battaloglu, E, Polyakov, Av, Timmerman, V, Schroder, Jm, Vance, Jm, 2004, Mutations in the mitochondrial GTPase mitofusin 2 cause CharcotMarie-Tooth neuropathy type 2A, Nat Genet, 36: 449–51. Zunino, R, Schauss, A, Rippstein, P, Andrade-Navarro, M, Mcbride, Hm, 2007, The SUMO protease SENP5 is required to maintain mitochondrial morphology and function, J Cell Sci, 120: 1178–88.

Chapter 9

Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate Hongmin Wang, Mariusz Karbowski, and Mervyn J. Monteiro

Abstract  Huntington’s disease (HD) is a devastating neurodegenerative disorder whose age of onset can vary quite widely, with the most severe forms striking as early as the first decade of life. The disease is caused by an abnormal expansion of a polyglutamine tract in the huntingtin (htt) protein. The mechanism by which polyglutamine expansions in htt cause disease is still not understood, although many different mechanisms have been proposed. One theory is that HD is caused by an impairment of mitochondria function. Mitochondria are vital for cell survival. They are the main powerhouse for ATP generation and are the repository of many important molecules that govern whether a cell lives or dies. Studies have shown that mitochondria are not static structures, but are highly dynamic, undergoing constant cycles of fusion and fission. Accumulating evidence suggests that HD is associated with a disturbance of mitochondrial dynamics. Here we review the published data linking mitochondrial dysfunction in HD, concentrating on the role that defects in mitochondrial dynamics might play in the disease. Keywords  Mitochondrial dynamics • Mitochondrial fusion • Mitochondrial fission • Huntingtin • Polyglutamine expansion • Cell death H. Wang Division of Basic Biomedical Sciences, University of South Dakota, Vermillion, SD 57069, USA M. Karbowski Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD 21201, USA and Department of Molecular Biology and Biochemistry, University of Maryland School of Medicine, Baltimore, MD 21201, USA M.J. Monteiro () Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD 21201, USA and Department of Anatomy and Neurobiology, University of Maryland School of Medicine, Baltimore 21201, MD, USA e-mail: [email protected] B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1_9, © Springer Science+Business Media B.V. 2011

235

236

H. Wang et al.

Abbreviations AAA ATPases associated with diverse cellular activities ATP adenosine triphosphate Bcl-2 B-cell lymphoma 2 Bax Bcl2-associated X protein C. elegans Caenorhabditis elegans CAG cytosine adenine guanine Drp1 dynamin-related protein-1 EM electron microscope Fis1 mitochondrial fission-1 protein GFP green fluorescent protein GST glutathione S-transferase GTP guanosine triphosphate HEAT Huntingtin, elongation factor 3, protein phosphatase 2A, and the yeast PI3-kinase TOR1 HD Huntington’s disease Htt Huntingtin IMM inner mitochondrial membrane Mff mitochondrial fission factor Mfn1 and 2 Mitofusin 1 and 2 mito-PAGFP mitochondrial targeted photoactivatable GFP mRFP monomeric red fluorescent protein NMR nuclear magnetic resonance spectroscopy 3-NP 3-nitropropionic acid Opa1 optic atrophy 1 OMM outer mitochondrial membrane RING really interesting new gene

9.1 Huntington’s Disease: Molecular Genetics Huntington’s disease (HD) is a devastating neurodegenerative disorder that typically strikes between the ages of 30 and 40 years and whose symptoms include involuntary choreoathetotic movements of the limbs and head, severe behavioral and emotional disturbances, and a decline in cognition and memory. The disease is inherited in an autosomal dominant manner, and typically affects approximately 1 in 10,000 people, although extremely higher incidences of the disease are found in certain regions of the world, like near the Lake Maracaibo region in Venezuela (Wexler et al. 2004). The duration of the disease typically lasts about 15–20 years and is usually fatal. Currently, there is no effective treatment to alleviate or prevent HD. The mutation responsible for HD was identified in 1993 in a gene called huntingtin (htt), located on chromosome 4 p16.3 (1993). The mutation is caused by an expansion of a trinucleotide tract composed of reiterated CAG sequence that is

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

237

found in exon-1 of the htt gene (Bates 2005; Imarisio et  al. 2008). The htt open reading frame encodes a protein of 3,144 amino acids and the CAG tract occurs in-frame after the 17th codon following the initiator methionine. The length of the CAG tract varies in individuals with most unaffected individuals containing 15–26 CAG repeats (Myers 2004). The reason for the variation is not known, and neither is its function. Extensive genetic analysis has now shown that people carrying tracts of fewer than 35 CAG repeats will not develop HD, whereas those carrying tracts of between 35 and 39 repeats show incomplete penetrance, but alleles containing 40 or more repeats are fully penetrant (Rubinsztein et al. 1996; Myers 2004). It is now well established that there is an inverse correlation between the number of CAG repeats and the age of onset of disease, with very long repeats (>76) causing disease as early as 10 years of age (Walker 2007). Moreover, studies of HD patients have shown that carriers who possess shorter CAG tracts have greater variation when disease first manifests than those carrying longer tracts leading to the theory that genetic and/or environmental factors influence disease onset especially for carriers with shorter tracts, whereas the longer tracts because of their higher intrinsic toxicity may mask the influence of these factors (Wexler et al. 2004; Gusella and MacDonald 2009). The CAG repeats in htt are translated into a reiterated stretch of glutamine amino acids, commonly referred to as the polyglutamine tract (or polyQ). In fact, HD is one of nine neurological disorders that are associated with an expansion of polyglutamine tracts in otherwise unrelated proteins (Gusella and MacDonald 2006). The exact mechanism by which polyglutamine expansions in htt cause disease is still unclear, although numerous hypotheses have been advanced as to the potential mechanism(s) (for reviews see Landles and Bates 2004; Gatchel and Zoghbi 2005; Di Prospero and Fischbeck 2005; Walker 2007; Imarisio et al. 2008; Zuccato et al. 2010). A central issue regarding the mechanism is whether polyglutamine expansions cause a gain or loss of htt function. While, it is mostly agreed that the polyglutamine expansion causes a gain-of-toxic function (Landles and Bates 2004; Walker 2007), the loss of htt function cannot be discounted because conditional knockout of htt expression in mice leads to neurodegeneration (Dragatsis et al. 2000). Therefore, it is possible that HD pathogenesis could arise from both a gain and loss of htt function. This issue becomes especially important when considering therapeutic strategies for treating HD. For example, any method that eliminates both the mutant and wild type htt protein (e.g. RNAi efforts) could potentially pose problems unless of course only the mutant htt gene were specially targeted, or if expression of the normal htt gene can be restored.

9.2 Htt Protein: Normal and Pathological Function Full-length htt protein is composed of 3,144 amino acids (using 23 glutamines as the length of the polyQ tract). Despite this length, an examination of the protein sequence reveals few distinguishable structural motifs, providing little clues of its function. The only conspicuous motif is multiple HEAT (Huntingtin, elongation

238

H. Wang et al.

factor 3 (EF3), protein phosphatase 2A (PP2A), and the yeast PI3-kinase TOR1) repeats dispersed throughout the polypeptide. The HEAT repeat domain is a short ~47 amino acid sequence that is rich in alpha-helical content and which is thought to function in mediating protein-protein interaction. Proteins containing multiple HEAT repeats have been shown to function in intracellular transport. Aside from these repeats, the other most distinguishing feature about the protein is that it contains the polyglutamine tract, which starts at amino acid 18 in the polypeptide and is then immediately followed by a proline-rich sequence between amino acids 41 and 78 of the protein. The function of the proline-rich sequence is not known, although studies suggest it might prevent aggregation of polyglutamine tract in htt protein, but only when the polyglutamine tract is below the mutant range (Qin et al. 2004). Support for this idea has been obtained by structural studies of the NH2terminal region of the htt protein (Kim et al. 2009). Full-length htt protein is a stable protein, with a half-life of over 20 h (Persichetti et al. 1996). The protein is ubiquitously expressed (Sharp et al. 1995) and localizes mainly to the cytoplasm, although a fraction of the protein is also found in the nucleus (Kegel et al. 2002). The shuttling of the protein between the cytoplasm and nucleus appears to be regulated by a nuclear export signal located close to its C-terminus (Xia et  al. 2003). In the nucleus, the protein associates with factors involved in regulating transcription and DNA repair, while in the cytoplasm the protein associates with several different structures including microtubules, the plasma membrane, endoplasmic reticulum (ER), the Golgi complex, endosomes, synaptic vesicles, autophagosomes, ubiquitin-proteasome system, lysosomes, and mitochondria (Imarisio et al. 2008). The widespread distribution of htt protein in different subcellular compartments provides little clarification of its function. Instead it was hoped that its function might be learnt from the proteins it interacts. However, these screens have yielded an even larger variety of molecules with diverse cellular functions, leaving unresolved what htt’s true function is (Li and Li 2004; Kaltenbach et al. 2007). A clear indication that htt is vital for survival is that molecular disruption of the htt gene in mouse is lethal and embryos homozygous for the deletion die by embryonic day 8.5 (Nasir et  al. 1995; Zeitlin et  al. 1995; Duyao et al. 1995). While the function of wild-type htt protein containing a polyglutamine tract in the nonpathogenic range remains enigmatic, the mutant protein containing an expanded polyglutamine tract displays some unique properties. Using antibodies to different regions of the htt protein it was found that the NH2-terminal fragment (~40 kDa) containing the polyglutamine tract aggregates and forms nuclear inclusions that are ubiquitin positive and only visible in patients afflicted with HD (DiFiglia et al. 1997). The abnormal aggregation of the NH2-terminal htt fragment is thought to arise by self-assembly of the fragments into amyloid-like fibrils, the speed of whose assembly is directly governed by the length of the polyglutamine tract (Scherzinger et al. 1997, 1999). Other studies have demonstrated that expression of the NH2-terminal fragment of htt, particularly the exon 1 fragment, containing the expanded polyglutamine tract, is sufficient to induce toxicity in a variety of

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

239

organisms including rodents, Caenorhabditis elegans, Danio rerio, Drosophila melanogaster and yeast (Mangiarini et al. 1996; Zuccato et al. 2010). The production of the NH2-terminal fragment is thought to arise by abnormal proteolytic cleavage of htt protein. Indeed, several different proteases including caplains and caspases have been implicated in this abnormal cleavage (Imarisio et  al. 2008; Zuccato et al. 2010). Interestingly, the proteolytic cleavage appears to be regulated by changes in post-translational modification of htt protein: phosphorylation, fatty acid modification (palmitoylation), ubiquitination and/or sumoylation have all been implicated in affecting the cleavage (Luo et al. 2005; Imarisio et al. 2008; Gu et al. 2009; Zuccato et al. 2010). The general consensus that emerges from these studies is that htt cleavage is required for inducing toxicity, and not surprisingly, toxicity appears to be mainly confined to the NH2-terminal portion of htt protein, in the fragment containing the expanded polyglutamine tract. An important and unresolved issue in HD research regards the exact mechanism by which expanded polyglutamine tracts in htt induce pathogenicity. Several different mechanisms have been proposed, including a disruption in the normal function of critical factors (such as transcription factors and chaperones) because of their sequestration with mutant htt proteins in aggregates, or because the aggregates and/or mutant proteins block the normal transport of proteins and factors vital for cellular survival, or because the mutant proteins block and/or interfere with protein disposal through the proteasome and/or autophagy degradation pathways, or because the mutant proteins bind and interfere with the function of organelles such as the ER and mitochondria, to name a few (Imarisio et  al. 2008). Another unresolved issue in this regard is the identity of the culprit species involved in causing disease. Of particular debate is whether htt aggregates are toxic or whether toxicity is caused by some other species of this protein. While there appears to be a good correlation between the appearance of htt aggregates and HD progression recent studies in cell culture to examine this connection have cast doubts about this conclusion, and instead suggests that the aggregates (particularly large aggregates) might in fact be protective (Imarisio et  al. 2008; Arrasate et al. 2004; Miller et al. 2010). Regardless of the mechanism(s) involved in toxicity, it is clear that methods to rid cells of mutant htt protein aggregates are in general protective (Ravikumar et al. 2004; Wang et al. 2006; Yamamoto et al. 2006; Jeong et al. 2009; Gu et al. 2009). Another important issue in trying to relate mechanisms to pathogenicity is the identity of the cells and tissues that are affected in HD. While earlier studies suggested that the g-amino butyric acid-releasing neurons in the striatal regions of the brain that control movement may be specifically affected in HD, it is now becoming clear that pathogenesis is more widespread affecting many different parts of the body including the brain, muscle, as well as peripheral tissues (Gusella and MacDonald 2006; Henley et al. 2009; Kloppel et al. 2009; Moffitt et  al. 2009; Johnson and Davidson 2010). Therefore efforts to treat HD should also take into consideration that an ideal treatment should alleviate symptoms of all the affected tissues.

240

H. Wang et al.

9.3 Structural and Functional Defects of Mitochondria in Huntington’s Disease While several different mechanisms have been implicated in HD pathogenesis, the one that is receiving increasing attention, and the subject of this review, is the role that defects in mitochondria play in the pathogenesis. Here we highlight some of the evidence linking defects in mitochondrial function in HD, concentrating on disturbances in mitochondrial dynamics. Several excellent reviews have been published on related aspects of mitochondrial dysfunction in HD, which should also be considered as they most likely influence mitochondrial dynamics (Browne 2008; Yang et al. 2008a; Beal 2007; Zuccato et al. 2010). Some of the first visual indications that mitochondria in HD possess structural defects were uncovered by electron microscope (EM) examination of tissue from HD cases (Tellez-Nagel et al. 1974; Goebel et al. 1978). Tellez-Nagel and coauthors showed that mitochondria in patients with HD looked abnormal in that they contained very few cristae, which they noted were typically displaced onto one side (Tellez-Nagel et al. 1974). Cristae are distinguishable as invaginations of the inner mitochondrial membrane and it is there where many important complexes reside, including Complexes I-to-V that function in ATP generation by oxidative phosphorylation. Interestingly, Tellez-Nagel et al. also highlighted the unusual presence of dense granules in the nucleus, which could be the nuclear inclusions that are now known to be widespread in the disease. Similarly, EM studies conducted by Goebel et al. reported that mitochondria in neurons and glia in tissue sample of a 20 yearold patient with juvenile HD displayed gross abnormalities in both their size and structure (Goebel et al. 1978). These investigators too found that the mitochondria in the HD patient contained few cristae, but they also described abnormal accumulations of different shaped electron-dense inclusions within some mitochondria. Additional evidence that ultrastructural abnormalities in mitochondria are more widespread in the body of HD patients was provided by EM examination of lympoblasts from subjects that were either heterozygous or homogenous carriers of the HD mutation (Squitieri et al. 2006). In this study too, mitochondria from the HD patients were also found to contain gross derangements of their matrix and cristae. Inclusions were not observed, but the matrix was shown to be highly translucent, indicative of a disruption in mitochondria structure. The general conclusion from these early pathological studies of HD tissue suggested that HD is associated with major structural defects in mitochondrial organization. Because mitochondria are the main cellular site of ATP production, one prediction is that if the structural changes that are seen in mitochondria are an integral part of HD disease progression then defects in bioenergetics and/or mitochondrial respiration should manifest in HD patients. Several studies have examined energy metabolism in brains of presymptomatic and symptomatic HD patients as well as in mitochondria isolated from tissues of HD patients. These studies have shown that mitochondrial bioenergetics and energy production are indeed compromised in HD (Lin and Beal 2006; Beal 2007). Positron emission tomography (PET) studies

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

241

revealed a 15% reduction in glucose consumption in cortical and subcortical regions of the brain in HD patients (Kuwert et al. 1990; Martin et al. 1992), whereas nuclear magnetic resonance spectroscopy (NMR) revealed that lactate production is increased in occipital cortex and basal ganglia in HD (Jenkins et al. 1993). The increased lactate production is thought to arise from anaerobic glycolysis due to suppression of mitochondrial ATP production. Indeed, measurement of ATP levels in muscle tissue of symptomatic and presymptomatic patients have revealed that after exercise ATP production is reduced as much as 35–44% compared to that found in control patients (Lodi et al. 2000; Saft et al. 2005). The decrease in ATP production in muscle is an accord with the ubiquitous expression of the htt protein and the widespread pathology of the disease, including the muscle. Consistent with decreased mitochondrial ATP production, activity measurements of the different steps of mitochondria respiration have documented general decreases in the activities of respiratory complexes I, II, III and IV (Brennan et  al. 1985; Mann et  al. 1990; Gu et  al. 1996; Browne et  al. 1997; Arenas et  al. 1998; Browne 2008; Zuccato et al. 2010). However, several other studies have not found the aforementioned defects in mitochondrial respiration in HD leading some to speculate that they might arise because of technical issues related to the measurements or from artifacts produced due to increased cell loss in HD. Three other lines of investigation lend further support for the idea that HD pathogenesis might arise from defects in mitochondrial function. The first comes from indirect evidence showing that administration of the fungal mitochondrial toxin called 3-nitropropionic acid (3-NP) in rats and apes, induces loss of striatal spiny neurons and produce phenotypes that share many HD-like symptoms (Brouillet et  al. 1993, 1995). 3-NP is an irreversible inhibitor of mitochondrial respiratory complex II. The implication from these studies is that inhibition of mitochondrial respiration induces HD-like symptoms, but it does not necessarily prove that HD is caused by similar mechanisms. The second line of evidence is that mitochondria isolated from lymphoblasts of HD patients and from brain tissue of a transgenic mouse model of HD display defects in calcium handling (Panov et al. 2002). Specifically the authors found that mitochondria from the HD samples displayed lower mitochondrial membrane potential and that they depolarize at lower calcium levels than mitochondria from control subjects. In agreement with these finding others have also reported disturbances in calcium regulation in mitochondria (Choo et  al. 2004; Bezprozvanny and Hayden 2004; Tang et  al. 2005). These results suggest that mitochondria in HD are somehow compromised in their ability to properly regulate calcium homeostasis, although the precise mechanism of how this defect arises is not fully understood. The third line of evidence suggesting a defect in mitochondrial function may be involved in HD pathogenesis is the observation that mutant HD proteins are associated with a dramatic alteration in the dynamics of mitochondrial fusion and fission (Wang et al. 2009a; Kim et  al. 2010). In order to appreciate why an alteration in mitochondrial dynamics could be important in HD pathogenesis we have provided a brief description of the process, highlighting how disturbances in the process could affect neuronal survival.

242

H. Wang et al.

9.4 Mitochondria Dynamics of Fusion and Fission Mitochondria are highly dynamic organelles and they frequently change their shape and location in response to various intracellular and extracellular stimuli. Changes in mitochondrial shape are regulated by two opposite processes fusion and fission. In most cells the two processes occur simultaneously, the balance of which dictates the overall morphology of the mitochondrial network at any given time. In their extremes, fusion produces long tubules and fission small round vesicles. Studies have shown that although the organization of the mitochondrial network can differ markedly between different cell types, it is very strongly influenced by both the metabolic rate of the cell and by exposure to cellular stress. Furthermore, recent studies have indicated that defects in mitochondrial fusion, fission, mislocalization of mitochondria, or a disruption of the mitochondrial network can all profoundly affect cellular homeostasis, and which is now believed could play an important role in the development of different diseases. Mitochondrial fusion and fission are regulated by a number of proteins, which is summarized below. For more extensive coverage of the subject see the following reviews (Westermann 2008; Benard and Karbowski 2009; Chen and Chan 2009).

9.5 Mitochondrial Fusion Mitofusin 1 and 2 proteins (Mfn1 and 2) (Chen et  al. 2003) and optic atrophy 1 (Opa1) protein (Delettre et al. 2000) are known to regulate mitochondrial fusion. Mfn 1 and Mfn2 are two large closely related integral membrane GTPases that localize to the outer mitochondrial membrane (OMM) whereas Opa1 is a dynaminrelated protein located in the inner mitochondrial membrane (IMM). Both Mfn1 and Mfn2 appear to be required for the proper maintenance of mitochondrial fusion, because single knockouts of either Mfn1 or Mfn2 reduce the rate of mitochondrial fusion (Chen et al. 2003; Karbowski et al. 2004). However, studies suggest that the two proteins might function in different steps during mitochondrial fusion. These studies suggest Mfn1 might be specifically required for GTP hydrolysis-dependent tethering of mitochondria prior to fusion, whereas Mfn2, which is less efficient in this process, might act as a signaling GTPase to somehow stimulate assembly of the fusion complexes (Ishihara et  al. 2004; Neuspiel et  al. 2005). As discussed later, further indication of a functional difference between Mfn1 and Mfn2 is supported by differences in development and functional defects found upon ablation of the Mfn1 and Mfn2 genes. The precise function of Opa1 is not clear but studies suggest it functions together with Mfn1, probably by direct binding of the proteins, to regulate fusion (Guillery et al. 2008). Notably, as in case of Mfns, reduction of Opa1 expression leads to inhibition of mitochondrial fusion and increased mitochondrial fragmentation (Olichon et al. 2003). In addition to Mfn1, Mfn2 and Opa1, a number of accessory proteins have been reported to regulate mitochondrial fusion in mammalian cells, including mitofusin binding protein (MiB) (Eura et al. 2006), mitochondria associated phospholipase D (mito-PLD) (Choi et al. 2006), and stomatin-like protein 2 (Stoml2 also known as

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

243

SLP2) (Hajek et al. 2007). Furthermore, Bcl-xL, Bak and Bax, Bcl-2 family proteins involved in the regulation of apoptosis have also been implicated in the regulation of mitochondrial fusion. Accordingly in both, Bcl-xL−/− cortical neurons, as well as in a number of Bak−/−/Bax−/− cells, including mouse embryonic fibroblasts, HCT116 cells, as well as primary cortical neurons, mitochondrial size was significantly reduced, as compared to wild type cells (Karbowski et al. 2006; Berman et al. 2009; Cleland et al. 2010). In summary, although there is clear indication that the abovementioned proteins influence mitochondrial fusion, the molecular mechanisms by which they regulate the process still needs to be clarified.

9.6 Mitochondrial Fission The most critical protein known to regulate mitochondrial fission is dynaminrelated protein-1 (Drp1), a dynamin-like GTPase. Loss of Drp1 expression through either gene disruption, knockdown of its expression by RNA interference, or expression of dominant negative mutants of the protein (e.g. GTPase domain mutant Drp1K38A, or phosphomimetic mutant Drp1S637D) all lead to inhibition of mitochondrial fission and formation of abnormally elongated and interconnected mitochondria (Chang and Blackstone 2007a; Smirnova et al. 2001; Ishihara et al. 2009). Under normal growth conditions most Drp1 is localized in the cytoplasm, but to initiate mitochondrial fission Drp1 translocates to the OMM where it often accumulates in discreet foci that often correspond to the eventual sites of mitochondrial fission (Smirnova et  al. 2001). The recruitment of Drp1 to mitochondria appears to be mediated by interaction with Fis1, a OMM anchored protein, the interaction of which was first identified in yeast (James et  al. 2003; Yoon et  al. 2003). Evidence that Fis1 protein functions in fission comes from studies of the overexpression or inactivation of the protein in mammalian cells, yet its precise role in fission is not fully understood (James et al. 2003; Yoon et al. 2003). It is likely that a signal is involved in triggering binding of Drp1 to Fis1, or to some other protein located on the mitochondria. The support for this idea come from studies showing submitochondrial foci of Drp1 are stabilized by sumoylation (Harder et al. 2004) and destabilized by ubiquitination (Karbowski et al. 2007), suggesting that these two posttranslational modifications of Drp1 play opposing roles in the formation of mitochondrial fission protein complex. In addition to these modifications, phosphorylation of Drp1 by cAMP-dependent protein kinase, Ca2+/calmodulindependent protein kinase I alpha (CaMKIalpha), and cyclin-dependent kinase (Cdk1/cyclin B) have been shown to regulate mitochondrial fission (Cribbs and Strack 2007; Chang and Blackstone 2007b; Han et al. 2008; Taguchi et al. 2007). Several additional proteins have been found to regulate mitochondrial fission. These include Mff (for mitochondrial fission factor) (Gandre-Babbe and van der Bliek 2008) and MARCH5, a RING finger domain E3 ubiquitin ligase of the OMM (Karbowski et  al. 2007; Yonashiro et  al. 2006). Furthermore, like mitochondrial fusion, mitochondrial fission appears to be regulated by the Bcl-2 family of proteins. As discussed next, Bcl-w, an antiapoptotic mitochondria-localized protein regulates

244

H. Wang et al.

mitochondrial length in Purkinje cell processes (Liu and Shio 2008). Mitochondria in Bcl-w−/− mice often contain points where they become constricted, suggesting that this abnormal constriction in the mitochondria might be due to the inhibition or a delay in mitochondrial fission.

9.7 Disruption of Mitochondrial Fusion and Fission Balance Leads to Neurodegeneration A correct balance in the rates of mitochondrial fusion and fission in health was first illuminated by reports showing that mutations in Opa1 and Mfn2 fusion proteins, which result in defects in mitochondrial fusion, are associated with two different neurodegenerative diseases, dominant optic atrophy and Charcot-Marie-Tooth disease type 2A, respectively (Zuchner et  al. 2004; Alexander et  al. 2000). In addition, mutations of paraplegin, an m-AAA-metalloprotease that is required for mitochondrial processing of Opa1 are linked to another peripheral neuropathy, spastic paraplegia. Aberrations in the organization of the mitochondrial network are also seen in the so called “mutator mouse”, considered a genetic model of premature aging, that arises because of proof reading mutation in mitochondrial DNA polymerase-g (Trifunovic et  al. 2004). Fibroblasts isolated from the “mutator mouse” possess a fragmented mitochondrial network, as well as alterations in proteolytic processing of the fusion protein Opa1 (Duvezin-Caubet et al. 2006). Collectively, these data suggest that defects in the dynamics of mitochondrial fusion and fission can cause disease. In other chapters in this book, defects in mitochondrial dynamics are discussed in relation to other neurodegenerative diseases including Parkinson’s disease (Yang et  al. 2008b) and Alzheimer’s disease (Wang et al. 2009b). Collectively these findings suggest that proper balance of mitochondrial fusion and fission might be particularly important for maintenance of neuronal homeostasis and that disruption of this balance can lead to neurodegeneration.

9.8 Relationship of Mitochondrial Fusion and Fission in Neuronal and Synaptic Physiology On the other hand, genetic alteration of genes involved in mitochondrial fusion and fission has shed new insight into the role that the mitochondrial network plays in neuronal physiology. For example, genetic ablation of Mfn2 in mice leads to defective dendritic arborization, spine formation, and Purkinje cell loss that correlates not only with an aberrant distribution of mitochondria in the cells but diminished ATP generation due to perturbation in oxidative phosphorylation (Chen et al. 2007). The results suggest that mitochondrial ATP generation is somehow linked to proper dynamics of mitochondrial fusion, which is in turn important for dendritic stability and function. The importance of a proper balance in mitochondrial fusion and fission in dendritic physiology is further illustrated by studies of the overexpression

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

245

of the dominant-negative Drp1 (Drp1K38A) construct, which was found to lead to a depletion of mitochondria in dendrites and a 80% reduction in spine density (Li et al. 2004). Conversely, overexpression of wild-type Drp1 strongly increased both mitochondria and spine density in dendrites. These studies highlight the importance that mitochondrial fission and fusion play in the proper distribution of mitochondria in dendrites and in the formation and/or maintenance of synapses. Other studies conducted in Drosophila suggested that improper regulation of mitochondrial dynamics could ultimately lead to defects in synaptic transmission. Thus, a forward genetic screen for genes affecting neurotransmission in Drosophila, identified mutations in Drp1 that affected high frequency induced-stimulation of neurotransmission (Verstreken et  al. 2005). Interestingly, mitochondria were generally absent from synapses of neurons in the Drp1-mutant flies. The defect in neurotransmission was traced to aberrant formation or mobilization of the reserve pool of vesicles required for release during high-frequency stimulation. The study suggests that mitochondrial dynamics are involved in modulating the strength of the synaptic response. Consistent with this notion, synaptic function has been found to be compromised in neuronal cultures prepared from Drp1−/− mice (Ishihara et al. 2009). Apart from the core mitochondrial fusion and fission factors, Bcl-2 family members have also been implicated in regulating neuronal function by altering the organization and dynamics of the mitochondrial network. For example, Bcl-xL, an antiapoptotic protein belonging to the Bcl-2 family of proteins has been implicated in regulating synapse formation and mitochondrial dynamics in cortical neurons (Berman et  al. 2009). Paradoxically however, the authors found that Bcl-xL increased the rates of both fusion and fission, leaving uncertain exactly how mitochondrial dynamics are regulated by Bcl-xL. More recent evidence obtained by Cleland et al. (2010) suggests that both Bcl-xL and the proapoptotic Bax protein bind and interfere with the function of Mfn1 and Mfn2 proteins, providing a possible explanation of how mitochondrial dynamics is altered by Bcl-2 related proteins. Further support that Bcl-2 proteins regulate mitochondrial dynamics come from studies showing a significant increase in mitochondrial length in dendritic processes of Purkinje cells of mice disrupted of Bcl-w expression, a pro-survival Bcl-2-related protein (Liu and Shio 2008). Interestingly, the long mitochondria in the Bcl-w−/− mice contained discreet regions of constriction, suggesting that the abnormally long mitochondria accumulate due to a defect in fission. Taken together the results suggest that proper mitochondria dynamics plays an important role in neuronal function, particularly in dendrite development and synapse formation.

9.9 Mutant htt Impairs Mitochondrial Dynamics by Increasing Mitochondrial Fragmentation As discussed earlier, htt protein is known to bind to mitochondria as well as other organelles. Through deletion analysis, the mitochondrial-binding region was mapped to the NH2-terminal portion of htt protein (Panov et al. 2002; Choo et al. 2004;

246

H. Wang et al.

Orr et  al. 2008). Moreover, several different groups reported that polyglutamine expansions in htt protein affects axonal transport (Szebenyi et al. 2003; Gunawardena et al. 2003; Lee et al. 2004; Gunawardena and Goldstein 2005; Sinadinos et al. 2009) and impairs mitochondrial trafficking (Trushina et al. 2004; Chang et al. 2006; Orr et  al. 2008). We conducted experiments to examine how htt proteins containing different lengths of polyglutamine expansions affect mitochondrial structure and dynamics (Wang et al. 2009a). A summary of our findings follows. To examine if htt proteins containing expanded polyglutamine tracts affects mitochondrial structure and dynamics we used a photoactivatable green fluorescent protein (GFP) reporter that is targeted to mitochondria (mito-PAGFP) (Karbowski et  al. 2004) in combination with live cell imaging to examine if mitochondrial dynamics are altered in cells overexpressing htt proteins containing different lengths of polyglutamine expansions (Wang et al. 2009a). Previous studies showed that following focal activation of the mito-PAGFP, GFP fluorescence illuminates only the mitochondria in the region that is activated, but that the fluorescence dissipates over time throughout the whole mitochondrial network due to the normal dynamics of mitochondrial fusion and fission (Karbowski et  al. 2004). With this technique, we monitored changes in GFP fluorescence in HeLa cells cotransfected with mito-PAGFP and expression constructs encoding either monomeric red fluorescent protein (mRFP), or mRFP-tagged htt exon 1 fragment containing 28 polyglutamine repeats (mRFP-Htt28Q), or mRFP-Htt74Q. Through these experiments we found that cells transfected with mRFP and mRFP-Htt28Q remodeled their mitochondria rapidly over a 30  min-interval as evidenced by rapid movement of mitochondria and dissipation of GFP fluorescence over the entire mitochondrial network, whereas, mitochondria in cells transfected with mRFP-Htt74Q construct appeared motionless and retained fluorescence mainly in the photoactivated area over the same period of time. The experiments suggested that overexpression of htt proteins with extended polyglutamine tracts might inhibit mitochondrial fusion. Another possibility was that it increased mitochondrial fragmentation or fission. To examine this issue, we cultured HeLa cell lines stably expressing either GFPHtt28Q or GFP-Htt74Q in medium containing either 25 mM H2O2 or in low serum (1.25% fetal bovine serum instead of 10%), to induce oxidative stress (Atabay et al. 1996). Examination of mitochondria length by staining of the cells with antibodies against Tom20, an outer mitochondrial membrane protein, revealed that many more cells expressing GFP-Htt74Q had fragmented mitochondria than cells expressing GFP-Htt28Q, under both experimental conditions of oxidative stress. Interestingly, even under basal growth conditions (without any challenge of oxidative stress), the GFP-Htt74Q cell line contained a low but noticeable population of cells that had fragmented mitochondria compared with the GFP-Htt28Q cell line (Wang et  al. 2009a). To confirm these findings, we utilized electron microscopy to examine mitochondria structure in the two cell lines. By this analysis we found that the cell line expressing GFP-Htt74Q was more sensitized to oxidative stress as assessed by damage to their mitochondria than the GFP-Htt28Q cell line. Remarkably, almost 90% of cells from the line expressing GFP-Htt74Q had abnormal mitochondria with displaced and disorganized cristae and a less-electron dense matrix, which

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

247

Fig. 9.1  Cells expressing huntingtin proteins with expanded polyglutamine repeats are more sensitized to oxidative stress induced mitochondrial fragmentation. Immunofluorescence staining of HeLa cells transfected with a 549 amino acid N-terminal untagged htt protein fragment expression construct containing either 17 polyglutamines (Htt17Q) or 138 polyglutamines (Htt138Q). The cells were cultured in 1.25% fetal bovine serum, which is known to induce oxidative stress, and after 24 h incubation they were fixed and stained with antibodies against huntingtin and Tom20 to reveal huntingtin expression and mitochondria structure, respectively. Note the increase in mitochondrial fragmentation in the cell expressing Htt138Q compared to the cell expressing Htt17Q. Bar: 10 mm

appeared reminiscent of the mitochondrial abnormalities seen in specimens of HD afflicted patients (see above). By contrast, only 12% of cells in the GFP-Htt28Q cell line displayed this defect. We confirmed that the increase in fragmentation induced by overexpression of htt proteins containing expanded polyglutamine repeats was not unique to the GFP-Htt74Q protein that contained a short portion of the htt protein by showing that longer htt protein fragments (548 amino acids long) also induced increased mitochondrial fragmentation, but only if they contained an expanded polyglutamine tract known to be in the pathological range (Fig. 9.1). Because mitochondria are the main source of ATP production in cells we examined whether the structural alterations seen in cells expressing htt proteins containing expanded polyglutamine repeats is associated with an alteration in ATP levels in the cells. Accordingly we measured ATP levels in cells transfected with the GFP-Htt28Q and GFP-Htt74Q constructs and found the cells transfected with the htt construct expressing 74 polyglutamines had 30% lower ATP levels than the cells transfected with the construct containing the shorter polyglutamine tract (Wang et  al. 2009a). These results suggested that in addition to causing mitochondrial fragmentation, mutant htt proteins also trigger mitochondrial dysfunction as reflected by a reduction in ATP levels. Again, this mimics the ATP deficit that is seen in HD patients.

248

H. Wang et al.

9.10 Changing Mitochondrial Dynamics by Increasing Mitochondrial Fusion Reduces Cell Death Caused by Mutant htt Proteins We considered that the increase in mitochondria fragmentation induced by expanded polyglutamine proteins might predispose cells, or that it might be a precursor, to execution of cell death programs because mitochondria play a central role in the regulation of cell survival. They contain proteins such as Bcl-2 family of proteins and cytochrome c that are involved in the execution of the apoptosis. Mitochondria are also a major source of production of reactive oxygen species, which can induce oxidative damage to cells. As outlined earlier, in response to various metabolic demands and stimuli, cells frequently change the shape and structure of their mitochondria, of which fusion and fission play key role in the remodeling (Chan 2006). Of particular relevance to our findings is that mitochondrial fragmentation appears to be an early event in the apoptotic cascade as it precedes caspase activation, and furthermore, studies have shown that inhibition of mitochondrial fission reduces or delays cell death (Youle and Karbowski 2005). Also, studies have shown that mitochondrial fission is linked to oxidative stressinduced cell death and that cell death can be mitigated by forcibly increasing mitochondrial fusion through overexpression of Mfn proteins, or by expression of the dominant-negative Drp1K38A mutant (Neuspiel et al. 2005; Barsoum et al. 2006; Yuan et al. 2007; Jahani-Asl et al. 2007; Zhang et al. 2010; Ong et al. 2010). Accordingly, we designed experiments to test whether changing mitochondrial dynamics by increasing mitochondrial fusion would reduce cell death and the ATP deficit caused by overexpression of expanded polyglutamine proteins. Thus, we transfected HeLa cells with either GFP-Htt28Q or GFP-Htt74Q expression constructs together with or without a Drp1K38A expression construct, which promotes mitochondrial fusion, and quantified mitochondrial fragmentation, ATP levels, and cell death. Through these experiments we found that coexpression of Drp1K38A not only reduced GFP-Htt74Q-induced mitochondrial fragmentation, but it also restored ATP levels and reduced cell death to levels similar to those seen in untransfected cells or to cells transfected with the GFP-Htt28Q construct. A repetition of these experiments, but this time overexpressing Myc-tagged Mfn2 instead of Drp1K38A led to a similar outcome, showing that excessive mitochondrial fragmentation is the main driving force that reduces ATP levels and increases cell death in cell overexpressing expanded polyglutamine proteins, especially because Mfn2 and Drp1K38A function through different mechanisms to promote mitochondrial fusion. Interestingly, in these experiments we found that caspase-3 activation that is induced by overexpression of GFP-Htt74Q was suppressed by coexpression of Myc-Mfn2, suggesting that the protective effect exerted by Mfn2 might indeed suppresses execution of the apoptotic cell death pathway. Taken together, our results suggest that suppression of excessive mitochondrial fragmentation by manipulation of the core machinery involved in mitochondrial fusion and fission can protect cells against deficits in ATP levels and the induction of cell death propagated by htt proteins containing expanded polyglutamine proteins.

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

249

9.11 Reduction of Drp1 Levels in C. elegans by RNA Interference Rescues a Motility Defect Caused by Overexpression of Expanded Htt Proteins in Muscles of the Worm While the above experiments clearly illustrated that manipulation of mitochondrial dynamics can protect cells against the toxicity caused by mutant htt proteins it was important to know whether such manipulation would provide any benefit in an animal model of HD. To examine this possibility, we performed RNA interference (RNAi) studies in C. elegans, using two worm lines that stably express either GFP-Htt28Q or GFP-Htt74Q proteins in muscle cells (Wang et  al. 2006). In previous studies, we showed that expression of the GFP-tagged htt proteins causes a polyglutamine-lengthdependent motility defect in the worms, which can be quantified by differences in the frequency of body bends exhibited by the animals (Wang et al. 2006). To examine how changes in Drp1 expression affect worm motility, we fed the two worm lines with Escherichia coli transformed with plasmids to either genetically target or not target Drp1 for RNAi. Immunoblotting confirmed that Drp1 levels were reduced by over 70% in worms in which Drp1 was specifically targeted for RNAi. Remarkably, motility assays revealed a dramatic 200% improvement in worm motility in the GFP-Htt74Q worms that had been targeted for RNAi compared to animals in which Drp1 had not been targeted for RNAi. These data indicate that reduction of Drp1 levels in C. elegans by RNAi rescues the motility defect caused by expression of mutant htt proteins.

9.12 Potential Interference of Mitochondrial Dynamics by Expanded htt Proteins Although our studies suggest that htt proteins with expanded polyglutamine tracts increase mitochondrial fragmentation we do not know the mechanism by which it induces the increase. There are several likely possibilities. One possibility is that mutant htt proteins could bind and directly interfere with the normal function of proteins involved in regulating mitochondrial fusion and/or fission, either by inhibiting fusion or promoting fission. A second possibility is that mutant htt proteins might bind and/or block mitochondria movement (Orr et al. 2008; Trushina et al. 2004; Chang et al. 2006), indirectly perturbing the ability of mitochondria to fuse and divide. A third possibility is that mutant htt proteins could affect the expression of proteins required for mitochondrial function, particularly those involved in regulating the function and dynamics of mitochondria fission and fusion. A fourth possibility is that mutant htt proteins could affect the clearance of mitochondria (Martinez-Vicente et al. 2010) that could affect mitochondria dynamics. To examine whether mutant htt proteins might bind and interfere with proteins involved in regulating mitochondria dynamics we performed glutathione S-transferase (GST) pull-down assays, to examine whether mitofusin proteins expressed in HeLa cells bind

250

H. Wang et al.

differently to GST-tagged htt proteins containing different lengths of polyglutamine tracts. Interestingly, we found that cell lysates prepared from HeLa cells transfected with a Myc-Mfn2 expression construct bound preferentially with GST-Htt74Q fusion protein, rather than with GST-Htt28Q or GST proteins (Wang et al. 2009a). These results hint at the possibility that mutant htt containing expanded polyQ stretch may alter mitochondrial fragmentation/fission by binding and interfering with mitofusin protein function. However, the alternate possibility that mutant htt proteins enhance mitochondrial fission cannot be discounted because of the recent findings by Nakamura et  al. (Nakamura et  al. 2010) who recently showed that excessive nitric oxide build-up, which can lead to cell damage and death, is associated with S-Nitrosylation modification of Drp1, and that the modification enhances Drp1 activity in promoting excessive mitochondrial fission/fragmentation (Nakamura et al. 2010). It remains unknown whether mutant htt proteins increase mitochondrial fragmentation by a similar mechanism.

9.13 Evidence That HD Progression Is Associated with Changes in Mitochondria Size and in the Expression of Genes Involved in Regulating Mitochondrial Dynamics Strong evidence indicating that changes in mitochondrial dynamics is associated with HD pathogenesis comes from an elegant study in which Kim et al. quantified changes in mitochondria size and gene expression in spiny striatal neurons of HD patients with moderate and severe cases of the disease and compared them to those of age-matched controls without any known neurological disorder (Kim et  al. 2010). Their analysis revealed a significant reduction in the number of mitochondria during HD progression, and interestingly, they found that both large (0.4 mm) and medium size mitochondria (0.32 mm) are preferentially lost during early stages of HD. Furthermore, they found that Drp1 protein levels increase progressively with HD severity, while Mfn1 protein levels initially increase and then rapidly declined as disease progresses. Taken together the results are consistent with the idea that HD is associated with a change in mitochondrial dynamics in a direction that tips the balance toward an increase in mitochondrial fission. The initial increase in Mfn1 levels that is seen during early stages of disease could represent a futile effort of cells to restore mitochondrial fusion by increasing expression of genes involved in mitochondrial fusion. Further studies are needed to establish the mechanism and connection between changes in mitochondrial dynamics and HD.

9.14 Outlook and Therapeutic Efforts to Treat HD Hopefully, what should become clear from considering all of the information present above is of the strong connection between perturbation of mitochondrial function and dynamics and development of HD. However, many questions still remain to be

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

251

answered. For example, we do not know the exact mechanism by which wild type and mutant htt proteins affect mitochondrial dynamics and function? Are the defects caused by loss or a gain in the wild type and mutant htt protein functions? If so, what specific interactions, or loss of interaction, lead to the disruption in mitochondrial function? Do the mitochondrial abnormalities that are seen in HD arise from a direct affect on mitochondria or a indirect consequence of perturbation of other signaling pathways? How does the loss of mitochondria function and dynamics cause disease? Based on the current progress in HD research it should not be long before answers to many of these questions are known and rationale therapies are designed that could lead to an effective treatment to cure and/or prevent HD. Acknowledgments  This work was funded by grants from the National Institutes of Health (AG016839 and GM066287) to MJM.

References (1993) A novel gene containing a trinucleotide repeat that is expanded and unstable on Huntington’s disease chromosomes. The Huntington’s Disease Collaborative Research Group. Cell, 72, 971–83. Alexander, C., Votruba, M., Pesch, U. E., Thiselton, D. L., Mayer, S., Moore, A., Rodriguez, M., Kellner, U., Leo-Kottler, B., Auburger, G., Bhattacharya, S. S. & Wissinger, B. (2000) OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat Genet, 26, 211–5. Arenas, J., Campos, Y., Ribacoba, R., Martin, M. A., Rubio, J. C., Ablanedo, P. & Cabello, A. (1998) Complex I defect in muscle from patients with Huntington’s disease. Ann Neurol, 43, 397–400. Arrasate, M., Mitra, S., Schweitzer, E. S., Segal, M. R. & Finkbeiner, S. (2004) Inclusion body formation reduces levels of mutant huntingtin and the risk of neuronal death. Nature, 431, 805–10. Atabay, C., Cagnoli, C. M., Kharlamov, E., Ikonomovic, M. D. & Manev, H. (1996) Removal of serum from primary cultures of cerebellar granule neurons induces oxidative stress and DNA fragmentation: protection with antioxidants and glutamate receptor antagonists. J Neurosci Res, 43, 465–75. Barsoum, M. J., Yuan, H., Gerencser, A. A., Liot, G., Kushnareva, Y., Graber, S., Kovacs, I., Lee, W. D., Waggoner, J., Cui, J., White, A. D., Bossy, B., Martinou, J. C., Youle, R. J., Lipton, S. A., Ellisman, M. H., Perkins, G. A. & Bossy-Wetzel, E. (2006) Nitric oxide-induced mitochondrial fission is regulated by dynamin-related GTPases in neurons. Embo J, 25, 3900–11. Bates, G. P. (2005) History of genetic disease: the molecular genetics of Huntington disease a history. Nat Rev Genet, 6, 766–73. Beal, M. F. (2007) Mitochondria and neurodegeneration. Novartis Found Symp, 287, 183–92; discussion 192–6. Benard, G. & Karbowski, M. (2009) Mitochondrial fusion and division: Regulation and role in cell viability. Semin Cell Dev Biol, 20, 365–74. Berman, S. B., Chen, Y. B., QI, B., Mccaffery, J. M., Rucker, E. B., 3 rd, Goebbels, S., Nave, K. A., Arnold, B. A., Jonas, E. A., Pineda, F. J. & Hardwick, J. M. (2009) Bcl-x L increases mitochondrial fission, fusion, and biomass in neurons. J Cell Biol, 184, 707–19. Bezprozvanny, I. & Hayden, M. R. (2004) Deranged neuronal calcium signaling and Huntington disease. Biochem Biophys Res Commun, 322, 1310–7. Brennan, W. A., Jr., Bird, E. D. & Aprille, J. R. (1985) Regional mitochondrial respiratory activity in Huntington’s disease brain. J Neurochem, 44, 1948–50.

252

H. Wang et al.

Brouillet, E., Hantraye, P., Ferrante, R. J., Dolan, R., Leroy-Willig, A., Kowall, N. W. & Beal, M. F. (1995) Chronic mitochondrial energy impairment produces selective striatal degeneration and abnormal choreiform movements in primates. Proc Natl Acad Sci U S A, 92, 7105–9. Brouillet, E., Jenkins, B. G., Hyman, B. T., Ferrante, R. J., Kowall, N. W., Srivastava, R., Roy, D. S., Rosen, B. R. & Beal, M. F. (1993) Age-dependent vulnerability of the striatum to the mitochondrial toxin 3-nitropropionic acid. J Neurochem, 60, 356–9. Browne, S. E. (2008) Mitochondria and Huntington’s disease pathogenesis: insight from genetic and chemical models. Ann N Y Acad Sci, 1147, 358–82. Browne, S. E., Bowling, A. C., Macgarvey, U., Baik, M. J., Berger, S. C., Muqit, M. M., Bird, E. D. & Beal, M. F. (1997) Oxidative damage and metabolic dysfunction in Huntington’s disease: selective vulnerability of the basal ganglia. Ann Neurol, 41, 646–53. Chan, D. C. (2006) Mitochondria: dynamic organelles in disease, aging, and development. Cell, 125, 1241–52. Chang, C. R. & Blackstone, C. (2007a) Cyclic AMP-dependent protein kinase phosphorylation of Drp1 regulates its GTPase activity and mitochondrial morphology. J Biol Chem, 282, 21583–7. Chang, C. R. & Blackstone, C. (2007b) Drp1 phosphorylation and mitochondrial regulation. EMBO Rep, 8, 1088–9; author reply 1089–90. Chang, D. T., Rintoul, G. L., Pandipati, S. & Reynolds, I. J. (2006) Mutant huntingtin aggregates impair mitochondrial movement and trafficking in cortical neurons. Neurobiol Dis, 22, 388–400. Chen, H. & Chan, D. C. (2009) Mitochondrial dynamics--fusion, fission, movement, and mitophagy--in neurodegenerative diseases. Hum Mol Genet, 18, R169-76. Chen, H., Detmer, S. A., Ewald, A. J., Griffin, E. E., Fraser, S. E. & Chan, D. C. (2003) Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J Cell Biol, 160, 189–200. Chen, H., Mccaffery, J. M. & Chan, D. C. (2007) Mitochondrial fusion protects against neurodegeneration in the cerebellum. Cell, 130, 548–62. Choi, S. Y., Huang, P., Jenkins, G. M., Chan, D. C., Schiller, J. & Frohman, M. A. (2006) A common lipid links Mfn-mediated mitochondrial fusion and SNARE-regulated exocytosis. Nat Cell Biol, 8, 1255–62. Choo, Y. S., Johnson, G. V., Macdonald, M., Detloff, P. J. & Lesort, M. (2004) Mutant huntingtin directly increases susceptibility of mitochondria to the calcium-induced permeability transition and cytochrome c release. Hum Mol Genet, 13, 1407–20. Cleland, M. M., Norris, K. L., Karbowski, M., Wang, C., Suen, D. F., Jiao, S., George, N. M., Luo, X., Li, Z. & Youle, R. J. (2010) Bcl-2 family interaction with the mitochondrial morphogenesis machinery. Cell Death Differ. Cribbs, J. T. & Strack, S. (2007) Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein kinase and calcineurin regulates mitochondrial fission and cell death. EMBO Rep, 8, 939–44. Delettre, C., Lenaers, G., Griffoin, J. M., Gigarel, N., Lorenzo, C., Belenguer, P., Pelloquin, L., Grosgeorge, J., Turc-Carel, C., Perret, E., Astarie-Dequeker, C., Lasquellec, L., Arnaud, B., Ducommun, B., Kaplan, J. & Hamel, C. P. (2000) Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat Genet, 26, 207–10. Di Prospero, N. A. & Fischbeck, K. H. (2005) Therapeutics development for triplet repeat expansion diseases. Nat Rev Genet, 6, 756–65. Difiglia, M., Sapp, E., Chase, K. O., Davies, S. W., Bates, G. P., Vonsattel, J. P. & Aronin, N. (1997) Aggregation of huntingtin in neuronal intranuclear inclusions and dystrophic neurites in brain. Science, 277, 1990–3. Dragatsis, I., Levine, M. S. & Zeitlin, S. (2000) Inactivation of Hdh in the brain and testis results in progressive neurodegeneration and sterility in mice. Nat Genet, 26, 300–6. Duvezin-Caubet, S., Jagasia, R., Wagener, J., Hofmann, S., Trifunovic, A., Hansson, A., Chomyn, A., Bauer, M. F., Attardi, G., Larsson, N. G., Neupert, W. & Reichert, A. S. (2006) Proteolytic processing of OPA1 links mitochondrial dysfunction to alterations in mitochondrial morphology. J Biol Chem, 281, 37972–9.

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

253

Duyao, M. P., Auerbach, A. B., Ryan, A., Persichetti, F., Barnes, G. T., Mcneil, S. M., Ge, P., Vonsattel, J. P., Gusella, J. F., Joyner, A. L. & Et al. (1995) Inactivation of the mouse Huntington’s disease gene homolog Hdh. Science, 269, 407–10. Eura, Y., Ishihara, N., Oka, T. & Mihara, K. (2006) Identification of a novel protein that regulates mitochondrial fusion by modulating mitofusin (Mfn) protein function. J Cell Sci, 119, 4913–25. Gandre-Babbe, S. & Van Der Bliek, A. M. (2008) The novel tail-anchored membrane protein Mff controls mitochondrial and peroxisomal fission in mammalian cells. Mol Biol Cell, 19, 2402–12. Gatchel, J. R. & Zoghbi, H. Y. (2005) Diseases of unstable repeat expansion: mechanisms and common principles. Nat Rev Genet, 6, 743–55. Goebel, H. H., Heipertz, R., Scholz, W., Iqbal, K. & Tellez-Nagel, I. (1978) Juvenile Huntington chorea: clinical, ultrastructural, and biochemical studies. Neurology, 28, 23–31. Gu, M., Gash, M. T., Mann, V. M., Javoy-Agid, F., Cooper, J. M. & Schapira, A. H. (1996) Mitochondrial defect in Huntington’s disease caudate nucleus. Ann Neurol, 39, 385–9. Gu, X., Greiner, E. R., Mishra, R., Kodali, R., Osmand, A., Finkbeiner, S., Steffan, J. S., Thompson, L. M., Wetzel, R. & Yang, X. W. (2009) Serines 13 and 16 are critical determinants of full-length human mutant huntingtin induced disease pathogenesis in HD mice. Neuron, 64, 828–40. Guillery, O., Malka, F., Landes, T., Guillou, E., Blackstone, C., Lombes, A., Belenguer, P., Arnoult, D. & Rojo, M. (2008) Metalloprotease-mediated OPA1 processing is modulated by the mitochondrial membrane potential. Biol Cell, 100, 315–25. Gunawardena, S. & Goldstein, L. S. (2005) Polyglutamine diseases and transport problems: deadly traffic jams on neuronal highways. Arch Neurol, 62, 46–51. Gunawardena, S., Her, L. S., Brusch, R. G., Laymon, R. A., Niesman, I. R., Gordesky-Gold, B., Sintasath, L., Bonini, N. M. & Goldstein, L. S. (2003) Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila. Neuron, 40, 25–40. Gusella, J. F. & Macdonald, M. E. (2006) Huntington’s disease: seeing the pathogenic process through a genetic lens. Trends Biochem Sci, 31, 533–40. Gusella, J. F. & Macdonald, M. E. (2009) Huntington’s disease: the case for genetic modifiers. Genome Med, 1, 80. Hajek, P., Chomyn, A. & Attardi, G. (2007) Identification of a novel mitochondrial complex containing mitofusin 2 and stomatin-like protein 2. J Biol Chem, 282, 5670–81. Han, X. J., Lu, Y. F., Li, S. A., Kaitsuka, T., Sato, Y., Tomizawa, K., Nairn, A. C., Takei, K., Matsui, H. & Matsushita, M. (2008) CaM kinase I alpha-induced phosphorylation of Drp1 regulates mitochondrial morphology. J Cell Biol, 182, 573–85. Harder, Z., Zunino, R. & Mcbride, H. (2004) Sumo1 conjugates mitochondrial substrates and participates in mitochondrial fission. Curr Biol, 14, 340–5. Henley, S. M., Wild, E. J., Hobbs, N. Z., Frost, C., Macmanus, D. G., Barker, R. A., Fox, N. C. & Tabrizi, S. J. (2009) Whole-brain atrophy as a measure of progression in premanifest and early Huntington’s disease. Mov Disord, 24, 932–6. Imarisio, S., Carmichael, J., Korolchuk, V., Chen, C. W., Saiki, S., Rose, C., Krishna, G., Davies, J. E., Ttofi, E., Underwood, B. R. & Rubinsztein, D. C. (2008) Huntington’s disease: from pathology and genetics to potential therapies. Biochem J, 412, 191–209. Ishihara, N., Eura, Y. & Mihara, K. (2004) Mitofusin 1 and 2 play distinct roles in mitochondrial fusion reactions via GTPase activity. J Cell Sci, 117, 6535–46. Ishihara, N., Nomura, M., Jofuku, A., Kato, H., Suzuki, S. O., Masuda, K., Otera, H., Nakanishi, Y., Nonaka, I., Goto, Y., Taguchi, N., Morinaga, H., Maeda, M., Takayanagi, R., Yokota, S. & Mihara, K. (2009) Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nat Cell Biol, 11, 958–66. Jahani-Asl, A., Cheung, E. C., Neuspiel, M., Maclaurin, J. G., Fortin, A., Park, D. S., Mcbride, H. M. & Slack, R. S. (2007) Mitofusin 2 protects cerebellar granule neurons against injuryinduced cell death. J Biol Chem, 282, 23788–98. James, D. I., Parone, P. A., Mattenberger, Y. & Martinou, J. C. (2003) hFis1, a novel component of the mammalian mitochondrial fission machinery. J Biol Chem, 278, 36373–9.

254

H. Wang et al.

Jenkins, B. G., Koroshetz, W. J., Beal, M. F. & Rosen, B. R. (1993) Evidence for impairment of energy metabolism in vivo in Huntington’s disease using localized 1 H NMR spectroscopy. Neurology, 43, 2689–95. Jeong, H., Then, F., Melia, T. J., Jr., Mazzulli, J. R., Cui, L., Savas, J. N., Voisine, C., Paganetti, P., Tanese, N., Hart, A. C., Yamamoto, A. & Krainc, D. (2009) Acetylation targets mutant huntingtin to autophagosomes for degradation. Cell, 137, 60–72. Johnson, C. D. & Davidson, B. L. (2010) Huntington’s disease: progress toward effective diseasemodifying treatments and a cure. Hum Mol Genet, 19, R98-R102. Kaltenbach, L. S., Romero, E., Becklin, R. R., Chettier, R., Bell, R., Phansalkar, A., Strand, A., Torcassi, C., Savage, J., Hurlburt, A., Cha, G. H., Ukani, L., Chepanoske, C. L., Zhen, Y., Sahasrabudhe, S., Olson, J., Kurschner, C., Ellerby, L. M., Peltier, J. M., Botas, J. & Hughes, R. E. (2007) Huntingtin interacting proteins are genetic modifiers of neurodegeneration. PLoS Genet, 3, e82. Karbowski, M., Arnoult, D., Chen, H., Chan, D. C., Smith, C. L. & Youle, R. J. (2004) Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J Cell Biol, 164, 493–9. Karbowski, M., Neutzner, A. & Youle, R. J. (2007) The mitochondrial E3 ubiquitin ligase MARCH5 is required for Drp1 dependent mitochondrial division. J Cell Biol, 178, 71–84. Karbowski, M., Norris, K. L., Cleland, M. M., Jeong, S. Y. & Youle, R. J. (2006) Role of Bax and Bak in mitochondrial morphogenesis. Nature, 443, 658–62. Kegel, K. B., Meloni, A. R., Yi, Y., Kim, Y. J., Doyle, E., Cuiffo, B. G., Sapp, E., Wang, Y., Qin, Z. H., Chen, J. D., Nevins, J. R., Aronin, N. & Difiglia, M. (2002) Huntingtin is present in the nucleus, interacts with the transcriptional corepressor C-terminal binding protein, and represses transcription. J Biol Chem, 277, 7466–76. Kim, J., Moody, J. P., Edgerly, C. K., Bordiuk, O. L., Cormier, K., Smith, K., Beal, M. F. & Ferrante, R. J. (2010) Mitochondrial loss, dysfunction and altered dynamics in Huntington’s disease. Hum Mol Genet. Kim, M. W., Chelliah, Y., Kim, S. W., Otwinowski, Z. & Bezprozvanny, I. (2009) Secondary structure of Huntingtin amino-terminal region. Structure, 17, 1205–12. Kloppel, S., Henley, S. M., Hobbs, N. Z., Wolf, R. C., Kassubek, J., Tabrizi, S. J. & Frackowiak, R. S. (2009) Magnetic resonance imaging of Huntington’s disease: preparing for clinical trials. Neuroscience, 164, 205–19. Kuwert, T., Lange, H. W., Langen, K. J., Herzog, H., Aulich, A. & Feinendegen, L. E. (1990) Cortical and subcortical glucose consumption measured by PET in patients with Huntington’s disease. Brain, 113 (Pt 5), 1405–23. Landles, C. & Bates, G. P. (2004) Huntingtin and the molecular pathogenesis of Huntington’s disease. Fourth in molecular medicine review series. EMBO Rep, 5, 958–63. Lee, W. C., Yoshihara, M. & Littleton, J. T. (2004) Cytoplasmic aggregates trap polyglutaminecontaining proteins and block axonal transport in a Drosophila model of Huntington’s disease. Proc Natl Acad Sci U S A, 101, 3224–9. Li, S. H. & Li, X. J. (2004) Huntingtin-protein interactions and the pathogenesis of Huntington’s disease. Trends Genet, 20, 146–54. Li, Z., Okamoto, K., Hayashi, Y. & Sheng, M. (2004) The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses. Cell, 119, 873–87. Lin, M. T. & Beal, M. F. (2006) Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature, 443, 787–95. Liu, Q. A. & Shio, H. (2008) Mitochondrial morphogenesis, dendrite development, and synapse formation in cerebellum require both Bcl-w and the glutamate receptor delta2. PLoS Genet, 4, e1000097. Lodi, R., Schapira, A. H., Manners, D., Styles, P., Wood, N. W., Taylor, D. J. & Warner, T. T. (2000) Abnormal in  vivo skeletal muscle energy metabolism in Huntington’s disease and dentatorubropallidoluysian atrophy. Ann Neurol, 48, 72–6. Luo, S., Vacher, C., Davies, J. E. & Rubinsztein, D. C. (2005) Cdk5 phosphorylation of huntingtin reduces its cleavage by caspases: implications for mutant huntingtin toxicity. J Cell Biol, 169, 647–56.

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

255

Mangiarini, L., Sathasivam, K., Seller, M., Cozens, B., Harper, A., Hetherington, C., Lawton, M., Trottier, Y., Lehrach, H., Davies, S. W. & Bates, G. P. (1996) Exon 1 of the HD gene with an expanded CAG repeat is sufficient to cause a progressive neurological phenotype in transgenic mice. Cell, 87, 493–506. Mann, V. M., Cooper, J. M., Javoy-Agid, F., Agid, Y., Jenner, P. & Schapira, A. H. (1990) Mitochondrial function and parental sex effect in Huntington’s disease. Lancet, 336, 749. Martin, W. R., Clark, C., Ammann, W., Stoessl, A. J., Shtybel, W. & Hayden, M. R. (1992) Cortical glucose metabolism in Huntington’s disease. Neurology, 42, 223–9. Martinez-Vicente, M., Talloczy, Z., Wong, E., Tang, G., Koga, H., Kaushik, S., De Vries, R., Arias, E., Harris, S., Sulzer, D. & Cuervo, A. M. (2010) Cargo recognition failure is responsible for inefficient autophagy in Huntington’s disease. Nat Neurosci, 13, 567–76. Miller, J., Arrasate, M., Shaby, B. A., Mitra, S., Masliah, E. & Finkbeiner, S. (2010) Quantitative relationships between huntingtin levels, polyglutamine length, inclusion body formation, and neuronal death provide novel insight into huntington’s disease molecular pathogenesis. J Neurosci, 30, 10541–50. Moffitt, H., Mcphail, G. D., Woodman, B., Hobbs, C. & Bates, G. P. (2009) Formation of polyglutamine inclusions in a wide range of non-CNS tissues in the HdhQ150 knock-in mouse model of Huntington’s disease. PLoS One, 4, e8025. Myers, R. H. (2004) Huntington’s disease genetics. NeuroRx, 1, 255–62. Nakamura, T., Cieplak, P., Cho, D. H., Godzik, A. & Lipton, S. A. (2010) S-nitrosylation of Drp1 links excessive mitochondrial fission to neuronal injury in neurodegeneration. Mitochondrion, 10, 573–8. Nasir, J., Floresco, S. B., O’kusky, J. R., Diewert, V. M., Richman, J. M., Zeisler, J., Borowski, A., Marth, J. D., Phillips, A. G. & Hayden, M. R. (1995) Targeted disruption of the Huntington’s disease gene results in embryonic lethality and behavioral and morphological changes in heterozygotes. Cell, 81, 811–23. Neuspiel, M., Zunino, R., Gangaraju, S., Rippstein, P. & Mcbride, H. (2005) Activated mitofusin 2 signals mitochondrial fusion, interferes with Bax activation, and reduces susceptibility to radical induced depolarization. J Biol Chem, 280, 25060–70. Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P. & Lenaers, G. (2003) Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J Biol Chem, 278, 7743–6. Ong, S. B., Subrayan, S., Lim, S. Y., Yellon, D. M., Davidson, S. M. & Hausenloy, D. J. (2010) Inhibiting mitochondrial fission protects the heart against ischemia/reperfusion injury. Circulation, 121, 2012–22. Orr, A. L., Li, S., Wang, C. E., Li, H., Wang, J., Rong, J., Xu, X., Mastroberardino, P. G., Greenamyre, J. T. & Li, X. J. (2008) N-terminal mutant huntingtin associates with mitochondria and impairs mitochondrial trafficking. J Neurosci, 28, 2783–92. Panov, A. V., Gutekunst, C. A., Leavitt, B. R., Hayden, M. R., Burke, J. R., Strittmatter, W. J. & Greenamyre, J. T. (2002) Early mitochondrial calcium defects in Huntington’s disease are a direct effect of polyglutamines. Nat Neurosci, 5, 731–6. Persichetti, F., Carlee, L., Faber, P. W., Mcneil, S. M., Ambrose, C. M., Srinidhi, J., Anderson, M., Barnes, G. T., Gusella, J. F. & Macdonald, M. E. (1996) Differential expression of normal and mutant Huntington’s disease gene alleles. Neurobiol Dis, 3, 183–90. Qin, Z. H., Wang, Y., Sapp, E., Cuiffo, B., Wanker, E., Hayden, M. R., Kegel, K. B., Aronin, N. & Difiglia, M. (2004) Huntingtin bodies sequester vesicle-associated proteins by a polyproline-dependent interaction. J Neurosci, 24, 269–81. Ravikumar, B., Vacher, C., Berger, Z., Davies, J. E., Luo, S., Oroz, L. G., Scaravilli, F., Easton, D. F., Duden, R., O’kane, C. J. & Rubinsztein, D. C. (2004) Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat Genet, 36, 585–95. Rubinsztein, D. C., Leggo, J., Coles, R., Almqvist, E., Biancalana, V., Cassiman, J. J., Chotai, K., Connarty, M., Crauford, D., Curtis, A., Curtis, D., Davidson, M. J., Differ, A. M., Dode, C., Dodge, A., Frontali, M., Ranen, N. G., Stine, O. C., Sherr, M., Abbott, M. H., Franz, M. L., Graham, C. A., Harper, P. S., Hedreen, J. C., Hayden, M. R. & Et al. (1996) Phenotypic

256

H. Wang et al.

characterization of individuals with 30–40 CAG repeats in the Huntington disease (HD) gene reveals HD cases with 36 repeats and apparently normal elderly individuals with 36–39 repeats. Am J Hum Genet, 59, 16–22. Saft, C., Zange, J., Andrich, J., Muller, K., Lindenberg, K., Landwehrmeyer, B., Vorgerd, M., Kraus, P. H., Przuntek, H. & Schols, L. (2005) Mitochondrial impairment in patients and asymptomatic mutation carriers of Huntington’s disease. Mov Disord, 20, 674–9. Scherzinger, E., Lurz, R., Turmaine, M., Mangiarini, L., Hollenbach, B., Hasenbank, R., Bates, G. P., Davies, S. W., Lehrach, H. & Wanker, E. E. (1997) Huntingtin-encoded polyglutamine expansions form amyloid-like protein aggregates in vitro and in vivo. Cell, 90, 549–58. Scherzinger, E., Sittler, A., Schweiger, K., Heiser, V., Lurz, R., Hasenbank, R., Bates, G. P., Lehrach, H. & Wanker, E. E. (1999) Self-assembly of polyglutamine-containing huntingtin fragments into amyloid-like fibrils: implications for Huntington’s disease pathology. Proc Natl Acad Sci U S A, 96, 4604–9. Sharp, A. H., Loev, S. J., Schilling, G., Li, S. H., Li, X. J., Bao, J., Wagster, M. V., Kotzuk, J. A., Steiner, J. P., Lo, A. & Et al. (1995) Widespread expression of Huntington’s disease gene (IT15) protein product. Neuron, 14, 1065–74. Sinadinos, C., Burbidge-King, T., Soh, D., Thompson, L. M., Marsh, J. L., Wyttenbach, A. & Mudher, A. K. (2009) Live axonal transport disruption by mutant huntingtin fragments in Drosophila motor neuron axons. Neurobiol Dis, 34, 389–95. Smirnova, E., Griparic, L., Shurland, D. L. & Van Der Bliek, A. M. (2001) Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells. Mol Biol Cell, 12, 2245–56. Squitieri, F., Cannella, M., Sgarbi, G., Maglione, V., Falleni, A., Lenzi, P., Baracca, A., Cislaghi, G., Saft, C., Ragona, G., Russo, M. A., Thompson, L. M., Solaini, G. & Fornai, F. (2006) Severe ultrastructural mitochondrial changes in lymphoblasts homozygous for Huntington disease mutation. Mech Ageing Dev, 127, 217–20. Szebenyi, G., Morfini, G. A., Babcock, A., Gould, M., Selkoe, K., Stenoien, D. L., Young, M., Faber, P. W., Macdonald, M. E., Mcphaul, M. J. & Brady, S. T. (2003) Neuropathogenic forms of huntingtin and androgen receptor inhibit fast axonal transport. Neuron, 40, 41–52. Taguchi, N., Ishihara, N., Jofuku, A., Oka, T. & Mihara, K. (2007) Mitotic phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial fission. J Biol Chem, 282, 11521–9. Tang, T. S., Slow, E., Lupu, V., Stavrovskaya, I. G., Sugimori, M., Llinas, R., kristal, B. S., Hayden, M. R. & Bezprozvanny, I. (2005) Disturbed Ca2+ signaling and apoptosis of medium spiny neurons in Huntington’s disease. Proc Natl Acad Sci U S A, 102, 2602–7. Tellez-Nagel, I., Johnson, A. B. & Terry, R. D. (1974) Studies on brain biopsies of patients with Huntington’s chorea. J Neuropathol Exp Neurol, 33, 308–32. Trifunovic, A., Wredenberg, A., Falkenberg, M., Spelbrink, J. N., Rovio, A. T., Bruder, C. E., Bohlooly, Y. M., Gidlof, S., Oldfors, A., Wibom, R., Tornell, J., Jacobs, H. T. & Larsson, N. G. (2004) Premature ageing in mice expressing defective mitochondrial DNA polymerase. Nature, 429, 417–23. Trushina, E., Dyer, R. B., Badger, J. D., 2nd, Ure, D., Eide, L., Tran, D. D., Vrieze, B. T., Legendre-Guillemin, V., Mcpherson, P. S., Mandavilli, B. S., Van Houten, B., Zeitlin, S., Mcniven, M., Aebersold, R., Hayden, M., Parisi, J. E., Seeberg, E., Dragatsis, I., Doyle, K., Bender, A., Chacko, C. & Mcmurray, C. T. (2004) Mutant huntingtin impairs axonal trafficking in mammalian neurons in vivo and in vitro. Mol Cell Biol, 24, 8195–209. Verstreken, P., Ly, C. V., Venken, K. J., Koh, T. W., Zhou, Y. & Bellen, H. J. (2005) Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions. Neuron, 47, 365–78. Walker, F. O. (2007) Huntington’s disease. Lancet, 369, 218–28. Wang, H., Lim, P. J., Karbowski, M. & Monteiro, M. J. (2009a) Effects of overexpression of huntingtin proteins on mitochondrial integrity. Hum Mol Genet, 18, 737–52. Wang, H., Lim, P. J., Yin, C., Rieckher, M., Vogel, B. E. & Monteiro, M. J. (2006) Suppression of polyglutamine-induced toxicity in cell and animal models of Huntington’s disease by ubiquilin. Hum Mol Genet, 15, 1025–41.

9  Mitochondrial Dynamics and Huntington’s Disease: A Dance of Fate

257

Wang, X., Su, B., Lee, H. G., Li, X., Perry, G., Smith, M. A. & Zhu, X. (2009b) Impaired balance of mitochondrial fission and fusion in Alzheimer’s disease. J Neurosci, 29, 9090–103. Westermann, B. (2008) Molecular machinery of mitochondrial fusion and fission. J Biol Chem, 283, 13501–05. Wexler, N. S., Lorimer, J., Porter, J., Gomez, F., Moskowitz, C., Shackell, E., Marder, K., Penchaszadeh, G., Roberts, S. A., Gayan, J., Brocklebank, D., Cherny, S. S., Cardon, L. R., Gray, J., Dlouhy, S. R., Wiktorski, S., Hodes, M. E., Conneally, P. M., Penney, J. B., Gusella, J., Cha, J. H., Irizarry, M., Rosas, D., Hersch, S., Hollingsworth, Z., Macdonald, M., Young, A. B., Andresen, J. M., Housman, D. E., De Young, M. M., Bonilla, E., Stillings, T., Negrette, A., Snodgrass, S. R., Martinez-Jaurrieta, M. D., Ramos-Arroyo, M. A., Bickham, J., Ramos, J. S., Marshall, F., Shoulson, I., Rey, G. J., Feigin, A., Arnheim, N., Acevedo-Cruz, A., Acosta, L., Alvir, J., Fischbeck, K., Thompson, L. M., Young, A., Dure, L., O’brien, C. J., Paulsen, J., Brickman, A., Krch, D., Peery, S., Hogarth, P., Higgins, D. S., Jr. & Landwehrmeyer, B. (2004) Venezuelan kindreds reveal that genetic and environmental factors modulate Huntington’s disease age of onset. Proc Natl Acad Sci U S A, 101, 3498–503. Xia, J., Lee, D. H., Taylor, J., Vandelft, M. & Truant, R. (2003) Huntingtin contains a highly conserved nuclear export signal. Hum Mol Genet, 12, 1393–403. Yamamoto, A., Cremona, M. L. & Rothman, J. E. (2006) Autophagy-mediated clearance of huntingtin aggregates triggered by the insulin-signaling pathway. J Cell Biol, 172, 719–31. Yang, J. L., Weissman, L., Bohr, V. A. & Mattson, M. P. (2008a) Mitochondrial DNA damage and repair in neurodegenerative disorders. DNA Repair (Amst), 7, 1110–20. Yang, Y., Ouyang, Y., Yang, L., Beal, M. F., Mcquibban, A., Vogel, H. & Lu, B. (2008b) Pink1 regulates mitochondrial dynamics through interaction with the fission/fusion machinery. Proc Natl Acad Sci U S A, 105, 7070–5. Yonashiro, R., Ishido, S., Kyo, S., Fukuda, T., Goto, E., Matsuki, Y., Ohmura-Hoshino, M., Sada, K., Hotta, H., Yamamura, H., Inatome, R. & Yanagi, S. (2006) A novel mitochondrial ubiquitin ligase plays a critical role in mitochondrial dynamics. EMBO J, 25, 3618–26. Yoon, Y., Krueger, E. W., Oswald, B. J. & Mcniven, M. A. (2003) The mitochondrial protein hFis1 regulates mitochondrial fission in mammalian cells through an interaction with the dynaminlike protein DLP1. Mol Cell Biol, 23, 5409–20. Youle, R. J. & Karbowski, M. (2005) Mitochondrial fission in apoptosis. Nat Rev Mol Cell Biol, 6, 657–63. Yuan, H., Gerencser, A. A., Liot, G., Lipton, S. A., Ellisman, M., Perkins, G. A. & Bossy-Wetzel, E. (2007) Mitochondrial fission is an upstream and required event for bax foci formation in response to nitric oxide in cortical neurons. Cell Death Differ, 14, 462–71. Zeitlin, S., Liu, J. P., Chapman, D. L., Papaioannou, V. E. & Efstratiadis, A. (1995) Increased apoptosis and early embryonic lethality in mice nullizygous for the Huntington’s disease gene homologue. Nat Genet, 11, 155–63. Zhang, W., Chen, Y., Yang, Q., Che, H., Chen, X., Yao, T., Zhao, F., Liu, M., Ke, T., Chen, J. & Luo, W. (2010) Mitofusin-2 protects against cold stress-induced cell injury in HEK293 cells. Biochem Biophys Res Commun, 397, 270–6. Zuccato, C., Valenza, M. & Cattaneo, E. (2010) Molecular mechanisms and potential therapeutical targets in Huntington’s disease. Physiol Rev, 90, 905–81. Zuchner, S., Mersiyanova, I. V., Muglia, M., Bissar-Tadmouri, N., Rochelle, J., Dadali, E. L., Zappia, M., Nelis, E., Patitucci, A., Senderek, J., Parman, Y., Evgrafov, O., Jonghe, P. D., Takahashi, Y., Tsuji, S., Pericak-Vance, M. A., Quattrone, A., Battaloglu, E., Polyakov, A. V., Timmerman, V., Schroder, J. M. & Vance, J. M. (2004) Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat Genet, 36, 449–51.

Index

A Aging, 41, 59, 72, 93–100, 202, 244 Anterograde transport, 141, 143, 145, 149, 154, 156 Apoptosis, 7, 9, 15, 23, 26, 28, 32, 34–38, 42, 49, 53, 56, 58, 71, 72, 81, 94, 109–129, 173, 176, 180, 181, 184, 207, 217, 224, 225, 243, 248 Axonal transport, 92, 139–162, 173, 175, 183, 246 B Bioenergetics, 47–63, 71, 216, 221, 223, 240 C Cell death, 7, 28, 35, 38, 58, 78, 81, 110, 112, 113, 116–120, 126–129, 180, 181, 198, 202–205, 207, 216, 217, 223–226, 248 Charcot-Marie-Tooth (CMT) disease, 24, 26, 63, 72, 124, 174–175, 178, 217, 244 D Docking, 20, 141, 142, 147, 150–153, 156, 157, 159–162 Dominant optic atrophy (DOA), 118, 124, 170–176, 178–189, 244 Dynamics, 3–17, 20, 34–42, 47–63, 69–100, 109–129, 139–162, 171, 174, 180, 182, 183, 199–208, 217–219, 222–228, 235–251 Dynamin-related protein 1 (Drp1), 12, 13, 15–17, 19, 21–25, 30–36, 39, 54–57, 61, 63, 71, 80, 84–86, 89, 90, 92, 94, 95, 111–118, 122, 124, 127–129, 157, 158, 202–204, 215–228, 243, 245, 248–250 Dynamin-related proteins (DRPs), 16, 25, 113

G Genetic screens, 13, 245 H Huntingtin (htt), 236–239, 241, 245–251 K Kinesin, 91, 92, 141–150, 156, 158–159 M Mechanoenzymes, 16, 17, 20, 28 Membranes and lipid homeostasis, 14, 31, 39–40 Membrane scission and fusion, 13, 20, 25 Mitochondria dynamics, 3–17, 20, 34–42, 47–63, 69–100, 109–129, 139–162, 171, 174, 180, 182, 183, 199–208, 217–219, 222–228, 235–251 fission, 1–42, 49, 53–57, 59, 61, 71, 72, 83, 84, 86–89, 91, 93, 94, 97, 111, 113–118, 127, 129, 156, 180, 181, 197–208, 215–228, 241–246, 248, 250 fusion, 1–42, 49, 53–63, 71, 72, 83–86, 88, 89, 91–94, 97, 112, 113, 118–121, 123–129, 157, 169–189, 201–203, 207, 217–219, 224, 225, 228, 241–246, 248–250 morphology, 11–15, 23, 24, 29, 32, 34, 71, 84–86, 90–92, 95, 111, 117, 120, 127–129, 161, 180, 199–201, 204, 205, 216, 222–224, 227, 228 quality control, 79, 84, 87, 89, 92–94, 97, 199, 206, 207 Mitochondrial complex I inhibitors, 198, 201, 203, 227

B. Lu (ed.), Mitochondrial Dynamics and Neurodegeneration, DOI 10.1007/978-94-007-1291-1, © Springer Science+Business Media B.V. 2011

259

260 Mitochondrial mobility, 141, 150–159, 161, 162 Mitofusin (Mfn), 12, 15, 23, 26, 31, 33, 39, 53, 57, 58, 71, 84, 85, 119, 123–126, 128, 157, 174, 179, 183, 202, 204, 206, 217, 219, 228, 248–250 Mitofusin 2 (Mfn2), 12, 15, 17, 18, 26, 30, 36, 53–58, 61, 63, 71, 72, 85, 86, 90, 92, 111, 112, 115, 118, 123–129, 157, 173–189, 217, 219, 224, 225, 242, 244, 245, 248, 250 Mitophagy, 24, 36–38, 40, 78–100, 156, 162, 176, 187, 188, 200, 201, 203–207, 227, 228 Motor adaptor, 141, 146, 147, 150, 154, 156, 162 N Neurodegeneration, 7, 36–38, 61, 72, 78, 82, 83, 93–100, 161, 198, 199, 205–208, 215–228, 237, 244 Neurodegenerative diseases, 5, 7, 15, 37, 39, 72, 82, 98, 140, 151, 161, 162, 217, 225–228, 244 Neuronal cell death, 38, 202, 223, 226 O Optic atrophy 1 (OPA1), 11, 14, 15, 18, 20, 26–31, 36, 39, 53–58, 62, 63, 71, 72, 84, 86–90, 92, 111–113, 118–124, 126, 127, 129, 170–189, 217–219, 223, 224, 226, 228, 242, 244

Index Oxidative phosphorylation (OXPHOS), 6, 15, 37, 49, 50, 52, 56–60, 62, 63, 71, 80, 111, 112, 172, 180, 181, 240, 244 P Parkin, 31, 63, 83–87, 92, 96, 97, 126, 187, 188, 199, 200, 205–207, 227, 228 Parkinson disease (PD), 39, 63, 78, 82, 83, 95–98, 187, 197–208, 223, 225–228, 244 Polyglutamine expansion, 237, 246 Proteolytic processing, 13, 20, 27, 30, 62, 120–123, 217, 244 PTEN induced putative kinase 1 (PINK1), 39, 83–87, 90, 92, 93, 96, 97, 126, 187, 199, 200, 203–207, 227, 228 R Retrograde transport, 145, 146, 147, 150, 155 S Stationary mitochondria, 91, 150–153 Synaptic damage, 215–228 Synaptic plasticity, 140, 158–159