International Review Of Cell and Molecular Biology, Volume 285 (International Review of Cell & Molecular Biology)

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INTERNATIONAL REVIEW OF

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INTERNATIONAL REVIEW OF CELL AND MOLECULAR BIOLOGY Series Editors

GEOFFREY H. BOURNE JAMES F. DANIELLI KWANG W. JEON MARTIN FRIEDLANDER JONATHAN JARVIK

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ISAIAH ARKIN PETER L. BEECH ROBERT A. BLOODGOOD DEAN BOK KEITH BURRIDGE HIROO FUKUDA RAY H. GAVIN MAY GRIFFITH WILLIAM R. JEFFERY KEITH LATHAM

WALLACE F. MARSHALL BRUCE D. MCKEE MICHAEL MELKONIAN KEITH E. MOSTOV ANDREAS OKSCHE MANFRED SCHLIWA TERUO SHIMMEN ROBERT A. SMITH ALEXEY TOMILIN

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KWANG W. JEON Department of Biochemistry University of Tennessee Knoxville, Tennessee

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Front Cover Photography: Cover figure by Timo Mu¨hlhaus and Michael Schroda Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32 Jamestown Road, London NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands First edition 2010 Copyright # 2010, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@elsevier. com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material. Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress For information on all Academic Press publications visit our website at elsevierdirect.com

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CONTENTS

Contributors

1. Cell and Molecular Biology of Microtubule Plus End Tracking Proteins: End Binding Proteins and Their Partners

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Susana Montenegro Gouveia and Anna Akhmanova 1. Introduction 2. Microtubules (MTs) 3. MT Regulators 4. MT Plus End Tracking Proteins (þTIPs) 5. þ TIP Families 6. Conclusions and Future Directions Acknowledgments References

2. New Insights into the Roles of Molecular Chaperones in Chlamydomonas and Volvox

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¨hlhaus, and Michael Schroda Andre´ Nordhues, Stephen M. Miller, Timo Mu 1. Introduction 2. Cytosol/Nucleus 3. Flagella 4. Endoplasmic Reticulum 5. Chloroplast 6. Mitochondrion 7. Conclusions and outlook Acknowledgments References

3. Unique Functions of Repetitive Transcriptomes

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Gerald G. Schumann, Elena V. Gogvadze, Mizuko Osanai-Futahashi, ¨nk, Haruko Fujiwara, Zoltan Ivics, Azusa Kuroki, Carsten Mu and Anton A. Buzdin 1. Introduction 2. Eukaryotic Retrotransposons 3. Mechanisms of Intracellular Defense against TEs

116 119 123 v

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4. The Use of Transposable Elements in Biotechnology and in Fundamental Studies 5. Domestication of Mobile DNA by the Host Genomes 6. Retrotransposons as Drivers of Mammalian Genome Evolution 7. Concluding Remarks Acknowledgments References Index

134 149 162 167 167 167 189

CONTRIBUTORS

Anna Akhmanova Department of Cell Biology, Erasmus Medical Center, Rotterdam, The Netherlands Anton A. Buzdin Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, Russia Haruko Fujiwara Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Japan Elena V. Gogvadze Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, Russia Susana Montenegro Gouveia Department of Cell Biology, Erasmus Medical Center, Rotterdam, The Netherlands Zoltan Ivics Max Delbruck Center for Molecular Medicine, Berlin, Germany; and University of Debrecen, Debrecen, Hungary Azusa Kuroki Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Japan Stephen M. Miller Department of Biological Sciences, University of Maryland, Baltimore County, Baltimore, Maryland, USA ¨hlhaus Timo Mu Max Planck Institute of Molecular Plant Physiology, Am Muehlenberg, PotsdamGolm, Germany ¨nk Carsten Mu Clinic for Gastroenterology, Hepatology and Infectiology, Medical Faculty, Heinrich-Heine-University, Du¨sseldorf, Germany Andre´ Nordhues Max Planck Institute of Molecular Plant Physiology, Am Muehlenberg, PotsdamGolm, Germany

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Contributors

Mizuko Osanai-Futahashi Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Japan Michael Schroda Max Planck Institute of Molecular Plant Physiology, Am Muehlenberg, PotsdamGolm, Germany Gerald G. Schumann Paul-Ehrlich-Institut, Federal Institute for Vaccines and Biomedicines, Langen, Germany

C H A P T E R

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Cell and Molecular Biology of Microtubule Plus End Tracking Proteins: End Binding Proteins and Their Partners Susana Montenegro Gouveia and Anna Akhmanova Contents 2 2 2 3 4 6 7 8 9 9 10 18 25 36 43 49 50 51 52

1. Introduction 2. Microtubules (MTs) 2.1. Tubulin 2.2. MT organization and structure 2.3. MT dynamics 2.4. Role of GTP hydrolysis in MT dynamics 2.5. MT plus end structure and the GTP cap 3. MT Regulators 4. MT Plus End Tracking Proteins (þTIPs) 5. þ TIP Families 5.1. EB family 5.2. CAP-Gly proteins 5.3. Proteins with basic and serine-rich regions 5.4. TOG and TOG-like domain proteins 5.5. Motor proteins 5.6. Other þ TIPs 6. Conclusions and Future Directions Acknowledgments References

Abstract The microtubule plus end is a crucial site for the regulation of microtubule dynamics and microtubule association with different cellular organelles and macromolecular complexes. Several evolutionarily conserved groups of proteins form comet-like accumulations at the growing microtubule plus ends. These proteins belong to functionally diverse and structurally unrelated Department of Cell Biology, Erasmus Medical Center, Rotterdam, The Netherlands International Review of Cell and Molecular Biology, Volume 285 ISSN 1937-6448, DOI: 10.1016/S1937-6448(10)85001-4

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2010 Elsevier Inc. All rights reserved.

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families: they include motors, nonmotor proteins, microtubule polymerases, and depolymerases as well as regulatory and adaptor proteins. Here, we provide an overview of microtubule plus end binding proteins, describe what is known about the mechanisms of their association with growing microtubule tips, and discuss their functional properties in relation to microtubule plus end accumulation. Key Words: Microtubule, EB1, CLIP-170, APC, Spectraplakin, STIM1, Kinesin, Dynein. ß 2010 Elsevier Inc.

1. Introduction The cytoskeleton is a scaffold made of proteinaceous fibers that is required for the majority of essential cellular functions, such as maintenance of cellular shape, cell motility, intracellular transport, and cell division. It is a complex and often highly dynamic structure composed of several major components. One of them are microtubules (MTs), which are among the most ubiquitous cytoskeletal elements present in all eukaryotic cells. MTs are hollow asymmetric tubes that are built and broken down from their ends. In this review, we zoom in on a tiny part of the MT—its growing end. This structure, which is 25 nm in diameter and not more than 1 or 2 mm in length, is surprisingly complex: it concentrates a large set of structurally diverse proteins and serves as a site of convergence of numerous cellular processes. Here, we summarize what is known about a peculiar and highly conserved group of proteins that form comet-like accumulations at the growing MT tips, discuss the molecular mechanisms of MT plus end localization, and describe how MT end localization relates to the functions of these proteins.

2. Microtubules (MTs) 2.1. Tubulin MTs are cylindrical protein filaments found in all eukaryotes. The structural subunit of MTs is tubulin, a heterodimeric protein composed of two polypeptide chains designated a and b tubulin (Krauhs et al., 1981; Ponstingl et al., 1981). The a and b monomers have similar masses of 55 kDa and interact noncovalently to form a very stable heterodimer, the functional form of the protein (Wade, 2009). Individual tubulin heterodimers are 8 nm in length. Each tubulin monomer can be divided into three functional domains: the N-terminal domain containing the GTP-binding

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site, an intermediate domain containing the taxol-binding site in b tubulin, and the C-terminal domain, which probably forms the binding surface for many MT-associated proteins (MAPs) (Nogales and Wang, 2006b; Nogales et al., 1998). Each tubulin monomer binds a guanine nucleotide. The GTP bound to a tubulin does not get hydrolyzed during polymerization and is nonexchangeable; the site to which it binds is therefore referred to as the N-site (Desai and Mitchison, 1997). In contrast, GTP bound to b tubulin at the E-site is exposed on the surface of the dimer and can exchange but gets locked in the protofilament as GDP after undergoing hydrolysis during polymerization. When a MT depolymerizes, the GDP-bound b tubulin exchanges GDP to GTP in solution. In most eukaryotes, the tubulin gene family encodes multiple isoforms or isotypes (Wade, 2007). The expression of multiple genes leads to a variety of slightly different a and b tubulins. Each tubulin isotype differs from the others in its amino acid sequence and/or the temporal, tissue, and subcellular distribution (Luduena, 1998). In addition, both a and b tubulin undergo a variety of posttranslational covalent modifications, including acetylation, phosphorylation, detyrosynation, polyglutamylation, and polyglycylation, many of which target the C-terminal tail of tubulin (Hammond et al., 2008).

2.2. MT organization and structure MTs within cells can be organized in different types of networks, such as a radial array in interphase fibroblasts, a bipolar spindle in mitosis and meiosis, a parallel array in polarized epithelial cells, and a linear array in neuronal extensions. In addition, MTs form highly stable and precisely organized structures, such as centrioles, basal bodies, and axonemes of cilia and flagella. MTs are cylinders of 25 nm in diameter but can have variable length. Their basic building blocks, a and b tubulin heterodimers, are arranged in a head-to-tail fashion into linear protofilaments, which associate laterally to form MTs. MTs commonly found in vivo have 13 protofilaments (Ledbetter and Porter, 1964; Tilney et al., 1973), although exceptions have been described, such as 15-protofilament MTs in certain neuronal cells of Caenorhabditis elegans (Bounoutas et al., 2009). In vitro, the protofilament number in MTs spontaneously assembled from mammalian brain tubulin can vary from 9 to 17, even within the same MT (Chretien and Wade, 1991; Chretien et al., 1992). MTs in centrioles and axonemes represent a special case, as they can form doublets or triplets, in which some MTs are incomplete (Unger et al., 1990). Two distinct MT lattice structures are possible: an A-type and a B-type lattice. In an A-type lattice, a monomers from one protofilament associate with b monomers from the adjacent one, while in a B-type lattice, the a–a

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and b–b contacts are formed between the adjacent protofilaments (Desai and Mitchison, 1997). Ultrastructural analysis showed that the B-type lattice is the naturally occurring protofilament arrangement (Mandelkow et al., 1986; Song and Mandelkow, 1993). This implies that MTs have a lattice discontinuity, called the seam, where a and b monomers from adjacent protofilaments undergo lateral interactions. In the B-type lattice, the helical pattern is created by a–a and b–b lateral bonds, with a–b bonds present at the seam (Desai and Mitchison, 1997). It was proposed that MTs grow as a sheet and later zipper into a closed tube at the seam (Chretien et al., 1995). Because at the seam the lateral interactions are weaker, it might also play a role in MT disassembly (Wade, 2007). Interestingly, A lattice formation was recently observed in vitro in the presence of a yeast MT end binding protein Mal3 (see below; des Georges et al., 2008). However, careful reexamination of MT lattice structure confirmed earlier findings that 13-protofilament B-lattice MTs with a seam is the predominant form in cultured mammalian cells (McIntosh et al., 2009). MTs display intrinsic polarity, generated by the head-to-tail assembly of a/b tubulin dimers, with one end growing more rapidly than the other in vitro. The dynamic and faster growing end of a MT is termed the plus end, and the slower growing and less dynamic one the minus end. In vitro experiments showed that the opposite ends of free MTs have different sensitivities to MT depolymerizing agents such as low temperature, Ca2þ, or colchicine (Summers and Kirschner, 1979). MT minus ends are usually anchored at the centrosome or other minus end stabilizing sites (Dammermann et al., 2003), while the plus ends explore the cytoplasmic space and often interact with the cell cortex and other cellular structures. The polarity of MTs is central to the mechanism of action of motor proteins, kinesins, and dyneins, which use ATP hydrolysis to transport various cargos along MTs. Motor proteins are unidirectional: they can move either toward MT plus ends (kinesins), or toward the minus ends (a few kinesins and cytoplasmic dynein) (Mallik and Gross, 2004). Together with asymmetric MT organization, MT-based motors thus contribute to cell polarity and morphogenesis.

2.3. MT dynamics MTs can assemble spontaneously in solutions of purified tubulin. Observation of a population of fixed MTs led to the discovery of MT dynamic instability, a process whereby MTs interconvert between phases of polymerization and depolymerization (Mitchison and Kirschner, 1984). Transitions between MT growth and shortening are abrupt and stochastic and are defined as catastrophe for the switch from growth to shortening and rescue for the switch from shortening to growth (Desai and Mitchison, 1997). Subsequent extensive studies using dark field and differential

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interference contrast (DIC) microscopy have established this phenomenon both in vitro and in vivo (Cassimeris et al., 1988; Horio and Hotani, 1986; Hotani and Horio, 1988; Walker et al., 1988). One additional observation was that MTs sometimes neither polymerize nor depolymerize, they just pause; this behavior is much more frequent in vivo but also occurs in vitro with pure tubulin (Desai and Mitchison, 1997; Walker et al., 1988). Four parameters have been proposed to describe dynamic instability in vitro: growth velocity, shrinking velocity, and frequencies of catastrophe and rescue (Walker et al., 1988). The catastrophe frequency is determined as the number of catastrophes observed during the total time MTs spend in the growing state. Rescues are defined in a similar way. Because of the frequent pausing in vivo, additional parameters describing the frequencies of entry and exit from the pausing state are needed for quantitative description of MT dynamics in cells. MT growth velocity depends on the soluble tubulin concentration and on the rate constants for GTP-tubulin association and dissociation during growth. The shortening velocity is independent of tubulin concentration and is characterized only by the dissociation rate of GDP-tubulin at the depolymerizing end (Erickson and O’Brien, 1992). The growth velocity of a MT in vitro is typically 1–5 mm/min depending on several factors such as tubulin concentration, temperature, and presence or absence of MAPs. Several studies suggested that a MT end structure might regulate the rates of polymerization and depolymerization, based on the fact that individual MTs exhibit significant variability in their assembly and disassembly rates (Gildersleeve et al., 1992; O’Brien et al., 1990). Some propagating structural feature such as the protofilament number (Chretien et al., 1992) or lattice organization might be responsible for this variability. The shortening velocity is up to 10-fold higher than growth, 10–50 mm/min, and rescues are rarely observed in vitro. Catastrophes are assumed to be stochastic events, and their frequency is largely dependent on tubulin concentration and temperature. In spite of their stochasticity, catastrophes do not display first-order kinetics (Odde et al., 1995), and their mechanism might be complex. MT dynamics plays an essential role in the organization and rapid remodeling of the cytoskeleton, allowing MTs to search the 3D cytoplasmic space (Holy and Leibler, 1994; Kirschner and Mitchison, 1986). MT dynamics is regulated during the cell cycle: MTs in interphase turn over in several minutes, while in mitosis, this occurs in seconds (McNally, 1996). Overall, MTs in living cells display rapid polymerization rates and high frequencies of rescues and catastrophes (Cassimeris et al., 1988). This implies the existence of cellular factors that independently modulate MT dynamics in vivo, either by promoting growth, increasing transition frequencies, or affecting MT stability (Cassimeris, 1993; Howard and Hyman, 2007; van der Vaart et al., 2009).

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2.4. Role of GTP hydrolysis in MT dynamics GTP hydrolysis provides the energy source for dynamic instability (Hyman et al., 1992). To address the exact role of GTP hydrolysis, several studies were performed with nonhydrolysable analogues of GTP, GMPPNP, and GMPPCP (Kirschner, 1978; Mejillano et al., 1990a,b). However, these analogues have a low affinity for b tubulin (Desai and Mitchison, 1997). Studies performed with a slowly hydrolysable analogue of GTP, guanylyl(a,b)-methylene-diphosphonate (GMPCPP), lead to the important conclusion that GTP hydrolysis is not required for polymerization but is essential for dynamic instability (Hyman et al., 1992). GMPCPP binds to the E-site of tubulin with a significantly lower affinity than GTP, and promotes MT nucleation and polymerization. GMPCPP MTs do not show dynamic instability, but continuously grow without transitions to a shrinking phase. Interestingly, GMPCPP hydrolysis in the MT lattice could be triggered by specific buffer conditions, resulting in the generation of relatively unstable GMPCP MTs and thus providing a direct indication that GTP hydrolysis is important for MT dynamics (Caplow et al., 1994). Further investigations attempted to clarify the role of GTP hydrolysis in the mechanism of catastrophes. Cryoeletron microscopy (cryoEM) studies revealed that protofilaments within a MT are straight, whereas depolymerizing ends contain highly curved protofilaments (Kirschner, 1978; Mandelkow and Mandelkow, 1985; Mandelkow et al., 1991). Observation of MT disassembly induced by cold and calcium showed that depolymerizing ends of GDP MTs were consistently more curved than those of GMPCPP MTs (Muller-Reichert et al., 1998). These results support a model in which GTP hydrolysis increases the intrinsic protofilament curvature. To address the structural basis of this phenomenon, Wang and Nogales solved the structure of GMPCPP tubulin and showed that GMPCPP protofilaments curve outward, albeit to a lesser extent than GDP protofilaments (Wang and Nogales, 2005). They also observed that GMPCPP tubulin forms helical ribbons when polymerized at low temperatures and at high concentration of Mg2þ, and that an increase in temperature results in the direct conversion of these structures into MTs. The authors suggested that the observed helical ribbons of tubulin correspond to the early stages of MT assembly that could be regarded, from a structural point of view, as an intermediate state between tubulin sheet and the MT tube (Wang et al., 2005). Sheet intermediates were also observed by cryoEM during MT growth in vitro (Chretien et al., 1995; Erickson, 1974) and in vivo (Zovko et al., 2008). Existence of sheet intermediates is also supported by theoretical analysis (Wu et al., 2009). The function of the sheet-like intermediates remains an open question, as they were proposed to be stabilizing and destabilizing MT end structures (Arnal et al., 2000; VanBuren et al., 2005). The situation in vivo

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is even more complex with the existence of MAPs that modify the process of MT assembly and disassembly.

2.5. MT plus end structure and the GTP cap GTP hydrolysis is supposed to occur rapidly after tubulin polymerization, generating a MT lattice of GDP-tubulin that is less stable. Polymerizing MTs are believed to be stable due to a cap of GTP-bound b tubulin in which GTP hydrolysis has not yet occurred, preventing the MT from shrinking. As the GTP hydrolysis does not occur in the last dimer added to the MT plus end but only in the penultimate ones, there is a lag between polymerization and GTP hydrolysis ensuring MT stability (Downing, 2000; Nogales, 2000). However, if hydrolysis is faster than polymerization, the GTP cap is lost resulting in a catastrophe. Several groups attempted to measure or estimate the size of the GTP cap. First evidence based on the analysis of the rate of disassembly of diluted MTs estimated that the maximum size of the GTP cap would be 40 and 20 subunits at the plus and minus ends, respectively (Voter et al., 1991). Next, studies of MTs composed of a mixture of GMPCPP (a slowly hydrolysable analogue of GTP) and GDP-tubulin proposed that a single layer of GTP-tubulin is necessary and sufficient to stabilize MTs (Caplow and Shanks, 1996; Drechsel and Kirschner, 1994). Recently, computational simulations and theoretical models have been generated on the basis of existing experimental data. Taking into account the bending properties of GDP-bound tubulin and the elasticity of the MT, theoretical models show that a relatively short or even incomplete structural cap can stabilize MTs ( Janosi et al., 2002). A molecular-mechanical model, in which structural and biochemical properties of tubulin were used to predict the shape and stability of MTs, suggested that the MT plus end is stabilized by at least two layers of GTP-tubulin subunits, whereas the minus end requires at least one layer (Molodtsov et al., 2005). Another theoretical study also concluded that a very small cap, equal to a single GTP-tubulin ring at the growing end is necessary and sufficient to maintain a metastable equilibrium (Hunyadi and Janosi, 2007). It is important to note that these studies provide estimates for the minimal, and not necessarily the actual size of the cap, which might be more extended (Piette et al., 2009; VanBuren et al., 2005). Importantly, all theoretical models reach the same conclusion: the energy released from GTP hydrolysis becomes constrained in the MT lattice generating force to perform work or cause rapid depolymerization when the GTP cap is lost (Nogales and Wang, 2006a). New insights emerged with recent experiments that use optical tweezers to track MT growth. An important finding achieved by these experiments was that MTs exhibit growth fluctuations on subsecond timescales (Gardner et al., 2008b; Kerssemakers et al., 2006; Schek et al., 2007). It was suggested

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that these growth fluctuations are explained by the addition of tubulin oligomers from solution (Kerssemakers et al., 2006). This view, which goes against the idea that MT growth occurs by addition of single tubulin heterodimers, is supported by experiments that determined the interaction energy of tubulin dimers and by EM analysis and fluorescence correlation spectroscopy measurements indicating that tubulin dimers oligomerize before incorporation into the MT (Mozziconacci et al., 2008). This view is, however, contradicted by direct single molecule level observations of MT growth in the presence of MT polymerase XMAP215 (Brouhard et al., 2008). An alternative explanation for the growth fluctuations would be a dynamic GTP cap, which is spatially extended through the MT lattice, allowing nanoscale depolymerization events but preventing “true” catastrophes (Schek et al., 2007). It is unclear how all these studies can be correlated with the observation of MT sheets at the growing MT ends by cryoEM (Arnal et al., 2000; Chretien et al., 1995). A recent paper visualized for the first time the putative GTP cap in vivo, based on a new anti-tubulin antibody that specifically recognizes the GTPtubulin conformation in MTs (Dimitrov et al., 2008). Unexpectedly, it was found that besides colocalization with the plus ends of MTs, GTP-tubulin remnants were also present in the MT lattice, suggesting that GTP hydrolysis might not always be complete (Howard and Hyman, 2009). Another explanation is that the MT lattice can “breathe,” allowing subunits to enter and exit from the middle of the lattice (Cassimeris, 2009). Finally, in agreement with the dynamic GTP cap model, a recent computational thermodynamic model of MT dynamics proposed that the GTP cap can fluctuate and be several micrometers long (Piette et al., 2009). Thus, although the concept of a stabilizing cap at the MT plus end is generally accepted, its structural and biochemical details require further elucidation.

3. MT Regulators MTs are primarily controlled by factors that can directly regulate different parameters of MT polymerization dynamics, sever or bundle existing MTs or affect MTs indirectly, through influencing other proteins (Lyle et al., 2009a,b). MTs can be stabilized in several ways: by preventing catastrophes, by rescuing depolymerizing MTs, or by decreasing shrinkage velocity. MT regulators that induce catastrophes, prevent rescues, or increase shrinkage speeds destabilize MTs (Desai and Mitchison, 1997; van der Vaart et al., 2009). All MT regulators are MAPs that were initially defined as proteins that copurify with tubulin (Olmsted, 1986). Nowadays, the definition evolved and includes all proteins that specifically bind to tubulin or MTs, either

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in vitro or in vivo. Their activities cover a wide range of regulatory processes. Classical MAPs, such as tau and MAP2, which are highly abundant in neurons, stabilize the MT lattice (Dehmelt and Halpain, 2005). There are MAPs that nucleate MTs, like g tubulin-associated proteins (Wiese and Zheng, 2006), or promote MT assembly by adding tubulin subunits to MT ends, such as XMAP215 (Brouhard et al., 2008). MAPs can induce MT bundling (e.g., MAP65; van Damme et al., 2004b), or, conversely, sever MTs, like katanin and spastin, creating new plus and minus ends (McNally and Vale, 1993). Stathmin ( Jourdain et al., 1997) and MCAK (Hunter et al., 2003) are known to promote MT disassembly. Kinesins and dyneins walk along MTs and transport different cargo, but in some cases, they can also affect MT dynamics (Hunter and Wordeman, 2000).

4. MT Plus End Tracking Proteins (þTIPs) MT plus end tracking proteins (þTIPs) belong to a distinct class of MAPs that accumulate at the growing MT plus ends (Schuyler and Pellman, 2001). Their MT end localization is the distinguishing feature when compared to other MAPs. Most þTIPs associate preferentially with growing and not depolymerizing MT ends. Interestingly, some þTIPs are also able to track shrinking MT ends (“backtracking”). This phenomenon is characteristic for þTIPs from budding yeast (Carvalho et al., 2004; Salmon, 2005), but has also been observed in Drosophila (Mennella et al., 2005), Xenopus (Brouhard et al., 2008), and mammalian cells (Langford et al., 2006). It could be explained by biased diffusion along the shrinking MT lattice, by specific recognition of shrinking MT ends or by kinesindependent MT plus end accumulation.

5. þ TIP Families Since the discovery of the first þ TIP, CLIP-170 (Perez et al., 1999) multiple proteins showing this behavior have been identified. Importantly, some þTIPs localize in a comet-like pattern at MT ends: they are highly concentrated at or close to the freshly polymerized MT tip, while their accumulation decreases exponentially on the older MT lattice. In mammalian cells, such þTIP comets are typically 1–2 mm in length. Since 1 mm of a 13-protofilament MT contains 1600 tubulin dimers, hundreds of þTIP molecules can potentially be bound to each MT end. þTIPs showing this “comet-like” behavior are exemplified by EB proteins and their partners, which are the main focus of this review. Other þTIPs, such as Dam1/ DASH complex on yeast kinetochores are likely to be present in a few copies

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close to MT ends and do not form comets ( Joglekar et al., 2010). Because several excellent recent reviews summarize current knowledge on MTkinetochore association ( Joglekar et al., 2010; Kops et al., 2010; McEwen and Dong, 2010), these proteins will not be discussed here in any detail. Finally, all kinesin motors that display processive plus end-directed motion and do not detach rapidly once they reach the MT end are potential þ TIPs. Important members of this þTIP group are kinesin-8 motors, autonomous þTIPs that can control MT dynamics (Gardner et al., 2008c). Here, we will only briefly discuss kinesin families that have been functionally and biochemically linked to the EB protein interaction network. Even if one considers only the “comet-making” þTIPs, they represent a very heterogeneous group that is difficult to categorize according to functional aspects. We therefore subdivide it based on structure and homologies: presence of conserved binding domains, repeat sequences, and linear motifs (LMs).

5.1. EB family 5.1.1. Structure and evolutionary conservation EB family members are evolutionary conserved proteins, which in mammals are encoded by the MAPRE gene family and include EB1, EB2 (RP1), and EB3 (EBF3). EB1 was initially identified in a yeast two-hybrid screen as a protein that interacts with the C-terminus of the adenomatous polyposis coli (APC) tumor suppressor protein, hence it was named End Binding protein (Su et al., 1995). In mammals, there is a single isoform of EB1 but there are two isoforms of EB2 translated from different initiation codons, and two isoforms of EB3 translated from two alternatively spliced mRNAs (Su and Qi, 2001). EB1 and EB3 are ubiquitously expressed, while expression levels of EB2 vary between cell lines (Su and Qi, 2001). EB3 is especially abundant in the central nervous system and in muscles (Nakagawa et al., 2000; Straube and Merdes, 2007). Among the EB homologues in other species, the yeast proteins, Mal3 in Schizosaccharomyces pombe (Beinhauer et al., 1997) and Bim1 in Saccharomyces cerevisiae (Schwartz et al., 1997), were the most extensively studied. The EB family is also conserved in plants and comprises three members in Arabidopsis: AtEB1a/AtEB1-Homolog2, AtEB1b/AtEB1, and AtEB1c/AtEB1-Homolog1 (Chan et al., 2003; Mathur et al., 2003). EB family proteins are usually dimers that contain two highly conserved domains connected by a linker sequence (Fig. 1.1). The N-terminal part of the EBs consists of a calponin homology (CH) domain, the crystal structure of which has been determined showing a highly conserved fold (Hayashi and Ikura, 2003; Komarova et al., 2009; Slep and Vale, 2007). This domain is necessary and sufficient for binding to MTs and recognizing growing MT ends (Hayashi and Ikura, 2003; Komarova et al., 2009; Skube et al., 2010). The CH domain is followed by a positively charged but poorly

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EB1

Calponin homology domain

Linker

EEY/F motif EB1-homology Acidic domain tail Coiled coil

Microtubules

+TIP partners

Figure 1.1 Schematic representation of EB1 protein sequence. Interaction sites with MTs and þ TIP partners are indicated.

conserved linker region that contributes to MT affinity and is essential for binding to MT ends in some family members, such as the budding yeast Bim1 (des Georges et al., 2008; Komarova et al., 2009; Zimniak et al., 2009). The C-terminal region of EB1 (EB1c) contains a coiled coil, which is responsible for dimerization (Su and Qi, 2001). The coiled coil domain of EB proteins partially overlaps with the unique motif, referred to as end binding homology (EBH) domain, which is implicated in the interaction with numerous EB binding partners (Akhmanova and Steinmetz, 2008). The structure of the EB1c was solved by X-ray crystallography; it was shown that it is a coiled coil, which ends with a four helix bundle; it comprises a deep hydrophobic cavity and a polar rim (Honnappa et al., 2005; Slep et al., 2005). The unstructured acidic tail, the C-terminal sequence of 20–30 residues, was implicated in EBs self-inhibition and interaction with other proteins (Hayashi et al., 2005; Honnappa et al., 2005). It contains a highly conserved C-terminal EEY/F sequence motif similar to those found in a tubulin and CLIP-170 (Komarova et al., 2005; Miller et al., 2006; Weisbrich et al., 2007). It should be noted that plant EB homologues lack the EEY/F sequence; furthermore, one of the three EB1 family members in A. thaliana, AtEB1c, contains a basic rather than an acidic tail, which is responsible for the nuclear localization of the protein (Komaki et al., 2010). All mammalian EB proteins are very similar in structure, but it appears that EB2 is the most divergent family member showing differences in expression, as well as interactions with MTs and binding partners (Morrison, 2007). The C-terminal tail of EB2 differs considerably from those of EB1 and EB3 by having fewer acidic residues; this probably explains why EB2 has a reduced affinity for several partners, such as APC, CLIP-170, and MCAK (Bu and Su, 2003; Komarova et al., 2005; Lee et al., 2008). In addition, EB2 differs from EB1 and EB3 by the presence of an 40 amino acid N-terminal extension (Komarova et al., 2009). The residue differences found between the N-terminal regions of EB3 and EB2 CH domains cluster around a conserved sequence SRHD, which is critical for

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MT binding (Komarova et al., 2009; Slep and Vale, 2007). In addition, EB1 and EB3 form heterodimers in vivo and in vitro, while this feature is less pronounced for EB2 (De Groot et al., 2009; Komarova et al., 2009). The EB1–EB3 heterodimer generates an additional EB variant with distinct properties; consequently, in cells that coexpress different EB species, their functions cannot be addressed separately from each other (De Groot et al., 2009). Interestingly, the chain exchange between the EBs can be suppressed by EBs interaction partners (De Groot et al., 2009). Heterodimer formation was also observed for plant EBs: AtEB1a and AtEB1b heterodimerize with each other, but not with AtEB1c (Komaki et al., 2010). EB proteins bind to multiple partners and until now two major binding modes have been described: through the association of a linear SxIP motif with the hydrophobic cavity of the EBs and through the binding of CAPGly domains to the C-terminal EEY motif of the EBs. The first mode is used by a broad group of proteins including APC, MT-actin crosslinking factor (MACF), CLIP associating proteins (CLASPs), stromal interaction molecule 1 (STIM1), and MCAK, while the second one applies to the CLIPs and dynactin large subunit p150Glued. The specific details of these interactions will be discussed below. 5.1.2. Mechanism of MT plus end tracking by EB proteins In vitro studies using EB homologues from both yeast and vertebrates convincingly demonstrated that they are autonomous þTIPs, which can track growing but not depolymerizing plus and minus ends. This behavior was reconstituted for Mal3 (Bieling et al., 2007), vertebrate EB1 and EB3, (Bieling et al., 2008; Dixit et al., 2009; Komarova et al., 2009) and Bim1 (Blake-Hodek et al., 2010; Zimniak et al., 2009). Preference of EB proteins for MT plus ends in cells is thus based on the fact that in vivo, MT minus ends never grow. In vitro experiments further showed that a monomeric EB3 fusion protein, containing a single CH domain and the linker region, is able to recognize growing MT ends in the absence of other þTIPs, proving that dimerization is not essential for þTIP behavior, although dimerization improves the affinity for the MT ends (Komarova et al., 2009). In contrast, dimerization might be essential for plus end tracking of Drosophila EB1 (Slep and Vale, 2007). Furthermore, it was shown that a monomeric Bim1 fragment containing the CH and the linker domain failed to plus end track in vitro, suggesting that dimerized CH domains and the presence of the linker are both necessary for its autonomous plus end tracking (Zimniak et al., 2009). Interestingly, phosphorylation of Bim1 linker by Aurora/Ipl1 removes Bim1 from MTs in vitro (Zimniak et al., 2009), supporting the importance of this region for the interaction with MTs. In vitro studies showed that Mal3 does not bind to tubulin dimers and its accumulation at MT tips is not dependent on tubulin concentration, indicating that it does not copolymerize with tubulin (Bieling et al., 2007).

Microtubule Plus End Tracking Proteins

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Single molecule experiments in vitro showed that the dwell time of individual Mal3 or EB1 molecules on the MT tip is short, typically less than a second, while MT tips are decorated by these proteins for several seconds, indicating that EB proteins exchange rapidly on the binding sites at the MT tips (Bieling et al., 2007, 2008; Dixit et al., 2009). Fluorescence recovery after photobleaching (FRAP) experiments in cells fully support the idea that EBs and at least some of their partners turn over rapidly at the MT ends (Dragestein et al., 2008); importantly signal recovery after FRAP is observed at both proximal and distal part of the þTIP comet, which is inconsistent with a copolymerization mechanism. If the EBs do not copolymerize with tubulin, then they must bind to the growing MT tips by recognizing some specific feature that is distinct from the rest of the MT lattice. The question is, what is this feature? An obvious hypothesis is the GTP cap. In support of this model, EB1 was found to preferentially recognize the GMPCPP MT lattice as opposed to the GDP lattice (Zanic et al., 2009). One argument against it is that the GTP cap is thought to be smaller than the region decorated by EB comets, which encompass hundreds of tubulin subunits (Bieling et al., 2007; Schwan et al., 2009; Wittmann, 2008). In order to explain EB distribution, the GTP cap should decay exponentially. Recent observations, both in vivo and in vitro, support the existence of a dynamic and spatially extended GTP cap (Dimitrov et al., 2008; Schek et al., 2007). Alternatively, EBs might bind to tubulin sheets, individual protofilaments, or hidden tubulin sites at the MT tip. This idea is supported by a recent EM study, which showed that the yeast EB homologue Mal3 specifically binds to the MT seam, suggesting that EB proteins play a role in the sheet closure by a zippering activity (Sandblad et al., 2006). In addition, it was proposed that EB proteins might bind preferentially to the edges of the tubulin sheet (Vitre et al., 2008), or change the structure of the MT lattice, by supporting preferential formation of an A lattice that has seam-like contacts between protofilaments (des Georges et al., 2008). Taken together, these results show a close relationship between EB proteins and regulation of MT structure. It should be noted that in cases where it has been investigated in detail by electron tomography, tubulin sheets or protofilament extensions at the MT plus end are typically quite short, shorter that 0.5 mm (Hoog et al., 2007; McIntosh et al., 2008; Zovko et al., 2008). It thus remains unclear if tubulin sheets or any other structural deviations at the MT ends can extend for several micrometers and explain size and the exponential profile of EB comets. 5.1.3. EB proteins in regulation of MT dynamics EB proteins localize to all cellular MTs with a higher concentration at the growing plus ends. Moreover, they bind to centrosomes and the midbody (Berrueta et al., 1998; Mimori-Kiyosue et al., 2000; Morrison et al., 1998).

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In plants, AtEB1a and AtEB1b were observed to behave as þTIPs, localizing to MT plus ends. However, AtEB1c differs from the other homologues, as it localizes to the nucleus in interphase and at the preprophase band, spindle and phragmoplast in mitosis (Chan et al., 2003; Mathur et al., 2003; Van Damme et al., 2004a; Komaki et al., 2010). It is now clear that EB proteins share a common role in regulating MT dynamics in vitro and in vivo, but their precise influence on each parameter of dynamic instability is still controversial. First, in budding yeast and Drosophila cells, EBs have been shown to stimulate both catastrophes and rescues, making MTs more dynamic and suppressing pauses (Rogers et al., 2002; Tirnauer et al., 1999). In agreement, additional experiments in S. cerevisiae reported similar changes in MT dynamics, but only in G1 cells and not in pre-anaphase cells (Wolyniak et al., 2006). This result suggests that plus end localization per se is not sufficient to stimulate dynamics, since Bim1 localizes to MT plus ends throughout the cell cycle. Along similar lines, depletion of EB1 in mouse fibroblasts promotes MT pausing and decreases the time MTs spent in growth (Kita et al., 2006). It should be noted, however, that another study in mammalian cells implicated EB1 in MT stabilization at the cortex (Wen et al., 2004). In Xenopus extracts, EB1 stimulates MT polymerization, promotes MT rescues, and inhibits catastrophes (Tirnauer et al., 2002b). In plants, MT stabilization was observed by overexpression of AtEB1a-GFP, but not by AtEB1b-GFP (Chan et al., 2003; Van Damme et al., 2004a). Experiments with the fission yeast homologue of EB1, Mal3, showed that it inhibits catastrophes and stimulates the initiation of MT growth (Busch and Brunner, 2004). In line with these results, simultaneous inactivation of different EB species in mammalian cells showed that EB1 and EB3 promote persistent MT growth by suppressing catastrophes, with little effect on MT growth rate or rescues (Komarova et al., 2009). In agreement with catastrophe suppressing function in vivo, in vitro EB binding is likely to coincide with the stabilization of growing tubulin sheets and/or the lattice seam, and may possibly exert an overall effect on MT lattice structure (des Georges et al., 2008; Sandblad et al., 2006; Vitre et al., 2008). Mal3 was shown to strongly promote MT assembly in vitro into predominantly 13-protofilament MTs with a high proportion of A lattice (des Georges et al., 2008). Preference for assembly of a 13-protofilament lattice was also observed for EB1 (Vitre et al., 2008). Another in vitro study provided evidence that Mal3 stabilizes MTs not by preventing catastrophes at the MT tip but by inhibiting lattice depolymerization and enhancing rescues (Katsuki et al., 2009). These findings suggest different modes of Mal3 binding to MT tips and the lattice. Suppression of shortening rate and the extent of shortening were also observed in vitro with purified EB1; however, in this study, catastrophe frequency was decreased while rescue frequency and MT growth rate were unaffected (Manna et al., 2008). It should be

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noted that it is uncertain if EB homologues were tracking MT plus ends in the assay conditions used. Importantly, in cells all EB proteins specifically accumulate at growing MT ends. Surprisingly, even in well-defined conditions when EB proteins track the growing ends in vitro, a broad variety of effects on MT dynamics have been reported, ranging from a complete absence of visible effects (Dixit et al., 2009) to strong enhancement of MT polymerization velocity, catastrophes and rescues (Komarova et al., 2009), or suppression of catastrophes (Blake-Hodek et al., 2010). The discrepancies between in vitro and in vivo results for the same EB species could be explained by the fact that EB proteins in cells might counteract the function of MT destabilizing factors or cooperate with MT stabilizers, while in vitro only their intrinsic properties are measured. The divergent in vitro effects of different EB proteins are most likely due to variability in protein preparations, salt concentration, tubulin sources, and other features of MT polymerization assays used in different laboratories. It is possible that EB proteins strongly affect MT tip structure, making it more sensitive to variations in assay conditions. 5.1.4. EBs in mitosis Interestingly, EB1 binds to MT plus ends at the interface between kinetochores and polymerizing MTs, suggesting that it could modulate their dynamic behavior during mitosis (Mimori-Kiyosue and Tsukita, 2003; Tirnauer et al., 2002a). Supporting this idea, deletion of Mal3 and Bim1 in yeast leads to aberrant spindles, lagging chromosomes, and chromosome missegregation (Beinhauer et al., 1997; Goldstone et al., 2010; Schwartz et al., 1997). Depletion of EB1 by RNAi from Drosophila cells as well as a null mutant of Dictyostelium EB1 (DdEB1) confirmed that EB proteins are required for proper spindle formation during mitosis (Rehberg and Graf, 2002; Rogers et al., 2002). Misalignment of spindle and phragmoplast MTs and chromosome separation defects were also described in cells with reduced levels of Arabidopsis AtEb1c (Komaki et al., 2010). In Drosophila cells, it was shown that Ncd, a minus end-directed motor, binds to EB1 and accumulates at growing MT plus ends in an EB1-dependent manner; this localization is important for Ncd-mediated focusing of kinetochore fibers into spindle poles (Goshima et al., 2005). In mammalian cells, the mitotic function of the EBs may at least partly depend on the interaction with APC: siRNA-mediated inhibition of APC, EB1, or APC and EB1 together gave rise to similar defects in mitotic spindles and chromosome alignment without arresting cells in mitosis (Green et al., 2005). Further, Aurora B, an important mitotic kinase and chromosome passenger protein, binds to EB1 in vivo and in vitro (Sun et al., 2008). This interaction promotes the kinase activity of Aurora B through blocking its inactivation by the phosphatase PP2A (Sun et al., 2008). While EB1 appears to regulate Aurora B, EB3 is a target of Aurora A and B kinases: phosphorylation causes EB3 stabilization

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during mitosis (Ban et al., 2009). Physical and functional interaction between EB1 and Aurora B is highly conserved, as their budding yeast homologues, Bim1 and Ipl1p bind to each other (Zimniak et al., 2009). Bim1 stabilizes the anaphase spindle (Gardner et al., 2008a). Ipl1 phosphorylates Bim1 and reduces its accumulation on MTs, a process needed for normal spindle elongation and proper disassembly of the spindle midzone (Zimniak et al., 2009). Bim1 also participates in controlling spindle orientation. It directly interacts with Kar9, which in its turn binds to myosin Myo2 that transports MTs into the bud along actin cables (Korinek et al., 2000; Lee et al., 2000; Liakopoulos et al., 2003). In HeLa cells, EB1 plays a role in spindle positioning by stabilizing astral MTs and affecting their interaction with the cortex (Toyoshima and Nishida, 2007). Also in dividing sea urchin eggs, injection of antibodies against EB1 and p150Glued showed that the interaction between EB1 and dynactin promotes elongation of astral MTs at the anaphase onset, suggesting a function in communication between the mitotic spindle and the cell cortex and a role in cytokinesis (Strickland et al., 2005). Astral MTs and spindle positioning were also affected after EB1 depletion in Drosophila cells (Rogers et al., 2002). Taken together, these data indicate that EBs are intimately involved in multiple aspects of MT function during mitosis. 5.1.5. EBs at the centrosomes and in cilia EB proteins also prominently localize to the centrosomes. Overexpression of a dominant negative EB1 mutant revealed a severe defect in MT anchorage at the centrosome (Askham et al., 2000). Later, a more detailed analysis indicated that EB1 localizes to centrosomes via its C-terminal region independently of MTs (Louie et al., 2004). It was also reported that EB1 preferentially localizes to the mother centriole forming a cap at its end. Depletion of EB1 by RNAi reduced MT minus end anchoring at centrosomes and delayed MT regrowth from centrosomes. In addition, EB1 localization at the centriole/basal body is required for primary cilia assembly in fibroblasts (Schroder et al., 2007). Interestingly, the function of EB proteins in flagella assembly may not be restricted to basal bodies: in Chlamydomonas, EB1 also localizes to the flagellar tip, and its mutation causes accumulation of intraflagellar transport particles near the tip, suggesting a role in switching between anterograde and retrograde transport (Pedersen et al., 2003). At the centrosome, EB1 cooperates with a centrosomal complex containing CAP350 and FOP (FGFR1 oncogene partner), which is needed for MT anchoring (Yan et al., 2006). FOP binds to EB1 and is required for centrosomal localization of EB1 (Yan et al., 2006). Depletion of CAP350, FOP, or EB1 by siRNA perturbed MT anchoring and caused disorganization of the MT network. Moreover, another important centrosomal protein, CDK5RAP2/Cep215 binds to EB1, and the CDK5RAP2–EB1

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complex stimulates MT assembly and MT bundle formation in vitro (Fong et al., 2009). Other proteins implicated in EB1-dependent centrosomal MT organization, are APC and p150Glued (see below). The potential cross talk between different EB1 partners at the centrosome and their redundant and nonredundant properties require further elucidation. 5.1.6. EBs in other cellular structures þTIPs are in a perfect position to mediate interactions between MTs and other structures such as the cell cortex. EB proteins have multiple binding partners that are known for their ability to link MTs to the cell cortex. MACF/ACF7 and Short Stop are members of the spectraplakin family that through their binding to EB1 reinforce links between F-actin and MTs, thus acting as integrators of MT-actin dynamics (Applewhite et al., 2010; Kodama et al., 2003). EB1, APC, and the formin family protein mDia were proposed to form a triple complex, which stabilizes MTs at the leading cell edges and promotes cell migration (Wen et al., 2004). The CLASPs bind to EB1 and regulate MT plus end dynamics at the cell cortex acting as local rescue factors (Mimori-Kiyosue et al., 2005). EB1-positive MT tips make contacts with adherens junctions and might play a role in directional delivery of proteins such as connexins to these sites (Shaw et al., 2007; Smyth et al., 2010). Also Drosophila RhoGEF2 accumulates at the tips of growing MTs in an EB1-dependent manner and regulates cortical actomyosin contraction leading to morphogenic movements during development (Rogers et al., 2004). Melanophilin and myosin Va form a triple complex with EB1 at MT plus ends, suggesting a role in the transfer of melanosomes from MTs to actin at the MT plus ends (Wu et al., 2005). More recently, it was shown that EB1 links ER tubules to MT tips through targeting STIM1, a transmembrane protein, to the growing MT plus ends; this mechanism promotes endoplasmic reticulum (ER) remodeling by growing MTs (Grigoriev et al., 2008). In Arabidopsis, AtEB1a and AtEB1b localize to MT nucleation sites in cortical MT arrays and to endomembrane system, respectively (Chan et al., 2003, 2005; Mathur et al., 2003). Thus, several lines of evidence show that EB proteins together with their partners can form crucial links between MT plus ends and cellular membranes (Lansbergen and Akhmanova, 2006; Mimori-Kiyosue and Tsukita, 2003; Vaughan, 2005). Since the EBs appear to form the core of þTIP interaction networks, they can be ultimately expected to participate in the majority of MT-membrane connections that involve growing MT tips. Studies of EB protein involvement in cell differentiation and morphogenesis in higher organisms have been limited. In plants, AtEB1 proteins were found to colocalize with MTs in roots, and EB1 mutants displayed root growth defects: mutant roots exhibit leftward deviation during growth, probably due to abnormalities in cortical MT arrays (Bisgrove et al., 2008). However, these findings found no support in a more recent study, which

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reported no root defects in Arabidopsis plants lacking all three EB1 family members but showed that they exhibited an increased sensitivity to MT-active herbicide oryzalin (Komaki et al., 2010). In Drosophila, a hypomorphic mutant in the EB1 genes exhibited neuromuscular defects, including inability to fly and uncoordinated movement, probably due to malfunction of the chordotonal mechanosensory organs (Elliott et al., 2005). In mammals, addressing the functions of the EBs in embryonic development and tissue maintenance awaits generation of mouse knockout models.

5.2. CAP-Gly proteins The cytoskeleton-associated protein-glycine-rich (CAP-Gly) domain is a specialized protein module of 80 residues that is highly conserved in eukaryotes (Riehemann and Sorg, 1993). CAP-Gly proteins include cytoplasmic linker proteins (CLIPs) and the large subunit of the dynactin complex (p150Glued). CAP-Gly domains are found in single or multiple copies and are primarily involved in protein–protein interactions, in particular with tubulin monomers, dimers, MTs, and EB proteins (Steinmetz and Akhmanova, 2008). The crystal structures of several CAP-Gly domains have been solved (Honnappa et al., 2006; Li et al., 2002; Mishima et al., 2007; Weisbrich et al., 2007) and provide insight into their function. It was demonstrated that the conserved GKNDG motif of CAP-Gly domains is responsible for targeting to the C-terminal EEY/F sequence motifs of CLIP-170 itself, EB proteins and a tubulin (Steinmetz and Akhmanova, 2008). The GKNDG motif is located next to a cluster of conserved aromatic residues, which are packed against each other forming a hydrophobic cavity that serves as a binding site for the EEY/F sequence (Steinmetz and Akhmanova, 2008). In most eukaryotic cells, the C-terminal residue of a tubulin is aromatic, a tyrosine (Tyr) in mammals and a phenylalanine (Phe) in S. cerevisiae. In a tubulin from higher eukaryotes it can be removed and added back; its readdition is catalyzed by the enzyme tubulin-tyrosine ligase (TTL) (Verhey and Gaertig, 2007; Westermann and Weber, 2003). Recent studies have demonstrated that this Tyr is an important factor affecting the recruitment of CAP-Gly proteins to MT plus ends (Badin-Larcon et al., 2004; Erck et al., 2005; Weisbrich et al., 2007). It was shown that CAP-Gly domains bind more efficiently to tyrosinated MTs compared to detyrosinated ones, suggesting that tubulin tyrosination can in principle regulate MT interactions with CAP-Gly proteins (Bieling et al., 2008; Erck et al., 2005; Peris et al., 2006). Importantly, the binding sites of CAP-Gly domains are not confined to EEY/F motifs, and additional binding surfaces of their partners, such as the EBH domain of EB1, zinc finger motifs of CLIP-170, or helices of a and b tubulin (Gupta et al., 2010; Honnappa et al., 2006; Weisbrich et al., 2007) are involved in the interactions.

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5.2.1. CLIPs 5.2.1.1. Structure and evolutionary conservation CLIP-170 was the first protein revealing the interaction of CAP-Gly domains and MTs (Pierre et al., 1992); it was also the first protein for which plus end tracking behavior was described (Perez et al., 1999). It was independently cloned as a protein highly expressed in the Reed–Sternberg cells of the Hodgkin’s disease and named restin (Bilbe et al., 1992); in this study, the pattern formed by the overexpressed protein was interpreted as intermediate filaments, but later studies proved that it localizes to MTs. Localization of the CLIPs to MTs and MT plus ends depends on CAP-Gly domains (Gupta et al., 2009a; Honnappa et al., 2006; Mishima et al., 2007; Steinmetz and Akhmanova, 2008). CAP-Gly domains are surrounded by regions rich in serines and basic residues, which contribute to interactions with MTs (Gupta et al., 2009a; Hoogenraad et al., 2000). CLIP-170 forms a parallel dimer and at its N-terminus each monomeric subunit contains two similar CAP-Gly domains, scoring four in total (Scheel et al., 1999) (Fig. 1.2). At the C-terminus, CLIP-170 contains two metal binding domains (the zinc fingers) and the C-terminal EEY/ F-like sequence. These domains are important for CLIP-170 autoregulation, because the CAP-Gly domains at the N-terminus of CLIP-170 can bind to the zinc fingers at the C-terminus creating a donut-shaped molecule (Lansbergen et al., 2004; Fig. 1.2). This conformational change of CLIP170 negatively regulates its interaction with MTs. Multiple splice isoforms of CLIP-170 exist in different tissues; these predominantly cause variability in the central coiled coil region (Akhmanova et al., 2005; Griparic et al., 1998), but a testis-specific isoform encoding only the C-terminal tail of the

CLIP-170 Self-inhibition CAP-Gly domains S

S

Zinc fingers

S

Microtubules EBs IQGAP1

EEY/F-like motif

Coiled coil

CLASPs

Dynactin, LIS1

Figure 1.2 Schematic representation of CLIP-170 protein sequence. S, basic and serine-rich regions surrounding CAP-Gly domains. Autoinhibitory interaction between the N- and C-terminus of CLIP-170 is indicated by a two-sided arrow. Interaction sites with MTs, IQGAP1 and þ TIP partners are indicated.

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protein has also been described (Tarsounas et al., 2001). CLIP-115, a brain specific CLIP, is very similar to the N-terminal part of CLIP-170 but lacks the C-terminal region, zinc fingers included (De Zeeuw et al., 1997). CLIP homologues are present in yeast (Bik1 in S. cerevisiae (Berlin et al., 1990) and Tip1 in S. pombe (Brunner and Nurse, 2000)). Both proteins are dimers with a single CAP-Gly domain and a single zinc finger motif present in each monomeric subunit (Miller et al., 2006). CLIP homologues also exist in invertebrates, such as CLIP-190 in flies (Lantz and Miller, 1998); in contrast, higher plants lack CLIPs. This might be due to the fact that proteins of this family primarily act as accessory factors of the cytoplasmic dynein pathway, which is absent from plant cells (Wickstead and Gull, 2007). 5.2.1.2. Mechanisms of plus end tracking by CLIPs Plus end tracking of CLIP homologues was investigated in detail for yeast and vertebrate proteins. Although biochemical studies suggested copolymerization with tubulin dimers (Folker et al., 2005), FRAP experiments indicate that CLIP-170 exchanges fast at the growing MT ends, similar to EB1 (Dragestein et al., 2008). In vitro reconstitution also showed that CLIP-170 exchanges rapidly at MT ends (Bieling et al., 2008). Importantly, it requires intact EB1 and tubulin EEY/F motifs for efficient MT plus end accumulation indicating that it recognizes composite binding sites at the MT tips (Bieling et al., 2008; Komarova et al., 2005; Mishima et al., 2007; Peris et al., 2006). This might explain why the residence time of CLIP-170 at the MT ends is longer than that of EB1 (Bieling et al., 2008; Dixit et al., 2009). Whether CLIP170 uses different CAP-Gly domains to interact with EB1 and tubulin is unclear; studies in cells showed that a fragment consisting of the second CAP-Gly domain with its adjacent serine-rich region is sufficient to track MT plus ends (Gupta et al., 2009b). Also in budding yeast, Bik1 requires Bim1 and the C-terminal aromatic residue of a tubulin for efficient MT tip accumulation (Badin-Larcon et al., 2004; Blake-Hodek et al., 2010; Caudron et al., 2008). In fission yeast, MT plus end tracking of Tip1 depends on Mal3 (Busch and Brunner, 2004). Importantly, in both budding and fission yeast, plus end-directed kinesins Tea2/Kip2 are involved in the delivery of CLIP homologues to MT plus ends, a phenomenon that has been reconstituted in vitro using purified proteins (Bieling et al., 2007; Busch et al., 2004; Carvalho et al., 2004). In contrast, no kinesin-based delivery of CLIPs has been observed in animal cells. 5.2.1.3. CLIPs in regulation of MT dynamics The CLIP-170 family members are clearly involved in MT stabilization; they have been described acting as anticatastrophe factors in fission yeast (Brunner and Nurse, 2000) and rescue factors in mammalian cells (Komarova et al., 2002). In fission yeast, Tip1 has been shown to prevent MT catastrophes at the cell cortex

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(Brunner and Nurse, 2000). If Tip1 degradation pathway is perturbed, MTs do not undergo catastrophes at the cell tips but continue to grow and curve around the cell end (Martin-Garcia and Mulvihill, 2009); Tip1 is thus one of the major fission yeast factors shaping the interphase MT network in fission yeast (Brunner and Nurse, 2000; Daga et al., 2006a,b). Bik1 in budding yeast also has a role in MT dynamics since null mutants are viable but contain very short MTs (Berlin et al., 1990). Furthermore, CLIPs were implicated in MT nucleation both in vitro and in cells (Arnal et al., 2004; Rogers et al., 2008; Slep and Vale, 2007). Early in vitro studies suggested that the N-terminal region of CLIP-170 promotes rescues (Arnal et al., 2004). More recently, EB-dependent MT plus end tracking behavior was reconstituted in vitro for a number of CLIP homologues (Bieling et al., 2007, 2008; Blake-Hodek et al., 2010; Dixit et al., 2009). These studies revealed that CLIPs directly interact with the EBs and tubulin, and thus can potentially affect both EBs and MTs by their accumulation at the MT ends. Measurements of MT dynamics parameters yielded variable results, ranging from a lack of effect for CLIP-170 (Dixit et al., 2009) to catastrophe promoting activity for Bik1 (Blake-Hodek et al., 2010). Similar to the EBs, additional studies and more complex reconstitution experiments will be needed to get a full picture of how these proteins control MT dynamics. Importantly, a rescue activity, promotion of transitions from shrinking to growing ends, seems to be paradoxal for a þTIP like CLIP-170 because it implies a localization at depolymerizing MT ends, which has not been reported for CLIPs in mammalian cells. The situation is different in budding yeast, where Bik1 follows not only growing but also shrinking MTs (Carvalho et al., 2004). A possible explanation for the MT rescue activity of CLIP-170 could be the stabilization of curved protofilaments observed by the MT binding domain of CLIP-170 in vitro (Arnal et al., 2004). Alternatively, small amounts of CLIP-170 could bind to GTP remnants (Dimitrov et al., 2008) or other lattice defects and promote MT rescue once a depolymerization event reaches this point. 5.2.1.4. CLIPs in the dynein pathway Through its C-terminal zinc finger-containing region, CLIP-170 directly interacts with two major players in the cytoplasmic dynein pathway, the dynactin large subunit p150Glued and lissencephaly 1 (LIS1); these interactions are mutually exclusive (Coquelle et al., 2002; Lansbergen et al., 2004). In mammalian cells, these interactions are important for recruitment of these two proteins as well as cytoplasmic dynein to MT tips (Coquelle et al., 2002; Lansbergen et al., 2004; Vaughan et al., 1999; Watson and Stephens, 2006). In budding yeast, the CLIP-170 homologue Bik1 also participates in the dynein pathway; its interaction with the LIS1 homologue Pac1 is important for this function (Huisman and Segal, 2005); however, Bik1-dependent and -independent pathways for dynein localization at MT tips appear to exist (Caudron et al., 2008).

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It was proposed that dynein–dynactin accumulation at the plus ends can promote loading of cargo for minus end-directed transport (Vaughan et al., 2002). Interestingly, in Xenopus melanophores CLIP-170 directly participates in loading dynein cargo, melanosomes, on MT plus ends during stimulus-induced melanosome aggregation (Lomakin et al., 2009). The originally proposed function of CLIP-170, to link endosomes to MTs in mammalian cells (Pierre et al., 1992), has thus found unexpected support, albeit in a different cell system. 5.2.1.5. CLIPs in mitosis Mitotic functions of CLIP homologues seem to be related to their interactions with dynein cofactors. Budding yeast Bik1 plays an important role in dynein-mediated spindle positioning pathway (Huisman and Segal, 2005; Miller et al., 2006). In mitotic mammalian cells, CLIP-170 localizes to MT plus ends and accumulates strongly at kinetochores in prometaphase, but only weakly in metaphase, similar to dynein and dynactin (Dujardin et al., 1998; Lin et al., 2001; Tai et al., 2002). CLIP170 interaction with kinetochores is independent of its CAP-Gly domains, relies on its C-terminal region (Dujardin et al., 1998) and is mediated by dynein–dynactin and LIS1 (Coquelle et al., 2002; Faulkner et al., 2000; Tai et al., 2002). A similar CLIP localization pathway also exists in fly cells (Dzhindzhev et al., 2005). CLIP-170 preferentially localizes to kinetochores that lack bound MTs and is involved in the formation and stabilization of kinetochore-MT attachments (Tanenbaum et al., 2006). The importance of this pathway might vary between cell types, because although it is essential in HeLa cells (Tanenbaum et al., 2006; Wieland et al., 2004), no significant mitotic defects were found in CLIP-170 knockout mice or cultured mouse embryonic fibroblasts (Akhmanova et al., 2005), suggesting that additional redundant pathways for kinetochore-MT attachment exist in mammals. In fission yeast, absence of Tip1 affects chromosome congression into the metaphase plate and causes appearance of lagging chromosomes; however, Tip1 and its partner kinesin Tea2 are difficult to detect in the spindle and the mechanisms underlying their mitotic activities remain unclear (Goldstone et al., 2010). 5.2.1.6. CLIPs in other cellular pathways Several studies linked the CLIPs to MT-actin cross talk. CLIP-170 binds to IQGAP1 and through this interaction seems to target special cortical sites marked by Rac1/Cdc42, enabling MT capture at the leading edge (Fukata et al., 2002). Further, it participates in controlling actin nucleation by the formin mDia1 during phagocytosis in macrophages (Lewkowicz et al., 2008). Also, fission yeast CLIP homologue Tip1 participates in actin organization at the cell ends by acting in a complex with MT end-associated factors Tea1, Tea4, and a formin For3 (Chang et al., 2005; Martin et al., 2005). Physical and functional links between the CLIPs and myosin motors have also been

Microtubule Plus End Tracking Proteins

23

described: the Drosophila, CLIP-190 interacts with myosin VI in neurons and fly embryos (Lantz and Miller, 1998), while myosin V Myo52 controls Tip1 stability through an ubiquitination pathway in fission yeast (Martin-Garcia and Mulvihill, 2009). CLIP-170 has long been known to be a phosphoprotein (Rickard and Kreis, 1991). Both Bik1 and CLIP-170 can be phosphorylated by TOR and were suggested to organize MTs and MT-based processes downstream of this kinase (Choi et al., 2000, 2002; Jiang and Yeung, 2006). Recently, Cdc2 was suggested to control CLIP-170 function in centrosome reduplication (Yang et al., 2009). Cell cycle-mediated regulation of CLIP phosphorylation is likely to be evolutionary conserved, as CLIP-190 accumulation at MT tips is strongly attenuated in mitotic Drosophila cells (Dzhindzhev et al., 2005). An important regulatory region in CLIP-170 is the third serine-rich stretch, which can be phosphorylated by protein kinase A (PKA) and AMPactivated protein kinase (AMPK) (Lee et al., 2010; Nakano et al., 2010). Phosphorylation in this region can inhibit the interaction with MTs by two mechanisms: directly, through reducing electrostatic interaction of CLIP170 with negatively charged MTs, and indirectly, by promoting self-folding of the protein (Lee et al., 2010). The downstream consequences of CLIP170 phosphorylation require further elucidation but might include effects on directional cell migration through regulation of focal adhesions by dynamic MTs (Nakano et al., 2010). CLIPs are abundantly expressed in neurons and are likely to have some specific functions in these highly differentiated cells. This notion is supported by the fact that CLIP-115 haploinsufficiency was linked to neurodevelopmental features of Williams syndrome (Hoogenraad et al., 2002) and also by observation of MT dynamics defects in CLIP-115 and CLIP-170deficient cultured neurons (Stepanova et al., 2010). CLIP-170 also has a very specific role in spermatogenesis: during spermatid development, it stabilizes a highly specialized and transient MT structure involved in shaping the sperm nucleus, the manchette (Akhmanova et al., 2005). Importantly, in this case, the binding of CLIP-170 to MTs seems to be quite stable and appears not to involve tracking of growing MT ends. 5.2.2. Dynactin large subunit p150Glued Another þ TIP that has a CAP-Gly domain at its N-terminus is the already mentioned p150Glued, the large subunit of the dynactin complex (Holzbaur et al., 1991). The CAP-Gly domain is part of the MT binding domain of p150Glued, which also contains a basic and serine-rich region (Fig. 1.3). Two coiled coil domains are also present in the molecule and are required for dimerization and for interaction with other dynactin subunits and with the dynein intermediate chain (Schroer, 2004). Dynactin complex is essential for virtually all functions of cytoplasmic dynein, and there is a vast body of

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p150Glued

Basic/serine rich CAP-gly domain S

Coiled coil

Coiled coil

Microtubules EBs

Figure 1.3 Schematic representation of p150Glued protein sequence. Interaction sites with MTs and EB1 are indicated.

literature describing its structure and function. In this review, we will briefly summarize only the studies related to MT plus end binding properties of this protein complex. Like the CLIPs, p150Glued uses its CAP-Gly domain to bind to EB proteins, a tubulin, and CLIP-170. Interaction with the EBs is quite specific: p150Glued binds to EB1 and EB3, but not to EB2 (Bu and Su, 2003). The crystal structure of the complex of EB1 C-terminus with the CAP-Gly domain of p150Glued has shown that p150Glued binds to the flexible tail of EB1 via its EEY/F motif (Honnappa et al., 2006). In addition, complex formation involves residues shaping the hydrophobic cavity and the polar rim of EB1c, and the b2–b3 loop of the CAP-Gly domain of p150Glued and some adjacent areas of its N-terminal region (Honnappa et al., 2006). Another interaction important for p150Glued function is with CLIP-170, which is required for p150Glued recruitment to the growing MT plus ends (Lansbergen et al., 2004). This interaction also involves the binding of the CAP-Gly domain of p150Glued to an EEY/F motif, in this case the one at the C-terminus of CLIP-170 (Hayashi et al., 2007; Weisbrich et al., 2007). p150Glued can thus be recruited to the MT plus ends via two different pathways: through EB1 and through CLIP-170. Also, it should be emphasized that p150Glued contains a short basic sequence that is implicated in direct MT binding (Culver-Hanlon et al., 2006). The dynactin complex is required for targeting dynein to cargo and for dynein motor processivity (King and Schroer, 2000; Schroer, 2004). In vitro experiments have shown that while the CAP-Gly domain of p150Glued can inhibit dynein motility acting as a brake, the basic domain significantly increases dynein processivity (Culver-Hanlon et al., 2006). The length of the basic domain and MT lattice binding properties can vary between different splice isoforms of p150Glued (Dixit et al., 2008; Zhapparova et al., 2009). It should be noted that another p150Glued isoform, which is abundant in the brain (p135), lacks the MT binding domain altogether

Microtubule Plus End Tracking Proteins

25

(Tokito et al., 1996), and the MT binding domain of p150Glued is not required for dynein-dependent organelle movement (Dixit et al., 2008; Kim et al., 2007). In contrast, the MT binding capacity of dynactin is important for MT organization (Kim et al., 2007); in line with this view, p150Glued localizes to centrosomes where its interaction with EB1 is required for the formation and maintenance of a radial MT array (Askham et al., 2002; Quintyne et al., 1999). MT affinity of p150Glued can be negatively regulated by phosphorylation, and several kinases, including PKA (Vaughan et al., 2002) and Aurora A (Rome et al., 2010), have been shown to phosphorylate the MT binding domain of p150Glued in cells. In humans, a missense G59S point mutation in the CAP-Gly domain of p150Glued is implicated in an autosomal dominant form of a motor neuron disease (distal bulbar muscular atrophy) (Puls et al., 2003). This amino acid substitution in p150Glued impairs proper folding of the CAP-Gly domain, largely abrogates its binding to MTs and CLIP-170 and causes protein aggregation and defects in dynactin function (Levy et al., 2006), a conclusion that was confirmed by studies using genetically modified mice (Chevalier-Larsen et al., 2008; Lai et al., 2007; Laird et al., 2008). Another type of human neurological disease, Perry’s syndrome, is also associated with missense mutations in the p150Glued CAP-Gly domain (Farrer et al., 2009). These mutations do not cause misfolding but affect the stability of the CAP-Gly domain and inhibit its association with EB1 although not with MTs, suggesting that plus end tracking behavior of dynactin might be important for brain function in humans (Ahmed et al., 2010).

5.3. Proteins with basic and serine-rich regions Unstructured sequence regions enriched in serines and basic residues have been identified within a large group of diverse þTIPs. These regions, which are predicted to be flexible, mediate interactions with MTs and EB proteins (Akhmanova and Steinmetz, 2008) and often contain one or several copies of a short, conserved, and phosphorylation controllable motif, the Ser/Thr-XIle-Pro (SxIP), where X is any amino acid. This motif can target þTIPs to the growing MT ends in an EB1-dependent manner, a notion that is supported by both in vitro and in vivo experiments (Honnappa et al., 2009). It is currently unclear whether additional LMs can perform a similar function. Representatives of this group of proteins are APC, MACF and STIM1. Many of these proteins contain multiple conserved domains of different evolutionary origins. In addition, CLASPs and MCAK (mitotic centromere associated kinesin) also target MT ends by association with EB family members through SxIP motifs, but will be discussed in subsequent review sections, together with TOG domain and motor proteins, respectively. The hypothesis that the Ile–Pro dipeptide is involved in binding to the EB1 C-terminus has initially emerged based on its presence in the EB1

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binding sites of APC and spectraplakins (Honnappa et al., 2005; Slep et al., 2005). Curiously, indications for the importance of this site were provided by a crystallographic artifact: within the EB1c crystal, the Ile–Pro residues from one EB1c molecule made contact with hydrophobic cavity of another EB1c molecule (Honnappa et al., 2005). Since an Ile–Pro dipeptide was present in the core of the EB1-interacting fragment of APC, the authors hypothesized that it might be involved in the interaction, mutated it and proved that it was indeed the case (Honnappa et al., 2005). Subsequently, a combination of biochemical assays, live cell imaging, and in vitro reconstitution assays demonstrated that this motif is indeed used by numerous þTIPs for localization to MT tips in an EB-dependent manner (Honnappa et al., 2009). Structural studies, by both X-ray crystallography and NMR, revealed the details of the SxIP–EB1c complex, and confirmed the involvement of the hydrophobic cavity of EB1c in the interaction (Honnappa et al., 2009). Mutations within SxIP motif (such as substitution of isoleucine and proline for asparagines) abrogate EB binding and consequently plus end tracking. Although some SxIP-containing þTIPs, such as RhoGEF2 and melanophilin, do not bind to MTs directly, most other þTIPs do, and therefore, similar to CLIP-170, they are likely to recognize at MT tips composite binding sites that are shaped by EB1 C-terminus, tubulin, and other þTIPs. This might explain why the comet profiles of EB partners may differ from those of the EBs themselves (Wittmann and Waterman-Storer, 2005). It should be noted that þ TIPs that do not use SxIP motifs for MT tip binding, like CLIPs and EBs, as well as many unrelated MAPs also contain unstructured basic and serine-rich peptide stretches, making this type of sequence a very common feature involved in electrostatic association with MTs. 5.3.1. APC APC is a large multidomain tumor suppressor protein of  300 kDa that is conserved from Drosophila to humans (Mimori-Kiyosue and Tsukita, 2001). It has an important role in the regulation of the Wnt signaling, and its involvement in this pathway is the primary basis of its tumor suppressor properties (Aoki and Taketo, 2007). However, APC proteins shuttle between several subcellular destinations, where they participate in multiple processes (Bienz, 2002). These include cell migration and adhesion, spindle assembly, chromosome segregation, neuronal differentiation, apoptosis, and MT stabilization (Aoki and Taketo, 2007; Hanson and Miller, 2005; Nathke, 2004). APC localizes to the MT cytoskeleton and promotes MT polymerization and stabilization in vitro (Munemitsu et al., 1994; Nakamura et al., 2001; Zumbrunn et al., 2001). A second mammalian gene, APC2, encodes a protein that is predominantly expressed in the nervous system, and regulates MTs during neuronal development (Nakagawa et al., 2000; Shintani et al., 2009). Homologues of APC are present in invertebrates,

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such as flies, where APC is also encoded by two genes (Rusan et al., 2008), but not in plants or yeast. Because of its important role in tumorigenesis, APC is very intensively studied, and a vast literature on this topic has been subject to numerous excellent recent reviews (Aoki and Taketo, 2007; Bahmanyar et al., 2009; Hanson and Miller, 2005; Morrison, 2009; Nathke, 2004). Here, we will briefly focus only on the þTIP-related properties of APC. EB1 was initially identified in a yeast two-hybrid screen as a protein that interacts with the C-terminus of APC (Su et al., 1995). Subsequent biochemical studies have mapped the EB1–APC interaction to a basic and serine-rich sequence of 39 residues in APC C-terminus, named APCp1, which can track growing MT ends in cells (Bu and Su, 2003; Honnappa et al., 2005; Fig. 1.4). This interaction is mediated by the SxIP motif, which is anchored in the hydrophobic cavity of EB proteins (Honnappa et al., 2009). These results strongly suggest that APC “hitchhikes” on EB1 to plus end track growing MT ends. However, another study showed that APC association with MT plus ends can occur independently of its association with EB1 suggesting distinct mechanisms for APC and EB1 accumulation at MT plus ends (Kita et al., 2006). APC can be transported to MT tips by kinesin II ( Jimbo et al., 2002), and “backtracking” of APC on depolymerizing MT ends has also been reported (Langford et al., 2006). These observations suggest cooperation between different mechanisms, which APC uses to reach the MT tip. The interaction between EB1 and APC is likely to play a role in tumorigenesis of intestinal epithelial cells probably due to effects on cell polarity and/or cell division. During mitosis, APC localizes to kinetochores,

APC SxIP

SAMP repeats

Basic/serine rich

Armadillo repeats CC

S

CC 15 aa repeats

S

20 aa repeats

Kinesin-2

EB1 Microtubules

Figure 1.4 Schematic representation of APC protein sequence. CC, coiled coil region; SAMP, Ser-Ala-Met-Pro repeat. The scheme illustrates interaction domains with MTs, EB1 and kinesin-2, but not with the other numerous partners such as the components of the Wnt signaling pathway.

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centrosomes, and the cell cortex, and mutations in APC might contribute to mitotic abnormalities and chromosome instability through different pathways (Bahmanyar et al., 2009). In addition, the APC–EB1 interaction is thought to play a role in several MT capture and capping mechanisms at the cell cortex (Morrison, 2009). APC stabilizes MTs and stimulates their polymerization, and in association with actin-binding proteins like IQGAP1 and mDia it appears to link MT plus ends to the cortex (Watanabe et al., 2004; Wen et al., 2004). However, experiments with APC mutant MEFs lead to the conclusion that the APC–EB1 interaction is not essential for the regulation of MT stabilization (Drabek et al., 2006). Another pathway operating in migrating astrocytes involves APC-mediated attachment of MT tips to the plasma membrane in conjunction with Dlg1 (Etienne-Manneville et al., 2005). Importance of APC in MT stabilization in glia cells was supported by a mouse knockout study (Yokota et al., 2009). The functions of APC proteins in cell polarization were also addressed in flies, with somewhat contradictory results (see Rusan et al., 2008 and references therein). Muliple signaling pathways have been reported to regulate APC. For example, the EB1–APC interaction may be attenuated during mitosis due to APC phosphorylation by Cdc2 in the vicinity of SxIP motifs (Askham et al., 2000; Honnappa et al., 2005; Nakamura et al., 2001). Another important kinase directly involved in APC phosphorylation in the context of both Wnt signaling and MT-related functions is GSK3b (Zumbrunn et al., 2001). Like with most other MAPs, phosphorylation negatively affects the electrostatic component of APC interaction with EB1 and MTs, and thus might be a means for spatial and/or temporal restriction of MT-stabilizing properties of APC. 5.3.2. Kar9 Kar9 is a budding yeast linker protein, which plays an important role in mitotic spindle positioning. It contains a central coiled coil region, an acidic N-terminal region, and a basic and proline-rich C-terminal part (Miller and Rose, 1998). It is often referred to as the counterpart of the mammalian APC (which is the reason why it is discussed in this section), but in fact the sequence similarity between Kar9 and APC is very limited. Kar9 localizes to MT tips through the interaction with the C-terminus of Bim1, an EB1 homologue (Miller et al., 2000), but the molecular basis of this interaction remains to be determined. In addition, Kar9 also interacts with two other þTIPs, Bik1 (CLIP-170 homologue), and Stu2 (XMAP215 homologue) (Miller et al., 2000; Moore et al., 2006). Importantly, Kar9 can also interact with the myosin V Myo2, which promotes the interaction of Kar9-associated astral MTs with actin, a process important for the spindle positioning in the bud neck of the dividing yeast cell (Beach et al., 2000; Miller et al., 2000; Yin et al., 2000).

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Similar to other yeast þTIPs, Kar9 is transported to the MT plus ends by kinesin-7 Kip2 (Liakopoulos et al., 2003; Maekawa et al., 2003; Moore et al., 2006). It associates with both growing and shrinking MT ends, and photoconversion experiments show that its dissociation from MTs is slow (Kammerer et al., 2010). The transport is initiated at the spindle pole body (SPB), and in fact, Kar9 associates with only one of the two SPB, an asymmetry important for the proper spindle orientation (Liakopoulos et al., 2003). Posttranslational modifications of Kar9, such as phosphorylation, sumoylation, and ubiquitylation, have been studied in detail (Kammerer et al., 2010; Leisner et al., 2008; Meednu et al., 2008; Moore and Miller, 2007). Asymmetric localization of Kar9 is dependent on its phosphorylation by the yeast cyclin dependent kinase Cdc28 (see Kammerer et al., 2010; Moore and Miller, 2007 and references therein). Kar9 degradation and thus the association of astral MTs with the bud neck are controlled by ubiquitylation (Kammerer et al., 2010). 5.3.3. Spectraplakins Spectraplakins are represented in mammals by two paralogues, ACF7/ MACF1 and dystonin/BPAG1/MACF2. These are huge multidomain proteins that can associate with both actin filaments and MTs and are related to plakin and spectrin families (Leung et al., 1999; Roper et al., 2002). The N-terminal region of spectraplakins typically contains a calponin type actinbinding domain and a plakin domain, a central rod composed of dystrophin-like spectrin repeats, a EF-hand motif, a MT binding GAS2 domain and a basic and serine-rich C-terminus (Sun et al., 2001) (Fig. 1.5). In addition, ACF7/MACF1 was recently reported to contain an ATPase

MACF/ACF7 Calponin homology Plakin domains domain

Spectrin repeats CC

Actin

SxIP EF-hand GAS2 S

Basic/serine rich

Microtubules EBs

Figure 1.5 Schematic representation of MACF/ACF7 (spectraplakin) protein sequence. CC, coiled coil region. EF-hand, calcium-binding motif; GAS2, growtharrest-specific protein 2. The scheme illustrates interaction domains with MTs, EB1, and actin filaments.

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domain (Wu et al., 2008). A single spectraplakin gene, Short Stop (Shot), is present in Drosophila (Roper et al., 2002). Both Drosophila and mammalian spectraplakins directly interact with EB1 (Honnappa et al., 2009; Slep et al., 2005) and track growing MT ends in cells (Kodama et al., 2003; Slep et al., 2005). A small fragment of 43 residues (MACF43) derived from the unstructured C-terminal region of MACF2 contains an SxIP motif, interacts with EB1 and tracks growing MT ends in vivo and in vitro (Honnappa et al., 2009). Remarkably, robust plus end tracking was still observed with a dimer of a 12 residues MACF2 peptide encompassing the SxIP motif. X-ray crystallography of a complex of the MACF2 C-terminal peptide with EB1 C-terminal domain confirmed the importance of the SxIP motif for EB1 binding by showing that the most prominent contacts involve the Ser, Ile, and Pro residues (Honnappa et al., 2009). Subsequently, a detailed study of the fly homologue Shot also identified two SxIP motifs within the C-terminal part of this protein involved in EB1 binding and MT plus end tracking (Applewhite et al., 2010). The function of spectraplakins was extensively investigated in flies and mammals. Shot was implicated in epithelial cell adhesion (Roper and Brown, 2003), as well as in MT organization and oocyte specification during oogenesis (Roper and Brown, 2004). By acting together with EB1 and APC1, Shot was also shown to participate in MT organization at the muscle–tendon junction (Subramanian et al., 2003). A more recent analysis revealed that the CH and plakin domains of Shot and its MT-actin crosslinking function are essential in neurons but not in tendon cells (Bottenberg et al., 2009). A direct role of spectraplakins in MT-actin coupling was demonstrated in cultured cells (Applewhite et al., 2010; Kodama et al., 2003). The function of dystonin/BPAG1/MACF2 has been extensively studied: it is required for proper cytoskeletal organization and maintenance of sensory neurons in mice (Sonnenberg and Liem, 2007). ACF7/MACF1 is essential for mouse embryonic development (Kodama et al., 2003), and conditional targeting of this gene in the mouse epidermis causes abnormalities in wound repair and cell migration (Wu et al., 2008). Interestingly, MACF1 deficiency appears to impair the targeting of MTs to focal adhesions and focal adhesion turnover (Wu et al., 2008). Moreover, attempts to rescue the defects in focal adhesion dynamics and cell migration with the individual MACF domains that bind to F-actin, MTs, or EB1 failed, indicating the importance of MACF function as a cytoskeletal crosslinker (Wu et al., 2008). ACF7/MACF1 also plays a role in MT stabilization, which occurs specifically at the leading edge of migrating cells (Kodama et al., 2003). In this process, it cooperates with CLASPs (see below) (Drabek et al., 2006). Furthermore, similar to Shot, ACF7/MACF1 is required for proper organization of neuronal MTs and axon extension (Sanchez-Soriano et al., 2009).

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5.3.4. STIM1 STIM1 is a multidomain transmembrane protein that is essential for storeoperated Ca2þ entry, a process of extracellular Ca2þ influx in response to the depletion of Ca2þ stores in the ER (Wu et al., 2007). The STIM1 N-terminal region is located in the ER lumen and contains a SAM domain (sterile a motif domain, a protein–protein interaction module) and an EF-hand motif (calcium-binding motif) (Cahalan, 2009; Fig. 1.6). In the middle of the protein, there is a transmembrane domain, which is followed by a cytoplasmic C-terminal region, including a coiled coil, an ERM domain (ezrin–radixin– moesin) and a basic/serine/proline region (Cahalan, 2009). The essential function of STIM1 is to act as a calcium sensor in the ER lumen. When ER Ca2þ stores are intact, STIM1 is localized diffusely in the ER membrane. Depletion of Ca2þ from the ER induces STIM1 oligomerization and triggers translocation of STIM1 to the plasma membrane and activation of Ca2þ release-activated Ca2þ channels (Cahalan, 2009). This function of STIM1 is extensively studied and is beyond the scope of this review. Besides store-operated Ca2þ entry, STIM1 is also involved in MTdependent ER remodeling (Grigoriev et al., 2008). Through the SxIP motif located within its cytoplasmic basic and serine-rich region, STIM1 binds to EB1 and tracks MT ends at sites where growing MT tips contact the ER membrane (Grigoriev et al., 2008). Mutations in the SxIP motif of STIM1 abrogate both binding to EB1 and MT tip tracking (Honnappa et al., 2009). An in vitro reconstitution showed that a GFP-tagged Cterminal fragment of STIM1 encompassing the SxIP motif is able to track growing MT ends in the presence of EB1 (Honnappa et al., 2009). STIM1 and EB1 are required for MT growth-mediated ER tubule extension

STIM1

EF-hand

SAM domain

SxIP Basic/serine rich

TM Coiled coil

S

Ca2+ ER lumen

EB1 Cytosol

Figure 1.6 Schematic representation of STIM1 protein sequence. EF-hand, calciumbinding motif; TM, transmembrane region; SAM, sterile a motif domain. The scheme illustrates the site of interaction with EB1, but not with the other partners. The part located N-terminally of the transmembrane region is in the ER lumen, while the C-terminal part protrudes into the cytosol.

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(Cahalan, 2009; Grigoriev et al., 2008). It is tempting to speculate that this process might contribute to quick translocation of the ER to the plasma membrane and store-operated Ca2þ entry; however, experimental support for this idea is currently lacking. 5.3.5. p140Cap p140Cap, also known as SNIP (SNAP-25 interacting protein), comprises two predicted coiled coil domains, two highly charged domains, and two proline-rich domains (Chin et al., 2000). Based on this domain structure, p140Cap is thought to function as an adaptor protein. It was identified as an interacting partner of p130Cas (Crk-associated substrate) involved in integrin and epidermal growth factor signaling (Di Stefano et al., 2004). It was also recently described as a Src-binding protein which inhibits Src kinase activity, regulates the actin cytoskeleton, and suppresses tumor growth (Di Stefano et al., 2007). Besides playing important roles in cell adhesion, growth, and Src tyrosine kinase signaling in nonneuronal cells, p140Cap is abundant in the brain and is tightly associated with the neuronal cytoskeleton; it colocalizes with synaptic sites as well as actin (Chin et al., 2000; Ito et al., 2008). These findings suggest a potential role for p140Cap in neuronal cells, particularly in neurotransmitter release, synapse formation or maintenance, and signaling. p140Cap was recently found to interact with all three mammalian EB proteins in vitro through its C-terminal basic region, which contains a good match to the SxIP motif (Honnappa et al., 2009; Jaworski et al., 2009; Fig. 1.7). Indeed, in nonneuronal cells p140Cap colocalizes with EBs at growing MT plus ends ( Jaworski et al., 2009). p140Cap is abundantly present in dendritic spines, and together with EB3 plays a role in regulating dendritic spine morphology. Various depletion and overexpression experiments in hippocampal neurons support a model in which EB3-decorated SxIP

p140Cap

Basic/ serine rich P - Tyr

P

CC

P

S

EB3

Figure 1.7 Schematic representation of p140Cap protein sequence. P, proline-rich regions, P-Tyr, a stretch of phosphorylated tyrosine residues. The scheme illustrates interaction site with EB3 but not with other partners.

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MT plus ends grow into dendritic spines and influence their morphology by controlling the actin cytoskeleton through p140Cap ( Jaworski et al., 2009). This pathway likely involves p140Cap binding partner cortactin, a Src kinase substrate which was implicated in stabilization and branching of actin filaments (Ammer and Weed, 2008; Jaworski et al., 2009). 5.3.6. Navigators Neuron Navigators (NAV1, NAV2, and NAV3) are mammalian homologues of the C. elegans unc-53 protein that is important for cell migration and axon guidance (Maes et al., 2002). Navigators are large proteins of 250 kDa that contain a MT binding domain at the N-terminus and an AAA ATPase domain at the C-terminus (Frickey and Lupas, 2004; Fig. 1.8). NAV2 and NAV3 also contain an N-terminal CH domain of the actin-binding type (Peeters et al., 2004). NAV1 is mainly expressed in the heart and skeletal muscle, while NAV2 is ubiquitously expressed and NAV3 is mostly brain specific (Coy et al., 2002; Maes et al., 2002; Peeters et al., 2004). Members of this protein family were shown to be involved in a variety of cellular processes (see van Haren et al., 2009, and references therein). NAV1 was the first family member that was described to localize to growing MT plus ends in vivo (Martinez-Lopez et al., 2005). More recently, all tree mammalian Navigators were shown to be þTIPs involved in cytoskeletal organization (van Haren et al., 2009). Interestingly, the authors also reported that in HeLa cells overexpression of Navigators delocalized both CLIP-170 and dynactin from MT tips, indicating that the MT tip binding sites for Navigators and CAP-Gly proteins might overlap (van Haren et al., 2009). In agreement with this observation, the presence of basic and serinerich regions in Navigator sequences suggests that their localization at the

Navigator 2/3 Calponin homology domain

AAA ATPase

Basic/serine rich S

CC

Microtubules

Figure 1.8 Schematic representation of Navigator 2 and 3 protein sequences. CC, coiled coil region; S, basic and serine-rich regions; AAA, ATPase family associated with various cellular activities. MT-interacting region is indicated. Navigator 1 has a similar structure, but lacks the CH domain.

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MT ends might be dependent on binding to EB proteins. Further studies will be needed to better understand the mechanism for plus end tracking by Navigators and the functional importance of their MT plus end localization. 5.3.7. Melanophilin Melanophilin was identified as the mutated gene in leaden mice (Matesic et al., 2001). It is considered as a critical component of the melanosome transport machinery by forming a bridge between Rab27a on the melanosome surface and the actin-based motor myosin Va (Nagashima et al., 2002; Wu et al., 2002). It belongs to the Slac2 family of Rab27-binding proteins and contains conserved Rab-binding and coiled coil domains (Fukuda, 2005; Fig. 1.9). Melanophilin is also a þTIP that tracks growing MT ends by binding to EB1, and EB1 depletion abolishes its MT plus end accumulation (Wu et al., 2005). Melanophilin has a basic region containing an SxIP motif the mutation of which disrupts the interaction with EB1 (Hume et al., 2007); therefore, its recruitment to MT tips is likely similar to other SxIP-containing EB1 partners. Melanophilin can promote MT end targeting of Myosin Va, and might thus help to transfer melanosomes between the MT and actin networks (Wu et al., 2005). 5.3.8. RhoGEF2 Drosophila RhoGEF2 (Rho-type guanine nucleotide exchange factor) is a member of the Dbl family of Rho GEFs, which work as activators of the Rho/Rac/Cdc42 superfamily of GTPases. RhoGEF2 contains a PDZ domain at the N-terminus and a cysteine-rich C1 motif in the central region (Hacker and Perrimon, 1998; Fig. 1.10). The C-terminal region of RhoGEF2 contains two motifs: a Dbl homology domain (DH domain) and a pleckstrin homology domain (PH domain) (Hacker and Perrimon, 1998). Melanophilin SxIP Synaptotagmin homology domains SHD

SHD

Myosin Va binding domain

Actin binding domain

Basic

CC

Zinc finger EB1

Figure 1.9 Schematic representation of melanophilin protein sequence, with different partner-interacting domains. EB1 interaction site is indicated.

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RhoGEF2 Basic/serine rich PDZ domain

C1 domain

RhoGEF/DH domain PH domain

S

Figure 1.10 Schematic representation of Drosophila RhoGEF2 protein sequence. PDZ, a domain found in post synaptic density protein PSD95, Drosophila disc large tumor suppressor Dlg, and zonula occludens-1 protein (zo-1); C1, protein kinase C conserved region 1; RhoGEF/DH, guanine nucleotide exchange factor for Rho/Rac/ Cdc42-like GTPases, also called Dbl-homologous domain; PH, pleckstrin homology domain. EB1-binding domain is not precisely mapped but is expected to be within the basic and serine-rich region.

RhoGEF2 is implicated in altering actin and MT behavior and is considered to be a key regulator of morphogenesis in Drosophila. Interestingly, it accumulates at the tips of growing MTs through an interaction with EB1 (Rogers et al., 2004). RhoGEF2 contains a basic and serine-rich region with distinct SxIP motifs, suggesting that its plus end binding mechanism might be similar to those of APC and spectraplakins. The significance of plus end tracking behavior for the signaling function of RhoGEF2 in contractility remains to be determined. 5.3.9. TIP150 Recently, a new þTIP, TIP150 was shown to bind to EB1 in vitro and colocalize with it at the MT plus ends in cells ( Jiang et al., 2009). The EB1 binding domain of TIP150 has been mapped to the middle part of the protein. It consists of a basic, serine, and proline-rich region with one SxLP motif (Fig. 1.11). Depletion of EB1 eliminates the plus end tracking behavior of TIP150 ( Jiang et al., 2009). The protein has no other conspicuous domains with the exception of the C-terminal coiled coil, is conserved in vertebrates but has no clear counterparts in more distant taxa, and its exact function is currently unknown. Interestingly, TIP150 also binds to MT depolymerizing kinesin MCAK, and this interaction is negatively regulated by Aurora B ( Jiang et al., 2009). After TIP150 knockdown, the plus end localization of MCAK was diminished, suggesting that TIP150 could play an additional role in targeting MCAK to the MT plus ends. Further experiments will be needed to better understand the functional mechanism behind the cooperation of TIP150, MCAK, and EB1, which is likely to be evolutionary conserved, since the Xenopus homologue of TIP150, ICIS (inner centromere KinI stimulator) was shown to regulate the activity of the Xenopus MCAK homologue (Ohi et al., 2003).

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TIP150 SxLP Basic/serine rich S

Coiled coil

EB1

Figure 1.11 Schematic representation of TIP150 protein sequence, with EB1-binding site indicated.

5.3.10. CDK5RAP2 CDK5RAP2 (cyclin dependent kinase 5 regulatory subunit associated protein 2, also known as Cep215) is a centrosomal protein associated with a neurogenic disorder, the autosomal recessive primary microcephaly (Bond et al., 2005; Ching et al., 2000). It has a Drosophila homologue, centrosomin, with which it shares evolutionary conserved and functionally important N- and C-terminal domains, and more distantly related fission yeast counterpart Mod20 (Barr et al., 2010). CDK5RAP2 binds to the g tubulin ring complex, is required for centrosome assembly and cell division and its functions are intensely investigated (Barr et al., 2010; Bond et al., 2005; Lizarraga et al., 2010). Human CDK5RAP2 contains a basic and serine-rich region with a sequence SRLP, which represents a variant of SxIP motif; CDK5RAP2 binds to EB1 and mutation of the Leu–Pro dipeptide abolishes EB1 interaction (Fong et al., 2009; Fig. 1.12). As could be expected, CDK5RAP2 tracks MT plus ends; together with EB1 it promotes MT assembly in vitro, and its depletion induced mild MT growth promoting and pause-suppressing effect in cells (Fong et al., 2009). Interestingly, some mammalian homologues of CDK5RAP2 (such as those from rodents) miss the SRLP motif and do not bind to EB1, illustrating that plus end tracking dependent on EB1 association can be a feature that is altered rapidly during evolution (Fong et al., 2009).

5.4. TOG and TOG-like domain proteins Proteins from this family are characterized by the presence at their N-terminus of a variable number of tumor overexpressed gene (TOG) domains, which comprise six HEAT repeats (Neuwald and Hirano, 2000; Slep, 2009). Multiple TOG domains are present in the XMAP215/Dis1 family members, while a single canonical TOG domain and two conserved

37

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CDK5RAP2 SxLP

Centrosomin motif 1

Basic/ serine rich Coiled coils

Coiled coils S

EB1

Figure 1.12 Schematic representation of CDK5RAP2 protein sequence. The scheme illustrates interaction site with EB1 but not with other partners.

helical domains with an organization reminiscent of TOG domains are present in CLASPs. Crystallographic studies showed that TOG domains consist of 12 major helices arranged into an oblong structure with some highly conserved surfaces that can interact with tubulin dimers (Slep, 2009). 5.4.1. XMAP215/Dis1 family 5.4.1.1. Structure, evolutionary conservation, and MT plus end tracking mechanisms The XMAP215/Dis1 family proteins are widely conserved and have emerged as versatile regulators of MT dynamics (Gard et al., 2004; Howard and Hyman, 2007; Ohkura et al., 2001; Popov and Karsenti, 2003; Slep, 2009). Since a large number of excellent reviews covered different aspects of their function, they will be only briefly discussed here. XMAP215 was originally purified from Xenopus eggs as a factor that can promote rapid MT plus end growth (Gard and Kirschner, 1987). Homologues of XMAP215 have since been found in most eukaryotes including ch-TOG in humans (Charrasse et al., 1995, 1998), Stu2 in budding yeast (Wang and Huffaker, 1997), Dis1 and Alp14 in fission yeast (Ohkura et al., 1988), ZYG-9 in C. elegans (Matthews et al., 1998), Msps in Drosophila (Cullen et al., 1999), DdCP224 from Dictyostelium (Graf et al., 2000) and MOR1 from Arabidopsis (Kawamura and Wasteneys, 2008). Two to five TOG domains present in different family members are implicated in binding to tubulin and MTs (Gard et al., 2004; Fig. 1.13). XMAP215 is a thin, elongated protein that can span up to eight tubulin dimers along a protofilament (Cassimeris et al., 2001). XMAP215 was proposed to accelerate MT growth by targeting preassembled tubulin oligomers to MT plus ends (Kerssemakers et al., 2006; Slep and Vale, 2007). However, not all experimental observations fit with this model. First, Stu2 from budding yeast binds just one tubulin dimer at a time

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XMAP215 TOG domains

Basic

Tubulin

Helical

TACC

Figure 1.13 Schematic representation of XMAP215 protein sequence. TOG domains can mediate interaction with tubulin dimers, but it is unclear if all five TOG domains of XMAP215 perform this function. The domain interacting with TACC (transforming acidic coiled coil proteins) is indicated.

(Al-Bassam et al., 2006). Second, it was shown that under certain conditions, XMAP215 can promote MT shortening (Shirasu-Hiza et al., 2003; Vasquez et al., 1994). Recently, a new mechanism that can explain these observations was proposed (Brouhard et al., 2008). The authors observed that similar to Stu2, XMAP215 binds to tubulin dimers in a 1:1 ratio, and that GFP-XMAP215 associates with the tips of growing and shortening MTs in vitro. XMAP215 as well as Stu2 wraps around a tubulin dimer and facilitates its incorporation into a growing MT plus end. This activity is purely catalytic and therefore can also promote the reverse reaction under conditions that favor depolymerization such as absence of tubulin (Brouhard et al., 2008). Interestingly, single molecule imaging showed that XMAP215 moves with the growing MT plus end, acting as a processive MT polymerase. XMAP215 binds free tubulin and targets the MT plus end by 1D diffusion. After reaching the MT plus end it remains attached to its tip, facilitating several rounds of tubulin dimer addition. This new mechanism is a form of processive tip tracking; therefore, similar to the EBs, XMAP215 is an autonomous þTIP. How XMAP215 stays attached to MT tips is not known, but interestingly, the yeast homologue Stu2 preferentially binds to the plus ends of taxol-stabilized MTs (van Breugel et al., 2003). Importantly, the in vivo conditions are quite different from those in vitro due to the presence of other MAPs that could cooperate or compete with XMAP215 at the MT plus end (van der Vaart et al., 2009). For example, binding to EBs, the core proteins of the MT plus end could facilitate XMAP215 plus end tracking. Accordingly, several reports provide evidence for interaction between XMAP215 family members and EBs: Stu2 interacts with Bim1 (Chen et al., 1998; Wolyniak et al., 2006); DdEB1 is present in a complex with DdCP224 (Rehberg and Graf, 2002) and XMAP215 with EB1 (Kronja et al., 2009; Niethammer et al., 2007). The interaction between XMAP215 and EB proteins is probably cell cycle regulated and might involve additional factors.

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5.4.1.2. Cellular roles of XMAP215/Dis1 proteins All Dis1/TOG family members localize to interphase MTs, centrosomes (spindle pole bodies in yeast) and are found throughout the spindle during mitosis. While it is generally accepted that these proteins are major regulators of MT polymer levels and turnover (Holmfeldt et al., 2009; Howard and Hyman, 2007), the exact effect of these proteins on different parameters of dynamic instability in vivo is not yet entirely clear and can differ between model systems, cell types, and phases of the cell cycle. XMAP215 strongly accelerates MT polymerization rate in solutions of purified tubulin, and has a similar, albeit weaker effect in Xenopus egg extracts (Tournebize et al., 2000). In line with these observations, depletion of ZYG-9 inhibits MT polymerization rate in C. elegans embryos (Srayko et al., 2005). Mammalian ch-TOG can promote MT elongation (Bonfils et al., 2007); however, in mitotic mammalian cells, ch-TOG has little effect on MT growth rate although it does affect MT turnover (Cassimeris et al., 2009). Several studies support the idea that XMAP215/Dis1 proteins play a role in MT plus end stabilization (Charrasse et al., 1998; Dionne et al., 2000; Tournebize et al., 2000). One interesting model suggested that XMAP215 strongly stabilizes MTs during interphase by opposing MT destabilizers, like XKCM1 (Xenopus homologue of MCAK) (Kinoshita et al., 2001, 2002). In mitosis, the stabilizing activity of XMAP215 is strongly decreased by phosphorylation, whereas XKCM1 remains constant. This change in the balance of activities of the two factors would then result in more dynamic MTs during mitosis (Andersen and Wittmann, 2002). This model was supported by in vitro studies using three purified components, tubulin, XMAP215, and XKCM1, that were sufficient to reconstitute in vitro the essential features of cellular MT dynamics (Kinoshita et al., 2001). Although it is clear that the situation in cells is much more complex, this study was an important milestone in using an in vitro reconstitution system as a tool to understand MT dynamics in living cells. XMAP215 homologues are abundantly present at the centrosomes, to which they localize through an interaction of their C-terminal domains with the proteins of the TACC family (Peset and Vernos, 2008), and are required for proper spindle assembly and spindle pole integrity (Barr and Gergely, 2008; Cassimeris and Morabito, 2004; Cassimeris et al., 2009; Gergely et al., 2003; Holmfeldt et al., 2004). In plants, MOR1 is required for spatial organization of the acentrosomal MT arrays (Kawamura and Wasteneys, 2008). In budding yeast, Stu2 is present at kinetochores and regulates kinetochore MT dynamics (see Kitamura et al., 2010 and references therein), and Dis1 in fission yeast can also participate in MT-kinetochore attachment (Aoki et al., 2006). Much less is known about the function of XMAP215/Dis1 proteins in differentiated cells, but a genetic screen recently identified a role for the fly homologue, Msps, in axon guidance, where Msps might antagonize the effects of another TOG

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domain protein CLASP/Orbit/MAST (see below; Lowery et al., 2010). Taken together, these data suggest that XMAP215/Dis1 proteins exert a complex set of effects on MT polymerization and organization of MT plus and minus ends. 5.4.2. CLASPs 5.4.2.1. Structure, evolutionary conservation, and plus end tracking mechanisms The CLASP family is conserved between yeasts, plants, and animals. Mammalian CLASPs (CLASP1 and CLASP2) were named after being identified as CLIP associating proteins (Akhmanova et al., 2001) and are homologues of Drosophila Orbit/MAST (Inoue et al., 2000; Lemos et al., 2000), Xenopus Xorbit (Hannak and Heald, 2006), S. cerevisiae Stu1 (Pasqualone and Huffaker, 1994; Yin et al., 2002), S. pombe Peg1/CLASP (Bratman and Chang, 2007; Grallert et al., 2006), C. elegans Claspcls2 (Gonczy et al., 2000) and the A. thaliana AtClasp (Ambrose and Wasteneys, 2008; Gardiner and Marc, 2003). Analysis of CLASP sequences reveals one conventional TOG domain at the N-terminus, and two additional TOG-like domains (Slep, 2009), as well as a basic and serine-rich region in the middle of the protein (Mimori-Kiyosue et al., 2005; Fig. 1.14). The C-terminal domain of CLASPs, which is also highly conserved, mediates interactions with CLIPs, with the cortical protein LL5b, with a Golgi protein GCC185, and with a kinetochore-bound kinesin CENP-E (Akhmanova et al., 2001; Efimov et al., 2007; Hannak and Heald, 2006; Lansbergen et al., 2006; Maffini et al., 2009). The central basic and serine-rich region is required for binding to MTs and EB1 CLASP1/2a SxIP SxIP

TOG domain

TOGlike

Basic/serine rich

TOGlike

Helical

S

Microtubules EBs IQGAP1

CLIPs, LL5b, GCC185, CENP-E

Figure 1.14 Schematic representation of CLASP1/2a protein sequences. Shorter CLASP isoforms lack the C-terminal TOG domain. Interaction sites with various binding partners are indicated.

Microtubule Plus End Tracking Proteins

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(Mimori-Kiyosue et al., 2005). In human CLASP2, this part contains two tandemly repeated SxIP motifs, which are required for EB1 binding and targeting to MT ends (Honnappa et al., 2009). CLASPs thus depend on EB proteins for MT plus end accumulation, while they attach to various cellular structures through their C-terminus. Interaction with the CLIPs can also contribute to MT plus end targeting, even in the absence of the EB1binding region (Mimori-Kiyosue et al., 2005). The function of the TOG and TOG-like domains in CLASPs is currently less clear, but it might be pertinent for the MT stabilization and/or rescue function. It should be noted that the N-terminal TOG domain is present in the long (a) isoforms of mammalian CLASP1 and CLASP2, but absent from the shorter isoforms. Interestingly, one of such shorter isoforms, CLASP2b, which is highly expressed in neurons, contains a palmitoylation sequence at its N-terminus, suggesting a direct link to membranes (Akhmanova et al., 2001). 5.4.2.2. CLASPs in regulation of MT dynamics CLASPs are potent MT-stabilizing factors. They induce formation of stable MT bundles when overexpressed (Akhmanova et al., 2001; Bratman and Chang, 2007; Maiato et al., 2003), while injection of anti-CLASP antibodies or CLASP depletion prevents the formation of stabilized MTs and promotes MT dynamicity (Akhmanova et al., 2001; Drabek et al., 2006; MimoriKiyosue et al., 2005; Sousa et al., 2007). CLASPs stabilize MTs by promoting MT rescues and/or pauses (Bratman and Chang, 2007; Mimori-Kiyosue et al., 2005; Sousa et al., 2007). Studies in HeLa cells showed that CLASPs require the EB1 binding basic and serine-rich domain for their MT rescue activity (Mimori-Kiyosue et al., 2005), but since this region can also bind to MTs directly, it is not clear if the interaction with EBs is mechanistically important for the rescue function. The activity of CLASPs is tightly spatially regulated: for example, they rescue and stabilize MT plus ends specifically at the leading edge of migrating cells (Akhmanova et al., 2001; Drabek et al., 2006; Mimori-Kiyosue et al., 2005; Wittmann and Waterman-Storer, 2005). This is in part due to the action of GSK3b kinase, which phosphorylates serine residues around SxIP motifs, and thus prevents the interaction of CLASPs with the EBs and MTs throughout the cell, except for the leading edge where GSK3b is inhibited (Akhmanova et al., 2001; Kumar et al., 2009; Watanabe et al., 2009). These data provide a nice example of how phosphorylation in the vicinity of SxIP motifs negatively regulates the localization of þTIPs to MT ends by decreasing their affinity to EB1 (Honnappa et al., 2009; Kumar et al., 2009). Importantly, CLASPs not only stabilize MTs against depolymerization but also physically attach them to the cell cortex through a direct interaction of their C-terminal domain with the cortical protein LL5b, which is also located at the leading edge of migrating cells (Lansbergen et al., 2006). The spectraplakin ACF7/MACF1 acts in the

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same MT stabilization pathway and regulates cortical CLASP localization (Drabek et al., 2006). Interestingly, based on proteomics and genetic data from Drosophila, CLASP might physically interact with spectraplakin (Shot in flies), as well as XMAP215 homologue Msps (Lowery et al., 2010). The interaction of CLASP with the cell cortex was also observed in plants where in its absence MTs partially detach from the cortex (Ambrose and Wasteneys, 2008). Besides MT plus ends and the cortex, mammalian CLASPs also localize to centrosomes and the Golgi apparatus (Akhmanova et al., 2001; MimoriKiyosue et al., 2005). Attachment to the Golgi depends on the interaction of the C-terminal domain of CLASPs with the golgin GCC185 and is required for MT outgrowth from the Golgi membranes (Efimov et al., 2007). Whether CLASPs participate in MT nucleation or anchoring at the centrosome is currently unknown, as CLASPs are very difficult to deplete from the centrosomes. In fission yeast, CLASP localizes to and stabilizes the regions of antiparallel MT overlap, to which it is targeted by the MT bundling protein Ase1 (Bratman and Chang, 2007). Taken together, these studies show that although CLASPs can be localized by their partners to various subcellular sites, their function at these different locations is mechanistically similar—they decorate and stabilize specific MT stretches without capping them, allowing MT plus end outgrowth from the stabilized MT segments. 5.4.2.3. CLASPs in mitosis CLASPs are essential for cell division in a broad variety of organisms. During mitosis, CLASPs localize to the mitotic spindle, kinetochores, and the midbody (Inoue et al., 2004; Lemos et al., 2000; Maiato et al., 2002, 2003). In yeast, CLASP homologues participate in formation of the mitotic spindle (Bratman and Chang, 2007; Grallert et al., 2006; Yin et al., 2002); furthermore, the S. cerevisiae Stu1 localizes to unattached kinetochores and facilitates their capture in prometaphase (Ortiz et al., 2009). In animal cells, CLASPs are also needed for normal kinetochore-MT attachment, as well as chromosome congression and maintenance of spindle bipolarity (Hannak and Heald, 2006; Inoue et al., 2004; Lemos et al., 2000; Maiato et al., 2002, 2003). Human CLASP1 and CLASP2 play redundant roles during mitosis suggesting a possible mechanism to prevent aneuploidy in mammals (Pereira et al., 2006). Detailed mitotic studies of human CLASP1 showed that it accumulates at the outer kinetochore corona (Maiato et al., 2003). Kinetochore targeting in vertebrate cells requires the interaction of CLASP C-terminus with kinesin CENP-E (Hannak and Heald, 2006; Maffini et al., 2009); importantly, although CENP-E is a plus end-directed motor, CENP-E dependent CLASP targeting to kinetochores is MT-independent. Although specific kinetochore interaction of CLASP homologues is conserved in fungi and invertebrates (Cheeseman et al., 2005; Ortiz et al., 2009), the binding

Microtubule Plus End Tracking Proteins

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partners might be different, as in C. elegans CLASP homologue is targeted to kinetochore by CENP-F like proteins HCP-1/2 (Cheeseman et al., 2005). In mammalian cells, CLASPs promote MT turnover at the kinetochores (Maffini et al., 2009). Drosophila CLASP homologue Orbit/MAST also promotes incorporation of new tubulin subunits at the plus ends of kinetochore MTs (Maiato et al., 2005), and in the absence of Orbit/MAST, the poleward flux of MTs is strongly reduced (Buster et al., 2007). Codepletion of Drosophila CLASP and dynein rescues spindle bipolarity in CLASPdepleted fly cells, suggesting antagonistic roles for these proteins in the local regulation of MT plus end dynamics at kinetochores (Reis et al., 2009). CLASPs also contribute to the later stages of cell division, by stabilizing the internal part of the central spindle, which in its turn is important for the formation of the cleavage furrow (Inoue et al., 2004). 5.4.2.4. CLASPs in other cellular structures CLASPs might directly interact with actin (Tsvetkov et al., 2007), as well as IQGAP1 and some other actin-binding proteins (Lowery et al., 2010; Watanabe et al., 2009), and in Drosophila, CLASPs were implicated in oocyte differentiation, playing a role in the organization of the polarized MT network and participating in the interaction between the cleavage furrow and the fusome, an actinrich structure (Mathe et al., 2003). Establishment of polarized cortically attached MT arrays is also an important function of CLASPs in migrating fibroblasts and polarized epithelial cells (Akhmanova et al., 2001; Drabek et al., 2006; Hotta et al., 2010; Wittmann and Waterman-Storer, 2005). An interesting example of this function is provided by migrating Drosophila macrophages, which form a CLASP-dependent MT bundle (“arm”) that points in the direction of migration and participates in cell–cell repulsion (Stramer et al., 2010). In developing neurons, CLASPs localize to the growth cones, and Drosophila CLASP homologue participates in the regulation of axon guidance downstream of Abelson (Abl) nonreceptor tyrosine kinase, with which it interacts both genetically and physically (Lee et al., 2004; Lowery et al., 2010).

5.5. Motor proteins MT motor proteins, kinesins, and dyneins, provide the driving force for MT-based transport and play essential roles in controlling MT network organization and dynamics during interphase and mitosis. Several motor proteins, including minus end-directed motors such as cytoplasmic dynein, and plus end-directed motors, like kinesin-7 family, strongly accumulate at the growing MT ends by interacting with comet-making þTIP machinery.

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5.5.1. Kinesins Kinesins can be classified according to the position of their highly conserved motor domain, which can be located at the N- or C-terminus, or in the middle of the molecule (Hirokawa et al., 2009; Lawrence et al., 2004; Miki et al., 2005). Kinesins with an N-terminal motor domain move to MT plus ends, while those with a C-terminal motor walk to MT minus ends. Kinesin-13 family members, which have the motor domain in the middle of the proteins, are nonmotile and use their motor activity to depolymerize MTs. 5.5.1.1. MT plus end-directed kinesins Yeast kinesins Kip2 and Tea2 are plus end-directed motors that belong to the kinesin-7 family. They are the best-known examples of kinesins that track growing MT plus ends (Akhmanova and Steinmetz, 2008). Kip2 stabilizes MTs by targeting Bik1, the CLIP-170 homologue in budding yeast, to MT plus ends (Carvalho et al., 2004). Bik1 forms a complex with Kip2 and comigrates with it along MTs; because the rate of migration exceeds MT polymerization speed, Bik1 accumulates at the plus ends (Carvalho et al., 2004). Kip2 also targets dynein to the MT plus end; Kip2 levels are regulated during the cell cycle, and Kip2-dependent transport is therefore important for the cell cycle control of MT dynamics and dynein-dependent processes (Carvalho et al., 2004). Additional clues about Kip2 involvement in þTIP targeting in budding yeast were revealed by removing the last amino acid of a tubulin. In such “Glu-tubulin” yeast strains, despite the presence of robust Kip2 comets at MT plus ends, Bik1 failed to track the plus ends. However, dynein positioning at the same plus ends was unperturbed (Caudron et al., 2008). These results suggest that besides interaction with Kip2, Bik1 associated with the tubulin C-terminus (Caudron et al., 2008). Moreover, Kip2 also interacts with Kar9 and moves it along cytoplasmic MTs to their plus ends where it accumulates in a manner dependent on Cdc28, the yeast Cdk1 kinase (Maekawa et al., 2003). Tea2, the fission yeast homologue of Kip2, is located at the MT tips and promotes MT polymerization, which is important for establishing and maintaining polarized growth along the long axis of the cell (Browning et al., 2000). Tea2 is responsible for the plus end accumulation of Tip1, the CLIP-170 homologue (Browning et al., 2003; Busch and Brunner, 2004). Tea2 forms a complex with Tip1 and moves it along MTs toward their growing tips, similar to Kip2–Bik1 complex (Busch and Brunner, 2004). Interestingly, Tea2 requires Mal3, the yeast homologue of EB1, for MT loading, accumulation at MT plus ends, and/or processivity (Bieling et al., 2007; Browning et al., 2003; Busch et al., 2004). Mal3 directly binds to Tea2 and stimulates its ATPase activity (Browning and Hackney, 2005).

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KipA, a kinesin-7 homologue in Aspergillus nidulans also localizes to the plus ends of MTs (Konzack et al., 2005) and contributes to MT tip localization of the CLIP-170 homologue in certain conditions (Efimov et al., 2006). Kinesin-7 in vertebrates, CENP-E, has important mitotic functions, as it interacts with kinetochore MTs and also targets CLASPs to kinetochores (Maffini et al., 2009). However, the latter function is MT independent and there are no indications that CENP-E participates in þTIP transport to MT ends. Another family of plus end-directed kinesins with þTIP properties are kinesin-8 family members, which can depolymerize MTs or “dampen” MT dynamics (Gardner et al., 2008c; Varga et al., 2009). They can localize to MT tips independently of other þTIPs, and will not be discussed here. 5.5.1.2. MT minus end-directed kinesins Several members of kinesin-14 family, minus end-directed kinesins with a C-terminal motor domain, track MT plus ends. Kar3, a S. cerevisiae kinesin 14, is essential for karyogamy (nuclear fusion), meiosis as well as certain aspects of mitotic spindle organization (Gardner et al., 2008a; Meluh and Rose, 1990; Sproul et al., 2005). The budding yeast shmoo tip is a model for the regulation of dynamic MT attachment: MT plus ends interact with the cell cortex at the shmoo tip, positioning the nucleus for karyogamy (Maddox, 2005). High-resolution imaging and genetic manipulation have shown that Kar3 is required for depolymerization of MT plus ends at the shmoo tip while they remain attached to the cortex (Maddox et al., 2003). In vitro assays showed that Kar3 forms a heterodimer with Cik1, a noncatalytic polypeptide, which targets Kar3 to MT plus ends (Sproul et al., 2005). Kar3 and Cik1 were also shown to play a role in mitosis, where they contribute to spindle stability, most likely by promoting MT bundling (Gardner et al., 2008a). The Drosophila kinesin-14 Ncd also shows minus end-directed motility and binds to MT ends (Sproul et al., 2005). RNA interference and highresolution microscopy showed that Ncd localizes to the growing MT plus ends through an interaction with EB1 (Goshima et al., 2005). The plus end localization of Ncd facilitates the capture and transport of kinetochore fibers along MTs, a process important for maintaining bipolar mitotic spindles (Goshima et al., 2005). A similar mechanism might apply to a kinesin-14 from A. thaliana, ATK5, which was shown to target MT plus ends and is involved in spindle assembly (Ambrose and Cyr, 2007; Ambrose et al., 2005). Another kinesin-14, Klp2p from fission yeast, also binds to MT tips and participates in organizing antiparallel MT bundles through its MT sliding activity. Since it preferentially acts at MT ends, the sliding force that it can produce is limited; when it acts in combination with MT bundling proteins, it can transport short but not long MTs, and can thus position short MTs at their correct location ( Janson et al., 2007).

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Finally, Drosophila Nod, a nonmotile kinesin important for chromosome segregation in meiosis, has a strong preference for binding to MTs plus ends in vitro, possibly because it displays a differential binding mode at MT tips (Cochran et al., 2009; Cui et al., 2005). 5.5.1.3. Kinesin-13 family: MCAK MCAK was initially classified as a member of the Kin I subfamily of kinesins and recognized as a MT depolymerase influencing MT stability by inducing a destabilizing conformational change at MT plus ends (Desai et al., 1999). More recently, the kinesin nomenclature was standardized and MCAK (KIF2C) now belongs to the kinesin-13 family, which includes mammalian KIF2A and KIF2B, Xenopus XKCM1 and XKIF2 and Drosophila KLP10A, KLP59C, and KLP59D (Lawrence et al., 2004; Moores et al., 2006). This family has a unique feature among kinesins because rather than walk along MTs, they utilize ATP hydrolysis to depolymerize them from both ends (Desai et al., 1999; Helenius et al., 2006; Hunter et al., 2003). Interestingly, MCAK can depolymerize MTs in the absence of ATPase activity, but only when there is a 1:1 stoichiometry of motor and tubulin (Moore and Wordeman, 2004). MCAK is a homodimeric molecule with the centrally located motor domain, which together with the neck domain at the N-terminus is sufficient for MT depolymerization in vivo and in vitro (Maney et al., 2001; Moores et al., 2006; Newton et al., 2004; Ogawa et al., 2004; Fig. 1.15). The N-terminal region of MCAK also includes a basic and serine-rich stretch, which influences the subcellular localization of MCAK (Hertzer et al., 2006). The C-terminal region contains a coiled coil that is responsible for dimerization and plays a role in regulating the ATPase activity of MCAK (Ems-McClung et al., 2007).

MCAK SxIP Basic/serine rich S

Motor domain Neck

Coiled coil

Helical domain EBs

Microtubules

Figure 1.15 Schematic representation of MCAK protein sequence. The kinesin motor domain interacts with MTs; the positively charged neck region contributes to this interaction. The site of interaction with EB1 is indicated.

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Surprisingly, in spite of being a potent MT depolymerase, MCAK tracks growing MT plus ends in living cells (Moore et al., 2005). In vitro, the targeting of MCAK to MT ends is extremely rapid, as compared to standard kinesin kinetics. Single molecule microscopy assays showed that MCAK moves along the MT lattice by 1D diffusion targeting both MT ends more rapidly than could occur based on direct binding from solution (Helenius et al., 2006). Furthermore, MCAK associates with EB1 and EB3 through its N-terminus and colocalizes with EB1 at growing MT ends in cells (Lee et al., 2008). The basic and serine-rich region at the N-terminus of MCAK responsible for EB1 binding contains one SxIP motif (Honnappa et al., 2009). An MCAK mutant, in which the SxIP was disrupted displayed no binding to EB1 and no plus end tracking, although it could still bind to the MT lattice (Honnappa et al., 2009). A mitotic kinase Aurora B can phosphorylate MCAK at several sites and thus affect its function (Andrews et al., 2004; Lan et al., 2004). These sites mostly cluster near the SxIP motif, and phosphomimicking mutations in MCAK abolish its binding to EB1 and plus end tracking (Honnappa et al., 2009; Moore et al., 2005). KIF2A and KIF2B, which contain no SxIP motifs, are unable to accumulate at polymerizing MTs ends although they weakly bind to the MT lattice (Honnappa et al., 2009). Drosophila KLP10A also associates with EB1, and this interaction is involved in targeting KLP10A to MTs plus ends (Mennella et al., 2005). KLP10A does not colocalize with EB1 completely, and instead appears to trail just behind EB1 at the MT tip (Mennella et al., 2005). Together with the fact that kinesin-13s have MT lattice affinity, this suggests that they recognize composite binding sites on MT tips formed by EB1 and tubulin, similar to CLIPs. In line with this view, the interaction of MCAK with MTs is affected by tyrosination status of a tubulin (Peris et al., 2009). MCAK plays a role in MT depolymerization by triggering catastrophes (Howard and Hyman, 2007). This MCAK feature confers a crucial role in mitosis, where it participates in spindle assembly, chromosome congression and segregation, and correction of improper MT-kinetochore attachments (Ems-McClung and Walczak, 2010). 5.5.2. Cytoplasmic dynein Cytoplasmic dynein is a large multisubunit complex of 1 MDa composed of two heavy chains as well as intermediate, light intermediate, and light chains. The heavy chains contain the motor domain, which comprises six AAA-type ATPase units and a MT binding stalk (Hook and Vallee, 2006). Similar to kinesins, dynein uses the energy of ATP hydrolysis to move along MTs toward their minus ends. Cytoplasmic dynein activity in vivo requires the multisubunit dynactin complex (Schroer, 2004), the function of which was already discussed above. Dynein has important roles in transport of a large variety of cargo along MTs, formation and orientation of the mitotic

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spindles as well as other MT arrays, and in nuclear migration. These functions have been subject of numerous reviews (Kardon and Vale, 2009; Vallee et al., 2004). Here, we will briefly summarize what is known about the dynein association with MT tips. A pool of cytoplasmic dynein can be found at the growing MT ends in different species ranging from fungi to mammals. In mammalian cells, endogenous dynein has been observed at MT ends using confocal and TIRF microscopy (Dujardin et al., 2003; Vaughan et al., 1999). The expression of GFP-tagged dynein intermediate chain as a functional probe for cytoplasmic dynein showed comet-like structures that colocalized with EB1, indicating that it is indeed a þTIP in mammalian cells (Kobayashi and Murayama, 2009). In mammals, dynein is likely targeted to MT ends by dynactin or by LIS1, which both require CLIP-170 for efficient plus end accumulation (Akhmanova and Steinmetz, 2008). The functions of dynein at MT ends are not entirely clear, but loading of cargo for minus end transport (Vaughan et al., 2002) and MT interactions with the cell cortex (Dujardin and Vallee, 2002) are the most obvious possibilities. The function of cortical dynein was particularly extensively explored in C. elegans, where it participates in creating centripetal forces for asymmetric spindle positioning during the first cell division (Nguyen-Ngoc et al., 2007). Dynein and its cofactors are also targeted to MT plus ends in fungi. In A. nidulans, the genes encoding dynein and dynactin subunits were identified in a genetic screen for mutants deficient in nuclear distribution (NUD) (Xiang et al., 1999). Both NUDA (cytoplasmic dynein heavy chain) and its cofactor NUDF, the homologue of LIS1, localize to MT plus ends as comet-like structures and affect MT dynamics in vivo (Han et al., 2001). Similarly, GFP-tagged NUDM, the p150Glued homologue, accumulates at the plus ends, and the analysis of different loss-of-function mutants demonstrated that NUDM and NUDA require each other for MT plus end accumulation (Zhang et al., 2003). Interestingly, KINA, the conventional kinesin-1, is required for the þTIP behavior of both dynein and dynactin, but not NUDF in Aspergillus (Zhang et al., 2003). Similarly, in budding yeast, kinesin-7 Kip2 is required for the plus end targeting of dynein as well as the CLIP-170 homologue Bik1 (Carvalho et al., 2004). In the absence of Bik1, dynein is lost from the MT plus ends, suggesting that the Kip2–Bik1 complex is required for dynein plus end tracking (Sheeman et al., 2003). In addition, the plus end localization of dynein requires Pac1, the LIS1 homologue in budding yeast (Lee et al., 2003; Sheeman et al., 2003). As already mentioned, in yeast strains where the last aromatic residue of a tubulin is deleted and Bik1 cannot accumulate efficiently at MT ends, dynein is still present at MT tips, suggesting that Bik1 might be needed for transport but not for retention of dynein at MT tips (Caudron et al., 2008). Recently, a two-step model was proposed to explain dynein accumulation at the plus ends in yeast (Moore et al., 2009).

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First, dynein is targeted to the plus ends by Bik1, the CLIP-170 homologue, and second, it is retained by Pac1 and its binding partner NudEL/Ndl1. Ndl1 might facilitate the recruitment of Pac1 to the MT tip, where Pac1 then acts cooperatively with Kip2–Bik1 to drive the localization of dynein (Moore et al., 2009). Furthermore, a recent study showed that dynein motor domain is necessary and sufficient for dynein accumulation at MT plus ends in a Bik1 and Pac1 dependent manner (Markus et al., 2009). Dynein accumulation at the tips of astral MTs is important for spindle positioning in the bud neck (Moore et al., 2009).

5.6. Other þ TIPs 5.6.1. LIS1 LIS1 protein is involved in neuronal migration and has an essential role during embryonic brain development. LIS1 deficiency results in an impaired ability of neurons to migrate to their correct destination in the cerebral cortex probably due to defects in the dynein pathway (Dobyns et al., 1993; Vallee et al., 2001). LIS1 also has an independent function as a subunit of the brain cytosolic platelet activating factor acetylhydrolase (see Tarricone et al., 2004 for discussion). LIS1 comprises an N-terminal LIS homology domain (LisH) responsible for dimerization and a C-terminal WD40 repeat containing a b propeller domain (Kim et al., 2004; Tarricone et al., 2004; Fig. 1.16). LIS1 is also a þTIP, which interacts with CLIP-170 in a phosphorylation-dependent manner (Coquelle et al., 2002). In addition, LIS1 was also reported to interact with tubulin and MTs (Sapir et al., 1997), although it is not clear whether it directly interacts with MT lattice at the tips. In animal cells, the second zinc finger of CLIP-170 is crucial for LIS1 binding and recruitment to MT tips (Coquelle et al., 2002). Interestingly, the same zinc finger is also implicated in binding to the CAP-Gly domain of p150Glued, and LIS1 and p150Glued compete for the interaction with CLIP-170 and LIS1 LisH domain

WD40 domain Coiled coil

CLIP-170, dynein

Figure 1.16 Schematic representation of LIS1 protein sequence. WD40 region is required for binding other þTIPs, CLIP-170 and cytoplasmic dynein, as indicated.

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MT ends, suggesting that LIS1 might act to release dynactin from the MT tips (Lansbergen et al., 2004). LIS1 can bind to the dynein motor domain, and together with another dynein cofactor, NudE, it promotes persistent dynein force production, which is likely to be important when dynein works under high load during neuronal migration (McKenney et al., 2010). LIS1 also interacts through the WD40 repeat domain with dynein intermediate chains and dynactin (Tai et al., 2002). In mammalian cells, LIS1 is recruited to kinetochores in a dynein–dynactin dependent manner; LIS1 overexpression interferes with mitotic progression and leads to spindle misorientation (Coquelle et al., 2002; Faulkner et al., 2000). In budding yeast, the LIS1 homologue Pac1 also acts in the dynein pathway and participates in its targeting to MT tips, as described above (Sheeman et al., 2003; Xiang, 2003). Together with dynein and dynactin, LIS1 is also required for spindle positioning and proper cell division in C. elegans embryos (Cockell et al., 2004; Nguyen-Ngoc et al., 2007). In A. nidulans, LIS1 homologue NUDF affects MT plus end dynamics, possibly through one of its binding partners (Han et al., 2001).

6. Conclusions and Future Directions þTIPs have emerged as important MT regulators and consequently as key factors in a wide range of vital cellular processes. What started by observation of a peculiar dynamic behavior, quickly put þTIPs in the spotlight. The field made remarkable progress, not just in the discovery of a notably large number of new þTIPs but also in understanding the mechanisms of their action and their interactions with MTs and other þTIPs. It is clear that EB family proteins form the core of the comet-making þTIP network. Surprisingly, the EBs are not required for viability neither in fungi nor in plants. It would be interesting to know whether they are essential in animals, particularly in mammals. Furthermore, the mechanism underlying recognition of growing MT tips by EBs needs to be determined, as it provides the basis of MT plus end tracking of a large group of proteins. The mechanisms of EB-dependent MT plus end association are understood better than that of EBs themselves. In particular, the finding that a short sequence motif, SxIP, embedded in basic and serine-rich sequence binds to EB1 and is responsible for targeting a broad variety of otherwise structurally unrelated þTIPs opened the way to identify new þTIPs and to dissect the functional significance of MT plus end tracking (Honnappa et al., 2009). The SxIP sequence is a typical representative of LMs, short peptide sequences of 3–10 residues, of which usually just two or three are functionally important. The function of LMs strongly depends on the flanking regions, which are often flexible and accessible for interactions with globular

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domains of their partners (Chica et al., 2009). LMs are very likely to arise or disappear as a result of point mutations and are rarely conserved over long evolutionary distances (Saito et al., 2007). An interesting example of such rapid LM evolution is provided by CDK5RAP2, which contains an SRLP motif and binds to EB1 in humans and dogs, but not in mice and rats (Fong et al., 2009). It is likely that more of such “evolutionary flexible” SxIPcontaining þTIPs, specific for particular organisms, will be found in future. Another important theme is cooperation between different þTIPs. This includes not only EBs and their partners, but also seemingly autonomous þTIPs. For example, XMAP215 can track growing MT ends by itself; however, XMAP215 family proteins can also bind EB proteins, which, in their turn, might contribute to targeting them to the MT ends. Based on the existing structural studies, EB C-terminus is a docking platform for a large number of binding partners participating in MT plus end functions. The symmetric structure of the EB1 dimerization domain affords two binding sites, indicating that, in principle, two factors can be recruited simultaneously. It is, however, unlikely that more than two partners will interact with an EB molecule at the same time, because comparison of SxIP motif and CAP-Gly binding determinants reveals an overlap indicating that single site occupancy is mutually exclusive (Etienne-Manneville, 2009). Different EB binding partners from the same or different families will thus likely compete for interaction with the EBs. Furthermore, although the number of þTIP binding sites at the growing MT end may be large (up to several thousand), it is still limited, and þTIPs might compete for them. For example, EB1 and EB3 compete and displace EB2 from the MT tips; also, monomeric EB versions can be displaced by dimeric ones (Komarova et al., 2009) Competition has also been observed previously between the CLIPs (Hoogenraad et al., 2002); it thus appears to be a common determinant of the formation of þTIP complexes at the MT end. An important and still relatively poorly explored topic is þTIP regulation. The existing studies indicate that many signaling pathways can converge on þTIPs to regulate their MT association or activity, and many more examples of such regulation are expected to be discovered in future. To summarize, we can conclude that the composition of þTIPs at each MT end can be remarkably complex: it is shaped by both cooperation and competition of individual molecules, can be organism and cell type specific, and can be tightly regulated.

ACKNOWLEDGMENTS We thank Babet van der Vaart for critically reading the chapter. This work was supported by the Netherlands Organization for Scientific Research grants ALW-VICI and ZonMW-TOP to A.A. and by Fundac¸a˜o para a Cieˆncia e a Tecnologia fellowship to S.M.G.

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New Insights into the Roles of Molecular Chaperones in Chlamydomonas and Volvox ¨hlhaus,* and Andre´ Nordhues,* Stephen M. Miller,† Timo Mu Michael Schroda* Contents 1. Introduction 1.1. Hsp100/Clp 1.2. Hsp90 1.3. Hsp70 and its cochaperones 1.4. Hsp60/GroEL/Cpn60 1.5. sHSPs 1.6. Chlamydomonas reinhardtii as a model organism for studying plant molecular chaperone functions 2. Cytosol/Nucleus 2.1. Chaperone components 2.2. Function of cytosolic/nuclear chaperones 3. Flagella 3.1. Chaperone components 3.2. Function of flagellar chaperones 4. Endoplasmic Reticulum 4.1. Chaperone components 4.2. Function of ER-localized chaperones 5. Chloroplast 5.1. Chaperone components 5.2. Function of chloroplast (co)chaperones 6. Mitochondrion 6.1. Chaperone components 6.2. Function of mitochondrial chaperones 7. Conclusions and Outlook Acknowledgments References

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* Max Planck Institute of Molecular Plant Physiology, Am Muehlenberg, Potsdam-Golm, Germany { Department of Biological Sciences, University of Maryland, Baltimore County, Baltimore, Maryland, USA International Review of Cell and Molecular Biology, Volume 285 ISSN 1937-6448, DOI: 10.1016/S1937-6448(10)85002-6

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2010 Elsevier Inc. All rights reserved.

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Abstract The unicellular green alga Chlamydomonas reinhardtii has been used as a model organism for many decades, mainly to study photosynthesis and flagella/cilia. Only recently, Chlamydomonas has received much attention because of its ability to produce hydrogen and nonpolar lipids that have promise as biofuels. The best-studied multicellular cousin of Chlamydomonas reinhardtii is Volvox carteri, whose life cycle comprises events that have clear parallels in higher plants and/or animals, making it an excellent system in which to study fundamental developmental processes. Molecular chaperones are proteins that guide other cellular proteins through their life cycle. They assist in de novo folding of nascent chains, mediate assembly and disassembly of protein complexes, facilitate protein transport across membranes, disassemble protein aggregates, fold denatured proteins back to the native state, and transfer unfoldable proteins to proteolytic degradation. Hence, molecular chaperones regulate protein function under all growth conditions and play important roles in many basic cellular and developmental processes. The aim of this chapter is to describe recent advances toward understanding molecular chaperone biology in Chlamydomonas and Volvox. Key Words: Heat shock proteins, Heat stress response, Chloroplast chaperones, Thylakoid biogenesis, Flagellar chaperones, Asymmetric cell division. ß 2010 Elsevier Inc.

1. Introduction Molecular chaperones guide almost all cellular proteins through their life cycle (Frydman, 2001; Hartl and Hayer-Hartl, 2009). They protect nascent chains from the crowded cellular environment as they emerge from the ribosome; they help (particularly) multidomain proteins in folding to the native state; they mediate assembly and disassembly of protein complexes; they facilitate protein transport across membranes; they disassemble protein aggregates and fold denatured proteins back to the native state; and finally, they transfer unfoldable proteins to the machineries that process proteolytic degradation. Thus, although the specialized function that many chaperones carry out with respect to restoring protein homeostasis during heat shock has been eponymous for the family as a whole (heat shock proteins, Hsps), molecular chaperones in fact regulate protein function under all growth conditions and play important roles in many basic cellular and developmental processes. The purpose of this chapter is to highlight recent progress toward understanding chaperone biology in two particularly useful model systems, the unicellular green alga Chlamydomonas reinhardtii and its multicellular cousin Volvox carteri. We start with a brief introduction

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to the five major molecular-mass grouped families of chaperones—the Hsp100, Hsp90, Hsp70, Hsp60, and small Hsps, and their cochaperones— then describe new insights gained into chaperone function through recent studies involving these two green algae.

1.1. Hsp100/Clp The Hsp100/Clp family can be divided into two classes of proteins that are structurally distinct (Schirmer et al., 1996). Class 1 members contain two AAAþ nucleotide binding domains (NBD1 and NBD2) and comprise the ClpA–E subclasses. Plant HSP101 and yeast HSP104 represent cytosolic ClpB homologs. Class 2 members contain only a single NBD and comprise the HslU, ClpX, and ClpY subclasses. Hsp100/Clp family members assemble into hexameric rings and use the energy released by ATP hydrolysis to unfold substrate proteins and thread them through the central pore of the ring (Weibezahn et al., 2004). ClpBs bind to protein aggregates and extract polypeptide chains that may then be refolded by the Hsp70 chaperone system (Glover and Lindquist, 1998). The ability of ClpB proteins to disassemble protein aggregates is fundamental for induced thermotolerance mediated by cytosolic ClpBs (Sanchez and Lindquist, 1990). Non-ClpB members of the Hsp100 family may transfer the polypeptide thread into the proteolytic chamber of a diffusible protease (ClpP or HslV) (Wickner et al., 1999). However, they may also be involved in specialized functions like the import of proteins via the TOC/TIC translocon into chloroplasts (Akita et al., 1997; Nielsen et al., 1997).

1.2. Hsp90 Members of the Hsp90 family contain three conserved domains. The N-terminal domain realizes nucleotide binding, the middle domain is involved in client protein binding, and the C-terminal domain mediates dimerization (Csermely et al., 1998). Hsp90s interact closely with the Hsp70 system, which performs early folding steps (Wegele et al., 2006). The various folding paths carried out by Hsp90s are guided by a plethora of pathway-specific cochaperones, like Hop, p23, Cdc37, or Aha1. Hsp90s maintain client proteins in a nearly completely folded conformation poised to respond to an activation signal, such as ligand binding or phosphorylation (Zuehlke and Johnson, 2009). Consequently, Hsp90s are essential for the maturation of many hormone receptors and kinases (Pratt and Toft, 2003), but have also been shown to prevent aggregation of heat denatured citrate synthase in vitro ( Jakob et al., 1995).

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1.3. Hsp70 and its cochaperones Chaperones of the Hsp70 family are among the most highly conserved proteins known (Mayer and Bukau, 2005). They consist of an N-terminal ATPase domain and a C-terminal substrate-binding domain. ATP hydrolysis at the ATPase domain regulates substrate binding and release. Substrate proteins recognized by Hsp70s expose hydrophobic amino acids flanked by basic residues (Rudiger et al., 1997), a characteristic feature of nonnative proteins, but also of native Hsp70 substrates. Binding of Hsp70 to hydrophobic regions prevents the formation of aggregates. In addition, the ATP-driven unfolding activity of Hsp70s may introduce conformational changes to bound substrates that eventually allow nonnative proteins to assume the native state (Sharma et al., 2009). Thus, Hsp70s play a major role in the folding of nascent chains and in the renaturation of nonnative proteins that have accumulated during stress situations such as heat shock. However, they are also involved in many highly specialized functions like the regulation of the general stress response in bacteria (Tomoyasu et al., 1998), the uncoating of clathrin-coated vesicles (Ungewickell et al., 1995), or the translocation of proteins across membranes (Kang et al., 1990). Hsp70s function in concert with different cochaperones, of which the J-domain proteins represent an important class (Craig et al., 2006). J-domain proteins stimulate the ATPase activity of their Hsp70 partner (Liberek et al., 1991) and lock it onto specific substrates (Han and Christen, 2003). In this way, J-domain proteins mediate substrate specificity and thereby the function of their Hsp70 partner. The number of J-domain proteins an organism possesses exceeds the number of Hsp70 chaperones, indicating that one Hsp70 may be recruited by multiple J-domain proteins to different targets within a cellular compartment. Another class of Hsp70 cochaperones are the GrpE-type nucleotideexchange factors, which regulate the activities of bacterial Hsp70 homologs (DnaKs) and of the major Hsp70s in mitochondria and chloroplasts (Harrison, 2003). All GrpE homologs form dimers that interact with their Hsp70 partners in the ADP-state and catalyze the release of ADP to allow for rebinding of ATP (Liberek et al., 1991). Also the Hsp70-like proteins of the Hsp110 family act as nucleotide-exchange factors for cytosolic and endoplasmic reticulum (ER)-localized DnaK-type Hsp70s (Polier et al., 2008). As already mentioned above, Hsp70 systems do not act in isolation from the other cellular chaperone systems. They cooperate with ClpBs in protein disaggregation and refolding (Haslberger et al., 2007), with GroEL/ Hsp60 in protein folding (Langer et al., 1992), with the proteasome in quality control (Ballinger et al., 1999), and with the Hsp90 system in protein folding and signal transduction (Pratt and Toft, 2003; Wegele et al., 2004).

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1.4. Hsp60/GroEL/Cpn60 Chaperones of the Hsp60 family, also termed type I chaperonins, are found in eubacteria, mitochondria, and chloroplasts. They assemble into two stacked rings, each containing seven subunits of about 60 kDa (Bukau and Horwich, 1998). In bacteria, the chaperonin is called GroEL, while the plastid and mitochondrial forms are named Cpn60 or Hsp60 to account for their heat shock inducibility (Viitanen et al., 1995). The chaperonin cooperates with a cochaperonin, which forms one heptameric ring consisting of 10-kDa subunits termed GroES in bacteria, and Hsp10 and Cpn10 in mitochondria and plastids, respectively. Eubacteria and mitochondria contain only one type of chaperonin and cochaperonin, while plastids have several types. The 60-kDa subunits of the tetradecameric double ring may be of the a- or b-type, which share only about 50% identity. The b-subunits may autoassemble into homotetradecamers, while the a-subunits oligomerize only in the presence of b-subunits (Dickson et al., 2000). Two types of cochaperonins have been described, the conventional Cpn10 (Koumoto et al., 2001) and a head-to-tail fusion of two Cpn10 domains that give a subunit of 21 kDa, which is termed Cpn20 (Bertsch et al., 1992). Protein folding takes place within the confined cavity formed by the chaperonin and cochaperonin rings, where the chaperonin subunits cycle ATP dependently between binding-active and folding-active states (Bukau and Horwich, 1998). The most renowned chloroplast protein that requires folding by chaperonins to assume the native, oligomerization-competent state is the Rubisco large subunit (Boston et al., 1996).

1.5. sHSPs The characteristic feature of members of the small Hsps (sHsps) is the a-crystallin domain of  90 amino acids at their C-termini (Haslbeck et al., 2005; Sun and MacRae, 2005). sHSPs are strongly induced by heat shock and bind partially denatured proteins, thereby preventing irreversible protein aggregation during stress. The presence of sHSPs in protein aggregates facilitates the access of ATP-dependent chaperones, like ClpB and Hsp70, which catalyze the refolding of the complexed, denatured proteins back to the native state (Mogk et al., 2003). Arabidopsis possesses at least 19 sHSPs (Scharf et al., 2001), which arose from duplication and subsequent diversification of a nuclear sHSP gene in a process that is thought to have occurred only in the green lineage (Waters et al., 1996). The sHSPs in Chlamydomonas are more homologous to one another than to any of the sHSPs in Arabidopsis, suggesting that diversification occurred after divergence of the ancestors of algae and higher plants (Schroda, 2004; Waters and Rioflorido, 2007; Waters and Vierling, 1999; Waters et al., 1996). In contrast, members of the Hsp60/Hsp10, Hsp70/Hsp70 cochaperones,

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and Hsp100 families in Chlamydomonas are more closely related to the Arabidopsis homologs of same intracellular localization than to other members of that family found in other compartments (Schroda, 2004). In the case of the organellar chaperone families, this is because they are derived from genes transferred from the a-purple bacterial and cyanobacterial ancestors of mitochondria and chloroplasts, respectively, to the nucleus of the eukaryotic host.

1.6. Chlamydomonas reinhardtii as a model organism for studying plant molecular chaperone functions Chlamydomonas reinhardtii has been used as a model organism for many decades, mainly to study photosynthesis and flagella/cilia, as the photosynthetic apparatus and flagella of Chlamydomonas are closely related to the corresponding structures of higher plants and metazoans, respectively (Rochaix, 2002; Snell et al., 2004). Moreover, Chlamydomonas has received much attention recently because of its elaborated fermentation pathways (Mus et al., 2007) and because of its ability to produce hydrogen (Melis et al., 2000) and nonpolar lipids (Hu et al., 2008) that have promise as biofuels. All three Chlamydomonas genomes are sequenced (nuclear, chloroplast, and mitochondrial), and it is the only organism in which all three genomes may be genetically manipulated (Merchant et al., 2007). A large number of molecular tools have been established for Chlamydomonas, including selectable marker genes (Lumbreras et al., 1998; Sizova et al., 2001), strong promoters (Fischer and Rochaix, 2001; Schroda et al., 2000), codon-adapted reporter genes (Fuhrmann et al., 1999; Shao and Bock, 2008), insertional mutagenesis methods (Tam and Lefebvre, 1993), and vectors to induce RNAi (Schroda, 2006) or to express artificial microRNAs (Molnar et al., 2009; Schmollinger et al., 2010; Zhao et al., 2009). An important consideration is that gene families in general are much smaller in Chlamydomonas than in higher plants. For example, the members of the five major chaperone systems, Hsp100, Hsp90, Hsp70, Hsp60, and sHsps, are encoded by a total of 74 genes in Arabidopsis but only 39 genes in Chlamydomonas (Schroda, 2004; Schroda and Vallon, 2008). Finally, protein biochemistry approaches are facilitated by the ease and rapidity with which large amounts of Chlamydomonas biomass can be harvested, owing to the short generation time of 8 h (Harris, 2008). Hence, Chlamydomonas is an ideal model organism to study plant molecular chaperone functions and stress response pathways. Another advantage of Chlamydomonas as a model system is that it is closely related to other algae that lend themselves to the study of the role of chaperones in multicellular development and its evolution. The beststudied multicellular cousin of Chlamydomonas reinhardtii is Volvox carteri, which possesses 2000 cells but just two cell types: many small, somatic cells that closely resemble Chlamydomonas unicells and that are specialized

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for motility, and 16 large reproductive cells called gonidia (Kirk, 1998; Starr, 1970). The Volvox life cycle comprises events that have clear parallels in higher plants and/or animals, making it an excellent system in which to study fundamental developmental processes. For instance, when mature, gonidia initiate a program of embryogenesis that includes several rounds of asymmetric cell divisions that generate large cells that are the gonidial precursors of the next generation (Kirk et al., 1991). Following cell division, the embryo turns itself inside out in a process that resembles gastrulation in metazoans (Kirk and Nishii, 2001; Viamontes et al., 1979). Soon thereafter, the small cells begin to cytodifferentiate as somatic cells, while the large cells become specialized for reproduction (Starr, 1970). Importantly, each of these events, and in fact virtually every major aspect of Volvox development, is highly amenable to molecular genetic analysis. This is because mutants disrupted for normal development are easy to isolate, and because, as for Chlamydomonas, many molecular tools have already been developed for Volvox, including nuclear transformation, reporter genes, a transposon tagging (insertional mutagenesis) system, selectable marker genes, inducible promoters as well as strong constitutive promoters, and vectors for RNAi (Cheng et al., 2006; Gruber et al., 1996; Hallmann and Rappel, 1999; Hallmann and Sumper, 1994, 1996; Jakobiak et al., 2004; Mages et al., 1988; Miller et al., 1993; Schiedlmeier et al., 1994; Ueki and Nishii, 2008). Furthermore, the Volvox nuclear genome has also been sequenced (Prochnik et al., 2010). Hence, the stage is set for analyzing the role of chaperones in Volvox development, and for subsequent comparative studies involving Chlamydomonas to learn how chaperone function may have been co-opted to permit the multicellular way of life in this family. In the sections that follow, we describe recent advances toward understanding chaperone biology in Chlamydomonas and Volvox, integrated where appropriate with recent findings from other systems that complement the algal advances that are our main focus. Our discussion is organized according to the five major cell compartments/organelles in which chaperones are known to function in these algae: cytosol/nucleus, flagella, ER, chloroplast, and mitochondria.

2. Cytosol/Nucleus 2.1. Chaperone components The nucleocytosol of Chlamydomonas appears to contain a single Hsp100 (CLPB1 or HSP101), one Hsp90 (HSP90A), two Hsp70s (DnaK-type HSP70A and Hsp110-like HSP70E) and presumably three small Hsps (HSP22A, HSP22B, and HSP22H). One homolog each of the Hsp90 cochaperones p23 and Hop1 (HOP1) are encoded by the Chlamydomonas

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genome, so it is likely that these interact with HSP90A and that HOP1 organizes complex formation between HSP90A and HSP70A. Strikingly, of the 63 J-domain proteins encoded by the Chlamydomonas genome more than half appear to be localized to the cytosol/nucleus (M. Schroda, unpublished data). As HSP70A is the only DnaK-type Hsp70 in this compartment, HSP70A must be directed to an amazingly large number of diverse functions. Volvox chaperone genes are not as completely annotated as their Chlamydomonas counterparts, but it is known that Volvox possesses three nucleocytosolic Hsp70s, including two DnaK-types, Hsp70A and Hsp70H. The fact that Chlamydomonas does not possess an Hsp70H ortholog raises the question of whether Volvox gained a nucleocytosolic Hsp70 or Chlamydomonas lost one in the time since the two species diverged from a common ancestor. Since Hsp70H is much less similar to Hsp70A and HSP70A than those proteins are to each other (80% identity compared to 98%), and since the unicellular chlorophyte green alga Chlorella, like Volvox, possesses two DnaK-type nucleocytosolic Hsp70s (S. Miller, unpublished data), it seems likely that Chlamydomonas lost an HSP70 gene that existed in the most recent common ancestor of Chlamydomonas and Volvox. However, a more definitive answer to this question will have to await analysis of HSP70 gene families in additional volvocine/chlorophycean species.

2.2. Function of cytosolic/nuclear chaperones 2.2.1. Regulation of the stress response Integration of stress signals and triggering of the cellular stress response take place in the nucleocytosol. In Chlamydomonas, the stress response is mediated to a very large extent by heat shock factor 1 (HSF1) (Schulz-Raffelt et al., 2007). This is inferred from the observation that the induction of many heat shock genes and Hsps is abolished in Chlamydomonas strains in which HSF1 is strongly downregulated by RNAi. As a consequence, HSF1-RNAi strains are thermosensitive. HSP101 and HSP22A are among the chaperones most strongly upregulated by heat shock in Chlamydomonas (T. Mu¨hlhaus, J. Weiss, D. Hemme, F. Sommer, M. Schroda, unpublished data) and therefore, in accordance with data from other organisms (Queitsch et al., 2000; Sanchez and Lindquist, 1990), appear to be central for thermotolerance also in Chlamydomonas. Under nonstress conditions, HSP101 and HSP22A are barely detectable, whereas HSP70A and HSP90A are expressed constitutively at high levels (Schulz-Raffelt et al., 2007). Hence, HSP90A and HSP70A appear to be the chaperones in the nucleocytosol that carry out housekeeping functions and that maintain protein homeostasis under nonstress conditions in this compartment. Moreover, their inducibility by heat shock certainly suggests that HSP70A and HSP90A also contribute to refolding of stress-denatured proteins.

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Some facets of the HSF-mediated stress response in Chlamydomonas and Volvox are typical for developmentally simple eukaryotes like yeast, while others are typical for plants. Chlamydomonas and Volvox are both yeast-like in that they contain only one canonical HSF (HSF1) and also (at least in the case of Chlamydomonas) that HSF1 forms constitutive trimers (SchulzRaffelt et al., 2007; Sorger and Nelson, 1989). In contrast, Arabidopsis possesses 21 HSFs (Nover et al., 2001) and these are present as monomers under nonstress conditions and need to trimerize under stress to gain transcriptional competence (Lee et al., 1995). However, Chlamydomonas HSF1 is plant-like in its sequence homology with plant (class A) HSFs and its heat shock inducibility (Hubel and Schoffl, 1994; Scharf et al., 1990; SchulzRaffelt et al., unpublished data). In contrast, HSFs from yeast and nonplant higher eukaryotes are not inducible by heat shock (Wiederrecht et al., 1988). It appears to be a universal feature of plants, animals, and fungi that HSF transcriptional competence is activated by hyperphosphorylation (Holmberg et al., 2002). Accordingly, heat shock gene expression in Chlamydomonas was also found to strictly correlate with HSF1 hyperphosphorylation (SchulzRaffelt et al., 2007). Feeding of Chlamydomonas cells with kinase inhibitor staurosporine led to a delay of the stress response (Schulz-Raffelt et al., unpublished data). The activation of stress kinase(s) that hyperphosphorylate HSF1 appears to be mediated by the accumulation of unfolded proteins, since feeding Chlamydomonas cells with arginine analog canavanine elicited a strong stress response at ambient temperatures (Schulz-Raffelt et al., unpublished data). Combining heat stress and canavanine feeding led to a very strong stress response that did not attenuate over time. This phenomenon was also observed when heat stress was combined with feeding of cytosolic translation inhibitor cycloheximide. We concluded from these observations that synthesis of additional chaperones upon heat shock is essential for restoring protein homeostasis—and hence for attenuation of the stress response at elevated temperatures. The addition of low concentrations of specific Hsp90 inhibitors radicicol and geldanamycin to Chlamydomonas cultures did not induce a stress response, but resulted in delayed attenuation. These results suggested that Hsp90s in Chlamydomonas play an important role in restoring protein homeostasis, obviously by refolding heat-denatured proteins. In contrast to the response of Chlamydomonas to minor disruption of Hsp90 function, exposure to high concentrations of Hsp90 inhibitors triggered a stress response. This was in line with observations made in other eukaryotes, which suggested that Hsp90 is regulating the stress response (Ali et al., 1998; Duina et al., 1998; Zou et al., 1998). While in Chlamydomonas HSP70A was found to specifically interact with HSF1 (Schulz-Raffelt et al., 2007), it is not entirely clear whether this is the case also for HSP90A. Hence, the two chaperones might regulate activation of HSF1 also in Chlamydomonas, for example, by controlling accessibility of important Ser/Thr stress kinase targets or of their nuclear localization

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sequences. Alternatively, Hsp90 inhibitors might trigger the stress response by eliminating HSP90A as a chaperone component that plays an important role in maintaining protein homeostasis. According to the latter hypothesis, stress kinase(s) might become active in response to disturbances of protein homeostasis and hyperphosphorylate HSF1 during the time period in which protein homeostasis is out of balance. During this time period, temporally active kinases may exceed the function of phosphatase(s) that constitutively dephosphorylate HSF1. The connection between protein homeostasis and kinases might be mediated by chaperones (e.g., HSP90A and HSP70A) that maintain stress kinase(s) in an inactive state. The latter hypothesis is attractive because many kinases are client proteins of cytosolic Hsp90 (Pratt and Toft, 2003; Wegele et al., 2004). Interestingly, when Chlamydomonas cells that express HSP70B-antisense constructs were exposed to heat shock, they displayed a retarded general stress response that resembled the staurosporine feeding phenotype (SchulzRaffelt et al., unpublished data). Presumably, the disturbance of plastidic protein homeostasis that is a consequence of HSP70B underexpression is perceived by the nucleus and triggers a weak stress response, eventually resulting in a desensitizing of the HSF1-dependent stress response pathway. If HSP70B were involved in chloroplast protein import, as recently suggested for Physcomitrella (Shi and Theg, 2010), cytosolic protein homeostasis in HSP70B-antisense strains might also be disturbed by the accumulation of import precursors. We have constructed a mathematical model of the stress response in Chlamydomonas that explains all of the observations detailed above (SchulzRaffelt et al., unpublished data): delay of the stress response by kinase inhibitors, triggering of the response by canavanine; lack of attenuation by canavanine or cycloheximide feeding; time lag required for regaining competence for a full stress response following prior exposure to stress (Schroda et al., 2000). Challenging the predictions made by the model by experiments will reveal whether our understanding of the stress response in Chlamydomonas is correct. 2.2.2. Hsp70A and division symmetry in Volvox embryos Whereas no more functions have been identified for chaperones in the nucleocytosol of Chlamydomonas, a highly specific function in the regulation of unequal division of embryonic cells has been reported for the V. carteri J protein GlsA and its Hsp70 partner, Hsp70A. The glsA (gonidialessA) gene was identified in a transposon-tagging screen for mutants defective for the asymmetric divisions that generate large cells that are the progenitors of gonidia, the asexual reproductive cells of Volvox (Miller and Kirk, 1999). GlsA is a zuotin-family J protein that possesses three evolutionarily conserved domains in addition to the N-terminal J domain—a central M domain and two C-terminal SANT (SWI3, ADA2, N-CoR, and

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TFIIIB) domains. Zuotin family members have been implicated in regulation of both translation and transcription. For instance, numerous SANT-domain proteins bind histones and/or are involved in chromatin structure (Badri et al., 2008; Boyer et al., 2002, 2004; Guelman et al., 2006; Humphrey et al., 2001; Mo et al., 2005; Shi et al., 2005a; Wang et al., 2008), and mouse SANT-domain protein MIDA1 enhances transcription of reporter genes when coexpressed with its Hsp70 partner in a cell culture transient expression assay (Yoshida et al., 2004). However, yeast zuotin and human MPP11 are essential members of a ribosome-associated chaperone complex (RAC) that localizes to the exit tunnel of the ribosome and is required for normal translation fidelity and termination; consistent with this fact, zuotin mutants are ultra sensitive to the ribosome-binding aminoglycoside antibiotic paromomycin, which compromises translation accuracy (Gautschi et al., 2002; Otto et al., 2005; Rakwalska and Rospert, 2004). Interestingly, glsA mutants are also ultrasensitive to paromomycin, suggesting that like zuotin and MPP11, GlsA is very likely a ribosome-associated chaperone that regulates translation (Pappas and Miller, 2009). GlsA also interacts with histones in coimmunoprecipitation assays, with the association dependent on the C-terminal SANT domain. Variants defective for this SANT domain are just as resistant to paromomycin as wild-type GlsA, but are not able to rescue the asymmetric division defect of glsA mutants. This result suggests that the translation function of GlsA may not be important for asymmetric division, and implies that GlsA likely regulates the activity of one or more transcription factors that regulate cell division symmetry. Consistent with this view, the majority of GlsA in cleaving embryos localizes to the nucleus (Cheng et al., 2005; Miller and Kirk, 1999). However, in asymmetric division-stage embryos, GlsA appears to be just as abundant in cells that divide symmetrically as it is in those that cleave asymmetrically. So how might GlsA be involved in the decision to divide asymmetrically versus symmetrically? The answer might be that the Hsp70 partner of GlsA, Hsp70A, has the more important say in this decision. Like GlsA, Hsp70A colocalizes with chromatin and is equally abundant in all cells of early embryos, but at the 32-cell stage, just prior to the stage when cells in the anterior half of the embryo undergo asymmetric division, Hsp70A becomes significantly more abundant in anterior cells than in posterior ones (Cheng et al., 2005). It is unknown how this Hsp70A asymmetry is effected, or what the substrates of GlsA/Hsp70A action are that determine which cells divide asymmetrically, but it is intriguing that a typically ubiquitous chaperone like Hsp70A may accumulate in temporal and spatial-specific fashion to regulate an important developmental process, such as asymmetric cell division.

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3. Flagella 3.1. Chaperone components Using an indirect immunolocalization approach, Johnson and coworkers (Bloch and Johnson, 1995; Shapiro et al., 2005) found HSP70A in Chlamydomonas flagella. A potential cochaperone of HSP70A could be the radial spoke J-domain protein radial spoke protein 16 (RSP16) (Yang et al., 2006) (Fig. 2.1). HSP90A was also identified in Chlamydomonas flagella, via a proteomics approach (Pazour et al., 2005).

3.2. Function of flagellar chaperones 3.2.1. HSP70A in folding and delivery of flagellar proteins Not surprisingly, transcripts that encode flagellar proteins accumulate at elevated levels during flagellar regeneration, but interestingly HSP70A and HSP90A transcripts are also induced by deflagellation (Baker et al., A

B 1, 4, 6, 9, 10 5 A ID

xin Ne A OD

A B

Head

5

Central sheath

C1

16 16

Stalk

RS

23 23 2

Figure 2.1 Half cross section of a flagellar axoneme. (a) A flagellum consists of nine peripheral doublet microtubules (A and B) and two central microtubules (C1 and C2). This highly conserved “9þ2” structure forms the core of the axoneme, or flagellar cytoskeleton. Each A microtubule anchors three outer dynein arms (ODAs) and two inner dynein arms (IDAs), which are responsible for flagellar locomotion. The elastic protein, nexin, maintains this organization during the motion of the doublet microtubules. RS, radial spoke. (b) The T-shaped RSs are associated with A microtubules and transiently interact with the central sheath, which protects the central tubules. It is postulated that the RSs are involved in controlling flagellar motility (Yang et al., 2005). The RSs consist of 23 proteins (radial spoke protein (RSP) 1–RSP23) (Yang et al., 2001, 2005). RSP1, 4, 6, 9, and 10 form the spoke head, and the rest of the RSPs are integrated in the spoke stalk (Yang et al., 2005). Homodimeric J-domain protein RSP16 may interact with dimeric RSP2 and/or RSP23 in the stalk (Yang et al., 2008).

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1986; Stolc et al., 2005). However, transcripts coding for other cellular chaperones like chloroplast HSP70B are also induced during regeneration, presumably resulting from a general stress response triggered by pH shock-induced deflagellation (Stolc et al., 2005). Immunofluorescence analysis of a transgenic strain that expresses epitope tagged HSP70A revealed that HSP70A localizes in the cell body and, with a discontinuous, punctuate pattern, along the flagellum. Remarkably, highest HSP70A concentrations were observed in the flagellar tip (Bloch and Johnson, 1995; Shapiro et al., 2005) where flagellar assembly takes place ( Johnson and Rosenbaum, 1992). Bloch and Johnson speculated that tubulin or other axonemal proteins may preassemble with HSP70A in the cytoplasm and are transported by IFT to the flagellar tip. Dual labeling of HSP70A and the anterograde IFT kinesin-II, FLA-10, revealed that the distribution of both proteins overlaps along the length of the flagellum, although the label stoichiometry varied considerably. This variation could occur because HSP70A is not an integral part of the IFT machinery but rather is carried by IFT as a cargo (Shapiro et al., 2005). After deflagellation, Chlamydomonas is able to reassemble its flagella within minutes (Randall et al., 1969; Rosenbaum et al., 1969). Such a fast assembly process is made possible by storage of flagellar precursor proteins and IFT particles in the cell body. These proteins are concentrated in the region of the basal body where they are preassembled before entering the flagellum (Rosenbaum and Witman, 2002; Snell et al., 2004). For example, the exclusively cytoplasmic protein PF13 is believed to be involved in the preassembly of outer dynein arm (ODA) heavy chains with other subunits (like IC1 and IC2) (Omran et al., 2008). Flagella from the Chlamydomonas pf13 mutant are paralyzed due to cytoplasmic depletion of all three ODA heavy chains and an important motor subunit (HC9) of inner dynein arms (IDAs) (Huang et al., 1979; Kamiya, 2002; Omran et al., 2008). Interestingly, Ktu, a homolog of PF13 in vertebrates, coimmunoprecipitates with an Hsp70. It was suggested that Ktu/PF13 might cooperate with Hsp70 in dynein preassembly (Omran et al., 2008), an idea that is consistent with the notion that Chlamydomonas HSP70A preassembles with axonemal proteins in the cytoplasm. At the distal tip of the flagella, a capping complex controls the elongation process of the axonemal microtubules (Dentler and LeCluyse, 1982). Presumably, HSP70A is transiently associated with the capping complex while releasing fully folded flagellar proteins for incorporation into the distal ends of elongating axonemes. Another fraction of HSP70A is associated with the central pair protein CPC1 in the central pair C1b projection that anchors enzymes for ATP synthesis. Mitchell et al. (2005) propose that this fraction of HSP70A might mediate a conformational switch for central pair regulation of flagellar motility.

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3.2.2. Radial spoke protein 16 RSP16 has all characteristics of type II J-domain proteins, in that it contains at its N-terminus the highly conserved DnaJ J domain followed by a G/Frich region and a DnaJ C-terminal domain, but lacks the cysteine-rich domain typical for type I J-domain proteins (Cheetham and Caplan, 1998; Satouh et al., 2005; Yang et al., 2005). The DnaJ C-terminal domain is responsible for dimerization and for substrate protein binding (Sha et al., 2000; Shi et al., 2005b). Yang and coworkers identified homodimeric RSP16 as a structural component of the T-shaped radial spokes (Qin et al., 2004; Yang et al., 2005). Before assembling at the distal tip, RSP16 is transported separately from the 12S spoke precursors, which are first preassembled in the cell body. Yang et al. (2005) speculated that HSP70A and its supposed cochaperone RSP16 convert the 12S spoke precursor into the bigger 20S structural complex. However, their recent data from an RNAi-knockdown experiment indicate that RSP16 is presumably not involved in flagellar biogenesis (Yang et al., 2008). The flagella of RSP16-RNAi lines have normal length, with the only noticeable structural phenotype being that the spokehead region of the radial spokes appears less defined. The motility defect of rsp16 knockdown flagella, which display an abnormal twitching phenotype, is also immediately rescued by electroporated his-tagged RSP16 protein (RSP16-his). These findings imply that one or more other J-domain proteins aid HSP70A to fulfill its function in flagellar assembly. Interestingly, the J domain of RSP16 is not essential for flagellar motility, as rsp16 cells were complemented with electroporated RSP16-his and RSP16DJ-his (Yang et al., 2008). However, RSP16DJ-his-complemented cells exhibited a slightly slower swimming velocity compared to RSP16his-complemented cells. The fact that the normally highly conserved HPD motif of the J domain, which is essential for interaction with Hsp70, is poorly conserved among the spoke RSP16 orthologs also supports the idea that the J domain of RSP16 is not important for its function. As to what that function is, RSP16 may be important for stabilizing the radial stalk and head. RSP16 appears to be localized at the junction between spokehead and stalk, where it might interact with RSP2 and/or RSP23 (Huang et al., 1981; Piperno et al., 1981) (Fig. 2.1). This suggestion is based on the result that depletion of RSP2 also leads to the reduction of RSP16 and RSP23 (Yang et al., 2004). Moreover, RSP16 could be important for the coordination between the central pair, radial spoke, and the dynein motors. Previous structural and genetic analyses indicated that the radial spoke interacts with the central pair apparatus to control the dynein motors by transmitting signals from the central pair through mechanical and/or mechanochemical interactions with the microtubules (Mitchell and Nakatsugawa, 2004; Smith and Yang, 2004). If components of the radial

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spoke are depleted, the flagella are paralyzed or show abnormal motility, presumably caused by a perturbation in signaling (Witman et al., 1978). 3.2.3. Radial spoke protein 12 Using mass spectrometry, Yang et al. (2006) identified several RSPs, including RSP12, a predicted member of the cyclophilin family of peptidyl-prolyl cis–trans isomerases (PPIs). The function of PPIs is to assist protein folding by catalyzing proline cis–trans isomerization (Fink, 1999). By specifically binding to Ser-Pro or Thr-Pro motifs, when serine or threonine is phosphorylated (Yaffe et al., 1997), PPI affects substrate conformation and thereby alters the activity of target proteins (Zacchi et al., 2002; Zheng et al., 2002). Several RSPs (RSP2, 3, 5, and 17) have Ser-Pro/Thr-Pro motifs, which are also targets of phosphorylation. Therefore Yang and coworkers suggested that RSP12 might be a protein with regulatory function.

4. Endoplasmic Reticulum 4.1. Chaperone components The ER of Chlamydomonas appears to contain three Hsp70 members, two DnaK-like proteins termed BIP1 and BIP2, and one Hsp110-like protein termed HSP70G. In addition, there appears to be one Hsp90 member in the ER, termed HSP90B (Willmund and Schroda, 2005). Moreover, approximately six J-domain cochaperones presumably specific to the BIPs might be present in the ER (M. Schroda, unpublished data). The amino acid sequences of BIP1 and BIP2 are 92% identical and 95% similar, suggesting that their genes were duplicated rather recently. Accordingly, both genes are located in a cluster, which also includes the HSP90B gene (Schroda, 2004; Schroda and Vallon, 2008).

4.2. Function of ER-localized chaperones Expression of the BIP1 gene is strongly induced by light (Vasileuskaya et al., 2004) and by heat shock (Schulz-Raffelt et al., unpublished data). Interestingly, heat shock inducibility of BIP1 was observed also in HSF1-RNAi strains, suggesting that the stress response in the ER of Chlamydomonas is not HSF1-dependent, but might be mediated by the unfolded protein response (UPR) (Spear and Ng, 2001). Accordingly, a homolog of inositol-requiring kinase 1 (IRE1)—an essential component of the UPR—is encoded by the Chlamydomonas genome.

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5. Chloroplast 5.1. Chaperone components Chlamydomonas chloroplasts contain at least one member of each of the five major chaperone classes (Schroda, 2004; Schroda and Vallon, 2008): at least three Hsp100s (CLPB3 and/or CLPB4, CLPC, and CLPD), one Hsp90 (HSP90C), up to three Hsp70s (HSP70B and perhaps HSP70D and/or HSP70F), three chaperonins and three cochaperonins (CPN60A, CPN60B1, CPN60B2, CPN11, CPN20, and CPN23), and up to four small Hsps (HSP22C–F). Moreover, there exist two GrpE-type cochaperones of chloroplast Hsp70s (chloroplast GrpE; CGE1 and CGE2), one Hsp70 cofactor (HSP70 Escort Protein 2, HEP2), and up to 13 J-domain proteins. Of the latter, five (CDJ1–5) have been confirmed to be chloroplast-targeted (see below). As judged from cDNA coverage and proteomics results (T. Mu¨hlhaus, J. Weiss, D. Hemme, F. Sommer, M. Schroda, unpublished data), HSP70B is by far the most abundant Hsp70 in the chloroplast of Chlamydomonas. Though poorly abundant, cDNAs for HSP70D and F have been detected, and hence these species must be expressed. Their localization to the chloroplast has not been demonstrated, but is inferred from the fact that the best BLAST hits for both are Hsp70s from plant stroma and cyanobacteria, and that HSP70D and F have N-terminal extensions that might be chloroplast transit peptides (Schroda, 2004; Schroda and Vallon, 2008). Also supporting the idea that Chlamydomonas has at least two chloroplast Hsp70s is the presence of two GrpE homologs (CGE1 and CGE2) with signatures typical for chloroplast members of this type of Hsp70 cochaperone (Schroda, 2004; Schroda et al., 2001b). Whereas CGE1 is the major cochaperone of HSP70B, CGE2 might cooperate with HSP70D and/or HSP70F.

5.2. Function of chloroplast (co)chaperones 5.2.1. Mystery concerning stress protection of photosystem II by HSP22 Early studies reported that HSP22A is localized in thylakoid membranes and plays a role in the protection of photosystem II (PS II) against photoinhibition (Kloppstech et al., 1985; Schuster et al., 1988). The import of HSP22A into the chloroplast was suggested to take place without a processing step (Grimm et al., 1989). The idea that sHSPs are able to protect PS II activity was supported by a report showing that a tomato chloroplast-targeted sHSP protected PS II electron transport during heat stress (Heckathorn et al., 1998). However, this finding could not be reproduced with recombinant chloroplast-targeted pea sHSP (Harndahl and Sundby, 2001). Moreover,

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thermoprotection of PS II activity as measured by oxygen evolving capacity in Chlamydomonas appears to be mediated by proteins induced at a temperature of 35  C, which apparently is below that required to induce HSP22A (Tanaka et al., 2000). Later work revealed that Chlamydomonas HSP22A is a cytosolic protein that becomes associated with the chloroplast outer envelope during heat shock and dissociates during recovery (EisenbergDomovich et al., 1994). Hence, the proposed thylakoid association of HSP22A might have arisen from contamination of thylakoids by envelopes. However, in Synechocystis a sHSP has been shown to specifically associate with thylakoid membranes upon heat shock, leading to membrane rigidification and potentially reduced thermodamage of PS II (Tsvetkova et al., 2002). In summary, while no role has been firmly established for Chlamydomonas HSP22A in protecting PS II against photoinhibition, its association with the chloroplast outer envelope might at least improve the thermotolerance of processes occurring at this membrane, such as protein import. More recent studies using microarrays have shown that the putatively chloroplast-targeted HSP22E and HSP22F are induced by oxidative stress (Fischer et al., 2005), sulfur starvation (Zhang et al., 2004), and phosphorus starvation (Moseley et al., 2006). These findings suggest that Chlamydomonas sHSPs appear to protect plastidic processes also from stresses other than heat stress. In any event, the exact role of HSP22 species in protecting PS II and other chloroplast functions from heat and/or other stresses remains unresolved. 5.2.2. CPN60 functions Chloroplast CPN60A, CPN60B1, and CPN60B2 all are induced by heat shock, both at the mRNA and protein levels (Tanaka et al., 2000; Thompson et al., 1995), suggesting a role of these chaperonins in refolding of stress-denatured proteins. Interestingly, CPN60A was also found to specifically interact with group II intron RNA via its two ATPase domains and was suggested to play a role as a general organelle splicing factor (Balczun et al., 2006). 5.2.3. Comparison of CGE1 and bacterial nucleotide-exchange factor GrpE CGE1 is a chloroplast GrpE homolog and, being the most prominent interaction partner of chloroplast HSP70B in its ADP-bound state (Schroda et al., 2001b), appears to be its nucleotide-exchange factor (Fig. 2.2). HSP70B (68 kDa) and CGE1 (24 kDa) constitute about 0.19% and 0.01% of Chlamydomonas total cell protein, respectively (Liu et al., 2007; Schroda et al., 2001b). When taking into account the molecular weights of the proteins, this yields a molar ratio of 6.7:1. Somewhat surprisingly, CGE1 is only 32% identical at the primary amino acid sequence level to its Escherichia coli homolog, GrpE. Yet CGE1 shares a

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Figure 2.2 The components and substrates of the chloroplast HSP70B/HSP90C chaperone system in Chlamydomonas. The major Hsp70 in the chloroplast of Chlamydomonas is HSP70B (light blue) (Drzymalla et al., 1996). It requires HEP2 (HSP70 Escort Protein 2, blue) to attain the functional state (Willmund et al., 2008b). Most likely, HEP2 acts by folding HSP70B to the native state following cleavage of its transit peptide after import (1). HSP70B was identified as a thioredoxin (trx) substrate (Lemaire et al., 2004) and may become glutathionylated (GS) (2) (Michelet et al., 2008). HSP70B appears to form constitutive complexes with dimeric HSP90C (purple), the chloroplast HSP90 homolog of Chlamydomonas (3) (Willmund and Schroda, 2005; Willmund et al., 2008a). In ATP-driven cycles, Hsp70B, presumably supported by HSP90C, induces conformational changes into substrate proteins that are important e.g. protein folding or assembly/disassembly of protein complexes. Substrates are delivered to HSP70B by one of its CDJ (Chloroplast DnaJ-like protein, green) cochaperones (4). CDJ1 is a functional homolog of bacterial DnaJ and thus likely delivers unfolded substrates (gray) for general folding to HSP70B/HSP90C. Chloroplast proteins involved in (light) signal transduction may depend on chaperoning by HSP70B/ HSP90C (5a) (Cao et al., 2003; Schroda and M€ uhlhaus, 2009). CDJ2 is a specialized J-domain protein that delivers the various assembly states of VIPP1 (Vesicle-Inducing Protein in Plastids, light red) to HSP70B and HSP90C. The latter may catalyze the assembly of VIPP1 monomers/dimers to rings and the disassembly of rods and rings (5b) (Liu et al., 2005, 2007). CDJ3–5 contain redox-active Fe–S clusters and CDJ3 is in complex with RNA, presumably to recruit HSP70B for mediating redox-dependent

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number of important structural features with GrpE (Willmund et al., 2007), including the ability to form dimers, and a three-dimensional architecture that consists of a paired a-helix, a four-helix bundle, and a b-sheet domain arranged in the same N-to-C configuration (Fig. 2.3) (Harrison, 2003). Moreover, CGE1 complements the temperature sensitive growth phenotype of an E. coli strain lacking GrpE and interacts with E. coli DnaK/Hsp70 (Schroda et al., 2001b; Willmund et al., 2007). Therefore, despite the low sequence conservation and substantial evolutionary distance between CGE1 and bacterial GrpE, the proteins are quite similar at both the structural and functional levels. However, CGE1 and GrpE also differ in at least three important respects. First, CGE1 exists as two isoforms, a and b, which differ in that CGE1b possesses a valine–glutamine dipeptide at positions 4 and 5 of the mature protein, which CGE1a lacks (Schroda et al., 2001b). This difference is due to a temperature-dependent alternative splicing of the CGE1 transcript, with CGE1b transcript and protein levels increasing upon heat shock (Willmund et al., 2007). Curiously, the two isoforms have different affinities for HSP70B: the affinity of CGE1b is about 25% higher than that of CGE1a, indicating that the CGE1 extreme N-terminus plays an important role in determining the affinity of the cochaperone for HSP70B. However, the functional significance of this finding is not yet understood. Two additional important differences between CGE1 and GrpE relate to their N-termini and dimer formation. According to homology modeling the N-terminus of CGE1 contains a coiled-coil motif (Willmund et al., 2007), as opposed to the unstructured N-terminus of GrpE (Fig. 2.3). Since coiled-coils mediate tight protein–protein interactions, it seemed reasonable to surmise that CGE1 dimer formation might be dependent on this motif. Note that dimerization of E. coli GrpE is mediated by the four-helix bundle at the posterior part of the molecule (Harrison et al., 1997). In fact, removal of half of CGE1’s N-terminal coiled-coil domain was sufficient to abolish its ability to form dimers at 30  C, and to complement the growth phenotype of a DgrpE E. coli mutant. Deletion of the entire coiled-coil led to a complete abolishment of CGE1 dimer formation and loss of a-helical secondary structure, whereas neither was impaired upon removal of the four-helix bundle. Hence, although general structural and functional properties of GrpE and CGE1 appear to be conserved, the proteins have clearly evolved remodeling of translation initiation complexes (5c) (Dorn et al., 2010). CGE1 (Chloroplast GrpE homolog 1, orange) binds to HSP70B in the ADP-state and catalyzes the exchange of ADP by ATP (6) (Schroda et al., 2001b). CGE1 dimers form via coiled-coil interactions of their N-terminal a-helices (Willmund et al., 2007).

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b-Sheet domains

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Figure 2.3 Structural models of dimeric CGE1 and GrpE. The GrpE model is based on a crystal structure of the E. coli GrpE dimer bound to its HSP70 chaperone partner DnaK (PDB entry 1DKG.pdb) (Harrison et al., 1997). Since the N-terminal 33 amino acids of GrpE were unstructured and had to be removed for crystallization, they are not shown here; because of their low resolution in the crystal structure, loops connecting helices of the four-helix bundle in GrpE are also not shown. The CGE1 model was generated by homology modeling using structures of E. coli GrpE and tropomyosin (Whitby and Phillips, 2000; Willmund et al., 2007). Due to ambiguous predictions of the secondary structure formed by the residues between Pro13 and Pro21, this region was drawn as a random coil. Also the positions of the N-terminal a-helices predicted between Ala1 and Ala12 are drawn arbitrarily.

somewhat differently. It seems quite possible that at least some of these differences, such as the modification of CGE1’s extreme N-terminus by differential splicing (and therefore fine-tuning of its affinity for HSP70B) may represent adaptations of CGE1 to the specialized folding requirements of HSP70B–CGE1 in the chloroplast. 5.2.4. HEP2 activates chloroplast HSP70B For a number of years, the biochemical characterization of HSP70B was severely hampered because recombinant protein expressed in E. coli, unlike HSP70B purified from Chlamydomonas, could not form complexes with

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Zn2+ S.cerevisiae Hep1p

Figure 2.4 Structural model of yeast mitochondrial Hep1p. The Hep1p model is based on an NMR structure of Hep1p (PDB entry 2E2Z.pdb) (Momose et al., 2007).

CGE1 (Willmund et al., 2007). Sichting et al. (2005) reported that a specialized protein termed Hsp70 escort protein 1 (Hep1) is essential for maintaining the functionality of yeast mitochondrial Hsp70s Ssq1 and Ssc1, by preventing their aggregation. NMR studies have revealed that Hep1 is an L-shaped molecule (Momose et al., 2007). One leg of the “L” is formed by two two-stranded, antiparallel b-sheets which, at their connecting loops, each contain two cysteines that together coordinate one Zn2þ ion (Fig. 2.4). The latter appears to stabilize the structure of the protein. The existence of an Hsp70 escort protein in mitochondria suggested the existence of a Hep homolog also in chloroplasts, which would explain why recombinantly expressed chloroplast HSP70B was nonfunctional. Database searches indeed revealed potentially chloroplast-targeted Hep1 homologs in algae, moss, and higher plants (Willmund et al., 2008b). The homolog in Chlamydomonas was termed HEP2. HEP2 is a constitutively (but weakly) accumulating stromal protein with a calculated molecular weight of 14 kDa and apparent molecular weight of 21 kDa. HEP2 appears to form dimers and compared to CGE1 interacts as a minor partner with HSP70B, preferably in the ADP-bound state. As hoped, active HSP70B, that is, HSP70B capable of interacting with CGE1 and in a proteaseresistant conformation, could be quantitatively produced in E. coli when coexpressed with HEP2. HEP2 binds to active and inactive HSP70B, but cannot activate inactive forms of HSP70B such as HSP70B expressed in E. coli without HEP2. The need of mitochondrial and chloroplast Hsp70s for escort proteins is surprising, as no such proteins are required for ensuring functionality of the homologous Hsp70s in bacteria.

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Hep1 was proposed to interact with mitochondrial Hsp70s to maintain them in a functional conformation, by preventing their aggregation (Momose et al., 2007; Sanjuan Szklarz et al., 2005; Sichting et al., 2005). This view was not supported by data reported for chloroplast HSP70B– HEP2 (Willmund et al., 2008b). Instead of aggregating, HSP70B expressed in the absence of HEP2 assumed a protease-sensitive, inactive configuration. Active HSP70B that was coexpressed with HEP2 remained active for 48 h in the absence of HEP2, suggesting that HEP2 was not involved in maintaining HSP70B in an active state. Moreover, some HSP70B expressed with an N-terminal extension became activated if HEP2 was present during cleavage of the fusion protein, suggesting that in vivo HEP2 might be required for de novo folding of HSP70B after transit peptide cleavage (Fig. 2.2). Clearly, in order to better understand Hep1 and HEP2 function, more investigation is required to elucidate the molecular mechanism(s) by which these Hsp70 escort proteins activate their Hsp70 partners in mitochondria and chloroplasts, and to learn why such activation is required. 5.2.5. HSP70B activity might be redox-controlled Oxidative stress has been shown to cause multiple protein modifications (Berlett and Stadtman, 1997) that lead to the increased expression of molecular chaperones and proteases (VanBogelen et al., 1987). However, in yeast and other organisms, oxidative stress also results in a dramatic drop in cellular ATP levels, which precludes ATP-dependent folding by molecular chaperones (Osorio et al., 2003; Winter et al., 2005). Accordingly, mammalian cytosolic Hsc70 in the nucleotide-free state performed significantly better in preventing protein aggregation when it was glutathionylated compared to its unmodified conformation (Hoppe et al., 2004). In the presence of ATP, the performance of glutathionylated and unmodified Hsc70 were similar. Recent studies aimed at the identification of Chlamydomonas proteins that are targets for thioredoxins and glutathionylation revealed chloroplast HSP70B as a target for both (Lemaire et al., 2004; Michelet et al., 2008). This finding suggests that the activity of chloroplast HSP70B might be regulated by the redox-state of the chloroplast (Fig. 2.2). 5.2.6. HSP70B and CDJ3–5 as redox switches controlling translation initiation? CDJ3–5 are chloroplast-targeted J-domain proteins that contain bacterial 4Fe–4S clusters (Dorn et al., 2010). Genes encoding homologs of CDJ3–5 are present in all members of the green lineage that have been sequenced so far, but are not present in cyanobacterial genomes. Phylogenetically, CDJ3–5 homologs split into two clades, and at least one representative of each clade is encoded by every member of the green lineage examined. In Chlamydomonas, CDJ3–4 are members of one clade, and CDJ5 is a member of the second. CDJ3–5 transcripts and proteins are rather weakly

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expressed. While expression levels increase in the light, they decrease after heat shock, suggesting that CDJ3–5 do not play roles in folding of stress-denatured proteins. The Arabidopsis CDJ5-like gene is strongly regulated by the circadian clock, and the Arabidopsis CDJ4-like gene is induced by iron deficiency. Redox-active Fe–S clusters are assembled in E. coliexpressed CDJ3–4, and both proteins interact in vitro with purified HSP70B only when it is in the ATP-bound state. CDJ3 is localized to the stroma, whereas CDJ4 is associated with thylakoid membranes. The suborganellar localization of CDJ5 is not known. Interestingly, genes encoding proteins that contain a J-domain and a bacterial-type Fe–S cluster within the same polypeptide have also been found in mesophilic Crenarchaeota (or Thaumarchaeota), albeit with different domain organization than encoded by the CDJ3–5 genes of the green lineage (Dorn et al., 2010). Most likely, the CDJ3–5 genes were transferred to the archaebacteria by horizontal gene transfer, and in those organisms carry out functions similar to those of CDJ3–5 in the green lineage. CDJ3 associates with RNA (Dorn et al., 2010), a function that might point to a chaperone-mediated remodeling of RNA-binding protein complexes that, for example, are involved in translation initiation/elongation or mRNA stability. Such complexes are found in the stroma and associated with thylakoids (Marı´n-Navarro et al., 2007), where CDJ3–4 and HSP70B are also located. Well-studied examples for chaperone-mediated remodeling of replication initiation occur in E. coli, where DnaK and DnaJ monomerize RepA dimers and dissociate DnaB-helicase–Lambda P complexes to trigger replication of plasmid P1 and lambda phage, respectively (Alfano and McMacken, 1989; Wickner et al., 1991). As posttranscriptional regulation of the expression of many chloroplast genes is strongly regulated by light (Marı´n-Navarro et al., 2007), it is possible that CDJ3–5 represent nuclearencoded factors that act as redox switches by recruiting HSP70B for the reorganization of regulatory protein complexes (Fig. 2.2). 5.2.7. HSP70B–CDJ2–CGE1 in assembly and disassembly of VIPP1 complexes VIPP1 (Vesicle-Inducing Protein in Plastids) is named after the phenotype of Arabidopsis mutants that strongly underexpress VIPP1 (Kroll et al., 2001): mutant plants were deficient in the formation of vesicles and unable to grow photoautotrophically because they displayed a distorted thylakoid structure. In wild-type plants, vesicles bud off from the inner envelope presumably to transport galactolipids to the thylakoids (Benning et al., 2006; Vothknecht and Soll, 2005). Like its homolog, the phage shock protein A (PspA) (Hankamer et al., 2004), the 28-kDa VIPP1 was shown to assemble into rotationally symmetric rings of >1 MDa (Aseeva et al., 2004). In contrast to PspA, Chlamydomonas VIPP1 rings (and to some extent also Synechocystis VIPP1 rings (Fuhrmann et al., 2009a)) were able to assemble

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into rod-like supercomplexes (Liu et al., 2007). While rod-like supercomplexes were also detected in cell extracts, it is not clear whether they exert a function in the chloroplast or simply represent a storage form of VIPP1. CDJ2 is a specialized J-domain protein that is mainly localized to the chloroplast stroma (Liu et al., 2005). Like HEP2 and CDJ3–5, CDJ2 is present only in the green lineage and in its C-terminus contains conserved motifs predicted to mediate coiled-coil interactions. Presumably via these motifs CDJ2 interacts with VIPP1. CDJ2 is expressed constitutively but weakly—while VIPP1 accounts for 0.05% of Chlamydomonas total cell protein, CDJ2 accounts only for 0.006%. When taking into account the molecular weights of CDJ2 (32 kDa) and VIPP1 (28 kDa), the molar ratio of CDJ2:VIPP1 is 1:10 (Liu et al., 2007). Both CDJ2 and VIPP1 were found to interact with HSP70B and CGE1 (Liu et al., 2005, 2007). The HSP70B–CDJ2–CGE1 chaperone system was found to catalyze the ATP-dependent assembly of VIPP1 monomers/dimers to rings and the disassembly of rods and rings to monomers/dimers in vitro. Moreover, the chaperones formed complexes with VIPP1 (dis)assembly intermediates ( 670 kDa) in ATP-replete cells extracts, whereas they interacted with large (670 kDa) and small (< 230 kDa) VIPP1 assembly states in ATP-depleted cells. Finally, very large VIPP1 oligomers in cell extracts were partially disassembled by the addition of ATP and essentially completely disassembled by the addition of ATP and purified chaperones (Liu et al., 2007). Hence, CDJ2 appears to be a specialized J-domain protein that recruits HSP70B–CGE1 to VIPP1 for dynamic remodeling of its assembly state (Fig. 2.2). The dynamic remodeling of VIPP1 complexes by chaperones suggests an important function for this protein in the chloroplast. Previously, rod-shaped supercomplexes, also termed microtubule-like structures (MTLs), were observed in plastids from algae and various plant tissues by electron microscopy (Lawrence and Possingham, 1984; Lunney et al., 1975; Newcomb, 1967; Pickett-Heaps, 1968; Rivera and Arnott, 1982; Schnepf, 1961; Sprey, 1968; Whatley et al., 1982). With outer diameters measuring ˚ , a central cavity 100 A ˚ across, lengths ranging from 0.05 to 150–400 A several micrometers, rough margins, and transverse striations originating from the fusion of particles, MTLs very much resemble VIPP1 rods. MTLs were often observed to originate at the inner chloroplast membrane and to run perpendicularly into the stroma, possibly contacting thylakoids. This complex localization pattern is consistent with the localization of Chlamydomonas VIPP1 in stroma, low density membranes, and thylakoids (Liu et al., 2005). Based on their similar morphologies and intraplastid distributions, it is tempting to speculate that VIPP1 rods and MTLs are one and the same. Possible functions attributed to MTLs are diverse: it has been suggested that they might be involved in plastid division (Rivera and Arnott, 1982; Sprey, 1968), in the formation of the prolamellar body

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(Newcomb, 1967), in the formation of thylakoids during greening (Sprey, 1968), or in transport of cargo between the plastid envelope and internal lamellae and vesicles (Carde et al., 1982; Lunney et al., 1975; Whatley et al., 1982). A structural role for VIPP1 rods as tracks that transport proteins and/or lipids would be consistent with the thylakoid biogenesis defects exhibited by Arabidopsis and cyanobacterial vipp1 mutants (Fuhrmann et al., 2009b; Kroll et al., 2001; Westphal et al., 2001). However, several findings suggest that VIPP1 may have different or additional functions: first, expression of PspA significantly improves the performance of the bacterial Sec and TAT protein export pathways and of the thylakoidal TAT protein export pathway (DeLisa et al., 2004; Vrancken et al., 2007). Likewise, VIPP1 improves export via bacterial and thylakoidal TAT pathways (DeLisa et al., 2004). Consistent with these observations, mutational disruption of the Sec and TAT pathways produces similar defects in thylakoid structure as those observed in the Arabidopsis vipp1 mutant (Settles et al., 1997). These findings suggest that the vipp1 mutant phenotype might well be caused by inefficient protein translocation rather than defective vesicle formation. Second, some proteins of the bacterial plasma membrane and of the Chlamydomonas thylakoid membrane require YidC and Alb3, respectively, for efficient membrane insertion (Kuhn, 2009). E. coli yidC mutants overexpress the psp operon ( Jones et al., 2003), and Chlamydomonas alb3.2 knockdown mutants have elevated levels of VIPP1, HSP70B, and CDJ2 (Gohre et al., 2006). These results suggest involvement of PspA and of VIPP1–HSP70B– CDJ2 in processes related to membrane protein sorting. 5.2.8. HSP70B in protecting photosystem II from damage by high light One of the earliest investigations of chloroplast HSP70B revealed that its gene is highly induced by light (von Gromoff et al., 1989), resulting in an approximately twofold increase in HSP70B protein levels (Drzymalla et al., 1996). This finding suggested a possible role for this chaperone in processes that help the cell to cope with photodamage. Accordingly, cells overexpressing HSP70B exhibited less severe damage to PS II and recovered PS II activity faster after photoinhibition, compared to wild-type cells. The opposite effect (more severe damage and slower recovery) was observed for cells underexpressing HSP70B (Schroda et al., 1999). It was hypothesized that HSP70B might facilitate a coordinated exchange of damaged D1 protein by de novo-synthesized D1 protein (Schroda et al., 2001a). In support of this hypothesis, HSP70B in the green alga Dunaliella salina was found to be part of a 320-kDa complex containing photodamaged D1, D2, and CP47 proteins (Yokthongwattana et al., 2001). Alternatively, HSP70B–CDJ2 might play an indirect role in the assembly/maintenance of PS I and PS II, by facilitating Alb3.2-mediated protein sorting via VIPP1.

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5.2.9. Chloroplast HSP70B and HSP90C as a functional couple It is well known that cytosolic Hsp70 and Hsp90 constitute the core of a multichaperone complex, also referred to as the “foldosome” (Dittmar et al., 1998), that is involved in the maturation of numerous client proteins (Pratt and Toft, 2003; Wandinger et al., 2008). Typical client proteins are hormone receptors and kinases, that is, components of signal transduction pathways. The minimal folding system contains Hsp90, Hop, Hsp70, and Hsp40, and folding efficiency is increased by Hsp90 cochaperones p23 and Cdc37 for hormone receptors and kinases, respectively. Hop mediates the contact between Hsp90 and Hsp70, and Hsp40 recognizes unfolded substrate proteins and delivers them to Hsp70. Interestingly, a multichaperone complex resembling the “foldosome” (Fig. 2.2) is also present in the chloroplast. This complex contains HSP70B, HSP90C, CDJ1, and CGE1 (Willmund and Schroda, 2005; Willmund et al., 2008a). The 81-kDa HSP90C is one of three Hsp90 paralogs encoded by the Chlamydomonas genome and the only one targeted to the chloroplast (Willmund and Schroda, 2005). Like HSP70B, HSP90C is inducible by light and heat shock and is localized to stroma, low-density membranes, and thylakoids. HSP90C is about three times less abundant than HSP70B, forms labile dimers, and displays a weak ATPase activity (0.71 ATP hydrolyzed per HSP90C per minute) that is completely inhibited by radicicol. In contrast to chloroplast Hsp70 and chloroplast DnaJ and GrpE homologs, chloroplast Hsp90 is not derived from the ancestral cyanobacterial endocymbiont. Rather, chloroplast Hsp90 most likely originated from the duplication of a gene encoding an ER-targeted Hsp90, which in a secondary event acquired a chloroplast transit peptide coding sequence, while the cyanobacterial htpG gene was lost (Emelyanov, 2002). The 40-kDa CDJ1 is the chloroplast DnaJ homolog and can also functionally substitute for E. coli DnaJ (Willmund et al., 2008a). CDJ1 is induced by heat shock and light (Liu et al., 2005) and is localized mainly to the stroma, but also to low density membranes and thylakoids. CDJ1 forms stable dimers and is roughly as abundant as HSP70B (Willmund et al., 2008a). In parallel with the cytosolic “foldosome” (Dittmar et al., 1998), HSP90C and HSP70B appear to represent the core complex of the chloroplast HSP90C–HSP70B–CDJ1–CGE1 multichaperone complex, from which CDJ1 and CGE1 dynamically cycle in and out (Willmund et al., 2008a). In the case of CDJ1 this cycling likely serves the purpose of supplying the chaperones with substrates, whereas cycling of CGE1 is likely to mediate nucleotide exchange from HSP70B (Fig. 2.2). However, there are no potentially chloroplast-targeted homologs of Hop, p23, or Cdc37 (Schroda and Vallon, 2008). It was speculated that in analogy to the cytosolic “foldsome”, the HSP90C–HSP70B–CDJ1–CGE1 multichaperone complex in the

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chloroplast also might be required for the folding of proteins involved in signal transduction (Fig. 2.2) (Schroda and Mu¨hlhaus, 2009). This idea was supported by signaling defects in the Arabidopsis cr88 (chlorate resistant 88) mutant, which contains a point mutation in the C-terminal dimerization domain of chloroplast Hsp90 (Cao et al., 2003; Lin and Cheng, 1997). This mutant displays long hypocotyls and retarded deetiolation in red light and has a yellow-green appearance due to the retarded development of chloroplasts, particularly in young leaves. The mutant also exhibits reduced lightinducible expression of the nuclear NR2, CAB, and RBCS genes, but not of NR1, NiR, and CHS (Cao et al., 2000; Lin and Cheng, 1997). It was proposed that chloroplast Hsp90 might be involved in the transduction of a light signal responsible for the regulation of a subset of photosynthesisrelated genes (Cao et al., 2003). Chlorophyll biogenesis intermediates have been shown to serve as chloroplast signaling molecules for the regulation of nuclear genes by light (Beck, 2005; Woodson and Chory, 2008). As the cyanobacterial Hsp90 homolog HtpG was shown to control chlorophyll biogenesis by controlling the activity of uroporphyrinogen decarboxylase (the HemE protein), this might also be the mechanism by which light induction of specific nuclear genes is regulated by chloroplast Hsp90 (Saito et al., 2008; Watanabe et al., 2007). Through a novel approach for the unequivocal identification of protein– protein interactions in Chlamydomonas (termed QUICK-X), VIPP1 was identified as a specific substrate of HSP90C (Heide et al., 2009) (Fig. 2.2). Hence, in light of the role of VIPP1 in the biogenesis/maintenance of thylakoid membranes, it is possible that at least part of the Arabidopsis chloroplast development defect that Lin and Cheng (1997) reported for chloroplastic Hsp90 mutants is due to inefficient remodeling of VIPP1 oligomers by an impaired chloroplast chaperone complex.

6. Mitochondrion 6.1. Chaperone components Database searches suggest that Chlamydomonas mitochondria contain at least two members of the Hsp100 class (CLPX and perhaps CLPB3 or CLPB4), up to three Hsp70s (HSP70C and perhaps HSP70D or HSP70F), one chaperonin and one cochaperonin (CPN60C and CPN10), and presumably one sHSP (HSP22G). Moreover, several Hsp70 cochaperones/cofactors were also found, including a GrpE homolog (MGE1), a DnaJ homolog (MDJ1), and an Hsp70 escort protein (Hep1) (Schroda, 2004; Schroda and Vallon, 2008). A proteomic analysis confirmed the presence of CPN60C, CPN10, HSP70C, and MGE1 in mitochondria (Atteia et al., 2009). Interestingly, it appears that Chlamydomonas mitochondria do not contain

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an Hsp90 homolog. Although HSP90C was among the Chlamydomonas mitochondrial proteins found by the proteomics approach (Atteia et al., 2009) that protein probably originated from lysed chloroplasts and was detected because it adhered to the mitochondrial outer membrane (Willmund and Schroda, 2005).

6.2. Function of mitochondrial chaperones An interesting early observation was that the gene encoding mitochondrial HSP70C is light inducible (von Gromoff et al., 1989). However, no more information on functions of mitochondrial chaperones in Chlamydomonas or Volvox is available.

7. Conclusions and Outlook Although we have gained considerable insights to the composition of the chaperone outfit of Chlamydomonas and Volvox and have gained important insights into some of their functions, many open questions remain to be answered in the future. Have we correctly understood the regulation of the stress response in Chlamydomonas? By which mechanisms do Hsp70A and GlsA decide on the developmental fate of Volvox embryonic cells? Which roles do HSP70A and HSP90A play in flagella? How are organellar Hsp70s activated by their escort proteins and why is this activation necessary? What impact does the plastidic redox-state have on chloroplast (co)chaperone function? What purpose(s) does chaperone-mediated remodeling of VIPP1 oligomeric states serve? Does the chloroplast HSP90C–HSP70B chaperone pair fold proteins involved in retrograde signaling and does it play roles in protein quality control? Without any doubt, Chlamydomonas and Volvox are ideal model systems to tackle these questions and the many more that need to be addressed to understand the complex functions of molecular chaperones in algal systems.

ACKNOWLEDGMENTS This work was supported by the Max Planck Society and grants from the Deutsche Forschungsgemeinschaft (Schr 617/6-1) and the Bundesministerium fu¨r Bildung und Forschung (Systems Biology Initiative FORSYS, project GoFORSYS).

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C H A P T E R

T H R E E

Unique Functions of Repetitive Transcriptomes Gerald G. Schumann,* Elena V. Gogvadze,† ¨nk,§ Mizuko Osanai-Futahashi,‡ Azusa Kuroki,‡ Carsten Mu ‡ },k Haruko Fujiwara, Zoltan Ivics, and Anton A. Buzdin† Contents 1. Introduction 2. Eukaryotic Retrotransposons 2.1. LINE retrotransposons 2.2. SINE retrotransposons 2.3. SVA elements 2.4. Processed pseudogenes 2.5. LTR retrotransposons and ERVs 2.6. Penelope-like elements 3. Mechanisms of Intracellular Defense Against TEs 3.1. Impact of AID on retrotransposition 3.2. APOBEC3 proteins 3.3. Evidence for ADAR editing of Alu elements 3.4. piRNAs and PIWI proteins as regulators of mammalian retrotransposon activity 4. The Use of Transposable Elements in Biotechnology and in Fundamental Studies 4.1. DNA transposons as genetic tools 4.2. Retrotransposons as genetic tools 5. Domestication of Mobile DNA by the Host Genomes 5.1. Genomic repeats as transcriptional promoters 5.2. REs as enhancers for host cell gene transcription 5.3. REs as providers of new splice sites for the host genes

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* Paul-Ehrlich-Institut, Federal Institute for Vaccines and Biomedicines, Langen, Germany { Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, Russia { Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Japan } Clinic for Gastroenterology, Hepatology and Infectiology, Medical Faculty, Heinrich-Heine-University, Du¨sseldorf, Germany } Max Delbruck Center for Molecular Medicine, Berlin, Germany k University of Debrecen, Debrecen, Hungary International Review of Cell and Molecular Biology, Volume 285 ISSN 1937-6448, DOI: 10.1016/S1937-6448(10)85003-8

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2010 Elsevier Inc. All rights reserved.

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5.4. REs as sources of novel polyadenylation signals 5.5. REs as transcriptional silencers 5.6. REs as antisense regulators of the host gene transcription 5.7. REs as insulator elements 5.8. REs as regulators of translation 6. Retrotransposons as Drivers of Mammalian Genome Evolution 6.1. REs generate new REs 6.2. REs and recombination events 6.3. Transduction of flanking sequences 6.4. Formation of processed pseudogenes 6.5. Chimeric retrogene formation during reverse transcription 7. Concluding Remarks Acknowledgments References

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Abstract Repetitive sequences occupy a huge fraction of essentially every eukaryotic genome. Repetitive sequences cover more than 50% of mammalian genomic DNAs, whereas gene exons and protein-coding sequences occupy only  3% and 1%, respectively. Numerous genomic repeats include genes themselves. They generally encode “selfish” proteins necessary for the proliferation of transposable elements (TEs) in the host genome. The major part of evolutionary “older” TEs accumulated mutations over time and fails to encode functional proteins. However, repeats have important functions also on the RNA level. Repetitive transcripts may serve as multifunctional RNAs by participating in the antisense regulation of gene activity and by competing with the host-encoded transcripts for cellular factors. In addition, genomic repeats include regulatory sequences like promoters, enhancers, splice sites, polyadenylation signals, and insulators, which actively reshape cellular transcriptomes. TE expression is tightly controlled by the host cells, and some mechanisms of this regulation were recently decoded. Finally, capacity of TEs to proliferate in the host genome led to the development of multiple biotechnological applications. Key Words: Repetitive sequences, Transposable elements, Retrotransposons, APOBEC 3 proteins, RNA interference, Gene delivery Genome evolution. ß 2010 Elsevier Inc.

1. Introduction The eukaryotic genome is a complex and dynamic structure. Only about 3% of the mammalian genome is composed of protein-coding sequences compared to  50% constituted by transposable elements (TEs). Transposable or mobile genetic elements are DNA sequences that are able to jump into new locations within genomes (Bohne et al., 2008). They can

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reach very high copy numbers and represent the major fraction of eukaryotic genomes. Since their initial discovery in the maize genome by Barbara McClintock in 1956 (McClintock, 1956), mobile elements have been found in genomes of almost all organisms. They constitute more than 50% of the maize genome (Wessler, 2006), 22% of the Drosophila genome (Kapitonov and Jurka, 2003), and 42% of human DNA (Lander et al., 2001). Initially considered as “junk” DNA or genomic parasites, mobile elements are now suggested to be “functional genome reshapers,” which are able to alter gene expression and promote genome evolution (Beauregard et al., 2008; Goodier and Kazazian, 2008; Han and Boeke, 2005). TEs can be grouped in two major classes (Kazazian, 2004). Class II elements or DNA transposons comprise about 3% of the human genome and most move by a so-called cut-and-paste mechanism. No currently active DNA transposons have been identified in mammals to date (Bohne et al., 2008). Class I elements are termed retrotransposons or retroelements (REs). They move by a “copy-and-paste” mechanism involving reverse transcription of an RNA intermediate and insertion of its cDNA copy at a new site in the host genome. This process is termed retrotransposition. Retrotransposons can be grouped into two major subclasses (Kazazian, 2004). Retroviral-like or long terminal repeat (LTR) retrotransposons include endogenous retroviruses (ERVs), which are relics of past rounds of germline infection by exogenous retroviruses that lost their ability to reinfect and became trapped in the genome because they harbor inactivating mutations that render them replication defective. These elements undergo reverse transcription in virus-like particles (VLPs) by a complex multistep process. LTR-containing REs account for 10% of the mammalian genomes and their life cycle includes the formation of VLPs that, in several instances—but not systematically—can remain strictly intracellular as observed for the well-characterized murine intracisternal A-particle (IAP) and MusD elements (the so-called intracellularized ERVs; Dewannieux et al., 2004; Ribet et al., 2008), or that can bud at the cell membrane to replicate via an extracellular infection cycle as observed for the recently identified murine intracisternal A-particle-related envelope-encoding element (IAPE; Ribet et al. 2008) and for the ‘reconstituted’ infectious, human progenitor of the HERV-K(HML2) family members (Dewannieux et al. 2006; Lee and Bienasz, 2007). The second major subclass comprises the strictly intracellular non-LTR retrotransposons and is represented in the mammalian genome by long interspersed nuclear elements (LINEs), short interspersed nuclear elements (SINEs), and processed pseudogenes accounting for 30% of each mammalian genome. Only primate genomes harbor the fourth group of non-LTR retrotransposons termed SVA (SINE–variable number of tandem repeats–Alu-like). The transposition process for non-LTR retrotransposons is fundamentally different from the process observed for LTR

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LTR-containing retrotransposons

Non-LTR retrotransposons

retrotransposons. RNA copies of non-LTR retrotransposons become part of a ribonucleoprotein (RNP) complex and are thought to be carried back into the nucleus where their reverse transcription and integration occur in a single step on the genomic target DNA itself (Goodier and Kazazian, 2008). Major groups of the LTR- and non-LTR retrotransposons are schematized in Fig. 3.1.

Line

5⬘UTR

ORF1

Processed pseudogene

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Endogenous retrovirus

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Pol IN

Gag

RT RH

Pol RT RH

IN

Env

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PAT-like

Penelope-like retrotransposons

Gag

YR

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RT

EN

Figure 3.1 Schematic representation of the different types of retrotransposons. White triangles, short direct repeats (target site duplications); UTR, untranslated region; ORF, open reading frame; LTR, long terminal repeat; PR, protease; RT, reverse transcriptase; RH, ribonuclease H; IN, integrase; Env, envelope; YR, tyrosine recombinase; EN, endonuclease.

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2. Eukaryotic Retrotransposons In this section, we focus on class I TEs (retrotransposons) because they generally constitute significant proportions of higher eukaryotic DNAs and are the only group of TEs actively proliferating in the mammalian genomes.

2.1. LINE retrotransposons LINEs are termed autonomous because they are coding for the protein machinery that is required for their mobilization. They are widely distributed in eukaryotes. About 21% of the human genome is covered by elements that belong to the families LINE1, LINE2, or CR1/LINE3 (Lander et al., 2001). LINE2 and CR1/LINE3 represent ancient inactive fossils that constitute 3% and 0.3% of the human genome, respectively, and 0.4% and 0.05% of the mouse genome, accordingly (Gentles et al., 2007). In spite of the low-copy number of LINE2 and LINE3 sequences, their presence may be valuable for the host. For example, a LINE-2 fragment was shown to be a potent T-cell-specific silencer regulatory sequence (Donnelly et al., 1999). The LINE-1 (L1) family is covering about 500,000 L1 copies occupying 18% of the haploid genome. L1 elements represent the only family of autonomous non-LTR retrotransposons harboring functional elements that are currently expanding in humans (Goodier and Kazazian, 2008). However, only 80–100 elements are functional and retrotransposition-competent (Brouha et al., 2003). A human full-length L1 is 6 kb long and has a 900-nt 50 -untranslated region (UTR) that functions as an RNA polymerase II internal promoter, two open reading frames (ORF1 and ORF2), a short 30 -UTR, and a poly(A) tail. ORF1 encodes a nucleic acid-binding protein that lacks sequence similarity with any other known protein (Goodier et al., 2007; Han and Boeke, 2005). The ORF2 protein contains endonuclease (EN) and reverse transcriptase (RT) activities as well as a Cys-rich domain, and all three domains are absolutely essential for retrotransposition (Moran et al., 1996). Usually, L1 sequences are flanked by short direct repeats called target site duplications (TSDs) (Fig. 3.1). L1 retrotransposition is thought to occur by a mechanism termed targetprimed reverse transcription (TPRT) (Luan et al., 1993). During TPRT, L1EN recognizes and cleaves the DNA consensus target sequence 50 TTTT/AA-30 which means that there are a multitude of potential genomic L1 integration sites (Feng et al., 1996; Jurka, 1997). Due to the cis-preference of the L1-encoded protein machinery for its own mRNA, L1 mobilizes preferentially itself (Wei et al., 2001). However, in very rare cases, L1s are able to mobilize Alu (Dewannieux et al., 2003) and SVA RNAs (Raiz and Schumann, unpublished data) as well as cellular mRNAs whose retrotransposition results in pseudogene formation (Esnault et al., 2000).

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2.2. SINE retrotransposons SINEs are reiterated, short (80–500 bp) sequences, comprise about 12% of the human genome, and do not code for proteins (Kramerov and Vassetzky, 2005). SINEs harbor an internal promoter, are pol III-transcribed, and possess at their 30 -end a pA-rich tail (Fig. 3.1) Most SINEs within a given family are full-length and are flanked by TSDs of varying length. Structural similarities between LINEs and SINEs suggested early that the LINEencoded protein machinery is responsible for SINE mobilization. SINEs “hijack” the RT encoded by an autonomous non-LTR retrotransposon for their own mobilization. It is generally accepted that LINEs are used as a source of RT for SINE proliferation (Eickbush, 1992). In the human genome, SINEs are represented by two major families termed MIR (mammalian-wide interspersed repeats) and Alu. MIR elements are tRNA-like SINEs that include 470,000 copies constituting about 2% of the human genome; while Alus are 7SL RNA-derived elements, include 1.1  106 elements occupying 10% of the genome (Kramerov and Vassetzky, 2005; Lander et al., 2001). Alu elements are the most abundant repeats in the human genome. The major burst of Alu retrotransposition took place 50–60 million years ago (mya) and has since dropped to a frequency of one new transposition event in every 20–125 births (Cordaux et al., 2006; Shen et al., 1991).

2.3. SVA elements SVA elements are primate-specific nonautonomous non-LTR retrotransposons which originated <25 mya and represent the youngest retrotransposon family in primates. So far their copy number has increased to 3000 in the human genome (Ostertag et al., 2003; Wang et al., 2005). SVA elements stand out from the group of human non-LTR retrotransposons due to their composite structure including modules derived from other primate repetitive elements. Starting at the 50 -end, a full-length SVA element is composed of a (CCCTCT)n hexamer repeat region; an Alulike region consisting of three antisense Alu fragments adjacent to an additional sequence of unknown origin; a variable number of tandem repeat (VNTR) region which is made up of copies of a 36- to 42-bp sequence or of a 49- to 51-bp sequence (Ostertag et al., 2003); and a short interspersed element of retroviral origin (SINE-R) region. The latter is derived from the 30 -end of the env gene and the 30 -LTR of the ERV HERV-K10. A poly(A) tail is positioned downstream of the predicted conserved polyadenylation signal AATAAA (Ostertag et al., 2003). Considering the number of diseasecausing insertions relative to their overall copy number, SVA elements are thought to represent a highly active retrotransposon family in humans (Ostertag et al., 2003; Wang et al., 2005).

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The origin of SVA elements can be traced back to the beginnings of hominid primate evolution, only 18–25 mya. Their very young evolutionary age represents a unique opportunity to study the entire evolutionary history of a human retrotransposon. In addition, SVA elements may be valuable as markers for primate or human phylogenetic and population genetic studies, as has been the case for Alu elements (Bamshad et al., 2003; Watkins et al., 2003; Xing et al., 2007).

2.4. Processed pseudogenes Processed pseudogenes are found in most mammalian genomes and their structure is that of an integrated cDNA copy of a cellular mRNA: They do contain introns, have lost the untranscribed part of the promoter, end with a polyA tail, and are flanked by TSDs (Fig. 3.1) (Brosius, 1999b; Esnault et al., 2000; Weiner et al., 1986). Similar to Alu retrotransposition, processed pseudogenes were demonstrated to be generated by trans-mobilization of cellular mRNAs by the protein machinery encoded by intact LINE retrotransposons (Esnault et al., 2000). In most cases, processed pseudogenes are not functional because they do not include complete promoters and/or because of the accumulation of mutations which occur in the absence of any selection pressure. On rare occasions, processed pseudogenes are functional due to the fortuitous presence of a promoter upstream of the insertion site and the conservation of an intact ORF with a new expression pattern. Generally, there are 1–10 (in some cases up to 100) processed pseudogenes for each human gene (Brosius, 1999b).

2.5. LTR retrotransposons and ERVs The group is also termed LTR-containing REs and combines LTR retrotransposons and ERVs which are all flanked by LTRs. All LTR-containing elements comprise about 8% of the human genome (Bannert and Kurth, 2004). The organization of LTR retrotransposons is similar to that of retroviruses except for the absence of the env (envelope) gene in all LTR retrotransposons (Eickbush and Jamburuthugoda, 2008). LTR-containing REs include a gag gene, coding for a structural protein with nucleic acidbinding activity, and a pol gene which encodes a polyprotein with protease, RT, RNaseH, and integrase activities. There are three distinct lineages of LTR retrotransposons in vertebrates: the Ty1/copia, Ty3/gypsy, and BEL (Fig. 3.1). ERVs are relics of past rounds of germline infection by exogenous retroviruses that lost their ability to reinfect and became trapped in the genome because they harbor inactivating mutations that render them replication defective. They were found in all vertebrate genomes and constitute about 8% of the human genomic DNA (Lander et al., 2001). Most ERV

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sequences have undergone extensive deletions and mutations, therefore becoming transpositionally deficient and transcriptionally silent (Sverdlov, 2000). Moreover, the majority of ERVs reside in the genome in the form of solitary LTRs, arisen most probably due to homologous recombination between two LTRs of a full-length element. Tyrosine-recombinase encoding retrotransposons (or YR-retrotransposons) represent an additional group of LTR-containing REs (Poulter and Goodwin, 2005). These elements have structures quite distinct from the REs described above. The major difference is that YR-retrotransposons do not code for an integrase, but for a tyrosine recombinase (YR) instead (Fig. 3.1). The first element of this group was identified in the slime mold, Dictyostelium discoideum, and was called DIRS (Cappello et al., 1985). Later on, related elements were found in the genomes of numerous fungi, plants, and animals (Goodwin and Poulter, 2004). All these elements could be divided into two basic groups: DIRS-like elements which are flanked by inverted repeats and contain an internal complementary region, and elements of the PAT and Ngaro type which have split direct repeats (Fig. 3.1). The unusual structure of the terminal repeats of the YR-retrotransposons was suggested to be required for their replication via a free circular intermediate (Cappello et al., 1985; Goodwin and Poulter, 2004). The circular intermediate is believed to be integrated into the genome by site-specific recombination without the formation of TSDs. The human genome contains a DNA sequence similar to a large fragment of a DIRS1-like recombinase gene. However, no full-length mammalian DIRS-like elements have been found to date (Poulter and Goodwin, 2005).

2.6. Penelope-like elements PLEs constitute a novel class of eukaryotic REs that are distinct from both non-LTR and LTR retrotransposons (Evgen’ev and Arkhipova, 2005) (Fig. 3.1). They were first discovered in Drosophila virilis as elements responsible for the hybrid dysgenesis syndrome, and characterized by simultaneous mobilization of several unrelated TE families in the progeny of dysgenic crosses. PLEs were further found in genome databases of various eukaryotes (Gladyshev and Arkhipova, 2007). They have a rather complex and highly variable organization. These elements were shown to contain an internal promoter (Schostak et al., 2008) and one ORF coding for RT and EN activities that differ from the corresponding proteins of LTR-containing and/or non-LTR retrotransposons (Evgen’ev and Arkhipova, 2005). The PLE EN belongs to the URI protein family, which includes, inter alia, catalytic modules of the GIY-YIG ENs of group I introns, as well as bacterial UvrC DNA repair proteins. The RT of PLEs mostly resembles the RT domain of telomerase. Both RT and EN domains encoded

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by D. virilis Penelope are functionally active, but the mechanism of their transposition remains unclear.

3. Mechanisms of Intracellular Defense Against TEs TEs have played an important role in evolution and speciation. However, mobilization of these elements can also be deleterious to the host and can result in various genetic disorders and cancer. Given these various deleterious effects, it is not surprising that the cell has generated multiple mechanisms controlling their proliferation. To limit the negative effects of retrotransposition, several strategies have been adopted to restrict mobility and potentially deleterious consequences of uncontrolled retrotransposition. Such host-encoded strategies include DNA methylation, RNA interference (RNAi), and inhibition of retrotransposition by the activity of members of the APOBEC (named after apolipoprotein B mRNA-editing enzyme catalytic polypeptide 1, APOBEC1) protein family which comprises 11 closely related DNA or RNA cytidine deaminases. Epigenetic modifications controlling the activity of TEs were initially reported more than 24 years ago (Chandler and Walbot, 1986). Since then many genes involved in epigenetic silencing of TEs (including DNA methyltransferases and demethylases, histone modifying enzymes, chromatin remodeling enzymes, and genes involved in small RNA metabolism) were characterized. Defects in different components of silencing mechanisms were shown to increase transposition events (Weil and Martienssen, 2008). Most of the methylated cytosines in mammalian genomes reside in repetitive elements and it has been proposed that DNA methylation evolved primarily to suppress the activity of TEs and to protect the host cell (Yoder et al., 1997). Hypomethylation of REs was demonstrated to be associated with genomic instability in cancer (Daskalos et al., 2009). On the one hand, chromatin condensation may suppress the activity of REs. On the other hand, DNA methylation, initiated within RE, may spread to the surrounding genomic regions and, hence, suppress their functional activity. Spreading of CpG methylation from SINEs into flanking genomic regions was suggested to create distal epigenetic modifications in plants (Arnaud et al., 2000). Human Alu elements were proposed as potential de novo methylation centers implicated in tumor suppressor gene silencing in neoplasia (Graff et al., 1997). Recent studies have shown the involvement of RNAi-related mechanisms in the control of TE activities, in particular, in DNA methylation of TE sequences and in the formation of heterochromatin. Plants, yeasts, and animals use different strategies to detect transposons and to generate small RNAs against them (Girard and Hannon, 2008; Slotkin and Martienssen, 2007).

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3.1. Impact of AID on retrotransposition The activation-induced cytidine deaminase (AID) gene is evolutionary quite old because it is part of the genomes of vertebrates down to jawless vertebrates (Rogozin et al., 2007). AID deaminates cytidines to uridines (C to U) in single-stranded DNA and is required for antibody maturation involving somatic hypermutation at the immunoglobulin variable regions, class switch recombination at the switch regions, and Ig gene conversion in some species. In general, the enzyme is mainly expressed in the B-cell compartment. In the mouse, AID is detectable in the spleen, ovaries, and oocytes (MacDuff et al., 2009; Morgan et al., 2004) and is moderately expressed in the heart (MacDuff et al., 2009) and in murine embryonic stem (ES) and germ cells (Morgan et al., 2004). Contrasting to the mouse, humans express AID only in B-cells and testes (MacDuff et al., 2009; Schreck et al., 2006). L1 retrotransposition reporter assays performed separately in HEK293 cells in the presence of overexpressed HA-tagged AID proteins from different species demonstrated that human L1 retrotransposition is inhibited by 24–68% by AID from human, mouse, rat, chicken, pufferfish, and zebrafish, while porcine AID restricted L1 activity by even 90% (MacDuff et al., 2009). Interestingly, neither mutations in the catalytically active site nor mutations in the predicted DNA-binding site of human AID had any consequences for the L1-inhibiting effect of AID. This is indicating that AID-mediated inhibition of L1 is both cytidine deaminase- and DNAbinding-independent (MacDuff et al., 2009). In an earlier report, inhibition of L1 retrotransposition activity by human AID could not be demonstrated (Niewiadomska et al., 2007). In the presence of overexpressed wild-type AID proteins from multiple species or the catalytically inactive mutant AID-E58Q, retrotransposition of the mouse LTR retrotransposon MusD was inhibited by only 13–50% in HeLa cells (Esnault et al., 2006; MacDuff et al., 2009). Under such experimental conditions, AID induced a small number of mutations in de novo MusD retrotransposition events (Esnault et al., 2006). In contrast to the MusD element, the yeast LTR retrotransposon Ty1 was not affected by coexpression of AIDs in yeast cells (MacDuff et al., 2009). It was reported that neither APOBEC1 nor APOBEC2 had any effect on MusD retrotransposition in HeLa cells (Chen et al., 2006; Esnault et al., 2006). Also, L1- and IAP retrotransposition were not impaired by A2 (Chen et al., 2006; Niewiadomska et al., 2007).

3.2. APOBEC3 proteins The APOBEC3 (A3; apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3) proteins are Zn2þ-dependent DNA cytidine deaminases, which were discovered to constitute a defensive network of proteins

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that restrict the replication of retroviruses (Bishop et al., 2004; Chiu and Greene, 2008) and an incredible range of mobile genetic elements (Chiu and Greene, 2008; Goila-Gaur and Strebel, 2008; Hultquist and Harris, 2009; Malim and Emerman, 2008). The A3 genes are only present in placental mammals (LaRue et al., 2008; Munk et al., 2008). Phylogenetic studies have indicated that the first A3 gene(s) arose from an AID-like ancestral gene through a series of duplication and diversification events (Conticello, 2008; Conticello et al., 2007; LaRue et al., 2008). The APOBEC3 gene family has proliferated during mammalian speciation and many members of which exhibit signs of positive (diversifying) selection in the primate and felid lineages and accordingly are highly polymorphic (Kidd et al., 2007; Munk et al., 2008; OhAinle et al., 2006, 2008; Sawyer et al., 2004; Zhang and Webb, 2004). The seven A3 genes are positioned in tandem on human chromosome 22: A3A, A3B, A3C, A3D (formerly A3DE), A3F, A3G, and A3H ( Jarmuz et al., 2002; OhAinle et al., 2006). A defining feature of each A3 gene is that it encodes a protein with one or two conserved zinc (Z)-coordinating deaminase domains. Each Z domain belongs to one of three distinct phylogenetic groups: Z1 (A3A and the C-terminal halves of A3B and A3G), Z2 (A3C, both domains of A3D and A3F, and the N-terminal halves of A3B and A3G), and Z3 (A3H) (LaRue et al., 2009). Based on the relatedness of these Z domains, the human A3 repertoire appears to be the result of a minimum of eight unequal crossing-over recombination events, which mostly occurred during the radiation of primates (LaRue et al., 2008; Munk et al., 2008). The net result is that the human A3 mRNAs share considerable identity, ranging from 30% to nearly 100% (Refsland et al., 2010). The members of the human A3 protein family differ from each other with respect to their intracellular localization after overexpression (Fig. 3.2). While A3B is found exclusively in the nucleus, A3A is predominantly located to the nucleus. A3C is equally distributed and A3H is located in the cytoplasm and in nucleoli, while A3F, A3D, and A3G appeared exclusively in the cytoplasmic compartment (Bogerd et al., 2006a; Kinomoto et al., 2007; Muckenfuss et al., 2006; Stenglein and Harris, 2006; Zielonka et al., 2009). 3.2.1. APOBEC3 deaminases as inhibitors of LTR retrotransposons APOBEC3 proteins do in fact function as inhibitors of LTR retrotransposons. Human A3A, A3B, A3C, A3F, A3G, and mA3 all effectively restrict mouse IAP and MusD elements (Bogerd et al., 2006a; Chen et al., 2006; Esnault et al., 2005, 2006), whereas hA3C, hA3F, and hA3G inhibit retrotransposition of the yeast Ty1 element (Dutko et al., 2005; Schumacher et al., 2005). APOBEC3 proteins exert dual inhibitory effects on these ERVs, involving both a decrease in the number of transposed cDNA copies and extensive editing of the transposed copies (Esnault et al., 2005, 2006).

Inhibitory effect of APOBEC proteins on

Subcellular localization

100 aa

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198 aa

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236 aa

hA1

224 aa

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+ −1

/

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/

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/

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++4,5,8

++4 −3

/

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++4

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++10,19 −9

/

/

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C

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/

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/

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1

++1,2,4,11,14 ++11 ++9,10,19 ++15,16 −5,17 ++1,2,11,14,16 +11,12,17 +10 −4,5,15,17,18,20 +21 −9,19 +12,16 −15,11 +13

+12 /

+10 −9 −10,19

++4

++4,8,10,18 ++4,3,18 ++6,7

/

/

/

++4,10,18 ++4,13,18 ++6

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These effects are reminiscent of the dual effects of hA3G in HIV-1 replication. In the mouse genome, many preexisting retrotransposon sequences bear mutations consistent with APOBEC3-mediated deamination (Esnault et al., 2005, 2006). Interestingly, hA3A effectively inhibits IAP and MusD retrotransposition through a novel deamination-independent mechanism (Bogerd et al., 2006a). One human ERV [HERV-K(HML-2)], which has replicated in humans for the past few million years but is now thought to be extinct, was recently reconstituted in a functional form in two separate laboratories by aligning human-specific proviruses and synthesizing a pseudo-ancestral proviral construct termed HERV-KCON and Phoenix, respectively (Dewannieux et al., 2006; Lee and Bieniasz, 2007). Of all tested human A3 proteins, only hA3A, hA3B, and hA3F were shown to be intrinsically capable of mutating and inhibiting infection by HERV-K CON in cell culture by up to 80% while hA3G led only to a marginal restriction of infection (Lee et al., 2008b). Differing from these results, Esnault and coworkers reported that the infectious HERV-K and murine IAPE elements (Ribet et al., 2008) are both restricted efficiently by the murine A3 protein and by hA3G in an ex vivo assay for infectivity (Esnault et al., 2008). They also demonstrated that hA3A, hA3B, hA3D, and hA3F restrict infectivity of HERV-K but do not affect IAPE infectivity. The same report presented evidence of strandspecific G-to-A editing of both proviruses (Esnault et al., 2008). For HERV-K, G-to-A editing was not observed with hA3A (Esnault et al., 2008). In silico analysis of the naturally occurring genomic copies of the corresponding endogenous elements performed on the mouse and human genomes discloses “traces” of A3-editing, with the specific signature of the murine A3 and human A3G enzymes, respectively, and to a variable extent depending on the family member (Esnault et al., 2006). There is striking evidence indicating that two HERV-K(HML-2) proviruses that are fixed in the modern human genome (HERV-K60 and HERV-KI) were subjected to hypermutation by A3G (Lee et al., 2008b). Figure 3.2 Members of the APOBEC protein family and their effects on the activity of retroelements. The domain organization of human and murine APOBEC proteins is depicted. Gray bars represent CDA motifs in each protein. The consensus amino acid sequence of the catalytic domains and the numbers of amino acids (aa) that compose the proteins are shown. Nuclear (N) and cytoplasmic (C) localization of the proteins as well as publications reporting major (þþ), modest (þ), minimal (þ/), and no () effects are indicated. /, not determined (n.d.). 1, MacDuff et al. (2009); 2, Niewiadomska et al. (2007); 3, Esnault et al. (2006); 4, Chen et al. (2006); 5, Bogerd et al. (2006b); 6, Dutko et al. (2005); 7, Schumacher et al. (2005); 8, Bogerd et al. (2006a); 9, Lee et al. (2008b); 10, Esnault et al. (2008); 11, Khatua et al. (2010); 12, Tan et al. (2009); 13, Lovsin and Peterlin (2009); 14, Stenglein and Harris (2006); 15, Muckenfuss et al. (2006); 16, Kinomoto et al. (2007); 17, Hulme et al. (2007); 18, Esnault et al. (2005); 19, Turelli et al. (2004); 20, Chiu et al. (2006).

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These are rare examples for the antiretroviral effects of A3G in the setting of natural human infection, whose consequences have been fossilized in human DNA. 3.2.2. APOBEC3 deaminases as inhibitors of L1 retrotransposition It was reported concordantly that A3A and A3B are potent inhibitors of human L1 retrotransposition causing a reduction of retrotransposition frequency by 85–99% and 75–90%, respectively, while A3C-mediated inhibition is less pronounced (40–75%) (Bogerd et al., 2006b; Chen et al., 2006; Khatua et al., 2010; Kinomoto et al., 2007; Lovsin and Peterlin, 2009; MacDuff et al., 2009; Muckenfuss et al., 2006; Niewiadomska et al., 2007; Stenglein and Harris, 2006; Tan et al., 2009). A3F-mediated L1 inhibition was shown to range from 66% to 85% (Chen et al., 2006; Khatua et al., 2010; Kinomoto et al., 2007; MacDuff et al., 2009; Muckenfuss et al., 2006; Niewiadomska et al., 2007; Stenglein and Harris, 2006), but these findings were questioned in two recent reports (Bogerd et al., 2006b; Hulme et al., 2007). The effect of A3G on L1 retrotransposition is more controversial because results presented in one-half of all studies argue against any A3G-mediated inhibitory effect (Bogerd et al., 2006b; Chen et al., 2006; Esnault et al., 2005; Hulme et al., 2007; Muckenfuss et al., 2006; Turelli et al., 2004) while the other half demonstrates L1 restriction by A3G by 30–90% (Khatua et al., 2010; Kinomoto et al., 2007; MacDuff et al., 2009; Niewiadomska et al., 2007). In the case of A3D, both minor inhibition by 35–45% (Kinomoto et al., 2007) and major inhibition by 95% (Niewiadomska et al., 2007) were referred. Previous reports have shown that A3H restricts L1 activity by 50% (Kinomoto et al., 2007) or not at all (Khatua et al., 2010; Muckenfuss et al., 2006). This was explained by the existence of human Z2-type cytidine deaminase A3H variants that have varying intrinsic abilities to restrict REs. It was found that in contrast to A3H, the variant A3H-Var is a highly effective inhibitor of L1 retrotransposition being almost as potent as A3A (OhAinle et al., 2008; Tan et al., 2009). The frequency of this A3H variant allele, A3H-Var, is the highest among sub-Saharan Africans and is significantly lower in Asian and European populations (Tan et al., 2009). Only little human L1 restriction by 32% is mediated by mA3 (Lovsin and Peterlin, 2009). The mechanisms responsible for A3-mediated L1 inhibition are unclear to date. So far, there is no direct evidence for L1 inhibition by cytidine deamination of L1 cDNA or any kind of editing of L1 nucleic acid sequences, strongly suggesting that cytidine deaminase-independent mechanisms are involved in A3-mediated L1 restriction. Niewiadomska and coauthors reported that A3A was associated with L1 RNA in high-molecular mass (HMM) complexes that presumably contain L1 RNPs. Consistently, A3A–HMM complexes were destroyed by RNase treatment (Niewiadomska et al., 2007). An interaction between L1 ORF1

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protein (L1 ORF1p) and A3A could not be demonstrated (Lovsin and Peterlin, 2009). A catalytically active cytidine deaminase domain (CDD) was shown to be essential for the interaction of A3A with L1 RNA and for the ability of A3A to inhibit L1 retrotransposition, even though no G-to-A hypermutations were detectable in L1 de novo retrotransposition events that occurred in the presence of A3A. It was concluded that A3A might indirectly interfere with L1 metabolism, probably by binding L1 RNA. A3A may also interfere with L1 reverse transcription/integration, similar to A3G-mediated restriction of Vif-deficient HIV-1 (Bishop et al., 2004). Alternatively, the association of A3A or other hA3 proteins with the L1 RNP could impede the intracellular movement of the L1 RNP and therefore its retrotransposition (Niewiadomska et al., 2007). Interaction between L1 RNA and hA3 proteins is also supported by the fact that it was not possible to date to demonstrate any direct interaction between hA3 proteins and L1 ORF1p in the absence of RNA (Lovsin and Peterlin, 2009). Data were presented indicating that A3B and mA3 bind to L1 ORF1p via an RNA bridge (Lovsin and Peterlin, 2009). 3.2.3. APOBEC3 deaminases as inhibitors of Alu retrotransposition Retrotransposition of Alu elements is mediated by the L1 ORF2 protein (L1 ORF2p) which has RT and EN activities but does not require the RNA-binding L1 ORF1p (Babushok et al., 2007; Dewannieux et al., 2003). Transient expression of hA3A, hA3B, hA3D, or hA3G was shown to restrict Alu retrotransposition frequency by 75–98% regardless of whether L1 ORF2p alone or both L1 ORF1p and L1 ORF2p were coexpressed (Chiu et al., 2006; Hulme et al., 2007; Khatua et al., 2010; Tan et al., 2009). A3C and A3H restrict Alu retrotransposition by only 50–70% and 65%, respectively (Bogerd et al., 2006b; Tan et al., 2009). Several groups have reported that A3G is localized to P-bodies and stress granules which are sites of mRNA storage and metabolism, raising the question of whether P-bodies and/or stress granules play a role in A3G-mediated Alu retrotransposition. A3G-mediated inhibition of Alu retrotransposition is thought to result from Alu RNA sequestration by A3G in cytoplasmic HMM complexes, particularly Staufen-containing RNA granules, denying these REs access to the nuclear L1 machinery. These effects appear to explain how hA3G interdicts the Alu retrotransposition cycle (Chiu et al., 2006; Hulme et al., 2007). This inhibitory mechanism does not involve editing of the Alu RNA and also differs from hA3A- and hA3B-mediated inhibition of Alu retrotransposition which is based on the alteration of the activity of the L1 machinery in the nucleus by the A3 protein. A3A and A3H which also act as potent Alu inhibitors have been reported to be localized to both cytoplasm and nucleus. Unlike A3G, both A3A and A3H interacted poorly with Alu RNAs. However, A3H

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associates with HMM complexes while A3A interacts poorly with P-bodies and mRNA-containing HMM complexes (Niewiadomska et al., 2007), suggesting that A3A-mediated suppression of Alu retrotransposition is not linked to P-bodies. Altogether, these results suggest that different A3 proteins may have evolved distinct inhibitory mechanisms against Alu REs. It is reasonable to hypothesize that A3 cytidine deaminases such as A3H and A3A have evolved to inhibit Alu mobilization by interfering with components of the L1 machinery and/or host factors that are required for Alu retrotransposition (Tan et al., 2009). Recently, exosomes secreted by CD4þ H9 T cells were reported to encapsidate A3G and A3F and to inhibit Alu retrotransposition by 92–96%, being almost equally potent inhibitors as A3G and A3F themselves (Khatua et al., 2010). Exosomes secreted by mature monocyte-derived dendritic cells (M-DC) that expressed A3G inhibited Alu retrotransposition by 55%. The data indicated that the inhibitory effect of exosomes against Alu mobilization is caused, at least in part, by the presence of encapsidated A3G. H9 exosomes that were originally found to encapsidate A3G and inhibit HIV-1 replication (Khatua et al., 2010) had a strong inhibitory effect against L1. They were also found to encapsidate mRNAs coding for A3C, A3F, and A3G. Since A3G mRNA isolated from exosomes was shown to be functional and supports protein synthesis in an in vitro translation system, the authors speculated that transfer of functional A3 proteins and corresponding mRNAs modulate or enable cells to resist invading or endogenous REs. It was suggested that exosomes with encapsidated A3 proteins may serve to fine tune the response against transiently expressed REs in germ cells, during early stages of human embryogenesis, or even in somatic cells (Khatua et al., 2010). 3.2.4. A3 expression profile in human tissues The expression profile of each of the seven human A3 genes was determined by RT-PCR, quantitative RT-PCR, and Northern blot analysis (Koning et al., 2009; Refsland et al., 2010; Schumann, 2007). The expressed A3 repertoire was profiled in 25 distinct human tissues, common T-cell lines, a variety of primary hematopoietic cell types, tumors, and tumor cell lines (Koning et al., 2009; Refsland et al., 2010; Schumann, 2007). It was demonstrated that multiple A3 genes are expressed constitutively in most types of cells and tissues, and that distinct A3 genes are induced upon T-cell activation and interferon treatment (Refsland et al., 2010). The relatively high expression levels of A3 proteins in human testis and ovary (hA3G, hA3F, and hA3C) ( Jarmuz et al., 2002; Koning et al., 2009; OhAinle et al., 2006) and in ES cells (hA3B) (Bogerd et al., 2006a) point to a physiologically relevant role for these DNA deaminases in the protection of these cells from the potentially deleterious effects of

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endogenous RE mobilization. Brain tissue is exceptional because it expresses virtually no A3 proteins (Refsland et al., 2010). This is consistent with the observation that endogenous L1 elements retrotranspose in neural progenitor cells (Coufal et al., 2009). More generally, nearly every cell type and tissue expresses multiple A3s, consistent with a model in which parasitic elements must evolve ways to cope with a constitutive set of restriction factors that can be further fortified by transcriptional induction.

3.3. Evidence for ADAR editing of Alu elements RNAs in higher eukaryotes can be subjected to a posttranscriptional modification called RNA editing by adenosine deaminases acting on RNA (ADARs). This process involves modification of individual adenosine bases to inosine in RNA molecules. Inosine acts as guanosine during translation, and A-to-I conversion in coding sequences leads to amino acid changes, alterations of transcriptional start and stop codons, as well as RNA splice sites. When comparing genomic with cDNA sequences, edited sites are identified by A-to-G transitions because inosine base pairs with cytosine and, therefore, is replaced by guanosine during reverse transcription and PCR amplification. RNA editing patterns characteristic of ADAR enzymes have been detected in several viral RNAs, including those of measles virus, influenza virus, hepatitis delta virus, and hepatitis C virus. To date, there is no evidence for any ADAR-mediated modulation of the activity of TEs from tissue culture experiments. However, in vivo findings described below indicate an intimate relationship between ADARs and retrotransposons. The total number of currently known A-to-I edited genes in mammals is small. ADAR editing is functionally crucial for the expression of some neurotransmitter receptors in the brain, and ADAR1-deficient mice show embryonic lethality. Usually, both the edited and the unedited versions of the RNA and/or protein coexist in the same cell. ADARs recognize and edit through their interaction with the complete or incomplete dsRNA structure formed between the edited site which is located within an exon sequence and its complementary sequence usually located in an intron sequence. Currently, it is not possible to predict if and to what extent a given RNA molecule is a substrate for A-to-I editing in vivo. Three distinct ADAR genes have been identified in mammals. ADAR1 is ubiquitously expressed in mammalian tissues; ADAR2 expression levels are highest in the brain. ADAR3, exclusively expressed in the brain has so far not shown any catalytic activity using synthetic dsRNA or known ADAR targets. Because the p150 isoform of ADAR1 is interferon inducible and upregulated in immune cells during inflammation, it is likely that ADAR1 is important for the cellular resistance to pathogens. Furthermore, ADAR1

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is involved in the RNAi pathway and is known to alter both the targeting and the processing of microRNAs (miRNA). Since 2004 several thousands of edited human mRNAs were identified (Athanasiadis et al., 2004). Clusters of A-to-G discrepancies in the cDNAs were found to be the result of RNA editing involving intermolecular pairs of inverted Alu repeat sequences (Athanasiadis et al., 2004). It was suggested that the vast majority of primary human gene transcripts are subject to A-to-I RNA editing (Athanasiadis et al., 2004; Blow et al., 2004; Kim et al., 2004). Signs of ADAR editing were detected in initial screens of small numbers of cDNAs of genes as well as in subsequent larger computational surveys ( 2000 different genes) of the majority of transcripts. In most of these cases, the location of the A-to-G cluster coincides with the position of a repetitive element, such as Alu and L1 present in introns or UTRs in the cDNAs (Athanasiadis et al., 2004; Blow et al., 2004; Kim et al., 2004; Levanon et al., 2004). These findings suggested that repetitive elements, such as Alu sequences, in RNAs might be frequent targets of ADAR editing which presumably requires the intramolecular pairing of two oppositely oriented base pairing repeat elements within the RNA molecule (Athanasiadis et al., 2004; Blow et al., 2004). It has been shown that many hyperedited, inosinecontaining RNAs are restrained in the nucleus by a protein complex containing the inosine-binding protein p54nrb (also known as NONO), PSF, and matrin3 (Zhang and Carmichael, 2001). In view of the widespread editing of Alu sequences, this offers an intriguing mechanism to mark nonstandard transcripts and preclude aberrantly spliced mRNAs and repetitive elements containing RNAs from exiting the nucleus (Athanasiadis et al., 2004; Kim et al., 2004). In mice, the ADAR editing sites are mainly found in B1 and B2 SINEs, in L1 and MaLR LTR sequences (Neeman et al., 2006; Riedmann et al., 2008).

3.4. piRNAs and PIWI proteins as regulators of mammalian retrotransposon activity DNA methylation and RNAi (Carmell and Hannon, 2004) are independent pathways that are restricting retrotransposons and can combine to form a powerful and redundant additional mechanism for keeping retrotransposons in check. This mechanism utilizes small noncoding RNAs (ncRNAs) that guide the effector complex, which is including members of the PIWI/ ARGONAUTE protein family to degrade and/or suppress target mRNAs encoded by LTR- and non-LTR retrotransposons. Members of the evolutionarily conserved PIWI/ARGONAUTE protein family are only expressed in germ cells and are key players in RNA silencing (Hutvagner and Simard, 2008). The PIWI/ARGONAUTE protein family can be subdivided into AGO and PIWI subfamilies. AGO proteins bind to small interfering RNAs (siRNAs) and miRNAs, and

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have been shown to play crucial roles in the siRNA and miRNA pathways in many tissues. PIWI proteins bind to a novel class of germ cell-specific ncRNAs called PIWI-interacting RNAs (piRNAs) (Aravin et al., 2006; Girard and Hannon, 2008; Lau et al., 2006; Watanabe et al., 2006) and have diverse functions in germline development and gametogenesis (Cox et al., 1998). Several lines of evidence indicate that PIWI proteins lead to epigenetic repression of retrotransposon-encoding regions presumably through piRNAs (Thomson and Lin, 2009). Mammalian PIWI family genes, including the three mouse PIWI homologs, Miwi, Mili, and Miwi2 (Carmell and Hannon, 2004; Deng and Lin, 2002; Kuramochi-Miyagawa et al., 2004), show germ cell-specific expression and are essential for spermatogenesis (Lin, 2007; Peters and Meister, 2007; Siomi and Kuramochi-Miyagawa, 2009). The phenotypes of Mili and Miwi2 gene-targeted mice were essentially the same and showed male sterility due to apoptosis of the germ cells at early pachytene phase (Carmell et al., 2007; Kuramochi-Miyagawa et al., 2004). MILI, which is expressed in primordial germ cells (PGCs) at embryonic day 12.5 to round spermatids, binds to 26- to 27-nucleotide (nt) piRNAs (Aravin et al., 2006; Kuramochi-Miyagawa et al., 2008). MIWI2, which is expressed in fetal gonocytes from embryonic day 15.5 until soon after birth, binds to 28- to 29-nt piRNAs (Aravin and Bourc’his, 2008; Aravin and Hannon, 2008; Kuramochi-Miyagawa et al., 2010; Thomson and Lin, 2009). About 25% of the piRNAs at the fetal stage were derived from LTR retrotransposon/ ERV (MER, ERVK, ERVL, ERV1, MaLR) and non-LTR retrotransposon sequences (L1, SINEs), and the production of piRNA was markedly impaired in MILI- and MIWI2-deficient mice (Kuramochi-Miyagawa et al., 2008). MILI and MIWI2 have been implicated in the repression of LTR retrotransposon IAP and non-LTR retrotransposon L1, with methylation of the L1 50 -UTR being reduced in newborn mice defective in these proteins (Aravin et al., 2007a; Carmell et al., 2007). These data suggest that MILI and MIWI2 are involved in piRNA production in the fetal male gonads, and that the piRNA production would play some important role(s) in gene silencing of retrotransposons via DNA methylation. MILI is central to the primary processing of sense piRNAs from retrotransposon mRNAs and other cellular transcripts (Aravin and Hannon, 2008; Aravin et al., 2007b; Kuramochi-Miyagawa et al., 2008). Then, the primary piRNAs guide the production of secondary piRNAs, which are loaded onto MIWI2, from mostly antisense RNAs transcribed from retrotransposons and other genomic elements (Aravin and Hannon, 2008). The loss of Mili leads to a gross reduction in total piRNAs and those loaded onto MIWI2. piRNAs are processed from long precursors encoded by large primary transcripts. Often piRNAs cluster to arrays that appear to be bidirectionally transcribed, while less often are primary transcripts derived from one strand. The clusters are transcribed as long transcripts that go through primary

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processing to give rise to mature piRNAs, but the mechanism of primary processing is not understood. piRNAs themselves may have a role in retrotransposon repression, as the deletion of a small piRNA cluster in mice leads to increased retrotransposon activity, consistent with a role for piRNAs in retrotransposon regulation (Xu et al., 2008). To date, two additional host-encoded factors were demonstrated to be involved in the PIWI-mediated restriction of L1 retrotransposons: MVH (mouse vasa homolog) is the homolog of the VASA protein which is an evolutionarily conserved RNA helicase essential for germ cell development in Drosophila. Expression of the murine MVH is restricted to the germ cell lineage and can be observed in male germ cells from embryonic day 10.5 to round spermatids (Toyooka et al., 2000), which covers the period of de novo DNA methylation of retrotransposons. Since MILI and MIWI were found to bind to MVH (Kuramochi-Miyagawa et al., 2004), it was postulated that MVH plays some role(s) in piRNA production and subsequent DNA methylation of retrotransposons. Indeed, essential roles for MVH in de novo methylation of retrotransposons could be confirmed, and it was demonstrated that MVH is an essential factor in the piRNA processing pathway (Kuramochi-Miyagawa et al., 2010). Also, TDRD1, a tudordomain-containing protein, associates with MILI, participates in the PIWI pathway to suppress retrotransposons (Reuter et al., 2009; Wang et al., 2009) and is essential for retrotransposon silencing and male meiosis in mice. During male germ cell development, Tdrd9 participates in ensuring a proper piRNA profile and in establishing DNA methylation of L1. The TDRD9 protein forms a discrete subcellular compartment with MIWI2 under the control of Mili in fetal prospermatogonia (Shoji et al., 2009).

4. The Use of Transposable Elements in Biotechnology and in Fundamental Studies 4.1. DNA transposons as genetic tools Class II TEs or DNA transposons are discrete pieces of DNA with the ability to change their positions within the genome via a “cut-and-paste” mechanism called transposition. In nature, these elements exist as single units containing the transposase gene flanked by inverted terminal repeats (ITRs) that carry transposase-binding sites (Fig. 3.3A). However, under laboratory conditions, it is possible to use transposons as bicomponent systems, in which virtually any DNA sequence of interest can be placed between the transposon ITRs and mobilized by trans-supplementing the transposase in form of an expression plasmid (Fig. 3.3B) or mRNA synthesized in vitro. In the transposition process, the transposase enzyme mediates the excision of the element from its donor plasmid, followed by

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A ITR

Transposase gene

ITR

ITR

DNA of interest

ITR

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C ITR

DNA of interest

P

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Figure 3.3 General organization of class II transposable elements and mechanism of transposition. (A) Autonomous transposable elements consist of inverted terminal repeats (ITR; black arrows) that flank the transposase gene. (B) Bicomponent transposon vector system for delivering transgenes that are maintained in plasmids. One component contains a DNA of interest between the transposon ITRs carried by a plasmid vector, while the other component is a transposase expression plasmid. Blue triangle represents the promoter (P) driving expression of the transposase. (C) The transposon carrying a DNA of interest is integrated at a chromosomal donor site.

reintegration of the transposon into a chromosomal locus (Fig. 3.3C). This feature makes transposons natural and easily controllable DNA delivery vehicles that can be used as tools for versatile applications ranging from somatic and germline transgenesis to functional genomics and gene therapy (Fig. 3.4). 4.1.1. The Sleeping Beauty transposon system Even though DNA transposons have been extensively harnessed as tools for genome manipulation in invertebrates (Cooley et al., 1988; Thibault et al., 2004; Zwaal et al., 1993), there was no known transposon that was active enough to be used for such purposes in vertebrates. In 1997, Ivics et al. succeeded to engineer the Sleeping Beauty (SB) transposon system by molecular reconstruction of an ancient, inactive Tc1/mariner-type transposon found in several fish genomes (Ivics et al., 1997). This newly reactivated element allowed highly efficient transposition-mediated gene transfer in major vertebrate model species without the potential risk of cross mobilization of endogenous transposon copies in host genomes. Indeed, SB has been successfully used as a tool for genetic modifications of a wide variety of vertebrate cell lines and species including humans (Ivics et al., 2009; Mates et al., 2007, 2009). However, although the resurrected SB element was active enough to be mobilized in vertebrate cells, its transpositional activity still presented a

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• Cell culture – Generating stable lines

• Transgenesis – Active in all vertebrate species

• Insertional mutagenesis – Zebrafish – Xenopus – Mouse, rat

• Gene therapy – New, nonviral delivery method

Figure 3.4 Broad applicability of Sleeping Beauty in vertebrate genetics.

bottleneck for some applications. Requirements for transfection of primary cells and other hard-to-transfect cell types or for remobilization of transposons from chromosomally resident single-copy donor sites demanded an enzyme with more robust activity. In the past years, significant efforts have been put into enhancing SB’s transpositional efficiency and engineering hyperactive versions by mutagenizing and modifying the transposon ITRs and the transposase-coding region (Baus et al., 2005; Cui et al., 2002; Geurts et al., 2003; Mates et al., 2009; Vigdal et al., 2002; Wilson et al., 2005; Zayed et al., 2004). These endeavors yielded a novel hyperactive SB transposase (referred to as SB100X) (Mates et al., 2009) that is up to 100 times more active than the originally reconstructed SB enzyme with its efficiency in transgene delivery reaching those of viral vectors. 4.1.2. Transposons and functional genomics The postgenomic era presented the scientific community with the new challenge of functional annotation of every gene and identification of elaborate genetic networks. Diverse methods have been employed to address this task, including mutational analysis that proved to be one of the most direct ways to decipher gene functions. There are versatile

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strategies for creating mutations, including insertional mutagenesis by discrete pieces of foreign DNA that has the advantage that the inserted DNA fragment can serve as a molecular tag that allows rapid, usually PCR-based, identification of the mutated allele. Since the function of the gene in which the insertion has occurred is often disturbed, such loss-of-function insertional mutagenesis is frequently followed by functional analysis of mutant phenotypes. In many instances, retroviral vectors were utilized to introduce mutagenic cassettes into genomes, but their chromosomal insertion bias does not allow full coverage of genes. The random integration pattern of the SB transposon combined with its ability to efficiently integrate versatile transgene cassettes into chromosomes established this system as a highly useful tool for insertional mutagenesis in both ES cells (Kokubu et al., 2009; Liang et al., 2009) as well as in somatic (Collier et al., 2005; Dupuy et al., 2005) and germline tissues (Carlson et al., 2003; Dupuy et al., 2001; Fischer et al., 2001; Geurts et al., 2006b; Horie et al., 2001; Kitada et al., 2007; Lu et al., 2007; Roberg-Perez et al., 2003) in animal models. Insertional mutagenesis can be applied in cultured, germline-competent stem cells including ES and spermatogonial stem (SS) cells. One advantage of this approach lies in the possibility to perform preselection of modified ES cell clones before generating mutant animals as well as in the possibility to differentiate selected clones into many different tissue types in vitro. It is possible to perform large-scale, SB-based, insertional mutagenesis screens in ES and SS cells by simply transfecting or electroporating transposon donor and transposase expression plasmids into the cells. The amounts of the delivered plasmids can be adjusted for obtaining the desired insertion frequencies per cell. In addition, SB transposons can also be remobilized from chromosomally resident loci and reintegrated somewhere else in the genome by transiently providing the transposase source; such excision–reintegration events can be monitored by using double selection systems, in which excision results in activation of the first and reintegration in activation of the second selection marker (Luo et al., 1998). Since several aspects of physiology in rats have evolved to be more similar to humans than that of mice, it would be highly desirable to link the rat into the process of annotating the human genome with function. However, the lack of technology for generating defined mutants in the rat genome has hindered the identification of causative relationships between genes and disease phenotypes. As an important step toward this goal, an approach of establishing SB transposon-mediated insertional mutagenesis in rat SS cells was recently reported (Izsvak et al., 2010). SB transposition can be used to tag and simultaneously mutate thousands of genes in culture, by taking advantage of gene trap cassettes. Importantly, culture conditions maintain the potential of genetically manipulated SS cells to produce viable sperm cells. The spermatogonial clones were transplanted to repopulate the

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testes of sterilized, wild-type recipient male rats. The stem cell genome is then passed on to transgenic offspring upon crossing the recipient males with wild-type females. Although transposition events in a given target gene occur by chance, the tissue culture conditions allow screening for a large number of events. Transposition-mediated gene insertion and cell culture conditions thus allow generation of libraries of gene knockouts in rat SS cells. This technology has the potential to develop powerful genomic tools for the rat, offering the opportunity to create a bridge between physiology and genomics. Another method, in which TEs are utilized for insertional mutagenesis in animal models, employs a “jumpstarter and mutator” scheme (Carlson et al., 2003; Dupuy et al., 2001; Horie et al., 2001). In this arrangement, mutator transgenic lines carry SB transposon-based gene-trapping vectors, while a jumpstarter line expresses the transposase preferably in the male germ line (Fischer et al., 2001; Horie et al., 2003). Crossing of the two lines results in transposition in the germline of the F1 double-transgenic males, which are then repeatedly crossed with wild-type females to segregate the transposition events that occurred in their sperm cells to separate F2 animals (Fig. 3.4). In the mouse system, a single sperm cell of an F1 male contains, on average, two transposon insertions (Dupuy et al., 2001), and up to 90% of the F2 progeny can carry transposon insertions (Horie et al., 2001). The applicability of this approach has been demonstrated by the identification of mouse genes with either ubiquitous or tissue-specific expression patterns (Carlson et al., 2003; Geurts et al., 2006a; Horie et al., 2003; Yae et al., 2006). Recently, a similar system for SB insertional mutagenesis was also established in rats (Kitada et al., 2007; Lu et al., 2007). 4.1.3. Transposons as vectors for gene therapy Considerable effort has been devoted to the development of gene delivery strategies for the treatment of inherited and acquired disorders in humans. A desirable gene therapy approach should (i) achieve delivery of therapeutic genes at high efficiency specifically into relevant cells, (ii) be adaptable to changing needs in terms of vector design, (iii) minimize the risk of genotoxicity, and (iv) be cost-effective. Adapting viruses for gene transfer is a popular approach; for example, g-retroviral and lentiviral vectors are efficient at integrating foreign DNA into the chromosomes of transduced cells and have enormous potential for lifelong gene expression. A major concern of using retroviral vectors is the potential for mutagenic effects at the sites of genomic integration (Baum et al., 2004; Hacein-Bey-Abina et al., 2003, 2008). Indeed, insertional mutagenesis has been observed in clinical trials using a retroviral vector for gene therapy of X-linked severe combined immunodeficiency (SCIDX1) (Hacein-Bey-Abina et al., 2003, 2008; Thrasher et al., 2006). The clinical use of retroviral vectors can be curtailed due to the limited size of the

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payload, as multiple or large transgenes compromise the efficiency of viral reverse transcription and packaging. Finally, regulatory issues and high costs associated with manufacture of clinical-grade retrovirus hamper their widespread translation into clinical practice. An ideal therapeutic vector would combine the favorable attributes of integrating viral vectors (i.e., stable chromosomal insertion) while significantly reducing the potential for adverse events. Transposons could potentially offer such an alternative (Fig. 3.4). The advantage of SB transposon-based gene delivery is that, due to stable genomic insertion of expression cassettes, it can lead to both longterm and efficient transgene expression in preclinical animal models (Ivics and Izsvak, 2006). Thus, the SB plasmid-based transposon system combines the advantages of viral vectors with those of naked DNA molecules. However, in contrast to viral vectors, transposon vectors can be maintained and propagated as plasmid DNA that makes them simple and inexpensive to manufacture—an important issue regarding the implementation of future clinical trials. The further advantages of the SB system include its reduced immunogenicity, no strict limitation of the size of expression cassettes (Zayed et al., 2004), and improved safety/toxicity profiles (Ivics et al., 2007; Mates et al., 2009; Moldt et al., 2007; Walisko et al., 2008). Since the transposition mechanism does not involve reverse transcription, DNAbased transposon vectors are not prone to incorporate mutations and can tolerate larger and more complex transgenes, including those containing repeated DNA motifs. Moreover, the use of SB-based gene delivery eliminates the risk of rearrangements of the expression cassette that, as part of a transposing unit of DNA, integrates into chromosomal DNA in an intact form (Ivics and Izsvak, 2006). In comparison to retroviral systems, the SB vectors have an inherently low enhancer/promoter activity (Moldt et al., 2007; Walisko et al., 2008). Inserting insulator sequences flanking the transcription units of the cargo to prevent accidental trans-activation of promoters of neighboring genes further increased the safety features of the SB system (Walisko et al., 2008). Notably, the transposase can be provided as mRNA, thereby reducing the risk of “re-hopping” of the transposonbased vector (Wilber et al., 2006). Chromosomal integration of SB transposons is precise and no SB-associated adverse effects have been reported (Fernando and Fletcher, 2006; Ivics and Izsvak, 2006). The past couple of years have seen a steady growth in interest in applying the SB system for the treatment of a number of conditions including hemophilia A and B (Liu et al., 2006; Ohlfest et al., 2005b), junctional epidermolysis bullosa (OrtizUrda et al., 2002), tyrosinemia I (Montini et al., 2002), glioblastoma (Ohlfest et al., 2005a), Huntington disease (Graepler et al., 2005), and type 1 diabetes (He et al., 2004). In addition, important steps have been made toward SB-mediated gene transfer in the lung for potential therapy of alpha-1-antitrypsin deficiency, cystic fibrosis, and a variety of cardiovascular diseases (Belur et al., 2003; Liu et al., 2004). Thus, the establishment of

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nonviral, integrating vectors generated considerable interest in developing efficient and safe vectors for human gene therapy (Essner et al., 2005; Hackett et al., 2005; Izsvak and Ivics, 2004). The first clinical application of the SB system (Fig. 3.4) will be tested using autologous T cells genetically modified to redirect specificity for B-lineage malignancies (Williams, 2008). Lymphocytes are a suitable initial platform for testing new gene transfer systems as there have been hundreds of infusions of clinical-grade T cells genetically modified using viral and nonviral approaches without apparent genotoxicity (Bonini et al., 2003). The SB transposon to be introduced in the first-in-human application carries a chimeric antigen receptor (CAR) to render the T cells specifically cytotoxic toward CD19þ lymphoid tumors (Huang et al., 2008; Tuteja et al., 2001). The advantage of using the SB system for the genetic modification of T cells includes the reduced cost associated with manufacturing clinical-grade DNA plasmids compared with recombinant viral vectors. This is important when one considers that trials infusing CARþ T cells are only now beginning to demonstrate antitumor effects (Pule et al., 2008; Till et al., 2004). The higher enzymatic activity of SB100X might enable to achieve integration efficiencies comparable to that of retroviral vectors for next-generation trials. The next phase of preclinical research will focus on further refinement in large animal models to undertake SB-mediated transposition in vivo and improving the safety profile of SB vectors by target-selected transgene integration into genomic “safe harbors.” While it remains to be seen whether the first clinical application of the SB system will result in an antitumor effect, this trial will help validate the safety of this approach and allow investigators to revisit the design of DNA vectors in general to help improve the therapeutic effect in subsequent next-generation trials.

4.2. Retrotransposons as genetic tools 4.2.1. General introduction DNA transposons, the mobile elements that move via a “cut-and-paste” mechanism, have been used in various types of biotechnology. However, substantial genome projects have revealed that the genomic proportion of retrotransposons, the mobile elements that move by a “copy-and-paste” mechanism, often exceeds that of DNA transposons (Table 3.1). The recent availability of genome information and the advances in understanding of retrotransposition mechanisms allow us to utilize retrotransposons for biotechnological applications such as genetic markers, insertional mutagenesis, and gene delivery vectors. Retrotransposons are widespread in metazoan genomes. Most integrate into random sites of the host genome, but some have a sequence preference. A few subclades of LINEs show highly sequence-specific integration into

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Table 3.1 Comparison of contents of transposable elements in the genome Retrotransposons

Human (Homo sapiens) Mouse (Mus musculus) Dog (Canis familiaris) Chicken (Gallus gallus) Fruit fly (Drosophila melanogaster) Silkworm (Bombyx mori) Mosquito (Aedes aegypti) Rice (Oryza sativa)

LINEs (%)

SINEs (%)

LTRs (%)

DNA transposons (%)

21.1 19.5 19.2 6.5 1.3

13.6 8.0 10.8 * *

8.6 10.4 3.9 1.3 3.3

3.0 0.9 2.0 0.8 0.5

13.8 14.4 1.2

12.8 1.9 0.1

1.7 10.5 5.4

3.0 3.0 9.3

*, not stated. The data sources are as follows: human, mouse, and dog, Lindblad-Toh et al. (2005); chicken, International Chicken Genome Sequencing Consortium (2004); fruit fly, Bergman et al. (2006); silkworm, Osanai-Futahashi et al. (2008); mosquito, Nene et al. (2007); rice, Yu et al. (2002).

the genome. In this chapter, we introduce retrotransposons whose transposition mechanisms have been well studied in various organisms and discuss retrotransposition assay systems and their availability for biotechnological tools, focusing especially on the sequence specificity used for site-specific gene delivery. Several genetic elements such as homing endonuclease genes (HEGs) (Edgell, 2009; Marcaida et al., 2010; Paques and Duchateau, 2007; Redondo et al., 2008; Remy et al., 2010) and engineered zinc-finger nucleases (ZFNs) (Beumer et al., 2008; Geurts et al., 2009; Le Provost et al., 2010; Shukla et al., 2009; Townsend et al., 2009) have been toward the practical use for the site-specific gene delivery or transgenic tools in various organisms. In this chapter, however, we do not deal these subjects because they are not retroelements. Mobile group II introns, retroTEs found in bacterial and organellar genomes, are also hoped to be used for site-specific DNA integration in gene targeting (Cui and Davis, 2007; Lambowitz and Zimmerly, 2004; Mastroianni et al., 2008; Yao et al., 2005; Zhuang et al., 2009). The group II introns recognize their target DNA sites mainly by base paring of the intron RNA, and thus it is possible to design the target site simply by modifying the RNA. Such programmable gene-targeting vectors named “targetrons” from Lactococcus lactis L1.LtrB introns have been used in various bacteria. Recently, the group II intron knockout technology is also expanding in application for eukaryote systems. There are several detailed reviews for group II introns toward application (Cui and Davis, 2007; Lambowitz and Zimmerly, 2004) and thus we focus other REs in this chapter.

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4.2.2. Retrotransposons in insects, focusing on sequence-specific integration 4.2.2.1. Retrotransposons in Drosophila Although many putative retrotransposons have been identified in the genomes of Drosophila and other insects, few of them have been shown to have retrotransposition activity. An LTR retrotransposon in Drosophila melanogaster, gypsy, has been shown to integrate to the ovo gene in a site-specific manner (Mevel-Ninio et al., 1989). Its retrotransposition activity was shown by a mating experiment (Mevel-Ninio et al., 1989), a microinjection experiment of egg plasm (Kim et al., 1994) or a feeding experiment (Song et al., 1994). Gypsy integrates into the ovo locus at a high frequency that is determined by multiple DNA-binding sites within the gene (Labrador and Corces, 2001). At the end of the gypsy unit, there is a short insulator sequence (Gdula et al., 1996) that has been used in various experiments (Gerasimova and Corces, 2001). Importantly, gypsy works in other organisms including Saccharomyces cerevisiae (Donze and Kamakaka, 2001) and mice (Yao et al., 2003). Recently, the importance of the relative orientation of two gypsy sequences was shown for enhancer blocking activity (Labrador et al., 2008). Gypsy was established as a useful general genetic tool for several organisms. Another LTR retrotransposon in D. melanogaster, ZAM, seems to have a sequence preference for its integration. The recombinant protein of EN domain recognizes CGCGCG within the white gene (Faye et al., 2008), although its retrotransposition activity was not directly shown. A randomly integrated LINE, I factor, was originally found in the I-R hybrid dysgenesis of D. melanogaster (Bucheton, 1990). The first direct evidence of LINE transposition intermediating RNA was shown with I factor ( Jensen and Heidmann, 1991; Pelisson et al., 1991). Various markers were tagged with I factor to investigate its retrotransposition activity. It shows similarity to the mammalian L1 family, is transcribed from an internal promoter by RNA polymerase II, is capped at the 50 -end, polyadenylated, and prefers A-rich regions for integration. Most studies have been done in vivo, crossing R females and I males, with a few in vitro studies in cell culture ( Jensen et al., 1994). The features of such experiments limited the application of I factor to Drosophila; however, the experiments showed the mechanism of retrotransposon silencing by disappearing female sterility through several generations (Chambeyron and Bucheton, 2005). 4.2.2.2. Target-specific LINEs in insects It is of interest that many LINEs found in insects have specific target sequences in the host genome. The existence of R1 and R2, inserted in the specific sites of 28S rDNA, was originally suggested in D. melanogaster (Roiha et al., 1981) and the silkworm Bombyx mori (Fujiwara et al., 1984), and was later confirmed as a LINE (Burke et al., 1987; Xiong and Eickbush, 1988). In particular, fine studies of

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R2 using in vitro (Luan and Eickbush, 1995; Luan et al., 1993) and in vivo retrotransposition assays (Eickbush et al., 2000) clarified general aspects of the TPRT mechanisms that are peculiar to LINE. More recently, several R elements (R4 (Burke et al., 1995), R5 (Burke et al., 2003), R6 (Kojima and Fujiwara, 2003), R7 (Kojima and Fujiwara, 2003), R8 (Kojima et al., 2006), and RT (Besansky et al., 1992)) that insert into different sites of rDNA than do R1 and R2, telomeric repeat-specific LINEs, SART (Takahashi et al., 1997), and TRAS (Kubo et al., 2001; Okazaki et al., 1995) and other several target-specific LINEs that integrate into microsatellite or other repetitive sequences in the genome, have been found, mainly in insect genomes (Kojima and Fujiwara, 2004). Of the 16 clades of LINE known to date, early branched groups occupy five clades including the well-studied R2 clade. While many early branched LINEs are target site specific, not all members have this specificity. Early branched LINEs encode only one ORF including a restriction endonuclease-like endonuclease (RLE) close to the C-terminus (Yang et al., 1999) (Fig. 3.5A). This RLE is involved in a sequence-specific digestion of the target, but its functional role is not certain. However, the N-terminal domain of R2 from B. mori genome has been shown to bind target DNA substrate in vitro (Christensen et al., 2005). The rest of the LINE clades are categorized as recently branched LINEs, which normally encode two ORFs and an AP endonuclease-like endonuclease (APE) at the N-terminus of ORF2 (Fig. 3.5A). Most are not target specific, but the R1 and Tx clades include target-specific elements (Kojima and Fujiwara, 2004, 2005a). The major domain involved in the target specificity of recently branched LINEs is thought to be APE, because it cuts the target DNA in a sequence-specific manner in vitro (shown for R1 (Feng et al., 1998) and TRAS1 (Anzai et al., 2001) elements). The comparison of the crystal structure of the isolated APE domains from target-specific LINEs TRAS1 and R1 and the nonspecific LINE human L1 clarify the putative amino acids involved in target-specific recognition and digestion (Maita et al., 2004, 2007). Regions other than the APE domain also may play a role in target-specific integration. The ORF1 (Gag) proteins of the telomere-specific LINEs TART, HeT-A (Rashkova et al., 2002), and SART1 (Matsumoto et al., 2004, 2006) show a punctate localization in nuclei, which may be involved in recruiting the LINE unit to the telomere region. In addition, read-through transcripts of R1 are necessary for its precise integration (Anzai et al., 2005). 4.2.2.3. Target-specific retrotransposition systems and their application In most transposition systems, plasmid DNA is injected into embryos or transfected into cells. For target-specific LINEs from insects, however, a novel retrotransposition system using baculovirus AcNPV has been developed (Takahashi and Fujiwara, 2002) (Fig. 3.5B).

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A

Typical structure of two types of LINE

Early branched LINE 5⬘UTR

3⬘UTR

ORF Reverse transcriptase RT

Poly A

RLE

Restriction endonucleaselike endonuclease

Zinc-finger

Recently branched LINE 5⬘UTR

ORF1

APE

RT

Zinc fingers AP endonucleaselike endonuclease

B

3⬘UTR

ORF2 Reverse transcriptase

Zinc-finger Poly A

RH RNase H

AcNPV-hyp mediated LINE retrotransposition assay system Target-specific LINE AcNPV vector

ORF1

AcNPV vector

ORF2

Sf9 cells (or other cells)

Virus particle

Target-specific retrotransposition

Chromosome

Infection and expression LINE machinery

Detection by PCR

Figure 3.5 Structure- and virus-mediated retrotransposition assay of target-specific LINE. (A) Schematic structures of two types of LINE are shown. Early branched LINE has only one open reading frame (ORF) and recently branched LINE two ORFs. There are some target-specific LINEs in both types. Vertical lines in ORF represent zincfinger motifs. (B) Target-specific LINE recombinated in the baculovirus (AcNPV) vector can infect Sf9 (insect) cells or other cells and be expressed effectively. Targetspecific LINE integrates into the specific sequence of the host chromosome and its integration can be easily detected by PCR.

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The telomere-specific LINEs TRAS and SART, when recombined in the AcNPV vector, can integrate into their respective specific targets when they are infected in Sf9 insect cells, and their target can be altered by swapping APEs between them (Takahashi and Fujiwara, 2002). This system is also available for other target-specific LINEs, 28S rDNA-specific LINEs, R1, R2 (Fujimoto et al., 2004), and R7 and R8 (unpublished data). Many target-specific LINEs recognize the 30 -UTR of LINE–RNA during the initial step of the TPRT process. A sequence such as a fluorescent protein reporter with an added 30 UTR sequence can be retrotransposed in a target-specific manner with a helper LINE by transcomplementation, a useful feature for a gene delivery tool (Osanai et al., 2004). This baculovirus-mediated retrotransposition system, which enables very high expression of the LINE machinery, also has the advantage that it is easily converted to an in vitro (Matsumoto et al., 2006) retrotransposition assay. The purified retrotransposition machinery for SART, after being expressed with AcNPV vector in Sf9 cells, can integrate specifically in vitro at (TTAGG) telomeric repeats. The in vitro assay contributes to understanding of not only the mechanism of target-specific retrotransposition but also the structural features of LINEs. The baculovirus AcNPV, which is derived from a moth, has a wide host range and can infect a broad range of species including human cells. Thus, the AcNPV-mediated system is thought to be available in vivo in various species. It has been shown that AcNPV recombined with R1 and SART1 can integrate at specific target sites in the genome of various organs when injected into larva of B. mori (Kawashima et al., 2007), and that this gene transfer system is also useful in the honeybee (Ando et al., 2007). This AcNPV-mediated LINE system is also applicable for human and fish cell lines as a target-specific gene delivery tool (Kawashima, unpublished data). 4.2.3. Retrotransposons in fish 4.2.3.1. Randomly integrated LINEs and SINEs in fish Several LINEs have been found in fish genomes and some of them have been shown to have retrotransposition activity. UnaL2, found in the eel genome, was shown to move in human HeLa cells by being tagged with an inverted neo gene divided by an intron (Kajikawa and Okada, 2002). UnaL2 integrates randomly into the genome. It also recognizes 30 -UTR sequences of specific “passenger” SINEs like UnaSINE1 and UnaSINE2 and mobilizes them by transcomplementation (Kajikawa and Okada, 2002; Kajikawa et al., 2005), which is a useful feature for a gene delivery tool. ZfL2-1 and ZfL2-2, found in the zebrafish genome, also can retrotranspose in HeLa cells (Sugano et al., 2006; Tamura et al., 2007). ZfL2-1 has two ORFs and ZfL2-2 has one ORF but both belong to the same clade L2. These LINEs have the ability to mobilize in zebrafish in vivo.

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4.2.3.2. Target-specific LINEs in fish, good candidates for gene therapy tools We have recently identified a 28S rDNA-specific LINE R2Ol from the genome of the medaka fish, Oryzias latipes (Kojima and Fujiwara, 2005b), which was inserted at the same site as R2 elements in insects. We have established a sequence-specific retrotransposition assay system for R2Ol in zebrafish embryos by microinjecting mRNA transcribed in vitro. R2Ol shows effective retrotransposition activity of around 50%, and we obtained transgenic F1 fish retaining integrated R2Ol (Yatabe, manuscript in preparation). This in vivo retrotransposition assay in zebrafish has advantages for investigating LINE mechanisms through development or through several generations. We also succeeded in retrotransposing R2Ol into a precise site of 28S rDNA of several human cell lines (Kawashima, manuscript in preparation), suggesting that R2Ol is a good candidate for a gene therapy tool that avoids undesirable integration into the human genome.

4.2.4. Mammalian retrotransposons and their application as genetic tools 4.2.4.1. Mammalian retrotransposons In many mammals, retrotransposons account for a notable portion of the genome. Genome sequencing has revealed that there are many inter- and intraspecies polymorphisms in retrotransposon insertion. The de novo insertion of a retrotransposon is stable compared to that of a DNA transposon, which enables us to investigate molecular phylogeny by analysis of retrotransposon insertions. Advances in understanding of the retrotransposition mechanisms of L1 and intracisternal A particle (IAP) elements allow the exploration of the possibility of utilizing these elements in insertional mutagenesis and gene delivery. 4.2.4.2. Genetic markers Retrotransposons have been used as genetic markers for nearly two decades. SINE insertions have been used to analyze phylogenetic relationships in fish (Murata et al., 1993; Takahashi et al., 2001), the evolution of whales (Nikaido et al., 1999; Shimamura et al., 1997), and the evolutionary history of humans and primates (Batzer et al., 1994; Li et al., 2009; Perna et al., 1992; Stoneking et al., 1997). The progress of genome sequencing in mammals is accelerating the use of retrotransposons as intraspecies genetic markers. In humans, some retrotransposons have been inserted so recently that they are polymorphic for presence or absence among populations and individuals. These polymorphisms are also being used as forensic tools (Ray et al., 2007). In dogs, it is reported that more than 10,000 loci are bimorphic for SINE insertions, which may be used as genotyping markers or for identifying the ancestral relationships between dog breeds (Wang and Kirkness, 2005). Active retrotransposons are also reported in the bovine genome (Adelson et al., 2009), so retrotransposons are promising as genetic markers in breed identification of cattle. Interspecies identification can also be conducted by detecting retrotransposon sequences:

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for instance, it can be identified whether the meat in sausages is derived from beef, pork, chicken, or ruminant materials (Ray et al., 2007). 4.2.4.3. Mutagenesis Attempts at using retrotransposons in random mutagenesis have been made in mammals. Insertional mutagenesis using retrotransposons has several advantages over methods based on chemicals such as ENU (N-ethyl-N-nitrosourea). One advantage is that while ENU creates point mutations that often do not affect gene function, retrotransposons can be used to deliver a gene trap that efficiently splices into genes and disrupts their function. Another is that the mutated genes can be cloned more easily when retrotransposons are used compared with when ENU is used. Retrotransposon-based mutagenesis is also likely to have some advantages over DNA transposon-based mutagenesis. DNA transposons have a tendency to insert novel copies close to their original genomic localization, so-called local hopping effect (Fischer et al., 2001; Guimond et al., 2003; Horie et al., 2003; Keng et al., 2005; Scali et al., 2007; Wang et al., 2008), but retrotransposons such as L1 do not have this tendency and insert randomly into the genome (An et al., 2006; Babushok et al., 2006). In addition, retrotransposon insertion appears to be more stable than DNA transposon insertion. Retrotransposon insertions are fixed in the genome, whereas the inserted DNA transposon has a possibility of being excised if a DNA transposase gene exists in the host genome. Intense studies of the application of human and mouse L1 retrotransposons as insertional mutagenesis tools (Ostertag et al., 2007; Rangwala and Kazazian, 2009) have recently been made. L1 is a LINE-type retrotransposon that inserts randomly into the genome without apparent locus specificity. L1 retrotransposition has been detected in cultured cells of human (Moran et al., 1996), mouse (Moran et al., 1996), rat (Kirilyuk et al., 2008; Muotri et al., 2005), hamster (Morrish et al., 2007), and chicken (Suzuki et al., 2009; Wallace et al., 2008) and also in transgenic mice (Kano et al., 2009). In a cultured cell assay, L1-expressing plasmids were transfected into the cultured cells. The rate of L1 retrotransposition varied from 0.04% to 10% of cells containing the L1 plasmid, depending on the kind of L1 element used (Han and Boeke, 2004; Moran et al., 1996). The codonoptimized mouse L1 has the highest retrotransposition rate so far detected (Han and Boeke, 2004). Transgenic mice have been generated for studying L1 retrotransposition in vivo (An et al., 2006, 2008; Babushok et al., 2006; Muotri et al., 2005; Ostertag et al., 2002; Prak et al., 2003). These transgenic mice contain the L1 retrotransposon that has a gene cassette for detecting de novo retrotransposition inside the L1 30 -UTR (L1/transgene). In many transgenic lines, the gene cassette is designed so that the EGFP gene will be expressed when L1 retrotransposition occurs. At the cell culture level, a gene cassette for gene-trapping purposes that consists of a bidirectional poly(A) signal

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sandwiched between two oppositely oriented splice acceptors was effective (An et al., 2006). Transcription of the L1/transgene in transgenic animals is driven by the endogenous promoter in the L1 50 -UTR (Muotri et al., 2005; Ostertag et al., 2002) or a heterologous promoter such as mouse RNA polymerase II large subunit promoter (Ostertag et al., 2002; Prak et al., 2003), the Hsp70-2 promoter (Babushok et al., 2006), or the CAG promoter (An et al., 2006, 2008). The use of a heterologous promoter increases the frequency of retrotransposition events (Ostertag et al., 2002). The highest germline retrotransposition frequency so far reported is that of the codon-optimized mouse L1, ORFeus. In a transgenic line that has 10 copies of the donor ORFeus/transgene (An et al., 2006), the germline insertion frequency was 33%, while a recently generated transgenic line that has a single copy of ORFeus/transgene had a germline insertion frequency of 9.4% (An et al., 2008). In transgenic mice carrying the L1/transgene, de novo insertions have been detected in all chromosomes (An et al., 2006). This contrasts strikingly with DNA transposons that have a tendency to insert close to their original genomic localization, a process called “local hopping” (Fischer et al., 2001; Horie et al., 2003; Keng et al., 2005; Wang et al., 2008). Tissue-specific activation systems for the L1 transgene using Cre/loxP have been developed, and specific activation in germline and pancreas has been reported (An et al., 2008). The tissue-specific activation system in combination with a gene trap cassette containing strong splice acceptors is useful for somatic mutagenesis. One application of L1-based somatic mutagenesis in mice may be a screening system for tumor suppressors and oncogenes (Ostertag et al., 2007). In addition to the L1 retrotransposon, there is also the possibility of using the IAP retrotransposon for insertional mutagenesis. IAP is a mouse LTR retrotransposon, and a cell culture assay has been established for examining IAP retrotransposition. IAP retrotransposition is detected in 0.2–1.0% of the cells transfected with an IAP construct (Dewannieux et al., 2004; Horie et al., 2007; Saito et al., 2008). 4.2.4.4. Gene delivery vectors There are also studies using L1 for a gene delivery vector in combination with helper-dependent adenoviruses (Kubo et al., 2006; Soifer and Kasahara, 2004; Soifer et al., 2001). Adenovirus lacks the machinery for efficient integration into host chromosomes, and rarely integrates into the genome, but efficiently infects many cell types. The helper-dependent adenovirus is engineered to lack all the coding sequences that could be toxic or immunogenic to the host (Kochanek et al., 1996; Mitani et al., 1995; Schiedner et al., 1998). There are several advantages in using L1–adenovirus rather than retrovirus in gene delivery. Retrovirus integrates preferentially into active genes, but L1 integrations are random, so there is a smaller possibility of disrupting

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other genes. Retroviral integration always includes the LTR sequences, which have strong promoter activity and have a risk of activating genes flanking the insertion site, but L1 usually loses its endogenous promoter during integration. In the L1–adenovirus hybrid vector, the adenovirus delivers the L1/ transgene element into the cells, and the L1 integrates the transgene into the genome. Helper-dependent adenovirus does not propagate in the infected cells. By the use of helper-dependent adenovirus, L1/transgene retrotransposition frequencies of up to 91% of infected cells have been observed (Kubo et al., 2006). This is markedly greater than the retrotransposition efficiency achieved by direct plasmid transfection. The maximum retrotransposition efficiency achieved by direct plasmid transfection is about one cell per 10 transfected cells using the highly active ORFeus construct (Han and Boeke, 2004). In the traditional plasmid transfection method, most of the L1 retrotransposition reported was in transformed or immortalized cells. In the L1/adenovirus system, the high transduction ability of the adenovirus allows L1/transgene retrotransposition in differentiated human primary somatic cells, including dermal fibroblasts and hepatocytes, and furthermore, in nondividing cells arrested in the G1/S cells, suggesting the potential for utilization in gene therapy. Besides GFP expression cassettes, short hairpin RNA (shRNA) expression cassettes have been delivered by L1 to the genomes of cultured cells (Yang et al., 2005). Although in this experiment L1/shRNA was transduced to cells by the traditional plasmid transfection method, it was demonstrated that both exogenous (GFP) and endogenous (GADPH) gene expression could be reduced by L1-transmitted shRNA. One copy of retrotransposed L1 with a GFP shRNA cassette was sufficient to reduce GFP fluorescence by 90%. This report suggests that the L1-based RNAi system is promising as a stable gene-silencing system in human cells.

5. Domestication of Mobile DNA by the Host Genomes 5.1. Genomic repeats as transcriptional promoters Recent whole-genome analysis revealed that about 25% of all human promoters contain REs in their sequence (van de Lagemaat et al., 2003). Moreover, 7–10% of experimentally characterized transcription factorbinding sites (TFBS) were shown to be derived from repetitive sequences including simple sequence repeats and TEs (Polavarapu et al., 2008). TFBS that originated from repeats evolve more rapidly than nonrepetitive TFBS but still show signs of sequence conservation on functionally critical bases.

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RE Serve as enhancers of gene transcription RE Disrupt gene exon-intronic structure

RE Provide alternative promoters Genomic repeats

RE Cause premature transciptional termination

enh RE Serve as insulator elements

RE Serve as transcriptional silencers

RE Generate antisense RNAs

Figure 3.6 Genomic repeats influence on transcription of the host genes. Red boxes, retroelements; green boxes, gene exons; green arrow, gene transcriptional start site; purple oval, enhancer element.

Such rapidly evolving TFBS are likely to direct species-specific regulation of gene expression, thus participating in evolutionary process (Fig. 3.6). In the majority of examples reported to date, REs act as alternative promoters. REs can either influence the level of a corresponding RNA transcription or change the tissue specificity of its expression. For example, LTR integration into CYP19 gene, encoding for aromatase P450, the key enzyme in estrogen biosynthesis, led to the formation of alternative promoter located 100 kb upstream of the coding region (van de Lagemaat et al., 2003). This event resulted in the primate-specific transcription of CYP19 in the syncytiotrophoblast layer of the placenta. Placental-specific expression might play an important role in controlling estrogen levels during pregnancy. Placental-specific transcription driven from endogenous retroviral promoters was also shown for Mid1 gene linked with inheritable Opitz syndrome (Landry et al., 2002), endothelin B receptor (Medstrand et al., 2001), and insulin-like growth factor INSL4 (Bieche et al., 2003). Solitary ERV-L LTR was shown to promote b3GAL-T5 transcription in various tissues, being especially active in colon, where it is responsible for the majority of gene transcripts (Dunn et al., 2005). b3GAL-T5 is involved in the synthesis of type 1 carbohydrate chains in gastrointestinal and pancreatic tissues. Interestingly, murine b3GAL-T5 gene is also expressed primarily in colon, despite the absence of an orthologous LTR in the mouse genome. It is likely that in humans the LTR adopted the function of an ancestral mammalian promoter active in colon (Dunn et al., 2005). An interesting example of gene transcriptional regulation by LTR was shown for NAIP (BIRC1) gene coding for neuronal apoptosis inhibitory protein (Romanish et al., 2007). Although human and rodent NAIP

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promoter regions share no similarity, in both cases LTR serve as an alternative promoter. Thus, two different LTR retrotransposons were recruited independently in primate and rodent genomes for the gene transcriptional regulation. REs may also represent the only known promoter for some human genes. For example, the only apparent promoter of the liver-specific BAAT gene recently implicated in familial hypercholanemia is an ancient LTR in human but not in mouse (Carlton et al., 2003). Antisense L1 and Alu sequences were shown to act as the unique promoter for HYAL-4 gene, necessary for hyaluronan catabolism (van de Lagemaat et al., 2003). The application of novel high-throughput techniques such as cap analysis of gene expression (CAGE) and paired-end ditag (PET) sequencing revealed 51,197 ERV-derived promoter sequences. In 1743 cases, ERVs were located in gene proximal or 50 -UTRs. In all, 114 ERV-derived transcription start sites can be demonstrated to drive transcription of 97 human genes, producing chimeric transcripts initiated within LTR and read-through into known gene sequences (Conley et al., 2008b). Recently, we found that at least 50% of human-specific HERV-K LTRs possess promoter activity and the level of their expression ranges from 0.001 to 3% of the beta-actin gene transcriptional level (Buzdin et al., 2006a,b). We have also shown that 50 -proviral LTR is more transcriptionally active than 30 -proviral or solitary LTRs and that the relative content of promoter-active LTRs in gene-rich regions is significantly higher than that in gene-poor loci.

5.2. REs as enhancers for host cell gene transcription One of the first striking reports of the involvement of REs in tissue-specific gene transcriptional regulation was for the human amylase locus (Meisler and Ting, 1993). In humans, amylase is produced in pancreas and in salivary glands. Human amylase locus includes two genes of pancreatic amylase (AMY2A and AMY2B) and three genes of salivary amylase (AMY1A, AMY1B, and AMY1C). The latter three genes are likely products of a recent triplication, because in the chimpanzee genome there is only one gene for AMY1. Exon-intronic structures of these genes are identical, except for an additional untranslated exon at the 50 -terminus of the salivary amylase genes. Moreover, all genes for salivary amylase contain a full-length insert of HERV-E upstream their transcription start site. It was hypothesized that the insertion of full-length ERV activated a cryptic promoter that drives the transcription of amylase in salivary glands. When there is a solitary LTR instead of full-length HERV-E provirus, cryptic promoter cannot be activated and the gene is expressed only in pancreas. There are several other well-supported examples of LTR involvement in gene regulation. For example, the ERV9 LTR element upstream of the

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DNase I hypersensitive site 5 (HS5) of the locus control region in the human b-globin cluster might be responsible for controlling expression of this cluster in erythroid cells (Long et al., 1998). It was suggested that the enhancer effect might be caused by LTR-initiated transcription driven in the direction of associated gene promoter (Ling et al., 2002, 2003). Another example is the mouse Slp (sex-limited protein) gene. ERV-located upstream of the Slp in antisense orientation was shown to direct androgen-specific expression of this gene in males (Loreni et al., 1988). LINEs and SINEs can also serve as transcriptional enhancers. The enhancer of human apoliprotein A was shown to reside within LINE element (Yang et al., 1998). Alu sequence is a part of enhancer element located in the last intron of the human CD8 alpha gene (Hambor et al., 1993). Expression of this gene is restricted to cells of lymphoid lineage and is developmentally regulated during thymopoesis. A CORE-SINE RE (ancient tRNA-derived SINE with a conserved core sequence) was found to represent a neuronal enhancer for the POMC (proopiomelanocortin) gene (Santangelo et al., 2007). POMC encodes a prohormone that gives rise to several bioactive peptides that participate in the stress response, skin and hair pigmentation, analgesia, and the regulation of food intake and energy balance. CORE-SINE was shown to be responsible for the expression of POMC in ventral hypothalamic neurons. Recently, AmnSINEs (a new SINE family identified in the genomes of Amniota) have been shown to act as distal transcriptional enhancers for FGF8 (fibroblast growth factor 8) and SATB2 genes in developing mouse forebrain (Sasaki et al., 2008).

5.3. REs as providers of new splice sites for the host genes Apart from the modulation of transcription, REs can also regulate splicing of pre-mRNA. An outstanding role here belongs to SINE elements, namely Alu retrotransposons in the case of human transcriptome. In a genome-wide comparison of the genomes of human and mouse, a total of 3,932,058 and 3,122,416 TEs have been identified in human and mouse, respectively. Interestingly, 60% of transposons in both human and mouse are located in intronic sequences, even though introns comprise only 24% of the human genome (Sela et al., 2007). All families of transposons in both human and mouse can “exonize,” that is, be included in the exons of mature mRNA. Transposons that are shared between human and mouse exhibit the same percentage of exonization in the two species, but the exonization level of a primate-specific RE Alu is far greater than that of other human transposons. This results in a higher overall level of transposon exonization in human than in mouse (1824 exons compared with 506 exons, respectively) (Sela et al., 2007). Alus are the most abundant repetitive elements in the human genome. The major burst of Alu retroposition took place 50–60 mya and has since dropped to a frequency of one new retroposition for every 20–125

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new births (Batzer et al., 1993; Cordaux et al., 2006). Alus are presented by more than 1.1 million of copies (Chen et al., 2009), and over 0.5 million of them reside in introns of human protein-coding genes (Levy et al., 2008). Alu elements have several sequence motifs resembling consensus splice sites in both sense and antisense orientations (Gotea and Makalowski, 2006), and the insertion of Alu elements into intronic regions may introduce new exons into existing functional genes. It has became a current opinion now that it is the exonization of Alu elements that plays a crucial role in birth of new exons in primate genomes (Corvelo and Eyras, 2008; Lin et al., 2008). Most Alu-derived exons are short (median length of the 330 exons is 121 nucleotides) (Lin et al., 2008). Almost all Alu-derived exons are alternatively spliced, and the vast majority of these exons have low transcript inclusion levels (are included only in the minor transcript isoforms). However, younger Alu-derived exons have weaker splice sites and lower absolute values for the relative abundance of putative splicing regulators between exonic and adjacent intronic regions. This relative abundance was shown to increase with exon age, leading to higher exon inclusion (Corvelo and Eyras, 2008). Furthermore, using exon array data of 330 Alu-derived exons in 11 human tissues and detailed RT-PCR analyses of 38 exons, it has been demonstrated that some Alu-derived exons are constitutively spliced in a broad range of human tissues, and some display strong tissue-specific switch in their transcript inclusion levels. Most of these latter exons were derived from ancient Alu elements in the genome (Lin et al., 2008). This is probably due to the fact that exons derived from older Alu elements had more evolutionary time to accumulate nucleotide substitutions that supported exon inclusion in the transcript products (Lin et al., 2008). Alu consists of two monomeres derived each from a truncated copy of 7SL RNA involved in protein sorting. In rodent genomes, there are also multiple copies of 7SL RNA-derived short retrotransposons, but of a monomeric structure. Why there are so many alternatively spliced Alus compared to rodent retrotransposons? The most probable explanation is its dimeric organization (Gal-Mark et al., 2008). Alus are composed of two related but distinct monomers, left and right arms. Most exonizations occur in right arms of antisense Alu elements. Without the left arm, exonization of the right arm shifts from alternative to constitutive splicing. This eliminates the evolutionary conserved isoform of the host gene and thus may be selected against. The insertion of the left arm downstream of a constitutively spliced non-Alu exon shifts splicing from constitutive to alternative. Although the two arms are highly similar, the left arm is characterized by weaker splicing signals and lower exonic splicing regulatory densities. Mutations that improve these potential splice signals activate exonization and shift splicing from the right to the left arm. Interplay between two or more putative splice signals renders the intronic left arm with a pseudoexon function. Thus, the dimeric form of the Alu element fortuitously provides it

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with an evolutionary advantage, allowing enrichment of the primate transcriptome without compromising its original repertoire (Gal-Mark et al., 2008). Overall, Alu-derived exons had significantly weaker splicing signals compared to nonrepetitive constitutively spliced exons and typical cassette exons (other alternatively spliced exons). This is most probably due to a lower density of exonic splicing regulatory elements in Alu-derived exons. Aluderived exons had much higher evolutionary rates during primate evolution, compared to constitutive exons and cassette exons. For exons present in both human and chimpanzee genomes, the overall nucleotide substitution rate of Alu-derived exons was 1.34% compared to 0.73% for cassette exons and 0.52% for constitutive exons. Similarly, between human and orangutan genomes, the overall nucleotide substitution rates of Alu-derived exons, cassette exons, and constitutive exons were 3.69%, 1.81%, and 1.31%, respectively. However, at least six Alu-containing exons (in genes FAM55C, NLRP1, ZNF611, ADAL, RPP38, and RSPH10B) are constitutively spliced in human tissues (Lin et al., 2008; Makalowski et al., 1994; Sorek et al., 2002). In addition, Alu sequence provided a donor splice site to one of the constitutive exons of the human gene encoding survivin (a member of the apoptosis inhibitor family that is overexpressed in many malignancies) (Mola et al., 2007). There is also an excess of Alu-derived internal exons in the 50 -UTRs of the genes compared to the 30 -UTRs. This phenomenon likely reflects stronger purifying selection pressure against exon creation in 30 -UTR because such exons may trigger mRNA nonsense-mediated decay (Lin et al., 2008). In addition, there is an “exclusion zone” in intron sequences flanking exons, where insertion of Alu elements is presumably under purifying selection. The length of this “exclusion zone” is similar to that of the human–mouse conserved sequences flanking alternatively spliced exons ( 80–150 nucleotides) (Lev-Maor et al., 2008). In some genes, Alu elements strikingly increased the average amount of sequence divergence between human and chimpanzee up to more than 2% in the 30 -UTRs. Moreover, 20 out of the 87 transcripts carrying Alu insert either in the 50 - or in the 30 -UTR contained more than 10% structural divergence in length. In particular, two-thirds of this structural divergence was found in the 30 -UTRs, and variable transcription start sites were conspicuous in the 50 -UTRs (Sakate et al., 2007). In both 50 - and 30 -UTR sequences, presence of an Alu element may be important for posttranscriptional regulation of gene expression, for example, by affecting protein translation, alternative splicing, and mRNA stability (Hasler et al., 2007). Interestingly, Alu exonization might have played a role in human speciation. For example, there is a muscle-specific inclusion of an Aluderived exon in mRNA of gene SEPN1 (gene implicated in a form of congenital muscular dystrophy), which appeared due to a human-specific

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splicing change after the divergence of humans and chimpanzees (Lev-Maor et al., 2008). The second example is the functional deletion of an exon within the protein-coding sequence of human gene CMP for syalic acid hydroxylase. Mutation, caused by the human-specific insertion of Alu RE into 92 bp-long CMP exon, disrupted normal ORF for this enzyme and resulted in the lack of N-glycolyl neuraminic acid (Neu5Gc) on a surface of cell membranes (Chou et al., 1998; Irie et al., 1998). Neu5Gc, thus, is replaced in humans by its precursor, N-acetyl neuraminic acid (Neu5Ac). This absence of Neu5Gc is the major biochemical distinction between human and chimpanzee, which, theoretically, may influence intercellular interactions and embryo development, for example, brain organogenesis. A subset of other Alu-derived exons, especially those derived from more ancient Alu elements in the genome, might have contributed to functional novelties during evolution of many other primates as well. Some novel polymorphic Alu inserts interfere with the normal pre-mRNA splicing by providing additional splicing enhancers, thus causing inheritable diseases (Gu et al., 2007). Noteworthy, canine SINE element SINEC_Cf is likely to play a major role in the evolution of dog genomes nowadays (Wang and Kirkness, 2005). Canine genomes harbor a high frequency of alleles that seem to differ only by the absence or presence of a SINEC_Cf repeat. Comparison of an individual dog (a poodle) DNA with a draft genome sequence of a distinct dog (a boxer) has revealed the chromosomal coordinates for >10,000 loci that are bimorphic for SINEC_Cf insertions. Further analysis of SINE insertion sites from the genomes of nine additional dogs indicates an additional 10,000 bimorphic loci could be readily identified in the general dog population. Approximately, half of all annotated canine genes contain SINEC_Cf repeats. When transcribed in the antisense orientation, they provide splice acceptor sites that can result in incorporation of novel exons. The high frequency of bimorphic SINE insertions in the dog population is predicted to provide numerous examples of allele-specific transcription patterns that may be valuable for the study of differential gene expression among the dog breeds (Wang and Kirkness, 2005). LINE elements may be involved in constitutive or alternative splicing of cellular RNAs too, although with relatively lower frequencies. For example, mammalian L1 elements contain numerous functional internal splice sites that generate a variety of processed L1 transcripts (most probably useless for the L1 retrotransposition) and also contribute to the generation of hybrid transcripts between L1 elements and host genes. Interestingly, L1 splicing is delayed during the course of L1 expression (Belancio et al., 2008b). This delay in L1 splicing may also serve to protect host genes from the excessive burden of L1 interference with their normal expression via aberrant splicing (Belancio et al., 2008b). However, an increased ratio of constitutively spliced L1s relatively to alternatively spliced ones has been reported

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compared to Alu elements. Proportion of L1 elements in gene introns is significantly lower than the one of Alu repeats, although both retrotransposons utilize the same retrotranspositional mechanism (Buzdin, 2004). This bias is probably due to a purifying selection acting against accumulation of L1s in genes. In other vertebrate genomes, LINEs also have been reported to generate new chimeric spliced mRNA variants for the host functional genes, for example, in zebrafish (Tamura et al., 2007) or in pig cells (Sironen et al., 2007). LTR retrotransposons also may contribute to a diversity of alternatively spliced RNAs (van de Lagemaat et al., 2006). For example, in the case of human gene VEGFR-3/FLT4 for endothelial angeogenesis controlling receptor, two different isoforms of this protein are encoded by the same gene. Polypeptide encoded by the shorter transcript lacks 65 C-terminal aminoacids. The short VEGFR-3 transcript is formed because of the use of a noncanonical acceptor splice site within the endogenous retroviral sequence located between the exons 1 and 2. These different forms of VEGFR-3 gene product probably have different biological functions (Hughes, 2001). Apart from animal DNA, retrotransposons comprise a significant fraction of plant genomes and are likely involved in gene regulation there too, although the effects of retrotransposon insertions in plants are not well understood. For example, one-sixth of rice genes is associated with retrotransposons, with insertions either in the gene itself or within its putative promoter region. Among genes with inserts in the promoter region, the likelihood of the gene being expressed was shown to be directly proportional to the distance of the retrotransposon from the translation start site. In addition, retrotransposon insertions in the transcribed region of the gene were found to be positively correlated with the presence of alternative splicing forms. Some of the retrotransposons that are embedded in cDNA contribute splice sites and give rise to novel exons (Krom et al., 2008). Yet, the effect of intronic repeats on splicing of the flanking exons is largely unknown. Importantly, more Alus flank alternatively spliced exons than constitutively spliced ones. This implies that Alu insertions may change the mode of splicing of the flanking exons; this is especially notable for those exons that have changed their mode of splicing from constitutive to alternative during primate genome evolution (Lev-Maor et al., 2008). LevMaor and colleagues demonstrated experimentally that two Alu elements that were inserted into an intron in opposite orientation undergo base pairing, as evident by RNA editing, and affect the splicing patterns of a downstream exon, shifting it from constitutive to alternative. It may also be possible that formation of a long and stable double-stranded structure in the upstream intron, especially near the splice site, reduces the ability of the splicing machinery to properly recognize the downstream exon, leading to slower splicing kinetics or suboptimal exon selection and, thus, to intron retention or exon skipping (Lev-Maor et al., 2008).

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Smalheiser and Torvik (2005) showed that a few mammalian miRNA precursors are derived from intronic insertion of two adjacent LINE retrotransposons in opposite orientation, creating a hairpin structure that serves as miRNA precursor. Some other elements have an intrinsic hairpin structure and/or serve as miRNA precursors when inserted into transcriptionally active genomic regions (Hernandez-Pinzon et al., 2009; Piriyapongsa and Jordan, 2007). Many of the newly identified piRNAs are derived from retrotransposons and play a role in transposon silencing in zebrafish germ cells (Houwing et al., 2007; Levy et al., 2008). In order to catalog the data on TEs that may have an impact in gene regulation and functioning, a comprehensive database termed “TranspoGene database” has been constructed that covers genomes of seven species: human, mouse, chicken, zebrafish, fruit fly, nematode, and sea squirt (Levy et al., 2008). The database includes information about repeat localization relative to gene: proximal promoter TEs, exonized TEs (insertion within an intron that led to exon creation), exonic TEs (insertion into an existing exon), or intronic TEs. A variant of this database termed “microTranspoGene” collects the data on human, mouse, zebrafish, and nematode TE-derived miRNAs (Levy et al., 2008). Overall, the proportion of proteins with retrotransposon-encoded fragments (0.1%), although probably underestimated, is much less than what the data at transcript level suggest (4%). In all cases, the RE cassettes are most frequently derived from older REs, in line with the hypothesis that incorporation of TE fragments into functional proteins requires long evolutionary periods. The role of evolutionary recent REs is probably limited to regulatory functions (Gotea and Makalowski, 2006).

5.4. REs as sources of novel polyadenylation signals mRNA polyadenylation is an essential step for the maturation of almost all eukaryotic mRNAs and is tightly coupled with termination of transcription in defining the 30 -end of genes. A polyadenylation signal (AAUAAA) nearby the 30 -end of pre-mRNA is required for poly(A) synthesis. The protein complex involved in the pre-mRNA polyadenylation is coupled with RNA polymerase II during the transcription of a gene, and only RNA polymerase II products are polyadenylated with the remarkable exception of two polyadenylated polymerase III-transcribed RNAs (Borodulina and Kramerov, 2008). Autonomous retrotransposons encode proteins and utilize functional poly(A) signals at the 30 -termini of their genes. Therefore, insertions of these elements in genes in the sense orientation can influence the expression of neighboring genes by providing new poly(A) signals. This is probably the right explanation for the clearly seen strong negative selective pressure on such elements oriented in the same transcriptional direction as the enclosing gene (Buzdin, 2007; Cutter et al., 2005; van de Lagemaat

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et al., 2006; Wheelan et al., 2005; Zemojtel et al., 2007). Indeed, all protein-coding intronic REs (including LINEs and LTR retrotransposons)-oriented sense to gene transcription are underrepresented in all investigated genomes compared to statistically expected ratio of sense/antisense inserts. In contrast, nonautonomous REs like Alu do not employ polyadenylation of their transcripts and, thus, may have only casual AAUAAA sequences. However, such poly(A) signals are very weak and are highly affected by the surrounding sequence (Roy-Engel et al., 2005). Even in the antisense direction relatively to enclosing genes, many retrotransposons provide poly(A) signals that may dynamically modify 30 -ends of genes through evolution. For example, in breast cancer cell line T47D, there were identified four mRNAs polyadenylated at the sequence of HERV-K retroviral LTR (Baust et al., 2000). Transcripts of gene NSBP1 can be alternatively polyadenylated at the retroviral sequences located in the 30 -UTR of that gene (King and Francomano, 2001). 50 -LTR of the retrovirus HERV-F may function as the alternative polyadenylation site for gene ZNF195 (Kjellman et al., 1999). Human genes HHLA2 and HHLA3 utilize HERV-H LTRs as the major polyadenylation signals. In the baboon genome, orthologous loci lack retroviral inserts and these genes recruit other polyadenylation sites (Mager et al., 1999). Interestingly, REs are mostly associated with nonconserved poly(A) sites (Lee et al., 2008a). Of the 1.1 million of human Alu retrotransposons, about 10,000 are inserted in the 30 -UTRs of protein-coding genes and 1% of these (107 events) are active as poly(A) sites (Chen et al., 2009). Alu inserts usually represent weak or cryptic poly(A) signals, but often constitute the major or the unique poly(A) site in a gene. Strikingly, although Alus in 30 -UTR are indifferently inserted in the forward or reverse direction, 99% of polyadenylation-active Alu sequences are forward oriented (Chen et al., 2009). Recently, it was estimated that 8% of all mammalian poly(A) sites are associated with TEs (Lee et al., 2008a). Interestingly, human poly(A) sites that are not conserved in mouse were found to be associated with TEs to a much greater extent than the conserved ones. This result suggests the involvement of TEs in creation or modulation of poly(A) sites in evolution.

5.5. REs as transcriptional silencers Some retrotransposons are known to function as transcriptional silencers by downregulating transcription of the enclosing genes. For example, one out of 44 Alu repeats located in human GH locus encoding for human growth hormone genes hGH-1 and hGH-2 harbors a regulatory element that most probably acts by decreasing the rate of promoter-associated histone acetylation, resulting in a significant decrease of RNA polymerase II recruitment to the promoter. This silencer likely provides for regulatory control of hGH gene expression in pituitary cells (Trujillo et al., 2006).

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Expression of the tumor suppressor protein BRCA2 is tightly regulated throughout development. Sharan et al. identified a transcriptional silencer at the distal part of the human BRCA2 gene promoter. This silencer was involved in the tissue-specific negative regulation of BRCA2 expression in breast cell lines. The former mapped 221-base pair long silencer region was also a part of a full-length Alu element (Sharan et al., 1999). Another example is the transcriptional regulation of a human gene Hpr for haptoglobin-related protein. Hpr sequence is 92% identical to haptoglobin gene HP (Maeda, 1985; Smith et al., 1995). Both genes are transcribed at the higher levels in liver. Hpr promoter is stronger than HP promoter (Oliviero et al., 1987), but the concentration of Hpr liver transcripts is 17fold lower than the one of HP mRNA (Hatada et al., 2003). The major distinction between these genes is the endogenous retroviral sequence RTVL-Ia in the intron of Hpr (Maeda and Kim, 1990). RTVL-Ia fragment has demonstrated significant silencer activity in a series of luciferase transient transfection experiments (Hatada et al., 2003). The mechanism of the negative Hpr regulation by the RTVL-Ia ERV is not clear, but the authors propose that this effect is due to an aberrant splicing of the Hpr transcript with the retroviral sequences. This hypothesis was supported by the identification of the corresponding abnormal transcripts (Hatada et al., 2003).

5.6. REs as antisense regulators of the host gene transcription It has been demonstrated that RE inserts in gene introns are preferentially fixed in the antisense orientation relatively to enclosing gene transcriptional direction (Medstrand et al., 2002; van de Lagemaat et al., 2006). Therefore, promoters of intronic retrotransposons may drive transcription of the RNAs that are complementary to gene introns and/or exons. Moreover, some retrotransposons are also known to possess bidirectional promoter (Copeland et al., 2007; Domansky et al., 2000; Dunn et al., 2006; Feuchter and Mager, 1990; Huh et al., 2008; Matlik et al., 2006), and even downstream insertions of these elements relatively to genes may result in production of the antisense RNA. These complementary RNAs may alter functional host gene expression. The possibility of retrotransposon involvement in antisense regulation of gene expression was suggested few years ago (Mack et al., 2004). Retroposition likely accounts for the origin of a significant number of functional sense–antisense pairs in eukaryotic genomes (Galante et al., 2007). Recently, applied CAGE technology identified 48,718 human gene antisense transcriptional start sites within TEs (Conley et al., 2008a). Gogvadze et al. found the first evidence for the human-specific antisense regulation of gene activity occurring due to promoter activity of HERV-K (HML-2) endogenous retroviral inserts (Gogvadze and Buzdin, 2009;

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Gogvadze et al., 2009). Human-specific LTRs located in the introns of genes SLC4A8 (for sodium bicarbonate cotransporter) and IFT172 (for intraflagellar transport protein 172) in vivo generate transcripts that are complementary to exons within the corresponding mRNAs in a variety of human tissues. As shown by using 50 -RACE technique (rapid amplification of cDNA ends), in both cases the LTR-promoted transcription starts within the same position of the LTR consensus sequence, which coincides with the previously reported HERV-K (HML-2) LTR transcriptional start site (Kovalskaya et al., 2006). The effect of the antisense transcript overexpression on the mRNA level of the corresponding genes was investigated using quantitative real-time RT-PCR. Almost fourfold increase in SLC-AS expression led to 3.9-fold decrease of SLC4A8 mRNA level, and overexpression of IFT-AS transcript 2.9-fold reduced the level of IFT172 mRNA. In all cases, the level of the antisense RNAs in the transfected cells was close to or lower than in many human tissues. Similarly, intronically located representatives of an LTR retrotransposon family from rice genome called Dasheng likely regulate tissue-specific expression of several adjacent functional genes via antisense transcripts driven by the LTRs (Kashkush and Khasdan, 2007). One possible mechanism of the antisense regulation on the pre-mRNA level is connected with the generation of alternatively spliced mRNAs. It has been shown previously that antisense transcripts can inhibit splicing of pre-mRNA in vitro and in vivo (Galante et al., 2007). The possible mechanism involves pairing of antisense transcript and a sense target RNA with the formation of double-stranded RNA that could induce the spliceosome to skip the paired region, thus forming an alternatively spliced transcript. This would result in the formation of nonfunctional RNAs containing multiple premature transcription termination codons. Normally, such RNAs are immediately degraded in the cytoplasm by nonsense-mediated decay machinery (Fasken and Corbett, 2005). Alternatively, antisense transcript base pairing to the target RNA can lead to its rapid enzymatic degradation directly in the nucleus. D. melanogaster genome has no active copies of telomerase gene. Remarkably, transcription of Drosophila retrotransposons HeT-A, TART, and TAHRE that have an important function of maintaining D. melanogaster telomere lengths instead of telomerase is tightly regulated by a specialized RNAi mechanism. This mechanism acts through so-called repeat-associated short interfering (rasi)RNAs. Telomeric retrotransposons are bidirectionally transcribed, and the antisense transcription in ovaries is regulated by a promoter localized within its 30 -UTR. The expression of antisense transcripts of telomeric elements is regulated by the RNA silencing machinery, suggesting rasiRNA-mediated interplay between sense and antisense transcripts in the cell (Shpiz et al., 2009). In the genome of yeasts, Ty1 retrotransposon is most likely regulated by the antisense transcripts

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encompassing its 50 -LTR, that mediate RNA-dependent gene silencing and repress Ty1 mobility. This Ty1 regulatory RNA was shown to repress Ty1 transcription and transposition in trans by acting on the de novo transcribed Ty1 RNA (Berretta et al., 2008). Smalheiser and Torvik (2005) showed that some mammalian miRNA precursors are derived from intronic insertion of two adjacent LINE retrotransposons oriented opposite to each other. Some other elements have an intrinsic hairpin structure and/or serve as miRNA precursors when inserted into transcriptionally active genomic regions (Hernandez-Pinzon et al., 2009; Piriyapongsa and Jordan, 2007).

5.7. REs as insulator elements The temporal and spatial regulation of gene expression is linked to the establishment of functional chromatin domains. Several lines of evidence have been provided recently that retrotransposons can serve in vivo as insulator sequences that distinguish blocks of active and transcriptionally silent chromatin. For example, a B2 SINE element located in the murine growth hormone locus is required for the correct spatio-temporal activation of that gene. This repeat serves as a boundary to block the influence of repressive chromatin modifications by generating short transcripts, which are necessary and sufficient to enable gene activation (Lunyak et al., 2007). Mammalian LINE elements are frequently found within matrix attachment regions (MARs) (Akopov et al., 2006; Purbowasito et al., 2004). Some Drosophila LTR retrotransposons have insulator activity and may block the activity of transcriptional enhancer elements when located between enhancer and promoter (Dorsett, 1993; Kostyuchenko et al., 2008). For example, in some fruitfly lineages, there is an insert of LTR retrotransposon gypsy into the 50 -region of the gene yellow that is responsible for the pigmentation of cuticula. Upstream of the gypsy element there are two enhancer elements that account for the transcription of yellow in different tissues; another enhancer that is responsible for the yellow expression in cilia is located downstream. In the lineage y2, gypsy insertion between the promoter and two upstream enhancers blocks these enhancers and downregulates yellow in the corresponding tissues, but the yellow expression in cilia remains unaffected (Dorsett, 1993).

5.8. REs as regulators of translation Although REs have been found in UTRs of many functional cellular genes, effect of their presence on the translational regulation of gene expression is still poorly investigated. Among the few known examples, there is human zinc-finger gene ZNF177, which incorporates Alu and L1 segments into the 50 -UTR of transcripts. The presence of the Alu and L1 segments which

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form one 50 -UTR exon modifies gene expression on the protein level by decreasing translation efficiency. Interestingly, the same Alu and L1 repeats in the 50 -UTR of ZNF177 exert a positive transcriptional enhancer effect, but repress translation (Landry et al., 2001). Approximately 4% of human 50 -UTRs harbor Alu sequences, indicating that the expression of many genes might be influenced by Alu repeats (Landry et al., 2001). In the mouse genome, there is a SINE retrotransposon-derived gene for neuronal dendrite-specific BC1 RNA. This small, nonprotein-coding RNA is thought to somehow regulate translation in dendritic microdomains. However, the mechanism of such a regulation remains a mystery, and further efforts are needed to investigate this phenomenon (Khanam et al., 2007).

6. Retrotransposons as Drivers of Mammalian Genome Evolution 6.1. REs generate new REs RE integrations into the genome can cause multiple effects and, among them, they may lead to the formation of novel REs, as in case of SVA elements. SVA is a composite element consisting of four parts: hexamer repeats (CCCTCT)n, Alu, 15–23 tandemly repeated sequences (VNTR), and SINE-R (SVA ¼ SINE-R þ VNTR þ Alu) (Ostertag et al., 2003; Wang et al., 2005). These elements are nonautonomous and are mobilized by L1-encoded proteins in trans. The SVA family that is thought to be the youngest genus of primate REs is presented by 3000 copies in the modern human genome (Ostertag et al., 2003). The first SVA probably appeared due to the integration of several elements into the same genomic locus (Wang et al., 2005). SVAs are flanked by TSDs, terminate in a poly(A) tail, and are occasionally truncated and inverted during their integration into the genome. SVA remain active in the human DNA. Several genetic diseases have been reported to be due to SVA insertions (Hancks and Kazazian, 2010). However, their impact in human genome diversity is not restricted to insertion mutagenesis. Evolution of this complex retrotransposon is still going on, first, via quantitative and qualitative changes in tandem repeats, oligomerization, and acquisition of new sequences. This acquisition of genomic sequences by SVA elements may occur in the middle part of an SVA (e.g., due to pseudogene insertion into SVA element), or on SVA termini. Recently, a novel human-specific family of TEs that consists of fused copies of the CpG island containing first exon of gene MAST2 and retrotransposon SVA was discovered (Bantysh and Buzdin, 2009; Damert et al., 2009; Hancks and Kazazian, 2010; Hancks et al., 2009). A mechanism proposed for generation of this family comprises an aberrant splicing event. After the divergence of human and chimpanzee ancestor lineages,

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retrotransposon SVA has inserted into the first intron of gene MAST2 in the sense orientation. Due to splicing of an aberrant RNA driven by MAST2 promoter, but terminally processed using SVA polyadenylation signal, the first exon of MAST2 has fused to a spliced 30 -terminal fragment of SVA retrotransposon. The above ancestor CpG–SVA element due to retrotranspositions of its own copies has formed a novel family presented in the human genome by 76 members. Recruitment of a MAST2 CpG island was probably beneficial to the hybrid retrotransposon as a positive transcriptional regulator. Furthermore, it is speculated that LTR-containing retrotransposons and SINEs themselves represent chimeric elements (Kramerov and Vassetzky, 2001, 2005; Malik and Eickbush, 2001; Ohshima et al., 1996). A phylogenetic analysis of the ribonuclease H domain revealed that LTR-containing REs might have been formed as a fusion between DNA transposon and non-LTR retrotransposon (Malik and Eickbush, 2001). tRNA-derived SINEs likely descended from retroviral strong-stop DNAs (Ohshima et al., 1996). They consist of two regions: a conservative, including a tRNA promoter and a core domain, and a variable one similar to 30 -terminal sequence of different LINE families. The core domain of tRNA-like SINEs has conservative regions similar to fragments of lysine tRNA-primed retroviral LTRs. On the basis of these structural peculiarities, it was suggested that tRNA-derived SINEs emerged due to the integration of retroviral strong-stop DNA into the LINE 30 -terminal part. The RE formed could be transcribed by RNA polymerase III and spread through the genome. Such a mechanism of SINE formation could also explain how these elements can transpose in the genome, namely, it seems very likely that they recruited the enzymatic machinery from LINEs through a common “tail” sequence (Ohshima et al., 1996).

6.2. REs and recombination events Recombination is a powerful factor of evolution that produces genetic variability by using already existing blocks of biological information (Makalowski, 2000). Because of their high copy number and sequence similarity, REs are the substrates for illegitimate homologous recombination, also called ectopic recombination. The chance that an ectopic recombination will occur depends on the number of homologous sequences and on the length of the elements (Boissinot et al., 2006; Song and Boissinot, 2007). Recombination causes genetic rearrangements that can be deleterious, advantageous, or null. There are numerous reported cases of human diseases caused by recombination between REs. For example, glycogen storage disease (Burwinkel and Kilimann, 1998), Alport syndrome (Segal et al., 1999) as a result of recombination between L1 elements and complete germ cell aplasia due to

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recombination between HERV-I (Kamp et al., 2000). Alu elements were implicated in almost 50 disease-causing recombination events (Belancio et al., 2008a; Xing et al., 2009). Apart from deleterious effects, recombination between REs can also have positive consequences. For example, human glycophorin gene family evolved through several duplication steps that involved recombination between Alu elements (Makalowski, 2000). Furthermore, Alu-derived ectopic recombination generated 492 human-specific deletions, the distribution of which is biased toward gene-rich regions of the genome (Sen et al., 2006). About 60% of Alu recombination-mediated deletions were shown to be located in genes and, in at least three cases, exons have been deleted in human genes relative to their chimpanzee orthologs. Finally, L1s were shown to join DNA breaks by inserting into the genome through EN-independent pathway, thus participating in DNA double-strand breaks repair (Morrish et al., 2002).

6.3. Transduction of flanking sequences The ability to transduce 30 -flanking DNA to new genomic loci was firstly shown for L1 elements (Goodier et al., 2000; Moran et al., 1999; Pickeral et al., 2000). L1s have a rather weak polyadenylation signal; therefore, RNA polymerase sometimes gets through it and terminates an RNA synthesis on any polyadenylation site-located downstream. It was estimated that 20% of all L1 inserts contain transduced DNA at the 30 -ends. The length of these sequences varies from few bases to over 1 kb. Taken together, such transduced DNA makes up 0.6–1% of the human genome. Therefore, L1-mediated transductions have the potential to shuffle exons and regulatory sequences to new genomic sites. Recently, it was shown that SVA elements are also able to transduce downstream sequence and it was estimated that about 10% of human SVA elements were involved in DNA transduction events (Ostertag et al., 2003; Wang et al., 2005). Moreover, SVA-mediated transduction can serve as a previously uncharacterized mechanism for gene duplication and the creation of new gene families (Xing et al., 2006). In the latter case, new sequences may appear either on the 50 - or on the 0 3 -terminus of an SVA (50 - and 30 -SVA transduction, respectively). 30 -Transduction mechanism is similar to that proposed for L1 retrotransposon. The size of genomic sequence transferred in such a way may differ from several base pairs to more than 1500 bp. Probably, the most striking example of this phenomenon is the transduction of a whole gene AMAC (acyl-malonyl condensing enzyme 1) in the great ape genomes (Xing et al., 2006). Due to SVA 30 -transduction, human genome has three functional 1.2 kb-long copies of AMAC gene, and at least two of them are transcribed in different human tissues.

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Another kind of transduction results in attaching of new sequences to the 50 -end of an SVA. RE transcription initiation may proceed from any promoter-located upstream in the genomic sequence. In this case, termination of transcription and RNA processing usually occur using normal polyadenylation signal of a RE. This results in a mature RNA having on its 50 -end an additional copy of flanking genomic sequence and a copy of RE at its 30 -end. Subsequent reverse transcription and integration into the genome of a nascent cDNA result in a new RE genomic insert carrying 50 -transduced part (Brosius, 1999a).

6.4. Formation of processed pseudogenes Genomes of all higher eukaryotes contain pseudogenes. These elements normally do not contain introns, end in a poly(A) tail, and are flanked by short direct repeats. Such pseudogenes are referred to as processed pseudogenes (Weiner et al., 1986) and are believed to be produced by the action of LINE retrotransposons (Esnault et al., 2000). As long as RNA polymerase II-transcribed genes generally lack any promoter sequence in their RNA, processed pseudogenes were classically thought to be transcriptionally silent. Indeed, there were not so many reported cases of active pseudogenes that happened to integrate within an existing transcription unit and gave rise to a novel gene or a novel transcriptional pattern of the existing ones. These include jingway element of Drosophila yakuba and D. melanogaster formed due to integration of alcohol dehydrogenase pseudogene into yellow-emperor gene (Long et al., 1999), mouse PMSE2b retrogene inserted into the L1 sequence under the control of LINE promoter (Zaiss and Kloetzel, 1999), mouse PHGP pseudogene, which is expressed from its 50 -adjacent sequence in a tissue-specific manner (Boschan et al., 2002), TRIMCyp gene of owl monkey, formed by retrotransposition of cyclophilin A transcript to intron 7 of TRIM5 ubiquitin ligase and shown to confer HIV-1 resistance in owl monkey (Babushok et al., 2007), and several others. However, recent genome-wide analysis of EST databases as well as transcriptional analyses of individual pseudogenes have revealed that up to a third of processed pseudogenes are transcribed, most of them specifically in testes (Babushok et al., 2007; Vinckenbosch et al., 2006). In humans, >1000 pseudogene transcripts were detected and the number of functionally active pseudogenes was estimated to be 120 (Vinckenbosch et al., 2006). Interestingly, a striking predominance of autosomal retrogenes, which are copies of X-linked parental genes, was shown. These autosomal substitutes probably sustain essential functions during male X chromosome inactivation in the process of spermatogenesis (Babushok et al., 2007; Vinckenbosch et al., 2006).

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6.5. Chimeric retrogene formation during reverse transcription Apart from RE retrotransposition and formation of pseudogenes, RT is also able to change templates during cDNA synthesis. This feature of RT is well known for retroviruses. The RT jumps from one place of the template to another are necessary for the synthesis of retroviral LTRs (Temin, 1993). Template switches can also occur during LINE-directed reverse transcription. Recently, bipartite and tripartite chimeric retrogenes were found in three mammalian and in one fungal genomes. A total of 82, 116, 66, and 31 elements were found in human, mouse, rat, and rice blast fungus Magnaporthe grisea DNAs, respectively (Buzdin et al., 2003, 2007; Fudal et al., 2005; Gogvadze et al., 2007). These elements are composed of DNA copies of different cellular transcripts either directly fused to each other or more frequently fused to the 30 -part of a LINE retrotransposon. The various cellular transcripts found in these chimeras correspond to messenger RNAs, ribosomal RNAs, small nuclear RNAs, 7SL RNA, and Alu retroposon. The chimeras have the following common features: (i) 50 -parts are fulllength copies of cellular RNAs, whereas 30 -parts are 50 -truncated copies of the corresponding RNAs (mostly LINEs); (ii) both parts are directly joined with the same transcriptional orientation; (iii) chimeras have a poly(A) tail at their 30 -end, and (iv) chimeras are flanked by short direct repeats. The last structural feature demonstrates that these elements were transposed as bipartite DNA copies. The simultaneous integration of both parts of these chimeras was confirmed by the data obtained from PCR-based multispecies insertion polymorphism assay (Buzdin et al., 2003). The chimeras were formed by a template switch during LINE reverse transcription. This mechanism was further supported by the direct analysis of LINE retrotranspositions in vitro and in vivo (Babushok et al., 2006; Gilbert et al., 2005). The presence of structurally similar chimeric elements in evolutionary distinct organism shows that template switching during LINE reverse transcription represents an evolutionary conserved mechanism of genome rearrangement. Moreover, many of the chimeras can be considered as new genes, as they were shown to be transcribed, some of them in a tissuespecific manner (Buzdin et al., 2003; Gogvadze et al., 2007). Except generating chimeric retrogenes, template switches during LINE reverse transcription could give rise to chimeric SINE elements (Nishihara et al., 2006) and to mosaic rodent L1 structures (Brosius, 1999a; Hayward et al., 1997). Evolution of certain LINE families might also involve change of a template during reverse transcription, resulting in fusion of the 30 -part of a LINE to a new sequence, as suggested by the observation that the 50 -UTRs of human, mouse, rat, and rabbit L1 families share no considerable sequence identity (Furano, 2000).

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7. Concluding Remarks In this chapter, we have tried to put together the major findings on the impact of TEs in both functioning of eukaryotic cells and in development of modern biotechnology. About 1000 papers on eukaryotic TEs were appearing annually during the past decade, and the total number of publications on the TEs is close to 20,000 for all years. Therefore, a lot of information left beyond the frameworks of this chapter. This ensures also that when this chapter will be published, many novel interesting and/or important related cases will be known. Moreover, the ongoing progress in sequencing technologies gives a realistic promise that not only a qualitative, but also an integrated quantitative figure of the TE impact on the eukaryotic organisms functioning in health and disease will become available in the nearest future.

ACKNOWLEDGMENTS E. V. G. and A. A. B. were supported by the Russian Foundation for Basic Research grants 09-04-12302 and 10-04-00593-a, by the President of the Russian Federation grant MD2010, and by the Program “Molecular and Cellular Biology” of the Presidium of the Russian Academy of Sciences. C. M. is grateful for constant support by Dieter Ha¨ussinger and the Heinz-Ansmann Foundation for AIDS Research. G. G. S was supported by grant DA 545/ 2-1 of the Deutsche Forschungsgemeinschaft. H. F. was supported by grants from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT) of Japan and the Program for Promotion of Basic Research Activities for Innovative Bioscience (PROBRAIN).

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Index

A Activation-induced cytidine deaminase (AID) gene, 124 Adenomatous polyposis coli (APC), 10–11 A3 genes, 125, 130–131 AID gene. See Activation-induced cytidine deaminase (AID) gene APOBEC3 proteins A3 expression profile, human tissues, 130–131 deaminases Alu retrotransposition inhibitors, 129–130 L1 retrotransposition inhibitors, 128–129 LTR retrotransposons inhibitors, 125, 127–128 members of, 124–126 C Chlamydomonas reinhardtii. See also Molecular chaperones asymmetric cell division, 81 chloroplast components, 90 function, 90–101 cytosol/nucleus chaperone components, 81–82 function, 82–85 endoplasmic reticulum components, 89 function, 89 flagella components, 86 function, 86–89 mitochondrion components, 101–102 function, 102 Chloroplast chaperones components, 90 function CDJ3–5, redox switches, 96–97 CGE1 vs. GrpE, 91–94 CPN60, 91 foldosome, 100–101 HSP70B activation, HEP2, 94–96 photosystem II (PS II) protection, 90–91, 99 redox-state, 96 VIPP1, 97–99 Cytoplasmic linker proteins (CLIPs)

cellular pathways, 22–23 dynein pathway, 21–22 mitosis, 22 MT dynamics regulation, 20–21 plus end tracking mechanisms, 20 structure and evolutionary conservation, 19–20 Cytoskeleton-associated protein-glycine-rich (CAP-Gly) cytoplasmic linker proteins (CLIPs), 19–23 dynactin large subunit p150Glued, 23–25 Cytosol/nucleus chaperones components Chlamydomonas, 82 nucleocytosol, 81 Volvox, 82 function Hsp70A, in Volvox embryos, 84–85 stress response regulation, 82–84 D Dynein pathway, 21–22 E EB family cellular structures, 17–18 centrosomes and cilia, 16–17 mitosis, 15–16 MT dynamics regulation, 13–15 MT plus end tracking mechanism, 12–13 structure and evolutionary conservation, 10–12 Endoplasmic reticulum, 89 Eukaryotic retrotransposons LINE retrotransposons, 119 LTR retrotransposons and ERVs, 121–122 penelope-like elements, 122–123 processed pseudogenes, 121 SINE retrotransposons, 120 SVA elements, 120–121 F Fish, retrotransposons randomly integrated LINEs and SINEs, 145 target-specific LINEs, 146 Flagellar chaperones components, 86 HSP70A

189

190

Index

Flagellar chaperones (cont.) capping complex, 87 deflagellation, 86 immunofluorescence analysis, 87 radial spoke protein 12 (RSP12), 89 radial spoke protein 16 (RSP16), 88–89 Foldosome, 100–101 G Genetic tools DNA transposons ITRs, 134–135 Sleeping Beauty transposon system, 135–136 transposons and functional genomics, 136–138 vectors, gene therapy, 138–140 retrotransposons (see also Retrotransposons) DNA transposons, 140–141 fish, retrotransposons, 145–146 insects, sequence-specific integration, 142–145 mammalian retrotransposons, 146–149 GroEL, 79 H Heat shock factor 1 (HSF1), 82 Heat shock proteins (HSPs) Hsp70 and cochaperones, 78 in Volvox embryos, 84–85 Hsp90, 77 Hsp100/Clp family, 77 Hsp60/GroEL/Cpn60, 79 small HSPs (sHSPs), 79–80 Heat stress response heat shock factor 1 (HSF1), 82 HSP70B-antisense, 84 Hsp90 inhibitors, 84 hyperphosphorylation, 83 kinase inhibitors, 84 HSPs. See Heat shock proteins I Insects, retrotransposons Drosophila, 142 target-specific LINEs, 142–143 target-specific retrotransposition systems and application, 143–145 J J-domain proteins, 78 K Kinesins 13 family, 46–47

MT minus end-directed, 45–46 MT plus end-directed, 44–45 L LIS1 protein, 49–50 Long interspersed nuclear elements (LINEs) retrotransposons, 119 LTR retrotransposons and ERVs, 121–122 M Mammalian retrotransposons activity regulators, 132–134 gene delivery vectors, 148–149 genetic markers, 146–147 mutagenesis, 147–148 retrotransposition mechanisms, 146 Microtubule plus end tracking proteins MTs dynamics, 4–5 and GTPcap, 7–8 GTP hydrolysis, in dynamics, 6–7 MT-associated proteins (MAPs), 9 organization and structure, 3–4 regulators, 8–9 tubulin, 2–3 þTIP families CAP-Gly proteins, 18–25 EB family, 10–18 LIS1 protein, 49–50 motor proteins, 43–49 serine-rich regions, 25–36 TOG and TOG-like domain proteins, 36–43 Microtubules (MTs) dynamics, 4–5 and GTPcap, 7–8 GTP hydrolysis, 6–7 MT-associated proteins (MAPs), 9 organization and structure, 3–4 regulators, 8–9 tubulin, 2–3 Molecular chaperones chloroplast components, 90 function, 90–101 cytosol/nucleus chaperone components, 81–82 function, 82–85 endoplasmic reticulum components, 89 function, 89 flagella components, 86 function, 86–89 Hsp70, 78 Hsp90, 77 Hsp100/Clp family, 77

191

Index

Hsp60/GroEL/Cpn60, 79 mitochondrion components, 101–102 function, 102 sHSPs, 79–80 Motor proteins cytoplasmic dynein, 47–49 kinesins 13 family, 46–47 MT minus end-directed, 45–46 MT plus end-directed, 44–45 MTs. See Microtubules P Penelope-like elements, 122–123 Peptidyl-prolyl cis-trans isomerases (PPIs), 89 Photosystem II (PS II) protection, 90–91, 99 Processed pseudogenes, 121, 165 R Radial spoke protein (RSP), 88–89 Repetitive transcriptomes eukaryotic retrotransposons LINE retrotransposons, 119 LTR retrotransposons and ERVs, 121–122 penelope-like elements, 122–123 processed pseudogenes, 121 SINE retrotransposons, 120 SVA elements, 120–121 genetic tools, retrotransposons DNA transposons, 140–141 Drosophila, retrotransposons, 142 gene delivery vectors, 148–149 genetic markers, 146–147 insects, target-specific LINEs, 142–143 mammalian retrotransposons, 146 mutagenesis, 147–148 randomly integrated LINEs and SINEs, fish, 145 target-specific LINEs, fish, 146 target-specific retrotransposition systems, 143–145 intracellular defense against TEs, mechanisms of AID, 124 Alu elements, ADAR editing of, 131–132 APOBEC, 123 APOBEC3 (see APOBEC3 proteins) piRNAs and PIWI proteins, 132–134 mammalian genome evolution chimeric retrogene formation, 166 flanking sequences, transduction of, 164–165 generate new REs, 162–163 processed pseudogenes, formation of, 165 REs and recombination events, 163–164 mobile DNA by host genomes domestication, REs

antisense regulators, 159–161 genomic repeats, 149–151 host cell gene transcription, 151–152 insulator elements, 161 novel polyadenylation signals, 157–158 splice sites, 152–157 transcriptional silencers, 158–159 translation regulators, 161–162 transposable elements, use genetic tools, 134–135 Sleeping Beauty transposon system, 135–136 transposons and functional genomics, 136–138 vectors, gene therapy, 138–140 types of, 118 Retrotransposons (REs). See also Eukaryotic retrotransposons enhancers, host cell gene transcription, 151–152 fish randomly integrated LINEs and SINEs, 145 target-specific LINEs, 146 host genes splice sites hairpin structure, 157 LINE elements, 155–156 LTR retrotransposons, 156 primate-specific RE Alu, 152 SEPN1, 154 SINEC_Cf, 155 7SL RNA, 153 host gene transcription, antisense regulators, 159–161 insects Drosophila, 142 target-specific LINEs, 142–143 target-specific retrotransposition systems and application, 143–145 insulator elements, 161 mammalian gene delivery vectors, 148–149 genetic markers, 146–147 mutagenesis, 147–148 retrotransposition mechanisms, 146 mammalian genome evolution formation, 162–163 recombination events, 163–164 novel polyadenylation signals, sources, 157–158 transcriptional silencers, 158–159 translation regulators, 161–162 Ribosome-associated chaperone complex (RAC), 85 S Serine-rich regions APC, 26–28 CDK5RAP2, 36 Kar9, 28–29

192

Index

Serine-rich regions (cont.) melanophilin, 34 neuron navigators, 33–34 p140Cap, 32–33 Rho-type guanine nucleotide exchange factor (RhoGEF2), 34–35 spectraplakins, 29–30 STIM1, 31–32 TIP150, 35–36 Short interspersed nuclear elements (SINEs) retrotransposons, 120 SINE–variable number of tandem repeats–Alu-like (SVA) elements, 120–121 Spectraplakin, 29–30 STIM1, 31–32 T TEs. See Transposable elements (TEs) Thylakoid biogenesis, 99 Transcriptional promoters, 149–151 Transcription factorbinding sites (TFBS), 149–150 Transcriptomes. See Repetitive transcriptomes Transduction, flanking sequences, 164–165 Transposable elements (TEs). See also Repetitive transcriptomes AID, 124 Alu elements, ADAR editing of, 131–132 APOBEC3 proteins A3 expression profile, human tissues, 130–131 Alu retrotransposition inhibitors, deaminases, 129–130 L1 retrotransposition inhibitors, deaminases, 128–129 LTR retrotransposons inhibitors, deaminases, 125, 127–128 members of, 124–126 DNA transposons, genetic tools ITRs, 134–135 Sleeping Beauty transposon system, 135–136

transposons and functional genomics, 136–138 vectors, gene therapy, 138–140 epigenetic modifications, 123 piRNAs and PIWI proteins, 132–134 retrotransposons, genetic tools (see also Retrotransposons) DNA transposons, 140–141 fish, retrotransposons, 145–146 insects, sequence-specific integration, 142–145 mammalian retrotransposons, 146–149 Tumor overexpressed gene (TOG) CLASPs, 40–43 XMAP215/Dis1 family, 37–40 V Vesicle-inducing protein in plastids (VIPP1) Arabidopsis vipp1 mutant, 99 CDJ2, 98 HSP70B-CDJ2-CGE1 chaperone system, 98 phage shock protein A (PspA), 97 rod-shaped supercomplexes, 98 thylakoid biogenesis, 99 Volvox. See also Molecular chaperones chloroplast components, 90 function, 90–101 cytosol/nucleus chaperone components, 81–82 function, 82–85 endoplasmic reticulum components, 89 function, 89 flagella components, 86 function, 86–89 mitochondrion components, 101–102 function, 102

A

B 1, 4, 6, 9, 10 5 A ID

xin Ne A OD

A B

Head

5

Central sheath

C1

16 16

Stalk

RS

23 23 2

Andre´ Nordhues et al., Figure 2.1 Half cross section of a flagellar axoneme. (a) A flagellum consists of nine peripheral doublet microtubules (A and B) and two central microtubules (C1 and C2). This highly conserved “9þ2” structure forms the core of the axoneme, or flagellar cytoskeleton. Each A microtubule anchors three outer dynein arms (ODAs) and two inner dynein arms (IDAs), which are responsible for flagellar locomotion. The elastic protein, nexin, maintains this organization during the motion of the doublet microtubules. RS, radial spoke. (b) The T-shaped RSs are associated with A microtubules and transiently interact with the central sheath, which protects the central tubules. It is postulated that the RSs are involved in controlling flagellar motility (Yang et al., 2005). The RSs consist of 23 proteins (radial spoke protein (RSP) 1–RSP23) (Yang et al., 2001, 2005). RSP1, 4, 6, 9, and 10 form the spoke head, and the rest of the RSPs are integrated in the spoke stalk (Yang et al., 2005). Homodimeric J-domain protein RSP16 may interact with dimeric RSP2 and/or RSP23 in the stalk (Yang et al., 2008).

Chloroplast 1 HEP2

P2

HE

70 ATP

trx

2

90

90

GSH

3

90

90

VIPP1 70 ATP

VIPP1

J

J

4 ADP

ATP 90

ATP 90

ATP 5c

J J E1

70 ADP

E1

6 VIPP1

5a VIPP1

5b

Signal transduction

Andre´ Nordhues et al., Figure 2.2 The components and substrates of the chloroplast HSP70B/HSP90C chaperone system in Chlamydomonas. The major Hsp70 in the chloroplast of Chlamydomonas is HSP70B (light blue) (Drzymalla et al., 1996). It requires HEP2 (HSP70 Escort Protein 2, blue) to attain the functional state (Willmund et al., 2008b). Most likely, HEP2 acts by folding HSP70B to the native state following cleavage of its transit peptide after import (1). HSP70B was identified as a thioredoxin (trx) substrate (Lemaire et al., 2004) and may become glutathionylated (GS) (2) (Michelet et al., 2008). HSP70B appears to form constitutive complexes with dimeric HSP90C (purple), the chloroplast HSP90 homolog of Chlamydomonas (3) (Willmund and Schroda, 2005; Willmund et al., 2008a). In ATP-driven cycles, Hsp70B, presumably supported by HSP90C, induces conformational changes into substrate proteins that are important e.g. protein folding or assembly/disassembly of protein complexes. Substrates are delivered to HSP70B by one of its CDJ (Chloroplast DnaJ-like protein, green) cochaperones (4). CDJ1 is a functional homolog of bacterial DnaJ and thus likely delivers unfolded substrates (gray) for general folding to HSP70B/ HSP90C. Chloroplast proteins involved in (light) signal transduction may depend on chaperoning by HSP70B/HSP90C (5a) (Cao et al., 2003; Schroda and M€ uhlhaus, 2009). CDJ2 is a specialized J-domain protein that delivers the various assembly states of VIPP1 (Vesicle-Inducing Protein in Plastids, light red) to HSP70B and HSP90C. The latter may catalyze the assembly of VIPP1 monomers/dimers to rings and the disassembly of rods and rings (5b) (Liu et al., 2005, 2007). CDJ3–5 contain redox-active Fe–S clusters and CDJ3 is in complex with RNA, presumably to recruit HSP70B for mediating redox-dependent remodeling of translation initiation complexes (5c) (Dorn et al., 2010). CGE1 (Chloroplast GrpE homolog 1, orange) binds to HSP70B in the ADP-state and catalyzes the exchange of ADP by ATP (6) (Schroda et al., 2001b). CGE1 dimers form via coiled-coil interactions of their N-terminal a-helices (Willmund et al., 2007).

b-Sheet domains

Four-helix bundles

Paired a-helices

E.coli GrpE Coiledcoil

C. reinhardtii CGE1

Andre´ Nordhues et al., Figure 2.3 Structural models of dimeric CGE1 and GrpE. The GrpE model is based on a crystal structure of the E. coli GrpE dimer bound to its HSP70 chaperone partner DnaK (PDB entry 1DKG.pdb) (Harrison et al., 1997). Since the N-terminal 33 amino acids of GrpE were unstructured and had to be removed for crystallization, they are not shown here; because of their low resolution in the crystal structure, loops connecting helices of the four-helix bundle in GrpE are also not shown. The CGE1 model was generated by homology modeling using structures of E. coli GrpE and tropomyosin (Whitby and Phillips, 2000; Willmund et al., 2007). Due to ambiguous predictions of the secondary structure formed by the residues between Pro13 and Pro21, this region was drawn as a random coil. Also the positions of the N-terminal a-helices predicted between Ala1 and Ala12 are drawn arbitrarily.

Zn2+ S.cerevisiae Hep1p

Andre´ Nordhues et al., Figure 2.4 Structural model of yeast mitochondrial Hep1p. The Hep1p model is based on an NMR structure of Hep1p (PDB entry 2E2Z.pdb) (Momose et al., 2007).

• Cell culture – Generating stable lines

• Transgenesis – Active in all vertebrate species

• Insertional mutagenesis – Zebrafish – Xenopus – Mouse, rat

• Gene therapy – New, nonviral delivery method

Gerald G. Schumann et al., Figure 3.4 Broad applicability of Sleeping Beauty in vertebrate genetics.