Ribonucleases (Nucleic Acids and Molecular Biology 26)

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Nucleic Acids and Molecular Biology

26

Series Editor Janusz M. Bujnicki

.

Allen W. Nicholson (Ed.)

Ribonucleases

Editor Allen W. Nicholson Temple University Department of Biology 1900 North 12th Street Philadelphia, PA 19122 USA [email protected] Series Editor Janusz M. Bujnicki International Institute of Molecular and Cell Biology Laboratory of Bioinformatics and Protein Engineering Trojdena 4 02-109 Warsaw Poland

ISSN 0933-1891 e-ISSN 1869-2486 ISBN 978-3-642-21077-8 e-ISBN 978-3-642-21078-5 DOI 10.1007/978-3-642-21078-5 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: 2011934623 # Springer-Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover design: deblik, Berlin, Germany Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

Ribonucleases continue to attract much interest and investigation in the basic and translational science arenas. Our present understanding of ribonuclease structures, mechanisms, and functions emerged from a myriad of pioneering investigations that employed (as well as led to the development of) diverse experimental approaches. These studies have shed light on the fundamental aspects of biological catalysis and protein folding and ribonuclease function in post-transcriptional regulatory pathways. Indeed, multiple volumes would be needed to provide a comprehensive coverage of ribonucleases. It is instead the intent of this single volume to present a focused collection of reviews on the major groups of ribonucleases, and how their structures and mechanisms relate to biological function. The first three chapters by D’Alessio, Rosenberg, Vilanova, and coauthors focus on the fascinating family of vertebrate secreted ribonucleases, within which pancreatic ribonuclease A has served as the founding member. The extraordinary functional and evolutionary diversity of these enzymes is discussed along with their promise as anticancer agents. The chapters contributed by MacIntosh, Ivanov, Anderson, Meyers, and coauthors focus on the ribonuclease T2 family enzymes. Here, only recently has there been an appreciation gained of the central involvement of T2 family members in stress responses, host defense, and strategies of viral infection. The chapter by Tong and coauthors examines the structures and functions of 50 –30 exoribonucleases, and the chapter by Arraiano and coauthors provides a comprehensive review of the diverse group of 30 –50 exoribonucleases. The multisubunit RNA exosome, with its 30 –50 exonuclease (and endonuclease) activity, is examined by Hopfner and Hartung, with a special focus on how specificity and regulation can be achieved in an otherwise nonselective manner of RNA breakdown. Condon and Gilet address the mechanistically and functionally intriguing metallob-lactamase family enzymes and their roles in processing tRNAs, mRNAs, and snRNAs. The structure, mechanism, and diverse functions of the double-strandspecific ribonuclease III is reviewed by Nicholson, and Hollis and Shaban next discuss the structures and functions of the ribonucleases H that cleave the RNA strand in RNA–DNA duplexes. Krasilnikov provides an in-depth examination of

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the ribonucleoprotein ribonucleases P and MRP, their central cellular roles in tRNA and rRNA processing, and the functions of the RNA and protein subunits in the catalytic mechanism. The chapter by Lo¨nnberg addresses the inherent reactivity of RNA toward metal ions, and summarizes studies of small molecule ribonuclease mimics that exhibit diverse structures. Finally, Scheraga reviews pioneering experimental studies on protein folding that have employed pancreatic ribonuclease A as the primary model. What is evident from these chapters is the integral involvement of ribonucleases in a broad array of physiological processes, and that the simple act of cutting an RNA molecule, either internally or by removal of one or more nucleotides from either end, has profound effects on cell phenotype. Finally, detailed knowledge of ribonuclease structure, catalytic mechanism, and interacting partners are spurring new approaches to the treatment of disease. It is hoped that this volume will inform and stimulate further investigations of ribonucleases and their involvement in cellular pathways.

Acknowledgements

I would like to thank Janusz Bujnicki for the invitation to edit a volume on ribonucleases as part of the Springer Nucleic Acids and Molecular Biology series, and Ursula Gramm of Springer for her assistance with the preparation of the volume.

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Contents

1

The Superfamily of Vertebrate-Secreted Ribonucleases . . . . . . . . . . . . . . . 1 Giuseppe D’Alessio

2

Vertebrate Secretory (RNase A) Ribonucleases and Host Defense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 Helene F. Rosenberg

3

Antitumor Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Marc Ribo´, Antoni Benito, and Maria Vilanova

4

RNase T2 Family: Enzymatic Properties, Functional Diversity, and Evolution of Ancient Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Gustavo C. MacIntosh

5

Stress-Induced Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Pavel Ivanov and Paul Anderson

6

Viral RNase Involvement in Strategies of Infection . . . . . . . . . . . . . . . . . . 135 Gregor Meyers, Tillmann Ru¨menapf, and John Ziebuhr

7

50 -30 Exoribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Jeong Ho Chang, Song Xiang, and Liang Tong

8

Structure and Degradation Mechanisms of 30 to 50 Exoribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Rute G. Matos, Vaˆnia Pobre, Filipa P. Reis, Michal Malecki, Jose´ M. Andrade, and Cecı´lia M. Arraiano

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The RNA Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223 Karl-Peter Hopfner and Sophia Hartung

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The Metallo-b-Lactamase Family of Ribonucleases . . . . . . . . . . . . . . . . . 245 Ciarań Condon and Laetitia Gilet

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Ribonuclease III and the Role of Double-Stranded RNA Processing in Bacterial Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 Allen W. Nicholson

12

Structure and Function of RNase H Enzymes . . . . . . . . . . . . . . . . . . . . . . . . 299 Thomas Hollis and Nadine M. Shaban

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Ribonucleoprotein Ribonucleases P and MRP . . . . . . . . . . . . . . . . . . . . . . . 319 Andrey S. Krasilnikov

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Nonenzymatic and Metal-Ion-Dependent RNA Cleavage, and RNase Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 343 Harri Lo¨nnberg

15

Ribonucleases as Models for Understanding Protein Folding . . . . . . . 367 Harold A. Scheraga

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399

Contributors

Paul Anderson Division of Rheumatology, Immunology and Allergy, Brigham and Women’s Hospital, Boston, MA, USA; Department of Medicine, Harvard Medical School, Boston, MA, USA Jose´ M. Andrade Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Cecı´lia M. Arraiano Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Antoni Benito Laboratori d’Enginyeria de Proteı¨nes, Departament de Biologia, Facultat de Cie`ncies, Universitat de Girona, Campus de Montilivi, Maria Aure`lia Capmany, Girona, Spain; Institut d’Investigacio´ Biome`dica de Girona Josep Trueta (IdIBGi), Girona, Spain Jeong Ho Chang Department of Biological Sciences, Columbia University, New York, NY, USA Ciarań Condon CNRS UPR 9073 (affiliated with Universite´ de Paris 7 - Denis Diderot), Institut de Biologie Physico-Chimique, Paris, France Giuseppe D’Alessio Department of Structural and Functional Biology, University of Naples Federico II, Via Cintia, Napoli, Italy Laetitia Gilet CNRS UPR 9073 (affiliated with Universite´ de Paris 7 - Denis Diderot), Institut de Biologie Physico-Chimique, Paris, France Sophia Hartung Life Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA

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Contributors

Thomas Hollis Department of Biochemistry, Center for Structural Biology, Wake Forest University School of Medicine, Winston-Salem, NC, USA Karl-Peter Hopfner Department of Biochemistry, Gene Center, Munich, Bavaria, Germany; Center for Integrated Protein Sciences, Ludwig-MaximiliansUniversity, Munich, Bavaria, Germany Pavel Ivanov Division of Rheumatology, Immunology and Allergy, Brigham and Women’s Hospital, Boston, MA, USA; Department of Medicine, Harvard Medical School, Boston, MA, USA Andrey S. Krasilnikov Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park, PA, USA Harri Lo¨nnberg Department of Chemistry, University of Turku, Turku, Finland Gustavo C. MacIntosh Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, IA, USA Michal Malecki Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Rute G. Matos Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Gregor Meyers Institut fu¨r Immunologie, Friedrich-Loeffler-Institut, Tu¨bingen, Germany Allen W. Nicholson Department of Biology, Temple University, Philadelphia, PA, USA Vaˆnia Pobre Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Filipa P. Reis Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Marc Ribo´ Laboratori d’Enginyeria de Proteı¨nes, Departament de Biologia, Facultat de Cie`ncies, Universitat de Girona, Campus de Montilivi, Maria Aure`lia Capmany, Girona, Spain; Institut d’Investigacio´ Biome`dica de Girona Josep Trueta (IdIBGi), Girona, Spain Helene F. Rosenberg Laboratory of Allergic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA

Contributors

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Tillmann Ru¨menapf Institut fu¨r Virologie, Fachbereich Veterina¨rmedizin, JustusLiebig-Universita¨t Giessen, Giessen, Germany Harold A. Scheraga Baker Laboratory of Chemistry and Chemical Biology, Cornell University, Ithaca, NY, USA Nadine M. Shaban Department of Biochemistry, Center for Structural Biology, Wake Forest University School of Medicine, Winston-Salem, NC, USA Liang Tong Department of Biological Sciences, Columbia University, New York, NY, USA Maria Vilanova Laboratori d’Enginyeria de Proteı¨nes, Departament de Biologia, Facultat de Cie`ncies, Universitat de Girona, Campus de Montilivi, Maria Aure`lia Capmany, Girona, Spain; Institut d’Investigacio´ Biome`dica de Girona Josep Trueta (IdIBGi), Girona, Spain Song Xiang Department of Biological Sciences, Columbia University, New York, NY, USA; Key Laboratory of Nutrition and Metabolism, Institute for Nutritional Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, P.R. China John Ziebuhr Institut fu¨r medizinische Virologie, Justus-Liebig-Universita¨t Giessen, Giessen, Germany

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Chapter 1

The Superfamily of Vertebrate-Secreted Ribonucleases Giuseppe D’Alessio

Contents 1.1 Premise . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Mammalian RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.1 Human RNases (Canonical, Noncanonical) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.2 Rodent RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.3 Bovine RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3 Non-mammalian RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.1 Bird RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.2 Reptile RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.3 Amphibian RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.4 Fish RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4 Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 10 11 19 19 19 20 21 23 24

Abstract Recent investigations on vertebrate proteomes have revealed the presence of a single vertebrate-specific enzyme group: that of the RNases homologous to RNase A, the historical RNase archetype studied for more than a century. The genes encoding these RNases are all phylogenetically linked, and the gene products are all secreted proteins, thus forming an impressively large superfamily of vertebrate-secreted RNases, formerly called “RNase A superfamily.” The vertebrate-secreted RNases display surprisingly different physiological functions, other than that of ribonucleolytic enzymes, including angiogenesis, host defense, immunosuppression, biogenesis of ribosomes, and stress response. Some of the RNases have antitumor activity, as they are capable of selectively killing malignant cells, and have inspired an intensely pursued research line of translational value.

G. D’Alessio Department of Structural and Functional Biology, University of Naples Federico II, Via Cintia, 80126 Napoli, Italy e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_1, # Springer-Verlag Berlin Heidelberg 2011

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A particular attention has been given in this chapter to the family of mammalian RNases, especially to RNase 5 (angiogenin) and microbicidal RNases 2 and 3, to the RNase inhibitor, and the recently investigated family of fish RNases.

1.1

Premise

Among the great successes of biochemistry in the twentieth century were: the determination of the first complete amino acid composition of a protein; the first complete amino acid sequence of a protein much larger than the insulin peptides; the recognition that the three-dimensional structure of a protein is determined by its amino acid sequence; and the complete chemical synthesis of a protein enzyme. These results, which not only changed our understanding of protein chemistry but also significantly contributed to set the foundations of modern biology, all employed the same experimental model: an enzyme protein from bovine pancreas, a ribonuclease, RNase A. The authors of these achievements were all awarded Nobel Prizes: William Stein and Stanford Moore, Christian Anfinsen, and Bruce Merrifield. Subsequently, RNase A became a convenient model for such innovative methodologies as protein X-ray crystallography, NMR, and calorimetry. Phylogenetic studies soon revealed (Beintema et al. 1997) that many other RNases present in a variety of organisms, from amphibians to reptiles, birds, mammals, were structurally and functionally close to RNase A. An RNase superfamily was constructed, also called “the RNase A superfamily” from the historical, first described, archetypical ribonuclease. Recently, homologous RNases have also been found in fishes, thus allowing a vertebrate superfamily to be defined. Furthermore, the vertebrate RNases of this superfamily are all secreted, so it may be convenient, and appropriate, to name the superfamily as the Vertebrate-Secreted-RNase-Superfamily. Interestingly, the sequencing of the human genome has unveiled an intriguing aspect. When vertebrate proteomes were explored, only a single vertebrate-specific enzyme group was found, that of RNases (Lander et al. 2001). Apparently, after the divergence of the vertebrate subphylum, about 500 million years ago, one or perhaps two new DNA sequences emerged (Cho and Zhang 2007), encoding a protein(s) absent in invertebrates, that rapidly evolved into many orthologs, following vertebrate speciation, to yield various numbers of paralogs within each evolving species. There are several features that exclusively define vertebrate-secreted RNases: (1) They contain in their amino acid sequence a short stretch of residues (CKxxNTF) known as the vertebrate RNase “signature,” at position 40–46 (the numbering of RNase A is used here and elsewhere in this chapter). (2) The reading frame of each protein is contained in a single exon. (3) The catalytic activity is carried out with the essential cooperation of His-12 and -119 and Lys-41. (4) Cleavage of the RNA P-diester bonds is initiated by the 30 -OH of a pyrimidine

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The Superfamily of Vertebrate-Secreted Ribonucleases

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Fig. 1.1 Ribbon diagram of the superimposition of the structure of ZF-RNase 5 from Danio rerio or zebrafish (green, PDB code 3LJE) and that of RNase A from Bos taurus (pink, PDB code 1KF3)

nucleotide, which through transphosphorolysis leads to the formation of a 20 :30 -cyclic-phosphodiester, which (with exceptions) is subsequently hydrolyzed into a 30 -phosphate (Cuchillo and Vilanova 1997). Vertebrate-secreted RNases are all structurally homologous, even when low identity scores are calculated between primary structures. They have an a/b structure arranged in a kidney shape by three alpha-helix stretches and a four-stranded antiparallel b-sheet. Figure 1.1 illustrates the 3D structures of archetypical RNase A, and that of a phylogenetically distant fish RNase, RNase-5 from zebrafish (Pizzo et al. 2010). Although the identity score is only 21%, the structural homology between the two proteins is strikingly apparent.

1.2 1.2.1

Mammalian RNases Human RNases (Canonical, Noncanonical)

The complete identification of human RNase genes was obtained only few years ago when human chromosome 14q11.2 could be read and analyzed (Cho et al. 2005). The chromosome was found to contain all human RNases. However, only eight of them (RNases 1–8) are true RNases, called “canonical RNases”; the remaining “noncanonical” five, numbered 9–13 (Cho et al. 2005), lack one or more residues of the catalytic triad (see above); thus, they may not catalyze RNA degradation. As 3D structures of these proteins are not available, and no RNase assays have been performed or published, the unlikely possibility cannot be excluded that the missing residues are replaced by residues identical or conservative with respect to the characteristic RNase catalytic triad, even located at different sequence positions.

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Most differences between noncanonical and canonical RNases are found at the C-terminal region of the proteins, but RNases 9 and 10 contain an insertion of 40–50 residues in their signal peptides (Castella et al. 2004; Penttinen et al. 2003). As for their function, RNases 9 and 10 appear to have a role in the reproductive tract, at least in some species, as they are expressed in mouse and porcine epididymis (Castella et al. 2004; Penttinen et al. 2003). A compact review of human canonical RNases has been recently published (Sorrentino 2010).

1.2.1.1

Human RNase 1

Human RNase 1 was originally studied in pancreas (Beintema et al. 1984; Seno et al. 1994) so that it is also termed HP-RNase or hPR (for Human Pancreatic RNase). However, it may not be a digestive enzyme, as the single gene encoding the RNase (Breukelman et al. 1993) is expressed not only in pancreas, but in a variety of tissues and organs, and especially in endothelial cells (Landre et al. 2002). Thus, it may be proposed instead that the enzyme is involved in the control of the homeostasis of extracellular RNAs. This hypothesis is supported by the following: (1) RNase 1 is actively and directly secreted by endothelial cells into blood vessels, and (2) it displays a powerful activity both on single- and double-stranded RNA (Libonati and Sorrentino 2001), as well as on RNA in RNA:DNA hybrids (Potenza et al. 2006).

1.2.1.2

Human RNases 2 and 3, and the Fortunes of Microbicidal RNases

Human RNase 2 is also called EDN (Eosinophils-Derived Neurotoxin) because it was first isolated from eosinophils with an assay for neurotoxicity (Snyder and Gleich 1997). Also human RNase 3 was first isolated from eosinophils and labeled ECP (Eosinophils Cationic Protein) for its high content of cationic residues. ECP/ RNase 3, however, is produced only in eosinophils, whereas the EDN/RNase 2 gene is expressed in a variety of organs and tissues (Beintema et al. 1988; Mizuta et al. 1990; Sorrentino et al. 1988). The catalytic properties of RNases 2 and 3 are unusual for vertebrate RNases. The two RNases do not hydrolyze the 20 :30 -cyclic-phosphodiesters produced in the first transphosphorolytic event of catalysis. Moreover, their base preference at the 30 side of the cleaved P-diester bond, like for human RNase 4 (see below), is for uracil rather than cytosine (Sorrentino and Libonati 1994). RNases 2 and 3 evolved through gene duplication and share about 70% of their amino acid sequence. Both have a pronounced neurotoxic activity and an antiviral activity that are dependent on their RNase activity (Domachowske et al. 1998; Rosenberg and Domachowske 2001; Sorrentino et al. 1992). EDN/RNase 2, but not ECP/RNase 3, has been found to activate human dendritic cells in response to pathogen stimulation (Yang et al. 2003, 2004). As this event

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results in the production of an array of cytokines and growth factors from the immune system, EDN has been classified an alarmin (Oppenheim and Yang 2005). ECP/RNase 3 is secreted by eosinophils activated by inflammation, and it was first identified as an antiparasitic agent (Hamann et al. 1990), then recognized as a powerful bactericidal protein (Lehrer et al. 1989), and later as an antiviral agent (Domachowske et al. 1998:532). These activities are not related to the RNase activity of the protein (Rosenberg 1995). M.V. Nogue´s, E. Boix and coworkers (Carreras et al. 2003; Torrent et al. 2007) have determined that the bactericidal activity of ECP is based on its ability to destabilize the bacterial membrane, an action in turn based on its high content of cationic and hydrophobic residues. More recently, ECP has been found to be cytotoxic also for eukaryotic cells (Navarro et al. 2008). It does not enter the cells, but it aggregates on the cell surface, thus affecting membrane permeability. These interactions with the membrane bring about in the affected cells a series of impressive morphological and biochemical changes, such as chromatin condensation, membrane reversion, production of reactive oxygen species, and activation of caspase-3-like activity through eventual cell death. It has been confirmed that the RNase catalytic activity of ECP plays no role in these processes. Two ECP polymorphic variants coexist in eosinophils, with Arg or Thr at position 97 of the protein sequence. Only the variant with Arg is cytotoxic for lung carcinoma cells (Rubin et al. 2009). Interestingly, the aggregation of ECP into amyloid-like fibrils has been recently reported (Torrent et al. 2010:745). Also, it was found that the isolated N-terminal peptide segment of the protein (Arg1-Asn19) promotes the formation of a fibrillar network, even more extensive than that induced by the protein itself. Fibrils are formed only at acidic pH, but protein hydrophobicity is also important. The most intriguing and elusive topic in a discussion on RNases 2 and 3 is the significance of the functionality of their ribonucleolytic active site for the various actions so far described for these proteins. As indicated above, the ribonucleolytic activity may be necessary (e.g., for the antiviral activity of EDN), or dispensable (e.g., for the bactericidal activity of ECP). In the former case, it is reasonable to imagine the damaging effects of an RNase on any cytosolic RNA, once the enzyme reaches the cell cytosol. In this case, the mechanism would be likely understood from the identification of the target RNA substrate. Here, research on the involvement of RNases with non-coding and interfering RNAs may provide new, unexpected clues. Different and more intriguing is the latter case, in which the ribonucleolytic activity appears to be redundant, as the protein does not need it to perform its function. Here the crucial question is: Why in such a long evolutionary timeframe was the global 3D structure of a vertebrate RNase, and a precisely poised RNase catalytic site preserved with no apparent advantage? Analysis of data from research on a bactericidal bird RNase (see also Sect. 1.3.2) has led Rosenberg and coauthors (Nitto et al. 2006) to consider the possibility that the RNase gene/protein structure is, for several RNases, merely a convenient source of biochemical, peptide material with toxic abilities. The hypothesis can be

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expanded and read as follows: These RNases are microbicidal because they comprise in their sequences the sequence(s) of a microbicidal peptide(s). When the RNase reaches a cell membrane, or it enters a cell, it is fragmented by a membrane and/or cytosolic protease. The resultant free peptides would exert their toxic activities (bactericidal, antiparasitic, antiviral, cytotoxic) and provide the cells with valuable host-defense agents. This scenario is especially convincing for RNases that conserve their bactericidal activity not only when the RNase activity is abolished, but also when they are completely and irreversibly denatured. There are several cases: (1) five zebrafish RNases, conserving their bactericidal activity after full denaturation (Pizzo et al. 2010), and (2) an active peptide contained within the sequence of bactericidal RNase 3 (Garcia-Mayoral et al. 2010); two Atlantic salmon RNases (Pizzo et al. 2008), and historical lysozyme as well (During et al. 1999; Ibrahim et al. 2001). The bactericidal activity of a zebrafish RNase (ZF-RNase-3) has been found to be due to a peptide fragment excised from the RNase by a membrane protease from the bacterium itself (Zanfardino et al. 2010). It is widely recognized that cationic/hydrophobic antimicrobial peptides have an important role in the host innate defense mechanisms against invading microorganisms (Boix and Nogues 2007). However, it is difficult to envisage a biomolecule that has undergone any length of evolutionary process without possessing a properly folded structure. It is possible instead that evolution has indeed taken advantage of an RNase, perhaps both structurally and catalytically, but its physiological role is still unknown today. Obviously, the artificially unfolded RNases described above have no relationships with the intrinsically destructured proteins that do not fold spontaneously into ordered and stable structures (Wright and Dyson 1999, 2009). These proteins, with key roles in the lives of cells, do acquire their structures when they interact and complex with their physiological partner(s) to perform their biological functions.

1.2.1.3

Human RNase 4

First isolated (Shapiro et al. 1986) from culture medium conditioned by an adenocarcinoma cell line, its characterization has been carried out in several laboratories (Seno et al. 1995; Vicentini et al. 1994, 1996; Zhou and Strydom 1993). The enzyme prefers uracil to cytosine as the base at the 30 side of the cleaved P-diester bond, an unusual feature, as discussed above, for a vertebrate RNase. Surprisingly, the uridine-specific preference of the enzyme was found to be readily changed to cytidine-specific when the Asp residue at position 80 was replaced by Ala (Hofsteenge et al. 1998). The finding was intriguing because Asp-80 is a highly conserved residue in all orthologous sequences of RNase 1 and several other RNases. The possible conclusion drawn by the authors was that a residue, although conserved in evolution, may not have the same structural/functional role in all homologous enzymes.

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An investigation (Rosenberg and Dyer 1995) on a genomic fragment from chromosome 14, where human RNase genes are located, has indicated that the mRNA encoding RNase 4 is much larger than those encoding closely related, homologous RNases. Surprisingly for a vertebrate RNase, two transcripts are identified in RNA from some organs (liver, kidney, pancreas). Furthermore, it has been established that RNase 4 is expressed in all tissues analyzed. However, the significance of these findings and the physiological role of RNase 4 remain to be investigated.

1.2.1.4

Human RNase 5

Human RNase 5 is most frequently known as “angiogenin,” because it is an angiogenic RNase first isolated from the conditioned medium of HT-29 colon adenocarcinoma cells (Fett et al. 1985). Extensive reviews on angiogenin are available (Gao and Xu 2008; Riordan 1997; Strydom 1998). The angiogenic activity of human angiogenin (ANG) depends on its activity as a ribonuclease, which is very low: With certain assays, it can be even one millionfold lower than that of RNase A, the archetype vertebrate RNase. The molecular basis for this low activity has been proposed to depend on the presence of a Gln at position 117 (substituting for Ala of RNase A), which hinders the access of the substrate to the pyrimidine binding site of the enzyme (Acharya et al. 1994; Russo et al. 1994). A recent kinetic analysis carried out with a rational series of substrates (Leland et al. 2002) has instead suggested that the low RNase activity of ANG is the result of a specific orientation of ANG catalytic residues, not favorable for cleavage of RNA. ANG is a very versatile bio-effector, capable of exerting several biological actions, besides the angiogenic activity. Its interactions with the endothelial cell surface can be ascribed to a specific stretch of residues (positions 60–68 of the amino acid chain), as well as to the ability of ANG to bind to a smooth muscle type a-actin (Hu et al. 1991, 1993), and a receptor, although the latter has not yet been described at the molecular level. Once in the cell, ANG is translocated to the nucleoli, where it eventually accumulates (Hu et al. 2000; Hu 1998; Moroianu and Riordan 1994b). This event requires the presence of a typical nuclear localization signal contained between Arg3I and Leu-35 in the ANG sequence (Moroianu and Riordan 1994a). In addition, or alternatively, the translocation of ANG to the nucleus may occur through passive diffusion, given the small size of the RNase (Lixin et al. 2001). Once ANG has reached the nucleoli, the site of ribosomal RNA (rRNA) transcription, ANG binds to the promoter of ribosomal DNA (rDNA), and rRNA transcription is stimulated (Kishimoto et al. 2005; Xu et al. 2002). The irony is that an RNA-degrading enzyme is implicated in RNA synthesis. In fact, the stimulation of rRNA biogenesis is essential for ANG angiogenic activity, and angiogenic factors, such as bFGF and VEGF, stimulate nuclear translocation of endogenous ANG (Hirukawa et al. 2005; Kishimoto et al. 2005). Silencing ANG

8

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expression in endothelial cells inhibits bFGF- and VEGF-induced cell proliferation and leads to a decrease of rRNA transcription, an effect reversed by addition of exogenous ANG. However, the way of ANG to angiogenesis described above poses a problem: How can ANG translocate to the nucleus moving through the cytosol in which there is a high concentration of RI, the RNase Inhibitor that binds ANG with exceedingly high affinity. Possibly, ANG travels in the cytosol fused to a carrier, which preserves, and obscures, the enzyme. However this carrier, if it exists, has not been found. The RI/ANG interrelationship has been studied instead under para-physiological conditions. Under environmental stress, the cell economy requires the arrest of protein synthesis, as translation is energetically costly. It has been found that the arrest is determined by ANG, which under stress blocks translation through the inactivation of tRNAs. The inactivation is performed through cleavage by ANG (with its ribonucleolytic ability) of P-diester bonds at the anticodon loops of tRNAs (Emara et al. 2010; Fu et al. 2009; Yamasaki et al. 2009). Under normal conditions, ANG cannot exert this function because it is bound and neutralized by RI, the RNase inhibitor. We can surmise how under stress RI dissociates from the complex and ANG is free to act. This certainly happens during oxidative stress, when RI is damaged and knocked down (Blazquez et al. 1996). In conclusion, ANG promotes translation through rRNA biosynthesis, although under stress conditions, it blocks translation. But there is no inconsistency between these events, as recent results (E. Pizzo, A. Furia and G. D’Alessio, unpublished data) have shown that under stress conditions, no ANG is detectable in nucleoli. Thus, while ANG arrests translation in the cytoplasm under stress, it simultaneously and coherently stops its nucleolar-based activation of rRNA synthesis. The significance of angiogenin has been considered for a multiplicity of diseases which could be related to angiogenesis (see (Gao and Xu 2008)). Here we shall review the role(s) of ANG in two most relevant diseases: cancer and amyotrophic lateral sclerosis (ALS). ALS is a disease that leads to a progressive degeneration of motor neurons. About 10% of the analyzed cases are familial, caused by alterations of a number of genes, including the ANG gene (Greenway et al. 2006; Wu et al. 2007). Furthermore, the ANG gene has been found to be strongly expressed in mouse CNS during development (Subramanian and Feng 2007), and in adult human spinal cord (Wu et al. 2007). Angiogenin variants present in ALS patients have been characterized and shown to affect neurite extension and pathfinding, and survival of motor neurons (Crabtree et al. 2007). Furthermore, they display poor RNase activity and/or impaired nuclear translocation (Wu et al. 2007). When ANG is administered to cultured motoneurons, the cells are protected from hypoxic injury (Sebastia et al. 2009); on the other hand, silencing of ANG leads to an increase in hypoxia-induced cell death. As for the involvement of ANG with cancer, its first role is that of providing tumors with the support needed for growth. Only a fraction of nascent tumors ever develop to detectable tumors unless they are provided with essential oxygen and

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nutrients, carried by blood. Vascularization of a micro-tumor is induced by tumor angiogenesis factors, and angiogenin is one of these factors. Inhibition of angiogenesis has formed the rational basis for anticancer therapy with anti-angiogenic peptides and proteins (Boehm et al. 1997; O’Reilly et al. 1996). An additional approach to cancer therapy has been based on the performance of ANG in the presence of neomycin, an antibiotic that inhibits the translocation of ANG to nuclei (Hu 1998). It has been found that neomycin has also direct, inhibitory effects on cell proliferation (Tsuji et al. 2005).

1.2.1.5

Human RNase 6

First labeled RNase k6 as the sixth human RNase, with “k” denoting its relationship to the orthologous RNase k2 from bovine kidney, this RNase is expressed in most tissues, but not in eosinophils (Rosenberg and Dyer 1996), although its amino acid sequence is most closely related to those of RNases 2 and 3 (see Sect. 1.2.1.2). But, likely, also RNase 6 has a role in host defense, as it has been found in neutrophils and monocytes.

1.2.1.6

Human RNase 7

RNase 7 was found while searching for antimicrobial proteins of human skin (Harder and Schroder 2002), during screening the human genome (Zhang et al. 2003). This RNase is expressed not only in skin, but in several tissues, especially in liver. It is endowed with a bactericidal activity against several pathogenic microorganisms, both Gram-negative and Gram-positive. It has been proposed (Huang et al. 2007) that a key role in the antibacterial mechanism of RNase 7 is played by Lys residues from flexible N- and C-terminal cationic clusters. Its bactericidal activity would be due to its ability to permeate the bacterial membrane, whereas its ribonucleolytic activity has no role. In contrast with RNase 3 (or ECP, see Sect. 1.2.1.2), RNase 7 does not initiate its toxic action through agglutination of the bacteria (Torrent et al. 2010). This suggests that beside disruption of the bacterial plasma membrane, a key factor in the mechanism are interactions of the protein with the bacterial cell wall. It has been found that the bactericidal activity of RNase 7 on P. aeruginosa is based first on its binding to a cell wall lipopolysaccharide, then to the oligomeric membrane lipoprotein OprI (Lin et al. 2010). RNase 7 is not only constitutively expressed in keratinocytes, but also induced by pro-inflammatory cytokines, such as interleukin-1b, interferon-g, and bacterial challenge (Harder and Schroder 2002). Abtin and coworkers (Abtin et al. 2009) recently reported that RNase 7, RNase 5 (angiogenin), and RI, the RNase inhibitor, are coordinated in a complex “system” with a role in the innate antimicrobial defense of the skin. In the differentiating keratinocytes of epidermis, the RNases are complexed to RI and inhibited, but when

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the keratinocytes mature into the stratum corneum, the RI is dislocated and then degraded by serine proteases. Thus, the released RNases are free to exert their antimicrobial activity. The authors have reported that the latter is dependent upon the ribonucleolytic activity of RNase 5, as this is inhibited also by small nonprotein inhibitors, such as diethylpyrocarbonate. This conclusion, and that proposed by other authors (Huang et al. 2007) on the irrelevance of the RNase activity for RNase 7 bactericidal activity, are surprisingly in contrast. Further experiments will clarify this issue.

1.2.1.7

Human RNase 8

RNase 8 (Harder and Schroder 2002; Zhang et al. 2002) has a high sequence identity with RNase 7 (78%), but the two evolutionarily related RNases are in several ways different: (1) RNase 7 is present in several tissues, whereas RNase 8 is produced only in placenta and (2) the disulfide bridge that links Cys residues at positions 84 and 26, conserved in all mammalian RNases, cannot form in RNase 8, because Cys-84 is not in the sequence. As a Cys residue, absent in all other RNases, is found at position 69, one should presume that disulfide 84/26 is replaced by a 69/ 26 disulfide. It would be interesting to examine the 3D structure of RNase 8. RNase 8 is active as an antibacterial agent against several Gram-positive and Gram-negative bacteria (Rudolph et al. 2006); its apparent function is to protect placenta from infections.

1.2.2

Rodent RNases

The pancreatic RNase from rat (Rattus norvegicus) was the next vertebrate RNase to be isolated (Beintema and Gruber 1965) following RNase A, the bovine pancreatic enzyme, by a quarter of a century (Kunitz 1940). The 3D structure of the protein, obtained by X-ray crystallography, was determined more recently (Gupta et al. 1999). Also recent is a detailed study on the characterization of mouse RNase 6 (Dyer et al. 2004), and mouse eosinophil-associated RNases (EAR). They have received a special attention for their rapid evolutionary expansion, characterized by gene duplication or deactivation (Cho et al. 2005; Zhang et al. 2000). As for murine angiogenins, the family includes six members (Brown et al. 1995; Strydom 1998), with angiogenin 1 from mouse (mAng-1) identified as the murine orthologue of human angiogenin (Bond and Vallee 1990; Holloway et al. 2005). mAng-1 is expressed in a wide variety of tissues during and after embryogenesis, whereas mAng-4 is expressed only in gut and pancreas (Crabtree et al. 2007). As mAng-4 is upregulated in the Paneth cells of the gut by bacteria (Hooper et al. 2003), it has been proposed that it is implicated not only in gut angiogenesis, but also in gut innate immunity.

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Site-directed mutagenesis studies and the 3D structure of mAng-4 have shown that Glu115 has a role in the low enzymatic activity of mAng-4 similar to that of Gln117 in human angiogenin (see above, Sect. 1.2.1.4) (Crabtree et al. 2007).

1.2.3

Bovine RNases

1.2.3.1

RNase A

The prehistory of the superfamily of secreted vertebrate RNases started with an investigation of Walter Jones from Johns Hopkins Medical School (Jones 1920) aimed at defining the nature of the internucleotide linkage in “yeast nucleic acid,” the predecessor of RNA in biochemical nomenclature. The enzyme that degraded RNA entered the stage of the scientific literature mainly for its high heat resistance: Surprisingly, an aqueous extract of pig pancreas after boiling could still degrade RNA into nucleotides. There was substrate specificity, because thymus nucleic acid (i.e., DNA) was not degraded. Many years later, Rene´ Dubos isolated the enzyme, called it ribonuclease, and confirmed that it was a protein very resistant to heat (Dubos and Thompson 1938). Almost simultaneously, Moses Kunitz crystallized the enzyme from bovine pancreas (Kunitz 1940). Then in the 1950s, the enzyme was further purified using the new, sophisticated chromatographic procedures offered to biochemists, mostly based on ion-exchange and affinity chromatography. The preparation of fully homogeneous enzyme led also to the identification of two RNases in bovine pancreas: RNase A and RNase B, the latter a glycosylated form of RNase A (Plummer and Hirs 1964). RNase A however, as illustrated in the Premise to this chapter, has become over the years a synonym of ribonuclease, after its use as a model protein/enzyme, and the discoveries that these studies produced. In fact, RNase A has been used as a name for the whole superfamily of tetrapod, later of all vertebrate RNases, and it is not unusual to read in scientific journals the expression “RNase A RNase” to denote an RNase from the so-called RNase A superfamily. Here, as indicated in the Premise, we use the more straightforward “vertebrate-secreted RNase superfamily”. Timely and exhaustive reviews have been produced on RNase A in a chronological order by: Frederic Richards and Harold Wyckoff; Peter Blackburn and Stanford Moore; Ronald T. Raines (Blackburn and Moore 1982; Raines 1998; Richards and Wyckoff 1971). In a book on ribonucleases (D’Alessio and Riordan 1997), several chapters were dedicated to various aspects of RNase A as an enzyme and as a protein. The identification of key residues in the mechanism of action and stability of RNase A has been successfully carried out for decades, first by modifying through chemistry the side-chains of the appropriate residues, then by heterologous production of recombinant variants.

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Yet, quite recently, an original methodology has been proposed (Smith and Raines 2006), in which RNase A and human angiogenin (RNase 5, see Sect. 1.2.1.4) were used as model proteins in a methodical search for residues important for structure and function of the two proteins. Libraries of mutated RNase A and angiogenin genes were obtained by using error-prone polymerase chain reactions. When a gene encoding an active RNase was expressed in a bacterial cell, engineered to allow for disulfide bond formation in the cytosol, the cell was killed by the active RNase. Thus, inactive variants could be readily selected. Many residues (about 10–15% of the total) were found to be not amenable to substitution in either RNase A or homologous angiogenin, and only a few were sensitive in both proteins. Although the importance of many among these residues was not clear, the identified genes may be suggestive targets for future studies. Investigations aimed at producing dimeric variants of RNase A have been favored in the last decades. The rational basis of this approach was twofold. It was prompted: (1) by the results with homologous, but dimeric, seminal RNase (see Sect. 1.2.3.2), endowed with many surprising, interesting bioactivities and (2) by the startling discovery that lyophilization of RNase A from a solution of acetic acid led to the formation of dimers (Crestfield et al. 1962). Furthermore, these RNase A dimers were found to be constructed by the exchange or swap of the N-terminal segments of the two protomers (Crestfield and Fruchter 1967). Years later, Mazzarella and coworkers determined the three-dimensional structure of seminal RNase (BS-RNase), and found that this dimeric protein was organized with the swap of N-terminal segments between subunits (Mazzarella et al. 1993). These data motivated Eisenberg and coworkers, working at that time on a similar exchange of parts between the subunits of diphtheria toxin (Bennett et al. 1994), to study the “Crestfield-type” dimers. They crystallized and determined the structure by X-ray crystallography of an RNase A dimer with N-terminal swap (Liu et al. 1998), and later of a dimer in which the swap involved segments from the C-terminal region of the protein (Liu et al. 2001). Eisenberg proposed the name of “three-dimensional domain swap” for the exchange of parts between subunits in an oligomeric protein, and demonstrated that the swap of structural domains between oligomers is a general solution to oligomer stability and function (Bennett et al. 1995). Libonati and coworkers later reported that when RNase A is lyophilized from dilute acetic acid, it not only forms dimers, but also trimers, tetramers, and pentamers (Gotte et al. 1999; Gotte and Libonati 2004). The ordered assemblage of RNase A into oligomeric structures upon treatment with acetic acid and lyophilization is surprising, as both the acid itself (LopezAlonso et al. 2010), and lyophilization (Griebenow and Klibanov 1995), induce a profound perturbation of the protein structure, especially its secondary structure. An alternative method to make dimers of RNase A consisted using bifunctional reagents to cross-link two RNase A molecules (Bartholeyns and Baudhuin 1976). Later, site-directed mutagenesis was used to demonstrate that RNase A could be transformed into a stable and active dimeric RNase when key residues were

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replaced by the corresponding residues present at the intersubunit surface of naturally dimeric seminal RNase (Di Donato et al. 1994). Another approach to make dimers of RNase A was based on the use of the poorly active enzyme variants H12A and H119A (Park and Raines 2000). When a 1:1 mixture of the two variants was incubated at pH 6.5 and 65 C, a large increase in ribonucleolytic activity resulted, suggesting that dimers were constructed with swap of parts between two enzyme units. Furthermore, when the mixture was lyophilized, active dimers were obtained. The results suggested the hypothesis of a monomer–dimer equilibrium, with a Kd 20-fold greater than the concentration of RNase A in the cow pancreas. Furthermore, these results, and physicochemical considerations, led the authors to two provocative conclusions: (1) There are RNase A dimers also in vivo even at 37 C, and (2) RNase A has an intrinsic, pre-evolved ability to form domain-swapped dimers. More recently, dimeric RNase A was constructed as a tandem protein (Leich et al. 2006) made up of two RNase A units fused through a peptide linker connecting the C terminus of one unit to the N terminus of another unit. Even though one of the dimer protomers was bound to and neutralized by the RNase inhibitor (see below), the tandem dimer was found to be active and strongly cytotoxic toward malignant cells. An even more radical solution toward protein dimerization has appeared (Simons et al. 2002), and applied to RNase A as a model protein (Simons et al. 2007). The protocol consists in linking covalently two RNase A molecules by creating an amide bond between the side-chain of Lys66 in one unit and that of Glu9 in a partner unit. The bond was generated by heating in vacuo at 85 C lyophilized preparations of RNase A. The linkage was dubbed “zero-length amide cross-link.” Later, the product of the heating reaction was found to be heterogeneous, with various amino and carboxyl groups forming amide bonds (Vottariello et al. 2010). Furthermore, the “zero-length” dimers of RNase A were found to have no cytotoxic activity toward tumor cells, unless “cationized” as described by J. Futami and coworkers (see below). However, the “cationized” dimers were as cytotoxic toward nonmalignant cells as toward malignant cells. A different approach to investigate the structure and the function of RNase A by X-ray crystallography and molecular dynamics simulations provided evidence that subtle b-sheet motions are essential in RNase A for substrate binding and product release (Berisio et al. 2002; Vitagliano et al. 2000, 2002), and that these motions corresponded to intrinsic dynamic properties of the native enzyme (Merlino et al. 2002; Merlino et al. 2003). The high quality of the electron density maps obtained for RNase A structures at six distinct pH values (Berisio et al. 1999, 2002) has allowed direct detection of the deprotonation of the catalytic His12 residue, corroborating the reaction mechanism proposed by kinetic and structural studies (Cuchillo and Vilanova 1997). Furthermore, this approach led to an accurate picture of the active site as well as to the observation of concerted structural changes in regions even remote from the active site.

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It has been pointed out above that the rational basis for the construction of RNase A dimers was largely due to the unusual dimeric structure of BS-RNase. But it was the antitumor action of dimeric BS-RNase, absent in its monomer (Vescia et al. 1980), that effectively stimulated the research on dimerized RNase A, especially for its translational value in innovative cancer therapies. RNase A had long been tested as an antitumor agent, but eventually, this research track was abandoned when it became clear that RNase A was antitumoric only when administered in very large amounts (Ledoux 1955). It has been hypothesized (Youle and D’Alessio 1997) that prolonged storage and/or repeated lyophilizations could have induced the production of dimeric forms of the enzyme, as recorded in the literature (Richards and Wyckoff 1971). Likely, these dimeric derivatives, present in old preparations of RNase A as minor protein contaminants, were the actual responsible determinants of the antitumor action of the RNase A preparations tested. All dimeric RNase A derivatives described above have been found to possess various degrees of cytotoxic activity, although it was not always determined that such cytotoxic activity was selective for tumor cells, with no adverse effects on nonmalignant cells. However, monomeric variants of seminal RNase have been found to have cytotoxic activity (Lee and Raines 2005). They will be discussed in Sect. 1.2.3.2. Beside dimerization of the protein, other experimental approaches were tested for providing RNase A with a cytotoxic activity. One approach has been to render the protein much more cationic through chemical modifications. This conferred a pronounced cytotoxic activity on RNase A toward a malignant fibroblast cell line (Futami et al. 2001). The observed cytotoxicity could be readily correlated with the net positive charge of the derivatized protein. On the other hand, the enzyme activity decreased and was only partially inhibited by the RNase inhibitor. Yet, the authors concluded that the cytotoxic activity of the “cationized” RNase was mainly due to its lower affinity for the acidic RNase inhibitor, and possibly also to its more efficient adsorption by the cells. No tests were carried out on nonmalignant cells, and the possibility was not verified that the derivatives were as toxic to cells as any super-cationic protein material might be, including polylysine and other cationic, nonbiological substances (Kornguth et al. 1961). Another approach to the construction of RNase A derivatives with antitumor activity has been proposed recently (Rutkoski et al. 2010), based on the production of RNase A multimers generated with thiol-reactive cross-linking reagents. Derivatives with various degrees of cytotoxic activity were obtained, depending on the affinity of the derivatives for RI, which in turn depended on the site of conjugation and the propinquity of the monomers within the conjugate. While in vitro the antitumor activity was hindered by the increased hydrodynamic radius of the derivatives, tests in vivo on laboratory mice gave more favorable results, likely for the higher levels of large derivatives in circulation.

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1.2.3.2

15

Seminal RNase and the Roles of the RNase Inhibitor

In the 1950s, the most prominent textbook of Biochemistry (Fruton and Simonds 1958) described RNase, obviously RNase A, as a very small and very heat-resistant protein. Now, given that heat-resistant, small proteins would be, in principle, more practical to study than large, heat-labile proteins, it is not surprising that an RNase was chosen for a new research line. The source selected for the study was bovine seminal plasma, a secretion of seminal vesicles, very rich in many enzyme activities, and very rich, it was soon discovered, in RNase activity (D’Alessio 1963). The enzyme was called bovine seminal RNase (BS-RNase). Seminal RNase was much larger and more cationic than RNase A (D’Alessio et al. 1997, 1991), as reported also by others who co-discovered the RNase (Hosokawa and Irie 1971). Furthermore, BS-RNase was not monomeric, but a homodimer with two disulfide bridges linking the two subunits (Di Donato and D’Alessio 1973). To date, BS-RNase is still the only dimeric RNase within the whole superfamily of secreted vertebrate RNases. Seminal RNase is less active as an RNase than many other RNases of the superfamily – e.g., it is about 50% as active as RNase A – but it displays several special, i.e., non-catalytic, biological actions: it is aspermatogenic, immunosuppressive, cytotoxic for tumor cells, and antiviral. This extraordinary multiplicity and variety of biological activities (D’Alessio 1993), and the protein dimeric structure, unique for an RNase and the structural basis for most of these bioactions, have directed the research on this RNase. It should be noted, however, that most of these special activities of BS-RNase are not physiologically significant. They are merely reflections off the mirrors proposed by the assay systems with which the RNase is confronted (D’Alessio 1993). A seminal-type gene evolved about 35 million years ago, at the time of the divergence of ruminants, likely through a duplication of the gene encoding bovine pancreatic RNase A. However, it remained a pseudogene in all evolved ruminants until about 5–10 million years ago, when the pseudogene was repaired (D’Alessio 1999; Sassi et al. 2007; Trabesinger Ruef et al. 1996), likely through gene conversion (Sassi et al. 2007; Trabesinger Ruef et al. 1996). Interestingly, in water buffalo, a seminal-type gene is expressed (Kleinedam et al. 1999), but the protein is not produced, apparently because one of the Cys residues engaged in the intersubunit disulfide bridges is replaced by a Phe. Thus, not only one of the two intersubunit disulfides cannot form in buffalo seminal RNase, but also the presence of a free thiol (the surviving Cys residue) can severely impair RNase survival in an extracellular, oxidizing environment. Considering the co-presence in seminal RNase of two structural arrangements (see below), and the ability of only one of them to exert a special bioaction(s), it has been surmised that the case of the rapid BS-RNase evolution is that of an “evolution in progress” toward a protein with a new physiological role (D’Alessio 1995, 1999; Trabesinger Ruef et al. 1996).

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This leads to the question of what is the physiological role of BS-RNase in Bos taurus, the only vertebrate in which such an unusual RNase is expressed. Given that some of the special bioactions of the seminal enzyme, such as the aspermatogenic and the antitumor actions, are unlikely to have any importance in bovine seminal vesicles or seminal plasma, it has been proposed that the role of BSRNase is that of an immunosuppressive agent (Tamburrini et al. 1990). The physiological significance of the immunosuppressive activity of BS-RNase (Soucek et al. 1983, 1996) can be based on the fact that male sperm cells are immunogenic when introduced in the female reproductive organs. Thus, a high concentration of an immunosuppressive agent in seminal plasma, where BS-RNase concentration is more than 1 mg/mL, can well protect the female genital area from the sperm attacks. When the amino acid sequences of BS-RNase and RNase A – the superfamily prototype, with no immunosuppressive or antitumor activities – are compared, 13 substitutions can be identified, with four of them found to have a key role in establishing the dimeric structure (Di Donato et al. 1994; Mazzarella et al. 1995). The residues involved are Cys31 and Cys32 forming the intersubunit disulfides with Cys32 and Cys31 of the partner subunit, respectively, and Leu28, which forms an essential hydrophobic contact at the intersubunit interface. The latter residue has an important role in another unusual structural property of this RNase: the peculiar exchange or swap of the N-terminal a-helical segment between the two subunits (Capasso et al. 1983). But the story of BS-RNase structure is even more complicated, because later it was found (Piccoli et al. 1992) that seminal RNase has two distinct quaternary arrangements: In one (called MXM), the dimer subunits exchange their N-terminal segments; in the other (called M¼M) there is no exchange. The structures of the isoforms MXM (Mazzarella et al. 1993) and M¼M (Berisio et al. 2003) of BS-RNase have been determined by X-ray crystallography (see Fig. 1.2). It should be noted that BS-RNase not only has access to two different structures (MXM and M¼M), but also that it exists in solution with both structures in a 2/1 equilibrium. When one of the two structures is removed from the equilibrium, the remaining structure is transformed into the other until equilibrium is again restored.

M=M

Fig. 1.2 The two isoforms of seminal RNase in equilibrium: the MXM isoform with exchange between subunits of the N-terminal a-helical arms, and the M¼M isoform without exchange

M×M

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This additional, unusual characteristic, and the finding that the antitumor action of the enzyme is not present in the M¼M dimer with no exchange of parts (Cafaro et al. 1995) are both consistent with the above hypothesis that BS-RNase is an example of evolution in progress. The swapping propensity of BS-RNase does not depend, as could be expected, on the primary structure of the 16–22 hinge loop that connects the main protein body to the N-terminal arm (Picone et al. 2005). Surprisingly, substitution by directed mutagenesis of Arg80 with Ser, while replacing the BS-RNase hinge with that of RNase A, induces a conformational change in the hinge structure such that the MM:M¼M equilibrium ratio is inverted from 2:1 to 1:2. This suggests that Arg80 may trigger the swapping of N-terminal ends in the native enzyme, thus stabilizing its swapped form (Ercole et al. 2007; Merlino et al. 2008). Once BS-RNase enters the cytosol, two events occur: reduction of its intersubunit disulfides by the reducing environment of the cytosol, and interactions of the RNase with the cytosolic RNase inhibitor. Upon reduction of the intersubunit disulfides, the M¼M dimer dissociates into monomers, which are readily bound and neutralized by the inhibitor. The MXM dimer instead remains dimeric also after reduction of the intersubunit disulfides, as in this non-covalent dimer (NCD-BS), the monomers are interconnected to each other by the swap of the N-terminal ends (Murthy et al. 1996). The structure of NCD-BS, stabilized by alkylation of the free thiols exposed by reduction of the intersubunit disulfides, has been found by X-ray crystallography to be close to that of the covalent MM isoform of the native RNase (Sica et al. 2004). Furthermore, model building of the complexes between NCD-BS and RI has indicated that the quaternary structure of NCD-BS appears to be designed to resist binding to RI (Merlino et al. 2008; Sica et al. 2004). These findings may well have a physiological significance, as resistance to neutralization by RI is essential for BS-RNase to display its cytotoxic activity. Recent findings on the biological effects of BS-RNase have shown that cells treated with BS-RNase undergo apoptosis; however, the effect is exerted both in tumor and in normal cells (Sinatra et al. 2000). Of interest is the finding that the effect is generated by a decreased telomerase activity (Viola et al. 2005). Special attention has been given in several laboratories to the mechanism of antitumor action of BS-RNase. The essential determinants for the antitumor action of BS-RNase can be: (1) the integrity of its catalytic activity (Vescia et al. 1980), (2) its dimeric structure (Vescia et al. 1980), (3) its ability to concentrate at the extracellular matrix of the target tumor cells (Mastronicola et al. 1995), and (4) its ability to bind target cell membranes, both plasmatic and intracellular (Notomista et al. 2006); its ability to translocate to the cytosol (Leich et al. 2007). The importance of crossing cell membranes is supported by results obtained also with other dimeric RNases (Notomista et al. 2006) selectively cytotoxic for tumor cells, including engineered BS-RNase variants, dimeric RNase 1 (Piccoli et al. 1999), and Crestfield-type RNase A dimers (Matousek et al. 2003). Another important determinant of BS-RNase antitumor action is the resistance of the RNase to RI, the RNase inhibitor. This is often considered the most important

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determinant of all, because: (1) onconase (see Sect. 1.3.3.1), the other naturally antitumor RNase, is also resistant to RI (Wu et al. 1993) and (2) RNases with no cytotoxic activity, such as bovine RNase A or human RNase 1, with a very high affinity for RI, become cytotoxic when they are engineered into RI-resistant RNases (Antignani et al. 2001; Haigis et al. 2003; Leland et al. 1998; Piccoli et al. 1999; Suzuki et al. 1999). These results have led to the description of RI role in the cell as a sentry to guard cells from the entry in the cytosol of cytotoxic foreign RNases such as BS-RNase (Leland et al. 1998) or onconase (Turcotte and Raines 2008). Consistently, cells manipulated to increase their RI levels become more resistant to RNase cytotoxicity (Rutkoski and Raines 2008). Even the dimeric structure has been found to be unnecessary as a structural determinant for RNase cytotoxicity, when BS-RNase monomers are engineered into RI-resistant RNase variants (Lee and Raines 2005). RI is in the cytosol, where most cellular RNA is. Hence, it is reasonable to assume that RI will protect cytosolic RNA by neutralizing any potentially damaging RNases, obviously on condition that these RNases enter the cytosol. However, RI resistance cannot be the only determinant of cytotoxicity, because RI-resistant RNases are cytotoxic even for cells deprived of RI (Monti and D’Alessio 2004), i.e., nontoxic RNases do not become toxic when the RI sentry is absent. Furthermore, there are cytotoxic RNases that are resistant to RI, such as Crestfield-type RNase A dimers (Matousek et al. 2003; Naddeo et al. 2005), and tandem RNase A dimers (Leich et al. 2006). On the other hand, it has been shown that: (1) RI protects cells from oxidative stress (Cui et al. 2003; Monti et al. 2007; Wang and Li 2006), (2) RI counteracts the effects of ANG when cells are under oxidative stress and ANG inactivates tRNAs to arrest translation (see Sect. 1.2.1.4), and (3) RI neutralizes RNases in epidermal cells until the epidermis matures into stratum corneum, and RI is eliminated and the RNases can exert their bactericidal action (see Sect. 1.2.1.6). These results indicate that any discussion about the cellular roles of RI should be simplified and dealt with within two distinct scenarios. One is that of RNases cytotoxicity; the other is that of the physiological role(s) of RI. In the scenario of RNases’ cytotoxic action, RI can certainly play the “sentry” role, although such role may not be always conclusive, as there are cases in which RI does not play this role, and other determinants of cytotoxicity, not described as yet, must be considered. As for the scenario in which the physiological roles of RI are discussed, we can try and consider the question in evolutionary terms. Given the importance to ensure protection to stressed cells, or prepare a bactericidal skin, it can be certainly accepted that evolution invested so much to maintain a physiological, valuable agent, obtained through a high concentration of RI in cytosol, with a high affinity for RNases with physiological roles. On the other hand, it would be difficult to accept that evolution invested so much just to avoid accidents, i.e., damages from RNases that accidentally entered cells.

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1.3 1.3.1

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Non-mammalian RNases Bird RNases

Only RNases from chicken (Gallus gallus) have been investigated. After early studies (Hayano et al. 1993; Klenova et al. 1992; Levy and Karpetsky 1980; Nakano and Graf 1992), three vertebrate RNases from chicken have been identified (Cho et al. 2005). Two of them, called RNases A-1 and A-2, have been recently studied (Nitto et al. 2006). The cationic RNase A-2, with a higher RNase activity and a very high pI of 11.0, has been found to possess angiogenic and bactericidal activities. An enzymatically inactive H110A variant of RNase A-2, obtained by mutagenesis of the catalytically essential His110, was found to retain the bactericidal activity. Surprisingly, human RNase inhibitor was found to effectively inhibit both RNases A-1 and A-2, despite the phylogenetic distance between Homo and Gallus, and the reported absence of inhibition by human RI of RNases from non-mammalian species: an amphibian (Wu et al. 1993) and a reptile (Nitto et al. 2005). Given the bactericidal activity of RNase A-2, and its localization in peripheral blood granulocytes, the RNase can be added to the impressive list of bactericidal RNases from the vertebrate-secreted RNase superfamily (Boix and Nogues 2007). Studies of the structural determinants necessary for the bactericidal activity of RNase A-2 (Nitto et al. 2006) identified two domains, defined by amino acid sequences 71–76 (domain II) and 89–104 (domain III), to be responsible for the activity. The activity was still present when the domains were isolated from the protein. The third domain was inactive. Surprisingly, the peptide from domain II was found to be active also in its scrambled form. However, as the authors suggest, the high content of basic and hydrophobic residues in the short, 6-residue peptide (Arg-Arg-His-Phe-Arg-Ile) may explain the apparently nonsensical result. It has already been found for other RNases, and discussed in Sect. 1.2.1.2, that the RNase activity and even the RNase structure are not essential for the mechanism of bactericidal action.

1.3.2

Reptile RNases

Only few RNases from reptiles have been described. One from Iguana iguana and two from turtles: Chelonia mydas, the marine green turtle, and Chelydra serpentine, the snapping turtle. They are structurally and phylogenetically close to mammalian angiogenins, as indicated also by the absence in their structure of the disulfide bond 65–72 (RNase A numbering). The Iguana iguana RNase has been sequenced (Zhao et al. 1994) and characterized (Nitto et al. 2005) showing that the enzyme has all the essential features of vertebrate RNases. It is not inhibited by the human RNase inhibitor, and it does not

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display any bactericidal or cytotoxic activity. In fact, it is expressed mainly in pancreas, so that it may be surmised that it has a digestive role. Recently, the RNase from the egg-white of the marine green turtle has been isolated, sequenced, and characterized (Katekaew et al. 2006, 2007). Furthermore, its 3D structure was determined by X-ray crystallography (Katekaew et al. 2010). The protein has the typical a/b folding of vertebrate RNases and an overall structure very similar to that of a murine angiogenin (PDB code 2bwl).

1.3.3

Amphibian RNases

Amphibian RNases have been studied exclusively from the Rana genus. The best studied member is onconase (ONC), or ranpirnase, from Rana pipiens (see next section). The other species investigated have been Rana catesbeiana (bullfrog) and Rana japonica (Japanese rice paddy frog) (Irie et al. 1998; Kamiya et al. 1990; Nitta et al. 1989; Titani et al. 1987). The latter RNases, with an amino acid sequence 50% identical to that of ONC, have features very similar to those of ONC: localization in frog oocytes, cytotoxicity, high stability, and disulfides arrangement (see below). The 3D structure of Rana catesbeiana, determined both in solution (Chang et al. 1998) and by X-ray crystallography (Leu et al. 2003), has been found to be similar to that of ONC. Its gene has been found to be expressed in the liver (Huang et al. 1998); then it moves to oocytes, where it is stored. An updated catalog of Rana family RNases has been recently proposed: It includes many members from Rana catesbeiana (Liao et al. 2000; Rosenberg et al. 2001), and a member from Rana pipiens, identified from a cDNA encoding an RNase expressed only in liver from female frogs (Chen et al. 2000).

1.3.3.1

Onconase

Onconase (ONC, generic name ranpirnase) is a ribonuclease from the oocytes of Northern Leopard frogs (Rana pipiens) with cytotoxic activity toward tumor cells in vitro as well as in vivo in animal models; furthermore, it enhances sensitivity of tumor cells to several standard cytotoxic agents (Ardelt et al. 2008; Arnold and Ulbrich-Hofmann 2006; Lee and Raines 2008; Rybak 2008). ONC has been previously tested in Phase III clinical trials in patients with malignant mesothelioma (Mikulski et al. 2002). At present, ONC is tested in Phase II clinical trials on patients affected by non-small cell lung cancer, in combination with antineoplastic drugs. It is also tested in preclinical tests against pathogenic viruses (Press Release from Tamir Biotechnology, Inc., formerly Alfacell Corporation). Onconase is a small RNase, containing only 104 residues, compared with 124 residues in RNase A, but its 3D structure (Mosimann et al. 1994) is that typical of vertebrate RNases. Differences are mainly localized in the loop regions, shorter

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than in the other RNase homologs, and at the C terminus, where a disulfide bond that links Cys104 to Cys87 is present, a bond absent in mammalian RNases. Another peculiar feature in the structure of ONC is the N-terminal pyro-Glu residue, found to be essential for optimal enzymatic and cytotoxic activity (Newton et al. 1998). The most relevant structural feature of ONC is its exceptional stability, recognized through physicochemical and proteolytic methodologies (Notomista et al. 2000). It has also been suggested that resistance to denaturation and proteolytic attacks may be at the basis of its persistence, hence toxicity, in kidneys of treated patients. Variants of ONC have been constructed in which the N-terminal cyclized glutamine and the C-terminal disulfide bridge have been removed. The results, obtained by using calorimetry and limited proteolysis (Notomista et al. 2001), as well as X-ray crystallography and molecular dynamic simulations (Merlino et al. 2005), have confirmed the key role of the C- and N-terminal regions in determining both the high stability, and the low catalytic activity of the enzyme. A recent detailed analysis of the structure of ONC has been carried out by studying folding and unfolding of the protein (Schulenburg et al. 2009, 2010). As for its mechanism of cytotoxic action, the ability of the enzyme to evade the neutralizing effect of the ribonuclease inhibitor (Wu et al. 1993) has been considered as determinant. However, ONC does bind RI, although with a much lower affinity than that measured for RNase A or angiogenin. The Ki for the inhibition, determined at low salt concentration, has been measured to be 0.15 mM (Turcotte and Raines 2008), about seven orders of magnitude weaker that that measured for the RI/RNase A complex. According to a recent hypothesis, the antitumor activity of ONC might be mediated through RNA interference. It has been found (Zhao et al. 2008) that the silencing in human lung adenocarcinoma cells of the gene encoding glyceraldehyde3-P-dehydrogenase could be effectively averted by ONC.

1.3.4

Fish RNases

For a long time, it has been debated whether homologs of archetypal RNase A were present only in tetrapods (mammals, birds, reptiles, and amphibians), i.e., that RNases were not present in the earliest class of vertebrates: fishes. However, the negative findings were simply due to the methodologies employed. First, the search went on with the classical biochemical approach: purification from animal tissues using assays of RNase activity. Then, when recombinant DNA techniques became available, Southern and Northern blots were carried out, generally using as probes oligonucleotides from the RNase A gene sequence. It is now clear that RNases are present in fish, thus completing the vertebrate RNase superfamily, as described in the Premise to this chapter. Clearly, the very low RNase activity of fish RNases (see below) has rendered their purification with

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standard procedures difficult if not impracticable. Also difficult was the identification of fish RNase genes homologous to that encoding RNase A, given the relatively low similarity in nucleotide sequences between bovine and fish genes. When fish genome sequences became available, undoubtedly comprising RNase genes, the first recombinant fish RNases could be obtained and studied, first from zebrafish (Danio rerio) (Cho and Zhang 2007; Kazakou et al. 2008; Pizzo et al. 2006, 2010), then from Atlantic salmon (Salmo salar) (Pizzo et al. 2008). More RNase sequences have been identified in several families of bony fishes, such as Cyprinidae, Ictaluridae, Moronidae, Nototheniidae, and others, as identified in the “vertebrate EST” database, inquired with the tBLASTIN algorithm. Ten RNase genes have been reported in the species Oryzias latipes (the common name is medaka) from the Adrianichthyidae (Cho and Zhang 2007). However, several of them are noncanonical members of the RNase superfamily because they lack one or more essential residues of the catalytic triad or one or more residues of the RNase signature (CKxxNTF). Furthermore, two of the medaka RNases have identical sequences. Thus, only four medaka RNases are canonical members of the superfamily. The fishes mentioned above are all bony fishes so that the possibility has been considered that no RNase genes are in the genomes of cartilaginous fishes. This may not be surprising, because the prospect can be considered that the most ancestral RNase gene(s) did not evolve when vertebrates separated from invertebrates, but when the separation occurred between cartilaginous and bony vertebrates. Certainly surprising instead is the absence of RNase sequences in certain bony fish genomes, such as those of Fugu rubripes, Tetraodon nigroviridis, and Gasterosteus aculeatus. However, the sequence of these genomes is not complete yet. The most studied fish RNases to date are those from zebrafish (Cho and Zhang 2007; Kazakou et al. 2008; Pizzo et al. 2006, 2010), which comprise in their genome a high degree of polymorphism (Kazakou et al. 2008). The latter finding is suggestive (Cho and Zhang 2007) of a rapid diversification through gene sorting of the RNase genes since the Devonian Period, when the early fish evolution took place. Based on their findings and analyses, the authors have suggested that at least two early RNase genes were present as last common ancestors of all vertebrate RNases. The main characteristic properties of the fish RNases studied so far can be summarized as follows. Fish RNases display: (1) a low or very low RNase activity, (2) an angiogenic activity, and (3) a bactericidal activity. Based on the latter finding, the hypothesis has been advanced that the ancestral members of the vertebrate RNases superfamily were involved in host-defense mechanisms (Cho and Zhang 2007). Consistent with this hypothesis is the finding of high levels of ZF-RNase-2 and -3 in zebrafish liver and gut, suggestive of their involvement in innate immunity just as it has been found for mouse angiogenin (Hooper et al. 2003) (RNase 5, see Sect. 1.2.1.4). Later, also the RNases from Atlantic salmon have been found to be bactericidal (Pizzo et al. 2008). However, other data do not fit to

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this hypothesis, particularly the findings on the conservation of the bactericidal activity in denatured RNases, as it may be difficult to recognize the evolution of active, functioning molecules lacking structure (see Sect. 1.2.1.2). Another hypothesis, based on the angiogenic activity of all fish RNases studied so far, and the importance of this activity in cellular growth, is that the ancestral RNases were angiogenins (Pizzo and D’Alessio 2007; Pizzo et al. 2008). Obviously, the possibility cannot be excluded that in a gene sharing manner (Piatigorsky and Wistow 1991) ancestral RNases were involved both in angiogenesis and host defense. As for their 3D structures, determined by X-ray crystallography and model building, it has been found (Kazakou et al. 2008; Pizzo et al. 2010) that in some of them, the active site is partly obstructed as described for human angiogenin (see Sect. 1.2.1.4), a structural feature that is consistent with the low catalytic activity of human angiogenin, and zebrafish RNases as well. In other zebrafish RNases, no obstruction is evident, so that the low activity must have a different structural basis. It has been proposed (Kazakou et al. 2008) that likely the ancestor(s) of all superfamily members were RNases with blocked active sites, later evolved into enzymes efficient for catalysis. This hypothesis is in line with the hypotheses above, as it assigns to ancestral RNases functions different from RNA degradation, such as angiogenic and/or host defense. Recent studies (Monti et al. 2009; Pizzo et al. 2010; Quarto et al. 2008) have revealed that the ZF-RNases may have different roles or functions, given their different expression in embryogenesis and adult life, and different fate in the cell. Only ZF-RNase-1 and -2 are very efficient as angiogenins even though also ZF-RNase-3 is able to induce phosphorylation of extracellular signal-regulated kinase 1/2 mitogen-activated protein kinase, and nuclear translocation. The case of bactericidal ZF-RNase-3 has been briefly described in Sect. 1.2.1.2. Its activity is due to the presence in its sequence of a peptide that is produced through a specific cleavage catalyzed by an OmpT outer membrane protease from the bacterium itself. Also two RNases from Atlantic salmon conserve their bactericidal activity after full denaturation (Pizzo et al. 2008), as previously discussed in Sect 1.2.1.2.

1.4

Conclusions and Perspectives

The story of the superfamily of vertebrate-secreted RNases started more than a century ago, and has been written by hundreds of scientists. Thus, it may not be surprising that after all the efforts dedicated to this superfamily of RNases, many questions on the function and structure of RNases have found answers. One important evidence is that evolution has played intensely with this simple molecule, and in many ways, apparently to confer to the RNase small structure different functions and abilities. Hence, it is not unexpected that when vertebrates evolved, these RNases readily evolved with them, to fully exploit their stable structure, and

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meet a series of physiological needs. Vertebrate-secreted RNases function, in all vertebrates, as valuable tools in many ways. These comprise, or apparently comprise, such key physiological needs as: production of blood vessels; RNA degradation in ruminants diet; biogenesis of rRNA; control of the arrest of translation under stress; host defense against microbes and/or parasites; immunosuppression in bovine reproduction. We still have no clear answer as to why in many cases only the RNase structural scaffold has had evolutionary success, while in other cases the ability of vertebrate RNases to degrade RNA has been conserved, although often we do not know the RNA target(s) of these RNases. The findings that some RNases have an angiogenic, or a microbicidal function, have certainly surprised the investigators who first discovered them, so we can expect more surprises. The suspicion remains that in cases in which the RNA degradation has no significance, this is because the real function and/or mechanism of the RNase has escaped the attention of the investigators. Certainly, several issues are open to investigation. Why some vertebrate genomes are rich in RNases, while in others, only few members have evolved? What are the biological implications of RNases apparently incapable to perform as RNases? How is angiogenin involved in the mechanism of amyotrophic lateral sclerosis? Is it possible that evolution has led to the production of RNases as mere containers of active peptide segments? Can we concretely exploit the translational value of angiogenin/neomycin, cytotoxic RNases, immunoRNases for novel approaches to cancer therapy? Looking from above with a bird’s eye at the vast landscape of vertebratesecreted RNases, the feeling is that we still know only a fraction of what we should know about them. Acknowledgments I wish to thank my colleagues Tina Giancola, Guo-Fu Hu, Lelio Mazzarella, Antonello Merlino, Elio Pizzo, James F. Riordan, and Filomena Sica, who kindly contributed to improve the manuscript with their suggestions.

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Soucek J, Marinov I, Benes J, Hilgert I, Matousek J, Raines RT (1996) Immunosuppressive activity of bovine seminal ribonuclease and its mode of action. Immunobiology 195(3): 271–285 Strydom DJ (1998) The angiogenins. Cell Mol Life Sci 54(8):811–824 Subramanian V, Feng Y (2007) A new role for angiogenin in neurite growth and pathfinding: implications for amyotrophic lateral sclerosis. Hum Mol Genet 16(12):1445–1453 Suzuki M, Saxena SK, Boix E, Prill RJ, Vasandani VM, Ladner JE, Sung C, Youle RJ (1999) Engineering receptor-mediated cytotoxicity into human ribonucleases by steric blockade of inhibitor interaction. Nat Biotechnol 17(3):265–270 Tamburrini M, Scala G, Verde C, Ruocco MR, Parente A, Venuta S, D’Alessio G, Tamburrini M, Scala G, Verde C, Ruocco MR, Parente A, Venuta S, D’Alessio G (1990) Immunosuppressive activity of bovine seminal RNase on T-cell proliferation. Eur J Biochem 190:145–148 Titani K, Takio K, Kuwada M, Nitta K, Sakakibara F, Kawauchi H, Takayanagi G, Hakomori S (1987) Amino acid sequence of sialic acid binding lectin from frog (Rana catesbeiana) eggs. Biochemistry 26(8):2189–2194 Torrent M, Cuyas E, Carreras E, Navarro S, Lopez O, de la Maza A, Nogues MV, Reshetnyak YK, Boix E (2007) Topography studies on the membrane interaction mechanism of the eosinophil cationic protein. Biochemistry 46(3):720–733 Torrent M, Badia M, Moussaoui M, Sanchez D, Nogues MV, Boix E (2010) Comparison of human RNase 3 and RNase 7 bactericidal action at the gram-negative and gram-positive bacterial cell wall. FEBS J 277(7):1713–1725 Trabesinger Ruef N, Jermann T, Zankel T, Durrant B, Frank G, Benner SA (1996) Pseudogenes in ribonuclease evolution: a source of new biomacromolecular function? FEBS Lett 382(3): 319–322 Tsuji T, Sun Y, Kishimoto K, Olson KA, Liu S, Hirukawa S, Hu GF (2005) Angiogenin is translocated to the nucleus of HeLa cells and is involved in ribosomal RNA transcription and cell proliferation. Cancer Res 65(4):1352–1360 Turcotte RF, Raines RT (2008) Interaction of onconase with the human ribonuclease inhibitor protein. Biochem Biophys Res Commun 377(2):512–514 Vescia S, Tramontano D, Augusti Tocco G, D’Alessio G (1980) In vitro studies on selective inhibition of tumor cell growth by seminal ribonuclease. Cancer Res 40(10):3740–3744 Vicentini AM, Hemmings BA, Hofsteenge J (1994) Residues 36–42 of liver RNase pl3 contribute to its uridine-preferring substrate specificity. Cloning of the cDNA and site-directed mutagenesis studies. Protein Sci 3(3):459–466 Vicentini AM, Kote-Jarai Z, Hofsteenge J (1996) Structural determinants of the uridine-preferring specificity of RNase pl3. Biochemistry 35(28):9128–9132 Viola M, Libra M, Callari D, Sinatra F, Spada D, Noto D, Emmanuele G, Romano F, Averna M, Pezzino FM, Stivala F, Travali S (2005) Bovine seminal ribonuclease is cytotoxic for both malignant and normal telomerase-positive cells. Int J Oncol 27(4):1071–1077 Vitagliano L, Merlino A, Zagari A, Mazzarella L (2000) Productive and nonproductive binding to ribonuclease A: x-ray structure of two complexes with uridylyl(20 ,50 )guanosine. Protein Sci 9 (6):1217–1225 Vitagliano L, Merlino A, Zagari A, Mazzarella L (2002) Reversible substrate-induced domain motions in ribonuclease A. Proteins 46(1):97–104 Vottariello F, Costanzo C, Gotte G, Libonati M (2010) “Zero-length” dimers of ribonuclease A: further characterization and no evidence of cytotoxicity. Bioconjug Chem 21(4):635–645 Wang S, Li H (2006) Radical scavenging activity of ribonuclease inhibitor from cow placenta. Biochemistry (Moscow) 71:520–524 Wright PE, Dyson HJ (1999) Intrinsically unstructured proteins: re-assessing the protein structurefunction paradigm. J Mol Biol 293(2):321–331 Wright PE, Dyson HJ (2009) Linking folding and binding. Curr Opin Struct Biol 19(1):31–38 Wu Y, Mikulski SM, Ardelt W, Rybak SM, Youle RJ (1993) A cytotoxic ribonuclease. Study of the mechanism of onconase cytotoxicity. J Biol Chem 268(14):10686–10693

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Wu D, Yu W, Kishikawa H, Folkerth RD, Iafrate AJ, Shen Y, Xin W, Sims K, Hu GF (2007) Angiogenin loss-of-function mutations in amyotrophic lateral sclerosis. Ann Neurol 62 (6):609–617 Xu ZP, Tsuji T, Riordan JF, Hu GF (2002) The nuclear function of angiogenin in endothelial cells is related to rRNA production. Biochem Biophys Res Commun 294(2):287–292 Yamasaki S, Ivanov P, Hu GF, Anderson P (2009) Angiogenin cleaves tRNA and promotes stressinduced translational repression. J Cell Biol 185(1):35–42 Yang D, Rosenberg HF, Chen Q, Dyer KD, Kurosaka K, Oppenheim JJ (2003) Eosinophil-derived neurotoxin (EDN), an antimicrobial protein with chemotactic activities for dendritic cells. Blood 102(9):3396–3403 Yang D, Chen Q, Rosenberg HF, Rybak SM, Newton DL, Wang ZY, Fu Q, Tchernev VT, Wang M, Schweitzer B, Kingsmore SF, Patel DD, Oppenheim JJ, Howard OM (2004) Human ribonuclease a superfamily members, eosinophil-derived neurotoxin and pancreatic ribonuclease, induce dendritic cell maturation and activation. J Immunol 173(10):6134–6142 Youle RJ, D’Alessio G (1997) Antitumor RNases. In: D’Alessio G, Riordan JF (eds) Ribonucleases: structures and function. Academic Press, San Diego, pp 491–514 Zanfardino A, Pizzo E, Di Maro A, Varcamonti M, D’Alessio G (2010) The bactericidal action on Escherichia coli of zf-RNase-3 is triggered by the suicidal action of the bacterium OmpT protease. FEBS J 277(8):1921–1928 Zhang J, Dyer KD, Rosenberg HF (2000) Evolution of the rodent eosinophil-associated RNase gene family by rapid gene sorting and positive selection. Proc Natl Acad Sci USA 97(9): 4701–4706 Zhang J, Dyer KD, Rosenberg HF (2002) RNase 8, a novel RNase A superfamily ribonuclease expressed uniquely in placenta. Nucleic Acids Res 30(5):1169–1175 Zhang J, Dyer KD, Rosenberg HF (2003) Human RNase 7: a new cationic ribonuclease of the RNase A superfamily. Nucleic Acids Res 31(2):602–607 Zhao W, Beintema JJ, Hofsteenge J (1994) The amino acid sequence of iguana (Iguana iguana) pancreatic ribonuclease. Eur J Biochem 219(1–2):641–646 Zhao H, Ardelt B, Ardelt W, Shogen K, Darzynkiewicz Z (2008) The cytotoxic ribonuclease onconase targets RNA interference (siRNA). Cell Cycle 7(20):3258–3261 Zhou H-M, Strydom DJ (1993) The amino acid sequence of human ribonuclease 4, a highly conserved ribonuclease that cleaves specifically on the 30 -side of uridine. Eur J Biochem 217:401–409

Chapter 2

Vertebrate Secretory (RNase A) Ribonucleases and Host Defense Helene F. Rosenberg

Contents 2.1 2.2 2.3 2.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Brief History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Host Defense: A Working Definition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vertebrate Secretory (RNase A family) RNases and Host Defense: General Questions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 The Eosinophil RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.1 Eosinophil Cationic Protein (ECP/RNase 3): Mechanism of Action . . . . . . . . . . . 2.5.2 Evolution of Eosinophil Ribonucleases and the Rodent Eosinophil-Associated Ribonucleases (Ears) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.3 EDN/RNase, Ears, and Host Defense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 RNases 7 and 8 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6.1 Structure and Evolution of RNase 7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6.2 RNase 7: Molecular Basis of Antimicrobial Action . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Angiogenin/RNase 5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Avian Leukocyte RNase A-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Zebrafish and Salmon RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

36 36 37 39 39 42 42 43 44 44 45 45 46 46 48 48

Abstract Bovine pancreatic ribonuclease, also known as RNase A, is the prototype of an extensive, multi-lineage family of vertebrate secretory proteins that share elements of structure and catalytic activity despite substantial functional divergence. In this review, we feature the RNase A family and its members that are implicated in promoting host defense – activities that include sustaining mucosal

H.F. Rosenberg Laboratory of Allergic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892, USA e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_2, # Springer-Verlag Berlin Heidelberg 2011

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barriers, as well as participating in various aspects of innate and acquired immunity – and explore relationships linking gene evolution, enzymatic activity, and physiologic function. Keywords Leukocytes • Eosinophil • Pathogen • Bacteria • Respiratory viruses

2.1

Introduction

In this chapter, we will review the actions of specific vertebrate secretory (RNase A) ribonucleases in promoting host defense, which is a general term that covers multiple aspects of mucosal, innate and acquired immunity. The reader is referred to several of earlier reviews on this subject (Dyer and Rosenberg 2006; Pizzo and D’Alessio 2007; Rosenberg 2008a; Sorrentino 2010), and also two reviews that focus on a related topic, the eosinophilic leukocyte and its complex interactions with respiratory virus pathogens (Jacoby 2004; Rosenberg et al. 2009).

2.2

A Brief History

The RNase A family of vertebrate secretory ribonucleases has long intrigued the research community and as such, it has an important place in the history of modern biology. As described in detail elsewhere (Beintema and Kleineidam 1998; Marshall et al. 2008) and here in this volume (Chap. 1,15), many of the principles of protein structure, protein folding, and enzyme catalysis emerged from studies of bovine pancreatic ribonuclease, also known as RNase A, the prototype, and founding member of this group. For the purposes of this review, we pick up the story in the mid-1970s, as medical science began to take an interest in serum ribonuclease activity as a marker for neoplastic disease (Reddi and Holland 1976; Maor and Mardiney 1978). Although serum ribonuclease levels alone proved to be insufficiently specific for diagnostic purposes (Peterson 1979), these explorations provided the first indication that there might be more than one human secretory ribonuclease. The medical and basic science literature suggested that there were at least two secreted ribonucleases: an acid type (from leukocytes) and an alkaline type (from pancreas) defined by their distinct catalytic properties (Biswas and Hindocha 1974; Akagi et al. 1978). At the same time, Jaap Beintema and colleagues (Welling et al. 1975; Beintema et al. 1988) initiated their studies on ribonuclease evolution, which provided insight into the structural diversity of the numerous orthologs of pancreatic-type ribonucleases in species throughout the animal kingdom. Equally important were the ongoing political events of this era, as they ultimately served to direct the course of scientific inquiry. The U.S. National Cancer

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Vertebrate Secretory (RNase A) Ribonucleases and Host Defense

37

Act of 1971 provided the scientific establishment with a tremendous economic boost, which is largely credited with funding the nascent field of Molecular Biology. In this environment, vertebrate ribonuclease research took another giant step forward. Specifically, upon characterization of the amino acid and cDNA sequences for angiogenin (ANG), a protein from HT-29 adenocarcinoma cells known to promote blood vessel growth (Fett et al. 1985; Strydom et al. 1985; Kurachi et al. 1985; Moenner et al. 1994), it became clear that this protein shared sequence features with vertebrate secretory (RNase A) ribonucleases. Likewise, the amino terminal sequences and cDNA clones of eosinophil-derived neurotoxin (EDN) and eosinophil cationic protein (ECP), otherwise unremarkable secretory proteins from human eosinophilic leukocytes, indicated that these two proteins might likewise be part of this emerging RNase A gene family (Gleich et al. 1986; Rosenberg et al. 1989a, b; Hamann et al. 1989; Barker et al. 1989). With the completion of the human genome sequencing project in 2003, interrelationships among several of the distinct vertebrate RNase lineages were clarified. Interestingly, in the report describing the initial findings relating to the human genome sequence, Lander and colleagues (2001) remarked specifically on the unique nature of the RNase A ribonucleases as the only known family of vertebratespecific enzymes, and likewise concurred with our interpretations regarding rapid evolution and the generation of antimicrobial properties (Rosenberg et al. 1995; Zhang et al. 2000; Cho and Zhang 2007). As shown in Fig. 2.1, human RNase 1 is the direct ortholog of bovine RNase A. Also shown are other canonical ribonucleases, which are secretory proteins with classical signal sequences, specific disulfide-bonded tertiary structure, a His-Lys-His triad in a catalytic crevice, and some degree of enzymatic activity against a single-stranded RNA target, including EDN/RNase 2, ECP/RNase 3, RNase 4, ANG/RNase 5, RNase 6 and RNase 7. Recent findings from our laboratory have suggested that we may need to reconsider some of our earlier assumptions regarding RNase 8 primary structure (Chan, Moser et al., manuscript in preparation). In contrast, the known noncanonical ribonucleases (RNases 9–13) are sequences that are clearly related to this family, but with divergent features (insertions, deletions, mutations in critical regions) that indicate that they are unlikely to function as active enzymes.

2.3

Host Defense: A Working Definition

In its simplest form, host defense is the general term that encompasses all the components of the immune system, working in concert to provide a barrier against incoming pathogens. Among its component parts, this includes physical surfaces, such as the skin, and the respiratory, gastrointestinal, and urogenital epithelia. In addition to a barrier function, these tissues express and release antimicrobial proteins and have specialized detection mechanisms (e.g., toll-like receptors) that serve to identify pathogens and to prevent invasion. Host defense also includes the actions of the innate immune system, those of complement proteins, and leukocytes

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Fig. 2.1 Phylogenetic tree documenting relationships among various vertebrate secretory (RNase A) ribonucleases, including those described in greater detail in this review. Neighbor-joining tree constructed with encoded amino acid sequences (Poisson correction, 2000 bootstrap replicates) using algorithms within MEGA 4.0 (Tamura et al. 2007)

including mast cells, natural killer cells, granulocytes (neutrophils, eosinophils, basophils), and macrophages. Resident dendritic cells present pathogen-specific antigens and thus serve to link the innate immune system to the adaptive immune system, the latter including the actions of T and B lymphocytes that are thus influenced by antigen exposure.

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2.4

39

Vertebrate Secretory (RNase A family) RNases and Host Defense: General Questions

To date, vertebrate secretory ribonucleases have been identified as participants in host defense at the skin and mucosal surfaces, and as secretory components from innate immune cells (eosinophils, heterophils, monocytes, dendritic cells, mast cells). Interestingly, there are no reports of vertebrate secretory RNases as functional components of T cells or B cells, but this may reflect the fact that no one has actually looked carefully for this possibility. The current state of the art is summarized in Table 2.1. Although each situation presents its own unique issues, there are some general questions that have promoted inquiry and that continue to guide research in this field. Among these questions, those of us working in this field have asked, which vertebrate secretory ribonucleases promote host defense, and how specifically is this accomplished? What strategies are used and what targets are involved? Is enzymatic activity crucial or even necessary for promoting all or at least some host defense functions? Similarly, is the characteristic disulfide-bonded tertiary structure a crucial feature, or is it completely dispensable for this activity? Finally, can we determine which aspects are incidental findings and which are absolutely necessary for vertebrate survival in vivo?

2.5

The Eosinophil RNases

Eosinophils are tissue leukocytes that develop in the bone marrow and expand in number in response to Th2 cytokines (primarily interleukin-5), which are produced and accumulate during allergic states and parasitic infection in vivo (Fig. 2.2). Despite years of study, the precise role of eosinophils in host defense remains uncertain, as studies performed with cytokine- and eosinophil-deficient mice have cast significant doubt on the long-held belief that eosinophils promote host defense against helminthic parasites (Klion and Nutman 2004; Jacobsen et al. 2007; Fabre et al. 2009). Recently, Yousefi and colleagues (2008) have shown that hypereosinophilic mice are protected against bacterial infection, although the full relationship between bacteria and eosinophils remains to be explored. The complex evolutionary divergence between human and mouse eosinophils (reviewed in Rosenberg et al. 2007) precludes strong conclusions in many of these experimental trials. The major eosinophil granule-secretory proteins were isolated and described by Gleich and colleagues (1984) and Venge and colleagues (1987). Eosinophil cationic protein (ECP) was described as a small ~16 kDa, highly charged protein with cytotoxicity to bacteria, parasites, and mammalian cells in vitro. Eosinophilderived neurotoxin (EDN, also known as eosinophil-protein-X) had a similar amino acid content to ECP, but was less cationic and less toxic in general, although

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Table 2.1 Summary of host defense functions identified by vertebrate secretory RNase A family ribonucleases RNase A ribonuclease Species pI Host defense activities Chemoattractant for immature human dendritic cells (Yang et al. 2003) Enhances maturation of dendritic cells (Yang et al. 2004)a Reduces infectivity for RNA viruses (HRSV, Eosinophil-derived HIV) in tissue culture assays neurotoxin (Domachowske et al. 1998; Rugeles et al. (RNase 2) Human 9.2 2003)b Eosinophil cationic Cytotoxin for helminthes, strong bactericidal protein (RNase 3) Human 10.5 activity (reviewed in Boix et al. 2008) Produced by alveolar macrophages in response to Th2 cytokine stimulation (Cormier et al. 2002) Produced in response to respiratory virus infection, upregulated in the absence of Eosinophil-associated type I interferon signaling (Garvey et al. RNase 11 Mouse 9.3 2005) Eosinophil-associated Moderate bactericidal activity against E. coli RNases 1 and 2 Rat 9.0, 9.9 and S. aureus (Ishihara et al. 2003) Moderate bactericidal activity against S. pneumoniae (Hooper et al. 2003); Angiogenin (RNase 5) Human 9.7 conflicting report (Avdeeva et al. 2006) Moderate bactericidal activity against L. monocytogenes and E. faecalis (Hooper et al. 2003) Produced in response to lipopolysaccharide (Hooper et al. 2003) Angiogenin-4 Mouse 9.2 Broad-spectrum strong bactericidal activity, very strong against E. faecium (Harder and Schroeder 2002) RNase 7 Human 9.8 Broad-spectrum moderate bactericidal RNase 8 Human 8.6 activity (Rudolph et al. 2006) Bactericidal against E. coli, S. aureus (Nitto et al. 2006) and Salmonella sps. (unpublished data) Leukocyte RNase A-2 Chicken 10.4 Broad-spectrum moderate bactericidal activity against gram-negative bacteria ZF-RNases 1–5 SSZebrafish 9.0–9.4 (Pizzo et al. 2006, 2008; Cho and Zhang RNases 1 and 2 salmon 9.3, 8.5 2007) Values for isoelectric points (pI) were calculated using the web-based ExPaSy tool (http://expasy. org/tools/pi_tool.html) using encoded amino acid sequence data. Moderate bactericidal activity, low micromolar concentrations reduce the colony count 10–100-fold in a standard overnight incubation assay; strong bactericidal activity, low micromolar concentrations reduce colony count 104–107-fold. Reprinted with modifications from Rosenberg (2008a)aActivity shared with human pancreatic ribonuclease, also known as RNase 1bActivity dependent on enzymatic (ribonuclease) activity

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41

Fig. 2.2 (a) Isolated human eosinophils stained with modified Giemsa (Diff Quik). The cells feature characteristic bilobed nucleus and the prominent cytoplasmic granules that contain the secretory ribonucleases, EDN/RNase 2 and ECP/RNase 3. (b) Recombinant EDN reduces the infectivity of the human respiratory syncytial virus (RSV-B) for target epithelial cells in a dosedependent fashion. Site-specific mutagenesis resulting in elimination of the catalytic lysine restores virus infectivity, consistent with ribonuclease-dependent antiviral activity. (Reprinted from Domachowske et al. 1998)

it promoted destruction of Purkinje cells when administered intrathecally to rodents and rabbits (Durack et al. 1981). Gleich and colleagues (Gleich et al. 1986; Slifman et al. 1986) were the first to recognize the homology between the amino terminal sequences of both EDN and ECP and human ribonuclease, and to document enzymatic activity; identification of cDNA clones encoding both proteins documented complete sequence homology to RNase A (Rosenberg et al. 1989a, b; Hamann et al. 1989; Barker et al. 1989).

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2.5.1

H.F. Rosenberg

Eosinophil Cationic Protein (ECP/RNase 3): Mechanism of Action

The earliest studies of ECP focused on its cationicity and its ability to disrupt membrane function (Young et al. 1986). However, once it became clear that ECP had dual potential, the role of ribonuclease activity and its relationship to the cytotoxic nature of ECP was addressed. Interestingly, in experiments performed with active site histidine and lysine mutations, it became quite clear that enzymatic activity played no significant role in the bactericidal activity promoted by ECP (Rosenberg 1995). This finding has been confirmed and extended many times over both directly and indirectly, as Torrent, Boix, Nogues and colleagues (Carreras et al. 2003; Torrent et al. 2007, 2008, 2009a, b, 2010a, b; Navarro et al. 2008; Boix et al. 2008; Garcia-Mayoral et al. 2010; Sanchez et al. 2010) examined the structure– function relationships of ECP with target pathogens and pathogenmimetic membranes. ECP and ECP-derived minimal cationic peptides promote cytotoxicity via interactions with target bacterial membrane and wall components, including peptidoglycan and lipopolysaccharides. Interestingly, ECP-mediated interactions elicit cytotoxicity via membrane aggregation more prominently than via cellular lysis. This finding – the emergence of ECP/RNase 3 as a cationic toxin with activity that is not dependent on ribonuclease activity – seems to be counterintuitive from an evolutionary perspective. However, this can be understood by applying the principles first introduced by the evolutionary biologist, Susumo Ohno (1970), specifically, that novel function can evolve by duplication of genetic material followed by relaxation of functional constraints. As shown in Fig. 2.1, ECP and EDN are paired genes, having undergone duplication from a single predecessor gene over 60 million years ago (described further in Sect. 2.5.2). As such, one might envision a case such that ECP might gain novel function (i.e., cationic cytotoxin) while EDN maintained the original enzymatic activity.

2.5.2

Evolution of Eosinophil Ribonucleases and the Rodent Eosinophil-Associated Ribonucleases (Ears)

As part of an attempt to obtain a better sense of the relationships connecting enzymatic activity, cationicity, and host defense, we proceeded to explore the evolutionary relationships among these proteins. In doing so, we uncovered the remarkable interspecies divergence within the EDN/ECP lineage, now known to be among the most rapidly evolving coding sequences among primate species (Rosenberg et al. 1995; Rosenberg and Dyer 1995; Zhang et al. 1998). All primate genomes encode a highly cationic ECP and a more neutral EDN, save for the new World monkey genomes, which maintain a single sequence, with properties similar to EDN, as predicted above (Sect. 2.5.1). Meanwhile, Larson and colleagues (1996)

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43

identified the first mouse orthologs of this lineage, a highly divergent cluster of eosinophil-associated ribonucleases, or Ears. Zhang and colleagues (2000) documented that the rodent Ears, similar to the human eosinophil ribonucleases, are undergoing rapid evolution and are diverging under unique constraints, via a pattern known as rapid-birth–death and gene sorting.

2.5.3

EDN/RNase, Ears, and Host Defense

Despite the rapid evolution, and clearly unusual constraints to which this gene lineage is responding, the EDN/RNase 2 genes maintain features consistent with their role as active enzymes. Taking into account (a) the role of eosinophils in promoting respiratory pathology in asthma, (b) the fact that RNA viruses such as respiratory syncytial virus (RSV) commonly incite asthmatic sequelae in susceptible individuals, we closed the circle and (c) considered the possibility that single-stranded RNA virus pathogens might be enzymatically susceptible, evolutionarily mobile targets of a ribonuclease such as EDN/RNase 2. To date, we have shown that EDN can reduce infectivity of RSV for target epithelial cells in culture, via a mechanism that is directly dependent on active ribonuclease activity (Domachowske et al. 1998; Rosenberg 2008b). Interestingly, EDN can also reduce infectivity of human immunodeficiency virus (HIV), a single-stranded RNA virus of the family Retroviridae, in similar in vitro culture assays (Rugeles et al. 2003; Bedoya et al. 2006). The precise mechanism of antiviral activity awaits further exploration. EDN has also been implicated in several other aspects of innate immunity. Yang and colleagues (Yang et al. 2003, 2004) reported that EDN elicited chemotaxis and enhanced maturation of human dendritic cells. In further studies, this group (Yang et al. 2008) examined the interaction of EDN with TLR2, and suggested that EDN might serve as an alarmin, one of a group of endogenous immunostimulant molecules that serve as signals of tissue damage (Oppenheim and Yang 2005). Rather little is known regarding the rodent Ears, their unique expression patterns, and their individual and/or collective roles in host defense. There are ~14 mEar coding sequences in the mouse genome; mEar 1 and mEar 2 are expressed most prominently in mouse eosinophils and in lung tissue (Moreau et al. 2003). Expression of mEar 11 was induced in mouse alveolar macrophages in response to Th2 cytokine stimulation (Cormier et al. 2002) and in virus-infected lung tissue in the absence of the type I IFN receptor (Garvey et al. 2005); mEar 6 was detected in liver in response to Schistosoma mansoni infection (Nitto et al. 2004). Likewise, Ishihara and colleagues (2003) demonstrated antimicrobial activity for rat Ears in standard in vitro assays, and Gaudreault and Gosselin (2007, 2008) detected release of mEars in virus-infected mouse lung tissue in response to leukotriene-B4.

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2.6

H.F. Rosenberg

RNases 7 and 8

Ribonuclease 7 (RNase 7) is another intriguing vertebrate secretory ribonuclease with unique structure and antimicrobial properties. Although RNase 7 was discovered after the emergence of the RNase A family, similar to ECP, it was identified initially as a host defense protein, emerging from a study by Harder and Schroeder (2002) who were evaluating healthy keratinocyte cultures for novel anti-pathogen molecules. In a parallel study, Zhang and colleagues (2003) identified RNase 7 as a novel RNase A family member from the first draft of the human genome sequence. RNase 7 is a cationic ribonuclease with wide-spectrum antimicrobial activity, particularly powerful against gram-negative bacteria including Pseudomonas and Enterococcus species (Harder and Schroeder 2002; K€ oten et al. 2009). Recently, Zanger and colleagues (2009) and Simanski and colleagues (2010) demonstrated that RNase 7 protects healthy skin against infection and colonization, respectively with Staphylococcus aureus. RNase 7 is expressed widely, and is induced in keratinocyte culture by prominent proinflammatory stimuli, including TNF, IFNg, IL-1b, and IL-1a (Harder and Schroeder 2002; Mohammed et al. 2010; Bando et al. 2007) as well as UVirradiation (Gl€aser et al. 2009) and bacterial components (Harder and Schroeder 2002). Ribonuclease 8 (RNase 8) is closely related to RNase 7. RNase 8 appears to have undergone “functional” pseudogenization in several primate species (i.e., several orthologs have mutations in elements that are crucial for enzymatic activity), although further evaluation of more samples is warranted prior to reaching a final conclusion on this subject. Although Zhang and colleagues (2002) reported no cytotoxicity in the bacterial cells synthesizing the recombinant protein, Rudolph and colleagues (2006) reported that recombinant RNase 8, modeled on the structure of RNase 7, displays moderate antimicrobial activity against several staphylococcal, enterococcal, and E. coli strains.

2.6.1

Structure and Evolution of RNase 7

RNase 7 is fairly typical for a member of the RNase A family. Its open reading frame encodes a 28-amino acid signal sequence, and the mature protein coding sequence includes eight cysteines, and appropriately localized histidines and lysine, the latter within the family signature motif. Human RNase 7 is relatively cationic, with a calculated isoelectric point (pI) of 9.8, and a predominance of lysines, rather than arginines, as was observed in the coding sequence of ECP/RNase 3. Compared to EDN/ECP, the divergence among primate RNase 7 genes is comparatively modest, with 23% sequence divergence observed in a comparison between human and common marmoset (New World monkey) C. jacchus RNase 7 sequences. Two RNase 7 genes have been identified in the genomes of the horse (E. caballus) and cow (B. taurus), at 30% and 35% sequence divergence from the human sequence, respectively; no orthologs of RNase 7 have been found in the mouse genome.

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2.6.2

45

RNase 7: Molecular Basis of Antimicrobial Action

Although RNase 7 shares some features with ECP/RNase 3 (structure, cationicity, ribonuclease activity), the molecular nature of its antimicrobial activity is unique and distinct. Similar to what has been observed for ECP, ribonuclease activity is not essential to RNase 7’s mechanism of antimicrobial action. Torrent and colleagues (2009b, 2010a) have compared the actions of ECP and RNase 7 using membrane phospholipid vesicles and bacterial cell walls. They have noted that ECP has a much larger tendency to induce aggregation of both targets, whereas RNase 7 results in vesicle leakage and release of bacterial cell contents without inducing substantial aggregation. Liao and colleagues (Huang et al. 2007; Lin et al. 2010) focused on the unique features of RNase 7, and concluded that the amino terminal lysines were crucial features promoting antimicrobial activity; an outer membrane protein of P. aeruginosa has been identified as a target for RNase 7/cell surface interactions preceding interaction with lipopolysaccharide and ultimately internalization.

2.7

Angiogenin/RNase 5

Angiogenin is best known for its role in promoting blood vessel growth (reviewed in Strydom 1998; Badet 1999); recently, polymorphisms in genes encoding angiogenin have been linked to susceptibility for developing amyotrophic lateral sclerosis (Bosco and Landers 2010). The human genome encodes only one functional angiogenin gene, while the mouse genome encodes six. Hooper and colleagues (2003) described an intriguing host defense function for angiogenin 4 (Ang 4), a member of the mouse cluster expressed in intestinal Paneth cells. Ang 4 expression is induced by normal microflora in conventionally raised mice, but interestingly, Ang4 is not expressed in germ-free mice, and can be induced in the latter by the introduction of the microflora component, B. thetaiotaomicron. Furthermore, human ANG, its mouse ortholog Ang1, and mouse Ang 4, all displayed antimicrobial activity at micromolar concentrations. While Avdeeva and colleagues (2006) presented a conflicting view, noting that human ANG was no more effective than control protein (bovine serum albumin) at inhibiting pathogen growth, it is crucial to recognize that one of the pathogens examined in this latter study, S. pneumoniae, is difficult to grow and quite sensitive to minimal levels of detergent contaminants. Since this first report, Lagishetty and colleagues (2010) presented their study of mice raised on vitamin-D deficient diets that were then induced to develop colitis. Interestingly, mouse Ang 4 expression was diminished more than twofold, a finding that correlated with a 50-fold elevation of bacteria in colonic tissue.

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2.8

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Avian Leukocyte RNase A-2

There are three RNase A family sequences in the genome of the chicken, Gallus gallus; two of these sequences are duplicated leukocyte-associated ribonuclease genes, which are renamed leukocyte RNase A-1 and A-2 (Nitto et al. 2006). Interestingly, both RNase A-1 and RNase A-2 are cationic (isoelectric points 9.8 and 10.4, respectively), both are angiogenic in an aortic ring assay (RNase A-2 > RNase A-1), and both immunoreactive proteins were found in both bone marrow progenitors and in circulating heterophils. In contrast, (and similar to the EDN/ECP pair) RNase A-1 was a more effective RNase against the standard tRNA substrate, while RNase A-2 was the powerful antimicrobial protein (Fig. 2.3). Consistent with earlier studies focusing on ECP, site-specific mutagenesis that eliminated the ribonuclease activity of RNase A-2 had no impact on the antimicrobial activity. Most intriguing was the fact that the unique tertiary structure likewise seemed to have no specific impact on antimicrobial function, as cationic domains within the structure of RNase A-2 were able to function as independent antimicrobial peptides. Specifically, domain III, a 16-amino acid peptide including amino acids 89–104 of the native protein reduced the bacterial colony count 105-fold when introduced at micromolar concentrations, matching the effectiveness of the full, native protein. These results suggest that not only is enzymatic activity dispensable, but the unique disulfide-bonded RNase A backbone may not be a crucial limiting constraint driving the evolution of this gene family.

2.9

Zebrafish and Salmon RNases

RNase A family genes from the zebrafish (Danio rerio) and Atlantic salmon (Salmo salar) have been identified and recombinant proteins generated and explored for antimicrobial activity. The salmon RNases 1 and 2 displayed moderate antibacterial activity at micromolar concentrations against both gram-negative and grampositive bacteria (Pizzo et al. 2008); analogous to ECP and leukocyte RNase A-2, ribonuclease activity is not essential, nor is tertiary structure, as fully denatured protein is as effective as folded ribonuclease at promoting anti-pathogen activity in tissue culture. Ribonucleases from zebrafish were isolated and characterized by D’Alessio and colleagues (Pizzo et al. 2006), Quarto et al. (2008), Kazakou and colleagues (2008), and Cho and Zhang (2007). Recombinant zebrafish ribonuclease proteins also display moderate antimicrobial activity against both Gram-negative and Grampositive bacteria; the detection of antimicrobial activity in among these proteins led Cho and Zhang (2007) to postulate that host defense was not only present but likely a primordial function of the RNase A gene family. Zanfardino and colleagues (2010) recently examined a molecular mechanism for the antimicrobial action of ZF-RNase 3, a zebrafish ribonuclease expressed in lung and gut tissue. As is typical

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47

Fig. 2.3 (a) Detection of leukocyte RNase A-1 and A-2 in heterophils from bone marrow from chicken (Gallus gallus). (b) Challenge with low micromolar concentrations of leukocyte RNase A-2 (but not leukocyte RNase A-1) results in a 107-fold reduction in E. coli colony count. (c) Similar concentrations of unstructured domain III peptide, corresponding to amino acids 89–104 of RNase A-2, are nearly as effective at reducing colony counts of E. coli. (Reprinted from Nitto et al. 2006)

among ribonucleases of this family, the antimicrobial activity of ZF-RNase 3 was not dependent on enzymatic activity, nor was it dependent on tertiary structure. ZF-RNase 3 itself is cleaved at Arg30-Arg31 by the E. coli OmpT protease, thereby releasing an antimicrobially active C-terminal proteolytic fragment. Likewise, Pizzo and colleagues (2010) characterized ZF-RNase 5, which has activity against Gram-negative, but not Gram-positive bacterial species.

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Of note, Balla and colleagues (2010) recently reported the isolation and characterization of zebrafish eosinophils, which are morphologically and functionally analogous to their mammalian counterparts. Although the zebrafish ribonuclease lineages are not directly related to the mammalian eosinophil ribonucleases, zebrafish eosinophils express transcripts encoding ZF-RNase 2, an antimicrobial protein with moderate activity against E. coli and P. aeruginosa in vitro. ZF-RNae 3 is expressed in liver, gut, and heart, and has relatively little enzymatic activity when compared to ZF/DR-RNases 1 and 3. Given the relative ease with which gene manipulations are carried out in zebrafish, this may represent the most powerful resource available for exploring the role of ribonucleases in promoting host defense in vivo.

2.10

Conclusions

Vertebrate secretory ribonucleases have long been intriguing subjects for the study of protein structure and enzyme biochemistry. We proceed to build on this profound body of knowledge which provides the basis for our study of the contribution of the proteins to mucosal, innate and acquired immune function. Acknowledgments Ongoing work in our laboratory is supported by funds from the NIAID Division of Intramural Research, Project AI000942.

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.

Chapter 3

Antitumor Ribonucleases Marc Ribo´, Antoni Benito, and Maria Vilanova

Contents 3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Natural Cytotoxic Ribonucleases and Their Mechanism of Action . . . . . . . . . . . . . . . . . . . . . . 3.2.1 Amphibian RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2 Bovine Seminal Ribonuclease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.3 Eosinophil Cationic Protein (ECP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.4 Ribonucleases of Different Origins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Approaches to Endow a Ribonuclease with Cytotoxic Properties . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1 Monomeric Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.2 Oligomeric and Tandem Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.3 Targeted Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Clinical Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

56 58 58 61 63 64 64 65 67 68 73 75 75

Abstract Ribonucleases are small basic proteins that have shown remarkable antitumor activity linked to their ability to destroy RNA. Therefore, they are a second line of cancer chemotherapeutics as they are not genotoxic. This chapter summarizes the main biochemical characteristics of these enzymes and the key factors responsible for their cytotoxic mechanism. Some of them are shared by most cytotoxins, but each RNase has particular cancer cell killing abilities. The effects on the cell cycle and the induced apoptosis mechanism are cell dependent. The knowledge obtained from the cytotoxic mechanism of natural cytotoxic RNases has been used to artificially engineer more potent and selective RNA-degrading

M. Ribo´ (*) • A. Benito (*) • M. Vilanova (*) Laboratori d’Enginyeria de Proteı¨nes, Departament de Biologia, Facultat de Cie`ncies, Universitat de Girona, Campus de Montilivi, Maria Aure`lia Capmany, 69, 17071 Girona, Spain Institut d’Investigacio´ Biome`dica de Girona Josep Trueta (IdIBGi), Girona, Spain e-mail: [email protected]; [email protected]; [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_3, # Springer-Verlag Berlin Heidelberg 2011

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56

enzymes. These approaches are also described. The chapter ends with a brief description of the results of the clinical trials performed with RNases.

3.1

Introduction

The expression of specific genes can be controlled at both the DNA and RNA levels (Fig. 3.1). The development of drugs with the ability to target nucleic acids (DNA, RNA) for degradation is a powerful way to control gene expression. In recent decades, this approach, among others, has been used to treat diseases and in particular cancer. However, drugs acting at the DNA level may have the disadvantage of being mutagenic (for a review, see Gurova 2009). On the contrary, destroying RNA may have similar effectiveness leaving DNA undamaged. There is a vast array of current available technologies for the destruction of RNA with therapeutic potential (for a review, see Tafech et al. 2006). The use of enzymes with ribonucleolytic activity is one of them. Until recently, the efficient turnover of RNA molecules in higher eukaryotes was thought to occur mainly through exonucleolytic activity. This view has been challenged by recent work on endoribonucleases (for a review, see Li et al. 2010). It is now possible to say that the endonucleolytic cleavage of RNA, as opposed to exonucleolytic decay, probably plays a larger role in RNA metabolism than has been previously imagined. Pancreatic ribonucleases (RNases) are endoribonucleases (Cuchillo et al. 1997), so their ability to cleave RNA internally and without high sequence specificity makes them exceptional candidates for antitumor drugs provided that they can be selectively and efficiently delivered to the cells. In fact, in a recent volume of the European Journal of Pharmacology dedicated to novel anticancer strategies, the introductory review (Los 2009) includes a member of the pancreatic RNase family, Onconase (ONC) (Ardelt et al. 1991), among the new, exciting developments in experimental

Fig. 3.1 Flow chart for transmission of biological information. The therapeutic approach of RNases is to destroy RNA; therefore, they are non-genotoxic antitumor drugs like conventional small anticancer drugs or radiotherapy

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therapies of the early twenty-first century. Bovine pancreatic ribonuclease A (RNase A, EC 3.1.27.5), the most well-known member of the RNase pancreatic family, both structurally and functionally (for a review, see Cuchillo et al. 1997; Raines 1998), was the first RNase to be tested in vitro (Ledoux and Baltus 1954; Ledoux and Revell 1955; Ledoux 1956) and in vivo (Ledoux 1955a, b; Aleksandrowicz 1958; Telford et al. 1959) as an anticancer drug. However, contradictory results were obtained (Roth 1963) and RNases were not considered as potential antitumor agents until the discovery of natural RNases with anticancer activities exhibited at much lower concentrations than RNase A. Cytotoxic RNases are not restricted to the pancreatic RNase family. Prokaryotic and eukaryotic microbial RNases (Irie 1997; Yoshida 2001) as well as plant RNases (Matousek 2010) are also cytotoxic enzymes. Although a significant number of studies have been carried out to elucidate the cytotoxic mechanism of antitumor RNases, the molecular basis is still not well understood. Nevertheless, there is a general consensus about their mechanism of action (for a review, see Makarov and Ilinskaya 2003; Benito et al. 2005; Makarov et al. 2008), which, in some parts, is shared by most cytotoxins. Briefly, to act as a cytotoxin, an RNase should follow the following steps (Fig. 3.2). It has to interact with a specific or a nonspecific component of the target cell surface in order to be

Fig. 3.2 Productive and nonproductive pathways of cytotoxic RNases. RNases interact with the cell surface nonspecifically or through a receptor either naturally or targeted by binding the RNase to a ligand (1) and they are endocytosed (2). From one of the endocytic vesicles, the RNase translocates to the cytosol (3) evading the lysosomal degradation. This process can be enhanced by RNase modification. Once in the cytosol, the RNase must retain its stability, and one of two conditions is necessary for it to be cytotoxic. To meet the first condition, it has to be, either naturally or artificially, resistant to the RI or, provided that enough molecules reach the cytosol, it has to saturate it. In this way, the RNase activity degrades cellular RNA and induces apoptosis (4). Otherwise, if the RNase is captured by the RI, it loses its ribonucleolytic activity leaving the RNA undamaged (5). To meet the second condition, it has to be sequestered by other molecules that hamper its interaction with the RI, driving it to an RNA-containing organelle free of RI like the cell nucleus (6). In this latter case, a-importin will sequester and release the RNase into the nucleus removing it from the RI-RNase and a-importin-RNase competing equilibriums and ultimately inducing cell death

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endocytosed, then to follow a productive intracellular route through the different membranous subcellular organelles. From one of these organelles, it has to translocate to reach the cytosol. Once there, it has to be stable enough and keep its ribonucleolytic activity in order to degrade RNA and induce cell death. In the cytosol, there are three different ways for an RNase to preserve its ability to cleave RNA: first, to be insensitive to the ribonuclease inhibitor protein (RI) present in the cytosol of all mammalian cells (Dickson et al. 2005); second, to saturate the RI, meaning that enough protein reaches the cytosol to leave free RNase molecules (Leich et al. 2007); and third, to be captured by other molecules, which impairs the interaction with the RI (Bosch et al. 2004). A walk through the antitumor members of the pancreatic RNases will unveil what is known about its cytotoxic mechanism.

3.2

3.2.1

Natural Cytotoxic Ribonucleases and Their Mechanism of Action Amphibian RNases

Onconase (ONC) and Amphinase (Amph) are two homologous RNases first isolated by Alfacell Corporation from Rana pipiens (leopard frog). The former was isolated two decades ago (Ardelt et al. 1991) and the latter much more recently (Singh et al. 2007). Their structure and function and their cytostatic and cytotoxic mechanisms, either alone or together with other cytotoxic RNases, have been the subject of several reviews (Youle and D’Alessio 1997; Leland and Raines 2001; Matousek 2001; Makarov and Ilinskaya 2003; Saxena et al. 2003; Benito et al. 2005, 2008b; Arnold and Ulbrich-Hofmann 2006; Ramos-Nino 2007; Ardelt et al. 2008, 2009; Arnold 2008; Lee and Raines 2008; Lu et al. 2008; Altomare et al. 2010). Although the main characteristic of these RNases will be briefly summarized, readers should refer to the reviews mentioned above for more detailed information. ONC has about 30% sequence identity with RNase A (Ardelt et al. 1991) and a similar fold (V or kidney-shaped), conserves three of the four disulfide bonds of RNase A, and is more compact. The N- and C-termini are blocked by pyroglutamic acid and a disulfide bond respectively (Mosimann et al. 1994). Its compact structure makes it a very stable protein and resistant to proteolysis (Leland et al. 2000; Notomista et al. 2000, 2001; Arnold et al. 2006; Schulenburg et al. 2010). Folding studies provide further explanation of this high conformational stability (Pradeep et al. 2006; Gahl et al. 2008; Gahl and Scheraga 2009). ONC is 102–105-fold less active against polymeric substrates and single-stranded RNA than RNase A (Ardelt et al. 1991; Boix et al. 1996) although it has the same key catalytic residues present in RNase A (Lee and Raines 2003). It possesses two additional active-s ite residues, Lys9 and an N-terminal pyroglutamic residue (Lee and Raines

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2003). The cytotoxic activity depends on its ability to degrade RNA (Ardelt et al. 1991; Wu et al. 1993). Its substrate specificity is different from that of RNase A (Singh et al. 2007) and depends on the structure (Lee et al. 2008). ONC evades the RI (for a review see Rutkoski and Raines 2008), a property not shared by the monomeric members of the pancreatic family, but which is critical for cytotoxicity. The dissociation constant of the complex formed between ONC and RI is about 107-fold greater than that of RNase A (Boix et al. 1996). This lower affinity has been attributed to a reduction of the length of some exposed loops responsible for the interaction with the RI (Mosimann et al. 1994; Kobe and Deisenhofer 1996), which precludes the complex formation at physiological salt concentration (Turcotte and Raines 2008). This in vivo evasion is supported by the absence of any effect on ONC cytotoxicity due to either increasing (Haigis et al. 2003) or silencing (Monti and D’Alessio 2004; Dickson and Raines 2009) the intracellular levels of RI. Nevertheless, ONC presents a disulfide bond (30–75), which in the reducing conditions of the cytosol could be cleaved, affecting the ONC-RI interaction (Torrent et al. 2008). ONC internalization is not completely understood (for a review, see Benito et al. 2008b). Early studies (Wu et al. 1993, 1995) and more recent ones (Haigis and Raines 2003) indicate that ONC enters the cell by energy-dependent endocytosis. Initial uptake takes place through the well-characterized clathrin/AP-2-mediated endocytic pathway (Rodriguez et al. 2007) although a previous study indicated that the entry was not dynamin dependent (Haigis and Raines 2003). This apparent contradiction might be related to the use of transiently or stably transfected cell lines expressing a dynamin-K44A dominant-negative mutant. The first evidence of the presence of a cell receptor was proposed by Wu et al. (1993). However, dosedependent internalization of Oregon-green-labeled D16C-ONC in HeLa cells showed a non-saturable process (Haigis and Raines 2003). It has also been reported that, due to its cationic nature, ONC and, in general, RNases bind to cell surfaces electrostatically, indicating that endocytosis is not mediated by a receptor (Notomista et al. 2006; Johnson et al. 2007a). Since the surface of most cancer cells is more electronegative compared to normal cells (Ran et al. 2002), this electrostatic interaction could explain their selectivity. It has recently been shown (Chao et al. 2010) that ONC interacts with membrane glycoproteins that do not seem to mediate a productive internalization. No correlation was apparent in the cell surface binding, internalization, and cytotoxicity of ONC. Nevertheless, the distribution of cationic residues on the surface of ONC might be responsible for an efficient translocation process, once endocytosed (Turcotte et al. 2009). One major question is from which endocytic vesicle this translocation is produced. The intracellular routing of ONC has been studied using drugs that disrupt intracellular trafficking (Wu et al. 1993, 1995; Newton et al. 1998; Haigis and Raines 2003) and by co-localization studies of fluorescent-labeled ONC with markers of different organelles (Rodriguez et al. 2007). The results indicate that the normal route for ONC to reach the cytosol is through an endosomal compartment, not from the Golgi or endoplasmic reticulum (ER). ONC is localized in the recycling endosomes from

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where it translocates, again suggesting the existence of a cellular receptor (Rodriguez et al. 2007). Once in the cytosol, ONC degrades RNA and precludes protein synthesis, inducing cell cycle arrest (initially described at the G1/S checkpoint later found to be cell dependent) and apoptosis (Darzynkiewicz et al. 1988; Deptala et al. 1998; Juan et al. 1998; Smith et al. 1999; Halicka et al. 2000; Iordanov et al. 2000b; Grabarek et al. 2002; Tsai et al. 2004). Early in vitro studies identified the rRNAs (28 S and 18 S) as the ONC targets (Wu et al. 1993), but now it is known that the tRNAs are the primary in vivo target of this enzyme (Lin et al. 1994; Iordanov et al. 2000a; Suhasini and Sirdeshmukh 2006, 2007). However, ONC-induced apoptosis presents features different from those of an indiscriminate translation inhibition (Iordanov et al. 2000b). Indeed, the upregulation or downregulation of genes that code for proteins involved in cell cycle control, or transcription factors, is observed after ONC cell treatment (Juan et al. 1998; Tsai et al. 2004; Altomare et al. 2010). Accordingly, two other targets for ONC have been proposed: the mRNA coding for the ubiquitous NF-kB transcription factor either directly or by degrading other RNAs involved in its turnover (Deptala et al. 1998; Juan et al. 1998; Tsai et al. 2004) and the non-coding RNAs (microRNAs and/or siRNAs) involved in gene expression control (Ardelt et al. 2003; Zhao et al. 2008). In support of these hypotheses, Saxena et al. (2009) have reported the degradation of double-stranded RNA by ONC. ONC-induced apoptotic effects are very dependent on the cell type. Thus, some apparently contradictory results are found in the literature. The activation of the stress-activated protein kinase c-Jun NH2 terminal kinase was described as an early event in the induction of apoptosis by ONC (Iordanov et al. 2000a). This process seems to promote the activation of pro-caspase-9, 3 and 7, but in the study, the involvement of the mitochondrial pathway was not clear (Iordanov et al. 2000a). Furthermore, in neuroblastoma cells, ONC induces autophagy by lysosomal activation, which leads to caspase-independent apoptosis (Michaelis et al. 2007). However, the typical mechanisms of the intrinsic apoptosis pathway, in addition to an activation of Ser-proteases, were observed in other cell lines (Grabarek et al. 2002; Ardelt et al. 2007b). Recently, it has been demonstrated that ONC-induced apoptosis is dependent on Apaf-1 and that ONC enhances apoptosis by reversing the inhibitory effect of the tRNAs on cytochrome c (Mei et al. 2010). These results could explain the synergistic effects with drugs that elicit the intrinsic apoptosis pathway. In addition, since tRNA expression is enhanced in tumor cells, its degradation could provide an explanation for ONC selectivity (Mei et al. 2010). The Amph variants present most of the characteristics of ONC; however, their catalytic efficiency, their substrate specificity, the N-terminal end, and the glycosylation state are different (Singh et al. 2007; Ardelt et al. 2008). All the variants present cytostatic and cytotoxic activity patterns that, to a certain extent, resemble those of ONC (Ardelt et al. 2007b). A recombinant form of an Amph that is as active as the natural enzyme suggests that the glycans are not involved in the cytotoxicity (Singh et al. 2007).

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Tumoricidal ribonucleases have been also found in Rana catesbeiana (RCRNase) and Rana japonica (RJ-RNase) oocytes. They present about 50% sequence identity with ONC, high stability, the same number of disulfide bonds, and both are pyrimidine base-specific RNases and do not present detectable carbohydrate moieties (for a review, see Youle and D’Alessio 1997; Irie et al. 1998). The structure is only known for RC-RNase and resembles that of ONC (Chang et al. 1998). Both are sialic acid–binding lectins that specifically agglutinate cancer cells (Sakakibara et al. 1979; Okabe et al. 1991; Nitta et al. 1994) binding to cell membrane glycoproteins with a high content of sialic acids (Sakakibara et al. 1979; Nitta et al. 1994). By contrast, ONC has been reported not to cause tumor cell agglutination (Ardelt et al. 1991) and shows minimal interaction with cell surface GAG and sialic acid–containing proteins (Chao et al. 2010). Preclinical studies not directly related to the elucidation of cytotoxic mechanisms have been carried out with amphibian RNases. ONC presents synergy, proved in vitro and/or in vivo, with a significant number of compounds including: tamoxifen (Mikulski et al. 1990a, 1992a), lovastatin (Mikulski et al. 1992b), cisplatin (Mikulski et al. 1992a; Lee et al. 2007a), AEBS/HIC-binding drugs (Mikulski et al. 1993a), vincristine, which was independent of P-gp expression (Rybak et al. 1996), proteasome inhibitors (Mikulski et al. 1998), interferons (Deptala et al. 1998; Vasandani et al. 1999b), doxorubicin (Mikulski et al. 1999), small molecule inhibitors of PI3-K (Ramos-Nino et al. 2005), cepharanthine (Ita et al. 2008), and rosiglitazone (Ramos-Nino and Littenberg 2008). Interestingly, ONC-induced apoptosis is independent of the p53 tumor-suppressor protein status (Iordanov et al. 2000b). In addition, ONC decreases the content of reactive oxygen intermediates (ROI), which may be an important element of its cytotoxicity toward cancer cells (Ardelt et al. 2007a). In vivo studies, using animal models, proved the efficacy of ONC in tumor suppression and prolonged survival (Mikulski et al. 1990b; Lee et al. 2000a; Lee and Shogen 2008). In vitro, ONC induces cellular radiation sensitivity by inhibiting cellular oxygen consumption (Lee et al. 2000b). In vivo, it changes tumor physiological parameters, significantly increasing tumor oxygenation (Lee et al. 2003). Because of that, ONC acts as a radiation sensitizer (Kim et al. 2007; Lee et al. 2007b). Apart from the positive effects, the main drawback of ONC is longer retention in the kidneys, which promotes nephrotoxicity, although it is reversed after drug withdrawal (Vasandani et al. 1996, 1999a). In addition, in vitro and in vivo assays have shown that ONC has stronger aspermatogenic, embryotoxic, and immunotoxic activity than other pancreatic RNases (Matousek et al. 2003) and that it also displays significant neurotoxicity when injected intracranially (Slager et al. 2009).

3.2.2

Bovine Seminal Ribonuclease

Bovine seminal ribonuclease (BS-RNase) is the only natural dimeric member of the pancreatic-type RNases. The enzyme was first discovered in 1963 in bull seminal

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plasma (for a review, see D’Alessio et al. 1991, 1997). Each subunit presents 83% amino acid sequence identity to RNase A and a similar fold. They are linked by two disulfide bonds (Mazzarella et al. 1993). BS-RNase exists as a mixture of two distinct forms, M ¼ M and M M. The latter swaps its N-terminal a-helices (for a comparative review on the structures, see Benito et al. 2008a) and at equilibrium exists in slight molar excess (Piccoli et al. 1992). BS-RNase conserves the catalytic residues of RNase A, although its dimeric structure endows it with allosteric properties (Piccoli et al. 1988), the ability to efficiently degrade dsRNA (Libonati and Floridi 1969), and the RNA moiety of RNA:DNA hybrids (Taniguchi and Libonati 1974; Libonati et al. 1975). The quaternary structure of BS-RNase is also responsible for its antitumor activity (for a review, see Youle and D’Alessio 1997). Enzymatic activity and the quaternary structure are necessary for its cytotoxicity (Vescia et al. 1980; Kim et al. 1995b; Antignani et al. 2001). The structure of the complex between RNase A and RI (Kobe and Deisenhofer 1996) shows that a BS-RNase-RI complex is not possible, due to steric hindrance, while a single-chain subunit is strongly inhibited (Murthy and Sirdeshmukh 1992). It is not clear whether the dimeric structure is maintained in the reducing conditions of the cytosol. Cells in which RI has been silenced are more sensitive to BS-RNase action (Monti and D’Alessio 2004). Nevertheless, it has been shown that the presence of RNA as a substrate stabilizes the M M reduced form of BS-RNase (Murthy et al. 1996), and may be counterbalance the conversion of the M M form to the M ¼ M form due to monomer sequestering by RI . Unlike frog RNases, results denying the existence of a cellular receptor for BS-RNase are more convincing. BS-RNase binds to the extracellular matrix (ECM) of different cell lines and this interaction seems to be important for the cytotoxic effect (Mastronicola et al. 1995; Bracale et al. 2002) but it does not bind to cell membranes, suggesting that it enters the cell by adsorption mechanisms (Kim et al. 1995a). Alternatively, it was proposed that BS-RNase binds to the cell membrane through sulfhydryl-disulfide interchange reactions between cell surface sulfhydryls and the disulfides that link both subunits (Bracale et al. 2003). However, this hypothesis is not sustained by results that demonstrate that a semisynthetic enzyme remains dimeric and conserves the cytotoxic properties of BS-RNase (Kim et al. 1995a). BS-RNase has been localized in endosomes and its cytotoxicity is blocked when this energy-dependent mechanism is inhibited (Bracale et al. 2002). The intracellular routing of BS-RNase, studied using drugs that disrupt this traffic and immunofluorescence methods, shows that it is localized in the trans-Golgi network of treated malignant cells, suggesting that this organelle, but not the ER, is an effective site for translocation and provides an explanation for its selectivity (Wu et al. 1995; Bracale et al. 2002). BS-RNase can destabilize artificial membranes (Mancheno et al. 1994; Notomista et al. 2006) and it is tempting to speculate that this mechanism is used to permeate the trans-Golgi network membranes, allowing BS-RNase to reach the cytosol. The main targets described for BS-RNase are the rRNAs (Mastronicola et al. 1995) resulting in protein synthesis inhibition and

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subsequent apoptosis. Interestingly, BS-RNase has also been found in the nucleolus of malignant cells (Bracale et al. 2002; Viola et al. 2005). It is not known how the enzyme reaches the nucleus or the significance of this localization. However, a correlation has been shown between cytotoxicity and a decrease of telomerase activity, as well as levels of RNA associated with the enzyme in proliferating cells treated with BS-RNase (Viola et al. 2005). Therefore, telomeric RNA might be another target of BS-RNase. BS-RNase induces apoptosis in a dose-dependent manner in a wide range of cells and reduces tumor growth in vivo (Marinov and Soucek 2000; Sinatra et al. 2000). It was shown to be selectively toxic to human neuroblastoma cell lines with or without the MDR phenotype (Cinalt et al. 1999) and to thyroid cancer cells. In vivo it induces significant tumor regression and shows no cytotoxicity for healthy mice (Kotchetkov et al. 2001; Spalletti-Cernia et al. 2003, 2004). The apoptotic mechanism has been studied in thyroid carcinoma cells and is associated with the activation of caspase-8, 9, and 3 accompanied by a reduced phosphorylation of Akt/protein kinase B (Spalletti-Cernia et al. 2003).

3.2.3

Eosinophil Cationic Protein (ECP)

Activated eosinophils release toxic proteins among which ECP and eosinophilderived neurotoxin (EDN) are members of pancreatic-type family (for a review see Venge and Bystrom 1998; Boix 2001). ECP is a potent cytotoxic protein that is able to kill normal and malignant mammalian cells (Motojima et al. 1989; Maeda et al. 2002a; Carreras et al. 2005), non-mammalian cells such as parasites and bacteria, as well as viruses (for a review, see Boix et al. 2008). The antipathogenic capacities have classified ECP as a human host defense RNase involved in inflammatory processes mediated by eosinophils (see Chap. 2). Although not described as an antitumor agent, its RNase nature and potent cytotoxicity merits a brief description. ECP is a highly stable protein (Maeda et al. 2002b; Nikolovski et al. 2006). Its structure has been solved by x-ray crystallography (Boix et al. 1999; MallorquiFernandez et al. 2000) and in solution (Laurents et al. 2009), and corresponds to the “RNase A fold,” although it has low ribonucleolytic activity (Barker et al. 1989). The relationship between the cytotoxic effect of ECP and its RNase activity is controversial. While its ability to kill bacteria and parasites (Rosenberg 1995) and some mammalian cells (Navarro et al. 2008; Chang et al. 2010) does not depend on the ribonucleolytic activity, the neurotoxic and antiviral activities do (Durack et al. 1979; Domachowske et al. 1998). ECP induces apoptosis in mammalian cells (Navarro et al. 2008, 2010; Chang et al. 2010) in a cell-dependent manner (Benito et al. 2008b). In the human bronchial epithelial cell line, Beas-2B, ECP is internalized following a clathrin- and caveolin-independent but lipid raft-dependent macropinocytosis, being the cell surface–bound heparin sulfate its major receptor (Fan et al. 2007). Recently, the ECP heparin binding affinity (Fan et al. 2008) has been shown to

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depend on the catalytic sites (Torrent et al. 2011). In Beas-2B, ECP causes apoptosis mainly through the TNF-amediated caspase-8 specific pathway in a mitochondria-independent manner (Chang et al. 2010). In contrast, apoptosis does not require internalization in human acute promyelocytic leukemia HL-60, cervix adenocarcinoma HeLa cell lines, and primary cultures of cerebellar granule cells and astrocytes. ECP binds and aggregates on the cell surface, altering membrane permeability, which results in a modification of the ionic intracellular equilibrium (Navarro et al. 2008, 2010). The membrane destabilization promoted by ECP, using synthetic lipid vesicles as membrane models, is mediated by a carpetlike mechanism (Torrent et al. 2007).

3.2.4

Ribonucleases of Different Origins

Over the years, not only the pancreatic-type RNases have drawn increasing attention due to their remarkable antitumor properties but RNases purified from multiple origins. Representative RNases of fungal, bacterial and plant origin have also shown their medicinal potential in the treatment of tumors. Among them, the most promising RNases are mushroom RNases (Ng 2004; Wong et al. 2010), a-sarcin from Aspergillus (Olmo et al. 2001), binase and barnase, two wellknown T1 ribonuclease members from Bacillus (Makarov et al. 2008), RNase Sa3 from Streptomyces (Sevcik et al. 2002), and ginseng RNases (Fang and Ng 2011). Besides, RNases from wheat leaf, mung bean, black pine pollen, tomato and hop have been reported to exhibit antitumor activities (Skvor et al. 2006; Soucek et al. 2006; Lipovova et al. 2008; Matousek 2010; Matousek et al. 2010). However, future studies on these RNases with prominent medicinal activities that would open a new perspective for them as potential antineoplastic drugs and their translation from the bench to the clinic are still needed. Fast developing protein engineering of these RNases, which display more potent cytotoxic activity and greater selectivity for malignant cells, is now the aim of researchers.

3.3

Approaches to Endow a Ribonuclease with Cytotoxic Properties

The discovery of the natural RNases described above has stimulated the construction over the past 15 years of other cytotoxic RNases with enhanced properties. The main objectives of these efforts have been to increase tumor cell selectivity and to construct more powerful antitumor drugs that could bypass the main drawbacks of ONC, which are its renal toxicity (Vasandani et al. 1996, 1999a) and its non-human origin. In addition, RNases are low molecular mass proteins that are rapidly cleared from the organism (Maack et al. 1979).

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Monomeric Ribonucleases

The cytotoxic pathway of RNases, described in the introduction section, has inspired different strategies to endow an RNase with cytotoxic activity. Researchers have tried to make a particular RNase more efficient to carry out some steps of this model in order to obtain more potent cytotoxins. The first artificial monomeric RNase to be described was constructed by Raines and coworkers. In this seminal work, changes were rationally introduced in the RNase A sequence to decrease its affinity for the RI (Leland et al. 1998). They replaced residue Gly88, which was positioned in a hydrophobic pocket of the RI in the three-dimensional structure of the complex (Kobe and Deisenhofer 1996), by different bulky charged residues (Arg or Asp). The replacement by Arg produced an RNase with a 104-fold less affinity for the RI, which was approximately only 20-fold less cytotoxic than ONC. The success of this approach was confirmed using other monomeric RNases like HP-RNase (Gaur et al. 2001; Leland et al. 2001) or monomeric BS-RNase (Antignani et al. 2001; Lee and Raines 2005). Further mutations were introduced in RNase A and HP-RNase to disturb their electrostatic attraction for the RI (Bretscher et al. 2000; Haigis et al. 2002; Rutkoski et al. 2005; Johnson et al. 2007b) obtaining new variants with an increased cytotoxicity. The substitutions D38R/R39D/N67R/G88R in RNase A yielded a variant that maintained ribonucleolytic activity and conformational stability but had 5.9 109-fold lower affinity for RI. This variant showed a cytotoxicity that was nearly equal to that of ONC (Rutkoski et al. 2005). However, when creating a new RNase variant, it is difficult to discern how a specific mutation may affect other characteristics important for the cytotoxicity. An illustrative example refers to variants evading the RI. In some cases, the introduced mutations produced variants that were more stable or that had less affinity to the RI but also were less catalytically active and therefore, the increase in cytotoxicity was minor (Bretscher et al. 2000; Dickson et al. 2003). An alternative strategy to bypass the RI action has been to redirect the RNase to the nucleus, which is believed to be devoid of this inhibitor (Roth and Juster 1972). An HP-RNase variant, named PE5, which carries a non-contiguous extended bipartite nuclear localization signal (NLS), has been reported (Rodriguez et al. 2006). This NLS is constituted by at least three different regions of the protein comprising residues 1, 31–33, and 89–91. This variant recognizes a-importin (Rodriguez et al. 2006) and cleaves nuclear but not cytoplasmic RNA in vivo. In addition, the modification of residues important for this NLS significantly decreases the variant’s cytotoxicity (Tubert et al. 2010). All these results show that this NLS endows this HP-RNase variant with cytotoxic activity (Bosch et al. 2004; Tubert et al. 2010). The variant is internalized and reaches the cytosol where it can interact with the RI and with the a-importin. It is likely that the concentrations of RI and a-importin are similar in the cytosol (RI concentration is 4 mM (Haigis et al. 2003) and a-importin concentration is 3 mM in Xenopus oocytes (Gorlich et al. 1994)), thus the affinity of the RNase for each protein will determine to which it will mainly

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bind. However, those RNase molecules captured by the a-importin will be released into the nucleus and therefore removed from the two competing equilibriums and PE5 will progressively accumulate into the nucleus (Fig. 3.2). The Gly88R RI-evading RNase A variant described above was modified by the addition of cystine bonds in order to increase its conformational stability (Klink and Raines 2000) resulting in a more cytotoxic variant. A different strategy to increase the stability and therefore the toxicity is glycosylation of the protein; the expression of ONC in Piscia factoris produces a glycosylated protein at Asn69 which is more stable, more resistant to protein degradation, and also is 50-fold more cytotoxic (Kim et al. 2004). The efficiency of internalization is another important determinant of cytotoxicity (Wu et al. 1995; Leich et al. 2007). One strategy to increase the internalization efficiency of the RNases is their cationization by chemical or genetic modification. This strategy is based on the rationale that a more efficient internalization could be achieved by using electrostatic interactions to efficiently adsorb highly cationic proteins into the negatively charged surface. It has been reported that the chemical modification of the carboxyl groups of RC-RNase with a watersoluble carbodiimide in the presence of nucleophiles or the amidation with ethylenediamine, 2-aminoethanol, taurine or ethylenediamine of HP-RNase and RNase A increases their cytotoxicity (Futami et al. 2001, 2002; Iwama et al. 2001). The enhancing effect was dependent on the increase in positive net charge and the higher cationic variants were more efficiently internalized into the cells. In addition, the preparation of RNase A and noncytotoxic cross-linked dimers of RNase A, both covalently linked to polyspermine to increase their basicity, has also been described (Pouckova et al. 2007). In this case, only the dimeric structures, which were much more basic, slightly increased the cytotoxicity exerted by the free polyspermine (two-fold increase). In some cases, the chemical modifications seriously compromised the ribonucleolytic activity of the modified enzyme (Futami et al. 2001, 2002) and were not specific, generating heterogeneous products that would be difficult to use as antitumor drugs. In any case, the effect of cationization has been confirmed by site-directed mutagenesis. In RC-RNase variants where Asp or Glu residues were replaced by Asn, Gln, or Arg (Ogawa et al. 2002), antitumor activity and internalization were enhanced. In addition, the replacement of acidic residues by positively charges residues increased the cytotoxicity of Streptomyces aureofaciens RNase Sa (Ilinskaya et al. 2002, 2004). However, it has been also shown that promoting the internalization of pancreatic RNases by introducing positive charges into the molecule can be counterbalanced by an increased affinity to the anionic RI in the cytosol of the resulting variants (Johnson et al. 2007a). To improve the internalization, Raines and colleagues (Fuchs et al. 2007) also replaced two residues of a cytotoxic variant of RNase A to create a patch of Arg residues on its surface. The cytotoxicity of the resulting variant was slightly improved (threefold increase). On the other hand, the addition of a protein translocation domain (nona-arginine) to improve the translocation of the RNase to the cytosol has been shown to increase the cytotoxicity of previously cytotoxic RNase A variants (Fuchs and Raines 2005; Fuchs et al. 2007). Recently, it has been reported that an

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alternative strategy to enhance the cellular internalization of the RNases consists in co-treating cells with a cationic 2 poly(amidoamine) dendrimer (Ellis et al. 2010). This treatment increases the cytotoxicity of the RNase probably by increasing endosomal escape. In addition, it avoids the deleterious consequence of decreasing ribonucleolytic activity or conformational stability observed upon cationization of the RNases. The engineering of RNases to improve their intracellular pathway has not been extensively explored, and the studies conducted until now have not given positive results. It has been shown that the introduction of a Lys-Asp-Glu-Leu consensus sequence to direct BS-RNase to the endoplasmic reticulum significantly reduces its cytotoxicity (Bracale et al. 2002). The alteration of intracellular routing has been also assayed with the Gly88R RI-evading RNase A variant to diminish its routing to the lysosomes and therefore favor the translocation of more protein. This variant carries a Lys-Phe-Glu-Arg-Gln sequence (residues 7–11) that targets it for lysosomal degradation. However, the replacement of Lys7 with Ala had no effect on the cytotoxicity (Haigis et al. 2002). Finally, formulations for improving tissue delivery have also been described. ONC has been encapsulated in biodegradable poly(ricinoleic-co-sebacic acid) for local controlled delivery in the parietal lobe of the brain in an attempt to overcome cerebellar neuronal toxicity while affecting glioma cells (Slager et al. 2009). ONC was released in a controlled manner and was cytotoxic against 9L glioma cells xenografted into the brain, while evading neurotoxicity in the cerebellum.

3.3.2

Oligomeric and Tandem Ribonucleases

BS-RNase has inspired the generation of other cytotoxic RNases through the formation of oligomeric structures that hinder the binding of the RI by steric hindrance (for a review see Libonati 2004; Libonati et al. 2008). The cytotoxicity of BS-RNase seems to be restricted to the swapped form (Cafaro et al. 1995; Mastronicola et al. 1995), which likely remains dimeric in the reductive conditions of the cytosol. However, the reasons for the cytotoxicity are not completely understood and the activity of different variants seems to contradict this model. For example, a dimeric HP-RNase variant in which the enzyme had been engineered to reproduce the sequence of BS-RNase helix-II and to eliminate the Glu111 charge on the surface was highly cytotoxic (Merlino et al. 2009) but this activity was also associated with the unswapped form of the protein. Moreover, RNase A dimeric variants covalently linked by two Cys at positions 31 and 32 with swapping propensities ranging between 14% and 60% are poorly cytotoxic (Ercole et al. 2009). This shows that other features must be needed for a dimeric RNase to be cytotoxic. Nevertheless, the design of RNase oligomers is attractive since they present more positive charges and therefore must interact more tightly with the anionic surface of cancer cells and, in most cases, they cleave dsRNA. Furthermore,

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the increase in molecular mass could help to avoid the clearance of the drug from the organism (Bartholeyns and Moore 1974). Cytotoxic HP-RNase (Piccoli et al. 1999; Di Gaetano et al. 2001); and RNase A (Di Donato et al. 1994) variants have been created by introducing some of the residues of BS-RNase that are believed to be important for the swapping process. In all these cases, two disulfide interchain bonds were established between the Cys introduced at positions 31 and 32. The deletion of five residues of the loop that connects the N-terminal a-helix of HP-RNase with the rest of the protein produced a dimeric enzyme (Russo et al. 2000), but until now, no cytotoxicity data have been reported. Other covalent linkers have been proposed to stabilize the RNase dimeric structure. Dimers of RNase A cross-linked with dimethyl suberimidate have been shown to display an antitumor action both in vitro and in vivo (Bartholeyns and Baudhuin 1976; Tarnowski et al. 1976). Higher oligomers cross-linked with the same strategy displayed higher cytotoxicity in vitro (Gotte et al. 1997). These variants were heterogeneous, and more specific cross-linking strategies were subsequently evaluated. They were based on the introduction of additional Cys residues to provide reaction sites for specific chemical reagents. Raines and colleagues (Kim et al. 1995b) introduced a thioether bond between both Cys31 residues of BS-RNase to increase the dimer stability, but they did not obtain an increase in cytotoxicity. On the other hand, the formation of covalent dimers by the introduction of a thioether bond between the cysteines of the RNase A Cys89 variant or the EDN Cys87 variant produced cytotoxic enzymes (Suzuki et al. 1999). Finally, an exhaustive evaluation of cross-linkers and of sites introducing the additional cysteine has recently been reported (Rutkoski et al. 2010). Some of these constructs were as effective as monomeric RNase A variants that highly evade RI to inhibit tumor growth. The construction of recombinant proteins with duplicated RNase genes is an alternative strategy to covalently link two RNase units. Arnold and colleagues (Leich et al. 2006) constructed tandem dimers of RNase A in which it was expected that one unit of the RNase tandem enzyme should remain unbound due to steric hindrance, but that the other could be trapped by the RI. Surprisingly, the tandem construction, although fully inhibited by the RI, was cytotoxic. As stated above, interpretations other than RI inhibition can explain the acquisition of cytotoxic properties of oligomeric or multimodular RNases.

3.3.3

Targeted Ribonucleases

Most of the natural RNases are devoid of innate anticancer activity and have not evolved mechanisms for efficiently entering and killing cells. However, nontoxic RNases can be linked to targeting molecules and acquire or perform cell typespecific cytotoxic activity. RNase-based targeted therapeutics, which have been developed in parallel with the so-called smart drugs or targeted drugs together with

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Fig. 3.3 Schematic drawing of targeted RNase constructs. Different strategies have been developed to target RNases by conjugation or fusion either to antibodies or ligands. RNases used in the different approaches are: (a) HP-RNase, EDN, ANG; (b) HP-RNase, EDN, ANG, BS-RNase, rapLRI and ONC; (c) rapLRI, ANG and HP-RNase (d)HP-RNase; (e) ANG; (f) RNase A, HPRNase, ECP, ANG; (g) HP-RNase. Cell surface targets used in the different constructions are: (a) TfR, CD30; (b) TfR, ErbB2, P-gp, hPLAP, ErbB2, P-gp, CD22, CD30; (c) CD22, CD30; (d) CD30, (e) TfR; (f) Tf, LHRH, EGF; (g) EGF, bFGF, IL-2. N and C indicate the amino and carboxyl terminus

monoclonal antibodies (mAb) and recombinant antibody technology, will be the aim of this section. Figure 3.3 shows schematically the strategies to target RNases by conjugation or fusion either to antibodies or ligands described below.

3.3.3.1

Chemical RNase Conjugates

One approach to improve the therapeutic efficacy of a drug is to combine a targeting molecule (hormone, interleukins, antibody, etc.) with the effector moiety (radioisotope, toxin, drug-activating enzyme, etc.) in the same molecule. In particular, the combination of the antigen-specific targeting abilities of antibodies with the toxicity of a payload toxin from plants such as ricin from Ricinus communis, or bacterial enzymes such as diphtheria toxin from Corynebacterium diphtheria or Pseudomonas exotoxin A, is referred to as “immunotoxins” (Pastan et al. 1986) (for review see Wu and Senter 2005; D€ ubel 2007).

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The most promising diphtheria toxin conjugates are the human transferrin receptor (TfR)–specific CRM107 conjugate (Johnson et al. 1989) and the CD3-specific A-dmDT390-bisFv(UCHT1) (Woo et al. 2008), which has entered phase I/II clinical trials. Also, ricin, a ribosome-inactivating protein from plants, has been thoroughly used as an immunotoxin payload, but it did not progress further than phase I trials due to product heterogeneity and individual differences in patients’ responses (Messmann et al. 2000). These immunotoxins demonstrated a potent in vitro tumor cell killing activity, but when applied in human patients, they caused toxic side effects and immunological antidrug responses that limited their therapeutic potential. In addition to the nonspecific binding and toxicity of immunotoxins using microbial or plant toxins that have resulted in several fatalities in clinical trials, the strong immunogenicity of the toxic heterologous compounds disallows repeated or long-term clinical applications and even short-term treatment requires prophylactic protocols. Although some successful clinical results may arise after continued refinement of the engineered plant and bacterial toxins, an alternative to all these problematic approaches is offered by the use of pancreatic-type RNases. The tumor targeting of RNases was first demonstrated by the chemical conjugation of RNase A to transferrin (Tf) or mAb against the TfR (Rybak et al. 1991) and against the T cell antigen CD5 (Newton et al. 1992). Soon after, ONC was compared to RNase A as a conjugated cargo of anti-TfR mAb 5E9 (Rybak et al. 1993) (for review, see Rybak 2008). The anti-TfR-RNase A conjugates were comparable in vivo to a ricin A chain conjugate, although in vitro, results had shown that the RNase conjugate was much less efficient. These studies also showed that the antibody conjugates were more efficient than the Tf conjugates, due likely to an enhanced binding to the cell and consequent internalization. Interestingly, the antibody conjugates linked by a reducible disulfide bond to ONC and RNase A were equally potent in the nanomolar range, even though ONC, which is cytotoxic in the micromolar range (0.1–0.8 mM for the majority of tested cell lines (Haigis et al. 2003)), is several orders of magnitude more cytotoxic than RNase A when assayed as non-conjugated drugs. In addition, while ONC did not change its internalization pathway, it is still not known whether RNase A evades the cytosolic RI, or saturates it. ONC has also been conjugated to the CD22-specific mAb LL2 and RFB4, which resulted in a many thousand-fold increase in cytotoxicity comparable to the specificity and potency of anti-CD22 immunotoxin conjugates with a plant or bacterial toxin payload (Newton et al. 2001), confirming that RNases are as potent as these toxins when properly targeted. ONC has also been conjugated to P-glycoprotein (P-gp) neutralizing mAb MRK16. When investigated against MDR1 overexpressing human carcinoma cells in vitro and in vivo, it was observed that the ONC conjugate increased its cytotoxicity and sensitized the multidrug-resistant cancer cells to vincristine in vivo (Newton et al. 1996). Although P-gp is not a rapidly internalizing transmembrane protein, the increase in cytotoxicity and the vincristine sensitizing effect of the MRK16-ONC conjugates could be explained by the P-gp cell-binding enhancement of the mAb, which at the same time diminishes the drug-expelling activity of the P-gp, and the internalizing capabilities of ONC by itself.

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RNase Fusion Proteins and ImmunoRNases

The progress of recombinant antibody engineering and fusion protein technology has led to the rapid expansion of drug-targeting approaches including the development of an antibody–RNase fusion protein. Recombinant antibody technology (D€ ubel 2007) has been used to improve novel antibody formats and drug-targeting devices with superior antigen binding, pharmacokinetic, and effector and production properties. The intended therapeutic purpose strongly influences the choice of a particular antibody format. Features such as molecular size, valence, additional domains, or chemical modifications must be carefully considered to achieve the most selective targeting, pharmacokinetic, and therapeutic efficacy. Generally speaking, small antibody or antibody fusions penetrate and distribute best into solid tissues and tumors. However, these constructions disappear from the blood circulation faster and, thus, have a reduced serum half-life. It has been proposed that engineering therapeutic molecules with a molecular size between 60 and 120 kDa provides the best equilibrium between tumor penetration and longest half-life (Hudson and Souriau 2003). As a proof of the concept, bivalent antibodies have shown remarkable enhanced tumor retention in comparison to monovalent counterparts (for a review of antibody architectures used in drug targeting, see (Rybak and Newton 2007). Several members of the RNase A superfamily have been used either as a scaffold onto which a targeting domain is engineered or fused to a targeting antibody. When choosing a particular vertebrate RNase, one has to consider that human RNases are believed to have the least immunogenic payload (De Lorenzo et al. 2004; Menzel et al. 2008) and also that RNases may display functions other than ribonucleolytic activity. Additionally, the connecting linker and the orientation of the RNase and antibody fragment have to be taken into account. For instance, ANG could promote remarkably enhanced selective tumor cell killing provided that the endothelial cell binding and angiogenic activity is maintained, which to a certain extent depends on the C-terminal residues of the molecule, while for amphibian RNases, the pyroglutamyl N-terminal residue has been proved essential. Regarding all the available clinical data for ONC (see Sects. 3.1 and 3.4), this amphibian RNase seems to be the most promising effector to use as an antibody payload, but constructs using HP-RNase or ANG or engineered variants have also shown potential for tumor therapy. Antibody RNase fusions using HP-RNase, ANG, or EDN have been tested in experimental sets targeting the TfR and have shown approximately 103 times more potency than the respective chemical antibody RNase conjugates (Rybak et al. 1992; Newton et al. 1994). However the (Fab)2-like CH2-ANG immunoRNase fusion construct exhibited serious production difficulties in myeloma cells with a yield of only 5 ng/mL. Yet, the same fusion protein could be produced in mammary glands of transgenic mice with a final yield of 0.8 mg/mL milk (Newton et al. 1999), indicating that yield is also dependent on the expression system used.

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Several studies have investigated different immunoRNase designs to overcome production concerns and to extend this approach to more promising clinical targets, using lineage antigens expressed on certain types of leukemia but not present on hematopoietic stem cells such as CD22 and CD30, or in particular internalizing tumor antigens such as ErbB-2 (Schirrmann et al. 2009). Chemical conjugates combining mice anti-CD22 mAbs and RNases have shown great potential as novel antitumor drugs. In animal models, these immunoRNases reduced tumors size with high efficacy while not showing appreciable toxicity. However, their heterogeneity affected their binding capacity, caused lot-to-lot variability, and prevented further development. As an alternative, ANG, HPRNase or RapLRI (Rana pipiens liver RNase I), a close relative of ONC, was fused to two CD22-specific scFv antibody fragments generated either by reengineering the variable domain core structure of mAb LL2, or by grafting the complementarity-determining regions of the clinically established mAb RFB4 into consistent human scFv scaffolds (Arndt et al. 2005; Krauss et al. 2005a, b). Among them, CD22-specific ANG-scFv(RFB4) immunoRNase was successfully produced in a mammalian myeloma cell line without degradation, and exhibited potent cytotoxicity with an IC50 in the nanomolar range (Krauss et al. 2005). Since bivalency improves retention to the tumor location (Hudson and Souriau 2003), dimeric secondgeneration derivatives were generated from monovalent first-generation anti-CD22 immunoRNases to improve the cytotoxicity and pharmacokinetics. Hence, scFv fragments LL2 and humRFB4 were engineered into a diabody format, fused either to ANG or RapLRI, and successfully produced in E. coli cells. Bivalent anti-CD22 immunoRNases showed a markedly superior cytotoxicity toward CD22+ tumor cells when compared with monovalent counterparts due to improved antigen binding by avidity effect and enhanced internalization (Arndt et al. 2005; Krauss et al. 2005). Also, different CD30 targeting approaches fusing HP-RNase or ANG to CD30specific murine or human scFv have been developed. While scFv(Ber-H2)HP-RNase fusion protein produced in insect cells inhibited tumor growth in vivo and in vitro (Braschoss et al. 2007), the entirely human bivalent immunoRNase ScFv-Fc-HP-RNase, consisting of human scFv, human IgG1 Fc part, and HP-RNase, showed better properties and inhibited growth of CD30+ Hodgkin lymphoma cells with an IC50 of 3 nM (Menzel et al. 2008). Even better results were observed for immunoRNases produced in HEK293T cells resulting from the fusion of CD30-specific scFv Ki4 to ANG which inhibited the growth of CD30+ lymphoma cells in vitro with an IC50 of 0.5 nM (Stocker et al. 2003). The first entirely human antibody RNase fusion protein consisting of an ErbB2 specific human scFv antibody fragment fused to HPR was described by D’Alessio and coworkers (De Lorenzo et al. 2004). The human anti-ErbB2 single-chain variable fragment, Erbicin, specifically distinguished ErbB2-positive cells with high affinity and was internalized upon specific antigen recognition by ErbB2-expressing cells. Erbicin strongly inhibited receptor autophosphorylation and displayed strong inhibitory activity on the growth of ErBB2-positive cell lines. The combination of these

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properties with the ribonucleolytic activity of HP-RNase resulted in a remarkable reduction (86%) of ErbB2-positive tumors in mice. Although the immunoRNase was inhibited by the cytosolic RNase inhibitor to an extent comparable to the free HPRNase (De Lorenzo et al. 2007), the amounts of immunoRNase entering the cytosol saturated the endogenous RNase inhibitor present in this cellular compartment. Based on bivalent immunoRNases being more powerful than monovalent ones, a dimeric variant of HP-RNase was fused to two Erbicin molecules, one per subunit (Riccio et al. 2008). The new immunoRNase, Er-HHP2-RNase, was found to selectively bind ErbB2-positive cancer cells with an increased avidity with respect to monovalent anti-ErbB2 scFv- HP-RNase, and exerted a more powerful cytotoxic activity, likely due to an increased RNase inhibitor evasion. Recently the human antitumor immunoconjugate engineered by the fusion of Erbicin with human RNase has been assayed on trastuzumab-resistant cells, and it has proven to be selectively cytotoxic for ErbB2-positive cancer cells both in vitro and vivo; targets an ErbB2 epitope different from that recognized by trastuzumab; and does not show cardiotoxic effects (Gelardi et al. 2010). Barnase manifests potent antitumor activity, but toxicity to the host cells limits its potential clinical application. Deyev and coworkers established a new plasmid for eukaryotic expression of a scFv 4D5-dibarnase, which consists of two barnase molecules fused serially to the single-chain variable fragment (scFv) of humanized 4D5 antibody. The 4D5 antibody is directed against the extracellular domain of human epidermal growth factor receptor 2 (HER2) and could assist the delivery of barnase to HER2/neu-positive cells and facilitate its penetration into the target cells (Glinka et al. 2006). They further evaluated its antitumor activity and toxicity in mice bearing HER2-overexpressing human breast cancer xenografts. This immunotoxin scFv 4D5-dibarnase manifested a specific apoptosis-associated cytotoxic effect on HER2-overexpressing SKBR-3 and BT-474 human breast carcinoma cells in vitro, and a significant inhibition of human breast cancer xenografts in nude mice without severe side effects (Balandin et al. 2011). Apart from being used as antibody payload, recombinant RNases have been either fused or chemically conjugated to various internalizing cell-binding ligands such as luteinizing hormone–releasing hormone (LHRH) (Gho and Chae 1999), basic fibroblast growth factor (bFGF) (Suzuki et al. 1999), epidermal growth factor (EGF) (Jinno et al. 1996a, b, 2002; Psarras et al. 1998; Yoon et al. 1999), or interleukin 2 (IL2) (Psarras et al. 2000). Although most of them overcame the concern of RNase inhibition when translocated into the cytosol and were cytotoxic to different tumor cell lines, the different constructs did not selectively target tumor cells.

3.4

Clinical Development

As mentioned above, ONC is the first RNase that reached clinical trials. Although most of them are completed, the full results have not yet been published (http:// www.cancer.gov/). Table 3.1 summarizes the clinical trials performed with RNases

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Table 3.1 Clinical trials performed with cytotoxic ribonucleases Main observations Protein: Onconase Two Phase I clinical trials with patients with solid tumors demonstrated that ONC was well tolerated. Dose limiting toxicity was renal but with reversal effects. MTD 960 mg/m2/week and recommended doses for Phase II 480 mg/m2/week In a Phase II trial, ONC increases the median survival time (7.7 months) of patients with advanced non-small cell lung cancer (NSCLC) compared with patients treated with a variety of chemotherapeutic regimes; Ongoing Phase II clinical trial with the addition of ONC to pemetrexed plus carboplatin in patients with NSCLC (http://www.alfacell.com/) A Phase I/II trial involving patients with advanced pancreatic carcinoma treated with ONC and tamoxifen suggested potential activity of the combination according to preclinical results. This trial was discontinued in 1998 because tolerated levels of ONC did not offer a significant advantage over gemcitabine (Gemzar®) In a Phase II trial, patients with prostate cancer recurrence were treated with a combination of ONC and tamoxifen with extremely low clinical benefit A Phase II study of ONC in patients with advanced breast cancer resulted in limited clinical benefit In a Phase II trial of ONC in patients with metastatic kidney cancer, it had minimal activity In a multicenter Phase II clinical trial carried out in patients with UMM, ONC demonstrated activity, including for those pretreated with one or more chemotherapeutic regimens, and tolerable toxicity. Overall median survival was 6 months for the intent-to-treat group and 8.3 months for the treatment target group. ONC acted as a cytostatic agent. Obtained results were the basis for the initiation of the randomized Phase III trial Initial Phase III clinical trial for patients with MM treated either with ONC or doxorubicin showed no significant differences. However, the group treated with ONC revealed an excess of poor prognosis patients. Retrospective analysis clearly favored ONC (median survival 11.6 months for ONC-treated group vs. 9.6 months for the doxorubicin group) setting the basis for another Phase III analysis A Phase IIIb study performed on a global scale in patients with UMM, to test the effects of combined use of ONC and doxorubicin vs. doxorubicin alone, seemed to favor the drug combination Protein: HP-RNase variant QBT-139 Ongoing Phase I clinical trial to evaluate the toxicity and tolerability (MTD) of QBT-139 in patients with advanced, refractory solid tumors

References

Mikulski et al. (1993b)

Mikulski et al. (1995)

Chun et al. (1995) Not published although completed Puccio et al. (1996) Vogelzang et al. (2001)

Mikulski et al. (2002)

Not published although completed Not published although completed

http://www.cancer.gov/

and some of the main published results. The reader is addressed to the following reviews for more detailed information (Costanzi et al. 2005; Favaretto 2005; Pavlakis and Vogelzang 2006; Ramos-Nino 2007; Beck et al. 2008; Lee 2008; Porta et al. 2008). The major advances of ONC have been in the treatment of unresectable malignant mesothelioma (UMM) (confirmatory Phase IIIb). However, in the annual report of the Alfacell Corporation (2009) (at present Tamir Biotechnology Inc.) (http://www.alfacell.com/annualreport2009.pdf), the company indicates that it decided not to pursue further clinical trials for the treatment of UMM based on

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recommendations of the FDA. The ongoing trials are now focused on the treatment of patients with non-small cell lung cancer. It is remarkable that an engineered HP-RNase that evades the RI (Evade™ Ribonuclease family from Quintessence Biosciences Inc; http://www.quintbio. com/whoIs.asp) is now in Phase I of clinical trials. This RNase of human origin presents a new challenge for research into the use of these enzymes as chemotherapeutics for cancer treatment.

3.5

Conclusions

Modern pharmacological anticancer drugs are no longer focused on “small molecules” and the pharmaceutical industry today is searching for a second line of cancer chemotherapeutics without genotoxic effects. In addition, it is exploring different options like cellular targets or new drug-delivery methods, even to a specific cellular compartment. RNases fall within this second line of anticancer drugs. Since they do not cleave a specific RNA molecule, their effects on gene expression are pleiotropic, ensuring a broad spectrum of synergistic interactions with other chemotherapeutics and, if used alone, making the appearance of resistance to the drug by cancer cells difficult. The efforts described in this chapter to understand their cytotoxic mechanism have led to engineering more potent and selective RNases with fewer side effects than conventional chemotherapeutic drugs. Acknowledgments This work has been supported by grants BFU2009-06935 from MICINN (Spain) and GRCT04 from the University of Girona.

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Ardelt B, Juan G, Burfeind P, Salomon T, Wu JM, Hsieh TC, Li X, Sperry R, Pozarowski P, Shogen K, Ardelt W, Darzynkiewicz Z (2007a) Onconase, an anti-tumor ribonuclease suppresses intracellular oxidative stress. Int J Oncol 31:663–669 Ardelt B, Ardelt W, Pozarowski P, Kunicki J, Shogen K, Darzynkiewicz Z (2007b) Cytostatic and cytotoxic properties of Amphinase: a novel cytotoxic ribonuclease from Rana pipiens oocytes. Cell Cycle 6:3097–3102 Ardelt W, Shogen K, Darzynkiewicz Z (2008) Onconase and amphinase, the antitumor ribonucleases from Rana pipiens oocytes. Curr Pharm Biotechnol 9:215–225 Ardelt W, Ardelt B, Darzynkiewicz Z (2009) Ribonucleases as potential modalities in anticancer therapy. Eur J Pharmacol 625:181–189 Arndt MA, Krauss J, Vu BK, Newton DL, Rybak SM (2005) A dimeric angiogenin immunofusion protein mediates selective toxicity toward CD22+ tumor cells. J Immunother 28:245–251 Arnold U (2008) Aspects of the cytotoxic action of ribonucleases. Curr Pharm Biotechnol 9:161–168 Arnold U, Ulbrich-Hofmann R (2006) Natural and engineered ribonucleases as potential cancer therapeutics. Biotechnol Lett 28:1615–1622 Arnold U, Schulenburg C, Schmidt D, Ulbrich-Hofmann R (2006) Contribution of structural peculiarities of onconase to its high stability and folding kinetics. Biochemistry 45:3580–3587 Balandin TG, Edelweiss E, Andronova NV, Treshalina EM, Sapozhnikov AM, Deyev SM (2011) Antitumor activity and toxicity of anti-HER2 immunoRNase scFv 4D5-dibarnase in mice bearing human breast cancer xenografts. Invest New Drugs 29(1):22–32, Epub 2009 Sep 30 Barker RL, Loegering DA, Ten RM, Hamann KJ, Pease LR, Gleich GJ (1989) Eosinophil cationic protein cDNA. Comparison with other toxic cationic proteins and ribonucleases. J Immunol 143:952–955 Bartholeyns J, Baudhuin P (1976) Inhibition of tumor cell proliferation by dimerized ribonuclease. Proc Natl Acad Sci USA 73:573–576 Bartholeyns J, Moore S (1974) Pancreatic ribonuclease: enzymic and physiological properties of a cross-linked dimer. Science 186:444–445 Beck AK, Pass HI, Carbone M, Yang H (2008) Ranpirnase as a potential antitumor ribonuclease treatment for mesothelioma and other malignancies. Future Oncol 4:341–349 Benito A, Ribo M, Vilanova M (2005) On the track of antitumour ribonucleases. Mol Biosyst 1:294–302 Benito A, Laurents DV, Ribo M, Vilanova M (2008a) The structural determinants that lead to the formation of particular oligomeric structures in the pancreatic-type ribonuclease family. Curr Protein Pept Sci 9:370–393 Benito A, Vilanova M, Ribo M (2008b) Intracellular routing of cytotoxic pancreatic-type ribonucleases. Curr Pharm Biotechnol 9:169–179 Boix E (2001) Eosinophil cationic protein. Methods Enzymol 341:287–305 Boix E, Wu Y, Vasandani VM, Saxena SK, Ardelt W, Ladner J, Youle RJ (1996) Role of the N terminus in RNase A homologues: differences in catalytic activity, ribonuclease inhibitor interaction and cytotoxicity. J Mol Biol 257:992–1007 Boix E, Leonidas DD, Nikolovski Z, Nogues MV, Cuchillo CM, Acharya KR (1999) Crystal structure of eosinophil cationic protein at 2.4 A resolution. Biochemistry 38:16794–16801 Boix E, Torrent M, Sanchez D, Nogues MV (2008) The antipathogen activities of eosinophil cationic protein. Curr Pharm Biotechnol 9:141–152 Bosch M, Benito A, Ribo M, Puig T, Beaumelle B, Vilanova M (2004) A nuclear localization sequence endows human pancreatic ribonuclease with cytotoxic activity. Biochemistry 43:2167–2177 Bracale A, Spalletti-Cernia D, Mastronicola M, Castaldi F, Mannucci R, Nitsch L, D’Alessio G (2002) Essential stations in the intracellular pathway of cytotoxic bovine seminal ribonuclease. Biochem J 362:553–560 Bracale A, Castaldi F, Nitsch L, D’Alessio G (2003) A role for the intersubunit disulfides of seminal RNase in the mechanism of its antitumor action. Eur J Biochem 270:1980–1987

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Chapter 4

RNase T2 Family: Enzymatic Properties, Functional Diversity, and Evolution of Ancient Ribonucleases Gustavo C. MacIntosh

Contents 4.1 4.2 4.3 4.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Protein Properties and Overall Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Enzymatic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological Roles and Evolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.1 Cytotoxic Ribonucleases, Defense Responses, and Self-Incompatibility . . . . . . . 4.4.2 Phosphate Scavenging and a Housekeeping Role in rRNA Recycling . . . . . . . . . 4.4.3 Catalysis-Independent Functions and Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract RNase T2 enzymes are transferase-type endoribonucleases that produce oligonucleotides and/or mononucleotides with a terminal 30 phosphate via a 20 ,30 cyclic phosphate intermediate. These RNases are found in all eukaryotes and also in bacteria and viruses, where they have a wide range of biological activities. Some have a housekeeping role, degrading rRNA, and mutations affecting this function result in alterations in cellular homeostasis and are associated with brain lesions in vertebrates. Others have a variety of specialized roles including antimicrobial defense, phosphate scavenging, rejection of “self” pollen, and even nitrogen storage. Members of this family have also acquired functions that are independent of their ribonuclease activity. One of these catalysis-independent functions is implicated in the control of cellular growth, and lack of RNASET2 protein in humans is correlated with several classes of tumors. This review will discuss the basic structure, enzymatic properties, and biological roles of this ancient RNase family.

G.C. MacIntosh Department of Biochemistry, Biophysics and Molecular Biology, lowa State University, Ames, IA 50011, USA e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_4, # Springer-Verlag Berlin Heidelberg 2011

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4.1

G.C. MacIntosh

Introduction

Prokaryotic and eukaryotic cells possess a large number of ribonucleases (RNases) that participate in many cellular functions, from DNA replication, to control of gene expression, to defense against microorganisms. Among those, a set of RNases are secreted or localized inside cellular structures associated with the secretory pathway, the vacuole, or lysosomes; thus, these enzymes are found in a space not normally associated with the presence of RNA. These RNases belong to a superfamily of enzymes that catalyze RNA cleavage via a 20 -30 -cyclic phosphate intermediate at the 30 -terminus of the resulting oligo- or mononucleotide products. Historically, 20 -30 -cyclizing RNases have been classified into three groups according to mass, base specificity, pH preference, and origin. According to Irie (1999), these correspond to the RNase T1, RNase A, and RNase T2 families. The RNase T1 family includes alkaline RNases with a molecular mass ~12 kDa and pH optima between 7 and 8 that are found in fungi and bacteria (Irie 1999; Deshpande and Shankar 2002). The vertebrate-specific RNase A family (discussed in Chaps. 1, 2, and 3) comprises proteins with a molecular mass between 13 and 14 kDa and either an alkaline (7–8) or weakly acidic (6.5–7) pH preference. Finally, the RNase T2 family includes RNases with an average molecular mass around 25 kDa that were originally categorized as acid RNases (Irie 1999). However, the RNase T2 family is very diverse, with members present in almost all eukaryotic genomes and many bacterial and even viral genomes. As will be discussed here, these proteins have a wide size range and broad pH preferences ranging from extreme acidic to very basic. Currently, the best criterion to classify a protein as a member of the RNase T2 family is the presence in its primary sequence of specific amino acid motifs associated with the RNase active site that are conserved in every T2 protein (Irie 1999). Studies on RNase T2 enzymes began almost a century ago, when Noguchi described the presence of nucleic-acid degrading enzymes in Takadiastase, an enzyme mixture prepared from the fungus Aspergillus oryzae (Noguchi 1924). Later, Sato and Egami (1957) purified RNase T1 and RNase T2 from this mixture and showed that they had different biochemical properties. Since then, many fungal, plant, and animal RNase T2 enzymes have been purified, cloned, and characterized. More recently, X-ray crystallographic analyses and mutagenesis studies allowed the identification of the catalysis mechanism and substrate specificity determinants, providing a very good understanding of the enzymological properties of the family. On the other hand, data on the biological roles of the RNase T2 family lagged well behind the large accumulation of biochemical data, in spite of the apparent importance of these enzymes that are conserved in almost all organisms. The discovery that proteins associated with self-incompatibility in several plants had RNase activity and were part of the RNase T2 family opened the door to a series of studies on the function of these enzymes. Since then, the spectrum of biological activities for this family of enzymes has increased exponentially. Some RNase T2

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enzymes are involved in a housekeeping role, turning over ribosomal RNA to maintain cellular homeostasis, while others have acquired specialized functions as defense against microorganisms, determinants of pathogenicity, phosphate scavenging, and even nitrogen storage. Moreover, these proteins have gained functions that are independent of their catalytic activity. These novel functions seem to be important to control cell proliferation and have a significant impact in tumorigenesis.

4.2

Protein Properties and Overall Structure

RNase T2 proteins have apparent molecular weights that fall in the range of 19 to ~97 kDa, with the majority between 20 and 40 kDa (Deshpande and Shankar 2002). In spite of this variability, protein and gene sequence analyses determined that most enzymes have polypeptide chains corresponding to ~20–30 kDa. In most cases, the size range is determined by glycosylation. Early analyses of RNase T2 showed that enzyme preparations were heterogeneous in molecular weight and separated into six fractions on gel filtrations. Amino acid and carbohydrate analyses revealed that in each of these fractions, the protein moiety corresponded to RNase T2 and the heterogeneities were due to the carbohydrate content, mainly galactose (Kanaya and Uchida 1981). Detailed analysis of the glycan moieties present on S-RNases from tobacco and wild tomato (both from the Solanaceae family) found that these proteins are N-glycosylated. The complex patterns containing glucose, mannose, and glucosamine varied largely among alleles of the S-RNase locus, even in a single species (Oxley et al. 1996; Parry et al. 1998). Similar analyses of S-RNase 4 from Pyrus pyrifolia (Rosaceae family) identified additional sugar chains, including xylose, fucose, N-acetylglucosamine, and chitobiose (Ishimizu et al. 1999). Effective use of N-Glycosidase F to remove sugar moieties from plant, animal, viral, and fungal enzymes indicated that N-glycosylation is a common modification of proteins in this family (Inada et al. 1991; MacIntosh et al. 2001; Langedijk et al. 2002; Campomenosi et al. 2006; Hillwig et al. 2011). In addition, O-glycosylation has been found in a few RNases. RNase Le37, from the fungus Lentinus edodes, has a C-terminal extension rich in Ser and Thr residues that are O-glycosylated, in addition to the common N-glycosylation observed on Asn residues (Inokuchi et al. 2000). A similar pattern of O-glycosylation is possibly present in another fungal protein with a C-terminal extension, RNase Irp1 (Watanabe et al. 1995). In addition to the C-terminal extension observed for the proteins RNase Le37 and RNase Irp1 from Basidiomycetes fungi, other enzymes with C-terminal extensions are Rny1 from yeast (MacIntosh et al. 2001), and Erns from classical swine fever virus [CSFV (Langedijk 2002)]; although in the latter cases, no O-glycosylation has been observed. In addition, Erns is the only RNase T2 enzyme to exist as a homodimer. In this case, the holoenzyme is a disulfide-linked

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homodimer of ~90 kDa, with approximately half of the molecular weight being contributed by the sugar moieties (Schneider et al. 1993; Langedijk et al. 2002). Crystal structures are available for Nicotiana alata and Pyrus pyrifolia S-RNases (Ida et al. 2001; Matsuura et al. 2001), wound-inducible RNases NW and NT from Nicotiana glutinosa leaves (Kawano et al. 2002, 2006), RNase LE from cultured tomato cells (Tanaka et al. 2000), RNase MC1 from bitter gourd (Nakagawa et al. 1999), trichomaglin from root tubers of Trichosanthes lepiniate (Gan et al. 2004), RNase Rh from the filamentous fungus R. niveus (Kurihara et al. 1996), and EcRNase I from Escherichia coli (Rodriguez et al. 2008). A conserved overall structure has been found for bacterial, fungal, and plant RNase T2 proteins (Fig. 4.1), despite the low sequence conservation between prokaryotic and eukaryotic enzymes [bacterial EcRNase I shows less than 35% sequence identity with plant RNase T2 family members and less than 15% sequence identity with fungal and animal members (Rodriguez et al. 2008)]. Although sequence identity is low, crystallographic analyses showed that all have a core of hydrophobic residues in similar positions (Deshpande and Shankar 2002; Rodriguez et al. 2008). In addition, the conserved structure of all characterized RNases consists of a four-stranded antiparallel b-sheet (strands b1, b2, b4, and b5), a small two-stranded antiparallel

Fig. 4.1 Overall structure of RNase T2 proteins. (a) Topology diagram of RNase T2 proteins showing structural elements. Elements conserved in all members of the family are shown in red (b-sheets, arrows) and dark blue (a-helixes, rectangles). Light blue indicates partial conservation. Other elements are not conserved in all proteins. Nomenclature follows that of RNase Rh and RNase LE, with the nomenclature used for EcRNase I in parentheses (figure modified from Rodriguez et al. 2008). (b) Ribbon diagram of RNase LE (PDB ID: 1DIX), using the same color coding as in (a)

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b-sheet (strands b3 and b7), and three a-helices (aB, aC, and aD) that are absolutely conserved. Another a-helix (aF) is conserved but with relative position and orientation more variable (Rodriguez et al. 2008). Other structural motifs are found in individual proteins, with different degrees of conservation (Fig. 4.1). Multiple alignments of available RNase T2 sequences showed very high conservation only for the amino acids belonging to the active site of these enzymes (Irie 1999; Rodriguez et al. 2008; MacIntosh et al. 2010), which reside mainly in strands b2 and b5 and helix aC (Kurihara et al. 1996; Tanaka et al. 2000; Rodriguez et al. 2008). All RNase T2 proteins have several highly conserved Cys residues, which form a variable number of disulfide bridges that stabilize the protein structure in an active conformation. The position of C-C bridges was determined by chemical methods (Kawata et al. 1988; Ishimizu et al. 1995), mass spectrometry (Oxley and Bacic 1996; Langedijk et al. 2002), and X-ray crystallography (Kurihara et al. 1996; Tanaka et al. 2000; Rodriguez et al. 2008). Two C-C bridges are conserved in all RNase T2 enzymes (Irie 1999). Two others are generally conserved in the plant/ animal subgroup, and the total number of C-C bridges varies between four in the self-incompatibility SF11-RNase from N. alata (Ida et al. 2001) to seven in trichomaglin (Gan et al. 2004). Similarly, three bridges different from those found in plants and animals are commonly conserved in fungal enzymes in addition to the two conserved in all RNase T2 proteins (Kurihara et al. 1996; Irie 1999). RNase I from E. coli has a total of four C-C bridges (Rodriguez et al. 2008), with the two non-conserved bridges positioned close in space to those found in eukaryotic enzymes. Erns, an envelope glycoprotein from CSFV and other pestiviruses and member of the RNase T2 family (Schneider et al. 1993; Hulst et al. 1994), has four C-C bridges, and one of these bridges is an unusual vicinal disulfide bridge between cysteines 68 and 69 (Langedijk et al. 2002). Since a C-C bridge between adjacent cysteine residues cannot have a long-range structural role, it was proposed that this bridge has another function, perhaps contributing to the formation of homodimers (Langedijk et al. 2002), as Erns is the only RNase T2 enzyme that has been described to exist as a dimer (Konig et al. 1995; Langedijk et al. 2002).

4.3

Enzymatic Activity

Enzymes from the RNase T2 family are transferase type endoribonucleases that produce oligonucleotides and/or mononucleotides with a terminal 30 phosphate via a 20 ,30 cyclic phosphate intermediate (Irie 1999). These enzymes do not have strict base specificity, although some can have base preferences that vary from enzyme to enzyme [an extensive review of individual enzyme preferences was published by Deshpande and Shankar (2002)]. The reaction mechanism was studied mostly using fungal enzymes, but since amino acid residues implicated in catalysis and the geometry of the active site are absolutely conserved in all RNase T2 enzymes so far characterized, it is safe to

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assume that the same reaction mechanism is employed by all enzymes in the family. Chemical modification experiments of fungal and plant enzymes showed that two histidine residues, His46 and His109 (using RNase Rh numbering), are essential for RNase activity (Kawata et al. 1990; Parry et al. 1997). These findings are consistent with the observation that all RNase T2 enzymes have two regions of almost absolute amino acid conservation. These regions correspond to the conserved active site (CAS) of the enzyme and are named CAS I and CAS II, with the consensus sequences F/WTL/IHGLWP and FWXHEWXKHGTC respectively (Irie 1999). The only His residue in CAS I corresponds to His49, and the second His in CAS II corresponds to His109 in RNase Rh. The residues corresponding to CAS I and CAS II are found in the conserved strand b2 and helix aC in all the crystal structures of RNase T2 proteins, with residues His46, Trp49, His 104, Glu105, Lys 108, and His 109 (RNase Rh numbering) forming the active site of the enzyme (Fig. 4.2a) (Kurihara et al. 1996; Tanaka et al. 2000). Based on enzyme properties and the geometry of the active site, a twostep (transphosphorylation and hydrolysis) general acid–base catalysis mechanism (Fig. 4.2a) was proposed (Irie 1999; Tanaka et al. 2000). In the first step, His46 acts as the general acid and His109 as the general base to generate the 20 ,30 cyclic phosphate intermediate. His46, with a higher pKa than His109 (Kawata et al. 1990; Ohgi et al. 1992), interacts with the 50 oxygen of the scissile phosphodiester bond and donates a proton to the released nucleotide. His109, as the general catalyst, removes the hydrogen of the 20 -OH of the ribose moiety. In the second step, the role of the two histidines is reversed; His46 acts as a general base and His109 as a general acid. Unprotonated His46 activates water, resulting in an activated hydroxyl group that attacks the P-O group of the cyclic phosphate. The proton is donated by His109, now acting as general acid. This mechanism is also confirmed by site-directed mutagenesis studies. Mutations in either His residue (H46F or H109F) resulted in virtually inactive enzymes, with less than 0.02% of WT RNase Rh activity (Ohgi et al. 1992). Similarly, mutations in the His residue equivalent to His109 in a Petunia inflata S-RNase resulted in an enzyme with no detectable activity (Huang et al. 1994) and loss of a histidine residue of S-locus ribonuclease equivalent to His46 was associated with loss of RNase activity in Lycopersicon peruvianum (Royo et al. 1994). Analysis of the crystal structure of RNase MC1 from bitter gourd seeds indicated that the putative catalytic residues of RNase MC1 can be easily superimposed with the catalytic residues of RNase Rh (Nakagawa et al. 1999), and site-directed mutagenesis confirmed the role of His46 in catalysis (Numata et al. 2000). However, mutations in the His residue corresponding to His109 from Rh resulted in a RNase MC1 mutant with about 20% of the WT activity, suggesting that in RNase MC1, other residues could replace this His role but less efficiently (Numata et al. 2000). Other conserved amino acid residues have also been implicated in catalysis. Trp49 mutants showed between 84% and 99% reduction in activity compared with WT, depending on the substitution and substrate used (Ohgi et al. 1996b, 1997b), and crystal structures indicated that its indole ring forms a hydrogen bond with the

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Fig. 4.2 Active site, substrate binding and mechanism of catalysis of RNase T2 enzymes. (a) First step of the proposed mechanism of catalysis of RNase T2 enzymes. In this step (transphosphorylation), His46 acts as general acid and His109 as general base. The other amino acid residues contribute to the reaction by stabilizing the pentacovalent phosphate intermediate or binding the substrate. In the second step (hydrolysis, not shown), the role of the two histidines is reversed, His46 acts as general base and His109 as general acid. Residue numbers refer to the RNase Rh sequence (modified from Irie 1999). (b) Space filling model of RNase LE showing the P1 (red), B1 (green), and B2 (green) sites

carboxyl group of Glu105 and has a partial stacking interaction with the imidazole ring of His109 (Kurihara et al. 1996; Tanaka et al. 2000). Thus, it was proposed that one of the roles of Trp49 is to fix the side-chains of active site residues

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(Tanaka et al. 2000). Glu105 seems to be catalytically crucial and probably operates to polarize the P¼O bond or to stabilize the pentacovalent intermediate (Tanaka et al. 2000). Glu105 mutants were strongly inactivated; however, while substitution for alanine resulted in a enzyme with only 0.07% of the WT activity, glutamine or aspartate at that position resulted in enzyme with about 1% of the WT activity(Ohgi et al. 1993), suggesting that Q and D can provide some catalytic function albeit with lower efficiency. His104 also seems to be critical for catalysis, as a H104F mutant only maintained about 1% of the WT activity (Ohgi et al. 1992). Since the Km for several substrates increased for this His104 mutant, H104 was considered to be a binding site for the negatively charged phosphate group of the substrate (Ohgi et al. 1992; Tanaka et al. 2000). It is important to note that many RNase T2 enzymes have natural substitutions at positions corresponding to His104 and Glu105. Many plant S-RNases contain a glutamine in the position corresponding to Glu105 and a histidine is almost never found at the position corresponding to His104 [see for example Richman et al. 1997; Vieira et al. 2008]. The absence of these residues may account for the low specific activity of many S-RNases (Ida et al. 2001). In most fish RNase T2 enzymes, the position corresponding to His104 is occupied by a tyrosine or a series of polar or charged amino acids, suggesting that these enzymes may also have lower specific activity (Suzuki et al. 2005; Hillwig et al. 2009). This finding led Suzuki et al. (2005) to propose that His104 may also be important for stabilization of the pentacovalent intermediate that would result in higher specific activity. In E. coli RNase I, this histidine is also replaced by tyrosine (Meador and Kennell 1990). Another residue important for substrate binding to the active site is Lys108. Crystal structures showed that the side-chain of this residue is directed toward the phosphate group (Tanaka et al. 2000), and site-directed mutagenesis experiments indicated that positively charged residues at this position maintain relatively high specific activity, and that residues with hydroxyl groups that can donate protons via hydrogen bonding are preferred (Ohgi et al. 1995, 1996a). Naturally occurring substitutions at this position also confirm this idea; a few fungal and plant RNases present threonine and arginine in place of this conserved lysine (Tanaka et al. 2000). By analogy to the nomenclature used to define the subsites of RNase A (Richards and Wyckoff 1971), the active site described above has been named “P1 site” (Fig. 4.2b) and contains the catalytic residues and the residues involved in binding to the substrate’s phosphate group. Also following this analogy, two other binding sites can be defined. These sites, “B1” and “B2”, correspond to hydrophobic pockets on either side of the P1 site and constitute the binding sites for the bases of the nucleotides on each side of the scissile phosphodiester bond (Fig. 4.2b). RNase T2 enzymes are base-nonspecific, but individual enzymes show unique base preferences, as shown by the release of mononucleotides from RNA, the rates of hydrolysis of homopolynucleotides, and the rates of hydrolysis of dinucleotide phosphates (Irie 1999; Deshpande and Shankar 2002). Some RNases are classified as adenylic acid preferential (RNase Rh), guanylic acid preferential (RNase LE and RNase NW), or uridylic acid preferential (RNase MC1 and RNase CL1),

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while others show comparable rate of hydrolysis at any base position (RNase NT and EcRNase I). Base preference is determined by the residues that form the B1 and B2 sites. The crystal structure of RNase Rh in complex with 20 -AMP showed that the adenine base forms hydrogen bonds with an aspartate and stacking interactions with a tyrosine and a tryptophan residues in the B1 site (Irie 1999). The tryptophan residue corresponds to Trp49; thus, this amino acid has dual functions, participating in the catalytic reaction and in substrate recognition. A D51N RNase Rh mutant showed markedly decreased enzymatic activity toward ApU but not toward UpU, and the substitution of Asp51 by Asn caused the enzyme to become more guanine nucleotide-preferential, suggesting that Asp51 is important for base recognition (Ohgi et al. 1993). Other mutations in the Asp, Tyr, and Trp positions also resulted in enzymes with altered preferences with respect to the 50 position composition of dinucleoside phosphate substrates (Ohgi et al. 1996b, c, 1997b). The B2 site of RNase Rh also presents some amino acids that are highly conserved among RNase T2 enzymes, such as Phe101 and Pro92. In the B2 site of crystals of the RNase Rh/d(ApC) complex, the cytosine base is stacked with the side-chain of Phe101 (Irie 1999). Site-directed mutagenesis of Phe101 (Ohgi et al. 2000) suggested that the side-chain of this amino acid interacts with the B2 base probably by p/p or CH/p interactions, and that these contacts are important to maintain enzymatic activity. These mutations also affected the substrate preference of the B2 site. The same effects on base preference were also observed in sitedirected mutagenesis experiments that altered two other B2 site residues, Ser 93 and Gln32 (Ohgi et al. 2003; Sanda et al. 2005). Although these experiments targeted residues that are part of the B2 site, small effects on the base preference of the B1 site were also observed. There is not yet an explanation for these observations, but it has been speculated that indirect conformational changes are responsible for this effect (Sanda et al. 2005). In plant RNases, the Trp and Tyr residues of the B1 site are also conserved (Tanaka et al. 2000; Kawano et al. 2002; Kawano et al. 2006), and may form stacking interactions with the B1 base as reported for RNase Rh. In tomato RNase LE, the position corresponding to Asp51 is occupied by Asn, and RNase LE shows guanine preference. Mutation of this residue to Asp changes the substrate preference to adenylic as in RNase Rh (Ohgi et al. 1997a), suggesting that the base recognition at the B1 site is similar in plant and fungal enzymes. However, in RNase MC1, which has a strong preference for uracil in the 30 position of the substrate, but does not discriminate based on the composition of the substrate at the 50 position (Irie et al. 1993), the B1 site is less conserved. The presence of Ser instead of Tyr in this position results in the lack of a hydrophobic pocket and probably explains the low selectivity in the B1 site (Nakagawa et al. 1999). Similar changes observed in S-RNases (Ida et al. 2001; Matsuura et al. 2001), which result in single-sided stacking, could also explain the lack of selectivity of these enzymes at this position (Matsuura et al. 2001). In E. coli RNase I, the B1 site is quite different from the one described in plants and fungi. In this nonselective RNase, only the Trp residue is conserved, and forms

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a one-sided stacking with the base. All other interactions, including van der Waals contacts and hydrogen bonds, are unique to E. coli and its closer relatives (Rodriguez et al. 2008). In addition, three water molecules at the bottom of the B1 cleft provide all the hydrogen bonds required for binding to the base (except one involving the side-chain of a B1 residue). Thus, this nonspecific binding site uses a series of water bridges to allow recognition of different bases in combination with base stacking to achieve broad specificity while maintaining good affinity (Rodriguez et al. 2008). This mechanism is different from the binding mechanism observed in the B1 site of other nonselective RNases like RNase NT from tobacco, which uses a series of different hydrogen bonds directly between protein and base (Kawano et al. 2006). Most mononucleotide complexes of plant RNases show the B2 site occupied. In most cases, a Phe residue, corresponding to Phe101 of RNase Rh, is conserved and participates in stacking interactions with the base, as described for the fungal B2 site (Suzuki et al. 2000; Kawano et al. 2002, 2006). Additional stacking interactions and an extensive network of hydrogen bonds between the base and side-chains of conserved and variable residues located in the B2 site provide additional contacts responsible for base preference. Mutation of amino acids involved in these hydrogen bonds resulted in enzymes with altered specificity (Numata et al. 2001, 2003). In some cases, the size of the B2 site also contributes to base preference. For example, mutation of an Asp in the B2 site of uridylic-preferential RNase MC1 to either Thr or Ser resulted in an enlarged site able to accept a guanine (Numata et al. 2003), and the large size of the B2 site of RNase NT has been proposed as one of the determinants of the purine preference of this enzyme (Kawano et al. 2006). The amino acids forming the B2 site of RNase I are well conserved among bacteria sequences but are different to those in plants and animal. However, stacking interactions (including a Phe residue) and an extensive hydrogen bond network are also observed in this site. Additionally, a water molecule bridging a B2 amino acid side-chain and a base group has also been observed (Rodriguez et al. 2008). The bacterial enzyme is the only one that has been crystallized with an oligonucleotide as a substrate mimic [the decadeoxynucleotide d(CGCGATCGCG)]. The structures of EcRNase I in its free and ligand-bound state are very similar, without any significant difference in backbone conformation, indicating a lock-and-key type of binding (Rodriguez et al. 2008). RNase T2 enzymes are generally regarded as having an acidic pH preference, between pH 4 and 5.5 (Irie 1999; Deshpande and Shankar 2002; Luhtala and Parker 2010). This is certainly the case of the fungal RNases first identified and characterized: RNase T2 optimum pH is 4.5 (Uchida 1966), and RNase Rh prefers pH 5.0 (Tomoyeda et al. 1969); and this is also true for a number of RNase T2 enzymes found in fungi and plants [summarized by Deshpande and Shankar (2002)], and also in animals (Inokuchi et al. 1997; Kusano et al. 1998; Suzuki et al. 2005; Campomenosi et al. 2006; Hillwig et al. 2009). However, many other RNase T2 enzymes have a near-neutral or basic pH preferences. For example, bacterial RNase I has a pH preference around 8.0 (Spahr and Hollingworth 1961),

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and plant S-RNases also prefer basic pH (McClure et al. 1989; Singh et al. 1991; Deshpande and Shankar 2002). Other RNases are more active near neutral pH, including viral Erns (Schneider et al. 1993), Arabidopsis RNS2 (Hillwig et al. 2011), and yeast RNY1 (G.C. MacIntosh, unpublished). The structural bases of these differences are unknown.

4.4

Biological Roles and Evolution

Members of the RNase T2 family are found in almost all groups of living organisms including virus, bacteria, fungi, plants, and animals (Irie 1999; Hillwig et al. 2009; MacIntosh et al. 2010), although they are not found in the Archaea (Condon and Putzer 2002). This family is particularly well conserved in eukaryotes; at least one gene has been found in every eukaryotic genome so far sequenced, with the exception of trypanosomes (Garcia-Silva et al. 2010; G.C. MacIntosh, unpublished) suggesting that RNase T2 enzymes perform an important biological role that has been conserved throughout evolution. Phylogenetic analyses of plant and animal RNase T2 proteins showed that this family has been very successful in plants, where it has undergone extensive expansion accompanied by high rates of gene duplication and gene loss, resulting in variable numbers of genes in different species (MacIntosh et al. 2010). On the other hand, RNase T2 has been maintained as a single copy gene in most animal species (Hillwig et al. 2009; R. Bailey and G.C. MacIntosh, unpublished). Evolutionary and expression analyses have found that plant RNase T2 genes can be classified in three classes (Igic and Kohn 2001; Steinbachs and Holsinger 2002; Roalson and McCubbin 2003; MacIntosh et al. 2010). Class I genes show tissue specificity and are generally regulated by stress. Gene duplication and deactivation occurring differentially among lineages [gene sorting (Zhang et al. 2000)] resulted in high diversification of Class I genes, possibly accompanied by the acquisition of novel functions after these duplication events (MacIntosh et al. 2010). Class III genes may have originated from a Class I gene, or at least they seem to share a common ancestor (Steinbachs and Holsinger 2002; Roalson and McCubbin 2003; MacIntosh et al. 2010). Class III genes correspond to the S-RNases and potential precursors of these enzymes (Igic and Kohn 2001; Steinbachs and Holsinger 2002; MacIntosh et al. 2010). S-RNases have a flower-specific role. In contrast to Classes I and III, Class II enzymes seem to have conserved more ancestral characteristics, and genes in this class are conserved in all plant species analyzed, with generally one Class II gene in each genome and an evolutionary pattern that follows organismal phylogenies. Most Class II genes are constitutively expressed (MacIntosh et al. 2010; K€ othke and K€ ock 2011). These evolutionary and expression characteristics suggested that plant Class II RNase T2 enzymes may have a housekeeping role (MacIntosh et al. 2010). Animal RNase T2 genes have not undergone extensive duplication events or diversification. Most animal genomes so far analyzed have only one RNase T2

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gene. A study on the phylogenetic relationships of RNase T2 proteins from deuterostomes showed that only one copy is present in each genome, with the exception of bony fishes that have two genes corresponding to this family (Hillwig et al. 2009), with one of the copies forming a fish-specific clade and the other belonging to the clade conserved in all chordates. Expression analyses showed the fish RNase in the conserved clade is expressed in all tissues and all developmental stages (Hillwig et al. 2009), and similar expression patterns have been reported for human RNASET2 (Henneke et al. 2009). These findings suggested that animal RNase T2 enzymes may be the counterpart of plant Class II RNases and also perform a housekeeping role (Hillwig et al. 2009) that could be conserved in plants and animals.

4.4.1

Cytotoxic Ribonucleases, Defense Responses, and Self-Incompatibility

While studies on RNase T2 enzymology go back more than half a century, little information on the biological role of these proteins, with a few exceptions, was known until recently. The first enzymes of this family with a definitive assigned function were the S-RNases. S-RNases are the pistil component of the mechanism of gametophytic self-incompatibility in three plant families (Solanaceae, Rosaceae, and Plantaginaceae). Self-incompatibility (S) prevents self-pollination, and in these families is determined by two components, the S-RNase in pistils and the S-locus F-box (SLF) protein expressed in pollen, that are genetically linked in the highly polymorphic S-locus. The different variants of the S-locus that occur in each organism are commonly known as haplotypes. If the haplotype of pollen, which is haploid, matches one of the two haplotypes of the diploid pistil, the pollen is recognized as self-pollen and pollen tube growth is inhibited. On the other hand, pollen that carries a haplotype different from the haplotypes of the pistil is recognized as nonself pollen and the pollen tube is allowed to grow through the style, resulting in fertilization of the ovule (reviewed by Kumar and McClure 2010; Meng et al. 2010). The pistil S-locus gene was originally cloned from Nicotiana alata, and described as a secreted glycoprotein (Anderson et al. 1986). Later, its homology to fungal RNase T2 enzymes was recognized, and its RNase activity was confirmed (McClure et al. 1989). Transgenic petunia plants expressing S-RNases with single amino acid mutations in the active site, which produced inactive enzymes, were used to demonstrate that the RNase activity is essential for rejection of self-pollen (Huang et al. 1994), and these experiments were confirmed by the identification of naturally occurring wild tomato mutants that had changes in the active site of S-RNases that resulted in lack of activity and loss of self-incompatibility (Royo et al. 1994). It has been shown that S-RNases are secreted from cells of the stigma, style, and ovary into the extracellular matrix that guides the pollen tube to the ovule

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(Cornish et al. 1987; Anderson et al. 1989), and they enter the pollen tubes, both in compatible and incompatible interactions (Luu et al. 2000; Goldraij et al. 2006). However, specific degradation of ribosomal RNA in pollen tubes occurs only during the incompatible interaction (McClure et al. 1990). Thus, it was hypothesized that the pollen S-locus component would inhibit S-RNase active during compatible interactions (Golz et al. 2001; Kao and Tsukamoto 2004). The identification of the S-pollen component proved more difficult; only recently, it was found that the product of the pollen S-locus is an F-box protein (SLF) that putatively is part of the multi-subunit E3 ubiquitin ligase complex (Qiao et al. 2004a, b; Sijacic et al. 2004). All current models for the mechanism of self-incompatibility agree that the S-RNase acts as a cytotoxin that inhibits pollen tube growth during incompatible interactions by degradation of rRNA in pollen. Specificity of this reaction is given by interaction of S-RNase with SFL. This interaction may result in the degradation of S-RNase in compatible but not in incompatible interactions, or in differential release of S-RNase from an intracellular compartment into the cytoplasm, depending on the model (Hua et al. 2008; Kumar and McClure 2010; Meng et al. 2010). Although the three plant families in which self-incompatibility is based on cytotoxic S-RNases are not closely related phylogenetically, most evidence suggests that this mechanism evolved only once and was the ancestral state in the majority of dicots (Igic and Kohn 2001; Steinbachs and Holsinger 2002; Vieira et al. 2008). The cytotoxic activity of S-RNases and their expression in flowers led to the hypothesis that gametophytic self-incompatibility evolved through the recruitment of an ancient flower ribonuclease involved in defense mechanisms against pathogens for the use in defense against “invasion” by self-pollen tubes (Lee et al. 1992; Hiscock et al. 1996; Nasrallah 2005). Plant RNase T2 proteins that do not participate in self-incompatibility are frequently called S-like RNases. Many of these S-like RNases have been, in fact, associated with defense responses. Most evidence for a defensive role in plants comes from gene expression analyses. For example, the expression of RNase NE, a S-like RNase from Nicotiana tabacum, is induced in response to the oomycete pathogen Phytophthora parasitica (Galiana et al. 1997). A highly similar gene from N. tabacum, RNase Nk1, is induced by cucumber mosaic virus infections (Ohno and Ehara 2005), and expression of RNase NW is induced in Nicotiana glutinosa plants infected with tobacco mosaic virus (Kurata et al. 2002). Microarray analyses indicated that several S-like RNase genes from rice are induced in response to infections by the bacterium Xanthomonas oryzae and the fungus Magnaporthe grisea (MacIntosh et al. 2010). These expression patterns suggested that RNase T2 enzymes could have an antimicrobial effect; however, only for RNase NE, this activity has been demonstrated. Addition of purified RNase NE inhibited hyphal growth from P. parasitica zoospores and from Fusarium oxysporum conidia in vitro, and infiltration of RNase NE into the extracellular space of tobacco leaves inhibited the development of P. parasitica in vivo (Hugot et al. 2002). This antimicrobial effect of RNase NE is dependent on the ribonuclease activity of the protein, since a protein with a point mutation in one of the active site His residues failed to inhibit hyphal growth in vitro (Hugot et al. 2002).

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This defense role supports the hypothesis that S-RNases evolved from an antimicrobial RNase. Even now, S-RNases may maintain a defense role in addition to their role in self-incompatibility. Petunia S-RNases are expressed in nectaries, the organ that produces floral nectar, together with other RNase T2 enzymes with intermediate characteristics between S- and S-like RNases (Hillwig et al. 2010a, b). Moreover, a proteomics analysis identified S-RNases as bona fide nectar proteins (Hillwig et al. 2010a). In nectar, these enzymes may also have an antimicrobial role to inhibit the growth of fungi and bacteria that would be favored in this rich medium. In fact, petunia nectar has a strong antibacterial activity (Hillwig et al. 2010b). Many S-like RNases are also induced in response to insect feeding and mechanical wounding, including Arabidopsis RNS1, tobacco RNase NW and RNase Nk1, tomato RNase LE, zinnia ZnRNaseII, and several rice and soybean RNase T2 genes (Ye and Droste 1996; Kariu et al. 1998; LeBrasseur et al. 2002; Gross et al. 2004; Bodenhausen and Reymond 2007; Hillwig et al. 2008; MacIntosh et al. 2010). Although the role of S-like RNases in the response to insects and wounding is not clear, it has been proposed that they could be antimicrobial enzymes that inhibit colonization by fungal, bacterial, and viral pathogens that use the wound as an entry site, or they could participate in phosphate remobilization during the healing process (LeBrasseur et al. 2002; Ohno and Ehara 2005). The extracellular nature of some of these RNases is consistent with an antimicrobial role (Jost et al. 1991; Bariola et al. 1999; Hugot et al. 2002). In addition to plant S-like RNases and S-RNases, other RNase T2 proteins have cytotoxic effects on different cells; however, in some cases, the mechanisms of action underlying cytotoxicity, and even the need for RNase activity, are not clear. Viral Erns acts as a virulence factor for pestiviruses. This protein is secreted from CSFV-infected cells (Rumenapf et al. 1993), and it displays cytotoxic effects against lymphocytes in cell cultures through induction of an apoptotic process (Bruschke et al. 1997); moreover, this effect is specific since Erns is not cytotoxic to epithelial cells. However, immunosuppression associated with virus persistence in the animal host seems to rely both on RNase activity of Erns and other protein features unrelated to its activity (Meyers et al. 1999; von Freyburg et al. 2004; Xia et al. 2007; Magkouras et al. 2008; Sainz et al. 2008; Tews et al. 2009). Omega-1, a glycoprotein secreted by eggs of the parasitic helminth Schistosoma mansoni, is an active RNase T2 enzyme (Fitzsimmons et al. 2005). This protein is the main elicitor of a potent CD4 T helper (Th) cell response that is necessary for parasite eggs to cross the endothelium and migrate to the intestinal lumen, from where they can exit the body to continue their lifecycle (Pearce 2005; Everts et al. 2009; Steinfelder et al. 2009). In immunosuppressed mice, omega-1 has a cytotoxic effect on hepatocytes, and it has been speculated that RNase activity is necessary for this effect (Dunne et al. 1991; Fitzsimmons et al. 2005). However, the ability of omega1 to promote Th2 lymphocyte differentiation is independent of its RNase activity (Everts et al. 2009; Steinfelder et al. 2009). Rny1, the only RNase T2 found in Saccharomyces cerevisiae (MacIntosh et al. 2001), is also involved in cytotoxic responses. Rny1 is localized to the vacuole and

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also secreted to the growth medium (MacIntosh et al. 2001; Thompson and Parker 2009); however, during oxidative stress, it is released from the vacuole to the cytoplasm, where it can cleave tRNA and rRNA (Thompson and Parker 2009), a function that clearly relies on its RNase activity. In addition, Rny1 is able to promote cell death through a mechanism that is independent of its catalytic activity (Thompson and Parker 2009).

4.4.2

Phosphate Scavenging and a Housekeeping Role in rRNA Recycling

The original discovery of RNase T2 as a secreted RNase from fungi suggested that it participates in scavenging of phosphate from RNA for nutritional purposes (Deshpande and Shankar 2002). Later, many plant RNases were found to be induced in response to phosphate starvation and during cell death processes when RNA is recycled, reinforcing this idea. The expression of two tomato RNases, RNase LE, and RNase LX, is induced when cultivated tomato cells or seedlings are grown in Pi-deficient media (Jost et al. 1991; Loffler et al. 1993; Kock et al. 1998, 2006). Two RNase T2 genes from Arabidopsis, RNS1, and RNS2, are also induced by Pi-starvation (Taylor et al. 1993; Bariola et al. 1994). The induction of RNase T2 genes as part of a phosphate scavenging system seems conserved in all plants. Other dicot genes regulated by Pi-starvation include tobacco RNase NE (Dodds et al. 1996), AhSL28 from Antirrhinum (Liang et al. 2002), and RNase PD2 from almond. Pi-starvation-regulated genes in monocots include OsRNS5, OsRNS7, and OsRNS8 from rice (MacIntosh et al. 2010), and WRN2 and WRN3 from wheat (Chang et al. 2005). Analysis of Arabidopsis seedlings grown on plates in which the only source of phosphate was RNA showed a strong increase in RNS1 activity and a weaker increase in RNS2 activity, and indicated that plants can use external RNA as a source of phosphate through the induction of a scavenging system (Chen et al. 2000). The presence of RNase T2 enzymes in the digestive liquid of the carnivorous plants Drosera adelae (Okabe et al. 2005) and Nepenthes ventricosa (Stephenson and Hogan 2006) also confirms a role in nutrition through phosphate scavenging for this family of RNases. While some of the enzymes induced by Pi-starvation are extracellular and could be released to the medium, such as RNase LE and RNS1 (Jost et al. 1991; Bariola et al. 1999), some intracellular proteins, such as RNS2 and RNase LX (Loffler et al. 1993; Hillwig et al. 2011), are also induced by this stress, suggesting that internal pools of RNA are also subjected to scavenging. In addition, the expression of RNS2 and RNase LX increases during senescence (Taylor et al. 1993; Lers et al. 1998; Lehmann et al. 2001), a regulation also observed for AhSL28 and for VRN1 from the green alga Volvox carteri (Shimizu et al. 2001; Liang et al. 2002). Moreover, ZRNaseI is expressed in the late stage of in vitro tracheary element differentiation

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in zinnia (Ye and Droste 1996) and RNase LX expression has been observed in xylem differentiation (Lehmann et al. 2001). These observations indicated that the corresponding proteins could be involved in recycling of phosphate during processes involving cell death. Antisense suppression of RNase LX expression resulted in delayed senescence and leaf abscission, suggesting that RNase LX may not only recycle phosphate during senescence and abscission, but could also participate in the control of these processes (Lers et al. 2006). Even more intriguing is the fact that some of the rice RNases induced by phosphate starvation (OsRNS5, OsRNS7) have accumulated mutations in the active site that most likely render the proteins inactive (MacIntosh et al. 2010). The function of these proteins is yet unknown. A scavenging role has also been proposed for non-plant RNase T2 enzymes. The edible mushroom Pholiota nameko grows well in phosphate-deficient conditions, and this characteristic was attributed to its ability to secrete RNases, including one RNase T2 enzyme, to the medium under Pi-limiting conditions (Tasaki et al. 2004). The human parasite Entamoeba histolytica is a nucleo-base auxotroph that needs to obtain purines and pyrimidines from its host. This parasite constitutively secretes two RNase T2 enzymes during axenic culture, and it was hypothesized that these proteins are utilized to scavenge nucleotides to support this organism’s growth (McGugan et al. 2007). The proposed housekeeping role of RNase T2 enzymes is also related to phosphate and/or nucleotide recycling. Although the Arabidopsis RNS2 gene is upregulated by Pi-starvation and during senescence, it is normally expressed at high levels in all tissues and developmental stages (Hillwig et al. 2011). Phylogenetic analysis determined that RNS2 belongs to the Class II category (Igic and Kohn 2001; MacIntosh et al. 2010), which suggested that it could have a housekeeping function. Fluorescently tagged protein was used to determine that RNS2 localizes to the endoplasmic reticulum and the vacuole (Hillwig et al. 2011). Mutants lacking RNS2 activity accumulate RNA intracellularly, mainly in the vacuole, and half-life analysis showed that rRNA decays more slowly in these mutants. Moreover, expression of RNS2 is necessary for maintenance of cellular homeostasis, since the rns2 mutants showed constitutive autophagy. Based on these results, it was suggested that the housekeeping role of RNase T2 enzymes is to participate in the degradation of ribosomes not only when plants are under nutritional stress but also under normal growth conditions (Hillwig et al. 2011). Cells lacking RNS2 activity may sense a nutritional imbalance that triggers autophagy to compensate for the lack of normal rRNA degradation. This housekeeping role seems to be conserved in all eukaryotes. The zebrafish genome contains two RNase T2 genes, with only one, RNASET2, conserved in other vertebrates (Hillwig et al. 2009). The RNASET2 protein of zebrafish was also found in the endoplasmic reticulum and lysosomes (Haud et al. 2011). Similarly, the homologous human protein, also RNASET2, is found in lysosomal fractions (Campomenosi et al. 2006). Mutant zebrafish that lack RNASET2 activity had enlarged lysosomes that accumulated rRNA in brain cells, and RNASET2 depletion in HEK 293 cells resulted in increased lysosome biogenesis (Haud et al. 2011). Moreover, magnetic resonance microimaging of mutant zebrafish revealed white

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matter lesions that may be the results of altered lysosomal function, suggesting that lack of RNASET2 also affects cellular homeostasis in this system (Haud et al. 2011). RNASET2 mutation in humans is linked with a leukoencephalopathy characterized by cortical cysts and multifocal white matter lesions (Henneke et al. 2009), similar to those found in zebrafish; these findings indicate that RNASET2 function is conserved in fish and humans. In these cases, it was also proposed that RNASET2 functions in normal rRNA decay (Haud et al. 2011). Analysis of mutant yeast cells lacking Rny1 activity also detected deficiencies in rRNA cleavage, although in this case, the effect was only observed when cells were subjected to oxidative stress, when a portion of this protein is released from the vacuole to the cytoplasm (Thompson and Parker 2009). Whether Rny1 participates in rRNA decay under normal conditions is not known.

4.4.3

Catalysis-Independent Functions and Cancer

Several lines of evidence point to a role for RNASET2 as a regulator of cell growth that can affect tumor progression in humans. The RNASET2 gene maps to a region of chromosome 6 that is frequently deleted in a variety of cancer cells (Trubia et al. 1997). In an extensive survey, no mutations were found in the RNASET2 gene in ovarian tumor tissues, but its expression was significantly reduced in 30% of primary ovarian tumors and in 75% of ovarian tumor cell lines (Acquati et al. 2001, 2005). Similarly, reduction of RNASET2 expression was found in lymphomas and melanomas (Steinemann et al. 2003; Monti et al. 2008). Cancer cell lines transfected with RNASET2 showed a reduction in clonogenicity, and reduced development of tumors and metastatic potential of cell lines in nude mice models was observed (Acquati et al. 2001, 2005; Smirnoff et al. 2006). While the effect of RNASET2 on growth of cancer cell cultures in vitro is controversial (Liu et al. 2002), a consistent effect as tumor antagonizing/malignancy suppressor has been found for this protein in vivo (Acquati et al. 2011). Remarkably, the tumor suppression function of RNASET2 is independent of its catalytic activity. Versions of the RNASET2 gene, in which the sequence encoding the active site histidines were mutated, were also effective in reducing tumorigenesis and metastasis (Acquati et al. 2005); and similar results were obtained with heat inactivated protein (Smirnoff et al. 2006). Moreover, the antitumorigenic, antiangiogenic, and antimetastatic effects of ACTIBIND, an RNase T2 protein isolated from Aspergillus niger, are also independent of its catalytic activity (Roiz et al. 2006; Schwartz et al. 2007). It has been shown that ACTIBIND is internalized by tumor cells (Schwartz et al. 2007). Both RNASET2 and ACTIBIND are able to bind actin, and ACTIBIND can disrupt the internal actin network and reduce cell motility (Roiz et al. 2006; Smirnoff et al. 2006). The catalysis-independent mechanism of action of these proteins in tumor suppression is not understood. In addition to binding the actin network, internalized ACTIBIND can be found localized in the nucleus, where it may act as a competitive

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inhibitor of angiogenin (Schwartz et al. 2007). Inhibition of angiogenin effects have also been reported for RNASET2 (Smirnoff et al. 2006); and it is known that in addition to its lysosomal localization, RNASET2 is secreted from cells to the extracellular space, where it could then find its target cells (Acquati et al. 2005). Overexpression of RNASET2 in SV40-immortalized fibroblasts resulted in reduction of colony-forming efficiency and growth rate accompanied by reduction in the expression of transcripts involved in Akt signaling, cell cycle control, and cell proliferation (Liu et al. 2010). Finally, RNASET2’s control of ovarian tumorigenesis seems to be the result of modification of the cellular microenvironment and the induction of immunocompetent cells of the monocyte/macrophage lineage (Acquati et al. 2011). Thus, it is possible that the tumor suppression effect is exerted by these proteins at several cellular levels. In addition to these antitumor roles, other catalysis-independent roles for RNase T2 proteins have been reported or hypothesized. RNase activity is not necessary for modulation of cell viability in yeast by Rny1 during oxidative stress responses (Thompson and Parker 2009). Some of the immunosuppressive effects of viral Erns might also be independent of RNase activity (Luhtala and Parker 2010); it has been hypothesized that an antimicrobial role of plant RNases may rely in part on properties of RNase T2 proteins separate from their catalytic activity (MacIntosh et al. 2010). In fact, several highly expressed plant members of the RNase T2 family have accumulated mutation in the active site resulting in inactive proteins and expression of these proteins is regulated by a variety of biotic and abiotic stress conditions (Gausing 2000; Chang et al. 2003; Wei et al. 2006; MacIntosh et al. 2010). The biological role of these plant inactive RNase T2 proteins is unknown. Inactive RNases are also used as storage proteins in plants; for example, an inactive RNase T2 protein is the main vegetative storage protein of resting rhizomes of Calystegia sepium (Van Damme et al. 2000). In this case, RNase activity may be dispensable since the protein is only used as a nitrogen reservoir.

4.5

Conclusion

The presence of the RNase T2 family in prokaryotes and eukaryotes and the almost absolute conservation in the former tells us that these ancient proteins have important functions. The last few years have brought advances in our understanding of this family. However, many questions remain to be answer to comprehend the mechanisms in which RNase T2 enzymes participate. Results from zebrafish and Arabidopsis link RNase T2 enzymes to ribosomal RNA decay; but the mechanism that determines when and why rRNA is targeted for degradation and how this substrate comes together with the enzyme needs to be investigated. The same is true for the effects of human RNASET2 and ACTIBIND on cell proliferation and tumor suppression. How do these proteins enter their target cells? What are their intracellular targets? How do they regulate those targets? Uncovering the properties of these proteins that allow them to regulate cell growth independently of their

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catalytic activity will have an important impact on future antitumorigenic therapies. It will be also important to investigate whether plant proteins that have lost their enzymatic activity have maintained a function as regulators of cell growth. The diversification of the RNase T2 family in plants also presents unique opportunities to learn more about the acquisition of new protein functions after gene duplication events. These are just a few of the questions that should keep us engaged in the exploration of this ancient and fascinating protein family.

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Ohgi K, Horiuchi H, Watanabe H, Iwama M, Takagi M, Irie M (1992) Evidence that three histidine residues of a base non-specific and adenylic acid preferential ribonuclease from Rhizopus niveus are involved in the catalytic function. J Biochem 112:132–138 Ohgi K, Horiuchi H, Watanabe H, Iwama M, Takagi M, Irie M (1993) Role of Asp51 and Glu105 in the enzymatic activity of a ribonuclease from Rhizopus niveus. J Biochem 113:219–224 Ohgi K, Iwama M, Tada K, Takizawa R, Irie M (1995) Role of Lys108 in the enzymatic activity of RNase Rh from Rhizopus niveus. J Biochem 117:27–33 Ohgi K, Iwama M, Ogawa Y, Hagiwara C, Ono E, Kawaguchi R, Kanazawa C, Irie M (1996a) Enzymatic activities of several K108 mutants of ribonuclease (RNase) isolated from Rhizopus niveus. Biol Pharm Bull 19:1080–1082 Ohgi K, Takeuchi M, Iwama M, Irie M (1996b) Enzymatic properties of mutant enzymes at Trp49 and Tyr57 of RNase Rh from Rhizopus niveus. J Biochem 119:9–15 Ohgi K, Takeuchi M, Iwama M, Irie M (1996c) Enzymatic properties of mutant forms of RNase Rh from Rhizopus niveus as to Asp51. J Biochem 119:548–552 Ohgi K, Shiratori Y, Nakajima A, Iwama M, Kobayashi H, Inokuchi N, Koyama T, Kock M, Loffler A, Glund K, Irie M (1997a) The base specificities of tomato ribonuclease (RNase LE) and its Asp44 mutant enzyme expressed from yeast cells. Biosci Biotechnol Biochem 61:432–438 Ohgi K, Takeuchi M, Iwama M, Irie M (1997b) Enzymatic properties of double mutant enzymes at Asp51 and Trp49 and Asp51 and Tyr57 of RNase Rh from Rhizopus niveus. Biosci Biotechnol Biochem 61:1913–1918 Ohgi K, Kudo S, Takeuchi M, Iwama M, Irie M (2000) Enzymatic properties of phenylalanine101 mutant enzyme of ribonuclease Rh from Rhizopus niveus. Biosci Biotechnol Biochem 64:2068–2074 Ohgi K, Iwama M, Inokuchi N, Irie M (2003) Enzymatic properties of glutamine 32 mutants of RNase Rh from Rhizopus niveus, a trial to alter the most preferential inter-nucleotidic linkages of RNase Rh. Biosci Biotechnol Biochem 67:570–576 Ohno H, Ehara Y (2005) Expression of ribonuclease gene in mechanically injured or virusinoculated Nicotiana tabacum leaves. Tohoku J Agric Res 55:99–109 Okabe T, Iwakiri Y, Mori H, Ogawa T, Ohyama T (2005) An S-like ribonuclease gene is used to generate a trap-leaf enzyme in the carnivorous plant Drosera adelae. FEBS Lett 579:5729–5733 Oxley D, Bacic A (1996) Disulphide bonding in a stylar self-incompatibility ribonuclease of Nicotiana alata. Eur J Biochem 242:75–80 Oxley D, Munro SLA, Craik DJ, Bacic A (1996) Structure of N-glycans on the S3- and S6-stylar self-incompatibility ribonucleases of Nicotiana alata. Glycobiology 6:611–618 Parry S, Newbigin E, Currie G, Bacic A, Oxley D (1997) Identification of active-site histidine residues of a self-incompatibility ribonuclease from a wild tomato. Plant Physiol 115:1421–1429 Parry S, Newbigin E, Craik D, Nakamura KT, Bacic A, Oxley D (1998) Structural analysis and molecular model of a self-incompatibility RNase from wild tomato. Plant Physiol 116:463–469 Pearce EJ (2005) Priming of the immune response by schistosome eggs. Parasite Immunol 27:265–270 Qiao H, Wang F, Zhao L, Zhou J, Lai Z, Zhang Y, Robbins TP, Xue Y (2004a) The F-box protein AhSLF-S2 controls the pollen function of S-RNase-based self-Incompatibility. Plant Cell 16:2307–2322 Qiao H, Wang H, Zhao L, Zhou J, Huang J, Zhang Y, Xue Y (2004b) The F-box protein AhSLF-S2 physically interacts with S-RNases that may be inhibited by the ubiquitin/26 S proteasome pathway of protein degradation during compatible pollination in antirrhinum. Plant Cell 16:582–595 Richards FM, Wyckoff HW (1971) Bovine pancreatic ribonuclease. In: Boyer PD (ed) The enzymes, vol 4. Academic, New York, pp 647–806 Richman A, Broothaerts W, Kohn J (1997) Self-incompatibility RNases from three plant families: homology or convergence? Am J Bot 84:912–917

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Suzuki R, Kanno S, Ogawa Y, Iwama M, Tsuji T, Ohgi K, Irie M (2005) On a salmon (Oncorhynchus keta) liver RNase, belonging to RNase T2 family: primary structure and some properties. Biosci Biotechnol Biochem 69:343–352 Tanaka N, Arai J, Inokuchi N, Koyama T, Ohgi K, Irie M, Nakamura KT (2000) Crystal structure of a plant ribonuclease, RNase LE. J Mol Biol 298:859–873 Tasaki Y, Azwan A, Hara T, Joh T (2004) Structure and expression of a phosphate deficiencyinducible ribonuclease gene in Pholiota nameko. Curr Genet 45:28–36 Taylor CB, Bariola PA, Delcardayre SB, Raines RT, Green PJ (1993) RNS2: a senescenceassociated rnase of Arabidopsis that diverged from the S-rnases before speciation. Proc Natl Acad Sci USA 90:5118–5122 Tews BA, Schurmann EM, Meyers G (2009) Mutation of cysteine 171 of pestivirus Erns RNase prevents homodimer formation and leads to attenuation of classical swine fever virus. J Virol 83:4823–4834 Thompson DM, Parker R (2009) The RNase Rny1p cleaves tRNAs and promotes cell death during oxidative stress in Saccharomyces cerevisiae. J Cell Biol 185:43–50 Tomoyeda M, Eto Y, Yoshino T (1969) Studies on ribonuclease produced by Rhizopus sp. I. Crystallization and some properties of the ribonuclease. Arch Biochem Biophys 131:191–202 Trubia M, Sessa L, Taramelli R (1997) Mammalian Rh/T2/S-glycoprotein ribonuclease family genes: cloning of a human member located in a region of chromosome 6 (6q27) frequently deleted in human malignancies. Genomics 42:342–344 Uchida T (1966) Purification and properties of RNase T2. J Biochem 60:115–132 Van Damme EJM, Hao Q, Barre A, Rouge P, Van Leuven F, Peumans WJ (2000) Major protein of resting rhizomes of Calystegia sepium (hedge bindweed) closely resembles plant RNases but has no enzymatic activity. Plant Physiol 122:433–445 Vieira J, Fonseca NA, Vieira CP (2008) An S-RNase-based gametophytic self-incompatibility system evolved only once in eudicots. J Mol Evol 67:179–190 von Freyburg M, Ege A, Saalmuller A, Meyers G (2004) Comparison of the effects of RNasenegative and wild-type classical swine fever virus on peripheral blood cells of infected pigs. J Gen Virol 85:1899–1908 Watanabe H, Fauzi H, Iwama M, Onda T, Ohgi K, Irie M (1995) Base non-specific acid ribonuclease from Irpex lacteus, primary structure and phylogenetic relationships in RNase T2 family enzyme. Biosci Biotechnol Biochem 59:2097–2103 Wei JY, Li AM, Li Y, Wang J, Liu XB, Liu LS, Xu ZF (2006) Cloning and characterization of an RNase-related protein gene preferentially expressed in rice stems. Biosci Biotechnol Biochem 70:1041–1045 Xia YH, Chen L, Pan ZS, Zhang CY (2007) A novel role of classical swine fever virus Erns glycoprotein in counteracting the newcastle disease virus (NDV)-mediated IFN-beta Induction. J Biochem Mol Biol 40:611–616 Ye ZH, Droste DL (1996) Isolation and characterization of cDNAs encoding xylogenesisassociated and wounding-induced ribonucleases in Zinnia elegans. Plant Mol Biol 30:697–709 Zhang JZ, Dyer KD, Rosenberg HF (2000) Evolution of the rodent eosinophil-associated RNase gene family by rapid gene sorting and positive selection. Proc Natl Acad Sci USA 97:4701–4706

Chapter 5

Stress-Induced Ribonucleases Pavel Ivanov and Paul Anderson

Contents 5.1 Stress-Induced Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.2 General Remarks About Stress-Induced Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.3 Stress-Induced Ribonucleases of Unicellular Organisms . . . . . . . . . . . . . . . . . . . . . . . . 5.1.4 Stress-Induced Ribonucleases of Multicellular Organisms . . . . . . . . . . . . . . . . . . . . . . 5.1.5 Concluding Remarks and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Ribonucleases are key regulators of RNA metabolism that play indispensable roles in cell physiology. A subset of ribonucleases that are regulated by stress help cells to adapt to adverse environmental conditions. Here, we review the structural and functional properties of stress-induced ribonucleases. We also summarize current thinking about the biological significance of stress-induced ribonucleases and discuss directions for future research.

5.1 5.1.1

Stress-Induced Ribonucleases Introduction

Cells are continually challenged by environmental changes that evoke acute or chronic stress. A network of cellular stress response programs operating at cellular

P. Ivanov (*) • P. Anderson (*) Division of Rheumatology, Immunology and Allergy, Brigham and Women’s Hospital, Boston, MA 02115, USA Department of Medicine, Harvard Medical School, Boston, MA 02115, USA e-mail: [email protected]; [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_5, # Springer-Verlag Berlin Heidelberg 2011

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and organismal levels helps cells to weather adverse environmental conditions to maintain cellular homeostasis. As a general feature, these control programs act at a global level by downregulating energy-intensive processes such as transcription and translation. As a consequence, stress reprograms gene expression to dampen the production of “housekeeping” proteins and selectively enhance the production of proteins that counter stress-induced damage and/or promote adaptation to altered conditions. Stress-induced reprogramming of gene expression involves the activation of a plethora of transcription factors that target genes involved in stress adaptation and repair of stress-induced damage. These transcription factors engage in intensive cross talk that can promote or suppress the expression of stress-response proteins. Although transcriptional programing produces a versatile set of expression profiles that can respond to a variety of internal or external stimuli, it cannot rapidly shift from one expression profile to another, a property required for a protective acute stress response. Rapid changes in protein expression are achieved by modulating the stability and translation of transcripts encoding stress-response proteins. Thus, posttranscriptional control mechanisms allow for rapid and flexible changes in protein expression that are required to protect cells from the deleterious effects of stress (Holcik and Sonenberg 2005). Ribonucleases (RNases) that catalyze the degradation of RNA into smaller fragments are ancient enzymes that regulate various aspects of RNA metabolism. While housekeeping ribonucleases play key roles in the maturation, quality control and turnover of cellular RNAs, stress-induced ribonucleases are activated in response to abiotic and biotic stresses. Cellular RNA levels are a function of both transcript synthesis and decay. Stress-induced activation or inactivation of RNases can rapidly alter RNA levels to alter cellular physiology. Here, we will describe the diversity of stress-induced ribonucleases and discuss their biological roles.

5.1.2

General Remarks About Stress-Induced Ribonucleases

Stress-induced ribonucleases are ancient enzymes found in most, if not all, organisms from prokaryotes to man (Table 5.1). Initially, part of an early “immune system” responsible for defending unicellular organisms from biotic stresses, such as viruses or nonself species, stress-induced ribonucleases, subsequently evolved to participate in a variety of biological processes. Induction of these ribonucleases by stress requires a tight control of their activity and/or their production under normal conditions. This is achieved by several mechanisms including physical compartmentalization/sequestration within membrane-bound organelles such as vacuoles or nuclei, secretion into the extracellular environment, or inactivation by RNase inhibitors (Fig. 5.1). Upon stress, intracellular stress-inducible RNases are released from sequestration and move into the cytoplasm where they gain access to cytosolic RNAs. In most cases, stress-induced RNases are present in large amounts in a latent

Colicin E5 RNA degradosome Ribotoxin RelE Ribotoxin YoeB Ribotoxin

Escherichia coli

Escherichia coli

Escherichia coli

RNase I RNase R ND ND ND

ND

Escherichia coli

Escherichia coli

Streptomyces coelicolor

Giardia lamblia Tetrahymena thermophila

Aspergillus fumigatus

Bacteria

Protozoa

Escherichia coli

MazF Ribotoxin ChpBK

Escherichia coli

Escherichia coli

RNase

Environmental changes Starvation Cold shock Starvation Developmental progression Starvation Encystation Starvation Starvation Developmental progression

Environmental changes

Environmental changes Environmental changes Heat shock Oxidative stress Amino acid starvation

Environmental changes

Cold shock

Suboptimal growing conditions

Bacteriophage infection Suboptimal growing conditions Suboptimal growing conditions

Inducer

tRNAs

tRNAs tRNAs

tRNAs

Structured RNAs

mRNAs rRNAs tRNAs

mRNAs

Ribosome-bound mRNAs

Ribosome-bound mRNAs

Jochl et al. (2008)

(continued)

Li et al. (2008) Lee and Collins (2005)

Haiser et al. (2008)

Chen and Deutscher (2005).

Ito and Ohnishi (1983)

Zhang et al. (2005)

Christensen-Dalsgaard and Gerdes (2008); Zhang et al. (2003)

Masaki et al. (1997); Masaki and Ogawa (2002) Polissi et al. (2003); Yamanaka and Inouye (2001) Christensen et al. (2001); Christensen and Gerdes (2003) Christensen-Dalsgaard and Gerdes (2008)

Reference Jiang et al. (2001, 2002) Masaki and Ogawa (2002) Ng et al. (2010)

Target tRNALys(UUU) tRNAArg 16 S rRNA tRNATyr(QUA) tRNAHis(QUG) tRNAAsn(QUU) tRNAAsp(QUC) mRNAs; Csp mRNAs

Taxonomy unit PrrC Colicin D Colicin E3

Table 5.1 Summary of stress-induced ribonucleases

Escherichia coli Escherichia coli Escherichia coli

Stress-Induced Ribonucleases

Organism

5 117

g-toxin

RNase LE

RNS1 RNase NE RNase LV-1, LV-2, LV-3; RNase LX

Kluyveromyces lactis

Solanum lycopersicum, Lycopersicum esculentum

Arbidopsis thaliana

Nicotiana alata

Homo sapiens; Cercopithecus acthiops; Mus musculus Homo sapiens, Mus musculus, Saccharomyces cerevisiae, etc. Homo sapiens Mus musculus, etc. RNase L

IRE1a

Angiogenin

Rny1

Saccharomyces cerevisiae

Lycopersicum esculentum

RNase

Organism

ND not determined

Vertebrate

Vertebrate, fungi

Vertebrate

Plants

Fungi

Taxonomy unit

Table 5.1 (continued)

ER stress Viral infection Oxidative and osmotic stress

Phosphate starvation Hypoxia Oxidative stress Heat Hypoxia UV light

Suboptimal growing conditions Phosphate starvation Mechanical injury Pathogen infection Phosphate starvation Mechanical injury Pathogen infection Phosphate starvation Mechanical injury

tRNAs

XBP-1 mRNA Viral RNAs Host rRNAs and mRNAs

tRNAs

Intracellular RNAs, rRNAs

Extracellular RNAs

Extracellular RNAs

rRNAs

rRNAs tRNAGlu(UUC) tRNALys(UUU) tRNAGln(UUG) Extracellular RNAs

Target

Inducer Oxidative stress, methionine and nitrogen starvation, heat, entry into stationary phase

Calfon et al. (2002; Yoshida et al. (2001) Pandey et al. (2004); Silverman (2007a)

Fu et al. (2009); Yamasaki et al. (2009)

Loffler et al. (1992)

Dodds et al. (1996)

Bariola et al. (1994)

Nurnberger et al. (1990)

Lu et al. (2008)

Thompson et al. (2008); Thompson and Parker (2009b)

Reference

118 P. Ivanov and P. Anderson

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Fig. 5.1 Stress-induced ribonucleases. Under normal conditions, stress-induced ribonucleases are inactivated by binding to specific ribonuclease inhibitors or by physical sequestration into membrane-bound organelles (e.g., nucleus, vacuoles, lysosomes, ER, periplasm (in bacteria)). Stress causes dissociation of ribonuclease from its inhibitor or releases sequestered ribonucleases into the cytoplasm where they cleave different RNA substrates (e.g., ribosome-free and ribosomebound mRNAs, tRNAs, and rRNAs). Secreted ribonucleases can scavenge extracellular RNAs or be taken up by adjacent cells. Upon internalization, secreted ribonucleases can exert cytotoxic or cytoprotective effects

state. Their activation allows rapid alterations in target RNA levels that can profoundly affect cellular physiology. In some cases, stress activates transcription of ribonuclease genes to upregulate ribonuclease expression. The synthesis of extracellular ribonucleases is tightly connected to their localization into secretory pathway organelles (vacuoles and lysosomes, in eukaryotes) or periplasm (in prokaryotes). Following their secretion, these ribonucleases can scavenge extracellular RNAs to provide nucleotides, or get internalized by other cells to activate cytotoxic or cytoprotective regulatory programs.

5.1.3

Stress-Induced Ribonucleases of Unicellular Organisms

Whereas multicellular organisms often have specialized cells or organs that function in the stress response, the survival of unicellular organisms relies on a molecular machinery that recognizes and responds to stress to promote cell survival. Here, we describe the role of stress-regulated ribonucleases in the stress-response program in these organisms.

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5.1.3.1


Cytotoxic Ribonucleases of Unicellular Organisms

Nuclease PrrC (EcoPrrC) found in the bacterium Escherichia coli is a latent tRNAspecific endoribonuclease that is activated in response to biotic stresses such as T4 bacteriophage infection (Kaufmann 2000). In uninfected cells, this ribonuclease physically associates with an enzyme complex comprising the host DNA restriction-modification system EcoprrI (encoded by prrA, prrB, and prrD genes of prr operon) (Levitz et al. 1990; Tyndall et al. 1994). Upon T4 bacteriophage infection, a phage-encoded peptide (Stp) binds to and inactivates EcoprrI (Amitsur et al. 1989, 1992, 2003). Paradoxically, Stp-induced inactivation of the host restriction-modification system activates PrrC ribonuclease. Activated PrrC cleaves bacterial tRNALys(UUU) directly 50 of a modified wobble uridine residue (5-methylaminomethyl-2-thiouridine) within the anticodon loop (Jiang et al. 2001, 2002). By depleting the tRNALys pool, PrrC disables both cellular and phage protein synthesis leading to cell suicide, and thus preventing the bacteriophage from spreading to adjacent bacteria. T4 phage counteracts this tRNA cleavage-mediated host defense by expressing an RNA repair system that rejoins cleaved tRNA fragments (Amitsur et al. 1987). Since PrrC homologues are found in diverse bacterial species, antiviral tRNA-specific endoribonucleases may be a common defense mechanism in diverse microorganisms. A similar phenomenon is also observed in eukaryotes in which RNaseL-mediated cleavage of self-RNA is required for the activation of antiviral interferon responses (Malathi et al. 2007; Silverman 2007a). Suboptimal growing conditions stimulate competition between different microorganisms for limited resources and often cause nutritional stress. In order to survive, some bacteria and fungi produce and secrete specific ribotoxins into the surrounding environment to promote irreversible growth arrest and/or death of nonself-species. For example, certain strains of the dairy yeast Kluyveromyces lactis secrete a heterotrimeric protein toxin that inhibits growth of non-self yeast species such as Saccharomyces cerevisiae (Jablonowski and Schaffrath 2007; Keppetipola et al. 2009; Lu et al. 2008). This killer toxin or zymogen consists of three subunits, a, b, and g, that are encoded on cytoplasmic episomes subject to cytoplasmic inheritance (Jablonowski and Schaffrath 2007). a- and b- subunits of the secreted zymogen interact with the cell wall of susceptible yeast cells and facilitate transport of the cytotoxic g-subunit into the cytoplasm of target cells. In the cytoplasm, g-toxin functions as an anticondon nuclease to selectively cleave tRNAGlu(UUC), tRNALys(UUU), and tRNAGln(UUG) between positions 34 and 35. All of these substrate tRNAs possess a 5-methoxycarbonylmethyl-2-thiouridine (mcm5s2U) residue at wobble position 34 of the anticodon loop providing a basis for cleavage specificity by g-toxin (Lu et al. 2008). As a consequence, g-toxin depletes pools of functional tRNAGlu, tRNALys, and tRNAGln, and causes growth arrest of target cells, thus eliminating non-self yeast cells from the competition for limited resources. Interestingly, a similar strategy to restrict the growth of competitor species is employed by the bacterium Escherichia coli. Certain strains of this bacterium carry

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Col plasmids that encode microbicidal proteins called colicins (Cascales et al. 2007). Colicins are a functionally diverse group of toxins that are encoded by a gene locus that includes the toxin gene and an immunity gene. The antitoxin gene encodes a protein that binds to colicin and inhibits its activity. Expression of colicin gene clusters is regulated by the SOS response, a major stress response program in prokaryotes that is activated by DNA damage (Cascales et al. 2007). Some colicins (e.g., colicins D, E3, and E5) are ribonucleases (Masaki and Ogawa 2002; Ng et al. 2010; Ogawa et al. 2006). Upon secretion into the medium, these colicins are able to translocate across the membrane of sensitive E.coli stains. Since cells of sensitive strains do not possess the immunity gene, they succumb to colicin-induced translational repression. While colicin E3 inhibits translation by cleavage of 16 S ribosomal RNA (Ng et al. 2010), colicins D and E5 cleave within the anticodon loop of tRNAs (Masaki et al. 1997; Masaki and Ogawa 2002; Ogawa et al. 2006). Colicin E3 binds to the A-site of the 70 S ribosome, the only place on the ribosome where 16 S rRNA is accessible, and cleaves between nucleotides A1493 and G1494 (Ng et al. 2010). While colicin D only hydrolyses isoacceptors of tRNAArg, colicin E5 specifically cleaves tRNAs that contain the modified nucleotide queosine in the wobble position of the anticodon loop, namely, tRNATyr(QUA), tRNAHis(QUG), tRNA,Asn(QUU) and tRNAAsp(QUC) (Masaki et al. 1997; Masaki and Ogawa, 2002; Ogawa et al. 2006). The use of colicin ribonucleases provides an example of a successful strategy used by bacteria to outcompete non-self species during unfavorable growth conditions.

5.1.3.2

Cytoprotective Ribonucleases of Unicellular Organisms

While some ribonucleases, such as colicins and PrrC nuclease, irreversibly alter the physiology of both host cells (bacterial suicide upon bacteriophage infection) and neighboring cells (causing cell death), other ribonucleases work as components of a stress-response program that reversibly alters cell metabolism to promote cell survival. The unicellular protozoan Giardia lamblia is an intestinal parasite that survives adverse environments by differentiating from a vegetative form into a dormant cyst (Adam 2001). The complex process of encystation is triggered by nutritional starvation encountered by the trophozoite (vegetative form) traversing the lower part of the host intestine (Adam 2001). During encystation, global tRNA cleavage without preferential hydrolysis of specific tRNA species is observed (Li et al. 2008). This cleavage occurs predominantly 50 of the anticodon loop to produce ~46 nucleotide RNA fragments. As less than 20% of tRNA is cleaved, global protein synthesis is only modestly affected (Li et al. 2008). However, this limited decrease of global translation can cause dramatic changes in gene expression and depresses cell metabolism as the cell enters the dormant stage of the protozoan life cycle. A similar tRNA cleavage phenomenon is observed in the protozoa Tetrahymena thermophila in response to starvation (Lee and Collins 2005), the bacterium Streptomyces coelicolor (Haiser et al. 2008), and the fungi Aspergillus fumigatus

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(Jochl et al. 2008) and Saccharomyces cerevisiae (Thompson et al. 2008; Thompson and Parker 2009b) in response to stress and/or during differentiation into resting states. However, in all these cases, cleavage occurs at or near to the anticodon loop of tRNA resulting in formation of two tRNA halves (Thompson and Parker 2009a). With the exception of budding yeast, the stress-induced RNases that cleave tRNA in these organisms have not been identified. In yeast, RNase Rny1, a member of the RNAse T2 family, cleaves tRNA and rRNA (5 S rRNA, 18 S rRNA and 25 S rRNA) in response to oxidative stress, heat shock, nitrogen starvation, or entry into stationary phase (Thompson et al. 2008; Thompson and Parker 2009b). This cleavage is not part of an RNA quality control pathway that degrades defective or incorrectly processed tRNAs. Instead, this endonucleolytic cleavage occurs on fully processed mature tRNAs in or near the anticodon loop. Notably, stress-induced cleavage is not a consequence of cell death since induction of apoptosis does not enhance tRNA or rRNA cleavage. Under normal conditions, Rny1 is sequestered within the vacuole. In stressed cells, Rny1 exits the vacuole to cleave cytoplasmic tRNAs (Thompson and Parker 2009b). Consequently, cytoplasmic Rny1 appears to reduce cell viability and even promote cell death in response to severe oxidative stress. The mechanism whereby Rny1mediated RNA cleavage causes reduced cell viability is not clear as rRNA/tRNA levels are not significantly depleted to inhibit translation. Moreover, a catalytically inactive mutant of Rny1 still affects cell survival suggesting that the ability of Rny1 to cleave RNA is separate from its ability to promote cell death (Thompson and Parker 2009b). Besides stress-induced cleavage of rRNA and tRNA, regulated changes in mRNA turnover also play a vital role in the adaptation to stress. In bacteria, regulating an mRNA’s accessibility to ribonucleases is a common means of modulating transcript stability and translatability under stress conditions (Anderson and Dunman 2009). Under normal conditions, bulk mRNA degradation in bacteria is catalyzed by the RNA degradosome (Bernstein et al. 2004), a dynamic complex of proteins that is proposed to bind to mRNA targets and catalyze both endonucleolytic cleavage and exonucleolytic decay (Rauhut and Klug 1999). The E.coli RNA degradosome (a typical bacterial RNA degradosome) consists of four core proteins: endoribonuclease E (RNase E), RNA helicase RhlB, enolase, and polynucleotide phosphorylase (PNPase) (Bernstein et al. 2004). RhlB is involved in loading the degradosome onto its mRNA target and removing secondary structures that hinder scanning to the cleavage site(s) (Liou et al. 2002). The initial mRNA cleavage is mediated by RNase E to produce mRNA fragments that are subsequently degraded by the 30 -50 -exonucleolytic activity of PNPase (Rauhut and Klug 1999). Enolase might participate in the degradation of mRNAs encoding metabolic enzymes (Chandran and Luisi 2006). The RNA degradosome is localized to the cell membrane where it communicates with other membrane-associated complexes (Liou et al. 2001; Taghbalout and Rothfield 2007, 2008). In response to stress, the RNA degradosome acquires new protein subunits. During cold shock conditions, the helicase RhlB is substituted for the cold shock RNA helicase CsdA (Prud’homme-Genereux et al. 2004). Additionally, another cold shock protein,

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CspE, can be recruited to the RNA degradosome (Feng et al. 2001). The remodeled RNA degradosome redirects its activity toward specific subsets of transcripts and, consequently, allows cells to adapt quickly to potentially lethal conditions. Interestingly, during the cold shock response, a transcript encoding PNPase is found within a small pool of upregulated mRNAs encoding cold shock proteins (Csp proteins 1–6) (Goldstein et al. 1990). Although the activity of PNPase is not changed during cold shock, this enzyme is specifically required to remodel the transcriptome from the poststress to the prestress state. After cold shock, PNPase specifically recognizes ribosomes translating Csp transcripts, degrades these transcripts and facilitates ribosome recycling (Polissi et al. 2003; Yamanaka and Inouye 2001). In addition to the cold shock response, PNPase is also involved in the oxidative stress response and the pH/acid shock response (Wu et al. 2009). In response to environmental changes, bacterial toxin–antitoxin systems allow cells to effectively shut down cellular processes. These systems are composed of closely linked genes encoding a stable toxin that can harm the cell and a short-lived antitoxin (Fozo et al. 2010). Under stress conditions, the level of antitoxin is quickly decreased causing activation of the cognate toxin. Some of these toxins act as mRNA endoribonucleases that bind to ribosomes and cleave transcripts undergoing translation (RelE (Christensen et al. 2001; Christensen and Gerdes 2003) and YoeB (Christensen-Dalsgaard and Gerdes 2008) toxins) or cleave mRNAs independently of their translation status (MazF (Christensen-Dalsgaard and Gerdes 2008; Zhang et al. 2003) and ChpBK (Zhang et al. 2005) toxins). As a consequence, these toxinmediated ribonucleases inhibit translation, and change gene expression patterns. Some of these toxin–antitoxin systems are modulated by a variety of stress conditions. For example, activation of MazF ribotoxin is modulated by different unrelated stress stimuli such as amino acid starvation, oxidative stress, heat shock, or antibiotic treatment. These conditions change the ratio of MazE antitoxin-MazF toxin levels in a ppGpp-dependent manner (Aizenman et al. 1996) (ppGpp is an alarmone produced during stress). Extracellular death factor EDF, a small peptide used for the regulation of gene expression in response to fluctuations in cellpopulation density, also influences the ratio of MazE/MazF levels (Kolodkin-Gal and Engelberg-Kulka 2008). Under severe stress conditions, derepression of MazF activity can cause cell death, and this altruistic programmed death is proposed to be beneficial for the bacterial population since dead cells provide required nutrients for their neighbors. In the absence of a specific protein inhibitor/regulator of ribonuclease activity, compartmentalization is often used as a means to regulate ribonuclease activity. For example, RNase I of E. coli is a nonspecific endoribonuclease that resides in the periplasmic space. In response to stress, RNase I moves from the periplasmic space into the cytoplasm where it degrades ribosomal and transfer RNA (Ito and Ohnishi 1983). Although exoribonuclease RNase R is considered to be a housekeeping enzyme involved in the degradation of highly structured RNA molecules such as rRNAs and tRNAs (Cheng and Deutscher 2005), its ribonuclease activity can be modulated by stress (Chen and Deutscher 2005). In response to cold shock or starvation, RNAse R

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activity is increased resulting in exonucleolytic degradation of structured transcripts including mRNAs (Chen and Deutscher 2005).

5.1.4

Stress-Induced Ribonucleases of Multicellular Organisms

5.1.4.1

Stress-Induced Ribonucleases in Plants

Plants have evolved a variety of morphological traits that facilitate survival under harsh environmental conditions. They have also evolved unique physiological responses to different stresses regulated by changes in hormonal balance. In some cases, this is achieved by altering RNase activity in response to endogenous and exogenous stimuli. Since inorganic phosphate (Pi) plays a fundamental role in the primary metabolism of plants, the availability of phosphates is one of the main determinants of plant growth. Several reports show that phosphate starvation can modulate expression and activity of plant ribonucleases. Several RNase T2 family ribonucleases are induced and often secreted from the cell upon phosphate starvation (Loffler et al. 1992; Nurnberger et al. 1990). Upon secretion, these ribonucleases are hypothesized to provide phosphates by scavenging extracellular nucleic acids. For example, both the expression level and the extracellular activity of RNase LE are increased during phosphate limitation in tomato cells (Solanum lycopersicum) (Nurnberger et al. 1990). Since induction of RNase LE is a function of extracellular but not intracellular Pi levels (Glund and Goldstein 1993), signal transduction pathway(s) sensing the availability of phosphates must be found on the plasma membrane. Similarly, phosphate starvation induces expression and extracellular activity of ribonucleases RNS1 and RNase NE in Arabidobsis thaliana and Nicotiana alata, respectively (Bariola et al. 1994; Dodds et al. 1996). A role for secreted extracellular ribonucleases in the scavenging of extracellular RNAs is supported by data showing that the addition of yeast tRNA to media can overcome phosphate starvation (Abel et al. 2000). Four other ribonucleases, three of which are vacuolar (RNase LV-1, RNase LV-2 and RNase LV-3) and one of which is endoplasmic reticulum associated (RNase LX), are co-induced together with RNase LE in response to Pi limitation (Loffler et al. 1992). Upon induction, these ribonucleases promote decay of intracellular RNA. Since vacuoles contain enzymes that degrade RNA (RNases, phosphodiesterases, and phosphatases) to nucleosides and Pi (Loffler et al. 1992), and contain short RNA oligonucleotides (Abel et al. 1990), these organelles may be specialized for intracellular RNA degradation. Uptake of RNA into vacuoles may occur by autophagy. Since expression of phosphate starvation–induced RNases is also observed during leaf senescence, this same subset of ribonucleases may promote phosphate remobilization by scavenging RNA from senescent organelles. Importantly, phosphate starvation–induced ribonucleases are not induced by reduced levels of nitrate or potassium (Bariola et al. 1994). However, induction

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of expression and activity of RNS1 and RNAse LE ribonucleases is observed in response to mechanical tissue injuries and in response to pathogen infection (LeBrasseur et al. 2002). In A. thaliana, induction of RNS1 expression is observed not only locally at the place of mechanical injury, but also at non-injured distal parts of the plant (LeBrasseur et al. 2002). This observation suggests that induction of ribonucleases might be a part of systemic stress response dedicated to phosphate remobilization, wound healing, and defense against pathogens.

5.1.4.2

Stress-Induced Ribonucleases of Vertebrates

Although certain ribonucleases such as RNase T2 family members are widely distributed and found in viruses, bacteria, protozoans, fungi, plants, and animals (Deshpande and Shankar 2002; Luhtala and Parker 2010), the RNase A family members are vertebrate specific (Cho et al. 2005; Lander et al. 2001). As a group, all RNase A family members are small secreted proteins, with a range of ribonucleolytic activities and cellular functions. RNase 5 or angiogenin, the only member of this family that is found outside of Mammalia (Cho et al. 2005), is a stress-induced ribonuclease with unique cellular functions (Fett et al. 1985). Angiogenin was initially isolated from the conditioned medium of human colon adenocarcinoma cells based on its ability to stimulate blood vessel formation in the chicken embryo chorioallantoic membrane angiogenesis assay (Fett et al. 1985). Further studies demonstrated that angiogenin is involved in tumor growth, and its expression and secretion are upregulated in many types of cancers (Olson et al. 1994, 1995; Tello-Montoliu et al. 2006). Mutational analysis shows that its ability to stimulate angiogenesis requires its ribonuclease activity as well as its ability to translocate into the nucleus where it accumulates in the nucleolus and stimulates rRNA transcription (Tsuji et al. 2005). Secreted angiogenin is readily taken up by target cells such as endothelial and smooth-muscle cells where it stimulates cell migration and invasion of the extracellular matrix, triggers second messenger production, promotes cell adhesion, and induces cell proliferation (Gao and Xu 2008). Angiogenin mutations that disrupt ribonuclease activity are found in patients suffering from amyotrophic lateral sclerosis, a fatal neurodegenerative disease (Greenway et al. 2004, 2006). Angiogenin is a stress-induced ribonuclease. Its expression is increased by hypoxia and acute inflammation (Hartmann et al. 1999; Nakamura et al. 2006; Olson et al. 1998). Since tumors are known to exist in a hypoxic microenvironment, the increase in both expression and secretion of angiogenin helps tumors to adapt to such conditions by stimulating new blood vessel formation to promote further tumor growth. Stressful stimuli also increase the activity of constitutively expressed angiogenin. Different stresses including heat shock, UV irradiation, and oxidative stress activate angiogenin to cleave tRNA (Fu et al. 2009; Yamasaki et al. 2009). The mechanism of angiogenin activation is not clear but it likely involves stressinduced inactivation of the angiogenin/ribonuclease inhibitor RNH1, angiogenin movement from nucleus to cytoplasm, or both. Angiogenin targets the anticodon

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loop of mature fully processed tRNA to produce 50 - and 30 -halves of tRNA with lengths of ~30 and 40 nucleotides, respectively (Fu et al. 2009; Yamasaki et al. 2009). These tRNA fragments have been designated as tiRNAs (50 - and 30 -tiRNAs, respectively) or tRNA-derived, stress-induced small RNAs (Yamasaki et al. 2009). Similar to the stress-induced tRNA cleavage mediated by Rny1 in yeast, angiogenin cleaves only a minor (less than 10%) fraction of mature tRNA without preferential cleavage of individual tRNA species or their isoforms (Fu et al. 2009; Thompson et al. 2008; Thompson and Parker 2009a; Yamasaki et al. 2009). However, in contrast to Rny1, angiogenin-induced tRNA cleavage causes transient inhibition of global protein synthesis and may promote cell survival. Inhibition of global protein synthesis in response to stress is an energy-saving mechanism that helps cells to survive under stressful conditions. Although regulation of translation can be exerted at any level, the well-characterized examples of translational regulation target the initiation machinery. This is achieved by exploiting the differential sensitivity of mRNAs to subtle changes in the availability or activity of general initiation factors such as eIF4E and eIF2a (Holcik and Sonenberg 2005; Yamasaki and Anderson 2008). As a consequence, selected mRNAs are preferentially translated during stress conditions, whereas the majority of transcripts are not. Translational reprogramming allows the production of proteins that repair stress-induced damage and promote cell survival or apoptosis (Holcik and Sonenberg 2005; Yamasaki and Anderson 2008). Phosphorylation of eIF2a is a major event in the global stress response pathway (Integrated Stress Response) that orchestrates stress-induced reprogramming at the posttranscriptional level (Holcik and Sonenberg 2005; Yamasaki and Anderson 2008). eIF2a phosphorylation promotes the assembly of stress granules (SGs), cytoplasmic foci at which translationally inhibited mRNPs are concentrated (Kedersha and Anderson 2002; Kedersha et al. 2002). Importantly, angiogenin and tiRNAs inhibit translation, and promote the assembly of SGs in a phosphoeIF2a-independent manner. Further analysis of stress-induced angiogeninmediated tRNA cleavage indicates that 50 - but not 30 -tiRNAs mediate inhibition of translation and stress granule assembly (Emara et al. 2010; Yamasaki et al. 2009). Since SGs play important roles in cellular adaptation and survival during stress (Anderson and Kedersha 2008; Arimoto et al. 2008), angiogenin/tiRNAinduced SG formation may contribute to the cytoprotective and pro-survival properties of angiogenin (Kieran et al. 2008; Subramanian et al. 2008). Significantly, an ALS-associated mutant of angiogenin (P112L mutant) does not efficiently induce tiRNA formation, translational repression, or stress granule assembly (Emara et al. 2010; Yamasaki et al. 2009). It is therefore possible that in ALS patients, the angiogenin-mediated stress response pathway is abrogated resulting in a failure to reprogram protein translation and to modulate expression of genes promoting cell survival and repair of stress-induced damage. Inositol requiring 1a (IRE1a) is a bifunctional transmembrane protein kinase/ endoribonuclease localized in the endoplasmic reticulum (ER) (Liu et al. 2002; Schroder 2008). Perturbations in ER, a cellular compartment where biosynthesis and folding of many secretory proteins occurs, lead to ER stress and disregulation

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of ER homeostasis. In turn, ER stress activates a complex cell signaling response collectively known as the unfolded protein response (UPR) that downregulates global protein synthesis and upregulates the expression of stress-responsive genes (Ron and Walter 2007; Rutkowski and Kaufman 2007). By decreasing overall cellular translation, cells decrease ER overload and promote degradation of abnormal misfolded proteins accumulated in the ER. In contrast, expression of genes coding for ER components, molecular chaperons, or proteins involved in the degradation of misfolded proteins helps normalize ER homeostasis (Schroder 2008). IRE1a ribonuclease is a central component of the UPR activated by ER stress (Calfon et al. 2002; Yoshida et al. 2001). Its C-terminus is localized in the cytoplasm, while its N-terminus is localized in the ER lumen where, in unstressed conditions, it interacts with the protein chaperone BiP. BiP also binds to unfolded/ misfolded proteins whereupon dissociation from IRE1a allows dimerization, autophosphorylation, and activation of IRE1a. In turn, activated IRE1a causes splicing of mRNA encoding X-box binding protein-1 (XBP-1) that leads to the synthesis and activation of transcription factor XBP-1. Activated XBP-1 upregulates expression of ER stress-responsive genes involved in protein folding, degradation, and secretion, thus restoring ER functions (Calfon et al. 2002; Yoshida et al. 2001). Ribonuclease L (RNase L) is one of the many proteins involved in the host response to viral infections. This ribonuclease is a central component of the innate cellular defense mechanism induced by type I interferons (Borden et al. 2007; Silverman 2007b). Under normal conditions, the amount of RNase L in cells is very low, but its expression is increased in response to interferon. RNase L is initially produced in an inactive form that is activated in response to virus infection. Its activation is dependent on the recognition of viral double-stranded RNA in the cytoplasm that stimulates production of unique unstable 20 -50 adenylic acid oligomers (20 -50 -A oligomers) from cellular ATP by 20 -50 oligoadenylate synthetase. In turn, 20 -50 -A oligomers bind to latent RNase L monomers causing their dimerization and activation (Silverman 2007a). Activated RNase L cleaves viral and host single-stranded RNAs. Interestingly, RNase L-mediated cleavage of host RNAs such as rRNA plays a signaling role where short self-RNAs are recognized by two pathogen recognition receptors, RIG-I (retinoic acid–inducible gene-I) and MDA5 (melanoma differentiation– associated gene-5), which further activate expression of alpha- and betainterferons (Malathi et al. 2007; Silverman 2007a). Moreover, depending on the level of viral infection, activation of RNase L can lead to the massive degradation of cellular RNA and consequently apoptosis (Castelli et al. 1997, 1998; Walczak et al. 2000; Zhou et al. 1997). Recent studies also show that activation of RNase L is not restricted to viral infections, but is also observed in response to oxidative and osmotic stress, indicating that RNase L is a broad range stress-responsive enzyme (Pandey et al. 2004). The biological significance of stress-induced RNase L activation requires further investigation, but might be connected to the ability of RNase L to promote apoptosis.

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5.1.5


Concluding Remarks and Perspectives

We describe here that stress-induced ribonucleases are a diverse group of proteins found in many organisms and involved in many biological processes. As the field matures, it is likely that additional biological functions of stress-responsive ribonucleases will be found. In many cases, however, the molecular details of stress-induced activation of ribonucleases and their functions are not well characterized. For example, the evolutionarily conserved response to stress that involves endonucleolytic cleavage of tRNA requires further investigation. An attractive possibility is that stress-induced tRNA fragments are directly involved in the regulation of gene expression in a manner similar to that of known small RNAs such as microRNAs, siRNAs, and piRNAs. This hypothesis needs experimental validation. In mammals, a number of different proteins have been shown to cleave specific mRNAs, and their activity can potentially be regulated by stress. For example, RAS GTPase-activating protein-SH3 domain binding protein (G3BP) and apurinic/ apyrimidinic endonuclease 1 (APE1) cleave c-myc mRNA (Barnes et al. 2009; Gallouzi et al. 1998); an estrogen-regulated polysomal endoribonuclease PMR1 cleaves vitellogenin and albumin mRNAs (Yang and Schoenberg 2004), and erythroid-enriched ribonuclease (ErEN) targets alpha-globin mRNA (Rodgers et al. 2002). Interestingly, in response to stress, both PMR1 and G3BP were shown to localize into stress granules (Tourriere et al. 2001; Yang et al. 2006), but whether localization into stress granules affects their ribonuclease activity is not known. APE1 is a key enzyme in the DNA damage response that repairs damaged or mismatched nucleotides in DNA. However, it is not known whether the endoribonuclease activity of APE1 is changed in response to stresses causing DNA damage such as oxidative stress. Further characterization of these endoribonucleases and their targets is required to understand the biological significance of endonucleolytic cleavage in the control of mRNA abundance and degradation. An improved understanding of stress-induced ribonucleases will have broad implications for both clinical and applied fields. For example, since many bacterial pathogens exposed to cold shock require stress-responsive ribonucleases for survival, interference with these enzymes could be a useful method to counter their toxic effects. The survival of bacteria in refrigerated food products is an example where inhibition of ribonuclease activity may have commercial applications. Further insights from studies of mammalian stress-induced ribonucleases may allow the development of new therapeutic agents targeting different diseases. For example, inhibition of angiogenin activity may promote the death of hypoxic cancer cells, while stimulation of angiogenin activity may promote survival of motor neurons in patients with amyotrophic lateral sclerosis. Modulation of IRE1a endoribonuclease activity, a regulatory checkpoint upstream of the unfolded protein response pathway, may be a useful strategy to approach diseases involving ER stress such as diabetes, cancer, neurodegenerative disorders, and cystic fibrosis. Clearly, further studies are required both to identify new stress-induced ribonucleases and to further characterize already known ones.

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Chapter 6

Viral RNase Involvement in Strategies of Infection Gregor Meyers, Tillmann R€ umenapf, and John Ziebuhr

Contents 6.1 RNases with Special Tasks in Viral Genome Replication and Gene Expression . . . . . . . 6.1.1 Endonucleases of Viruses with Segmented Negative-Strand RNA Genomes . . 6.1.2 RNases of Viruses Belonging to the Order Nidovirales . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Viral RNase Influencing Host Cellular Metabolism and the Host Immune Response: The Herpesvirus vhs Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Viral RNase Important for the Interaction Between Virus and Host Organism: The Pestivirus Erns RNase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

136 136 144 149 153 159

Abstract

The overwhelming majority of RNase activity is engaged in catabolic processes. Viruses have no metabolism of their own, but rely completely on host cellular energy and substrate provision to support the biochemical processes necessary for virus replication. It is therefore obvious that RNA hydrolysis does not represent an obligate step in the viral life cycle that would have to be governed by viral proteins. G. Meyers (*) Institut f€ur Immunologie, Friedrich-Loeffler-Institut, S€ udufer 10, D-17493 Greifswald – Insel Riems, Germany e-mail: [email protected] T. R€umenapf Institut f€ur Virologie, Fachbereich Veterin€armedizin, Justus-Liebig-Universit€at Giessen, D-35392 Giessen, Germany e-mail: [email protected] J. Ziebuhr Institut f€ur medizinische Virologie, Justus-Liebig-Universit€at Giessen, D-35392 Giessen, Germany e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_6, # Springer-Verlag Berlin Heidelberg 2011

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Accordingly, RNases are found only rarely in the viral proteomes and serve special functions. In this chapter, several virus-specific RNases will be described and their role in the viral life cycle discussed. The text will concentrate on RNases of members of the nidoviruses, herpesviruses, pestiviruses, and several viruses with segmented negative-strand RNA genome including influenza virus. These enzymes are involved in specific steps of viral gene expression, viral genome replication, shutoff of host cellular gene expression, and interference with the host’s immune response to virus infection.

6.1

RNases with Special Tasks in Viral Genome Replication and Gene Expression

6.1.1

Endonucleases of Viruses with Segmented Negative-Strand RNA Genomes

A group of negative-strand RNA viruses comprising the members of the families Orthomyxoviridae, Arenaviridae, and Bunyaviridae use a special type of RNA endonuclease for one step during production of their mRNAs. The genome segments of these viruses are associated with viral proteins giving rise to so-called ribonucleoprotein complexes (RNP) (Buchmeier 2007; Palese and Shaw 2007; Schmaljohn and Nichol 2007). These are replicated by a replicase complex with a core unit composed of 1–3 viral polypeptides. This replicase is also responsible for transcription of the viral mRNAs. In contrast to the de novo initiation of RNA synthesis during replication, transcription occurs as a primer-dependent process. 50 terminal segments of 1–15 nucleotides are cleaved from host cellular mRNAs by a viral endonuclease and serve as transcription primers carrying a 50 cap structure. Data on the endonucleases of the above-mentioned viruses are described in the following sections.

6.1.1.1

Influenza Virus

The influenza A virus of the family Orthomyxoviridae represents one of the best studied viruses. Influenza A virus particles contain a genome of eight RNA segments with negative polarity (Palese and Shaw 2007). The three largest segments 1, 2, and 3 encode the polypeptides PB2, PB1, and PA, respectively, which constitute the viral RNA–dependent RNA polymerase (RdRP) responsible for genome replication and mRNA transcription (Fig. 6.1). The RdRP is not only an RNA polymerase but is also responsible for polyadenylation of viral mRNA and the endonucleolytic generation of the transcription primers mentioned above. Initially, the endonuclease was thought to reside in PB2 (Shi et al. 1995), but recent results

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137 NA (tetramer) HA (trimer) M2 (tetramer) M1

Nucleocapsid segment

envelope membrane PB1 NP ssRNA genome PA endonuclease

PB2 cap binding

Fig. 6.1 On top, a schematic representation of an influenza A virus particle is shown with the envelope proteins neuraminidase (NA), hemagglutinin (HA), M1 and M2, and the nucleocapsid segments that are each composed of one of the eight single-stranded genome segments of negative polarity, a certain number of nucleoprotein (NP) molecules and a polymerase complex encompassing polypeptides PB1, PA, and PB2 (presentation of the scheme with kind permission from Springer Science and Business Media: Molekulare Virologie, chapter 16, 2010, page 357, Modrow et al. (2010), Fig. 16.6). As indicated, the PA protein contains the endonuclease domain, the active site of which is shown as a structure model at the bottom (left part). Helices and strands surrounding the active-site cleft are shown together with several residues important for coordination of the two catalytically active divalent metal cations (green spheres) shown as sticks (amino acids given in the one letter code). At the bottom on the right site, a structure model of the PB2

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provided evidence for PA harboring the enzymatic domain (Dias et al. 2009; Yuan et al. 2009). PA, a polypeptide of more than 700 amino acids, was expressed in insect cells in soluble form. Trypsin cleaves PA into N- and C-terminal domains of 25 and 55 kDa, respectively. The function of the latter is still unclear except for the fact that it mediates contacts with PB1 via a small C-terminal region (Ruigrok et al. 2010). The N-terminal ca. 250 amino acids of PA (PA-Nter) harbor the endonuclease activity of the influenza virus polymerase complex (Dias et al. 2009; Yuan et al. 2009). This fragment has been expressed in E. coli and subsequently crystallized. The overall structure shows an a/b architecture with a core of five mixed b strands forming a twisted plane surrounded by seven a-helices. Structure similarity searches gave no indications for entries with highly similar folds in the databases (Dias et al. 2009). The most similar structure identified in these searches was that of an archaeal Holliday junction resolvase (Hjc), obvious from superposition of helix a3 and strands b1–b5 encompassing a structural motif typically found in different nucleases including resolvases and type II restriction endonucleases (Knizewski et al. 2007; Kosinski et al. 2005). The motif conserved between Hjc and PA-Nter includes two acidic residues (Asp-108 and Glu-119 in the latter) that are known to be important for metal ion binding in the former enzyme. Residues Asp-108 and Glu-119 together with His-41 and Lys-134 of PA-Nter also align with catalytically important residues in the restriction endonuclease EcoRV (Glu -5, Asp-74, Asp-90 and Lys-92). Moreover, His-41, Asp-108, Glu-119, and Lys-134 together with Glu80, a third acidic residue in the active site, are conserved among influenza viruses. Members of the PD-(D/E)XK family of enzymes are known to contain up to three coordinately bound divalent metal ions that are involved in the catalytic process (Knizewski et al. 2007; Kosinski et al. 2005). Metal ions were also found in the influenza virus endonuclease. Within a negatively charged cavity surrounded by helices a2–a5 and strand b3, Yuan and coworkers identified an Mg2+ ion (Yuan et al. 2009). Dias et al. reported the presence of Mn2+ at the same position. In ˚ away suggesting a twoaddition, the latter group detected a second Mn2+ about 4 A metal based catalytic mechanism. A preference for Mn2+ over other divalent cations was found in stability and functional assays (Dias et al. 2009; Doan et al. 1999). Mutagenesis analyses supported the model described above. Yuan and coworkers mutated the residues H-41, E-80, L-106, P-107, D-108, and E-119 proposed to be important for metal ion coordination (Fig. 6.1) (Yuan et al. 2009). In addition, K-134 was also included in these experiments because this residue is located close to the proposed active site of the endonuclease, and exchanges at this position had

Fig. 6.1 (continued) region binding the 50 cap structure is shown. Again, important residues engaged in binding the m7GTP group of the cap via stacking of ring structures are shown as sticks. The PB2 moiety of the polymerase confers the specificity for the endonucleolytic cleavage of capped substrate RNAs. (The schemes of the structures are reprinted from publication Ruigrok et al. 2010, with permission from Elsevier)

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been shown before to specifically block transcription but not replication of the viral RNA (Hara et al. 2006). Polymerases with mutations E80A, D108A, E119A, and K134A retained significant genome replication efficiency, but were severely hampered in mRNA synthesis. The abrogation of transcription activity was indeed a consequence of blocked endonucleolytic activity of the polymerase since the mutants were able to perform transcription in the presence of ApG as a substitute for the cap-primer or after addition of a capped primer itself. In contrast, the L106A and P107A mutants retained some transcription activity but lost the ability to perform genome replication, while the H41A mutant was unable to perform RNA synthesis in general. The latter effects could not be circumvented in ApG-primed transcription assays. Finally, a direct endonuclease cleavage assay proved that all three exchanges affecting the acidic residues and the K134A mutation blocked the endonucleolytic activity (Yuan et al. 2009). Taken together the data proved that the endonucleolytic activity in the influenza virus RNA polymerase resides in the N-terminal region of the PA polypeptide and that the respective enzyme belongs to the PD-(D/E)XK family of nucleases with the characteristic motif occurring at 107-PDLYDYK. Special features of the PA-Nter with regard to the other family members are the close vicinity of the two acidic residues in the conserved motif, an unusual position of the presumably catalytically important lysine (Lys-134), the histidine residue in the active site and the fact that RNA is the substrate instead of DNA. A somewhat surprising result of the structure analysis was the fact that it did not provide information on how the RNA substrate could access the active site of the enzyme as it does not contain obvious determinants for RNA binding (Dias et al. 2009; Yuan et al. 2009). In fact, the endonuclease domain has a negatively charged surface with an even more acidic cavity containing the active site. The contact to the substrate RNA is presumably made by a number of positively charged residues that are located on the rim around the active site cavity. These residues are conserved and should help to position the substrate for cleavage. As mentioned above, within the PD-(D/E)XK family of nucleases, the influenza virus endonuclease is special for its substrate representing RNA. It has been shown that the endonuclease cleaves also single-stranded DNA with only slightly reduced activity (Klumpp et al. 2000), but the natural substrate of this enzyme consists of a 50 terminally capped RNA. The structural data do not provide any evidence how the substrate specificity for a 50 capped RNA could be achieved. Experimental evidence has been provided for PB2 representing the cap-binding domain of the polymerase (Guilligay et al. 2008). A structure of the PB2 residues 320–483 crystallized with m7GTP allowed identification of the residues responsible for binding the methylated guanine which are absolutely conserved among influenza A viruses. These residues include His-357 and Phe-404, which sandwich between their ring systems the purine of the methylated base, and Glu-361 and Lys-376 that specifically recognize the guanine base (Fig. 6.1). The mode of cap binding is very similar to what has been described for the nuclear cap-binding complex and the capbinding translation initiation factor eIF4E found in the cytoplasm (Guilligay et al. 2008; Fechter et al. 2003; Li et al. 2001).

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Fig. 6.2 Schematic diagram showing steps in cap-dependent transcription by influenza virus polymerase. (a) Binding of host pre-mRNAs (yellow) by the cap-binding domain located in the PB2 subunit. (b) Cleavage of the host mRNA 10–13 nucleotides downstream of the 50 cap by the endonuclease located in the PA subunit. (c) Elongation of the chimeric viral mRNA by the nucleotidyl-transferase site in the PB1 subunit using the vRNA as template. (d) Polyadenylation of the viral mRNA by polymerase stuttering at the oligo-U sequence near the 50 end of the vRNA. (The scheme is reprinted from the publication Ruigrok et al. 2010, with permission from Elsevier)

The function of the RNA endonuclease in the influenza virus life cycle is a special way to provide the viral mRNAs with a 50 cap structure via the so-called cap-snatching mechanisms (Fig. 6.2). The requirement for a 50 cap is a consequence of the mechanisms underlying translation initiation in eukaryotes (Sonenberg and Hinnebusch 2009). The cap structure represents the key element by which the translation system recognizes an mRNA to be translated. A so-called cap-binding complex assembles at the 50 terminal cap structure in the initial phase of the translation initiation process. Thus, an RNA without a cap structure is usually not accepted as a substrate for translation. Influenza viruses replicate in the nucleus of the infected cells, where also cellular mRNA synthesis and maturation takes place. However, because of the RNA nature of the viral genome, these viruses cannot employ the cellular machinery for the production of their mRNAs. To circumvent this problem, the viral RNP that includes the RNA polymerase binds capped cellular RNAs and cleaves these RNAs close to the 50 end. The cleavage product is a short RNA fragment with a 50 terminal cap and a free 30 hydroxyl that is used as a primer for the synthesis of viral mRNA.

6.1.1.2

Arena- and Bunyaviruses

In addition to the orthomyxoviruses, members of virus families Arenaviridae and Bunyaviridae also use a “cap-snatching” mechanism during synthesis of their

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ssRNA genome G1 envelope membrane

ribosome

L Z

S

L-protein endonuclease RNA polymerase

G2

NP 3′– 5′ exonuclease cap binding protein

Fig. 6.3 Schematic representation of an arenavirus particle. The different viral proteins and the two segments of single-stranded (ss) genomic RNA are marked. Highlighted are the locations of the two proteins with RNase, namely, the L-protein with the endonuclease engaged in cap snatching and the NP harboring the 30 –50 exonuclease, involved in blocking the innate immune response to virus infection of cells. (Presentation of the scheme with kind permission from Springer Science and Business Media: Molekulare Virologie, chapter 16, 2010, page 327, Modrow et al. (2010), Fig. 16.1)

mRNAs. Arena- and bunyaviruses are also enveloped viruses with segmented RNA genomes consisting of two or three segments, respectively (Fig. 6.3). Both families are classified as negative-strand RNA viruses even though the arenavirus genome segments both have an ambisense orientation (Buchmeier 2007; Schmaljohn and Nichol 2007). In contrast to the orthomyxoviruses, the RNA-dependent RNA polymerases of arena- and bunyaviruses, known as the L-protein, consist of only one polypeptide that encompasses all the necessary activities including the endonuclease providing the capped primers for mRNA transcription (Buchmeier 2007; Schmaljohn and Nichol 2007). In bunyaviruses, a quite heterogeneous group of mostly animal viruses comprising more than 300 species, the L-protein ranges in size from 240 to 460 kDa. It contains six motifs (preA, A-E) typical for RdRp in negative-strand RNA viruses, but specific functions could not be assigned to individual domains of the protein because of a lack of conserved sequence motifs (Schmaljohn and Nichol 2007). The endonucleolytic RNase activity crucial for cap snatching in bunyaviruses resides in the N-terminal ca. 180 amino acids of the L-protein (Reguera et al. 2010). The bunyavirus endonuclease has a very similar a-b topology as PA-Nter of influenza A virus even though the lengths of the helices are significantly different. This similarity is especially striking in the core structure with the active site even though

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there is basically no homology between the two amino acid sequences. A core region of 55 residues contains the motif characteristic for the PD-(D/E)xK family of endonucleases. Within this region, a one-to-one mapping between the ligands coordinating the two metal ions was found (Reguera et al. 2010). For site 1, His-34, Asp-79, Asp-92, and the carbonyl oxygen of Tyr-93 in the bunyavirus LaCross virus correspond to His-41, Asp-108, Glu-119, and Ile-120 in PA-Nter. For site 2, Asp-52 and Asp-79 in the bunyavirus sequence correspond to Glu-80 and Asp-108 in influenza virus PA N-ter. This high degree of similarity should reflect functional equivalence, and indeed the bunyavirus endonuclease can be blocked by the same inhibitor, 2, 4-dioxo-4-phenylbutanoic acid (DBPA), as the influenza A virus enzyme. Important differences between the two structures concern the substrate-binding pocket. In PA-Nter, helix 2 together with the following loop are found in a position enabling them to restrict substrate access to the active site, whereas in the bunyavirus enzyme a wider channel is found that should allow binding and cleavage of larger, more structured substrates. Indeed, the bunyavirus enzyme is more active with a largely double-stranded RNA substrate (Reguera et al. 2010). Mutagenesis analyses showed that the presence of divalent metal ions, which preferably represent Mn2+, is crucial both for thermal stability of the protein and catalysis. There is good evidence that thermal stability is already obtained after binding of the ion in site 1, whereas enzymatic activity depends on the presence of ions at both sites. Similarly, as described above for the influenza A virus enzyme, mutations could discriminate between the endonucleolytic/transcriptional activity and the genome-replicating polymerase (Reguera et al. 2010). The arenavirus L-protein is composed of approximately 2,200 amino acids and encompasses several conserved domains (Lopez et al. 2001) including one that contains the typical RdRp signature sequence motifs (Lukashevich et al. 1997; Vieth et al. 2004). Recently, the first part of an arenavirus L protein, namely, the N-terminal region of the lymphocytic choriomeningitis virus (LCMV), has been crystallized (Morin et al. 2010). The structure analysis of this fragment, denoted NL1, showed again a central fold composed of four mixed b-strands surrounded by seven a-helices. Also in this protein, the b-strands form a twisted plane creating a negatively charged cavity for binding of divalent cations. Even though the fifth beta strand is missing in NL1, a structure-based superimposition of the endonucleaseactive sites from LCMV and influenza A virus revealed a striking similarity of these regions which included the positioning of the residues important for metal ion binding in PA-Nter. As with the bunyavirus enzyme, also the LCMV endonuclease shows some differences with regard to the influenza virus PA-Nter. The analyses did not reveal structural matches for the influenza virus His-41 or Lys-134 in the LCMV protein which is especially important since His-41 was proposed to play a direct role in the catalytic process. Biochemical analyses revealed that the arenavirus endonuclease is most likely also an Mn2+-dependent enzyme, since it was at least ~90-fold more active in the presence of Mn2+ than with other divalent cations. Mutagenesis experiments proved

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that the endonucleolytic activity of the L protein was essential for arenavirus mRNA transcription but not for genome replication (Morin et al. 2010). RNA binding was shown by gel shift assays and cleavage reactions with synthetic RNA oligonucletides revealed a specificity for substrates containing uracil. RNA with adenosine in single-stranded regions is also cleaved, but with lower efficiency. The cleavage product was found to be unusually short, comprising only 1–4 nucleotides (Buchmeier 2007). The function of the arena- and bunyavirus endonucleases in the cap-snatching process is equivalent to that of the influenza virus enzyme. So far, there is no idea how the bunyavirus RNase identifies the main feature of its substrate, namely, the 50 -terminal cap structure. The functional experiments with the arenavirus enzyme showed that the presence of a 50 cap is not required for enzymatic activity in vitro. In fact, cleavage in close vicinity of a cap structure seemed to be not even preferred in these assays (Morin et al. 2010). It nevertheless has to be postulated that such substrate discriminating binding sites exist. Moreover, the activity of the endonucleases has to be regulated, for example, by allosteric effects in a way that it is activated only after binding of a cellular substrate RNA to prevent cleavage or decapping of nascent viral mRNAs during transcription. Recently, the cap-binding site in arenaviruses was demonstrated to reside in the N-terminal part of the arenavirus nucleoprotein (NP) (Qi et al. 2010). The respective protein domain adopts a completely new fold not related to other cap-binding proteins. Qi et al. (2010) also reported that the C-terminal part of the lassavirus NP contains a second RNase. This enzyme represents a 30 –50 exonuclease and was identified because it adopts a structure strikingly similar to known 30 –50 exonucleases/exoribonucleases in humans and bacteria. The enzymatic activity of this protein domain was demonstrated by biochemical analyses that also revealed its dependency on divalent cations. Again Mn2+ was found to represent the preferred ion. One cation was identified in the structure, but the authors proposed a two cation–dependent catalytic mechanism in analogy to the homologous enzymes (Qi et al. 2010). The lassavirus NP exonuclease efficiently degrades DNA and RNA substrates including different double-stranded RNAs (Qi et al. 2010). The latter is presumably important for the function of the exonuclease during the viral life cycle. It was already known before that NP is engaged in blocking the induction of type 1 interferon (IFN-1) expression in the arenavirus infected cell thereby impairing the innate immune response of the host. Qi and coworkers (2010) showed that the repression of the interferon response is dependent on the exonuclease activity in NP, whereas cap snatching and transcription is also detected when the exonuclease is inactivated by mutation. NP interacts with RIG-I and MDA5, two RNA helicases that serve as detectors of so-called pathogen-associated molecular patterns (PAMP) in intracellular interferon induction pathways (Zhou et al. 2010). Thus, it can be hypothesized that at least one function of the NP exonuclease is to associate with these helicases and degrade PAMP-containing RNAs that have been bound by them.

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RNases of Viruses Belonging to the Order Nidovirales

Members of the order Nidovirales form a phylogenetically compact cluster of plusstrand (+) RNA viruses that diverged profoundly from other RNA viruses. In line with this, the ribonucleases encoded by nidoviruses (Bhardwaj et al. 2004; Ivanov et al. 2004; Minskaia et al. 2006) are not closely related to any of the other viral ribonucleases discussed in this chapter. The order Nidovirales currently includes three families, Coronaviridae (subfamilies Coronavirinae and Torovirinae), Roniviridae, and Arteriviridae. Nidovirus RNA replication, modification, and processing involves a complex set of enzymes, including polymerase, primase, helicase, ADP-ribose 100 -phosphatase, methyltransferase, and ribonuclease activities (Snijder et al. 2003; Ziebuhr and Snijder 2007; Ziebuhr 2008). These enzymes are expressed as part of large polyproteins, pp1a (450 kDa) and pp1ab (750 kDa), that are either encoded by ORF1a alone (pp1a) or ORFs 1a and 1b together (pp1ab). Expression of the latter involves a programmed (–1) ribosomal frameshift occurring just upstream of the ORF1a stop codon (Brierley et al. 1987). The polyproteins are co- and posttranslationally cleaved by viral proteases to produce mature processing products called nonstructural proteins (nsp) 1–16 (in coronaviruses) and nsp1–nsp12 (in arteriviruses) (Ziebuhr et al. 2000; Ziebuhr 2008).

6.1.2.1

The Nidovirus Exoribonuclease, a Putative Proofreading Enzyme

Members of the Coronaviridae and Roniviridae encode in ORF1b an exoribonuclease domain called ExoN which, in coronaviruses, occupies the N-proximal two-thirds of nsp14 (Minskaia et al. 2006; Snijder et al. 2003), while the C-terminal domain of this protein harbors a cap- (Gppp-RNA-)specific N7-methyl transferase (N7-MT) activity (Bouvet et al. 2010; Chen et al. 2009). ExoN is related to the DEDD superfamily of metal-dependent exonucleases (Snijder et al. 2003; Zuo and Deutscher 2001) whose members contain four invariant acidic residues (three Asp and one Glu) that are part of three conserved motifs. The DEDD family also includes enzymes involved in DNA proofreading, such as dnaQ, the e subunit of E. coli DNA polymerase III (Echols et al. 1983; Scheuermann et al. 1983). In contrast to their cellular homologs, nidovirus ExoN domains contain a putative zinc-binding domain that is located between motifs I and II (Snijder et al. 2003). In roniviruses, a second putative zinc-binding domain was identified between motifs II and III (Sittidilokratna et al. 2008; Snijder et al. 2003). Bacterially expressed SARS-coronavirus (SARS-CoV) nsp14 was shown to possess 30 –50 exonuclease activity that, in contrast to many cellular DEDD exonucleases, acts exclusively on RNA (Minskaia et al. 2006). Ribonucleolytic activity was found to require Mg2+ or Mn2+ or low concentrations of Zn2+ as a cofactor, while Ca2+ or high concentrations of Zn2+ inhibited activity. Substitutions of any of the four strictly conserved active-site residues (D-E-D-D) by Ala were shown to abolish

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or significantly reduce nucleolytic activity, supporting previous predictions on the critical role of these residues in catalysis (Minskaia et al. 2006). The observation that ExoN is only conserved in “large nidoviruses” featuring genome sizes of about 30 kb (Coronaviridae, Roniviridae) but not in the much smaller Arteriviridae (genome sizes between 13 and 16 kb) and the phylogenetic relatedness of ExoN with cellular exonucleases acting as proofreading enzymes in DNA replication led to the speculation that ExoN may be a critical factor in the evolution of large RNA virus genomes (Gorbalenya et al. 2006), most likely by increasing RNA replication fidelity, thus keeping the mutation frequency below a postulated critical threshold (Biebricher and Eigen 2005; Crotty et al. 2001; Gorbalenya et al. 2006). Initial studies using human coronavirus 229E (HCoV-229E) ExoN active-site mutants generated by reverse genetics revealed a critical importance of ExoN in viral RNA synthesis (Minskaia et al. 2006). Accumulation of viral RNA was severely reduced, and viable virus progeny could not be obtained in these experiments. By contrast, viable ExoN active-site mutants could be obtained for SARSCoV and murine hepatitis virus A59 (MHV-A59), although RNA synthesis and virus reproduction was again found to be significantly reduced (Eckerle et al. 2007, 2010). Consistent with the proposed role of ExoN in proofreading, mutation frequencies (determined for viable MHV-A59 and SARS-CoV ExoN mutants) were reported to be significantly (more than 15-fold) increased in ExoN mutants when compared to wild-type virus(es) (Eckerle et al. 2007, 2010). SARS-CoV ExoN mutants also displayed a higher diversity within the virus population. Interestingly, the extent of genome diversity remained essentially unchanged when ExoN mutants were passaged in cell culture, suggesting counterselection of deleterious mutations. A higher-than-average replication fidelity was calculated for (wildtype) SARS-CoV and MHV (9.0 10–7 and 2.5 10–6 substitutions, respectively, per nucleotide per replication cycle) when compared to other RNA viruses, while replication fidelities of SARS-CoV and MHV ExoN mutants were similar to those reported for other RNA viruses (Eckerle et al. 2010). As calculations were done on infectious viruses in this study, mutation frequencies likely represent an underestimate of the actual number of mutations occurring during viral replication. Overall, however, the data support the idea that ExoN has a role in keeping replication fidelity at a (high) level typical for small DNA (rather than RNA) viruses (Cuevas et al. 2009). Although a direct proof for ExoN acting as a proofreading exoribonuclease remains to be obtained, the available information would support this hypothesis (Eckerle et al. 2007, 2010).

6.1.2.2

The Nidovirus Endoribonuclease, NendoU

Nidoviral endonucleases derive their name, NendoU, from Nidovirus endoribonuclease specific for U(ridylate). NendoU domains are related to a family of cellular enzymes prototyped by the Xenopus laevis endoribonuclease XendoU (Snijder et al. 2003), an endoribonuclease involved in the processing of small

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nucleolar (sno)RNAs (Laneve et al. 2003). NendoU is conserved across members of the Nidovirales but not in other RNA viruses, making the domain a genetic marker of nidoviruses (den Boon et al. 1991; Ivanov et al. 2004). NendoU resides in nsp15 in coronaviruses and nsp11 in arteriviruses. NendoU activities have been characterized for both coronavirus and arterivirus homologs (Bhardwaj et al. 2004; Cao et al. 2008; Ivanov et al. 2004; Kang et al. 2007; Nedialkova et al. 2009). The enzymes were reported to (1) cleave downstream of uridylate (and, in some cases and less efficiently, after cytidylate), (2) release products with 20 ,30 -cyclic phosphodiester ends, and (3) cleave ssRNA more efficiently than dsRNA (Bhardwaj et al. 2004, 2006; Ivanov et al. 2004; Nedialkova et al. 2009). Structural and biochemical studies revealed that coronavirus NendoUs (nsp15) and their cellular homolog XendoU possess a novel fold that is not found in other ribonucleases (Renzi et al. 2006; Ricagno et al. 2006). Coronavirus NendoUs form hexamers comprised of dimers of trimers (Bhardwaj et al. 2006, 2008; Ricagno et al. 2006; Xu et al. 2006). The monomers have an a + b structure comprised of three domains, with the nidovirus-wide conserved domain (den Boon et al. 1991) representing the C-terminal subdomain of the protein (Fig. 6.4). The NendoU hexamer forms a three petal-shaped surface that surrounds a small, predominantly ˚ . The two negatively charged central channel with an inner diameter of ~15 A trimers interact head to head, mainly involving the N-terminal domains, while the C-termini are located at the surface where they form six independent active sites. Inter-monomer interactions are largely mediated by residues of the N-terminal and central domains (Ricagno et al. 2006; Xu et al. 2006). Despite limited sequence and structural similarity, the active-site residues (His-234, His-242, and Lys-289 of SARS-CoV nsp15) can be superimposed with equivalent residues of the catalytic center of bovine RNase A (His-12, His-119 and Lys-41) (Ricagno et al. 2006), a well-characterized enzyme belonging to a different family of pyrimidine-specific ribonucleases. The proposed coronavirus NendoU catalytic His and Lys residues cluster inside a positively charged groove of the C-terminal domain (Fig. 6.4) (Ricagno et al. 2006; Xu et al. 2006) and mutagenesis data confirmed their critical role in nuclease activity (Bhardwaj et al. 2008; Ivanov et al. 2004; Kang et al. 2007). Additional active-site residues were implicated in binding of the substrate phosphate. These involve the side chain of a highly conserved Thr (Thr-340 in SARS-CoV nsp15, Fig. 6.4) and the main chain amide of a conserved Gly residue (Gly-247 in SARS-CoV nsp15, Fig. 6.4) (Bhardwaj et al. 2008; Ivanov et al. 2004; Kang et al. 2007; Xu et al. 2006). Although the NendoU reaction mechanism has not been analyzed in detail, it is generally thought to be similar to that of RNase A. This is supported by (1) similar spatial positions of the putative catalytic His and Lys residue(s) in the structure (see above), (2) the production of 20 ,30 cyclic phosphate-containing reaction products (Bhardwaj et al. 2008; Ivanov et al. 2004), and (3) the production of 30 -hydroxyl ends after extended reaction times using recombinant forms of NendoU. Similarities between the active sites of RNase A and nidovirus NendoUs extend to residues presumably involved in substrate binding and, more specifically, pyrimidine specificity. Thus, in the RNase A structure, Thr-45 and Phe-120, which are

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SLENVAYNVVNKGHFDGHAGEAPVSIINNAVYTKVDGIDVEIFENKTTLPVNVAFELWAKRNIKPV SLENVVYNLVNAGHFDGRAGELPCAVIGEKVIAKIQNEDVVVFKNNTPFPTNVAVELFAKRSIRPH SLENVAFNIVKKGAFTGLKGDLPTAVIADKIMVRDGPTDKCIFTNKTSLPTNVAFELYAKRKLGLT GLENIAFNVVNKGSFVGADGELPVAISGDKVFVRDGNTDNLVFVNKTSLPTNIAFELFAKRKVGLT SIDNIAYNMYKGGHYDAIAGEMPTVITGDKVFVIDQGVEKAVFVNQTTLPTSVAFELYAKRNIRTL α3

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PEIKILNNLGVDIAANTVIWDYKREAPAHVSTIGVCTMTDIAKKPTESACSSLTVLFDGRVEGQVD PELKLFRNLNIDVCWSHVLWDYAKDSVFCSSTYKVCKYTDLQ......CIESLNVLFDGRDNGALE PPLTILRNLGVVATYKFVLWDYEAERPFSNFTKQVCSYTDLD........SEVVTCFDNSIAGSFE PPLSILKNLGVVATYKFVLWDYEAERPLTSFTKSVCGYTDFA........EDVCTCYDNSIQGSYE PNNRILKGLGVDVTNGFVIWDYANQTPLYRNTVKVCAYTDIE.......PNGLVVLYDDR.YGDYQ α4

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LFRNARNGVLITEGSVKG....LTPSKGPAQASVNG..VTLIGES.....VKTQFNYFKKVDGIIQ AFKKCRNGVYINTTKIKS....LSMIKGPQRADLNGVVVEKVGDSD....VEFWFAVRKDGDDVIF RFTTTRDAVLISNNAVKG....LSAIKLQ.YGLLNDLPVSTVGN.....KPVTWYIYVRKNGEYVE RFTLSTNAVLFSATAVKTGGKSLPAIKLN.FGMLNGNAIATVKSEDGNIKNINWFVYVRKDGKPVD SFLAADNAVLVSTQCYKR....YSYVEIPSNLLVQNGMPLKDGAN........LYVYKRVNGAFVT η2

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QLP...............................ETYFTQSRDLEDFKPRSQMETDFLELAMDEFI SRTGSLEPSHYRSPQGNPGGNRVGDLSGNEALARGTIFTQSRLLSSFTPRSEMEKDFMDLDDDVFI QID................................SYYTQGRTFETFKPRSTMEEDFLSMDTTLFI HYD................................GFYTQGRNLQDFLPRSTMEEDFLNMDIGVFI LPN................................TINTQGRSYETFEPRSDIERDFLAMSEESFV α7

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QRYKLEGYAFEHIVYGDFSHGQLGGLHLMIGLAKRSQDSPLKLEDFIP.MDSTVKNYFITDAQTGS AKYSLQDYAFEHVVYGSFNQKIIGGLHLLIGLARRQQKSNLVIQEFVT.YDSSIHSYFITDENSGS QKYGLEDYGFEHVVFGDVSKTTIGGMHLLISQVRLAKMGLFSVQEFMNNSDSTLKSCCITYADDPS QKYGLEDFNFEHVVYGDVSKTTLGGLHLLISQVRLSKMGILKAEEFVAASDITLKCCTVTYLNDPS ERYG.KDLGLQHILYGEVDKPQLGGLHTVIGMYRLLRANKLNAKSVTN.SDSDVMQNYFVLSDNGS β18

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SKCVCSVIDLLLDDFVEIIK...SQDLSVISKVVKVTIDYAEISFMLWCKDGHVETFYPKLQ SKSVCTVIDLLLDDFVDIVK...SLNLKCVSKVVNVNVDFKDFQFMLWCNEEKVMTFYPRLQ SKNVCTYMDILLDDFVTIIK...SLDLNVVSKVVDVIVDCKAWRWMLWCENSHIKTFYPQLQ SKTVCTYMDLLLDDFVSVLK...SLDLTVVSKVHEVIIDNKPWRWMLWCKDNAVATFYPQLQ YKQVCTVVDLLLDDFLELLRNILKEYGTNKSKVVTVSIDYHSINFMTWFEDGSIKTCYPQLQ # #

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His-249 C N

Fig. 6.4 Coronavirus endoribonucleases. (a) Multiple sequence alignment of coronavirus nsp15 (NendoU) domains, representing the three coronavirus genera, Alphacoronavirus (HCoV-229E, TGEV), Betacoronavirus (SARS-CoV, MHV-A59), and Gammacoronavirus (IBV). Sequences

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known to contribute to pyrimidine specificity by forming hydrogen bonds (Thr-45) or stacking interactions (Phe-120) with the pyrimidine base (Raines 1998) occupy positions that are similar to those of two Ser/Thr and Tyr residues conserved in coronavirus and arterivirus NendoUs (Ser-293 and Tyr-342 in SARS-CoV nsp15; Fig. 6.4). Molecular modeling of uridine 30 -monophosphate binding to the SARSCoV NendoU active site and mutagenesis data provided further evidence for a role of Ser-293 and Tyr-342 in substrate binding and specificity (Bhardwaj et al. 2008; Nedialkova et al. 2009; Ricagno et al. 2006). Thus, substitutions with Ala of Ser-293 in SARS-CoV nsp15 and Ser-228 in EAV nsp11 resulted in enzymes that had essentially lost their preference for uridine over cytidine. By contrast, specificity and activity were restored when the same Ser residues were substituted with Thr (Bhardwaj et al. 2008; Nedialkova et al. 2009). In addition, Pro-343 and Leu-345 (SARS-CoV nsp15 numbering) were implicated in uridylate specificity, again supported by mutagenesis data (Bhardwaj et al. 2008). Thus far, the role of metal ions in NendoU activity has not been resolved conclusively. Whereas the nucleolytic activities of the cellular NendoU homologs, XendoU and human placental protein (PP11), and coronavirus NendoUs depend on (or are stimulated by) Mn2+ ions (Bhardwaj et al. 2004, 2008; Ivanov et al. 2004; Laneve et al. 2003, 2008), arterivirus NendoU activities do not appear to require metal ions (Nedialkova et al. 2009). At low concentrations of Mn2+, the activities of bacterially expressed equine arteritis virus (EAV) and porcine reproductive and respiratory syndrome virus (PRRSV) NendoUs were only slightly enhanced and, at higher concentrations, activities were found to be inhibited. The observed critical (or, at least, supportive) role of metal ions in most NendoU/XendoU homologs characterized to date is inconsistent with the proposed RNase A-like (metal-independent) reaction mechanism and structural studies did not reveal metal-binding sites in these enzymes (Renzi et al. 2006; Ricagno et al. 2006), thus questioning the requirement for Mn2+ ions in nuclease activity. Mn2+ ions were found to affect the intrinsic tryptophan fluorescence of SARS-CoV nsp15, suggesting that metal ion binding can induce conformational changes in the protein which were suggested to affect RNA binding (Bhardwaj et al. 2006; Guarino et al. 2005). Surprisingly, Mn2+ ions stimulated the RNA-binding activity of SARS-CoV nsp15 but not that of XendoU (Bhardwaj et al. 2006; Gioia et al. 2005).

Fig. 6.4 (continued) were aligned using ClustalW 2.0 (Larkin et al. 2007) and rendered with ESPript 2.2 (http://espript.ibcp.fr/ESPript/cgi-bin/ESPript.cgi). Abbreviations of virus names and accession numbers are as follows: SARS-CoV Severe acute respiratory syndrome coronavirus [AY291315], MHV-A59 Murine hepatitis virus A59 [NC_001846], TGEV Transmissible gastroenteritis virus [Z34093], HCoV-229E Human coronavirus 229E [NC_002645], IBV Avian infectious bronchitits virus [NC_001451]. Secondary structure elements of SARS-CoV nsp15 (PDB 2H85) are shown above the sequence. Catalytic residues are indicated by asterisks and key residues involved in uridylate specificity are indicated by hashes (see text for details). (b) Ribbon representation of a SARS-CoV nsp15 (NendoU) monomer (pdb 2H85). The catalytic residues, His234, His249, and Lys289, are shown in ball-and-stick representation, and amino and carboxyl termini are indicated (N, C)

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Biochemical studies suggest that the hexameric structure described above is the fully active form of coronavirus NendoUs (Bhardwaj et al. 2006; Guarino et al. 2005; Xu et al. 2006). This is supported by the conservation and functional relevance of residues predicted by structural and biochemical studies to stabilize intersubunit interactions in the trimer and hexamer. Also, a structure analysis of a truncated form of SARS-CoV nsp15, which lacked 28 N-terminal and 11 C-terminal residues and was monomeric, revealed that two loops of the catalytic domain were displaced compared to their location in the hexamer, causing major changes in the active site geometry and loss of activity (Joseph et al. 2007). On the basis of these observations, hexamerization has been suggested to act as an allosteric switch required to activate these enzymes. If confirmed, this regulatory mechanism would not apply to EAV nsp11 and XendoU, both of which were reported to be monomeric in solution (Nedialkova et al. 2009; Renzi et al. 2006). The biological significance of NendoU activities in the nidovirus life cycle has not been elucidated. Substitution of coronavirus NendoU active-site residues by reverse genetics resulted in a slight reduction of both genomic and subgenomic (sg) RNA synthesis in MHV (Kang et al. 2007) and a slight reduction in virus reproduction. No significant differences in plaque size were observed for NendoU mutants compared to wild-type MHV. In striking contrast, NendoU active-site substitutions caused profound defects in arterivirus reproduction, with virus titers being reduced by up to five log in a few cases. Several substitutions in the EAV NendoU resulted in a selective reduction of sgRNA synthesis compared to genome replication. Substitutions that, based on the available structure information for coronavirus NendoU, are predicted to cause major structural changes in NendoU (and, possibly, the polyprotein) abolished RNA synthesis in EAV and HCoV-229E completely (Ivanov et al. 2004; Posthuma et al. 2006). Taken together, the data suggest that NendoU domains may have nonidentical roles and/or substrates in different nidovirus families/subfamilies/genera, possibly reflecting adaptation to specific hosts and/or ecological niches (Lei et al. 2009).

6.2

Viral RNase Influencing Host Cellular Metabolism and the Host Immune Response: The Herpesvirus vhs Protein

Herpesviruses constitute a family of enveloped viruses with large double-stranded DNA genomes of more than 100 kb that encode a large number of viral proteins (Pellet and Roizman 2007). The genomic DNA is linear and contains both unique regions and long repeats. In viral particles, it is packaged into an icosahedral capsid that again is surrounded by a host cell–derived lipid bilayer into which the viral envelope proteins are inserted. The space between envelope and capsid is filled by the so-called tegument, a mixture containing a variety of viral proteins (Fig. 6.5). During membrane fusion in the course of infection, the tegument is delivered to the cytoplasm of the newly infected cell together with the viral capsid. In addition to,

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Fig. 6.5 Schematic representation of a herpes simplex virus particle. Basic elements including the envelope membrane, capsid, core, genome and tegument, and the different viral envelope glycoproteins are marked. As indicated, the tegument contains the vhs RNase and VP16 and VP22, two proteins involved in regulation of vhs activity. (The figure was kindly provided by H. Granzow and M. J€ orn, electron microscopy Friedrich-Loeffler-Institut, Insel Riems)

for example, proteins necessary for activation of viral gene expression, tegument of viruses belonging to the genus Alphaherpesvirus contains the “virus host shutoff” (vhs) protein, a factor involved in shutting down host cellular gene expression (Roizman et al. 2007). In herpes simplex virus 1 (HSV-1), vhs is a polypeptide of ca. 58 kDa that is encoded by the open reading frame UL41 (Roizman et al. 2007). It triggers a rapid shutoff of host cellular protein synthesis. This is achieved by disruption of preexisting polyribosomes and degradation of mRNAs. Since vhs is introduced into the infected cell as part of the tegument, viral gene expression is not necessary for shutoff (Smiley 2004). The observed effects of vhs on cellular RNA metabolism could easily be explained if vhs activated RNA degradation or represented an RNase itself. In fact, extracts of HSV-1-infected cells and, even more importantly, extracts from partially purified virions were shown to contain an RNase activity (Smiley 2004). RNA hydrolysis could be prevented by vhs-specific antibodies and different mutations affecting vhs. Further evidence for vhs representing an RNase came from studies in which vhs was expressed in the absence of other viral proteins and tested in vitro or in yeast cells (Elgadi et al. 1999; Zelus et al. 1996). In addition, a complex of vhs with eukaryotic translation initiation factor eIF4H (see also below), partially purified from recombinant E. coli, showed RNase activity (Everly et al. 2002). Finally, the RNase activity of vhs was demonstrated with recombinant protein that had been expressed in E. coli and purified to homogeneity (Taddeo and Roizman 2006; Taddeo et al. 2006, 2010). Similar to the RNases of the segmented negative-strand RNA viruses described above, the vhs RNAse seems not to belong to one of the known RNase families. Structural data are not available so far, but sequence comparison studies revealed a relationship to a family of cellular nucleases that are involved in DNA repair and

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replication (Doherty et al. 1996; Everly and Read 1997; Everly et al. 2002). One of these cellular homologs is FEN-1, a nuclease engaged in the removal of RNA primers used for synthesis of Okazaki fragments during eukaryotic DNA replication. This similarity with cellular nucleases is obviously also important on a functional level, since mutations affecting conserved residues known to be important for enzymatic activity of the cellular proteins also abrogate vhs-induced repression of gene expression (Everly et al. 2002). Isolated vhs has been shown to hydrolyze RNA with broad substrate specificity. It requires single-stranded RNA and cleaves 30 of C and U residues, a substrate specificity equivalent to that of RNase A (Taddeo and Roizman 2006). However, when expressed within a living cell, this unusual enzyme selectively targets cellular and viral mRNAs while sparing other ribonucleic acids. This substrate specificity under physiological conditions seems to be not mediated by direct recognition of one of the mRNA specific modifications 50 cap or 30 poly(A) tail. Instead, vhs interacts with cellular proteins that represent components of the translation initiation complex, such as initiation factor eIF4H was demonstrated (Feng et al. 2001). Since eIF4H associates with eIF4A, a component of the cap-binding complex, vhs could be attached to the 50 ends of mRNAs via an initiation factor bridge. In agreement with this hypothesis, it was reported for different mRNAs that vhs degradation starts close to the 50 end (Elgadi et al. 1999; Karr and Read 1999). Along those lines, it also has to be mentioned that vhs cleaves the cap-less picornavirus RNAs immediately downstream of the so-called internal ribosomal binding site (IRES), whereas other cap-less mRNAs like cellular IRES-containing mRNAs are not degraded (Elgadi and Smiley 1999). Picornavirus IRES recruit most of the canonical initiation factors to the viral RNAs, whereas the cellular IRES elements function without many of these factors. Thus, even though mere tethering of vhs to the cap-binding complex is not sufficient for mRNA degradation (Page and Read 2010), it can be hypothesized that the vhs specificity for mRNAs is mediated by components of the host cellular translation initiation system. Recent work has shown that the model of an initiation factor-mediated specific degradation of mRNAs is presumably oversimplified. Several mRNAs were found to be cleaved at a site close to the 30 end despite the presence of a 50 cap structure. According to these data, at least three classes of mRNAs can be distinguished based on their fate after infection (Corcoran et al. 2006; Smiley 2004; Taddeo et al. 2010). The mRNAs of housekeeping genes such as GAPDH or b-actin are rapidly degraded. At least some inducible genes generating mRNAs with AU-rich elements (AREs) in their 30 nontranslated regions are deadenylated and cleaved endonucleolytically close to their 30 end giving rise to 50 fragments that persist for hours before 30 –50 degradation occurs. Several stress-inducible genes coding for proteins with regulatory functions like IEX-1, c-fos, or the a subunit of the inhibitor of nuclear factor kB (IkBa) belong to this group. A third class of mRNAs, which are derived from inducible genes encoding, for example, tristetraprolin (TTP) is not degraded at all, so that the encoded proteins accumulate in the infected cells. Since TTP binds both to vhs and AREs of mRNAs (Esclatine et al. 2004), the vhs-induced cleavage

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of the latter was hypothesized to be mediated by TTP (Chen et al. 2001; LykkeAndersen and Wagner 2005). The RNase activity of vhs seems to be essentially modulated by at least three viral proteins. Two of these proteins, VP16 and VP22, are also present in the viral tegument (Fig. 6.5), and therefore already delivered to the newly infected cells. VP16 alone is able to associate with vhs, whereas VP22 cannot bind vhs but interacts with it only via VP16. The interaction of vhs with these two proteins is believed to interfere with the RNase activity and to be necessary for an appropriate level of viral gene expression in the early phase of the infection. However, it is not clear whether this effect is achieved alone by limiting mRNA degradation. Instead, there is evidence that these two proteins also function through enabling mRNA translation (Taddeo et al. 2007). In addition, a third protein termed ICP27 was shown to be also involved in modulation of vhs activity (Corcoran et al. 2006; Taddeo et al. 2010). ICP27 is an immediate early protein of HSV with different functions in regulation of viral and cellular gene expression. It stimulates transcription and translation of viral mRNA and contributes to HSV host shutoff by downregulation of host transcription and blocking of nuclear export of host cellular mRNAs. Moreover, ICP27 seems to be important for the specificity of vhs activity, but the exact role of ICP27 is still a matter of debate. It seems to interact with vhs bound to cap-and poly(A)-binding proteins. It is also essential for the synthesis of new vhs in the late phase of the viral replication cycle. The absence of ICP27 leads to degradation of mRNAs with AREs late after infection, which was proposed to result from a stabilizing effect of ICP27 on these RNAs (Corcoran et al. 2006) or the absence of the protective effect of vhs itself because of the absence of the newly synthesized vhs (Taddeo et al. 2010). The vhs function is dispensable for alphaherpesvirus replication, and vhs homologs are absent from beta- and gamma-herpesviruses (Smiley et al. 2001; Smiley 2004). Nevertheless, vhs is found in all alphaherpesviruses so that it likely plays an important role in the biology of these viruses. In fact, vhs-negative HSV is strongly attenuated. In accordance with its name, a primary function of vhs is certainly the shutoff of host cellular gene expression. At least the concentration of most stably expressed cellular mRNAs is lowered dramatically in the early phase of the replication cycle. Moreover, the function of the translational apparatus is altered so that the translation of most of the residual mRNAs is severely impaired. As a consequence, higher capacities of the translation system are available for the synthesis of viral proteins. However, it is obvious that vhs also degrades viral mRNAs so that maximization of viral gene expression seems to be not the final aim. In fact, vhs also ensures the rapid turnover of viral mRNAs, a process that is needed to regulate the progression of the replication cycle from the immediate early to the early and then to the late phase. In addition to gross effects on viral and cellular mRNA metabolism, vhs plays also a major role in alphaherpesvirus evasion of host immunity. It is involved in the loss of major histocompatibility complex (MHC) class I and class II from the surface of infected cells (Ambagala et al. 2000; Gopinath et al. 2002; Hinkley et al. 2000; Koppers-Lalic et al. 2001; Tigges et al. 1996). Moreover, the production

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of proinflammatory chemokines and cytokines is hampered by HSV infection (Suzutani et al. 2000). Recently, vhs has been shown to functionally inactivate human monocyte–derived dendritic cells (Samady et al. 2003). Collectively, vhs activity dampens the innate and both arms of the adaptive immune system. The effects of vhs on the host immune system are regarded as the major cause for the attenuation of vhs deletion mutants. Vhs deletion mutants have been shown to induce very robust immune responses in experimentally infected animals and, consequently, deletion of vhs is regarded as one possible step during the establishment of novel vaccines against alphaherpesviruses. It also has to be mentioned that its ability to help block the host immune response seems to be the major reason for conservation of vhs among alphaherpesviruses since the ability of vhs to induce host shutoff varies considerably from HSV-2 (very strong shutoff) to HSV-1 (strong shutoff), pseudorabies virus (moderate shutoff) and equine herpesvirus (no detectable host shutoff) (Smiley 2004). Taken together, the alphaherpesvirus vhs protein represents a new type of RNase with multiple functions in the viral life cycle and virus–host interaction. These different functions and the adaptation of vhs activity to changing demands in the course of the viral life cycle are achieved by a complex regulatory network established by different viral proteins controlling the activity of tegumentassociated vhs early in infection and expression of new vhs during the late phase of the cycle.

6.3

Viral RNase Important for the Interaction Between Virus and Host Organism: The Pestivirus Erns RNase

Pestiviruses represent a group of pathogens that are responsible for economically important diseases of farm animals (Lindenbach et al. 2007). Pestiviruses are enveloped positive-strand RNA viruses that are classified as one genus in the family Flaviviridae because of their general genome organization and strategy of gene expression. The pestiviral genome encodes a single polyprotein of roughly 4,000 amino acids that is cleaved into 12 mature viral proteins by cellular and viral proteases. Four of these proteins are found in the virus particle, a capsid protein C and three glycosylated proteins embedded into the viral envelope membrane (Fig. 6.6). The presence of three envelope proteins represents a special feature of pestiviruses, whereas the closely related human hepatitis C viruses (HCV) have only two envelope proteins. With regard to location in the polyprotein, biochemical properties, and presumed function, the glycoproteins E1 and E2 of pestiviruses are thought to be equivalent to E1 and E2 of HCV, respectively. Thus, the so-called Erns protein (envelope, ribonuclease secreted), formerly termed E0, located in the polyprotein between C and E1 seems to represent the additional acquirement of pestiviruses.

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Fig. 6.6 Schematic representation of a pestivirus particle. Basic elements like envelope membrane, singlestranded positive-sense RNA genome, and the viral structural proteins are marked. Below the virion, a blowup of the part of the Erns sequence containing the RNase motif is shown in comparison with the corresponding sequence from T2 RNase

envelope membrane Erns E1 E2 capsid protein

viral RNA

RNase Motif Erns: T2:

SLHGIWPEkiCkG…LqrHEWNKHGwCnwynIDP TIHGIWPDn-CdG…FweHEWNKHGtC-IntIEP

Erns exhibits several unusual features. It is highly glycosylated with more than 50% carbohydrate in its mature form, and usually forms disulfide-linked homodimers (Hulst and Moormann 2001; Thiel et al. 1991). The C-terminal part of Erns folds into a long amphipathic helix that associates with lipid bilayers in an in-plane configuration and thereby mediates membrane anchoring of the protein (Tews and Meyers 2007). This unusual form of membrane anchoring is believed to be responsible for the fact that part of the Erns protein synthesized within a cell is secreted into the cell-free supernatant. The most unusual feature of Erns, however, is its enzymatic function. Erns represents the only known viral surface protein that exhibits RNase activity (Hulst et al. 1994; Schneider et al. 1993). Erns was identified as ribonuclease because of two short stretches within the N-terminal half of the protein (Box A LH30GIWP and Box B HEWNKH79GWC) that displayed homology to ribonucleases of the T2 family (Fig. 6.6). Ribonucleases of the T2 family are transferase-type RNases and are classified by their similarity to RNase T2 from Aspergillus oryzae (Luhtala and Parker 2010). T2 RNases are widespread among bacteria, protozoa, plants, insects, vertebrates, and viruses. T2 RNases are endonucleases that typically have their enzymatic optima at acidic pH (pH 4–5). This acidic activity of T2 RNases correlates with their localization in lysosomal or vacuolar compartments. Most T2 RNases have little substrate specificity and cleave phosphodiester bonds at all four bases (Luhtala and Parker 2010). Biochemical characterization of the Erns activity revealed that various heteropolymeric RNA molecules such as ribosomal RNA or pestivirus genomic RNA were susceptible to Erns activity (Hulst et al. 1994; Schneider et al. 1993; Windisch et al. 1996). Of single-stranded homopolymeric RNA molecules, only polyrU was cleaved while polyrA, polyrG, and polyrC were resistant to Erns degradation (Schneider et al. 1993; Windisch et al. 1996). Addition of the complementary homopolymer (polyrA) to the polyrU resulted in massively reduced degradation indicating the inability of Erns to accept double-stranded RNA as substrate. In conflict with this

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observation is experimental evidence that suggests that Erns also degrades dsRNA such as poly IC (Iqbal et al. 2004) or double-stranded transcripts (Magkouras et al. 2008; M€atzener et al. 2009). Erns is insensitive to chelating agents such as EDTA or EGTA, and can be inhibited with Mn2+ and Zn2+ ions. Zn2+ leads at 15 mM concentration to a 50% reduction of Erns activity likely because of the interaction with histidine residues His-30 and His-79 that were shown to be essential for RNase activity (Meyers et al. 1999). As the function of Erns is enigmatic, the exact substrate specificity was analyzed by Hausmann et al. (Hausmann et al. 2004). Using radiolabeled substrates, the preference for uridine residues was confirmed for heteropolymeric substrates. Interestingly, the cleavage occurs at an Np/U site in which N can be formed by any nucleotide. While most T2 RNases have a specificity determined by the B1 site (Irie and Ohgi 2001), the determinant of the B2 position is not a unique property of Erns among the T2 RNases. RNase MC from the pumpkin Momorica charantia also has a specificity for Np/U (Irie et al. 1993). Labeling of the transcribed substrate RNAs that contained a single uridine residue with uridine triphosphate (32P alpha UTP) and separation of the cleavage product on a denaturing polyacrylamide gel revealed that the labeled phosphate was transfered to the nucleotide in the B1 position (the 30 end of the 50 cleavage product), while the nucleotide at the B2 position was dephosphorylated. This fits to the general mechanism of T2 RNases using transesterification and hydrolysis (Irie and Ohgi 2001). A 20 –30 cyclophosphate is an intermediate structure that in most T2 RNases is hydrolyzed to a 30 phosphate. Experiments by Windisch suggest that Erns is unable to modify 20 –30 cyclophosphate mononucleotides. Initial determination of kinetic parameters was reported by Windisch et al. (Windisch et al. 1996). Using polyrU as substrate and incubation times of 15–20 min a Km of about 872.5 10–6 M was determined. In a different approach using radioactively labeled single-stranded substrates with a single cleavage site each (GpU, CpU, ApU, and UpU), turnover was assayed for up to 60 s after mixing with the enzyme. These analyses revealed affinity constants of 83.8–258.7 10–9 M in the order UpU > GpU > CpU > ApU and that were 103- to 104-fold lower than described by Windisch. This is indicative for a high affinity of Erns for the substrate. New analyses using highly purified Erns and radiolabeled RNA oligonucleotides of 23 nt length revealed even higher affinities to the substrate (Km 2.17 10–9 M) and a umenapf, unpublished). Kcat of 11.5. (R€ T2 RNases are endonucleases that cleave polymeric RNA molecules internally. In the case of Erns, NpU sites represent the preferred substrate and the size of the cleavage products depends on the distribution and accessibility of uridine residues. Analysis of the cleavage products of Erns digested RNA molecules with known cleavage sites gave a different picture. Depending on the enzyme concentrations, the expected cleavage products were completely degraded to mononucleotides or shorter molecules forming a “ladder” of 50 coterminal fragments. This led to the assumption that the endonuclease Erns possesses an additional exonucleolytic activity. This exonucleolytic activity of Erns is only apparent with substrates that contain at least one uridine residue, and an endonucleolytic cleavage is required

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before exonucleolytic degradation ensues. Interestingly, a comparable activity has not been reported for other T2 RNases, and it is currently not understood how a uridine-specific endonuclease switches to a substrate-independent exonuclease. A hypothetical 3D structure modeled on the basis of an RNase Rh template was published (Langedijk et al. 2002), but this model does not allow conclusions with regard to the molecular basis of substrate recognition of Erns. However, crystal structure analysis of Erns is in progress (T. Krey, personal communication) so that deeper elucidation of this interesting enzyme will hopefully be possible soon. The identification of a sequence in the genome of an RNA virus which codes for an RNase raised the question how the viral genome is protected from this dangerous enzymatic activity. Experimental work showed that highly purified Erns protein was able to degrade viral genomic RNA isolated from virions demonstrating the absence of a modification preventing cleavage of the viral RNA (Windisch et al. 1996). Protection of the viral genome could also be achieved by separating the genomic RNA and the RNase through membranes. Since mature Erns is highly glycosylated and contains four intramolecular disulfide bonds, folding of the protein into its enzymatically active form could well be dependent on its translocation into the endoplasmatic reticulum (ER). As a matter of fact, prevention of translocation by expression of Erns without signal sequence resulted in an inactive form of the protein exhibiting no RNase activity (Meyers, unpublished results). Since membrane topology of viral envelope proteins and genome is conserved during budding and membrane fusion in the course of infection, it can be concluded that active RNase and viral genome are always separated by membranes which prevents degradation of the viral genome by its own RNase. The results and conclusions described above are also important for the question about the function of the Erns RNase. Since the active RNase is obviously never present within the cytoplasm of the infected cell, the enzymatic activity cannot have any function in the viral replication cycle. Indeed, mutation of predicted active-site residues of the RNase led to inactivation of the enzyme but allowed recovery of viable viruses with growth characteristics similar to wild-type viruses (Hulst et al. 1998; Meyer et al. 2002; Meyers et al. 1999). Thus, the ability to express an active Erns RNase seems to offer no significant advantage for virus replication in tissue culture cells. However, the RNase motifs and therefore most likely also the enzymatic activity of the protein have been conserved during evolution of pestiviruses indicating an important function of the RNase. Animal studies provided first hints pointing at a putative function of the Erns RNase. Viruses with inactivated RNase were shown to be attenuated in their natural hosts (Meyer et al. 2002; Meyers et al. 1999). Importantly, the initial reaction of the animals infected with the RNase-negative mutants was very similar to what was seen upon infection with a wild-type virus. However, around day 7–10 post infection, the animals were apparently able to control the RNase-negative virus and recovered whereas the amount of wild-type virus increased dramatically resulting in very severe symptoms of disease. Despite the early control of the RNase-negative virus, the animals infected with the mutant virus showed a potent antiviral immune response able to protect from a stringent challenge infection

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(Meyer et al. 2002; Meyers et al. 1999). Based on these data, it can be hypothesized that the function of the pestivirus RNase is to somehow interfere with the immune system of the host, thereby delaying the immunological control of the infection. The immune response to virus infection can be divided into the two functionally and temporally differentiated systems called the innate and the adaptive immune response. The latter type of response relies on a powerful, antigen-specific system designed for final clearance of the invading pathogen. Since, however, the development of an adaptive immune response takes a rather long time, a first line of defense is necessary to prevent uncontrolled amplification of the pathogen. This first line of defense is established by the innate immune system relying on the identification of pathogen-associated molecular patterns (PAMPs) (Janeway 1989). In RNA viruses, PAMPs are molecular structures like double-stranded RNA or cytoplasmic RNA with a 50 triphosphate structure which are not found in eukaryotic cells, and therefore specific for the pathogens (Hengel et al. 2005; Takeda and Akira 2004). Binding of such a molecular structure by a PAMP-specific receptor triggers a cascade of reactions among which the synthesis and secretion of type I interferon represents one of the earliest and most important steps. Secretion of IFN-1 induces an antiviral state in the infected as well as in neighboring cells characterized by a whole set of events aiming at repression of pathogen replication. This first line of defense represents a serious barrier for viruses so that most, if not all of them, have evolved mechanisms counteracting the innate immune system especially at the level of the IFN-1 response (Hengel et al. 2005). Because of the importance of the interferon response for the innate immune system and the fact that several types of RNA molecules represent PAMPs, it was proposed that the Erns RNase could be involved in blocking the interferon system. The high affinity to single-stranded RNA and the postulated exonuclease activity of Erns support the assumption that Erns is involved in degrading viral RNA molecules that could stimulate endosomal (Toll like receptors 3, 7, 8) or cytoplasmic innate immunity sensors (MDA5 and RIG-I). The low km facilitates interaction of minute concentrations of Erns with target RNA molecules, that is, in the endosomal compartment. Since endonucleolytic cleavage results in a molar increase of breakdown products that still are sensed as pathogen-associated molecular patterns (PAMPs), it is probably beneficial for the survival of pestiviruses to fully degrade these fragments by the exonucleolytic activity of Erns. Tissue culture experiments showed that supplementation of culture supernatant with Erns blocked the interferon response triggered by addition of double-stranded RNA (Poly-IC) (Iqbal et al. 2004; Magkouras et al. 2008; M€atzener et al. 2009). This was definitely an effect initiated extracellularly since Erns could not block the induction of interferon after transfection of Poly-IC. In vitro experiments showed that Erns was indeed able to bind and degrade dsRNA, even though the requirements necessary for hydrolysis of dsRNA were different in two labs (Iqbal et al. 2004; Magkouras et al. 2008; M€atzener et al. 2009). Thus, the hypothesis was put forward that secreted Erns was responsible for blocking an interferon response to extracellular dsRNA. This conclusion is in agreement with the published results, but has to be questioned in some points mainly because the origin of dsRNA in the extracellular

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milieu in an infected animal is obscure. This dsRNA should directly result from virus infection but since the overwhelming amount of pestiviruses are not cytopathic (Meyers and Thiel 1996; Thiel et al. 1996), lysis of considerable numbers of infected cells containing viral dsRNA is unlikely. It is therefore not clear where the dsRNA substrate to be degraded by the Erns RNase should come from. It also has to be stressed that Erns is much more active on single-stranded RNA and could therefore be regarded as a predominantly ssRNA-specific RNase. If dsRNA was the main substrate of this RNase, one could expect that evolution would have taken a different way. Pestiviruses express one further protein for which repression of the interferon response of an infected cell has been published. This protein named Npro is a nonstructural protein with protease activity that induces proteasomal degradation of interferon regulatory factor 3 (IRF3) within the infected cell (Bauhofer et al. 2007; Chen et al. 2007; Hilton et al. 2006; La Rocca et al. 2005; Ruggli et al. 2005, 2009; Seago et al. 2007). In contrast to the Erns RNase, Npro is able to block the expression of type 1 IFN upon transfection of dsRNA, supporting the idea that Npro is responsible for prevention of an innate immune response triggered from inside of the infected cell. Thus, if the Erns RNase was engaged in blocking a process leading to IFN-1 expression, this process should most likely take place outside of the infected cell. A formal proof for the involvement of the Erns RNase in the prevention of innate immunological control of virus infection was obtained in a special animal model. Pestiviruses are known to establish long-lasting persistent infections when introduced into the fetus in a pregnant host animal (Thiel et al. 1996). This process is best understood for the pestivirus bovine viral diarrhea virus (BVDV) (Fig. 6.7). Intrauterine infection of a fetus in the first trimester (day 40–120) of gestation by noncytopathic BVDV may lead to viral persistence accompanied by an acquired immunotolerance with high specificity for the infecting virus strain (Fig. 6.7). It is generally believed that self-reactive elements of the adaptive immune system, including the ones directed against the persisting virus, are inactivated in this developmental stage. Thus, the developing individuals do not produce an adaptive immune response against the persisting virus strain for their lifetime (Thiel et al. 1996). Furthermore, the virus is protected against the immune response of the mother cow by the barrier established by the bovine placenta that cannot be crossed by antibodies. To maintain a persistent infection for years, BVDV has to deal also with the innate immune system of the host. As a matter of fact, persistently infected fetuses or calves do not mount an IFN-1 response despite massive virus replication resulting in large amounts of PAMPs (Charleston et al. 2001). The absence of innate immune reactions is regarded as an important contribution to viral persistence. When fetuses were infected with an RNase-negative BVDV mutant, a significant IFN-1 response was observed in contrast to parallel experiments with the corresponding wild-type virus (Meyers et al. 2007). From these experiments, it can be concluded that the Erns RNase is involved in blocking the IFN-1 response to pestivirus infection in vivo. Similar results were also obtained for Npro deletion mutants. Most importantly, the combination of the Npro deletion and the

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infection of pregnant cow -day 50 to 100 of gestation -noncp BVDV diaplacental infection of fetus -persistent infection of fetus -immunotolerance

persistently infected (P.I.) animal P.I. calf may be clinically unaffected P.I. calf sheds large amounts of virus

-no antibodies against persisting virus -no T-cells specific for persisisting virus -Erns RNase involved in blocking of innate immune response

Fig. 6.7 The cartoon shows the principle pathway to virus-specific acquired immunotolerance and establishment of persistent infection by BVDV

RNase-inactivating mutation resulted in a virus that provoked an overboosting IFN-1 response in the fetus that finally led to its abortion (Meyers et al. 2007). Taken together the experimental evidence available so far shows that the Erns RNase is one of the pestiviral factors responsible for inhibition of the innate immune response. These viral factors are especially important for the establishment and maintenance of persistent pestivirus infections. Since persistently infected animals play a crucial role in the strategy that keeps pestiviruses within their host populations, the Erns RNase is of major importance for these viruses and was therefore conserved during virus evolution.

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.

Chapter 7

50 -30 Exoribonucleases Jeong Ho Chang, Song Xiang, and Liang Tong

Contents 7.1 7.2 7.3 7.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sequence Conservation of the XRNs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 -30 Exonuclease Activity of XRNs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Functions of Xrn1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4.1 Functions of Xrn1 in RNA Degradation and Turnover . . . . . . . . . . . . . . . . . . . . . . . . 7.4.2 Functions of Xrn1 in RNA Maturation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4.3 Functions of Xrn1 in DNA Recombination and Chromosome Stability . . . . . . 7.4.4 Other Functions of Xrn1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5 Functions of Xrn2/Rat1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5.1 Functions of Xrn2/Rat1 in RNA Processing and Degradation . . . . . . . . . . . . . . . . . 7.5.2 Functions of Xrn2/Rat1 in RNA Polymerase Transcription Termination . . . . . 7.5.3 Other Functions of Xrn2/Rat1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6 Protein Partners of XRNs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7 Functions of XRNs in Plants and Other Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.8 Overall Structure of Xrn2/Rat1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.9 Active Site of Xrn2/Rat1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.10 Structure of the Rat1-Rai1 Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.11 Rai1/Dom3Z and RNA 50 -End Capping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.12 The 50 -30 Exoribonuclease Rrp17 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.13 RNase J1/CPSF-73 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.14 Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

168 168 170 172 172 174 174 175 175 176 176 177 178 179 179 181 182 183 185 185 186 186

J.H. Chang • L. Tong (*) Department of Biological Sciences, Columbia University, New York, NY 10027, USA e-mail: [email protected] S. Xiang Department of Biological Sciences, Columbia University, New York, NY 10027, USA Key Laboratory of Nutrition and Metabolism, Institute for Nutritional Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200031, P.R. China A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_7, # Springer-Verlag Berlin Heidelberg 2011

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Abstract The 50 -30 exoribonucleases have important functions in RNA processing, RNA degradation, RNA interference, transcription, and other cellular processes. The Xrn1 and Xrn2/Rat1 family of enzymes are the best characterized 50 -30 exoribonucleases, and there has been significant recent progress in the understanding of their structure and function. Especially, the first structural information on Rat1 just became available. Other 50 -30 exoribonucleases have been identified recently, including yeast Rrp17 and B. subtilis RNase J1, the first enzyme with 50 -30 exoribonuclease activity found in prokaryotes. This review will summarize our current understanding of these enzymes, focusing on their sequence conservation, molecular structure, biochemical and cellular functions.

7.1

Introduction

Exoribonucleases are involved in RNA processing, RNA degradation, RNA interference, transcription, modulation of gene expression, antiviral defense, and other cellular processes. These enzymes can be simply classified based on the direction of their activity, hence 50 -30 or 30 -50 exoribonucleases. While a large number of 30 -50 exoribonucleases have been identified, in bacteria and eukaryotes (Zuo and Deutscher 2001) (see also Chap. 8), few 50 -30 exoribonucleases are currently known. The best characterized 50 -30 exoribonucleases are the Xrn1/Xrn2 family of enzymes (to be referred to as XRNs here), which have only been found in eukaryotes. Recently, Rrp17 was identified as another 50 -30 exoribonuclease, with an important role in the 50 -end processing of pre-ribosomal RNAs (Oeffinger et al. 2009). Several enzymes that possess both endo- and 50 -30 exoribonuclease activity have also been reported, including B. subtilis RNase J1 (Condon 2010), the first enzyme with 50 -30 exoribonuclease activity found in prokaryotes (see also Chap. 10). RNase J1 is structurally homologous to human CPSF-73 (Mandel et al. 2006), which has also been suggested to have 50 -30 exoribonuclease activity (Dominski et al. 2005) in addition to its endonuclease activity. In this chapter, we will focus on the sequence conservation, structure, and function of the XRNs (Sects. 7.2–7.11). We will also discuss the other 50 -30 exoribonucleases, including Rrp17 (Sect. 7.12) and RNase J1/CPSF-73 (Sect. 7.13).

7.2

Sequence Conservation of the XRNs

Yeast and most metazoans have two XRNs, with Xrn1 (175 kD) primarily in the cytoplasm and Xrn2 (115 kD, more commonly known as Rat1 in yeast) primarily in the nucleus. RAT1 is an essential gene in yeast, while deletion of XRN1 in yeast leads to slow growth, sporulation defect, DNA recombination defect, and other phenotypes. The plant Arabidopsis has three XRNs, two of which (AtXRN2 and

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Fig. 7.1 Sequence conservation of XRNs. Schematic drawing of the domain organization of human Xrn1, S. cerevisiae Xrn1, human Xrn2, S. cerevisiae Rat1, and S. pombe Rat1. The two highly conserved regions (CR1 and CR2) are labeled. The 570-residue weakly conserved segment in Xrn1 and a 120-residue segment in Xrn2/Rat1 are indicated. Small triangles in S. cerevisiae Xrn1 indicate protease-sensitive sites. The segment at the extreme C-terminus of these proteins is not required for activity

AtXRN3) are in the nucleus, while the third (AtXRN4) is in the cytoplasm (Kastenmayer and Green 2000). However, all three Arabidopsis XRNs are Xrn2 homologs, and a sequence homolog of Xrn1 may not exist in higher plants. The amino acid sequences of the XRNs contain two highly conserved regions (CR1 and CR2) in their N-terminal segment (Fig. 7.1). The sequence identity among Xrn2 homologs for these two regions is 50–60%, while that between Xrn1 and Xrn2 homologs is about 40–50%. In comparison, conservation of sequences outside of these two regions is much lower, especially between Xrn1 and Xrn2. In fact, the larger size of Xrn1 is due to an extensive C-terminal segment that is absent in Xrn2. The linker between CR1 and CR2 is also poorly conserved among the XRNs, both in sequence and in length (Fig. 7.1). Several protease-sensitive sites identified in S. cerevisiae Xrn1 map to the boundaries of these segments (Fig. 7.1) (Page et al. 1998). CR1 covers residues 1–354 of human Xrn1 and residues 1–407 of human Xrn2 (Fig. 7.1), as the latter has three small inserted segments. CR1 contains seven strictly conserved acidic residues (Asp35, Asp86, Glu176, Glu178, Asp206, Asp208, and Asp292 in human Xrn1), and it was recognized that these residues may be homologous to those in the active site of several other Mg2+-dependent nucleases (Solinger et al. 1999), even though CR1 shares little overall sequence conservation with these other enzymes. Therefore, CR1 may have a crucial role in the active site of the XRNs, which is supported by the fact that mutations of these acidic residues abolish the exonuclease activity (Johnson 1997; Page et al. 1998; Solinger et al. 1999). It is expected that the seven conserved acidic residues can coordinate two Mg2+ ions for catalysis (Yang et al. 2006).

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CR2 covers residues 426–595 of human Xrn1 and residues 509–679 of human Xrn2 (Fig. 7.1). This segment appears to be unique to the XRNs, and has an important role in defining the overall landscape of the active site of the XRNs (see Sect. 7.9). A 570-residue segment directly following CR2 shows weak sequence conservation among the Xrn1 enzymes (Fig. 7.1). For example, human and yeast Xrn1 share 26% sequence identity for this segment. In contrast, the remaining C-terminal segments of the Xrn1 enzymes have little sequence conservation. This C-terminal segment of yeast Xrn1 is dispensable for its exoribonuclease activity and in vivo function, while the 570-residue segment, though weakly conserved, is required for activity (Page et al. 1998). The Xrn2 enzymes have a roughly 240-residue C-terminal segment following CR2 (Fig. 7.1). Human Xrn2 and yeast Xrn2/Rat1 share 24% sequence identity for this segment. The last 125 residues of S. pombe Rat1 can be deleted without affecting its in vivo function at the permissive temperature (the truncation does lead to a ts phenotype). Further deletions, removing the C-terminal 204 residues, inactivated the protein (Shobuike et al. 2001). Observations on the C-terminal deletion mutants of Xrn1 and Xrn2/Rat1 described above suggest that CR1 and CR2, while highly conserved among the XRNs, are not sufficient for the activity of these enzymes. A segment following CR2 (roughly 570 residues for Xrn1 and 120 residues for Xrn2/Rat1) is required for activity, even though it is only weakly conserved. The segment of the XRNs containing CR1 and CR2 is generally acidic in nature, with a pI of 5.6 for this segment of yeast Xrn1. In contrast, the remaining C-terminal segments of the Xrn1 enzymes are much more basic, with a pI of 9.4 for yeast Xrn1 (Page et al. 1998).

7.3

50 -30 Exonuclease Activity of XRNs

The XRNs are Mg2+-dependent, processive 50 -30 exoribonucleases (Stevens 1978, 1980; Stevens and Poole 1995). Mn2+ can also support the catalytic activity of these enzymes. They generally prefer single-stranded RNA substrates with a 50 -end monophosphate group. RNAs with a hydroxyl, cap, or triphosphate group at the 50 -end are poor substrates for XRNs (Stevens 1978; Stevens and Poole 1995). Yeast Xrn1 and Rat1 also have weak exonuclease activity toward single-stranded DNA (Page et al. 1998; Solinger et al. 1999; Stevens and Poole 1995). Yeast Xrn1 can cleave G4 tetraplex DNA derived from guanine-rich sequences that are normally found in telomeres (Liu and Gilbert 1994), while mouse Xrn1 can also cleave G4 tetraplex RNA (Bashkirov et al. 1997). The presence of strong secondary structures in the RNA substrate can block or stall the exoribonuclease activity of yeast Xrn1 and Rat1 (Poole and Stevens 1997; Stevens and Poole 1995). A strong stem loop at the 50 -end of the genome of


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Narnavirus 20 S RNA, a persistent virus in yeast, protects it from degradation by Xrn1 (Esteban et al. 2008). The exoribonuclease activity of yeast Xrn1 and Rat1 is inhibited by adenosine 30 ,50 bisphosphate (pAp) (Dichtl et al. 1997). Nearly 80% inhibition of both Xrn1 and Rat1 can be achieved with 1 mM pAp. The inhibition of Xrn1 is not affected by the concentration of the RNA substrate, suggesting that pAp may not compete against RNA. pCp and pUp are as potent as pAp in inhibiting Xrn1, while 50 or 30 AMP is essentially inactive. pAp is a byproduct of the sulfate assimilation pathway, and is normally converted to 50 AMP and Pi by the enzyme 30 ,50 bisphosphate nucleotidase, Hal2/Met22 in yeast. Hal2 is inhibited by submillimolar concentrations of Li+, and the resulting increase in cellular pAp concentration (up to 3 mM) and the consequent inhibition of Xrn1 and Rat1 may be part of the mechanism for Li+ toxicity in yeast. A similar mechanism may contribute to the physiological effects of Li+ in other organisms, including the therapeutic effects of Li+ for the treatment of various neurological diseases in humans. The cellular functions of the XRNs are primarily linked to their exoribonuclease activity. Therefore, these enzymes are involved in the turnover of mRNAs and degradation of aberrant mRNAs (quality control) (Fig. 7.2). They are also involved in the maturation (50 trimming) of ribosomal RNAs (rRNAs), small nucleolar RNAs (snoRNAs), and others, as well as the degradation of hypomodified mature tRNAs and spacer RNA byproducts from rRNA processing. The exoribonuclease activity of Xrn2/Rat1 also contributes to transcription termination by nuclear RNA polymerases I and II (Pol I and Pol II). The physiological functions of Xrn1 and

Xrn1

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Fig. 7.2 Schematic drawing of the functions of XRNs

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Xrn2/Rat1 will be described in more detail in the following two sections, and the functions of the plant XRNs are decribed in Sect. 7.7. Some of the functional differences between Xrn1 and Xrn2/Rat1 are due to their different cellular localizations. However, a nuclear-targeted Xrn1 can rescue the lethal phenotype of rat1-1 (carrying a ts mutation in RAT1) yeast cells, suggesting that Xrn1 can complement the essential function of Rat1 (Johnson 1997). Conversely, RAT1 expressed from a high copy-number plasmid, as well as Rat1 without its nuclear localization sequence (NLS), can rescue the defects due to the loss of Xrn1 (Johnson 1997). The XRNs may have other functions that are independent of their exonuclease activity. For example, they may mediate protein–protein interactions to recruit other proteins or to be recruited by other proteins and/or RNA to proper locations in the cell. Especially, yeast Rat1 is known to form a stable complex with Rai1 (Rat1 interacting protein 1), which can stimulate the exoribonuclease activity of Rat1. Rat1 may also interact with other protein factors that are important for Pol II termination, including Rtt103. Yeast Xrn1 may interact directly with microtubules. The protein complexes for Xrn1 and Xrn2/Rat1 are described in a Sect. 7.6.

7.4

Functions of Xrn1

Xrn1 nuclease activity was first identified in yeast (Larimer et al. 1992; Stevens 1978). Later studies showed that the enzyme is identical to several other proteins isolated based on other biochemical and functional properties (Kearsey and Kipling 1991), DNA strand exchange protein 1 including (Sep1) (Tishkoff et al. 1991), DNA strand transferase 2 (Dst2) (Dykstra et al. 1991), Kar enhancing mutant 1 (Kem1) (Kim et al. 1990), and radiation-resistant on 5 (Rar5) (Kipling et al. 1991). Xrn1 may also be identical to the antiviral superkiller 1 (Ski1) protein (Johnson and Kolodner 1995). This illustrates the various functions for this enzyme other than RNA metabolism, such as DNA recombination, chromosome stability, microtubule association, nuclear fusion, meiosis, telomere maintenance, and cellular senescence. Defects in many of these processes are observed in cells lacking Xrn1 (Larimer and Stevens 1990). Xrn1 homologs in S. pombe (also known as Exo II) (Kaslin and Heyer 1994) and higher eukaryotes have also been cloned, including C. elegans (Newbury and Woollard 2004), Drosophila (Pacman) (Till et al. 1998), mouse (Bashkirov et al. 1997), and humans (Sato et al. 1998; Shimoyama et al. 2003).

7.4.1

Functions of Xrn1 in RNA Degradation and Turnover

Xrn1 has important roles in mRNA degradation and turnover. This subject has been reviewed extensively over the past few years (Conti and Izaurralde 2005; Doma and


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Fig. 7.3 Schematic drawing of mRNA turnover and mRNA degradation pathways

Parker 2007; Houseley and Tollervey 2009; Isken and Maquat 2007; Parker and Song 2004), and will only be discussed briefly here, focusing on the functions of Xrn1 in these processes. The basic mode of action is that RNAs with a 50 -end monophosphate are generated by decapping of mRNAs (possibly preceded by deadenylation) or by endonucleolytic cleavage, which are then rapidly degraded by Xrn1 (Fig. 7.3). The 30 -50 exosome also plays an important role in mRNA metabolism (see Chap. 9), although Xrn1 is the primary enzyme for mRNA degradation and turnover in yeast. The rate of mRNA turnover is reduced in yeast cells lacking Xrn1, leading to accumulation of non-polyadenylated mRNAs that also partially lack the 50 -end cap structure (Hsu and Stevens 1993). Xrn1 is predominantly localized to cytoplasmic foci known as P-bodies (processing bodies/GW bodies), which are the major location for mRNA decapping and 50 -30 degradation as well as for temporary storage of mRNAs during translation repression (Kulkarni et al. 2010; Parker and Sheth 2007). Recent studies show that decapping and Xrn1-mediated degradation of mRNAs can also occur on actively translating ribosomes (Hu et al. 2009), as does deadenylation-independent decapping initiated by nonsense-mediated decay (NMD) (Hu et al. 2010). Endonucleolytic cleavage of mRNAs can be initiated by no-go decay (NGD) and by the RNA-induced silencing complex (RISC) for RNA interference (RNAi) (Fig. 7.3) (Orban and Izaurralde 2005). In addition, endonucleolytic cleavage during maturational processing of many RNA precursors can produce byproducts that are degraded by Xrn1. For example, Xrn1 degrades the internal transcribed spacer ITS1 generated from pre-ribosomal RNA processing in yeast (Fig. 7.4) (Stevens et al. 1991).

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Fig. 7.4 Schematic drawing of the pre-ribosomal RNA processing pathways. The extent of the exonuclease trimming is indicated by the arrows

Recently, it has been found that Xrn1 and Rat1 can degrade hypomodified mature tRNAs in yeast, in the rapid tRNA decay (RTD) pathway (Chernyakov et al. 2008).

7.4.2

Functions of Xrn1 in RNA Maturation

Xrn1 plays a role in pre-ribosomal RNA processing and maturation, which may be especially important in the absence of Rat1 activity in yeast. This will be discussed in more detail in Sect. 7.5.1.

7.4.3

Functions of Xrn1 in DNA Recombination and Chromosome Stability

Xrn1 was identified in a biochemical search for DNA recombination proteins (and hence named Sep1 and Dst2) (Dykstra et al. 1991; Tishkoff et al. 1991). It has homologous pairing and strand exchange activities in vitro. Yeast cells lacking Xrn1 are defective for intrachromosomal recombination, sporulation, and trigger


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arrest at pachytene stage in the meiotic cell cycle (Solinger et al. 1999; Tishkoff et al. 1995). On the other hand, Xrn1 may not be involved in mitotic recombination or mating-type switching. Xrn1 was identified from a genetic screen for mutants that can enhance the nuclear fusion defect of yeast cells carrying the kar1-1 mutation (hence named Kem1) (Kim et al. 1990). Kem1 mutants also have reduced chromosome stability and are hypersensitive to the microtubule-destabilizing drug benomyl. Defective interactions with microtubules may be the basis of these phenotypes (see Sect. 7.6). Yeast cells lacking Xrn1 also show cellular senescence and telomere shortening (Liu et al. 1995), which may be related to the nuclease activity of this enzyme toward G4 tetraplex DNA (Liu and Gilbert 1994). Most of the defects in these nuclear processes (sporulation defect, arrest at pachytene, chromosome instability) due to loss of Xrn1 can be rescued by targeting Rat1 to the cytoplasm (Johnson 1997); consistent with the fact that Xrn1 is primarily a cytoplasmic protein. This also suggests the possibility that the effects of Xrn1 on these processes may not be direct.

7.4.4

Other Functions of Xrn1

Human Xrn1 may function as a tumor suppressor in osteogenic sarcoma, and its expression level is reduced in these tumors (Zhang et al. 2002). Mouse Xrn1 is highly expressed in testis, suggesting a functional role in gametogenesis (Shobuike et al. 1997). Drosophila Xrn1/Pacman is required for male fertility (Zabolotskaya et al. 2008). The expression level of Pacman is correlated with developmental stages in Drosophila (Till et al. 1998), and C. elegans Xrn1 is critical for ventral epithelial enclosure during embryogenesis (Newbury and Woollard 2004). Xrn1 is also involved in host antiviral response. It can suppress viral RNA recombination (Cheng et al. 2006), and down-regulate replication by HIV (Chable-Bessia et al. 2009) and HCV (Jones et al. 2010).

7.5

Functions of Xrn2/Rat1

Like Xrn1, Xrn2 was first identified from several independent studies, due to its different functions. It was found from a screen for ribonucleic acid trafficking defects in yeast, and hence named Rat1 (Amberg et al. 1992), and from a screen for protein translation defects (Hke1, homology to Kem1), which are more likely due to defects in RNA processing or trafficking (Kenna et al. 1993). It was also found to have functions in transcription activation (Tap1) (Aldrich et al. 1993; di Segni et al. 1993). In contrast to XRN1, RAT1 is an essential gene in yeast, although the exact function (or the substrate) of this protein that is required for cell viability is currently not known.

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Homologs of Rat1/Xrn2 in other organisms have also been cloned, including S. pombe (also named Dhp1) (Shobuike et al. 2001; Sugano et al. 1994), mouse (Dhm1) (Shobuike et al. 1995), and humans (Zhang et al. 1999).

7.5.1

Functions of Xrn2/Rat1 in RNA Processing and Degradation

Rat1 is required for 50 -end trimming during the maturation of the 5.8 S and 25 S rRNA, and Xrn1 can support this activity in the absence of Rat1 (El Hage et al. 2008; Fang et al. 2005; Fatica and Tollervey 2002; Geerlings et al. 2000; Henry et al. 1994). The 5.8 S, 18 S and 25 S ribosomal RNAs are made in a single transcript by Pol I in eukaryotes, which undergoes extensive endo and exonucleolytic processing (Fig. 7.4). The primary transcript includes two external transcribed spacers (50 - and 30 -ETS) and two internal transcribed spacers (ITS1 and ITS2) (Fig. 7.4). Rat1/Xrn1 is involved in the degradation of a fragment of ITS1 that is released during pre-rRNA processing. Recent studies identified Rrp17 as an independent 50 -30 exoribonuclease that can also process the 50 -ends of 5.8 S and 25 S rRNA (see Sect. 7.12) (Oeffinger et al. 2009). Rat1 is required for 50 -end processing of polycistronic and some intronic snoRNAs in yeast, and Xrn1 can (at least partially) support this activity (Lee et al. 2003; Petfalski et al. 1998; Qu et al. 1999; Villa et al. 1998). Rat1 and Xrn1 are involved in the degradation of some intron-containing unspliced pre-mRNAs and intron lariats (Danin-Kreiselman et al. 2003). The entry sites for the XRNs are produced by prior endonucleolytic cleavage or by debranching of the intron lariat. Rat1 degrades telomeric repeat-containing RNA (TERRA) in yeast (Luke et al. 2008). Telomeres are transcribed by Pol II and polyadenylated, and cells lacking Rat1 accumulate TERRA and have short telomeres. Therefore, Rat1 promotes telomere elongation and is important for telomerase regulation.

7.5.2

Functions of Xrn2/Rat1 in RNA Polymerase Transcription Termination

Xrn2/Rat1 has a central role in the torpedo model for transcription termination by RNA polymerases I and II. This area has been reviewed extensively over the past few years (Buratowski 2005; Ghazal et al. 2009; Gilmour and Fan 2008; Luo and Bentley 2004; Richard and Manley 2009; Rondon et al. 2009), and will only be briefly discussed here. The torpedo model suggests that the downstream RNA product, with a 50 -monophosphate, produced by an endonucleolytic cleavage of the primary transcript serves as the entry point for a 50 -30 exoribonuclease, which degrades this downstream RNA, catches up to the elongating (or paused) polymerase, and


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Fig. 7.5 Schematic drawing of the allosteric-torpedo (unified) model of Pol II termination. Changes in the phosphorylation state of the CTD and in the body of Pol II are indicated (Modified from Luo et al. 2006)

causes transcription termination (Connelly and Manley 1988). The 50 -30 exoribonuclease for Pol II termination was identified as Rat1 in yeast and Xrn2 in mammalian cells (Kim et al. 2004; West et al. 2004). It was shown more recently that Pol I transcription termination is also mediated by the torpedo model, with Rat1 being the 50 -30 exoribonuclease for this function in yeast (Fig. 7.5) (El Hage et al. 2008; Kawauchi et al. 2008). The molecular mechanism for how Xrn2/Rat1 brings about transcription termination once it catches up to the polymerase is still not clearly understood. Degradation of the downstream product is not sufficient for termination. Nuclear-targeted Xrn1 can degrade the downstream product in yeast cells lacking Rat1, but nuclear Xrn1 cannot cause Pol II termination (Luo et al. 2006). In addition, Rat1 alone is not sufficient for Pol II termination in an in vitro transcription system (Dengl and Cramer 2009). Therefore, other factors are also required for transcription termination by Rat1/Xrn2. The pre-mRNA 30 -end processing factor Pcf11 may be important for dismantling the polymerase elongation complex (Luo et al. 2006; West and Proudfoot 2008; Zhang et al. 2005).

7.5.3

Other Functions of Xrn2/Rat1

Xrn2 is a candidate gene for spontaneous lung tumor susceptibility based on a genome-wide association study in mice (Lu et al. 2010). In addition, polymorphisms in human Xrn2 are associated with human lung cancer, and over-expression of human Xrn2 can affect the differentiation of a leukemia cell line (Park et al. 2007).

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Protein Partners of XRNs

Xrn1 is associated with the decapping machinery in yeast and may directly interact with several of its components, including Dcp1/Dcp2, Pat1, Dhh1, and the Lsm1–7 complex (Coller and Parker 2004). This may facilitate the degradation of RNAs once they are decapped by this machinery. The region(s) of Xrn1 that is required for these interactions has not been identified. Yeast Xrn1 interacts directly with tubulin and promotes microtubule assembly (Interthal et al. 1995). Cells lacking Xrn1 show increased chromosome loss, defects in spindle pole body separation and karyogamy, and hypersensitivity to benomyl (Kim et al. 1990). The exonuclease activity of Xrn1 is not required for this interaction (Solinger et al. 1999). The benomyl sensitivity of cells lacking Xrn1 can be rescued by targeting Rat1 to the cytoplasm, although cytoplasmic Rat1 does not appear to be associated with microtubules (Johnson 1997). In yeast, Rat1 has direct and strong association with Rai1, and the Rat1-Rai1 complex was first purified from S. cerevisiae extract (Stevens and Poole 1995). A stable Rat1-Rai1 complex was also observed in S. pombe (Shobuike et al. 2001). Rai1 (45 kD) has orthologs in most eukaryotes, including plants, and the mammalian homolog is known as Dom3Z (Xue et al. 2000). The sequences of these orthologs are highly divergent, however, with only a few conserved residues. In contrast to Rai1, Dom3Z does not appear to interact with Xrn2. Rai1 is not essential for yeast cell viability, and does not have any nuclease activity (Xue et al. 2000). However, Rai1 can moderately stimulate the exoribonuclease activity of Rat1 (Xiang et al. 2009; Xue et al. 2000). This may be due in part to the stabilization of Rat1 by Rai1. Rat1 is unstable and quickly loses activity upon pre-incubation at 30 C, whereas the Rat1-Rai1 complex is able to retain most of its nuclease activity during this pre-incubation (Xue et al. 2000). Like Rat1, Rai1 is also required for 5.8 S rRNA processing. However, while Rat1 is only involved in the 50 -end processing of this RNA, Rai1 is also needed for 30 -end processing (Fang et al. 2005; Xue et al. 2000). The Drosophila genome contains two homologs of Rai1/Dom3Z: CG9125 and CG13190. CG13190, also known as Cutoff (Cuff), was first identified in a femalesterile screen. cuff mutations affect germline cyst development, produce ventralized eggs, and reduce female fecundity (Chen et al. 2007). Over-expressed Cuff is localized in the cytoplasm and in perinuclear puncta, and Cuff does not interact with Drosophila Xrn2. S. cerevisiae also has a homolog of Rai1, Ydr370c, which is poorly conserved with Rai1 at the sequence level (Xue et al. 2000). The function of this protein is currently not known. Sequence analysis suggests that this homolog is restricted to only a few of the fungal species, while most other eukaryotes contain only one homolog of Rai1/Dom3Z. Rtt103 (regulation of Ty1transposition 103) can interact with the Rat1-Rai1 complex in yeast (Dengl and Cramer 2009; Kim et al. 2004). Rtt103 was originally found by a screen for mutants that increased Ty1 transposon’s mobility (Scholes


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et al. 2001). Rtt103 has a RNA Pol II carboxy-terminal domain (CTD)-interacting domain (CID), and recognizes Ser2 phosphorylated CTD. Rtt103 may be involved in nuclear pre-mRNA regulation (Kim et al. 2004), and it localizes at the 30 -end of transcribing genes together with Rat1-Rai1 in vivo (Kim et al. 2004) and in vitro (Dengl and Cramer 2009). A functional interaction between Rat1 and the pre-mRNA 30 -end processing factor Pcf11 has been suggested (Luo et al. 2006; West and Proudfoot 2008), although currently there is no biochemical evidence for direct interaction between these two proteins. Pcf11 may be responsible for the recruitment of Rat1 to the 30 end of pre-mRNAs and/or vice versa.

7.7

Functions of XRNs in Plants and Other Organisms

In Arabidopsis, AtXRN2 is involved in 50 -end processing of 5.8 S and 25 S rRNAs (Zakrzewska-Placzek et al. 2010), a function similar to that of Rat1. In addition, both AtXRN2 and AtXRN3 can degrade miRNA loop and transgene for suppressing endogenous post-transcriptional gene silencing (Gy et al. 2007). The cytoplasmic AtXRN4 can degrade specific RNA transcripts but may not be a general RNA degradation enzyme, in contrast to Xrn1. It degrades 30 -end mRNA products derived from miRNA-mediated cleavage (Souret et al. 2004). Mutation of AtXRN4 leads to accumulation of decapped mRNAs that could be templates for facilitating the RNAi pathway, and AtXRN4 may link mRNA degradation and RNA silencing (Gazzani et al. 2004; Gregory et al. 2008). AtXRN4 also contributes to the regulation of the ethylene response pathway (and hence is also known as EIN5, ETHYLENE-INSENSITIVE5) (Olmedo et al. 2006; Potuschak et al. 2006). In Trypanosoma brucei and other kinetoplastids, four XRN-related proteins have been identified, XRNA, XRNB, XRNC, and XRND (Li et al. 2006). XRND is nuclear, XRNB and XRNC are cytoplasmic, and XRNA is present in both compartments. XRNA and XRND are essential for growth, and XRNA is required for degrading highly unstable, developmentally regulated mRNAs, while having little effect on more stable, unregulated mRNAs (Li et al. 2006).

7.8

Overall Structure of Xrn2/Rat1

Crystal structure of the S. pombe Rat1-Rai1 complex is the first structural information on the XRNs (Xiang et al. 2009). The structure of Rat1 indicates that CR1 and CR2 form a single, large domain (Fig. 7.6a). CR1 is composed of a seven-stranded (b1 through b7) mostly parallel b-sheet sandwiched by a-helices on both faces. Strands b2 through b7 are arranged similar to those in the Rossmann fold, but with strand b7 running in the opposite direction. A helix is inserted after b2 (aΒ) and b7 (aD). CR2 contains several helices and long loops, which wrap around the base

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Fig. 7.6 Structure of the S. pombe Rat1–Rai1 complex. (a) Schematic drawing of the structure of S. pombe Rat1–Rai1 complex (Xiang et al. 2009). The active site of Rat1 is indicated with the star, and the arrow points to the opening of the Rai1 active site pocket. A bound divalent metal cation in the active site of Rai1 is shown as a sphere. (b) Schematic drawing of the active site of S. pombe Rat1. Side chains of residues in the active site are shown and labeled. Overall molecular surface of (c) Rat1, (d) FEN-1 (Chapados et al. 2004), and (e) T4 RNase H (Devos et al. 2007). The active site is indicated with the star

(N-terminal end) of the aD helix. Residues in the linker between CR1 and CR2 are mostly disordered in the structure. The N- and C-termini of this segment are located ˚ of each other, suggesting that it is likely an inserted cassette between within 10 A the two conserved regions (Fig. 7.6a). A striking feature of the S. pombe Rat1 structure is the long aD helix, with its ˚ away from the rest of the structure (Fig. 7.6a). This C-terminus projected 30-A feature has been named the “tower domain.” The N-terminal residues of helix aD are strongly conserved among XRNs, and they contribute to the formation of the active site (see Sect. 7.9). The C-terminal residues of this helix are poorly


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conserved, and sequence analysis indicates that this helix is much shorter in Xrn1. Two temperature-sensitive mutations in XRNs, P90L in Xrn1 (Page et al. 1998), and Y657C in Rat1 (the rat1-1 mutation) (Luo et al. 2006), are located near the N-terminal end of helix aD. Both mutations may destabilize this helix at the nonpermissive temperature, supporting the functional importance of the tower domain. The structure of CR1 has many homologs, most of which are nucleases. These include the FEN-1 family of endonucleases (Chapados et al. 2004; Hwang et al. 1998; Sakurai et al. 2005; Sayers and Artymiuk 1998), the 50 exonuclease from the phage T5 (Ceska et al. 1996), RNase H from the phage T4 (Devos et al. 2007; Mueser et al. 1996), the 50 nuclease domain of Taq DNA polymerase (Kim et al. 1995; Murali et al. 1998), and other PIN domain-containing nucleases (Clissold and Ponting 2000; Glavan et al. 2006). The sequence conservation between Rat1 and these other enzymes is very low, between 8% and 15%. The structural homology is limited to strands b2-b7 in the central b-sheet and a few of the flanking helices. The tower domain in Rat1 is equivalent to the helical clamp in A. fulgidus FEN-1 (Chapados et al. 2004) and the helical arch in T5 exonuclease (Ceska et al. 1996), but the equivalent region is a long loop in M. jannaschii FEN-1 (Hwang et al. 1998) and is disordered in T4 RNase H (Devos et al. 2007; Mueser et al. 1996). The Rat1 structure covers residues 1–874, which are sufficient for the activity of this protein at the permissive temperature (Shobuike et al. 2001). The 120-residue segment following CR2 forms three distinct structural features (Fig. 7.6a). The N-terminal region (residues 752–840) of this segment adds four anti-parallel strands (b8–b11) to the central b-sheet of CR1, producing a highly twisted 11-stranded b-sheet. Residues 841–863 form a long loop that traverses the entire bottom face of the central b-sheet of CR1. Finally, the C-terminal region of this segment (residues 864–874) forms an a-helix that interacts with helices aA and aH in CR1. Therefore, despite being poorly conserved among XRNs, this segment has an important structural role, which may explain why it is required for the function of Rat1. The strong sequence conservation for CR1 and CR2 suggests that these two segments should have a similar structure in Xrn1 (with the exception of the tower domain). On the other hand, most of the 570-residue segment following CR2 is unique to Xrn1 and forms several distinct structural domains, as revealed by the structure of Xrn1 (unpublished data).

7.9

Active Site of Xrn2/Rat1

The active site of Rat1 is located at the top of the central b-sheet of CR1, with contributions from residues at the base of the aD helix (Fig. 7.6a). The seven conserved acidic residues in CR1 form a cluster, and are located in the center of the active site (Fig. 7.6b). In the structure of bacteriophage T4 RNase H, two metal ions are associated with these acidic residues (Mueser et al. 1996), consistent with the hypothesis that the two metal ions mediate the nuclease activity (Yang et al. 2006). Metal ions were not observed in the structure of Rat1, and there are some

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noticeable differences in the conformations of some of these acidic side chains between Rat1 and T4 RNase H. Three positively-charged (Lys111, Arg118, Arg119) and two polar (Gln114, Gln115) residues at the base of the aD helix, as well as His61, His65, and Asn57 in helix aB contribute their side chains to the active site (Fig. 7.6b). These residues form a steep wall at one side of the active site, and may be important for interacting with the phosphate backbone of the RNA substrate. Mutations of these residues, as well as several of the conserved acidic residues, disrupt Rat1’s exonuclease activity. Residues in CR2 encircle the base of helix aD, but contribute few residues to the Rat1 active site. The side-chain hydroxyl groups of Tyr627 and Tyr628 hydrogenbond with the acidic residues Glu205 and Asp237 in the active site, respectively, and the side chain of Gln671 is located in the cluster of polar side chains from the aB and aD helices (Fig. 7.6b). However, CR2 introduces a dramatic difference in the overall landscape of the active site of Rat1 as compared to other related nucleases. Due to the presence of CR2, the Rat1 active site is a pocket (Fig. 7.6c), while the active sites of related nucleases are more open (Figs. 7.6d,e). It has been suggested that the ssDNA substrate threads through the helical arch in T5 nuclease (Ceska et al. 1996). In T4 RNase H, a single-stranded DNA portion of its forked DNA substrate is also bound in this region (Devos et al. 2007). However, such a binding mode would not be possible in Rat1, as the substrate would clash with residues in CR2. This may be the explanation why Rat1 is an exonuclease. The poorly conserved C-terminal segment of Rat1, following CR2, is located away from the active site and does not have any direct contributions to it. However, this segment is important for recruiting Rai1, which can (indirectly) stimulate the exoribonuclease activity of Rat1.

7.10

Structure of the Rat1-Rai1 Complex

The structure of the Rat1-Rai1 complex shows that Rai1 is bound on the opposite face from the Rat1 active site (Fig. 7.6a), interacting primarily with the poorly conserved C-terminal loop that traverses the bottom of CR1 (Xiang et al. 2009). ˚ 2 of surface area of each The Rat1-Rai1 interface buries approximately 800 A protein, consistent with the stability of this complex. Ion-pair, hydrogen-bonding, as well as van der Waals interactions contribute to the formation of this complex. Mutations introduced in this interface can abolish the interaction as well as the stimulation of Rat1 by Rai1 (Xiang et al. 2009). Rai1 does not directly contribute to the active site of Rat1. Structural and biochemical studies indicate that Rai1 enhances Rat10 s exonuclease activity at least in part by increasing the enzyme’s stability (Xue et al. 2000). This is also supported by the observation that over-expressing Rai1 can rescue a temperaturesensitive phenotype of Rat1 (Shobuike et al. 2001). On the other hand, real-time


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measurements of exoribonuclease activities of Rat1 and Rat1-Rai1 complex suggest that the Rat1 enzyme is inherently less active (Sinturel et al. 2009). Therefore, Rai1 may also indirectly help to organize the active site of Rat1. The structure of Rat1 alone, and comparison with the Rat1–Rai1 complex, may reveal any changes in the active site that is induced by Rai1 binding. Residues at the Rat1–Rai1 interface are generally conserved among the fungal proteins, consistent with the observations that Rat1 and Rai1 form tight complexes in both S. cerevisiae and S. pombe (Shobuike et al. 2001; Stevens and Poole 1995). However, Rai1 residues that interact with Rat1 are not conserved in the mammalian Rai1 homolog Dom3Z, and Dom3Z does not interact with mammalian Xrn2. Therefore, the Rat1–Rai1 interaction appears to be unique to the fungal species. Whether mammalian Xrn2 also has a protein partner that can stimulate its activity is currently not known.

7.11

Rai1/Dom3Z and RNA 50 -End Capping

An unexpected discovery from the structure of Rai1 is that it contains a large pocket (Figs. 7.7a,b), and the few residues that are highly conserved among Rai1 orthologs are all located in this pocket (rather than in the interface with Rat1) (Xiang et al. 2009). Moreover, three conserved acidic side chains, Glu150, Asp201, and Glu239 (S. pombe Rai1 numbering), together with the main-chain carbonyl of Leu240 and two water molecules octahedrally coordinate a divalent cation (Mg2+ or Mn2+) (Fig. 7.7c), and this metal ion is located near the bottom of the pocket (Fig. 7.7b). Therefore, the structural information strongly suggests that Rai1 and its mammalian homolog Dom3Z may have enzymatic activity of its own. Biochemical studies demonstrate that Rai1 possesses RNA 50 -end pyrophosphohydrolase activity, being able to remove a pyrophosphate group from RNA with 50 -end triphosphate (pppRNA) (Xiang et al. 2009). Such an enzyme (RppH) was first characterized in bacteria (Deana et al. 2008), which is a member of the Nudix family of enzymes. Rai1/Dom3Z shares neither sequence nor structural homology with RppH. Remarkably, Rat1 can stimulate this pyrophosphohydrolase activity of Rai1, even though the binding site is located far from the active site of Rai1 (Fig. 7.7a) (Xiang et al. 2009). Further biochemical studies showed that Rai1 can also remove unmethylated 50 end cap of RNAs (GpppRNA) (Jiao et al. 2010). This activity is however distinct from the classical decapping enzymes. First, Rai1 has much lower activity toward methylated 50 -end cap. Second, the product released by Rai1 is GpppN, while the classical decapping enzymes release m7GDP. Therefore, Rai1 appears to have two distinct enzymatic activities. The amino acid sequence of the Drosophila Rai1 homolog Cuff contains mutations at several of the conserved acidic residues that are important for metal ion binding. It is possible that Cuff does not have RNA 50 -end pyrophosphohydrolase and decapping activities.

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Fig. 7.7 Rai1 possesses an active site of its own. (a) Schematic drawing of the structure of S. pombe Rai1 (Xiang et al. 2009). A bound divalent cation in the active site is shown as a sphere. The arrow points to residues in the interface with Rat1. (b) Molecular surface of the active site region of Rai1, showing a large pocket. The metal ion is located at the bottom of the pocket. (c) Overlays of the metal ion binding site in the structure of Rai1 (in black) and mouse Dom3Z (in gray). Residue numbers in parenthesis are for Dom3Z. The interaction between Glu192 in Dom3Z and the metal ion is mediated by a water molecule

The biochemical activities of Rai1 suggest a physiological function for this enzyme. Rai1 may be an mRNA 50 -end capping quality checkpoint. Both Rai1 substrates (pppRNA and GpppRNA) are intermediates in the mRNA 50 -end capping pathway. mRNAs with defective 50 -end capping cannot serve as template for translation. At the same time, these defective mRNAs cannot be degraded by Xrn1/Rat1, due to the protected 50 -end. Therefore, Rai1 can remove the 50 -end from such mRNAs, and the products can then be rapidly degraded by Xrn1/Rat1. Studies in yeast show that mRNAs with 50 -end capping defects are stabilized in cells lacking Rai1, consistent with this 50 -end capping quality checkpoint model (Jiao et al. 2010). In addition, mRNAs with aberrant 50 -end capping also accumulate under stress conditions (glucose deprivation or amino acid starvation) in cells lacking Rai1. Moreover, defective capping in yeast cells is linked to enhanced recruitment of Rat1 throughout the transcribing unit, and promotes Pol II termination upstream of the poly(A) site (Jimeno-Gonzalez et al. 2010). This suggests that Rai1 can remove the defective cap in such conditions, which then allows Rat1 to


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function as a torpedo to induce Pol II termination before the completion of transcription, in a mechanism equivalent to that of transcription termination at the 30 -end of the pre-mRNA. These studies provide the first demonstration of an mRNA 50 -end capping checkpoint (Jiao et al. 2010; Xiang et al. 2009). It was generally believed in the field that 50 -end capping always proceeds to completion. The data on Rai1 convincingly demonstrate such a checkpoint in yeast. Dom3Z has a conserved active site, and it remains to be seen whether such a checkpoint also functions in metazoans.

7.12

The 50 -30 Exoribonuclease Rrp17

Rrp17 (ribosomal RNA processing) is associated with pre-ribosomes and the nuclear pore complex (Oeffinger et al. 2009). It is an independent nuclease for the 50 -end trimming of the 5.8 S and 25 S rRNAs. Rrp17 is an essential gene in yeast, and has highly conserved orthologs in most eukaryotes. Rrp17 has 50 -30 exoribonuclease activity, with preference for a phosphate group at the 50 -end of the substrate, while a triphosphate group or cap structure inhibits the nuclease activity. In comparison to the XRNs, Rrp17 also has activity toward RNAs with a 50 -end hydroxyl group. The activity requires Mg2+ ions, while the enzyme is inactive with Mn2+.

7.13

RNase J1/CPSF-73

Earlier studies have only identified 50 -30 exoribonucleases in eukaryotes, leading to the general belief that these enzymes are not present in prokaryotes. However, it was recently discovered that the B. subtilis endoribonuclease RNase J1 also has 50 -30 exoribonuclease activity, establishing for the first time the presence of such activity in bacteria (Condon 2010; Mathy et al. 2007). The exoribonuclease activity is required for mRNA degradation and for 50 -end maturation of 16 S rRNA in B. subtilis. Structural studies show that the endo- and exonuclease activities share the same active site, and suggest that RNase J1 may switch from an endo mode to exo mode on the same RNA substrate (de la Sierra-Gallay et al. 2008). The exoribonuclease activity of RNase J1 is more permissive toward 50 -end modification of the RNA substrate as compared to Xrn1 (Mathy et al. 2007). The highest activity is observed for RNA with a 50 -end monophosphate or 50 -end hydroxyl group, although this activity is roughly tenfold lower than that of Xrn1, leading to the suggestion that RNase J1 may require a cofactor for full activity. RNA with a 50 -end triphosphate group can also be degraded, but with roughly fourfold weaker activity. The activity toward RNA with a 50 -end cap is even lower (Mathy et al. 2007). RNase J1 exists in a complex with RNase J2, which is a sequence homolog of RNase J1 but with little nuclease activity. RNase J1 homologs are found in bacteria (but not in E. coli), archaea (Clouet-d’Orval et al. 2010; de la Sierra-Gallay et al.

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2008), and they may also be present in the chloroplasts of plants (de la SierraGallay et al. 2008). RNase J1 is a structural homolog of CPSF-73 (Mandel et al. 2006), the endoribonuclease for the cleavage step in eukaryotic pre-mRNA 30 -end processing (Mandel et al. 2008; Proudfoot 2004; Wahle and Ruegsegger 1999; Zhao et al. 1999). Recent studies suggest that CPSF-73 may also have an exoribonuclease activity, degrading the downstream cleavage product of histone pre-mRNAs (Dominski and Marzluff 2007; Dominski et al. 2005; Yang et al. 2009). Analogous to the RNase J1/J2 heterodimer, the CPSF complex also contains CPSF-100, an inactive sequence homolog of CPSF-73. It may be possible that mammalian CPSF-73/CPSF-100 and B. subtilis RNase J1/J2 share a common evolutionary origin.

7.14

Perspectives

Studies over the past few years have greatly enhanced our understanding of the structure and function of 50 -30 exoribonucleases, as well as identified new proteins that possess this activity. It is anticipated that further characterization of these enzymes in the coming years, especially in higher eukaryotes, will lead to significant new insights into the biological significance of these enzymes. Moreover, studies on Rat1–Rai1 complex led to the discovery of a novel mRNA 50 -end capping quality checkpoint. There may be further exciting surprises from the studies of these exoribonucleases and their interaction partners. Acknowledgment This research is supported in part by grants from the NIH to LT (GM077175).

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Shobuike T, Sugano S, Yamashita T, Ikeda H (1997) Cloning and characterization of mouse Dhm2 cDNA, a functional homolog of budding yeast SEP1. Gene 191:161–166 Shobuike T, Tatebayashi K, Tani T, Sugano S, Ikeda H (2001) The dhp1+ gene, encoding a putative nuclear 50 ->30 exoribonuclease, is required for proper chromosome segregation in fission yeast. Nucleic Acids Res 29:1326–1333 Sinturel F, Pellegrini O, Xiang S, Tong L, Condon C, Benard L (2009) Real-time fluorescence detection of exoribonucleases. RNA 15:2057–2062 Solinger JA, Pascolini D, Heyer W-D (1999) Active-site mutations in the Xrn1p exoribonuclease of Saccharomyces cerevisiae reveal a specific role in meiosis. Mol Cell Biol 19:5930–5942 Souret FF, Kastenmayer JP, Green PJ (2004) AtXRN4 degrades mRNA in Arabidopsis and its substrates include selected miRNA targets. Mol Cell 15:173–183 Stevens A (1978) An exoribonuclease from Saccharomyces cerevisiae: effect of modifications of 50 end groups on the hydrolysis of substrates to 50 mononucleotides. Biochem Biophys Res Commun 81:656–661 Stevens A (1980) Purification and characterization of a Saccharomyces cerevisiae exoribonuclease which yields 50 -mononucleotides by a 50 ->30 mode of hydrolysis. J Biol Chem 255:3080–3085 Stevens A, Poole TL (1995) 50 -exonuclease-2 of Saccharomyces cerevisiae. Purification and features of ribonuclease activity with comparison to 50 -exonuclease-1. J Biol Chem 270:16063–16069 Stevens A, Hsu CL, Isham KR, Larimer FW (1991) Fragments of the internal transcribed spacer 1 of pre-rRNA accumulate in Saccharomyces cerevisiae lacking 50 -30 exoribonuclease 1. J Bacteriol 173:7024–7028 Sugano S, Shobuike T, Takeda T, Sugino A, Ikeda H (1994) Molecular analysis of the dhp1+ gene of Schizosaccharomyces pombe: an essential gene that has homology to the DST2 and RAT1 genes of Saccharomyces cerevisiae. Mol Gen Genet 243:1–8 Till DD, Linz B, Seago JE, Elgar SJ, Marujo PE, Elias ML, Arraiano CM, McClellan JA, McCarthy JE, Newbury SF (1998) Identification and developmental expression of a 50 -30 exoribonuclease from Drosophila melanogaster. Mech Dev 79:51–55 Tishkoff DX, Johnson AW, Kolodner RD (1991) Molecular and genetic analysis of the gene encoding the Saccharomyces cerevisiae strand exchange protein Sep1. Mol Cell Biol 11: 2593–2608 Tishkoff DX, Rockmill B, Roeder GS, Kolodner RD (1995) The sep1 mutant of Saccharomyces cerevisiae arrests in pachytene and is deficient in meiotic recombination. Genetics 139: 495–509 Villa T, Ceradini F, Presutti C, Bozzoni I (1998) Processing of the intron-encoded U18 small nucleolar RNA in the yeast Saccharomyces cerevisiae relies on both exo- and endonucleolytic activities. Mol Cell Biol 18:3376–3383 Wahle E, Ruegsegger U (1999) 30 -end processing of pre-mRNA in eukaryotes. FEMS Microbiol Rev 23:277–295 West S, Proudfoot NJ (2008) Human Pcf11 enhances degradation of RNA polymerase IIassociated nascent RNA and transcriptional termination. Nucleic Acids Res 36:905–914 West S, Gromak N, Proudfoot NJ (2004) Human 50 ->30 exonuclease Xrn2 promotes transcription termination at co-transcriptional cleavage sites. Nature 432:522–525 Xiang S, Cooper-Morgan A, Jiao X, Kiledjian M, Manley JL, Tong L (2009) Structure and function of the 50 ->30 exoribonuclease Rat1 and its activating partner Rai1. Nature 458: 784–788 Xue Y, Bai X, Lee I, Kallstrom G, Ho J, Brown J, Stevens A, Johnson AW (2000) Saccharomyces cerevisiae RAI1 (YGL246c) is homologous to human DOM3Z and encodes a protein that binds the nuclear exoribonuclease Rat1p. Mol Cell Biol 20:4006–4015 Yang W, Lee JY, Nowotny M (2006) Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity. Mol Cell 22:5–13 Yang X-C, Sullivan KD, Marzluff WF, Dominski Z (2009) Studies of the 50 exonuclease and endonuclease activities of CPSF-73 in histone pre-mRNA processing. Mol Cell Biol 29:31–42

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Zabolotskaya MV, Grima DP, Lin M-D, Chou T-B, Newbury SF (2008) The 50 -30 exoribonuclease Pacman is required for normal male fertility and is dynamically localized in cytoplasmic particles in Drosophila testis cells. Biochem J 416:327–335 Zakrzewska-Placzek M, Souret FF, Sobczyk GJ, Green PJ, Kufel J (2010) Arabidopsis thaliana XRN2 is required for primary cleavage in the pre-ribosomal RNA. Nucleic Acids Res 38:4487–4502 Zhang M, Yu L, Xin Y, Hu P, Fu Q, Yu C, Zhao S (1999) Cloning and mapping of the XRN2 gene to human chromosome 20p11.1-p11.2. Genomics 59:252–254 Zhang K, Dion N, Fuchs B, Damron T, Gitelis S, Irwin R, O’Connor M, Schwartz H, Scully SP, Rock MG et al (2002) The human homolog of yeast SEP1 is a novel candidate tumor suppressor gene in osteogenic sarcoma. Gene 298:121–127 Zhang Z, Fu J, Gilmour DS (2005) CTD-dependent dismantling of the RNA polymerase II elongation complex by the pre-mRNA 30 -end processing factor, Pcf11. Genes Dev 19: 1572–1580 Zhao J, Hyman L, Moore CL (1999) Formation of mRNA 30 ends in eukaryotes: mechanism, regulation, and interrelationships with other steps in mRNA synthesis. Microbiol Mol Biol Rev 63:405–445 Zuo Y, Deutscher MP (2001) Exoribonuclease superfamilies: structural analysis and phylogenetic distribution. Nucleic Acids Res 29:1017–1026

Chapter 8

Structure and Degradation Mechanisms of 30 to 50 Exoribonucleases Rute G. Matos, Vaˆnia Pobre, Filipa P. Reis, Michal Malecki, Jose´ M. Andrade, and Cecı´lia M. Arraiano

Contents 8.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 PDX Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.1 PNPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.2 RNase PH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.3 The Exosome and the PDX Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 RNB Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3.1 RNase II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3.2 RNase R . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3.3 Rrp44 (Dis3) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4 DEDD Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4.1 Oligoribonuclease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4.2 RNase D . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4.3 RNase T . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4.4 Deadenylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

194 195 195 199 199 200 202 204 207 209 210 211 213 213 214 215

Abstract Exoribonucleases are enzymes that cleave RNA molecules by removing terminal nucleotides from the 30 or 50 end of the RNA molecules. They are key factors in RNA metabolism and have a relevant role in the processing and degradation of all types of RNAs. The 30 to 50 exoribonucleases are divided into families, according to their sequence and structural characteristics. The PDX family contains phosphate-dependent degradative enzymes, which can also perform the synthesis of RNA tails when phosphate is limiting. The RNB family contains hydrolytic enzymes with a similar domain organization. All proteins from this widespread family R.G. Matos • V. Pobre • F.P. Reis • M. Malecki • J.M. Andrade • C.M. Arraiano (*) Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Apartado 127, 2781–901 Oeiras, Portugal e-mail: [email protected]; [email protected]; [email protected]; [email protected]; [email protected]; [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_8, # Springer-Verlag Berlin Heidelberg 2011

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present the characteristic RNB domain responsible for the 30 to 50 exoribonuclease activity. In eukaryotes they can act alone or in a complex, the exosome, where they are the only active component. Finally, the DEDD family includes both RNA and DNA exonucleases and they present a similar mechanism of action. In this chapter, we will summarize the available information regarding the 30 to 50 exoribonucleases and discuss their importance for the RNA metabolism.

8.1

Introduction

The RNA levels in the cell depend on the efficiency of the transcription, and the rate of degradation. Although transcription is important to determine RNA steady state levels, the processing and degradation of RNA are also key factors in the regulation of gene expression. Ribonucleases (RNases) are the enzymes that are able to process and degrade RNA. Moreover, they have a critical role in the maturation of ribosomal and transfer RNAs (Re´gnier and Arraiano 2000; Arraiano et al. 2010a). They are also involved in the quality control of all types of RNAs, allowing the recycling of the ribonucleotides in the cell (Li et al. 2002; Silva et al. 2011). RNases are present in all domains of life, and play a central role in the control of gene expression by determining the levels of functional RNAs in the cell (Re´gnier and Arraiano 2000; Arraiano and Maquat 2003; Parker and Song 2004). Many of the RNases in the cell are essential and others have overlapping functions (Re´gnier and Arraiano 2000). RNases can act alone or they can be part of RNA degradation complexes, namely, the degradosome and the exosome. Ribonucleases can be divided into endoribonucleases (which cleave the RNA molecules internally) and exoribonucleases (which degrade the RNA by removing terminal nucleotides from the 30 end or the 50 end of the RNA molecules). In this chapter, we will focus on exoribonucleases, namely, those which degrade the RNA from the 30 to the 50 end. Exoribonucleases can act hydrolytically, releasing nucleotide monophosphates, or phosphorolytically, if they use inorganic phosphate to cleave the molecules releasing nucleotide diphosphates (Zuo and Deutscher 2001). The nucleotides released after the action of exoribonucleases are very important for turnover since they can be reutilized for the synthesis of new RNA molecules. Exoribonucleases are involved in many RNA metabolic events, namely, in RNA maturation and degradation (Andrade et al. 2009b). According to their sequence and structural characteristics, exoribonucleases can be divided into five families: RNB, PDX, DEDD, RRP4, and 5PX (the last two families do not have any representatives in bacteria) (Table 8.1). In this chapter we will summarize the available information about all families of exoribonucleases that degrade RNA from the 30 to the 50 end, and discuss the latest findings and their relevance in RNA metabolism.

8

Structure and Degradation Mechanisms of 30 to 50 Exoribonucleases

Table 8.1 Family of 30 to 50 Exoribonucleases Eukaryotic Catalytic Family E. coli members members mechanism RNase II RNB

RNase R RNase D RNase T Oligoribonuclease

DEDD –

PNPase

PDX

8.2

RNase PH

195

Comments These enzymes are processive in the 30 to 50 direction Distributed in all domains of life

Rrp44 (Dis3) Tazman Rrp6 – Ynt20 (Rex2) Pan2, ERI-1 Rex1, 3, 4

Hydrolytic

– Rrp41–43, 45, 46 Mtr3 Csl4

The activity is distributive in the 30 to 50 direction and Phosphorolytic phosphate dependent

Hydrolytic

Proteins from this family are distributive in the 30 to 50 direction Some have DNase activity The activity is processive in the 30 to 50 direction and phosphate dependent

PDX Family

The PDX family of 30 -50 exoribonucleases includes PNPase, RNase PH from bacteria, and the core of the exosome in archaea and eukaryotes (Mian 1997; Zuo and Deutscher 2001; Pruijn 2005) (Fig. 8.1). These enzymes, contrary to the other exoribonucleases, are phosphate-dependent enzymes, and release a dinucleotide as an end product of degradation. Beside their role as exoribonucleases, the enzymes from this family can also catalyze other reactions like the addition of heteropolymeric tails to RNA substrates (Slomovic et al. 2008). In fact, the polymerization activity of PNPase was essential for the deciphering of the genetic code and this discovery led to the award of a Nobel Prize to Severo Ochoa in 1959 (GrunbergManago et al. 1955).

8.2.1

PNPase

PNPase is a multifunctional protein. Its main activity is the processive 30 -50 phosphorolytic degradation of single-stranded RNA with a minimal 30 overhang of 7–10 unpaired ribonucleotides (Spickler and Mackie 2000). However, under conditions of low inorganic phosphate concentration, PNPase catalyzes the inverse reaction, that is, polymerization of single-stranded RNA from nucleoside diphosphates (Littauer and Soreq 1982). Contrary to the homopolymeric poly(A) tails added by PAP I, the tails synthesized by PNPase are heteropolymeric, containing all four nucleotides (Slomovic et al. 2008). In spinach chloroplasts, cyanobacteria

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Fig. 8.1 The PDX-family of enzymes. (a) Linear representation of E. coli PNPase and RNase PH domains (note that the figures are not in scale). (b) View of PNPase crystal structure in the monomer organization (PDB ID 3CDI – this PDB does not include the KH and S1 domains) (Shi et al. 2008). (c) Comparison of PNPase trimer structure (PDB ID 3GCM – this PDB does not include the KH and S1 domains) (Nurmohamed et al. 2009) with the archaeal exosome ring from Sulfolobus solfataricus (Rrp41 is represented in light blue, Rrp42 in orange and Rrp4 in green; PDB ID 3L7Z) (Lu et al. 2010) and with the human exosome ring (Rrp4 is represented in orange, Rrp40 in red, Rrp41 in grey, Rrp42 in yellow, Rrp43 in light blue, Rrp45 in green, Rrp46 in magenta, Mtr3 in brown, and Csl4 in blue; PDB ID 2NN6) (Liu et al. 2006)

and Gram-positive bacteria, PNPase is suggested to be the main polyadenylating enzyme (Yehudai-Resheff et al. 2001; Rott et al. 2003; Sohlberg et al. 2003; Campos-Guilleń et al. 2005). The archaeal exosome, which is very similar to PNPase (see below) has also been demonstrated to be responsible for the addition of heteropolymeric tails in Sulfolobus (Portnoy et al. 2005). PNPase also catalyzes the exchange reaction between the b-phosphate group of nucleoside diphosphates and free orthophosphate. More recently, it was described that PNPase also regulates RNA translocation into mitochondria (Wang et al. 2010a). PNPase can form complexes with other enzymes to easily degrade RNA. The main complex in which PNPase is involved is the degradosome. The degradosome is a large, multiprotein complex involved in RNA degradation. In Escherichia coli, this multiprotein complex is composed of the endoribonuclease RNase E, the 30 -50 exoribonuclease PNPase, the DEAD-box RNA helicase B (RhlB), and the glycolytic enzyme enolase (Carpousis et al. 1994; Miczak et al. 1996; Py et al. 1996; Vanzo et al. 1998; Iost and Dreyfus 2006). RNase E provides the scaffold for

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the degradosome. Recent findings showed that E. coli PNPase and RNase E are present in the degradosome in an equimolar ratio (Nurmohamed et al. 2009). However, PNPase content can change in response to phosphosugar stress, temperature shock, and growth stage (Beran and Simons 2001). In other organisms, the degradosome content might be different. In Pseudomonas syringiae PNPase is substituted by RNase R, and the DEAD-box helicase present in the degradosome is RhlE (Purusharth et al. 2005). PNPase can also exist as a a3b2 complex where the b subunit has been identified as the helicase, RhlB (Lin and Lin-Chao 2005) and can form complexes with the host factor Hfq and PAP I (Mohanty et al. 2004). X-ray crystal structures of E. coli and Streptomyces antibioticus PNPase reveal a homotrimeric subunit organization with a ring-like architecture (Fig. 8.1) (Symmons et al. 2000; Shi et al. 2008; Nurmohamed et al. 2009). Each monomer exhibits a five-domain arrangement: at the N-terminus two RNase PH domains (PH1 and PH2) are linked by an a-helical domain, and at the C-terminal end there are two RNA-binding domains (KH and S1) (Fig. 8.1a, b). The three monomers associate via trimerization interfaces of the core domains, forming a central channel, where catalysis occurs (Fig. 8.1c). Structural and mutational analysis of E. coli and S. antibioticus PNPase and recent work on the spinach chloroplast and human PNPase (Sarkar et al. 2005) demonstrated that the catalytic site of PNPase is composed of structural elements of the first and second core domains (PH1 and PH2) (Symmons et al. 2000; Briani et al. 2007; Shi et al. 2008; Nurmohamed et al. 2009). The all-a-helical domain was shown to be important for the catalytic activity of E. coli PNPase (Briani et al. 2007). This domain is highly dynamic and may affect the access of nucleoside diphosphates and phosphate to the active site. PNPase catalytic activity is dependent on a metal ion coordinated in the active site by the conserved residues D486, D492, and K494 (Nurmohamed et al. 2009). In S. antibioticus PNPase, the metal used to identify the catalytic center was tungstate (a phosphate analogue), which is coordinated to the T462 and S463 side chains (Symmons et al. 2000). On the other hand, E. coli PNPase crystals were obtained in the presence of Mn2+, since this ion can substitute Mg2+ to support catalysis and is more easily identified in the crystal structure. Interestingly, several mutations in the core region did not affect phosphorolytic or polymerase activities, but affected RNA binding. One example is the substitution of the conserved residue Gly454 by an aspartate in E. coli PNPase (Regonesi et al. 2006). PNPase lacking either the S1 or KH domains retained phosphorolytic activity and still has some RNA-binding capacity, but the truncated enzymes are much less active. Although PNPase lacking the S1 domain, KH domain, or both domains could still be assembled in the degradosome and their presence in the degradosome is vital at low temperature, the domains were shown to be essential to support growth in the cold. Nevertheless, the presence of both KH and S1 domains are required for proper RNA binding (Goverde et al. 1998; Garcıá-Mena et al. 1999; Zangrossi et al. 2000; Briani et al. 2007; Matus-Ortega et al. 2007; Shi et al. 2008). Shi and coauthors demonstrated that these RNA-binding domains also have a major role in the formation of a more stable trimeric structure, and are essential for the

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constriction of the central channel (Shi et al. 2008). This is in agreement with a previous study where PNPase S1 domain was able to induce trimerization of an RNase II-PNPase chimeric protein (Amblar et al. 2007). Two constricted necks have been identified in the central channel (Symmons et al. 2000; Shi et al. 2008). Three arginine residues were identified in the PNPase neck region. The Arg102 and Arg103 residues are in the upper neck region close to the channel entrance and are involved in RNA binding. The third arginine residue (Arg106) is located in the lower neck region, closer to the active site, and apparently is involved in the processive RNA degradation (Shi et al. 2008). The upper neck and the crystal structure of the PNPase complexed with RNA support the hypothesis that the pathway followed by the RNA molecule is along the central pore in the direction to the active site (Symmons et al. 2000; Shi et al. 2008; Nurmohamed et al. 2009). The dynamic aperture of the central channel and the ability of its neighboring regions to undergo conformational changes probably are the key aspects that allow PNPase to constrain and translocate the substrates in a processive mode of action (Nurmohamed et al. 2009). PNPase is encoded by the pnp gene that is located downstream of the rpsO gene (encoding ribosomal protein S15) and is transcribed from two promoters (one upstream of the rpsO gene and another upstream of pnp gene). pnp expression is negatively autoregulated at the posttranscriptional level. This autoregulation only occurs after an initial cleavage by RNase III at the 50 end of the pnp message (Portier et al. 1987; Robert-Le Meur and Portier 1992; Jarrige et al. 2001; Carzaniga et al. 2009). PNPase levels are also affected by polyadenylation but not by Poly(A) polymerase (PAP I) itself (Jarrige et al. 2001). PNPase and RNase II are interregulated. In the absence of RNase II, PNPase levels are increased and PNPase overexpression leads to a decrease in RNase II activity (Zilhaõ et al. 1996a, b). More recently, it was shown that guanosine 50 -diphosphate 30 -diphosphate (ppGpp) inhibits Nonomuraea sp. and Streptomyces PNPase phosphorolytic and polymerization activities (Gatewood and Jones 2010; Siculella et al. 2010). In conclusion, PNPase has a complex regulation and its expression is finely controlled both at transcriptional and posttranscriptional levels (Andrade et al. 2009a). PNPase does not seem to be indispensable to E. coli at optimal temperature, unless either RNase II or RNase R are also missing (Donovan and Kushner 1986; Cheng et al. 1998). However, PNPase is essential for E. coli growth at low temperatures (Luttinger et al. 1996; Piazza et al. 1996; Zangrossi et al. 2000). It was shown that overexpression of RNase II could complement cold shock function of PNPase (Awano et al. 2008). PNPase has been implicated in the establishment of virulence in several pathogens, namely, in Salmonella, Dichelobacter nodosus, Dickeya dadantii, Yersinia, Campylobacter jejuni, and Streptococcus pyogenes (Arraiano et al. 2010a). However, PNPase role in virulence can be contradictory; while in some pathogens PNPase seems to act as a virulence repressor, in others PNPase is important for the establishment of virulence (Arraiano et al. 2010a).

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8.2.2

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RNase PH

RNase PH is encoded by the rph gene and is co-transcribed with pyrE, a gene necessary for pyrimidine synthesis, which is located upstream of rph (Ost and Deutscher 1991). RNase PH is not essential for cell growth, unless RNase T or PNPase are also missing. In fact, the rph mutation only results in inviability in a strain already lacking RNases I, II, D, BN, and T (Kelly et al. 1992). Contrary to PNPase, which is mainly involved in RNA degradation, RNase PH is involved in tRNA metabolism, namely, in the processing of tRNA precursors (Deutscher et al. 1988; Kelly and Deutscher 1992a), and in ribosome metabolism (Zhou and Deutscher 1997; Redko and Condon 2010). RNase PH can act as a phosphorolytic ribonuclease (removing nucleotides following the CCA terminus of tRNA) or as a nucleotidyltransferase (adding nucleotides to the ends of RNA molecules) (Kelly and Deutscher 1992a; Wen et al. 2005; Bralley et al. 2006). RNase PH can also modify the 30 end of other small RNAs, including M1, 6S, and 4.5S RNA (Li et al. 1998). The crystal structures of RNase PH from Aquifex aeolicus, Bacillus subtilis and P. aeruginosa have been determined. All the three proteins crystallized as a hexamer arranged as a trimer of dimers. The overall architecture of the three RNase PH is a baba fold, but the number of b-strands and a-helices are different between them (Ishii et al. 2003; Choi et al. 2004; Harlow et al. 2004). In the A. aeolicus RNase PH, the phosphate-binding site consists of four residues, and is located at the bottom of a deep cleft, and it was proposed that the narrow entrance of this cleft can discriminate between single- and double-strand RNA. Mutations of the conserved residues Arg86, Thr125, and Arg126 showed that these residues are very important for the phosphorolytic activity of the RNase PH. Based on structural and mutational analysis of the A. aeolicus RNase PH, Ishii and coworkers proposed that the RNase PH dimer only interacts with the tRNA acceptor stem, while the other parts of the tRNA remain unbound (Ishii et al. 2003). On the other hand, in the crystal structure of the B. subtilis RNase PH it was possible to identify a tRNA phosphate backbone-binding region. The active binding site (Thr125, Arg126 and Gly123) for the B. subtilis RNase PH corresponds to the binding of the phosphate ion in the A. aeolicus RNase PH structure. Harlow and coworkers also identified three conserved arginine residues (Arg68, Arg73, and Arg76) as very important residues for the maintenance of the RNase PH hexameric structure (Harlow et al. 2004). The formation of the RNase PH hexameric ring is essential for the binding of precursor tRNA and also for the exoribonucleolytic activity (Choi et al. 2004).

8.2.3

The Exosome and the PDX Family

The exosome is a multiprotein complex with a 30 -50 RNase activity that is involved in RNA degradation and processing. In Archaea, the exosome consists of two

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RNase PH subunits (Rrp41 and Rrp42) and two RNA-binding subunits (Rrp4 and Csl4) (Evguenieva-Hackenberg et al. 2003; Buttner et al. 2005). The crystal structures of the archaeal exosomes from the Sulfolobus solfataricus, Archaeoglobus fulgidus, and Pyrococcus abyssi have been solved. These structures revealed a hexameric ring-like assembly formed by three Rrp41 and Rrp42 subunits. This hexameric ring is capped by a trimer of the RNA-binding proteins Rrp4 and/or Csl4 (Buttner et al. 2005; Lorentzen et al. 2005) (Fig. 8.1c). Both Rrp41 and Rrp42 are involved in substrate binding; however, only the Rrp41 has catalytic activity (Lorentzen et al. 2005). The archaeal exosome concomitantly binds three singlestranded RNA that enter the exosome catalytic site from the top side of the RNase PH ring. The narrow constriction of the central channel might be the reason why the exosome can degrade single-stranded RNA but stalls with the secondary structures (Bonneau et al. 2009). In addition to trapping and directing the substrate to the catalytic site, the exosome central channel also appears to have an important role in the RNA stabilization and processing. A model for the RNA processing by the archaeal exosome has been proposed, which is very similar to the one proposed for RNase II (Frazaõ et al. 2006). In this model, the single-stranded RNA binds to the RNA-binding subunits and is then threaded to the catalytic site. After phosphorolytic cleavage, the nucleoside diphosphate undergoes a structural arrangement and is released through a conserved side channel. At the same time, there is the entrance of a new inorganic phosphate. The final 4–5 nucleotide products are no longer capable of maintaining the interactions between the RNA and the RNA recognition cleft (formed at the interface of the Rrp41-Rrp42 subunits), so the translocation is no longer possible and the substrate is released. The eukaryotic exosome is also formed by a six-subunit PH-domains ring, Rrp41, Rrp45, Rrp46, Rrp42, Rrp43, and Mtr3. However, and contrary to what happens in archaeal organisms, these proteins appear to lack catalytic activity. In fact, both yeast and human exosome cores do not present any catalytic activity (Liu et al. 2006; Dziembowski et al. 2007). In this case, the RNase PH homologues in Eukaryotes may have a role in substrate binding and recruitment.

8.3

RNB Family

The RNB family of enzymes is present in all domains of life and exhibit the same modular organization (Fig. 8.2a). They processively degrade RNA in the 30 to the 50 direction. They have a hydrolytic activity, releasing 50 -nucleotide monophosphates (Mian 1997; Zuo and Deutscher 2001). E. coli RNase II is the prototype of this family of enzymes, which also comprises RNase R and the eukaryotic Rrp44/Dis3 (Table 8.1). Members of this family play very important functions in the cell: they are essential for growth (Mitchell et al. 1997), they can be developmentally regulated (Cairraõ et al. 2005), and mutations in its gene have been linked with abnormal

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Fig. 8.2 The RNase II-family of enzymes. (a) Schematic representation of RNB-family members, which share a similar modular organization (note that the figures are not in scale). (b) RNase II 3D structure shows that it is comprised of two N-terminal Cold Shock Domains (CSD1 in orange and CSD 2 in red), a central RNB domain (in blue) and a C-terminal S1 domain (in green); the RNA molecule is also represented (in pink) (PDB ID 2IX0 and 2IX1) (Frazaõ et al. 2006). (c) In the catalytic cavity, several conserved residues interact with the RNA molecule (Frazaõ et al. 2006) Residues from E. coli RNase II are written in black, from E. coli RNase R in brown, and yeast Rrp44/Dis3 in dark blue. (d) The Rrp44 protein is the only active subunit in the yeast exosome and has both exo- and endonucleolytic activity (Dziembowski et al. 2007; Schaeffer et al. 2009). This protein can act as a member of the exosome and it is also possible that it may act alone. (e) Comparing the catalysis of E. coli RNase II and RNase R with different substrates. RNase II releases a 4 nt fragment as end product when degrading single-stranded substrates. For structured substrates, the product released by RNase II has an overhang of 4–7 nt before the duplex. RNase R is able to degrade both single- and double-stranded substrates releasing a 2 nt fragment as end product

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chloroplast biogenesis (Bollenbach et al. 2005), mitotic control, and cancer (Lim et al. 1997). They were also shown to be important for stress responses, RNA and protein quality control, and required for virulence in several organisms (Cheng et al. 1998; Cairraõ et al. 2003; Cheng and Deutscher 2003, 2005; Andrade et al. 2006; Cairraõ and Arraiano 2006; Purusharth et al. 2007; Charpentier et al. 2008; Erova et al. 2008). In eukaryotes, RNase II homologues can act independently or can be associated in multiprotein complexes, like the exosome, a complex of exoribonucleases crucial for the maintenance of the correct levels of RNAs in the cell (van Hoof and Parker 1999).

8.3.1

RNase II

E. coli RNase II is the prototype of this family of enzymes. This 72 kDa protein encoded by gene rnb is the major hydrolytic enzyme, responsible for 90% of the exoribonucleolytic activity in crude extracts (Deutscher and Reuven 1991). RNase II is expressed from two different promoters, P1 and P2, which implies a differential regulation of the rnb gene at the level of transcription (Zilhaõ et al. 1996b). This protein is also regulated at posttranscriptional levels (Cairraõ et al. 2001). Other ribonucleases, such as RNase III, RNase E, and PNPase, are involved in the modulation of RNase II levels (Zilhaõ et al. 1995; Zilhaõ et al. 1996a), and it was also shown that there is a posttranslational regulation of RNase II levels conferred by the gmr gene (Gene Modulating RNase II), which is located downstream of the rnb gene (Cairraõ et al. 2001). RNase II is a hydrolytic enzyme which processively degrades RNA in the 30 to 50 direction releasing 50 monophosphates, and the final product released is a 4 nt fragment. The activity of this protein is sequence independent but sensitive to secondary structures (RNase II stalls around seven nucleotides before it reaches the double-stranded region) (Fig. 8.2e) (Spickler and Mackie 2000). Ten nucleotides is the minimum of length of the RNA molecule needed for the processivity of the enzyme. For fragments less than 10 nt, the activity of RNase II becomes distributive (Cannistraro and Kennell 1994). Although being specific for RNA, RNase II is able to bind to DNA molecules without being able to cleave them. It seems that the DNA oligonucleotides can act as inhibitors of RNase II action since they bind to its specific binding site (Cannistraro and Kennell 1994). RNase II activity does not depend on the RNA sequence; however, it has a marked preference for poly(A) substrates. Polyadenylation is responsible for the control of mRNA stability in several organisms. The poly(A) tails are synthesized by poly(A) polymerase I (PAP I) to target RNAs to be degraded by exoribonucleases. Since RNase II has preference for poly(A) substrates, it will rapidly degrade these tails. The degradation process proceeds until a secondary structure, such as a Rhoindependent terminator is found. By rapidly degrading these Poly(A) tails, RNase II can impede the binding of other exoribonucleases to the RNA molecule, since

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no overhang is left to allow the binding of other ribonucleases, thus preventing the RNA degradation. With this behavior, RNase II is acting paradoxically by protecting some messages (Marujo et al. 2000; Andrade et al. 2009b). By sequence analysis, it was proposed that RNase II has a modular organization, with three conserved domains: a CSD at the N-terminal region, a central catalytic RNB domain, and a C-terminal S1 domain (Andrade et al. 2009b; Arraiano et al. 2010a, 2010b) (Fig. 8.2a). The RNB domain is well conserved and it is exclusive to RNase II-like proteins. It contains four highly conserved motifs (I to VI) with several invariant amino acids (Mian 1997). The function of each domain was determined and it was shown that the RNB domain is responsible for the catalytic activity, whereas CSD and S1 domains are responsible for the binding to RNA (Amblar et al. 2006). The resolution of the crystal structure of RNase II (the first of an RNB-family member) revealed the existence of four domains instead of the three previously proposed: two N-terminal CSD, the central RNB domain, and the S1 domain at the C-terminal region (Frazaõ et al. 2006). Moreover, it was possible to see that the RNB domain presents an unprecedented fold which is characteristic of this family. The RNA-binding domains (CSD1, CSD2, and S1) are grouped together on one side of the structure, while the active site is on the other side of the molecule (Frazaõ et al. 2006) (Fig. 8.2b). An RNase II mutant found in nature (D209N) was previously described and it was showed to encode an inactive protein still able to bind to RNA (Amblar and Arraiano 2005). The structure of this mutant was also solved, and the crystallization proceeded with a RNA molecule that was inside. This allowed the co-crystallization of the RNA molecule inside the protein, which was very important since it enabled the visualization of RNA-protein contacts, and explains the mode of action (Frazaõ et al. 2006). The RNA contacts RNase II at two different and noncontiguous regions, which act synergistically to provide a processive degradation: the anchoring region, constituted by the three RNA-binding domains, and the catalytic region, which is buried inside the catalytic domain and is flanked by the four RNase II conserved motifs (Frazaõ et al. 2006). The shortest RNA substrate that is able to contact both anchor and catalytic regions is a 10 nt fragment. In fact, this is the minimum size necessary for the enzyme to be processive. For shorter RNA fragments, fewer contacts with the protein are established and the enzyme becomes distributive (Frazaõ et al. 2006). The access to the catalytic pocket is restricted to ssRNA due to the steric hindrance at its entrance and explains why RNase II is not able to cleave double-stranded substrates (Spickler and Mackie 2000; Frazaõ et al. 2006). The first five nucleotides counting from the 30 end are stacked between the two aromatic residues Tyr253 e Phe358 (Fig. 8.2c), which helped us to understand why 4 nt is the final end product released by RNase II: when the RNA molecule is shorter than five nucleotides, the stacking no longer occurs, and the molecule is released (Frazaõ et al. 2006). A mutational analysis identified residue Tyr253 as responsible for setting the final end product in RNase II. The substitution of this residue by an alanine changed the RNase II product from 4 to 10 nt and it was also shown to be crucial for the RNA binding at the 30 -end (Barbas et al. 2008; Arraiano et al. 2010b; Matos et al. 2010). The active site of RNase II has four highly

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conserved aspartic acids in positions 201, 207, 209, and 210 and an arginine in position 500 (Fig. 8.2c), which were postulated to assist in catalysis, namely, by fixing the RNA molecule correctly. It was shown that the four aspartates are not equivalent in their function, with Asp209 being the only one essential for catalysis (Barbas et al. 2008; Arraiano et al. 2010b). The same result was obtained with other members of this family (Dziembowski et al. 2007; Matos et al. 2009). The Arg500 was also proved to have a crucial role in RNA degradation (Barbas et al. 2009; Arraiano et al. 2010b). The residue Glu542 is in close proximity with the leaving nucleotide (Fig. 8.2c), and it was proposed that it helped to release the last nucleotide after cleavage. However, its substitution in E. coli RNase II by an alanine led to a mutant, which is 110-fold more active and has a 20-fold higher RNA affinity when compared to the wild type protein (Barbas et al. 2009; Matos et al. 2010). This mutant protein constructed was described as an RNase II “Super-Enzyme.” This result showed that, in fact, Glu542 slows down the activity of the protein (Barbas et al. 2009; Arraiano et al. 2010b). As mentioned previously, RNase II is able to bind to DNA but it cannot cleave it. The resolution of RNase II structure allowed to see the interaction between Tyr313 and Asp201 and the O20 ribose oxygen of the second ribose and Glu390 with the O20 ribose oxygen of the fourth ribose (Frazaõ et al. 2006) (Fig. 8.2c). It was also demonstrated that RNase II has a strict requirement for a ribose at the second and/or the fourth nucleotide from the 30 -end of the molecule (Barbas et al. 2009; Arraiano et al. 2010b). Moreover, these contacts are responsible for the RNA specificity (Barbas et al. 2009).

8.3.2

RNase R

RNase R-like enzymes have a wide impact in RNA metabolism in many different organisms. RNase II stalls when reaching near a duplex, and PNPase needs association with a helicase to overcome such structures. However, RNase R has the remarkable feature of degrading double-stranded RNAs. The substrate list for RNase R includes defective tRNAs (Li et al. 2002) or rRNA (Cheng and Deutscher 2003) as well as mRNAs with REP-containing sequences (Cheng and Deutscher 2005) or small RNAs like the stable SsrA/tmRNA (Cairraõ et al. 2003). Degradation of a RNA duplex occurs provided there is a single-stranded 30 overhang of at least 7-nts (Vincent and Deutscher 2006) and RNase R was shown to be a key enzyme involved in the degradation of polyadenylated RNA (Andrade et al. 2009a). The activity of RNase R is modulated according to environmental stimuli and its protein levels are upregulated under several stresses, namely, stationary-phase of growth and in cold-shock (Andrade et al. 2006; Cairraõ and Arraiano 2006). RNase R was shown to be involved in the degradation of the ompA transcript specifically in the stationary phase of growth, while no effect is detected in exponentially growing cells (Andrade et al. 2006). E. coli RNase R deficient colonies are smaller especially in the cold (Cairraõ et al. 2003) and this enzyme is even essential for survival at low temperatures in pathogenic strains like P. syringae (Purusharth et al.

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2005) or Aeromonas hydrophila (Erova et al. 2008). In addition to its role in stress response, RNase R has also been implicated in virulence mechanisms (Tobe et al. 1992; Lalonde et al. 2007; Erova et al. 2008; Tsao et al. 2009). The stress-induction of RNase R levels can in fact be highly advantageous, enabling pathogens to adapt to environmental challenges imposed prior and during the infection process. Accordingly, most RNase R-deficient bacteria have been shown to be less virulent than the wild-type strains. How this is achieved is still unclear but it has been suggested that RNase R may control the export of proteins involved in virulence mechanisms (Tobe et al. 1992). A common trait of pathogenic RNase R mutants seems to be impaired motility (Erova et al. 2008; Tsao et al. 2009). RNase R also affects other cellular processes, like the development of competence in Legionella pneumophila (Charpentier et al. 2008) or the expression of apoptosis genes in Helicobacter pylori (Tsao et al. 2009). This is probably related to critical RNA degradation pathways mediated by RNase R. Currently, the structure of RNase R remains unknown, which represents a major drawback in understanding RNase R mechanism of action. Most of the knowledge on RNase R structure is inferred from the available structures of its close counterparts, RNase II and Rrp44. RNase R follows the typical modular organization found on RNase II-family members: a RNB catalytic domain flanked by RNAbinding domains (CSD1 and CSD2 located at the N-terminus and a C-terminal S1 domain). Furthermore, additional features are exclusively found in the RNase R sequence, namely, a predicted nucleic acid binding motif (Helix-turn-Helix) at the N-terminal and a highly basic extended region after the S1 domain (Fig. 8.2a). All these regions must combine in a way that makes RNase R unique amongst exoribonucleases. The majority of the residues interacting with the 30 end of the RNA are conserved throughout the RNase II-family of exoribonucleases, suggesting a similar mechanism for hydrolysis (Barbas et al. 2008; Bonneau et al. 2009). Structural differences might help explaining the divergence in substrate recognition between the members of RNase II-family. Despite the biochemical similarities many uncertainties remain concerning the pathway followed to effect the degradation of structured substrates. A three-dimensional model of RNase R has been proposed based on the structure of its paralogue RNase II (Barbas et al. 2008). Mutational analysis identified the highly conserved acid residues located in the active center responsible for catalysis: D272, D278, and D280 (Matos et al. 2009; Awano et al. 2010). As in the other members of the RNase II-family, these highly conserved Aspartates are involved in coordination of divalent metal ions (preferably Mg2+) essential to catalysis. In particular, the RNase R D280N mutant showed no exonucleolytic activity although RNA binding was not affected, analogous to what was reported with the D209N mutant in RNase II (Amblar and Arraiano 2005; Matos et al. 2009; Awano et al. 2010). The highly conserved Tyrosine Y324 was found to be responsible for setting the final end-product of RNase R. Mutation Y342A altered the final end product from 2 to 5 nucleotides, probably due to loose packing of the 30 -terminal nucleotides in the catalytic cavity (Matos et al. 2009). Overall, the structural model of RNase R when compared to RNase II and Rrp44 structures identified the critical

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residues located in similar catalytic environments (Barbas et al. 2008). However, mutagenic studies revealed that despite all similarities, the catalytic channel is not alike between these enzymes. RNase R was shown to bind RNA more tightly within its catalytic channel than does RNase II. The amino acid residue Arg572 located in the nuclease domain channel strongly affects RNase R catalytic properties. Mutation to a Lysine (the equivalent residue found in RNase II) greatly impaired RNase R activity on structured RNAs (Vincent and Deutscher 2009a). Surprisingly, just the nuclease domain of RNase R (but not the one from RNase II) without accessory domains is able to degrade a double-stranded RNA (Matos et al. 2009; Vincent and Deutscher 2009b). A truncated form of RNase R expressing only the RNB domain degrades a blunt dsRNA (Fig. 8.3), although with a low level of activity when compared to wild-type protein. However, the presence of the auxiliary domains for substrate binding completely inhibits this activity, probably by “blocking” the entrance of dsRNA into the catalytic channel (Fig. 8.3). In the presence of CSD1, CSD2, and S1 domains, a short 30 unpaired overhang is required to allow the degradation of dsRNA (Fig. 8.3). Available data indicates that RNA-binding domains actually discriminate the substrates that can be targeted by RNase R, favoring the selection of RNA molecules tagged with a 30 linear tail (Matos et al. 2009). Remarkably, RNA-binding domains may have an intrinsic ability to help to unwind the double-stranded RNA molecules. As an additional probe of the resourceful enzyme that RNase R is, it has been suggested that it can function both as an exoribonuclease as well as an RNA “helicase” (Awano et al. 2010). RNase R intrinsic “helicase” unwinding activity is dependent on RNA-binding regions (namely, on CDS2). In vitro studies showed that exonuclease and “helicase” activities are distinct and independent being located in separate domains. Like CsdA and other DEAD-box RNA helicases, RNase R was shown to be more active at

Fig. 8.3 Degradation of double-stranded substrates by E. coli RNase R. RNase R needs a 30 single-stranded overhang to degrade double-stranded substrates. In the absence of the RNAbinding domains, the protein can degrade “tail-less” RNAs

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unwinding short (10-bp) rather than longer RNA duplexes. Not surprisingly, only a double-stranded RNA with a 30 linear overhang was shown to be a suitable substrate for RNase R helicase action. Moreover, the additional motifs exclusively found in RNase R may expand the known functions of the anchor domains (Zuo et al. 2006; Andrade et al. 2009b). The helix-turn-helix predicted in the N-terminus is a potential nucleic acid-binding motif although there are no experimental evidence yet of this role. The extended region found in the C-terminal end contains a positively charged surface patch formed by basic arginine and lysine residues. This suggests a possible role in nucleic acid-binding through electrostatic interactions. These potential extra protein–RNA interactions might contribute to “melting” of secondary RNA structures, and help to explain the impressive RNase R mode of action. However, this is quite speculative at the moment. In fact, it was proven that these “extra” domains in RNase R are not there merely for cosmetic purposes; the basic region in the C-terminal region was found to control the stability of RNase R through interactions with components of the trans-translation machinery. RNase R is an unstable protein but is highly stabilized if SmpB and/or tmRNA are prevented from interacting with the C-terminal region of RNase R (Liang and Deutscher 2010). It is still not clear how binding of SmpB and tmRNA to RNase R leads to its destabilization. Possible structural changes of the complex might help explaining these observations. More recently, it was also shown that the C-terminal lysine-rich region is necessary for the recruitment of RNase R to stalled ribosomes and to the selective decay of defective transcripts (Ge et al. 2010). It was also shown that the S1 domain from RNase R, namely, the C-terminal region, is involved in the degradation of structured substrates in an RNase II context, probably by helping to unwind the substrate (Matos et al. 2011). A definitive model for RNA degradation by RNase R is still open, and it seems clear that only the resolution of RNase R structure will answer the many questions about its remarkable mode of action.

8.3.3

Rrp44 (Dis3)

Rrp44, a member of the RNB family of enzymes, degrades RNA hydrolytically from 30 to 50 in a processive manner to a final product of 3 to 5 nucleotides. In addition to single-stranded RNA, Rrp44 is able to degrade secondary structures provided that it has a 30 end overhang with at least 4 nucleotides (Dziembowski et al. 2007; Bonneau et al. 2009). In addition to the domains described for RNase II and RNase R, Rrp44 contains in the N-terminus a region with three conserved cysteines (CR3 region) and a highly conserved pilT N-terminal (PIN) domain with endonucleolytic activity (Cairraõ et al. 2005; Lebreton et al. 2008; Schaeffer et al. 2009; Schneider et al. 2009). The two active sites responsible for both exo- and endonucleolytic activity coordinate to degrade and process exosome substrates.

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In Saccharomyces cerevisiae, the crystallographic structure of a single aminoacid mutant (D551N) of Rrp44DPIN in complex with RNA was determined (Bonneau et al. 2009). Although, while in the RNase II D209N mutant the RNA molecule was fortuitously co-crystallized with the protein, in this case, the RNA fragment was forced to be co-crystallized. This mutant contains a point mutation (D551N) within the active site that allows RNA binding but prevents the exoribo nucleolytic cleavage (Dziembowski et al. 2007; Schneider et al. 2007). This mutation is equivalent to D209N in RNase II which impairs the coordination of one of the two magnesium ion essential for catalysis (Amblar and Arraiano 2005; Frazaõ et al. 2006). Similar to RNase II, the amino-terminal region starts with a a-helix followed by the two consecutive cold-shock domains (CSD1 and 2). At the carboxy terminus, there is a third RNA-binding domain with a typical S1 RNA-binding fold. Between the two CSD and the S1 domain, the RNB catalytic domain is centered around a core that is reminiscent of RNase H and is surrounded by several a-helices. The superposition of Rrp44 and RNase II showed that 85% of the residues of the RNB domain and more than 70% of the three OB-fold domains superpose in their a-carbon positions. Besides the structural conservation in all domains, there is a difference concerning the relative orientation of the binding domains, with implications on the route of RNA access to the catalytic site. In Rrp44, the RNA is threaded to the catalytic site by binding the CSD1 and the RNB domains, while in RNase II, the RNA is threaded by binding S1 and CSD2 domains. It was suggested that these different routes are responsible for the difference in RNA-unwinding properties and activity on structured RNA substrates between RNase II and Rrp44 enzymes (Bonneau et al. 2009). The crystal structure of the Rrp44 PIN domain was elucidated in a ternary complex constituted by Rrp44 and two other exosome proteins (Rrp41 and Rrp45). The final model includes residues 36–231 of the Rrp44 PIN domain and residues 253–1001 of the Rrp44 RNase II-like region (Bonneau et al. 2009). This domain folds in a twisted parallel b sheet, surrounded by helices, with the catalytic site in the C-terminus ends of the b strands (Malet et al. 2010). Rrp44 is the only active component of the yeast cytoplasmic exosome (Liu et al. 2006; Dziembowski et al. 2007; Schneider et al. 2007). Electron microscopic analysis of the S. cerevisiae exosome showed that Rrp44 binds to the bottom of the exosome PH ring through the interaction of the PIN domain and Rrp41, and the RNase II-like region contacts with Rrp41, Rrp43, and Rrp45 (see Fig. 8.2d) (Wang et al. 2007). The crystal structure of the ternary complex Rrp44-Rrp41-Rrp45 confirmed this finding and gave more details about the interaction between Rrp44 and the core exosome (Bonneau et al. 2009). The N-terminus of the PIN domain wraps around Rrp41, and the RNase II-like region of Rrp44 contacts both Rrp41Rrp45 and the PIN domain. Biochemical results showed that the interaction between the PIN domain and Rrp41 is the strongest between Rrp44 and the exosome. These interactions seem to be conserved in other species. Structural and biochemical data showed that the PIN domain faces the solvent rather than the exosome core and can be accessed from solvent without passing through the central channel of the core exosome. According to the structure, the exoribonuclease site of Rrp44 is also in principle accessible from solvent. However,

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a conformational change at residues 696–719 of the RNB domain inhibits this entrance, and seems to be stabilized by the binding of Rrp44 to the exosome core. This finding, together with nuclease and RNase protection assays indicate that a path where the 30 -end of the RNA is threaded through the central channel of the core exosome until the exoribonucleolytic site of Rrp44 is favored. However, there is no evidence in vivo that supports this model (Bonneau et al. 2009). Usually, cells present a single Rrp44 ortholog; however, there are cases where two or more orthologs of Rrp44 exist. In S. cerevisiae the unique Rrp44 is present in the nucleus and the cytoplasm (Houseley et al. 2006). In humans, there are two Rrp44 homologues, hDIS3 and hDIS3L, which interact with the exosome and display processive exonuclease activity (Staals et al. 2010; Tomecki et al. 2010). hDIS3 mainly localizes in the nucleoplasm, has endonucleolytic activity, and complements lower levels of yeast Rrp44, while hDIS3L is strictly cytoplasmic and has no endonucleolytic activity, probably due to several mutations in the PINc domain (Tomecki et al. 2010) (Fig. 8.2a). Recently, a protein that contains the RNase II-like region, without one or more of the CSD domains, but lacks the PIN domain was identified in Arabidopsis thaliana (Zhang et al. 2010) (Fig. 8.2a). This protein, named suppressor of varicose (SOV), is a major cytoplasmic protein with mRNA decay activity in vivo. Like RNase II-like proteins, it was proposed to have 30 to 50 exoribonuclease activity, but the absence of the PIN domain suggests that it is not one of the components of the exosome. SOV-like proteins have been conserved in distant lineages, namely, in Mus musculus, Drosophila melanogaster, Oryza sativa, Selaginella moellendorffii and Caenorhabditis elegans. In Drosophila, the Rrp44/Dis3 homologue was characterized and named Tazman. It was shown to be more similar to RNase R and Rrp44 than to RNase II, and similarly to Rrp44, it has an N-terminal PINc domain and a CR3 region (Cairraõ et al. 2005). This N-terminal region is sufficient for the endoribonucleolytic activity and also contributes to the interaction with the exosome and with protein localization (Mamolen et al. 2010). This protein was also shown to be differentially expressed during the Drosophila life cycle, suggesting that it may play an important role in the modulation of RNA levels important for development (Cairraõ et al. 2005).

8.4

DEDD Family

This family of enzymes includes both RNA and DNA exonucleases. These proteins have a characteristic core with four invariant acidic amino acids (which are responsible for the designation of this family) and other conserved residues which are distributed in three distinct sequence motifs. In motif III, the presence of a tyrosine or histidine led to the division of this family into two subgroups, DEDDy and DEDDh, respectively. All proteins of this family share a common mechanism of action which involves two metal ions (Zuo and Deutscher 2001).

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Oligoribonuclease

Pioneering work identified oligoribonuclease as the “finishing enzyme” in RNA metabolism. The degradative activity of other exoRNases result in final RNA fragments ranging from 2 to 5nts whose accumulation could be deleterious to the cell. Oligoribonuclease acts at the final steps of RNA degradation, converting the small oligoribonucleotides to mononucleotides (Ghosh and Deutscher 1999). Oligoribonuclease (OligoRNase/Orn) is a 30 -50 RNase member of the DnaQlike/DEDD exonuclease superfamily. It shows the typical DnaQ-fold containing five-stranded b-sheets flanked by a-helices (Fig. 8.4). The DnaQ-like exonuclease domain contains three well-conserved ExoI, ExoII, and ExoIII sequence motifs clustered around the active site. Like all members of this family, Orn contains four highly conserved acidic residues (DEDD) in the active center. These amino acids are proposed to be essential for binding divalent metal ions and thus for catalytic activity (Steitz and Steitz 1993). In addition to these invariant residues, Orn has other conserved residues with particular importance of a histidine in the ExoIII motif. Such a feature places Orn in the DEDDh subgroup, which includes both DNA- (like the e subunit of DNA polymerase III) and RNA-processing enzymes (like RNase T). Only a preliminary X-ray crystal study of E. coli oligoribonuclease was published (PDB 1YTA) (Fiedler et al. 2004), although a more refined structure is available in the PDB database (PDB 2IGI). A more detailed structural work concerning Orn was done with the plant pathogen Xanthomonas campestris, which oligoribonuclease XC847 shares a 52.6% identity with E. coli Orn (Chin et al. 2006). A general catalytic mechanism for oligoribonuclease was proposed involving all four conserved acidic residues and the conserved histidine present at a highly flexible loop in the ExoIII motif. All DEDD family exonucleases share common active site geometry with the four acidic side chains coordinating two divalent cations (preferably Mn2+ for E. coli Orn). Oligoribonuclease functions as a homodimeric enzyme (Zhang et al. 1998). The dimeric architecture of oligoribonuclease is very similar to the arrangement seen in

Fig. 8.4 Comparison of the structures of the three members of the DEDD family from E. coli: oligoribonuclease (PDB ID 2IGI) (Fiedler et al. to be published), RNase T (PDB ID 2IS3) (Zuo et al. 2007) and RNase D (PDB ID 1YT3) (Zuo et al. 2005)

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RNase T (Fig. 8.4). The two DEDD domains appear to complement each other with single monomer providing a substrate-binding surface leading into the catalytic center of the other monomer (Zuo et al. 2005). The resulting DEDD cavity is long enough to accommodate three to four nucleotides (Zuo et al. 2007). Destabilization of the dimer conformation can alter protein activity and possibly cause its inactivation. Oligoribonuclease is a processive enzyme and nuclease activity is inversely proportional to the length of the single-stranded substrate with a 5-mer oligoribonucleotide being a preferable substrate. Orn is insensitive to 50 RNA phosphorylation state but the molecule must have a free 30 -OH end (Datta and Niyogi 1975). Although Orn is a single-stranded specific exonuclease with strong affinity to small (2–5nts) RNA fragments, it has been reported that higher concentrations of the enzyme can degrade short DNA oligos (Mechold et al. 2006). Interestingly, the overall architecture of XC847 is quite similar to 30 -50 DNases. Oligoribonucleases are found in the proteobacteria (b and g divisions) and Actinomycetes in bacterial genomes and have not been detected in archaea, although are present in all eukaryotes (Zuo and Deutscher 2001). Oligoribonucleases are inhibited by the nucleotide 30 -phosphoadenosine-50 phosphate (pAp) that is generated in both prokaryotes and eukaryotes during the process of sulfur assimilation (Mechold et al. 2006). E. coli oligoribonuclease is encoded by the orn gene and is the only essential exoRNase required for cell viability in this organism (Ghosh and Deutscher 1999). B. subtilis lacks an Orn homologue (Mechold et al. 2006). However, B. subtilis has at least two functional analogues of Orn, termed nanoRNases NrnA and NrnB that can complement in vivo a defective E. coli orn mutant. The preferred substrate of these enzymes is a 3-mer instead of 5-mer RNA (Fang et al. 2009). Most likely, additional Nrn-like enzymes are present in B. subtilis genome as a double nrnA nrnB mutant is viable. Homologues of bacterial oligoribonuclease are found in many eukaryotes (Zhang et al. 1998). Yeast homologue Ynt20/Rex2 is localized in mitochondria whereas its function there remains unclear (Hanekamp and Thorsness 1999). It was also reported to function in the nucleus, namely, in the processing of some stable and small RNAs (van Hoof et al. 2000). The human homologue was proposed to exist in two isoforms that arise from alternatively spliced transcripts, one of them, Sfna, contains mitochondrial targeting sequence, while the other protein, referred to as Sfn, was shown to be oligoribonuclease in vitro and, in contrast to the bacterial enzyme, is able to digest both DNA and RNA substrates (Nguyen et al. 2000).

8.4.2

RNase D

RNase D is a 40 kDa protein encoded by the rnd gene. As a member of DEDD family of enzymes, it requires divalent metal ions for its activity and has a high degree of substrate specificity (Cudny et al. 1981). This enzyme was initially discovered through its action on “denatured” and damaged tRNAs but it also acts on tRNA precursors with extra 30 residues following the CCA sequence, 5S rRNA

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and other small-structured RNAs but not ssRNA (Cudny and Deutscher 1980; Zhang and Deutscher 1988b). RNase D is not an essential protein for the cell, however, is crucial for viability when RNase II, BN, T, and PH are absent, which may indicate that is a backup enzyme when the principal ribonucleases are missing (Kelly and Deutscher 1992b). RNase D overexpression seems to be deleterious for the cell (Zhang and Deutscher 1988a). In the cell, the rnd expression may be limited because: the chromosomal gene uses UUG as the initiation codon; it has an abnormally high level of rare codons; its expression is negatively regulated at the translational level by the initiation codon (Zhang and Deutscher 1989). The resolution of the RNase D crystal structure showed that this protein has one Cterminal DEDD catalytic domain and two HRDC domains at the N-terminal region. The interactions between HRDC2 and DEDD are responsible for bringing the three domains into a funnel-shaped ring structure, which is very flexible, and suggests a processive activity (Fig. 8.4). The DEDD domain forms a closed, very compact structure on one side, and an open pocket on the other side. The putative active site of RNase D (formed by the DEDD residues) is located inside the open pocket, which is surrounded by three flexible loops. RNase D is not able to degrade short oligonucleotides, possibly due to their weak binding at the active center (Zuo et al. 2005). RNase D homologues have been found in many bacterial organisms. In some genomes, it is possible to find more than one close homologue. In archeal genomes it was not possible to find any RNase D homologue, while in Eukaryotes there is at least one RNase D homologue, named Rrp6 (Zuo and Deutscher 2001). In contrast to the situation in the Archaea, in yeast and human cells the exosome ring proteins do not have phosphorolytic activity and the only active subunits of the complex are the two ring connected nucleases Rrp44 and Rrp6 (Liu et al. 2006; Dziembowski et al. 2007). Rrp6 structure of the yeast enzyme was solved showing similarity of nuclease domain to RNase D (Midtgaard et al. 2006) with the presence of one HRDC domain. Rrp6 is a distributive single-strand specific RNA nuclease. Recently, it was shown that the TRAMP complex accelerates the rate of RNA degradation by yeast Rrp6 in vitro. TRAMP is a complex which contains a polymerase, a helicase, and zinc knuckle proteins that add poly(A) tails to eukaryotic RNA substrates, targeting them to degradation (Callahan and Butler 2010). In yeast, Rrp6 plays a role in nuclear mRNA surveillance and in the degradation of rRNA maturation by-products or intergenic transcripts (Houseley et al. 2006). It is also involved in the final step in processing several noncoding RNAs (Allmang et al. 1999). The intracellular localization of Rrp6 in eukaryotes is intriguing. In yeast cells this nuclease is strictly nuclear; in human cells hRrp6 can be found in both the nucleus (where it seems to be enriched in nucleoli), and in the cytoplasm. In plants, it was shown that there are three different Rrp6-like proteins, which are localized in nucleus, nucleolus, and cytoplasm, respectively. Each of them seems to serve some specific and unique function (Lange et al. 2008). Nuclear Rrp6 co-purifies with exosome complex; however, Rrp6 was shown to have functions that are independent from the exosome. It will be interesting to determine whether cytoplasmatic Rrp6 also helps in exosome substrate degradation, or is an independent enzyme with specific functions.

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RNase T

RNase T, also a member of the DEDD superfamily of RNases, is a single-strandspecific exoribonuclease, which also has DNA exonuclease activity (Viswanathan et al. 1998). It has a distributive activity, which depends on the presence of divalent metal ions, such as Mg2+ or Mn2+, and an unusual base specificity, discriminating against pyrimidines and, particularly, C residues. This sequence specificity is determined by the last four nucleotides at the 30 end (Deutscher and Marlor 1985; Zuo and Deutscher 2002a, b). When compared to other ribonucleases, RNase T is the only enzyme capable of removing nucleotides near the duplex structure without unwinding the substrate, generating blunt-ended RNAs. RNase T is involved in the final step of maturation of many stable RNAs (Li and Deutscher 1995, 1996; Li et al. 1998): it is essential for the maturation of the 30 ends of 5S and 23S rRNA genes (Li and Deutscher 1995; Li et al. 1999), and it is also involved in the end turnover of tRNAs (Deutscher and Marlor 1985). In order to be functional, RNase T needs to form a dimer, both in vivo and in vitro (Li et al. 1996). Mutational analysis identified three nucleic acid-binding sequence (NBS) segments important for substrate binding (Zuo and Deutscher 2002a). The resolution of crystal structure of RNase T from both E. coli and P. aeruginosa shows that the protein has an oligoribonuclease-like homodimer architecture (Fig. 8.4) (Zuo et al. 2007). The two monomers face in opposite directions, that is, the NBS segment from one monomer is located in the vicinity of the DEDD active center pocket of the other monomer. This arrangement allows the binding of the RNA molecule from one monomer to be close to the active site of the other, and also helps us to explain why the enzyme requires the formation of the homodimer in order to be active (Zuo et al. 2007). Despite its critical role in RNA metabolism, RNase T orthologs are only found in the g division of Proteobacteria (Zuo and Deutscher 2001).

8.4.4

Deadenylases

In eukaryotic cells, deadenylation is the first step that precedes messenger degradation. Poly(A) tails are important not only in controlling the stability of the transcripts, but also in RNA processing, translation, or nuclear export processes. Deadenylation is performed by poly(A)-specific 30 -50 RNA exonucleases. In yeast, two different protein complexes were recognized to be involved in RNA deadenylation: Ccr4-Caf1 and Pan2-Pan3 (Tucker et al. 2001; Yamashita et al. 2005). Ccr4-Caf1 (or Ccr4-NOT) contains at least ten interacting proteins, among them Ccr4 and Caf1(Pop2) are the poly(A)-specific nucleases. Caf1 is the nuclease classified in the DEDD family, although the characteristic motif in this enzyme is replaced by SEDQ sequence (Thore et al. 2003). However, the human orthologs of Caf1 contain the conserved catalytic DEDD motif. The other nuclease in the complex-Crr4 represents a DNase-I like family of two metal ion dependent

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nucleases. The crystal structure of the catalytic domain of the human homologue of Ccr4-CNOT6L gave some insights into its poly(A) specificity and catalytic mechanism (Wang et al. 2010b). Ccr4 is suggested to be the main deadenylase in yeast Ccr4-Caf1 complex, but both nucleases of the complex seems to be involved in RNA degradation (Schwede et al. 2008). The active subunit of the Pan2-Pan3 deadenylation complex is Pan2, a DEDD family nuclease, while the protein in the second complex, Pan3, stimulates the nuclease activity (Brown et al. 1996). In yeast, the CCr4-Caf1 complex seems to be the main deadenylation factor; however, in mammalian cells, both complexes Ccr4-Caf1 and Pan2-Pan3 work sequentially in degradation of poly(A) tails with initial shortening by Pan2-Pan3 and subsequent degradation by Ccr4-Caf1 (Yamashita et al. 2005). Many eukaryotes contain also a third poly(A)-specific nuclease, named PARN. The PARN nuclease domain contains the characteristic DEDD motif and is structurally similar to Caf1. Moreover, PARN contains an RNA recognition motif (RRM) and an R3H domain. In addition to its nuclease activity, PARN is also a cap-binding protein and the interaction with RNA cap stimulates the RNase activity of the enzyme. PARN is a dimeric protein and its crystal structure, together with cap and with poly(A) RNA, showed that cap binding causes conformational changes, suggesting the existence of an open and closed form of the active site (Wu et al. 2005). Other yeast nucleases from the DEDD nuclease family are Rex1, Rex3, and Rex4. The two first were proved to be involved in RNA processing events. Homologues of these proteins can also be found in other eukaryotes, including humans (van Hoof et al. 2000). The ERI-1 exoribonuclease also belongs to this family of enzymes. The enzyme of Caenorhabditis elegans and Schizosaccharomyces pombe negatively regulates the RNA interference pathway, probably by degrading dsRNAs. In both organisms, Eri-1 also takes part in the processing of 30 end of 5.8S rRNA (Kennedy et al. 2004; Gabel and Ruvkun 2008). In humans, the homologue of Eri-1 is known to be involved in the metabolism of histone mRNA but its exact role in this and other processes in mammalian cells still remains to be elucidated (Yang et al. 2009).

8.5

Concluding Remarks

To maintain the correct RNA levels in the cell, RNA metabolism must be tightly controlled. One of the factors involved are ribonucleases, which will determine the degradation pathway for each RNA molecule. Exoribonucleases play an important role in the mechanism of RNA degradation, and they behave differently regarding RNA recognition and degradation. While some RNases are important for RNA degradation, others are specialized in the processing and maturation of some types of RNA. The determination of crystal structures of these enzymes has allowed investigators to better understand their mechanism of action and further our knowledge of RNA metabolism. It is also possible to verify that the similarities between prokaryotes and eukaryotes are much higher than initially expected.

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Acknowledgment We thank Miguel Luı´s for graphical assistance in Figs. 8.2 and 8.3.

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Chapter 9

The RNA Exosomes Karl-Peter Hopfner and Sophia Hartung

Contents 9.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Biology of Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.1 Exosomes in Eukaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.2 Exosomes in Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Exosome Architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.1 The Nine Subunit Core Exosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.2 The Eukaryotic Exosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4 Biochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.1 Archaeal Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.2 RNA Recognition, S1, ZnR, and KH Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.3 Eukaryotic Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5.1 Evolutionary Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5.2 Future Questions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract RNA exosomes are large multimeric 30 -50 exo- and endonucleases found in eukaryotes and many archaeal species. They represent the central RNA 30 -end processing factor and are implicated in processing, quality control, and turnover of both coding and noncoding RNAs. RNA exosomes are highly regulated and processive machineries, assembled as large macromolecular cages

K.-P. Hopfner (*) Department of Biochemistry at the Gene Center, Ludwig-Maximilians-University, Feodor-LynenStr. 25, 81377 Munich, Bavaria, Germany Center for Integrated Protein Sciences, Ludwig-Maximilians-University, Munich, Bavaria, Germany e-mail: [email protected] S. Hartung Life Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_9, # Springer-Verlag Berlin Heidelberg 2011

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that channel RNA to the ribonuclease sites. The primordial exosome – found in archaea and related to bacterial and organelle degradosomes – possesses a phosphorolytic active cage that can both degrade and polyadenylate RNA in RNA decay processes. Human and yeast exosomes lost phosphorolytic activities but gained ectopic subunits with hydrolytic activities, while preserving the RNA channeling function.

9.1

Introduction

In all kingdoms of life, the basal physiology, homeostasis, adaptation, and differentiation of cells is regulated at different levels, including transcription, translation, and mRNA decay. A critical quantity is the level of mRNA, which is controlled on one side by transcription by RNA polymerases, and degradation by ribonucleases on the other side. It is well known that transcription of different RNA species is a highly differential, regulated process and requires the temporal as well as spatial activity of a large number of transcription factors and co-activators as well as other features such as chromatin structure. Although mRNA decay appears much less regulated and fine tuned than transcription, it also has temporal as well as spatial components and it is of course important for cells to be able to either stabilize mRNAs or degrade them in a controlled, regulated manner that is coordinated with the synthesis machinery (Maniatis and Reed 2002). Furthermore, besides the turnover of “normal” RNA, surveillance mechanisms often check the functionality of RNA molecules. For example, mRNAs that lack a stop codon or contain premature stop codons are removed by nucleolytic degradation in processes called nonstop and nonsense-mediated decay, respectively (Belgrader et al. 1994; Frischmeyer et al. 2002; Whitfield et al. 1994). RNA is degraded by a large variety of nucleases, but the RNA exosome, a large multisubunit complex, emerged over the past decade as the key player in the regulated, controlled RNA processing and degradation in eukaryotes. After its original discovery in yeast and humans, exosome-like complexes have been identified in archaea (Allmang et al. 1999b; Estevez et al. 2001; Evguenieva-Hackenberg et al. 2003; Koonin et al. 2001; Mitchell et al. 1997). It was then established by structural biology results that eukaryotic and archaeal exosomes share many characteristic features and are related to the bacterial degradosome or polynucleotide phosphorylase (PNPase) as well as the phosphorolytic RNase PH (Buttner et al. 2005; Lorentzen et al. 2005; Symmons et al. 2000). The evolutionarily conserved core exosome is a particle of approx. 250–300 kDa molecular mass and consists of nine subunits, six of which are related to RNase-PH, and three subunits that contain RNA binding S1 or KH domains (Table 9.1, Fig. 9.1). The core shares much of the general fold and assembly with RNase-PH like polynucleotide-phosphatase (PNPase) as well as hexameric RNase-PH itself, both phosphorolytic ribonucleases in bacteria. Phosphorolytic nucleases use inorganic phosphate instead of a water molecule to attack the RNA phosphodiester bond. As

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Table 9.1 Exosome components in archaea and eukaryotes Exosome component Archaea Eukaryotes PH domain 1 PH domain 2 RNA-binding protein Active hydrolytic RNase Accessory or associated proteins

Rrp41 (active) Rrp42 Rrp4, CsI4 – DNAG, aNip7

Rrp41, Mtr3, Rrp46 Rrp42, Rrp43, Rrp45 Rrp4, Rrp40, CsI4 Rrp44(DIS3), Rrp6, DIS3L Nucleus: Rrp47, Nip7, TRAMP complexes, Mpp6, Ndr1-Nab3,Rnt1,AID Cytoplasm: TUTases, ARE-BP, ZAP, Dom34/Hbs1, Ski7, SKI complex

Fig. 9.1 Evolutionarily conserved exosome and PNPase architectures. Top panel: Structures are shown as surface representation with some annotated domains. In this “top view,” the reader views down the RNA entry pore. Middle panel: schematic and subunit annotated representations in top view. Bottom panel: ribbon models in side view

a result, nucleoside diphosphates are liberated from the 30 end of RNA. The reaction is energetically highly reversible, in contrast to hydrolytic RNA degradation and phosphorolytic RNases also catalyze the reverse reaction, adding nucleoside diphosphates to the 30 end and liberating inorganic phosphate. In archaea, the exosome contains three copies of each of the RNase PH-type subunits aRrp41 and aRrp42. They are assembled in a large macromolecular cage with three self-compartmentalized phosphorolytic active sites. This architectural principle, shielding active sites from unrestricted access, resembles the proteasome for protein degradation. RNA is channeled into the exosome through a pore,

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a feature that is well suited to provide means for tight regulation, but also to provide highly processive RNA degradation, once RNA is loaded into the exosome. The eukaryotic exosome contains six different RNase PH-type subunits (Rrp41, Rrp46, Mtr3, Rrp42, Rrp43, Rrp46 in S. cerevisiae) along with S1 and/or KH domain containing subunits Rrp4, Rrp40, and Csl4 (Table 9.1, Fig. 9.1). The RNase-PH type subunits have lost phosphorolytic activity in yeast and humans and the eukaryotic exosome contains the additional hydrolytic subunits Rrp6 and/ or Rrp44. Despite this shift in nuclease activities, the overall architecture and assembly of the exosome remains preserved and is clearly important as deduced from the essential nature of the “inactive” RNase PH subunits on yeast viability. In fact, the RNase-PH core still channels RNA Bonneau et al. 2009. From an evolutionary point of view the remarkable switch in activity from phosphorolytic to hydrolytic presumably might be a consequence or prerequisite to the more complexly regulated eukaryotic RNA degradation pathways. All in all, the emerging architectures reveal exosomes as a fascinating platform for targeted, regulated, and processive degradation of RNA molecules, and this chapter highlights the current understanding in terms of biology and molecular mechanism.

9.2

Biology of Exosomes

9.2.1

Exosomes in Eukaryotes

9.2.1.1

A General Machinery for 30 -50 Degradation

The “exosome” as a multisubunit RNase assembly was first described by David Tollervey and coworkers as large complex with 30 -50 exonuclease activity in S. cerevisiae (Mitchell et al. 1997). It was then shown that the human equivalent is the PM-Scl complex, a complex previously found to be the component of an antibody-autoantigen system in patients with polymyositis and scleroderma (Allmang et al. 1999b; Reimer et al. 1986). Exosomes were furthermore studied in trypanosomes (Estevez et al. 2001) and plants (Chekanova et al. 2000). The first identified activity of the yeast exosome was an essential role in rRNA biogenesis (Allmang et al. 2000; Mitchell et al. 1997; Zanchin and Goldfarb 1999b), but subsequent studies showed that the exosome is in fact a key player in most, if not all, pathways that require RNA processing. For instance, the exosome together with the Ski (superkiller) complex has been found to be a key enzyme in 30 -50 mRNA decay (Anderson and Parker 1998; Araki et al. 2001; van Hoof et al. 2000b; Wang and Kiledjian 2001). The involvement of the Ski complex provided a first insight that exosomes also need cofactors. The exosome is furthermore important for the biogenesis of the signal recognition particle (Grosshans et al. 2001), the turnover of small nuclear and nucleolar RNAs (van Hoof et al. 2000a), and rapid decay of AU-rich element (ARE) unstable mRNAs (Chen et al. 2001; Haile et al. 2003).

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Besides the processing of rRNA and decay or turnover of small RNAs and mRNA, it became rapidly clear that the exosome is also a key player in RNA quality control, for example, for degradation of RNAs that lack a stop codon (nonstop decay) (Frischmeyer et al. 2002; van Hoof et al. 2002) as well as nonsense-mediated decay, which degrades RNA with premature stop codons (Frischmeyer et al. 2002; Lejeune et al. 2003; Mitchell and Tollervey 2003; Takahashi et al. 2003). The exosome furthermore degrades splice-defective RNA in the cytoplasm (Hilleren and Parker 2003) and hypomodified tRNAs, rRNA precursors, and mRNAs with defective polyadenylation in the nucleus (Fang et al. 2005; Kadaba et al. 2004). The depletion or inactivation of exosome subunits not only identified their role in living cells and which of the RNA processing and degradation pathways requires the exosome, but also uncovered new RNA processing pathways and even new types of RNA molecules that were enriched in the absence of the exosome. This led, for instance, to the discovery of a pathway for nuclear pre-mRNA turnover (BousquetAntonelli et al. 2000). The mechanistic basis for this activity is a physical link between the exosome and elongating RNA polymerase in Drosophila (Andrulis et al. 2002). Furthermore, depletion or inactivation of exosomes in yeast and human cells led to the enrichment and identification of otherwise highly unstable, short RNAs that are normally rapidly degraded by exosomes. In yeast, these RNAs are called cryptic unstable transcripts (CUTs) and arise from transcription of intergenic regions (Wyers et al. 2005). In humans, related RNA molecules result from transcription upstream of active promoters and are called PROMPTs (promoter upstream transcripts) (Preker et al. 2008). Furthermore, RNAs arising from antisense transcription of the noncoding strand in transcriptional gene silencing are degraded by the exosome as well (Camblong et al. 2007). A recent review on these classes or RNA can be found, for example, in Carninci (2010). Despite its role in degrading coding and noncoding RNAs, the exosome also plays more complex regulatory roles in cells that perhaps go beyond simple RNA degradation. For instance, the exosome plays a role in Neurospora circadian gene expression (Guo et al. 2009), while in immune B cells, the exosome is involved in the generation of antibody diversity via somatic hypermutation and class switch recombination. Here, the exosome recruits the activation-induced cytidine deaminase (AID) to transcription elongation complexes at immunoglobulin loci (Basu et al. 2011).

9.2.1.2

Exosome Isoforms

In eukaryotes, the exosome exists in nuclear and cytoplasmic isoforms. Both isoforms share the nine core subunits as well as the RNase-R like Rrp44/Dis3 subunit. Rrp44/Dis3 carries the main constitutive exonuclease activity of the exosome (Dziembowski et al. 2007). Subsequent studies showed that Rrp44 also possesses endonuclease activity, which was a surprise to the field and created much

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excitement (Lebreton et al. 2008; Schaeffer et al. 2009; Schneider et al. 2009), because it uncovered a far more complex role of the exosome in degrading RNA substrates and drew exciting parallels to the combination of endo- and exonuclease activities of prokaryotic and organellar degradosomes (for a recent review, see, e.g., (Tomecki and Dziembowski 2010)). The nuclear isoform of the exosome also contains the Rrp6 protein, an RNase-D like hydrolytic nuclease. The activity of Rrp6 is for instance required to trim rRNA (Allmang et al. 1999a) and small nuclear and nucleolar RNAs (snRNAs and snoRNAs) (van Hoof et al. 2000a). In contrast, Rrp6 is not required for mRNA decay (cytoplasm), but required for nuclear mRNA quality control (Hilleren et al. 2001). In humans, the nuclear exosome associates with PM-Scl100, the homolog of yeast Rrp6. Thus the presence of a nuclear isoform containing an RNase-D activity appears to be evolutionarily preserved. Besides the specific association of the nuclear exosome with PM-Scl100/Rrp6, human exosomes associate with two different homologs of yeast Dis3p, denoted hDIS3 and hDIS3L (or DIS3-like 1) (Staals et al. 2010; Tomecki et al. 2010). Significantly, hDIS3 and hDIS3L are differently distributed in human cells: hDIS3 is nuclear, while hDIS3L is cytoplasmic. Consistently, hDIS3L is involved in cytoplasmic mRNA decay (Staals et al. 2010). While both proteins are active exonucleases, only hDIS3 has an additional endonuclease activity (Tomecki et al. 2010). In summary, the current data suggest that the human exosome exists with respect to both PM-Scl100 and DIS3 in distinct isoforms in the nucleus and cytoplasm, and hence possesses different endo- and exonuclease properties.

9.2.1.3

Cofactors

Since the exosome and its isoforms play key roles in numerous RNA processing and degradation pathways, they must be able to target RNA molecules in a manner independent of sequence, secondary structure, and even bound proteins. On the other hand, the activity of the exosome needs to be tightly controlled to avoid unregulated degradation of RNAs. There are several mechanisms that ensure broad but also regulated activity. One mechanism is the structure of the exosome itself. It channels RNA through a narrow pore that restricts entry to single-stranded unfolded RNA molecules (see below). Another aspect is the specific targeting of the exosome, for example, to transcription elongation complexes or stalled ribosomes (via Ski7). Finally, a variety of cofactors help to prepare RNA molecules for degradation by the exosome. It was not a surprise that Ski2 and Mtr4, two members of the large superfamily 2 helicase/nucleic acid translocases, were found to act as cofactors of the exosome (Anderson and Parker 1998; de la Cruz et al. 1998). Naturally, these enzymes can help unwind structured RNA for degradation and strip proteins from RNA. Ski2 exists in a hetero-trimeric or -tetrameric complex together with Ski3 and Ski8 (Synowsky and Heck 2008; Wang et al. 2005). The Ski complex is found in the cytoplasm and is required for several cytoplasmic activities of the exosome,

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including mRNA degradation (Anderson and Parker 1998; van Hoof et al. 2000a), degradation of mRNA targeted by the RNA-induced silencing complex (RISC) (Orban and Izaurralde 2005), and degradation of deadenylated mRNA in NMD (Mitchell and Tollervey 2003). Mechanistically, the activity of the Ski complex is not well understood, but the crystal structures of the Mtr4 homolog and more distantly related bacterial Ski-like helicases (which function in DNA replication and repair) as well as of the b-propeller protein Ski8 have been determined (Buttner et al. 2007; Cheng et al. 2004; Jackson et al. 2010; Madrona and Wilson 2004; Weir et al. 2010; Zhang et al. 2008). Ski2 is closely related to Mtr4, which is found in the nucleus and is involved in processing and degradation of rRNA, snRNAs, snoRNAs, and tRNAs (Allmang et al. 1999a; Cristodero and Clayton 2007; van Hoof et al. 2000a; Wang et al. 2008). Mtr4 has been biochemically characterized as a 30 -50 , ATP-dependent helicase. ATP hydrolysis is stimulated by, for example, tRNA but not by poly(A) RNA (Bernstein et al. 2008), indicating that Mtr4 prefers structured RNA. In this regard, Mtr4 possesses a peculiar arch domain that could help present structured RNA/ tRNA to the helicase core (Jackson et al. 2010; Weir et al. 2010). Mtr4 exists in the nucleus in a complex with two other polypeptides and forms the TRAMP4 (Mtr4-Air2-Trf4) and TRAMP5 (Mtr4-Ari2-Trf5) complexes (Houseley and Tollervey 2006; LaCava et al. 2005). TRAMP oligo/polyadenylates nuclear RNAs that are subsequently degraded by the exosome in nuclear pathways. These results established that adenylation is a common mechanism for RNA stability and turnover through exosome-mediated degradation in both the nucleus and the cytoplasm.

9.2.2

Exosomes in Archaea

While the exosome is a firmly established key player in the 30 degradation of RNA molecules in eukaryotes, the biology of archaeal exosomes is less clear. In fact, not all archaea contain homologs of exosome subunits, so other activities such as RNase-R (Portnoy and Schuster 2006) can perhaps compensate for a lack of exosome-like complexes. The archaeal exosome was first predicted by bioinformatic analysis of archaeal genomes (Koonin et al. 2001) and shortly after experimentally identified in Sulfolobus solfataricus (Evguenieva-Hackenberg et al. 2003) and Methanococcoides burtonii (Goodchild et al. 2004). The main biochemical difference between archaeal and eukaryotic exosomes is the presence of phosphorolytic activity in the former, while the latter appears to lack this activity and instead adopted additional subunits with hydrolytic activity. Initial insights into the role of exosomes in archaea came from the analysis of poly (A) tails on archaeal RNA. It was found that RNAs from a halophilic archaeon, which does not encode homologs for exosome subunits in its genome, do not show RNA poly(A) tails. It has been suggested that in the absence of the exosome, RNA degradation could be performed by an RNase R homolog (Portnoy and Schuster

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2006). Interestingly, RNase-R is related to Rrp44, a hydrolytic subunit of the eukaryotic exosome. While poly(A)-rich tails are not found in halophilic archaea, heteropolymeric, A-rich tails were found on RNA from S. solfataricus, which possesses an exosome (Portnoy et al. 2005). The presence of these tails in Sulfolobus and methanogens correlates with the presence on an exosome (Portnoy and Schuster 2006). The added tails are poly(A)-rich but heteropolymeric, suggesting that the composition perhaps reflects the nucleoside diphosphate concentrations in the cell. In other words, the exosome is not a specific poly(A) polymerase, but nondiscriminatorily adds nucleoside diphosphates in vivo similar to its biochemical properties in vitro. A role in both adding tails and degrading RNA is reminiscent of chloroplast PNPase, which also adds tails to RNA, followed by endonucleolytic cleavage and exonucleolytic degradation (Yehudai-Resheff et al. 2003). It is therefore possible that the archaeal exosome, like PNPases, acts in vivo both by adding tails and degradation of RNA (Slomovic et al. 2008), but it remains to be shown whether these activities are regulated. Recent single molecule studies showed that exosomes can easily switch between the two types of activities and there appears to be a memory effect for polymerization as well as degradation modes under conditions when both reactions are thermodynamically in equilibrium, that is, the exosome stochastically switches between degradation and polymerization but stays in one mode for a while (Lee et al. 2010). It is yet unclear how archaeal exosomes recognize their specific RNA targets and how the activity of the exosome is regulated in the archaeal cell. The core exosome is found associated with other polypeptides such as the archaeal DnaG homolog (Evguenieva-Hackenberg et al. 2003; Walter et al. 2006); however, the function of this protein in the context of the exosome is still unclear. DnaG is the primase in bacterial DNA replication. Archaea contain in addition to an archaeo-eukaryotic type of primase, which acts in DNA replication, a DnaG-like enzyme with a Toprim fold, and this protein is found in the exosome complex (Evguenieva-Hackenberg et al. 2003; Iyer et al. 2005; Walter et al. 2006). However, Sulfolobous DnaG also has a primase type activity in vitro, and the biochemical connection to exosomes is unclear (Zuo et al. 2010). Other interactors include archaeal Nip7, which inhibits degradation of poly-(A) and poly-(AU) RNA in vitro. Eukaryotic Nip7 also interacts with the eukaryotic exosome and is required for pre-rRNA processing, suggesting an evolutionary conserved targeting or regulation (Luz et al. 2010; Zanchin and Goldfarb 1999a). Recent exciting results show that the exosome is localized to membranes in archaea (Roppelt et al. 2010), a feature it shares with the degradosome (Khemici et al. 2008). In the case of degradosomes, membrane association is mediated by an amphipathic helix in the RNase-E subunit (Khemici et al. 2008). A potential candidate for membrane association is the DnaG subunit of the archaeal exosome (Evguenieva-Hackenberg et al. 2011). The functional importance of this subcellular localization remains to be established. In principal, it could be an important mechanism to target specific RNA molecules or a consequence of RNA localization to bacterial membranes (Kawamoto et al. 2005; Nevo-Dinur et al. 2011).

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In summary, the physiological isoforms of exosomes in archaea and their differential role in RNA processing and degradation require further study, although considerable progress in this respect has been made. For instance, archaeal exosomes can have different RNA binding (aRrp4 and aCsl4) caps, which confer different biochemical properties on the exosome. The role of different cap types in different archaeal RNA degradation or processing pathways remains to be studied. However, the cap proteins could be differentially expressed: While the aRrp4 gene is in the same operon as genes for aRrp41 and aRrp42, the gene for aCsl4 is located elsewhere in the genomes, offering the possibility of production of different exosome isoforms also in archaea by gene regulation.

9.3 9.3.1

Exosome Architecture The Nine Subunit Core Exosome

The pioneering studies on the archaeal exosome began with the structure of the 6-subunit RNase-PH-like ring that revealed the presence and location of three active sites and established the structural relation with PNPase (Lorentzen et al. 2005). Shortly after, structures of the full 9-subunit exosome in two different isoforms (with aRrp4- and aCsl4-type caps) (Buttner et al. 2005), and the 6-subunit and 9-subunit exosomes in complex with RNA were reported (Lorentzen and Conti 2005; Lorentzen et al. 2007). These studies showed that exosomes are “self-compartmentalized” RNases with an entry and exit pore for RNA and degradation products, respectively. Subsequent studies on different archaeal exosomes, with and without RNA, showed features of degradative processivity, conformational flexibility, and RNase as well as polymerization mechanisms (Hartung et al. 2010; Lu et al. 2010; Navarro et al. 2008). The archaeal exosome possesses a double-donut-like structure with a central cavity (Figs. 9.1 and 9.2). One donut is formed by six RNase PH-like domains. For RNase-PH, each of the six subunits is an active nuclease, while the archaeal exosome contains 3 “active” Rrp41 and three “inactive” Rrp42 type subunits. One Rrp41 and one Rrp42 together form a single phosphorolytic active site and the exosome can be viewed as trimer of Rrp41:42 dimers. The central cavity in the exosome, formed by the three pairs of circularly arranged Rrp41:Rrp42 dimers, therefore contains three phosphorolytic active sites. This structure is similar to the structure of bacterial PNPase, except that Rrp41 and Rrp42 are separated polypeptides, while in PNPase, both PH domains are contained in a single polypeptide chain. In fact, recent results also showed that despite the diverse genomic coding and architecture, RNA channeling into exosome and PNPase are related and proceed through analogous entry pores, although some differences in the narrow constrictions and channeling loops were observed (Shi et al. 2008; Symmons et al. 2000).

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Fig. 9.2 RNA channeling by the archaeal and eukaryotic exosomes. The location of the ribonuclease active site moved from inside the processing chamber (green star) in the bacterial PNPase and archaeal exosome to the tenth subunit Rrp44 in eukaryotes. Rrp44 contains two different active sites, an exonucleolytic active site (dark blue star) that is only reachable through the processing chamber of the exosome and an endonucleolytic active site in the PIN domain (light blue star). Although the location of the active site is changed, the RNA channeling mechanism is conserved

The (Rrp41:Rrp42)3 ring is bound in total by three copies of Csl4 and/or Rrp4 domains. Csl4 and Rrp4 both contain an S1 domain and the three copies of the S1 domains frame the “entry” pore for RNA into the active site. Although direct binding of the RNA to the S1 domains has not yet been observed, it is likely that these domains help to recruit and channel RNA into the central processing chamber. In fact, RNA could be visualized in the narrow neck between the S1 domains and the processing chamber using RNA with a single strand 30 tail and a stable hairpin structure at the 50 end (Lorentzen et al. 2007) and mutations in the neck also interfere with exosome activity (Buttner et al. 2005; Oddone et al. 2007). In summary, the narrow neck is the likely entry point for RNA and presumably functions to limit entry into the RNA processing chamber of only unfolded, protein-stripped RNAs. Besides the more central S1 domains that may guide RNA through the entry pore, both Csl4 and Rrp4 contain additional peripheral domains. Csl4 contains a ZR (zinc ribbon) domain and Rrp4 a KH domain. KH domains are well-known RNA interaction domains, suggesting that these domains are important elements of RNA targeting. The function of the ZR is less clear and it could mediate interactions either with specific RNA substrates or alternatively with other proteins. The eukaryotic RNase-PH-like ring is composed of three “aRrp41” homologs (Rrp41, Rrp46, Mtr3) and three “aRrp42” homologs (Rrp42, Rrp43, Rrp45). This core ring binds to Rrp4, Rrp40, and Csl4 and the positioning of S1, KH, and ZR is very similar to the position of equivalent domains on the archaeal exosome. The conservation of S1, KH, and ZR domains on the exosome surface between archaea and eukaryotes is remarkable and suggests that basal mechanism of RNA recognition and recruitment are preserved.

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The Eukaryotic Exosome

The 9-subunit eukaryotic core exosome associates with the hydrolytic nucleases Rrp44 and Rrp6, a feature that is very distinct from archaeal exosomes. The interaction of Rrp44 subunits with the core has been visualized by X-ray crystallography and by electron microscopy and additionally analyzed by mass spectrometry (Bonneau et al. 2009; Cristodero et al. 2008; Malet et al. 2010; Taverner et al. 2008). These results showed that Rrp44 binds to several RNase-PH subunits on the opposite site of the surface that is covered by the S1 domains (Figs. 9.1 and 9.2). This led to the idea that RNA exiting the core exosome is directly fed into the exonuclease active site of Rrp44. The PIN domain harboring the endonucleolytic site is located slightly to the outside of the exit pore of the core ring. It remains to be shown how this endoribunuclease acts in the context of the exosome. In contrast to Rrp44, the location of Rrp6 is less well understood. Electron microscopy on trypanosome exosomes suggests it binds to the outside and not near the exit pore (Cristodero et al. 2008). Thus, the mutual orientation of exo- and endoribonuclease active sites on the eukaryotic exosomes needs to be further addressed.

9.4 9.4.1

Biochemistry Archaeal Exosomes

In addition to the in vivo cell biology experiments on exosomes, biochemical studies have been performed on the processes of RNA recognition, binding, and degradation. Even before the structure of the archaeal exosome was determined, the active site was identified by homology with the phosphate binding sites and biochemical properties of bacterial PNPase and RNasePH (Harlow et al. 2004; Symmons et al. 2000). The archaeal exosome, like the bacterial PNPase (Deutscher and Reuven 1991), is a phosphorolytic RNase that cleaves the phosphodiester bond between the first and the second base at the 30 end of RNA by attacking the backbone with phosphate (instead of water in hydrolytic enzymes) and releasing a nucleoside 50 -diphosphate as product (Fig. 9.3). Several crystal structures of archaeal exosomes have been solved in complex with different RNA molecules (Hartung and Hopfner 2009; Lorentzen and Conti 2005; Lorentzen et al. 2007; Navarro et al. 2008). These structures showed that the active site binds the 30 -end of RNA through interactions with its backbone and base stacking. This sequence-unspecific recognition of the RNA substrate explains why the exosome can degrade mRNAs with any sequence and leaves the role of substrate selection to the additional RNA-binding domains and/or accessory proteins. Interactions between the 20 -hydroxy groups of two riboses and side chains of the exosome ensure RNA over DNA discrimination. A tyrosine residue in the active site of the exosome

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Fig. 9.3 Proposed RNA cleavage mechanisms for structurally characterized exosome active sites. Residues important for catalysis or substrate recognition are shown. Interactions responsible for RNA specificity are shown with dashed lines. (a) Archaeal phosphorolytic cleavage (b) hydrolytic cleavage in Rrp44 (c) two metal ion mechanism for hydrolytic cleavage in Rrp6. The Rrp6 structure was solved with bound product, not substrate

is stacked between the fourth and the fifth base of the RNA, thus positioning the RNA at the active site and enabling efficient cleavage (Hartung et al. 2010). Although the main active side residues are located in the Rrp41 protein, some residues (like the above-mentioned tyrosine) from the Rrp42 protein are important for RNA binding and influence degradation efficiency and processivity. All crystal structures of exosomes with RNA have one important aspect in common: even in the presence of longer RNA substrates, not more than 4–6 bases can be observed directly in the active sites, with clear electron density for an additional nucleotide in the neck region of the exosome ring. The conformation of all nucleotides in between is too flexible to be determined by crystallography. This observation raises the question whether (and if so, how) the three active sites within the processing chamber interact: does the 30 -end of the RNA molecule, after entering the processing chamber, bind to a single active site for degradation of the whole molecule? Or could RNA, while being fixed at the narrow neck, stochastically or in an ordered manner switch between the three active sites? The question still remains unanswered.

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The molecular mechanism of RNA polymerization and degradation has been extensively studied for Escherichia coli PNPase. Here, the detection of heteropolymeric tails after inactivation of the poly(A)-polymerase gene led to the identification of PNPase as the responsible enzyme (Mohanty and Kushner 2000). In accordance with these experiments it was also shown for other organisms, including plant chloroplasts, that PNPase in bacteria/eukaryotic organelles and the exosome in archaea are the only enzymes that produce heteropolymeric tails (Rott et al. 2003; Sohlberg et al. 2003; Yehudai-Resheff et al. 2003).

9.4.2

RNA Recognition, S1, ZnR, and KH Domains

Although the architecture of the processing chamber ensures that no RNA molecule with secondary structure can be degraded, there must be an additional regulatory mechanism directing some RNA molecules to the exosome for degradation while protecting others. Obvious candidates for some targeting are the Csl4 and Rrp4type subunits. It was indeed shown for the archaeal exosome that Rrp4 and Csl4 greatly increase the RNA degradation activity, whereas they are less important for efficient polymerization (Evguenieva-Hackenberg et al. 2008). Accessory factors like helicases can direct substrates to the exosome, but this is not the only mode of substrate selection. The genetic organization of the four archaeal exosome genes already indicated a possible regulatory function of the cap proteins: the genes rrp41, rrp42, and rrp4 are located in one operon whereas csl4 is under the control of a different promoter. Different promoters allow for specific expression of the Csl4 cap protein independent from the rest of the exosome. The composition of exosome caps can be varied based on expression levels, ranging from three Rrp4 proteins, via mixed complexes to three Csl4 proteins. Archaeal exosomes can be isolated as either Rrp4-only or Csl4-only complexes. Availability of the two different exosomes allows for the determination of different substrate specificity of the cap proteins: detailed kinetic studies on the degradation of poly(A) RNA showed clear differences between the mode and speed of degradation dependant on the cap proteins (Hartung et al. 2010; Niederberger et al. 2011). Additionally, the preference for various RNA substrates was examined and the exosome with Rrp4 was shown to degrade poly(A) RNA much better than heteropolymeric sequences, whereas exosomes with Csl4 prefer RNA with A-poor sequences. This hints to a possible role of Rrp4 in degradation of RNAs that are targeted by eukaryotic poly(A) polymerases such as Trf4/5.

9.4.2.1

Channeling and Processivity

Structural studies on different archaeal exosomes in complex with RNA led to the view that RNA binds to the RNA-binding domains of the cap proteins and the 30 -end is subsequently threaded through the narrow pore into the processing

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chamber and toward the active site (Fig. 9.2); this mechanism of binding was therefore named “channeling.” Mutagenesis experiments with bacterial PNPase suggest a similar channeling mechanism for RNA (Shi et al. 2008). The ring architecture of the exosome and the channeling mechanism seem to have advantages and disadvantages. Through arginine side chains in the narrow pore region, RNA is fixed to the exosome and processivity is ensured. This mechanism is also true for the eukaryotic exosome that uses a completely different protein as the catalytic component, but still uses the processing chamber as tong to hold on to the RNA (Bonneau et al. 2009). Still this architecture seems to hinder substrate binding: the 30 -end of a long RNA molecule has to thread through this narrow hole to reach into the processing chamber. This mechanism raises the questions: how the exosome can be such an efficient machine and how does the RNA find the correct path? There are two aspects to these questions. First, time-dependent experiments show that the rate for binding of the 30 -end of RNA to the active site of the exosome is considerably lower than the degradation rate (Hartung et al. 2010). This means that binding is slow, but the cleavage rate is dramatically high, so that altogether the exosome is still a very efficient machine. Second, comparison of various exosome complexes (with or without different caps; crosslinked or forced dimeric Rrp41/ Rrp42 complexes) led to the conclusion that the 6- and 9-subunit assembly of the exosome is variable, the “ring is breathing.” Without RNA, the three arginines in the neck region are not fixed, the diameter of ring increases, and a wider opening facilitates RNA threading. Only after the negatively charged RNA is bound does the neck tighten, and the exosome remains bound to its substrate until it is degraded.

9.4.3

Eukaryotic Exosomes

9.4.3.1

Loss of Phosphorolytic Active Sites

Based on structural, biochemical, and genetic work it is clear that the 9-subunit eukaryotic core exosome contains no enzymatically active center in human cells and yeast (Dziembowski et al. 2007; Liu et al. 2006) and that the two main proteins that are present as part of the eukaryotic complex must be responsible for the enzymatic activity in eukaryotes: Rrp44/DIS3 and Rrp6. In contrast to yeast and human exosomes, the Rrp41 protein from plant exosomes possibly retained its catalytic competence because phosphorolytic RNase activity could be detected for purified Rrp41 in vitro (Chekanova et al. 2000). Perhaps plant exosomes are evolutionary intermediates between the archaeal homolog and the yeast and human exosomes. Further research on plant exosomes confirmed this observation: First, Rrp44 could not be found as part of the exosome complex and second, Csl4 (in contrast to yeast) is not essential for viability and deletion of the csl4 gene affected only a very small subset of target RNAs (Chekanova et al. 2007).

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Two Different Hydrolytic Activities in Rrp44

The yeast complex and its corresponding enzymatically active subunit Rrp44 is from a biochemical standpoint the most intensively characterized eukaryotic exosome. Rrp44/Dis3 is homologous to the bacterial RNase II, a hydrolytic RNAdegrading enzyme (Fig. 9.3), and biochemical characterization of the isolated Rrp44 showed similar activities for the isolated and the exosome-bound enzyme (Dziembowski et al. 2007). This however is only true for single-stranded, unfolded RNA molecules. Rrp44 alone can unwind and degrade substrates with secondary structures, but this activity is strongly inhibited by the presence of the 9-subunit exosome complex (Bonneau et al. 2009; Lorentzen et al. 2008). RNase protection assays showed that isolated Rrp44 binds to a stretch of 9–12 nucleotides, whereas the 10-subunit exosome complex including Rrp44 binds 31–33 nucleotides. This is equivalent to the distance from the active site of Rrp44, through its substrate channel, the processing chamber of the exosome and to the neck region between the cap proteins. Only shortly after those discoveries, a new feature of Rrp44 was detected that was both surprising and intriguing: after the bacterial and the archaeal exosomes as well as Rrp44 were described as exonucleases, an additional endoribonucleolytic activity, mediated by a second domain of Rrp44, the PIN domain, was identified (Lebreton et al. 2008). Only the inactivation of both active sites within Rrp44 results in a synthetic growth phenotype, indicating that both activities have a physiological role. The cleavage of certain natural substrates could be assigned to this endonuclease activity (Schneider et al. 2009). Structural and mutagenesis studies helped to indentify the binding site of Rrp44 on the exosome (Bonneau et al. 2009): matching biochemical data already available, the RNA entry site of Rrp44 co-localizes with the RNA exit site of the processing chamber (Fig. 9.2a).

9.4.3.3

Hydrolytic Activities of Rrp6

The central part of Rrp6 is homologous to bacterial RNase D, which consists of an exonuclease and one or more helicase and RNase D C-terminal (HRDC) domains. Those RNases are characterized by at least four conserved acidic residues (DEDD) in the active site and were shown to require two divalent metal ions for the activation of a water molecule that attacks the last phosphodiester bond, resulting in 30 -to-50 exonucleolytic cleavage (Steitz and Steitz 1993). The eukaryotic RNaseD enzymes evolved from the bacterial homologs and have almost twice the size. The additional N-terminal domain of yeast Rrp6 covers the catalytic core and creates an interaction surface for the RNA-binding protein Rrp47 (Stead et al. 2007). The atomic structure of yeast Rrp6 shows the position of the two metal ions (Fig. 9.3), the specific ribonucleotide recognition mechanism and parts of the eukaryotic specific N-terminal extension (Midtgaard et al. 2006). The activating effect of the TRAMP complex on exosome activity in the nucleus could be ascribed to the hydrolytic activity from Rrp6, and is independent of Rrp44. Since the activating effect is

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not ATP dependent, neither the poly(A) polymerase activity of Trf4 nor the helicase activity of Mtr4 seem to be responsible (Callahan and Butler 2010). But why does the nuclear exosome need two different hydrolytic active sites? It can be speculated that in addition to specificity for certain substrates, accessibility might be a reason. From structural studies on Rrp44 we know that the active site is deeply buried in the protein and the 30 -tail of a structured RNA molecule cannot be degraded completely. No structural information on full-length Rrp6 is available; however, the solved structure of an Rrp6 fragment suggests a more surface-exposed active site, and thus Rrp6 could trim RNA overhangs for maturation in a way that Rrp44 is not capable of.

9.4.3.4

RNA Recognition and Channeling

Purification of stable human exosomes with fewer than 9 subunits has not been successful to date, and biochemical characterization of the cap proteins is therefore challenging (Liu et al. 2006). Recently, yeast exosomes could be purified as 7-subunit complexes containing only one of the cap proteins, and as 8-subunit complexes with two different caps (Malet et al. 2010). This is a very important step toward a better understanding of substrate selection and specificity of those proteins. RNA protection assays with all exosome variants determined a critical role for Rrp4 in the formation of an exosome-RNA complex and identified Rrp40 as the only cap protein that can alone form a stable RNase PH-like exosome. It seems that Rrp40 binds to the RNase PH-like ring in multiple copies, producing a structure more similar to the archaeal exosome. The channeling of RNA through the eukaryotic exosome could also help to select for RNAs with unfolded 30 -ends over folded RNAs or RNPs, explaining how poly/oligo(A) tails are an important structural feature for unstable RNAs. The RNA is believed to bind to the cap proteins Rrp4/Rrp40/Csl4, followed by a threading through the central channel of the RNase PH-like ring to the active site of Rrp44 (Fig. 9.2) (Malet et al. 2010). This mechanism not only maintains the roles of cap proteins and exosome ring-structure, but also protects the Rrp44 active site from solvent and regulates its enzymatic activity. In summary, RNA channeling seems to be an evolutionarily conserved mechanism for RNA degrading machines in all kingdoms of life.

9.5 9.5.1

Concluding Remarks Evolutionary Considerations

The conservation of exosomes in eukaryotes and many archaea, and the relationship to degradosomes in bacteria and organelles establish exosome-like complexes

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as quite ancient RNA degradation and polymerization machineries. Activities associated with eukaryotic exosomes and prokaryotic degradosomes show related overall mechanisms and pathways in RNA decay. These include the addition of oligo/poly-(A) tails to the 30 end of RNA molecules for stabilization (long tails in eukaryotes) or degradation (short tails in eukaryotes and tails in prokaryotes), endonucleolytic cleavage and processive 30 degradation. Nevertheless, it is remarkable that the exosome lost phosphorolytic activity in more complex organisms. In terms of cell physiology, a phosphorolytic enzyme may be viewed as a mediator between a free NDP pool and RNA polymers, and phosphorolytic activity is probably quite efficient in terms of energy conservation. Thus, it could be of substantial benefit to degrade RNA phosphorolytically, especially for fast growing organisms such as bacterial, or organisms in scant or extreme environments such as archaea. On the other hand, phosphorolytic degradation and polymerization is readily reversible and the cell might have little means to specifically control degradation versus polymerization although the reversible reaction could also be a simple means of regulation of mRNA stability based on the physiological state of the cell. It is therefore quite intuitive that in eukaryotes the phosphorolytic degradation and polymerization activities of the core exosome have been inactivated and mechanistically separated into two irreversible, highly specialized, and regulatable activities in the form of hydrolytic nucleases and poly(A) polymerases.

9.5.2

Future Questions

Despite the tremendous progress in the field over the past decade with respect to both pathways and structure, several important aspects still need to be addressed. From a structural and biochemical point of view, it is still unclear how substrates are chosen for degradation, what are the precise functions of Rrp4, Rrp40 and Csl4 caps in RNA recognition or interaction with other protein partners? How is the exosome targeted to the sites of RNA degradation, for instance in surveillance of nonfunctional RNA? How does it structurally or functionally interact with the Ski and Tramp complexes? What is the function of associated archaeal subunits such as DnaG? Is there also endonuclease activity associated with the archaeal exosome, and what are the functional equivalents of Ski2 and Mtr4 helicases in archaea? How are endo- and various exonuclease activities mechanistically coordinated? Finally, are there exosomes with both phosphorolytic and nucleolytic activities? There are of course many aspects to discover in the complex role of exosomes in eukaryotic and archaeal cell biology and we look forward to the next decade of exciting research on exosomes. Acknowledgment Work in KPHs laboratory on exosomes is supported by a grant from the German Research Council (DFG HO2489/3).

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Chapter 10

The Metallo-b-Lactamase Family of Ribonucleases Ciarań Condon and Laetitia Gilet

Contents 10.1 10.2 10.3 10.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . General Properties of Metallo-b-Lactamases Acting on Nucleic Acid . . . . . . . . . . . . . . . . The RNase Z Sub-family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The b-CASP Subfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.4.1 RNase J . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.4.2 CPSF-73 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.4.3 Int-11/RC-68 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.4.4 Archaeal and Bacterial Homologs of CPSF-73 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.5 Phylogeny of rMbLs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

246 248 250 253 253 259 260 261 262 264

Abstract The metallo-b-lactamases (MbLs) constitute a very ancient family of enzymes with a wide range of substrates, including RNA and DNA. The b-lactamases that act on RNA (here called rMbLs) can be grouped into two major families, the RNase Z family and the b-CASP family. Members of the RNase Z family are primarily involved in the maturation of the 30 end of tRNAs, whereas members of the b-CASP family have thus far been shown to have primarily mRNA and snRNA targets. Although they share a metallo-b-lactamase core and catalytic mechanism, the two families are easily distinguishable at the sequence level and by the presence of characteristic subdomains that play a key role in substrate recognition. These are principally the so-called flexible arm (about 40 amino-acids) of RNase Z and the b-CASP domain (>160 amino-acids) that gives its name to this family of enzymes. In this chapter, I will describe our current understanding of these two MbL families,

C. Condon (*) • L. Gilet CNRS UPR 9073 (affiliated with Universite´ de Paris Diderot, Sorbonne Paris Cite´), Institut de Biologie Physico-Chimique, 13 rue Pierre et Marie Curie, 75005 Paris, France e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_10, # Springer-Verlag Berlin Heidelberg 2011

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with particular emphasis on their structures, their substrates, and their phylogenic distribution. Keywords RNase J • RNase Z • CPSF • Int11/metallo-b-lactamase/b-CASP

10.1

Introduction

The metallo-b-lactamase (MbL) fold is a relatively recent addition to the list of domains found in proteins with ribonuclease (RNase) activity. Historically known for its role in the hydrolysis of the lactam rings in antibiotics of the penicillin family, the b-lactamase domain has been implicated in up to sixteen different types of hydrolysis reactions (Daiyasu et al. 2001). The MbL domain is thus a hydrolytic engine par excellence, with substrate specificity coming from the grafting of extra motifs or larger domains onto the catalytic core. As its name suggests, the MbL domain uses metal ions, often a pair of zinc ions, to promote hydrolysis. The proteins of this family have five signature motifs, consisting mostly of individual aspartic acid (motifs 1 and 4) and histidine residues (motifs 3 and 5), and a very recognizable histidine-motif HxHxDH (motif 2), which are involved both directly and indirectly in coordinating the metal ions and in the positioning of the substrate (Fig. 10.1a, b). The spatial positions of these key residues and the metal ions are highly superimposable in the members of the MbL family with RNase activity (here called rMbL) present in the Protein Data Base (PDB). The MbL hydrolysis reaction involves a general base (usually aspartate) that removes a proton from a neighboring water molecule, allowing the hydroxide ion to perform a nucleophilic attack on the substrate (often an ester linkage with a negative charge), polarized by the metal ions (Fig. 10.1c). Once the scissile bond is cleaved, the product is stabilized by taking a proton from a second adjacent water molecule. A role for the MbL domain in nucleic acid metabolism was first recognized by Aravind (Aravind 1999). An exhaustive search for members of the b-lactamase family identified a subgroup of these proteins known to be implicated in DNA repair (SNM1/PSO2) and RNA cleavage reactions (CPSF-73). This group of proteins was extended to include Artemis, involved in V(D)J recombination/DNA repair, and called the b-CASP subfamily (Callebaut et al. 2002). These proteins are characterized by an insertion of a large b-CASP domain (>160 amino acids) into a loop of the core b-lactamase fold (Figs. 10.2 and 10.3). Three extra motifs, A (Asp or Glu), B (His), and C (His), were proposed to characterize proteins of the b-CASP family. However, the resolution of the structures of some representatives of this family has made it clear that motif C and motif 5 are the same histidine residue in the b-CASP proteins that act on RNA. A second subfamily of b-lactamases involved in nucleic acid metabolism is defined by the RNase Z/ELAC group of enzymes. These proteins are primarily involved in transfer RNA (tRNA) maturation (for review see Redko et al. 2007) and lack the b-CASP domain. Rather, they have a characteristic domain called the

10

The Metallo-b-Lactamase Family of Ribonucleases

a (B) His247

247

b

(A) Glu231

(A) Asp204

(B) His376

(5) His398

(5) His269 UMP

(4) Asp211 (3) His140 Zn

Zn (2) His80

(3) His150 Zn

(2) His75 (2) His77

(2) Asp67

B. subtilis RNase Z (2FK6)

c

(1) Asp37

(1) Asp37

(2) His63 (2) His65

(4) Asp172 UMP

Zn (2) His68

T. thermophilus RNase J (3BK2)

WAT5 –

O

nt73

H+

H

(2) Asp79

O

P O–

O–

His269

His140

O nt74 Zn1

His68

O

O– – O

O

H+

His65

Zn2

Asp211

His63

H WAT147

Asp67

Fig. 10.1 Active site and cleavage mechanism of rMbLs. (a) Active site of Bacillus. subtilis RNase Z showing the relative positions of the amino acids of motifs 1–5 and motifs A and B. The position of the 30 -nucleotide of the tRNA cleavage product is shown. (b) Active site of Thermus thermophilus RNase J showing the relative positions of motifs 1–5 and motifs A and B. The position of bound UMP is shown. (c) Cleavage mechanism of metallo-b-lactamases, as represented by B. subtilis RNase Z. Asp67 removes a proton from water molecule 147. The resulting hydroxide ion performs a nucleophilic attack on the scissile phosphodiester bond between nt 73 and nt 74 of the tRNA precursor. The cleaved bond is stabilized by taking a proton from water molecule 5

flexible arm (also called the exosite) for substrate recognition, fused to a different loop of the catalytic core (Figs. 10.2 and 10.3). Although they lack the b-CASP domain, RNase Z enzymes have the signature Asp/Glu and His residues of motifs A and B at the equivalent positions in their 3D-structures. Motifs A and B are therefore not restricted to the b-CASP proteins, but are a general feature of b-lactamases that have nucleic acid substrates.

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a RNase Z *

flexible arm

metallo-βlactamase

Short form ( e.g. bacterial RNase Z, human ELAC1) *

metallo-βlactamase

metallo-βlactamase

Long form ( e.g. human ELAC2)

b β-CASP *

metallo-βlactamase

β-CASP

C-ter

Class 1 ( e.g. eukaryotic CPSF-73, Int11, bacterial RNase J)

*

metallo-βlactamase

β-CASP

Class 2 (e.g. archaeal RNase J, Tth-A0252, EF-2904)

β-CASP

Class 3 (e.g. Ph-1404, Mm-0695)

*

N-ter

metallo-βlactamase

Fig. 10.2 Domain structure of rMbLs. The motif 2 histidine signature (HxHxDH) of the active site is indicated by an asterisk. (a) Domain structures of the short and long forms of the RNase Z family. The location of the characteristic of the flexible arm is shown. In the long form of the enzyme, only the N-terminal domain contains a flexible arm, while the C-terminal domain contains the active site. (b) Three different classes of members of the b-CASP family. The position of the b-CASP insertion is shown. Class 1 represents the simplest form of this family of enzymes, containing just the core b-lactamase and b-CASP domains. Class 2 enzymes have a C-terminal extension such as that found in eukaryotic CPSF-73, CPSF100, Int11, and bacterial RNase J. A further insertion in the b-CASP domain is seen in CPSF100, indicated by the arrowhead. Class 3 proteins have an N-terminal extension. This corresponds to two KH domains in some archaeal forms of CPSF. A different N-terminal extension, whose function is not yet known, is found in some bacterial forms of RNase J (e.g., Helicobacter pylori)

10.2

General Properties of Metallo-b-Lactamases Acting on Nucleic Acid

Although historically classified as either endo- or exonucleases, the DNases and RNases of the MbL family often have both types of cleavage activities within the same polypeptide (Dutta and Deutscher 2009a; Yang et al. 2009) and, where examined, catalyzed by the same catalytic site (Li de la Sierra-Gallay et al. 2008). In some cases, the preference for one type of activity over the other can be regulated: binding of the autophosphorylated catalytic subunit of DNA-PK to Artemis, for example, shifts the balance from 50 -to-30 exonuclease to endonuclease activity (Goodarzi et al. 2006). The exonuclease activity of the b-CASP subfamily, whether DNase or RNase, is primarily oriented 50 -to-30 , whereas the only member of the RNase Z subfamily examined in this regard, Escherichia coli RNase Z, can degrade exonucleolytically in a 30 -to-50 orientation (Dutta and Deutscher 2009a, b). The RNase enzymes have some DNase activity and vice versa (our unpublished data for Bacillus subtilis RNase J1 and see (Ma et al. 2002; Clouet-d’Orval et al. 2010)), indicative of a relatively recent separation of these enzymes with regard to their

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C

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b

N

β14

β2 β1

α1

β3

α2

β4

β5

N β14

α3

β6

β2 β1

βB

M2 63-HMHGDH-68

M1 D37

α1

β3

α2

β4

M1 D38

βA

β9

β7 β8

α4

M3 H140

α5 β12

A E231 B H247

M4 D211

α4

β8

β13

αC

β9

β7

α6

β11

β10

M3 H150

M5 H269

βG

αD αE

βF βD

β10

β6

C βE

βC

Flexible Arm βB

α3

M2 75-HGHEDH-80

βA αA αB

β5

α5 β11

A

M4 D172

α6 β12

β13

M5 H398

B

D204 H376

αC

B. subtilis RNase Z β-CASP αC1

βC1

βC2 βC6 βC3 βC5 αC3 αC4 αC5

αC2

αCA

T. thermophilus RNase J

c

d

C-ter 227 aa

αB

αB

N β14

N β2

β1

α1

β3

β4

M1 D40

α2

β5

β14

α3

β6

β2 β1

M2 71-HFHLDH-76

β9 β8

β-CASP αC1

α4

M4 D179

βC1

β5

α3

β6

M2 59-HFHLDH-64

αA

β10

M3 H158

α2

β4

M1 D29

αA

β7

α1

β3

αC2

α5 β11

α6 β12

M5 H419

A B E204 H397

βC2

βC3

αC4

βC5

β9

β7 β13

β8 β

M3 H141

αC5

βC6

α5 β10 β

β-CASP αC1

M4 D162

βC1

αC2

β11 β

α6 β12 β

A B E186 H380

β13 β

M5 H400

βC2 βC6 βC3 βC5 αC3 αC4 αC5

αCE αCD

H. sapiens CPSF-73

αCE αCA

αCB αCC

αCD

T. thermophilus A0252

Fig. 10.3 Secondary structure topology of four representative rMbLs. Helices (a) are represented by rectangles, strands (b) by arrows. The secondary structure features of the core b-lactamase and core b-CASP domains are identified with numbers (a1, b1, . . . for the core MbL domain and aC1,

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substrate preference. However, whether the dual substrate specificities observed today are any more than an in vitro curiosity remains to be seen. Like many other members of the broader b-lactamase family, the proteins of ribonuclease subfamilies often form multimers (mostly dimers and tetramers). Multimerization can be through self-association or association with paralogous proteins. Interestingly, in cases where the enzyme associates with a paralogous protein, this protein often lacks some of the key residues for catalysis and is inactive. In one interesting variation on this theme, the so-called long form of RNase Z found in some eukaryotes, it is clear that two b-lactamase domains are fused together in a single polypeptide, but with only one of the domains containing the residues necessary for catalysis (Tavtigian et al. 2001). The crystal structures of representative members of each family have been solved (three RNase Z enzymes and seven b-CASP proteins to date). These enzymes all contain the classic b-lactamase fold consisting of a pair of b-sheets sandwiched between a-helices (an abba sandwich). While some b-lactamases have just six b-strands to a b-sheet, the RNases are among those b-lactamases with at least seven b-strands per sheet. In this chapter, I will describe our current understanding of these two subfamilies of proteins with (often essential) ribonuclease activity, with particular emphasis on those enzymes whose structures have been solved.

10.3

The RNase Z Sub-family

RNase Z was first identified in eukaryotes (Schiffer et al. 2002) and is known by multiple pseudonyms including tRNase Z, 30 tRNase, ZiPD, RNase BN, and ELAC. It is a ubiquitous and often essential enzyme that catalyses the ribonucleolytic maturation of the 30 end of tRNAs (for review see Hartmann et al. 2009). Two forms of the enzyme exist in nature. The so-called short form is found in both prokaryotes and eukaryotes and is usually a dimer in solution. The long form is found only in eukaryotes, is monomeric and consists of two fused b-lactamase domains in a single polypeptide. Mutations in the long form ELAC2 gene have been linked to prostate cancer in humans (Tavtigian et al. 2001). The RNase Z cleavage reaction is generally endonucleolytic and occurs after the discriminator nucleotide in most cases. The discriminator nucleotide is the unpaired nucleotide at the 30 end of the acceptor stem to which the universal CCA motif is added by nucleotidyl transferase. Exceptions to the cleavage mode and site are known, however. Although it acts primarily as an endoribonuclease in the presence Fig. 10.3 (continued) bC1, . . . for the core b-CASP domain). Insertions in either core domain are identified with letters (aA, bA, . . . for the core MbL domain and aCA, bCA, . . . for the core b-CASP domain). The nomenclature is kept consistent between members of the same family. The identities and positions of the key motifs (M1 to M5 and A, B) are indicated by stars. The N and C-termini are indicated as (N) and (C), respectively. Gaps in the trace indicate that these regions were not seen in the crystallographic structure and therefore their secondary structure is uncertain

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of physiologically relevant concentrations of divalent cations such as magnesium, E. coli RNase Z exhibits 30 -to-50 exonucleolytic activity in the presence of relatively high concentrations (200 mM) of cobalt (Dutta and Deutscher 2009a, b). Secondly, both Thermotoga maritima and E. coli RNase Z can cleave tRNAs endonucleolytically downstream of the CCA motif (Minagawa et al. 2004; Dutta and Deutscher 2009b). In most other organisms, the CCA motif acts as an antideterminant for RNase Z, preventing futile cycles of CCA addition and removal. The first crystallized rMbLs were the RNase Z proteins from B. subtilis (PDB: 1Y44) and T. maritima (PDB: 1WW1; 2E7Y) (Li de la Sierra-Gallay et al. 2005; Ishii et al. 2005). The structure of the E. coli enzyme was solved shortly thereafter (PDB: 2CBN) (Kostelecky et al. 2006). The general topology of representative rMbLs is shown in Fig 10.3. The MbL core consists of two opposing sevenstranded b-sheets wedged between three a-helices on either side, in a bbbbababab configuration. The first four b-strands of each sheet are antiparallel and the last three oriented parallel to each other. The flexible arm, which plays an important role in holding the tRNA in position, is inserted between the third and fourth strands (b9 and b10 in Fig. 10.3) of the second b-sheet. A second insertion consisting of just one a-helix (aC) is found after the fifth b-strand (b11) in the same sheet. Very interestingly, this is also where the b-CASP domain is inserted in the b-CASP family of proteins. RNase Z is known to be a dimer in solution (or a monomer consisting of two fused b-lactamase domains) and the asymmetric unit in the crystal structure of the B. subtilis enzyme is a dimer. The two subunits of RNase Z contact each other primarily through the first three a-helices (a1, 2 and 3). The structure of the B. subtilis enzyme has also been solved in complex with its tRNA cleavage product (PDB: 2FK6), giving the clearest picture yet of how this enzyme recognizes and cleaves its tRNA substrates (Li de la Sierra-Gallay et al. 2006). The tRNA is bound principally by one subunit and cleaved by the other (Fig. 10.4). It is clamped in place between the flexible arm and the protruding aChelix, resembling a ski-boot in its bindings (Redko et al. 2007). This is a wonderful example of how the grafting of extra domains to the b-lactamase core affords substrate recognition. In fact, RNase Z mutants lacking the flexible arm can still hydrolyze small substrates, such as bis-(p-nitrophenyl) phosphate (bpNPP), but not tRNA (Schilling et al. 2005). Two types classes of flexible arm have been described, a glycine/proline-rich arm found in E. coli, B. subtilis, and Methanococcus jannaschii and a flexible arm found in T. maritima in which the glycine/proline motif is replaced by a cluster of basic amino acids (Ishii et al. 2007). In the case of B. subtilis RNase Z, the protruding aC-helix has also been proposed to play a role of sensor that initiates a cascade of conformational changes ultimately leading to the correct positioning of the residues involved in zinc coordination and may account for the sigmoidal reaction kinetics of this enzyme (Li de la Sierra-Gallay et al. 2006). It is not clear whether this type of substrate activation is a common property of all short-form RNase Z enzymes, however. The tRNA is recognized via its stacked T-arm and acceptor stem; the D-arm and anti-codon loops do not make contact with the enzyme and indeed were not visible in the co-crystal structure (Li de la Sierra-Gallay et al. 2006). As expected, most of the

252


tRNA flexible arm

acceptor stem

β-lactamase domain

T-arm Zn++

β-lactamase domain α C-helix

B. subtilis RNase Z (2FK6)

Fig. 10.4 Crystallographic structure of Bacillus. subtilis RNase Z bound to tRNA. The structure is shown in its dimeric conformation. The anticodon loop of the tRNA was not visible in the crystal structure. The tRNA is clamped between the flexible arm and helix aC as shown. A mutation in the motif 2 histidine signature was made to favor substrate binding and results in the presence of only one zinc ion in the catalytic site

tRNA contacts with the enzyme are through the sugar-phosphate backbone, to accommodate the large variation in tRNA sequences. In the few cases where contacts with the bases were seen (e.g., G1 and G19), these were with highly conserved residues. Although the primary role of RNase Z is thought to be tRNA maturation, other substrates have been proposed for the enzyme. It is not immediately clear why RNase Z is present in some enterobacteria such as E. coli and Salmonella, for example, where all of the tRNAs have an encoded CCA motif, but is generally absent from other Proteobacteria. Although RNase Z can ensure tRNA maturation and cell viability in E. coli in the absence of the four major tRNA processing enzymes RNase T, RNase PH, RNase D, and RNase II (Li and Deutscher 1996), it seems unlikely that this is sufficient selective pressure to account for its presence in the E. coli genome. E. coli is a host for the phage T4, which does have some tRNAs lacking an encoded CCA on its genome and RNase Z/BN plays a key role in the maturation of these tRNAs (Seidman et al. 1975; Asha et al. 1983). Although intriguing, it seems unlikely that the enterobacteria possess RNase Z to allow the replication of T4-like phages unless there is some benefit to the host. A role in the turnover of about 150 E. coli mRNAs has been proposed by the Kushner laboratory (Perwez and Kushner 2006), but this was not seen by another group (Schilling et al. 2004) and the reason for the contradictory observations is not clear. In the Perwez et al. study (Perwez and Kushner 2006), the half-lives of some mRNAs was greater in strains lacking both RNase Z and RNase E than strains lacking either

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enzyme alone, suggesting some synergistic relationship between the two enzymes in mRNA decay. The nature of this relationship has not been elucidated, however. In addition to its role in tRNA 30 maturation in Archaea (Spath et al. 2008), RNase Z is involved in 5S rRNA processing in Haloferax volcanii (Holzle et al. 2008). The enzyme is thought to recognize a structure immediately upstream of 5S rRNA that mimics the T-arm and acceptor stem of a tRNA. Cleavage at the base of this mini-tRNA releases the mature 50 end of 5S rRNA. The long form of RNase Z can also accept non-tRNA substrates that mimic the stacked T-arm and acceptor stem of the tRNA (Schiffer et al. 2001). Recent publications have shown that RNase Z from several different organisms is also capable of cleaving unstructured RNA (Shibata et al. 2006; Dutta and Deutscher 2009a). However, these experiments were carried out at nonphysiological conditions (52 C or in the presence of high concentrations of cobalt). The Kd for unstructured RNA is orders of magnitude greater than that for tRNA precursors (Shibata et al. 2006) and therefore the relevance of these in vitro observations for RNase Z function in vivo remains to be determined.

10.4 10.4.1

The b-CASP Subfamily RNase J

RNase J1 was first identified in B. subtilis as an enzyme important for cleavage of the thrS leader mRNA and maturation of the 50 end of 16S ribosomal RNA (Even et al. 2005; Britton et al. 2007). Surprisingly, these two reactions employed different modes of enzyme activity, endonuclease and 50 -to-30 exonuclease activity, respectively (Mathy et al. 2007). Although DNases with dual activities were known in the MbL family of enzymes (Ma et al. 2002), this was the first evidence that such dual activity could also be found in a ribonuclease. The discovery that RNase J1 had 50 -to-30 exoribonuclease activity was also an important landmark in the field of bacterial RNA turnover, since up to that point, RNA degradation in the 50 -to-30 orientation was thought to be confined to eukaryotes, catalyzed by enzymes unrelated to RNase J1, namely, Xrn1 and Rat1. The 50 -to-30 exoribonucleolytic activity of RNase J1 is strongly inhibited by the 0 5 triphosphate group found on primary transcripts and is much more active on RNAs bearing 50 -monophosphate or 50 -OH groups (Mathy et al. 2007). The endonuclease activity appears to be independent of the phosphorylation status of the 50 end, however (Even et al. 2005; Li de la Sierra-Gallay et al. 2008). In vitro, the endonuclease activity can be relatively non-specific, with a clear preference for single-stranded conformation and this property can actually be exploited to determine RNA secondary structure (Daou-Chabo and Condon 2009). Clearly, this type of activity is not consistent with the enzyme’s function in vivo, however.

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Although RNase J1 is essential in B. subtilis, this organism also has a non-essential paralog called RNase J2 that is 49% identical to RNase J1. Recent evidence has shown that these two enzymes form a complex (a dimer of heterodimers) and are unlikely to exist as individual enzymes in wild-type cells (Mathy et al. 2010). While the individual enzymes have equivalent endonucleolytic activity in vitro, the RNase J1/J2 complex has altered cleavage site specificity. Thus the two enzymes behave synergistically to affect cleavage activity and choice of cleavage site. While RNase J2 has equivalent endonucleolytic activity to RNase J1 in vitro, its 50 -to-30 exoribonuclease is about two orders of magnitude weaker than that of RNase J1 and this presumably explains why RNase J1 is essential in B. subtilis. The essential nature of these proteins varies in other organisms, however. In group A Streptococci, both RNase J1 and J2 are essential (Bugrysheva and Scott 2010), while in Sinorhizobium meliloti, a deletion of the gene encoding the single RNase J ortholog is viable (Madhugiri and Evguenieva-Hackenberg 2009). B. subtilis RNase J1 has been localized to the ribosome (Hunt et al. 2006) presumably reflecting its role in 16S rRNA processing (Britton et al. 2007). Its cellular localization has not yet been determined in other organisms. It is not clear why RNase J2 has very weak 50 -to-30 exoribonuclease activity. Many RNase J2 orthologs have nonconsensus motif 2 histidine signatures (Fig. 10.5a) and are predicted to be unable to properly coordinate zinc in the catalytic site. However, rather than restore 50 -to-30 exoribonuclease activity of RNase J2, mutation of the RNase J2 motif 2 (HGHDEN) to that found in RNase J1 (HGHEDH) actually abolishes the enzyme’s residual activity (Fig. 10.5b, c). RNase J2 activity cannot be recovered by forming a complex with RNase J1; isolation of complexes containing wild-type RNase J2 and catalytically inactive RNase J1 show the same residual levels of activity as RNase J2 alone. These results suggest RNase J2 plays only a supporting role in RNase J1’s exonucleolytic mode of action. Since RNase J2 is a far more basic protein than RNase J1 (pI 9.2 compared to 5.9), this role could be related to substrate binding. The extent of the role of RNase J1/J2 in mRNA turnover in B. subtilis seems to depend on how the experiment is performed. A global analysis showed that reduced levels (about 9-fold) of RNase J1 or lack of RNase J2 had little effect by themselves on the B. subtilis transcriptome, whereas a combination of the two mutations led altered expression of hundreds of genes (Mader et al. 2008). On the other hand, when individual transcripts have been examined under conditions of a more severe RNase J1 depletion, inactivation of RNase J2 has little additional effect (Britton et al. 2007; Daou-Chabo et al. 2009). The interpretations of the data are different in these two cases. In the first case, the conclusion is that RNase J1 and J2 are redundant enzymes with overlapping function; in the second, RNase J1 plays the major role. Further experiments are needed to clear up this ambiguity. The crystal structure of RNase J from Thermus thermophilus has been solved (PDB 3BK2) (Li de la Sierra-Gallay et al. 2008). The asymmetric unit is a monomer (Fig. 10.6b), but dimers and tetramers can be built by crystallographic symmetry, consistent with forms seen in vitro (Mathy et al. 2010). Intriguingly, neither the dimerization nor tetramerization surfaces are the same as those found in RNase Z.

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a

b

c

Fig. 10.5 Neither binding to RNase J1 nor mutation of the catalytic site to consensus suffices to activate the 50 -to-30 exoribonuclease activity of RNase J2. (a) The motif 2 histidine signature of RNase J2 from different Gram-positive bacteria. (b) 50 -to-30 exoribonuclease assay of wild-type and mutant (H76A) RNase J1/J2 complexes and RNase J2 mutated to resemble motif 2 of RNase J1 (D77E, E78D, N79H). RNase J1/J2 complexes and RNase J2 protein were purified and assayed as described in (Mathy et al. 2010) using the 50 -labeled substrate shown. (c) Quantification of the rate of release of UMP in the gels shown in (b). Only the wild-type RNase J1/J2 and RNase J1/J2 (H76A) complexes show significant enzyme activity

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Each monomer of RNase J has a core MbL domain, with two inserted cassettes, and a C-terminal addition (Fig. 10.3). The first insertion is a single b-strand (bA) between strand b3 and helix a1. This strand forms part of an antiparallel b-sheet with two strands from the C-terminal extension. The second insertion is the b-CASP domain itself, which occurs between helix a5 and strand b11, the same site as the helix aC insertion in RNase Z. Two of the signature amino acids that play a key role in the catalytic reaction (motifs A and B) flank this insertion in both enzymes. The b-CASP domain of RNase J is a large globular domain, consisting of five b-strands surrounded by five a-helices. In related proteins of this family, additional secondary structure features are inserted into the b-CASP domain (Fig. 10.3), presumably bringing extra functions to the enzyme. Whereas the catalytic site of RNase Z is at the surface of the protein, the RNase J catalytic site is deep in a cleft between the b-lactamase and the b-CASP domain (Fig. 10.6). In the solved structure, this cleft is too narrow to allow passage of a strand of RNA and can therefore be considered in a “closed” conformation. Separation of the b-lactamase and b-CASP domains is required to open the cleft for RNA binding. The structure of RNase J has also been solved in complex with UMP, also in a closed conformation (Li de la Sierra-Gallay et al. 2008). The phosphate moiety of UMP is found in a phosphate-binding pocket located about one nucleotide distant from the catalytic zinc ions. This nicely explains the enzyme’s 50 end preference in 50 -to-30 exonucleolytic mode. A triphosphorylated species would not fit, or would otherwise place the scissile bond out of phase with the catalytic site. A 50 -OH is acceptable, but a 50 monophosphate is optimal. The structure also suggests how the enzyme works as a 50 -to-30 exoribonuclease: after the 50 nucleotide is cleaved and ejected (by a mechanism that is not yet clear, but that likely involves rotation of the nucleotide), the shortened substrate is moved into the phosphate-binding pocket for the next cleavage reaction. Although the structure does not shed any light on the endonucleolytic cleavage mechanism, it does suggest how the 50 monophosphorylated products of endonucleolytic cleavage could immediately become substrates for the 50 -to-30 exonucleolytic degradation. This type of “cut and run” mechanism has been proposed for the turnover of the trp leader mRNA (Deikus et al. 2008). The globular C-terminal domain is attached to a flexible linker. In the dimeric structure, the linkers of the two monomers cross each other, such that the C-terminal domain of one monomer is located opposite the main body of the other. This may explain why the RNase J1/J2 complex apparently cannot be assembled from individual components, but requires coexpression in the same cell (Condon, unpublished). Intriguingly, the overall shape and charge distribution of the RNase J monomer from T. thermophilus resembles that of E. coli RNase E (Li de la SierraGallay et al. 2008). Deletion of the C-terminal domain of B. subtilis RNase J1 abolishes endonuclease activity and dramatically reduces exonuclease activity (Li de la Sierra-Gallay et al. 2008). However, the protein no longer forms multimers, consistent with the structural data that shows the C-terminus accounts for about two thirds of the dimerization interface. For the moment, therefore, it is not possible to determine

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b b-CASP

C-terminal domain

α-helix Zn++ UMP

Zn++

β-lactamase domain flexible arm

β-lactamase domain

B. subtilis RNase Z (1Y44)

c

d

β-CASP

T. thermophilus RNase J (3BK2)

e

β-CASP

β-CASP

Zn++

Zn++

β-lactamase domain H. sapiens CPSF-73 (2I7T)

f

β-lactamase domain

β-lactamase domain

S. cerevisiae CPSF-100 (2I7X) T. thermophilus Tth-A0252 (3IEK)

g

β-CASP

β-CASP

90°

Zn++

KH2

RNA

β-lactamase domain

KH1 P. horikoshi PH-1404 (3AF6)

Zn++

β-lactamase domain E. faecalis EF-2904 (2AZ4)

Fig. 10.6 Structure of representative rMbLs in the Protein Data Base (PDB). (a) B. subtilis RNase Z; (b) T. thermophilus RNase J; (c) H. sapiens CPSF-73; (d) S. cerevisiae CPSF-100; (e) T. thermophilus Tth-0252; (f) P. horikoshi PH-1404; (g) E. faecalis EF-2904. All proteins are shown in the same orientation except that of Pyrococcus horikoshi Ph1404 (panel F), which has been rotated to better see the KH domains. PDB codes are given for each protein. The catalytic zinc ions are shown and ligands where available

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Fig. 10.7 Alignment of C-terminal domains of CPSF-73, Int11, and RNase J. The known secondary structure features of Thermus thermophilus (Tth) RNase J are shown on the last line of the alignment. Secondary structure features predicted by HHPRED are shown for the other enzymes. Positively charged amino acids are indicated in bold text. Organisms are as follows: Sau Staphylococcus aureus, Bsu Bacillus subtilis, Efa Enterococcus faecalis, h human, m mouse

whether the C-terminus plays a direct role in enzyme activity. It is intriguing to note however that the two a-helices of the C-terminal domain bear a high density of positively charged (mostly Arg and Lys) residues that could play a role in substrate binding (Fig. 10.7). In the dimer structure, the positively charged a-helices of the C-terminal domain of one subunit lie directly opposite the cleft leading to the active site of the other subunit. The archaeal form of RNase J lacks the C-terminal domain found in the bacterial protein. The activities of the RNase J orthologs from Pyrococcus abyssi and Thermococcus kodakaraensis have been tested in vitro (Clouet-d’Orval et al. 2010). Intriguingly, while both enzymes have 50 -to-30 exoribonuclease activity similar to bacterial RNase J, no endonucleolytic activity was detected. It is possible therefore that the C-terminal domain contributes to the endonuclease activity of this family of proteins. The archaeal RNase J has two extra conserved loops compared to the bacterial enzyme, both of which are important for enzyme activity (Clouet-d’Orval et al. 2010). Loop 1 creates a larger insertion just after motif 1 of the MbL core and loop 2 creates a larger insertion in the b-CASP domain between aC3 and aCA (Fig. 10.3b). Variants of RNase J can also be seen in bacteria. The Helicobacter

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pylori enzyme has an N-terminal extension of about 130 amino acids, for example, whose function is not yet known.

10.4.2

CPSF-73

The 30 ends of eukaryotic mRNAs are generated by endonucleolytic cleavage between a signature AAUAAA motif and a downstream G/U rich element (for reviews see Colgan and Manley 1997; Dominski 2007). Cleavage preferentially occurs at a CA motif 20–30 nucleotides downstream of the AAUAAA sequence. The upstream cleavage product is polyadenlylated and becomes the functional mRNA. The downstream cleavage product is a substrate for the nuclear 50 -to-30 exoribonuclease Rat1, which degrades the RNA until it catches up with RNA polymerase and causes it to terminate transcription (the so-called torpedo model (Connelly and Manley 1988; Proudfoot 1989)). Several subcomplexes combine to form a large protein complex that carries out the 30 processing reactions. One of the subcomplexes is called the cleavage and polyadenylation specificity factor, consisting of five proteins CPSF-30, -73, -100, -160, and Fip1 (Kaufmann et al. 2004). CPSF-73 is one of the founder members of the b-CASP family of MbLs (Callebaut et al. 2002) and although it was suspected of being the 30 processing endonuclease for some time (Aravind 1999), this was only recently shown directly (Ryan et al. 2004; Mandel et al. 2006). CPSF-73 forms a heterodimer with its paralog CPSF-100 (Dominski et al. 2005b), but CPSF-100 lacks some of the key residues for catalysis. This active/inactive subunit configuration is reminiscent of the RNase J1/J2 complex (Mathy et al. 2010) and is a recurring theme in b-lactamases. CPSF-73 is also part of the metazoan histone pre-mRNA 30 processing complex. Histone pre-mRNAs are cleaved downstream of a 30 stem-loop (30 -SL), but unlike other mRNAs, are not polyadenylated (for review see Dominski and Marzluff 1999). The cleavage reaction requires a purine-rich downstream element (HDE) that pairs with the 5’ end of U7 snRNA (Mowry and Steitz 1987). The fully assembled complex includes U7 snRNP, consisting of U7 snRNA bound to a heptameric SM/LSM ring (Pillai et al. 2003; Pillai et al. 2001), a zinc-finger protein called ZPF100 (Dominski et al. 2002), SLBP (Dominski et al. 1999) (which binds both the 30 -SL and ZPF100), symplekin (Kolev and Steitz 2005) (also found in the canonical mRNA processing complex) and CPSF-73/CPSF-100 (Kolev and Steitz 2005). Cleavage of the histone pre-mRNAs typically occurs 4–5 nucleotides downstream of the 30 -SL at a fixed distance from the HDE and, like that of the main class of mRNAs, occurs preferentially after a CA motif (Scharl and Steitz 1994). The upstream cleavage product corresponds to the mature histone mRNA and the downstream product is degraded by a 50 -to-30 exoribonuclease in a U7-dependent manner (Walther et al. 1998). UV cross-linking of CPSF-73 to both the upstream and downstream fragments suggested this enzyme is involved in both the endo- and exonucleolytic degradation reactions (Dominski et al. 2005a). This is highly

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reminiscent of the “cut and run” mechanism proposed for RNase J1 in the turnover of the trp leader mRNA (Deikus et al. 2008). The crystal structures of human CPSF-73 and yeast CPSF-100 have been solved (Fig. 10.6c, d; PDB: 2I7T and 2I7X, respectively) (Mandel et al. 2006). The C-terminal domains of each were removed to aid the crystallization process. As predicted from the amino acid sequence, CPSF-100 does not bind zinc and lacks enzyme activity. The core b-lactamase and b-CASP domains of CSPSF-73 and 100 are juxtaposed as in RNase J, with the catalytic site deep in the cleft between the two domains. This may explain why it is more difficult to strip the Zn ions from CPSF-73 than RNase Z upon addition of chelators such as EDTA. That the CPSF complex retained function in the presence of EDTA was the source of some discussion of whether the nuclease was metal dependent (e.g., Hirose and Manley 1997). Although the overall structures of the b-CASP domains of CPSF-73, CPSF-100, and RNase J are similar, CPSF-73 has one, and CPSF-100 two extra strands in the central b-sheet. Both CPSF proteins have a b-hairpin on the surface of the domain. The b-CASP domain from CPSF-100 is much larger than that of either CPSF-73 or RNase J, explained in part by the extra strand(s) in the central b-sheet and a much larger b-hairpin. The main difference, however, is a ~200 amino acid insertion consisting primarily of charged and hydrophilic residues that was removed by proteolysis during crystallization and whose structure is therefore not yet known. This insertion into the b-CASP domain accounts for most of the size difference between CPSF-73 and CPSF-100. Although the homology between the C-terminal domains of RNase J and CPSF73 is limited at the amino acid level, a secondary structure prediction of the C-terminal domain of CPSF-73 and Int11 (see below) compared to the known structure of RNase J, suggests they may be related (Fig. 10.7). The positively charged helices close to the C-terminal end are predicted to be particularly well conserved. Confirmation awaits resolution of the structure of the C-terminal domain of CPSF-73. Similar to its role in promoting the dimerization of RNase J, the C-terminal domain of CPSF-73 is required for dimerization with CPSF100 (Dominski et al. 2005b). However, its role in catalysis is not yet known.

10.4.3

Int-11/RC-68

A close paralog of CPSF-73, called Int11 (RC-68), is found in higher eukaryotes but is conspicuously absent in yeast (Dominski et al. 2005b). Int11 is a component of the so-called integrator complex (Baillat et al. 2005) involved in the maturation of small nuclear RNA (snRNAs) that play an important role in mRNA splicing. The snRNAs are transcribed by RNA polymerase II for the most part and, in vertebrates, processing requires a 30 stem-loop and a 13–16 nucleotide element called the 30 box located several nucleotides downstream of the mature 30 end (Hernandez 1985; Ach and Weiner 1987). Similar to 30 end processing of polyadenylated mRNAs and histone mRNAs, processing of snRNAs requires the assembly of a large complex.

10


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A key component of this complex, however, is the phosphorylated C-terminal domain of RNA polymerase II (Uguen and Murphy 2003), explaining why snRNAs are not efficiently processed when transcribed by other polymerases (de Vegvar et al. 1986; Hernandez and Weiner 1986). In addition to RNA polymerase II, the maturation complex includes the Integrator proteins 1–12, among which is the rMbL Int11 and its paralog Int9 (RC-74) (Baillat et al. 2005). Int11/RC-68 and Int9/RC-74 form a heterodimer in the now familiar pattern of one active, one inactive subunit (Dominski et al. 2005b). Although initially characterized for its role in the processing of the U1 and U2 snRNAs, it has been suggested that the integrator complex may play a much broader role in the processing of noncoding RNAs transcribed by PolII, such as the RNA component of telomerase and the individually transcribed small nucleolar RNAs (see Dominski 2007).

10.4.4

Archaeal and Bacterial Homologs of CPSF-73

b-CASP family members that are clearly more homologous to CPSF-73 than RNase J exist both in the bacteria and in the archaea, despite the fact that there is no evidence of an equivalent to the eukaryotic mRNA 30 processing pathway in these organisms. Although described as an RNase J ortholog (Hasenohrl et al. 2011), the recently characterized SSO0386 protein from the Crenarchaeum Sulfolobus solfataricus is more closely related to human CPSF-73 (26% identity/38% similarity). SSO0386 has 50 -to-30 exoribonuclease activity and is inhibited by the binding of the g-subunit of translation initiation factor a/eIF2 to the 50 end of its RNA substrates. Like most, if not all, archaeal b-CASP proteins, SSO0386 lacks the C-terminal domain found in CPSF-73 and bacterial RNase J. It has an additional 50 amino acids at its N-terminal end, however, whose function is not known. The crystal structures of P. horikoshi and Methanosarcina mazei homologs of CPSF-73 (Ph-1404: 29% identity/52% similarity and Mm-0695: 28% identity/52% similarity to human CPSF-73) have been solved (PDB: 3AF5 and 2XR1, respectively) (Nishida et al. 2010; Mir-Montazeri et al. 2010). Both Ph-1404 and Mm0695 lack the C-terminal domain of CPSF-73 and RNase J, but have two N-terminal KH-domains (Fig. 10.6f). The KH-domain (K homology) is found in many RNAbinding proteins, including PNPase, the exosome, NusA, and ribosomal protein S3. The structure of Ph-1404 has also been solved in complex with a 6-nucleotide RNA analog (PDB: 3AF6) that binds near the N-terminal KH-domain, but far from the active site. The enzyme has been reported to have Zn-dependent single-stranded RNA degrading capability in vitro (Nishida et al. 2010). However, its in vivo activity and how this relates to that of the RNase J ortholog in the same organism remains to be seen. The thermophilic bacterium T. thermophilus also has a homolog of CPSF-73 (30% identity/51% similarity to the human enzyme) called Tth-A0252, in addition to its RNase J ortholog. The structure of Tth-A0252 has also been solved (Fig. 10.6e; PDB: 3IEK) (Ishikawa et al. 2006) and it too lacks the C-terminal

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domain. It has been shown to have degradative activity on rRNA, but the directionality of the degradation reaction was not assessed.

10.5

Phylogeny of rMbLs

Ribonucleases of the MbL family are highly represented in all three kingdoms of life. Eukaryotes generally have RNase Z, CPSF-73 and integrator protein Int11/RC68 orthologs, although this last protein is so far absent in the fungi. RNase Z and CPSF-73 are also ubiquitous in the Archaea. Some have as many as three orthologs of CPSF-73, with all three copies containing a canonical motif 2 histidine signature HxHxDH and at least one copy usually having the N-terminal KH domains. RNase J, which is readily distinguishable from CPSF at the sequence level, is also well represented in the euryarchaeal branch of Archaea, but is confined to this phylum. Although ribonucleases with MbL domain architecture are also widespread in bacteria, they are remarkably under-represented in the Proteobacteria, particularly in the b- and g-subdivisions. As mentioned earlier, RNase Z is absent from the Proteobacteria except for the enterobacterial family of the g-Proteobacteria, that is, the close relatives of E. coli and Salmonella, while RNase J is principally only found in the a- and d/e-subdivisions of the Proteobacteria. The reason for this poor representation of the MbL family of RNases in the Proteobacteria is not clear. CPSF-73 homologs are not strictly confined to the eukaryotes and the archaea, but can also be found scattered throughout the bacterial kingdom. The aforementioned Tth-A0252 protein of T. thermophilus is one example, but there are many others, with the Clostridia alone representing about one third of the bacterial cases (28/90) and the Proteobacteria (g, b and d/e-subdivisions) accounting for another one third of occurrences (36/90). Enterococcus faecalis possesses a b-CASP enzyme, Ef-2904, that is not closely related to either RNase J or CPSF-73 and thus appears to represent yet another subfamily of b-CASP RNases (Fig. 10.8). The structure of this protein has been solved (Fig. 10.6g; PDB: 2AZ4). Like B. subtilis, E. faecalis has both RNase J1 and J2; thus, Ef-2904 is likely to play a different, as yet uncharacterized role in the cell. EF-2904 homologs are found primarily in the Firmicutes, with the Lactobacilli and the Clostridia alone accounting for over 80% of bacterial occurrences of this enzyme (103/131). Ten cases are found in the Archaea (notably in P. abyssi and P. horikoshi, which also have RNase J and CPSF-73 orthologs) and the few remaining examples are found scattered here and there among the bacteria. No close eukaryotic homologs are evident. The different ribonucleases of the rMbL family are clearly distinguishable by sequence alignment and form coherent groups on a phylogenetic tree (Fig. 10.8). Interestingly, the inactive forms of the different b-CASP family members cluster together in related subgroups, suggesting their ancestors arose from an early duplication event that subsequently led to an inactive protein. To generate the short and long forms of RNase Z, a first duplication event was likely followed by yet another,


1

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RNase J2

Fig. 10.8 Phylogenetic tree of rMbLs showing the relationships between the different proteins. The topological tree was made by Neighbor-Joining in MEGA 5.0. Sequence alignment was performed with Clustal W. Organisms are as follows: Ef Enterococcus faecalis, Pab Pyrococcus abyssi, Ph Pyrococcus horikoshi, Sso Sulfolobus solfataricus, Sau Staphylococcus aureus, Bsu B. subtilis, Eco Escherichia coli, m mouse, h human, y yeast (S. cerevisiae), Tth Thermus thermophilus, Sty Salmonella typhimurium

immediately adjacent on the chromosome. The N- and C-terminal halves of mouse and human long-form RNase Z (ELAC2) are more similar to each other than either is to its respective short form (ELAC1). The MbLs are truly a fascinating family of very ancient and important enzymes judging by their widespread distribution, the number of times the genes have been duplicated to allow the evolution of new functions, and their often essential nature. They will likely be the focus of intense study for some time to come. Acknowledgments This work was supported by funds from the CNRS (UPR 9073), Universite´ Paris VII-Denis Diderot and the Agence Nationale de la Recherche (ANR- SubtilRNA2). I thank colleagues and current and former lab members for their contributions to the data discussed in this chapter. I also thank Beátrice Clouet-d’Orval, Zbigniew Dominski, and Allen Nicholson for their helpful comments on the manuscript.

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.

Chapter 11

Ribonuclease III and the Role of Double-Stranded RNA Processing in Bacterial Systems Allen W. Nicholson

Contents 11.1 11.2 11.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RNase III Genomics and Phylogenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ribonuclease III Structure and Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.1 RNase IIII Catalytic Chemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.2 Other Ways to Produce 2 nt 30 -Overhangs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4 RNase III Substrates and Functions in RNA Maturation and Decay . . . . . . . . . . . . . . . . . . 11.4.1 Messenger RNA Maturation and Regulation by RNase III . . . . . . . . . . . . . . . . . . . 11.4.2 RNase III and Ribosomal RNA Maturation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.3 RNase III and Small Noncoding RNA Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.5 On the Site-Specificity of RNase III Action . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6 Noncatalytic Functions of RNase III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.7 Regulation of RNase III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.8 Summary and Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract The enzymatic cleavage of double-stranded(ds) RNA is an essential step in the maturation and decay of bacterial RNAs, and plays key roles in diverse posttranscriptional regulatory pathways that can include small noncoding RNAs. Ribonuclease III is a Mg2+-dependent endonuclease that specifically recognizes and cleaves dsRNA. RNase III is highly conserved in the bacteria and is the founding member of a diverse family that includes the eukaryotic orthologs, Dicer and Drosha. Significant insight has been recently gained on the structure, catalytic mechanism, and functions of RNase III. This review summarizes the current state of knowledge of RNase III, and relates the structural and mechanistic features of the endonuclease to its function in dsRNA-dependent gene expression and regulation.

A.W. Nicholson (*) Department of Biology, Temple University, Philadelphia, PA, USA e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_11, # Springer-Verlag Berlin Heidelberg 2011

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11.1

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Introduction

The discovery of RNA interference (RNAi) and related gene silencing pathways spurred intensive studies on the roles of double-stranded(ds) RNA processing in gene expression and regulation. It was known at that time that the bacterial enzyme ribonuclease III processed dsRNA and regulated gene expression, but it was unclear whether the mechanisms of action and functions of the eukaryotic dsRNA-specific nucleases Dicer and Drosha were similar to those of RNase III. The discovery of RNase III was unanticipated: investigations on the replication of the f2 RNA bacteriophage chromosome detected an activity in Escherichia coli cell-free extracts that could degrade phage dsRNA to acid-solubility (Robertson et al. 1967, 1968). Following isolation of an E. coli mutant that lacked the activity, it was determined that the enzyme, named RNase III, cleaved the ribosomal RNA precursor as well as a polycistronic mRNA precursor of bacteriophage T7 (Dunn and Studier 1973). RNase III function in gene silencing was established in studies on bacteriophage lambda gene expression. It was shown that during lytic growth, the synthesis of lambda Integrase (Int) protein is suppressed by RNase III cleavage of a hairpin structure immediately downstream of the Int cistron in the mRNA transcribed from the phage PL promoter (Guarneros et al. 1982; Gottesman et al. 1982). The action of RNase III destabilizes the transcript by creating an unstructured 30 -end, allowing efficient 30 !50 exonucleolytic degradation of the upstream sequence that contains the Int open reading frame. As a consequence, Integrase synthesis is blocked and the lytic pathway is sustained. Subsequent studies revealed an involvement of RNase III in destabilizing other RNAs through cleavage of sites within mRNA 50 and 30 -untranslated regions, as well as within coding sequences (Re´gnier and Grunberg-Manago 1990; Koraimann et al. 1993). This chapter summarizes the current state of knowledge on the structure, enzymology, and functions of RNase III. A particular focus will be on what has been learned since a crystal structure of Aquifex aeolicus RNase III was reported by Ji and coworkers 10 years ago (Blaszczyk et al. 2001). Studies since then have provided significant insight on RNase III involvement in diverse cellular phenotypes, including antibiotic production, morphogenesis, virulence factor production, and biofilm formation. Broadened investigations of bacterial small noncoding RNAs (sRNAs) and their modes of action have implicated RNase III as an essential accomplice in sRNAdependent gene regulation. These studies have underscored the highly specific action of RNase III in vivo. Determining how such specificity is achieved is key in understanding how RNase III regulates gene expression. Due to page constraints, this chapter will focus on RNase III, rather than the entire family. Eukaryotic RNase III orthologs will be discussed for comparative purposes where appropriate, and the reader is referred to reviews of eukaryotic RNase III family members (LaMontagne et al. 2001; Carmell and Hannon 2004; MacRae and Doudna 2007). Finally, as there have been many studies of RNase III over the 40 years following its discovery, and since these studies cannot be adequately covered here, the reader is referred to the following earlier reviews (Dunn 1982; Court 1993; Nicholson 2003; Drider and Condon 2004).

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Ribonuclease III and the Role of Double-Stranded RNA Processing

11.2

271

RNase III Genomics and Phylogenetics

RNase III is an intracellular nuclease that participates in RNA maturation and decay pathways in conjunction with other endo- and exoribonucleases. Ongoing genomic studies support the consensus that bacterial chromosomes typically contain a single gene (rnc) for RNase III as a highly conserved genetic element. The conservation is underscored by the presence of a rnc gene in the minimal-sized genome of Mycoplasma genitalium (~580,000 bp) that contains ~482 protein-coding genes (Glass et al. 2006). However, Deinococcus radiodurans lacks the rnc gene, but it also lacks the double-stranded stems of the 16S and 23S rRNA precursors that serve as conserved RNase III substrates (Saito et al. 2000) (see also below). Some firmicutes (Bacillus) and cyanobacteria (Synechocystis) encode an additional, truncated form of RNase III that functions in 23S rRNA maturation (see below). Archaeal genomes do not typically contain an RNase III gene. Instead, the “bulgehelix-bulge” (BHB) splicing endonuclease cleaves the rRNA precursors at BHB secondary structures (Xue et al. 2006). The BHB nuclease and RNase III have different structures and catalytic mechanisms. The sporadic presence of RNase III genes in sequenced but otherwise unannotated archaeal genomes may reflect horizontal gene transfer, and the function of RNase III in these organisms is not known. The chromosomal environment of the rnc gene is variable with respect to the identities of the flanking genes and transcriptional organization. However, as discussed below, RNase III cleavage of its mRNA appears to be a conserved posttranscriptional event with autoregulatory significance. Eukaryotic organisms encode multiple genes for RNase III-like polypeptides that contain additional domains, associate with other proteins, and have diverse roles. Fungal RNase III orthologs include Rnt1p (Saccharomyces) and Pac1p (Schizosaccharomyces) that process rRNA precursors as well as small noncoding RNAs (LaMontagne et al. 2001; Nazar 2004). The orthologs Dicer and Drosha participate in a coordinated fashion in microRNA maturation, and also are involved in other processing pathways, including rRNA maturation (Liang and Crooke 2011). An RNase III polypeptide in kinetoplast editosomes catalyzes endonucleolytic cleavages involved in uridine insertion in the pre-mRNA editing pathway (Carnes et al. 2005). A chloroplast RNase III polypeptide, RNC1, participates in Group II intron folding (Watkins et al. 2007) (see also Sect. 11.6). Figure 11.1 provides a gallery of ribonuclease III family members that is expected to have new additions in time.

11.3

Ribonuclease III Structure and Mechanism

RNase III polypeptides typically contain ~220 amino acids, with the minor variation in length confined to specific regions. The N-terminal ~150 amino acids comprises the Nuclease Domain (NucD) that (i) exhibits a unique, all ahelical fold, (ii) contains highly conserved carboxylic acid residues that bind the catalytic

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Fig. 11.1 Ribonuclease III family proteins. The Nuclease Domain (NucD) is highlighted in gray, while the double-stranded-RNA-binding domain (dsRBD) is shown in black. Other domains (unspecified in this figure) that are specific to various family members are shown in white. The symbol “X” in each NucD of the chloroplast RNC1 polypeptide indicates a natural substitution of a conserved, catalytically important residue, providing a protein that binds but does not cleave dsRNA (Watkins et al. 2007). The lengths of each polypeptide in the diagram are not strictly to scale

metal ions, and (iii) forms a stable homodimeric structure. RNase III polypeptides also contain a dsRNA-binding domain (dsRBD) that is joined to the NucD by a short (~7 amino acid) segment of nonconserved sequence, and contains a single copy of the dsRNA-binding motif (dsRBM) (Fig. 11.2). The dsRBM exhibits a conserved abbba fold and is prevalent in eukaryotic proteins that recognize dsRNA (Tian et al. 2004). RNase III is the only characterized bacterial protein with this motif. Gel filtration analysis of purified E. coli (Ec) RNase III indicated a ˚ crystal structure of A. aeolicus homodimeric structure (Dunn 1976), and a 2.1 A (Aa) RNase III confirmed this, and also showed that NucD self-association creates the dimer structure (Blaszczyk et al. 2001). The dimer interface is stabilized by hydrophobic interactions and hydrogen bonds involving conserved residues. The overall shape of the NucD resembles a shallow bowl, with the subunit interface providing a valley that binds dsRNA (Blaszczyk et al. 2001). The two catalytic sites are symmetrically positioned across the interface such that each site can accommodate a phosphodiester from each strand of a bound dsRNA. The scissile phosphodiesters are on opposite sides of the minor groove, and their cleavage provides 2 nt, 30 -overhang product ends that are characteristic of RNase III action. In summary,

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˚ structure of T. Fig. 11.2 RNase III of Thermotoga maritima. The ribbon diagram displays a 2.0 A maritima RNase III, as determined by the Joint Center for Structural Genomics, University of California at San Diego (PDB entry code: 1O0w). The diagram beneath the structure indicates the homodimeric structure of the holoenzyme, and uses the same color scheme in the ribbon structure to indicate the dsRBD (red/orange) and the NucD (green/blue)

the ability to bind dsRNA and provide the characteristic product ends reflects the unique structure of the dimeric NucD and the positions of the catalytic sites. Both dsRBDs are required for optimal RNase III function. Thus, removal of one dsRBD significantly reduces catalytic activity of Ec-RNase III, due to a weakened substrate binding (Meng and Nicholson 2008), while removal of both domains essentially inactivates the enzyme in physiological conditions in vitro (Sun et al. 2001). Structural studies show that the dsRBD exhibits positional mobility that reflects an inherent flexibility of the linker. The two dsRBDs of crystallized Thermotoga maritima RNase III (Fig. 11.2) are arrayed in a symmetric, extended manner with respect to the NucD, while crystals of Aa-RNase III bound to dsRNA show that the dsRBDs bind dsRNA and occupy different positions depending upon the experimental conditions (Blaszczyk et al. 2004; Gan et al. 2005). In an AaRNase III-dsRNA cocrystal structure that exhibits the likely attributes of a precatalytic complex, the dsRBDs assist in binding the dsRNA to the NucD (Gan ˚ crystal structure of Mycobacterium tuberculosis et al. 2008) (Fig. 11.3). A 2.1 A RNase III reveals a substantial residual motion of the dsRBDs, as evidenced by the absence of domain contributions to the diffraction pattern (Akey and Berger 2005). These findings support a model wherein RNase III action requires movement of the dsRBD as it binds dsRNA and then stabilizes the precatalytic complex. A proposed catalytic pathway for RNase III is outlined in Fig. 11.4, and is based on an original scheme by Ji and coworkers (Gan et al. 2005), with some modification. Substrate is initially bound by either dsRBD, perhaps at a rate near the diffusion-controlled limit. This event is followed by NucD engagement of dsRNA to form the precatalytic complex, followed by the chemical step (see also below). The two products disengage from the NucD, with each product still associated with

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Fig. 11.3 Two views of a crystal structure of Aquifex aeolicus RNase III bound to two molecules of a cleaved hairpin substrate (Gan et al. 2006). The enzyme contains the D44N mutation, which inhibits the catalytic step. The two dsRBDs are shown in yellow-gold, while the nuclease domain is shown in blue. The structures in (a) and (b) are related by a 90 rotation

+

+ H2O (Mg2+)

Fig. 11.4 Proposed pathway for dsRNA cleavage by RNase III. The pathway incorporates the following steps: (i) random-order initial recognition of dsRNA by either dsRBD; (ii) engagement of dsRNA by NucD and the other dsRBD to form the precatalytic complex; (iii) Mg2+-dependent hydrolysis of target site phosphodiesters; (iv) disengagement of the two products from the NucD; (v) product release by each dsRBD. The rate-limiting step under steady-state conditions involves one or more steps subsequent to the hydrolytic step (Campbell et al. 2002)

a dsRBD. The products then dissociate from the dsRBDs to provide free enzyme. Biochemical studies are consistent with this model, and highlight the involvement of the dsRBD in substrate recognition and product release. Removal of a dsRBD from Ec-RNase III increases the Km ~ threefold for the cleavage of a substrate (Meng and Nicholson 2008). This is consistent with the statistical aspect of substrate recognition that predicts a twofold increase in Km upon removal of a

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dsRBD, assuming no cooperativity between the two dsRBDs. Also, a requirement for linker flexibility is implied in the step(s) that connect substrate recognition with precatalytic complex formation. However, it has not been determined whether flexibility is strictly required, or whether the dsRBDs only need to act as entropic traps for dsRNA. The rate-limiting step under steady-state reaction conditions is associated with product release (Campbell et al. 2002). The slow step may therefore involve NucD disengagement of product, or dsRBD release of product. If dsRBD release of product is rate limiting then the intrinsic affinity of the dsRBD for substrate may represent a balance between optimal substrate affinity and optimal product release rate.

11.3.1

RNase IIII Catalytic Chemistry

The catalytic activity of Ec-RNase III is supported by Mg2+, Mn2+, Co2+, and Ni2+ ions, but not by Ca2+ or Zn2+ ions (Dunn 1982; Li et al. 1993; Campbell et al. 2002). The observation that Mg2+ best supports the activity, upholds the argument that this metal is the physiologically relevant cofactor. The divalent metal dependence reflects a catalytic rather than a substrate binding requirement (Dunn 1976; Li and Nicholson 1996). A role of metal ion in water nucleophile activation was first indicated by the correlation of the rate of hydrolysis with the metal pKa (Campbell et al. 2002). The structure of the NucD of Aa-RNase III (Blaszczyk et al. 2001) revealed a single metal ion (Mg2+ or Mn2+) coordinated to a conserved set of acidic side chains, as well as with water. A kinetic analysis of Ec-RNase III measured the rate of substrate cleavage as a function of Mg2+ concentration. Single-turnover reaction conditions were employed such that the observed rate reflected only the hydrolytic step, and the modeling assumed that catalysis would only occur if the full set of metal ions are bound. The results indicated that two Mg2+ ions are needed for phosphodiester hydrolysis (Sun et al. 2005). Independent biochemical support for a two Mg2+ ion mechanism was provided by the potent inhibitory action of N-hydroxyhomophthalimide toward Ec-RNase III, and which was shown to inhibit other ribonucleases that employ a two Mg2+ ion mechanism (Sun et al. 2005). The inhibitory ability of the compound is dependent upon the N-hydroxy group and ˚ flanking keto oxygens that can bind two divalent metal ions separated by ~4 A (Billamboz et al. 2008). Structural studies have verified the proposed interaction of this compound with two metal ions in the catalytic site of another ribonuclease (Klumpp and Mirzadegan 2006). N-hydroxyhomophthalimide does not inhibit RNase III in a competitive manner, suggesting that the catalytic site can accommodate the compound along with substrate (Sun et al. 2005). Structural evidence for a two ˚ structure of Aa-RNase III (containing Mg2+ ion mechanism was provided by a 2.05 A the D44N mutation, which reduces catalytic activity) bound to a cleaved dsRNA (Gan ˚ , with one of et al. 2006). The catalytic site contains two Mg2+ ions separated by 4 A the metals corresponding to the metal observed in the (dsRNA-free) Aa-RNase III ˚ crystal structure of Dicer from Giardia structure (Blaszczyk et al. 2001). A 3.3 A

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˚ in the two catalytic intestinalis revealed two Er3+ ions separated by 4.2 or 5.5 A sites (MacRae et al. 2006). The metals occupy positions corresponding to the two metals observed in the Aa-RNase III structure. Although Er3+ does not support Dicer catalytic activity, the conserved positions of the two metals and their interactions with conserved carboxylic acid side chains supports the conservation of the two Mg2+ ion mechanism across the RNase III family. A reaction pathway chemistry was proposed based on crystal structures of AaRNase III bound to dsRNA or the cleavage products, and also incorporated the basic features of the two Mg2+ ion mechanism (Gan et al. 2008; Ji 2008). Catalysis proceeds via an SN2(P) pathway, involving single displacement with inversion of configuration at phosphorus. One Mg2+ ion (Metal A) binds the water nucleophile and coordinates a nonbridging oxygen atom of the scissile phosphodiester. The second metal ion (Metal B) coordinates the 30 -bridging oxygen as well as the nonbridging oxygen atom coordinated by Metal A. Metal B is expected to lower the pKa of the 30 -oxygen leaving group, facilitating cleavage of the P–O bond, while nonbridging oxygen coordination by both metals would enhance phosphorus electrophilicity. It is not clear whether enzyme functional groups in addition to the conserved carboxylic acid side chains that bind the two metals are involved in the catalytic mechanism. However, it was noted in the structural analysis that a nonbridging oxygen atom of the 30 -penultimate phosphodiester closely approaches a water molecule bound to Metal A (Gan et al. 2008). While the functional importance of this interaction has not been critically tested, the oxygen atom may precisely position the water nucleophile and increase its reactivity in a manner similar to that proposed for several restriction enzymes (Jeltsch et al. 1995). However, theoretical calculations do not support a major role for this type of substrate-assisted catalysis for the BamHI restriction enzyme (Fuxreiter and Osman 2001). The Aa-RNase III•dsRNA cocrystal structures also reveal additional divalent metal ions bound to sites close to the Metals A and B (Gan et al. 2008). The function of the additional metal ion-binding sites is unclear, but there are several lines of evidence that suggest an involvement in the reaction pathway, but perhaps not directly associated with the hydrolytic step. First, the additional metal ions are coordinated by conserved carboxylic acid side chains that are important for cata˚ structure of lytic activity (Zhang et al. 2004; Sun et al. 2004). Second, a 2.1 A M. tuberculosis RNase III reveals a Ca2+ ion bound to one of the additional sites, and its proximity to the site of dsRNA binding suggests that a metal ion bound to this site participates in substrate recognition (Akey and Berger 2005).

11.3.2

Other Ways to Produce 2 nt 30 -Overhangs

The two metal ion mechanism is not the only enzymatic option for dsRNA processing. Several vertebrate secreted ribonucleases degrade dsRNA through local denaturation of the double helix, followed by scission of the single-strand segments (Yakoviev et al. 1997; Opitz et al. 1998; Sorrentino et al. 2003). In

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contrast to RNase III, these enzymes employ a transesterification mechanism involving the ribose 20 -hydroxyl group, and do not strictly create 2 nt 30 -overhang product ends. A more intriguing question is whether there are double-strandspecific nucleases that create the same product ends as RNase III, and if so, whether they exhibit structural or mechanistic similarities. Several restriction endonucleases create 2-nt 30 -overhang product ends. Of these, PacI of Pseudomonas alcaligenes has been subjected to structural and biochemical analyses. PacI was crystallized in the presence of an 18 bp DNA containing the 8 bp recognition sequence TTAAT•TAA (the scissile phosphodiester indicated by the dot), and in the presence ˚ structure shows of Ca2+ in order to suppress cleavage (Shen et al. 2010). The 2.0 A that PacI, like RNase III, is a homodimer and binds dsDNA such that the two scissile phosphodiesters on opposite sides of the minor groove are each accommodated in a separate catalytic site. However, in contrast to RNase III, PacI induces major structural changes in the DNA, including a 90 bend that stabilizes alternative base pairings, base mismatches, and also creates unpaired nucleotides, all of which are within the recognition sequence (Shen et al. 2010). The DNA structure exhibits an essentially identical conformation in a product complex obtained by including Mg2+ in the crystallization buffer (Shen et al. 2010). This suggests that phosphodiester cleavage is not accompanied by significant additional changes in the distorted DNA structure. In contrast, dsRNA undergoes only minor conformational changes upon binding Aa-RNase III, with the post-catalytic complex showing minor differences with the precatalytic-related complex (Gan et al. 2008). The conformationally conservative RNase III reaction pathway may reflect the necessity of avoiding an energetically costly conformational change in the structurally conservative A-form double helix. PacI also employs a catalytic chemistry involving a single Mg2+ ion that associates with a bba fold, and a tyrosine side chain may participate in catalysis, perhaps through formation of a transient phosphodiester (Shen et al. 2010). In summary, there is more than one pathway of double-strand nucleic acid processing that provides 2 nt 30 -overhangs. However, structural and enzymatic analyses of other nucleases are warranted in order to assess the scope of the mechanistic repertoire. For example, DNA homing endonucleases of the GIY-YIG family also provide products with 20 nt 30 -overhangs (Nord and Sj€ oberg 2008), and for one of these enzymes there is evidence that the monomeric form is responsible for cleaving both strands of the target site in a twostep process (Corina et al. 2009).

11.4

RNase III Substrates and Functions in RNA Maturation and Decay

Genome-wide studies have shown that RNase III is involved in the expression of genes whose products participate in select pathways. A DNA microarray analysis of E. coli transcripts expressed under conditions in which the level of RNase III ranged between 0.1 and 10 the normal amount showed that the amounts of ~100

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transcripts decreased by at least twofold when the RNase III levels increased (Sim et al. 2010). Proteins encoded by the transcripts are involved in sugar uptake and metabolism, energy production, protein modification, and protein turnover. The 100-fold variation in RNase III levels was shown not to affect the growth rate, which otherwise would have had additional effects on transcript levels. The same study showed that RNase III influences biofilm formation in part by downregulating the levels of the Bdm protein (Sim et al. 2010) (see below). Here, the mRNA of the bdm (biofilm-dependent modulation) gene is cleaved at sites within the coding sequence and the 50 -UTR, with the former cleavage event primarily responsible for initiating transcript decay and limiting Bdm protein production (Sim et al. 2010). Another study used a tiled DNA microarray with 20 nt resolution to interrogate transcripts expressed from either strand of the E. coli chromosome, and in the presence or absence of RNase III (Stead et al. 2010). In the absence of RNase III, approximately 12% of the transcripts exhibited either a significant increase or decrease in abundance. A Gene Ontology (GO)-based analysis revealed that RNase III influences the expression of genes whose products are involved in sulfate uptake and assimilation, iron transport, and the heat shock response, among other pathways. Eleven noncoding(nc) RNAs were more abundant in the absence of RNase III, while four others exhibited decreased levels (Stead et al. 2010). The increase in ncRNA levels in the absence of RNase III may in part reflect the normal processing of the ncRNAs by RNase III upon binding to their targets (see also below). The decrease in the levels of other ncRNAs in the absence of RNase III may reflect the accelerated decay of unprocessed (and therefore aberrant) precursors. This latter pathway also may change the levels of specific mRNAs that are targets of the ncRNAs. Interestingly, the levels of most of the examined polycistronic transcripts were unchanged in the absence of RNase III (Stead et al. 2010). This can be contrasted with the polycistronic pre-mRNAs of phage T7, which are cleaved at multiple sites by RNase III to provide the mature species (Dunn 1976; Dunn and Studier 1983). Given the 20 nt resolution of the tiled microarray, it was also possible to predict RNase III cleavage sites in transcripts by determining the abundance ratios of contiguous RNA sequences in the presence and absence of RNase III. One hitherto undetected site was located within the coding sequence of the nirB transcript (Stead et al. 2010). In summary, the studies by Sim et al. (2010) and Stead et al. (2010) directly implicate RNase III in the coordinated regulation of specific metabolic pathways. Since RNase III itself is controlled by specific factors (see below), the bacterial cell has a posttranscriptional mechanism for rapid physiological adaptation involving the regulated cleavage of dsRNA.

11.4.1

Messenger RNA Maturation and Regulation by RNase III

The bacteriophage lambda strategy of infection includes multiple points of posttranscriptional regulation involving RNase III (reviewed by Court 1993). Since Ec-RNase III levels are influenced by cellular growth rate (Britton et al. 1998), the

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lysis/lysogeny decision is coupled to growth rate via RNase III. Synthesis of the lambda transcription antiterminator N protein is regulated by the functional interplay of transcription and RNA processing (Wilson et al. 2002). The sustained synthesis of N is dependent upon RNase III cleavage of a hairpin element (rIII) adjacent to the ribosome binding site (rbs) of the N cistron (Kameyama et al. 1991). Immediately upstream of rIII is a cis-acting hairpin, NUTL, which is bound by N to form a termination-resistant transcription complex containing N. RNase III cleavage of rIII separates the downstream N cistron and rbs from the NUTL•N•RNA polymerase complex. The separation of the RNAs allows access of the 30S ribosomal subunit to the N rbs, since in the absence of RNase III the rbs is occluded by the adjacent N•NUTL complex (Wilson et al. 1997, 2002). This pathway provides a mechanism for N regulation, such that when RNase III levels fall, N levels decrease and antitermination is attenuated. At higher RNase III levels, N regulation would be minimal and antitermination would be maximal. However, as the authors point out, this model does not include other factors such as ribosome availability (Wilson et al. 2002). The Streptomyces are gram-positive bacteria that that are capable of morphological differentiation and synthesize diverse antibiotics. Studies on Streptomyces coelicolor (Sc) RNase III have revealed an integral function of the enzyme in both processes (Flardh and Buttner 2009; Jones 2010). Sc-RNase III is not essential for cell viability, and the gene is transcribed exclusively during exponential growth (Sello and Buttner 2008). Sc-RNase III dependence of antibiotic production was originally revealed by Champness and coworkers (Adamidis and Champness 1992; Aceti and Champness 1998; Price et al. 1999), and the specific requirement for enzyme catalytic activity was established by Gravenbeek and Jones (2008). A DNA microarray-based analysis of transcripts expressed in an RNase III mutant (absB) strain revealed that Sc-RNase III controls the levels of ~200 different transcripts (Huang et al. 2005). Of these, Sc-RNase III cleaves the rpsO-pnp transcript that encodes ribosomal protein S15 and polynucleotide phosphorylase (PNPase) (Chang et al. 2005), and also cleaves its own mRNA with autoregulatory consequences (see also below). While antibiotic production is not dependent upon the processing of either of these RNAs, other lines of evidence indicate how ScRNase III may function in this regard. First, Sc-RNase III affects the levels of the antibiotic pathway-specific regulators ActII-ORF4, which controls actinorhodin production, and RedD, which controls undecylprodigiosin production (Aceti and Champness 1998). The Sc-RNase III dependence of RedD levels may reflect ScRNase III activation of AfsR2, a higher-order regulator of antibiotic production (Huang et al. 2005). Second, analysis of the S. coelicolor abeABCD gene cluster that encodes proteins involved in actinorhodin synthesis revealed an involvement of Sc-RNase III in controlling the levels of an antisense transcript a-abeA that is transcribed from the strand opposite that of the abeABCD gene cluster, and which overlaps the abeA gene (Hindra et al. 2010). How Sc-RNase III affects a-abeA levels, and whether a-abeA controls AbeA production by an antisense mechanism are not yet established. In summary, these studies are revealing multiple points of

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involvement of RNase III in a complex, tightly controlled pathway of antibiotic biosynthesis. Sc-RNase III also influences the levels of mRNAs encoding factors involved in sporulation. Interestingly, the absence of RNase III enhances sporulation gene expression, while simultaneously suppressing antibiotic production (Xu et al. 2010). One of the mRNAs regulated by Sc-RNase III encodes the transcription factor AdpA that activates expression of sporulation pathway genes (Xu et al. 2010). The observation that AdpA overexpression inhibits antibiotic production while promoting sporulation suggests that AdpA directly or indirectly inhibits RNase III. The inhibition appears to be due to the AdpA-dependent induction of an as yet-unidentified protease that targets RNase III (Xu et al. 2010). In summary, these experiments indicate an additional, complex regulatory network in Streptomyces that includes RNase III, a transcription factor, and a protease. The chromosomes of the genus Neisseria frequently contain a group of short repetitive sequence elements, termed Nemis, or Correia Repeat Elements (CREE) (Correia et al. 1988; Mazzone et al. 2001; Buisine et al. 2002). CREE elements exhibit two size classes (~155 or ~120 bp), and exhibit terminal inverted repeat (TIR) sequences. CREE elements are preferentially located adjacent to genes, such that the 50 -termini of transcripts of the genes map within the TIR (Mazzone et al. 2001). Transcription of a CREE element at a start site within the TIR is predicted to create a 50 -end-proximal hairpin, formed through pairing of complementary sequences within the TIR (see Fig. 11.7). The 50 -ends of the mature RNAs are provided by RNase III cleavage of the hairpins (Mazzone et al. 2001; de Gregorio et al. 2002, 2003). Comparative sequence analysis of two Neisseria gonorrhoeae genomes indicate mobility of CREE elements and their preferential location near the 50 regions of genes important for virulence and survival in the infected host (Snyder et al. 2009). The apparent mobility of the CREE element indicates a genome-dynamic role of this element in influencing gene expression, and suggests diversified, CREE-dependent functions of RNase III in the Neisseriae (Snyder et al. 2009). RNase III may specifically participate in CREE-dependent gene regulation by controlling mRNA stability or translational efficiency. One N. meningitidis CREE element is located immediately upstream of the mtrCDE gene cluster that encodes an antibiotic efflux pump, and the 5’-end of the transcript corresponds to an RNase III cleavage site (Rouquette-Loughlin et al. 2004). Based on this observation it was proposed that the CREE element regulates antibiotic resistance (Rouquette-Loughlin et al. 2004). However, the absence of the CREE element did not significantly affect drug resistance (Enrı´quez et al. 2010), but it also noted in the study that the level and stability of the mtrCDE transcript were not analyzed, and that compensatory regulatory mechanisms may be operative in the absence of the CREE element at this particular locus. It is likely that RNase III-dependent CREE regulation of gene expression occurs under specific conditions, including growth in the infected host organism. In this regard, RNase III is required for N. meningitidis pathogenicity (Sun et al. 2000).

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RNase III and Ribosomal RNA Maturation

RNase III has a conserved role in ribosomal RNA maturation. The ~5,000–6,000 nt primary transcript containing the 16S, 23S, and 5S rRNAs is cleaved by several ribonucleases to provide the mature, functional rRNAs. The nucleolytic cleavages occur concomitantly with transcription, nucleotide modification and ribosomal protein binding (reviewed by Deutscher 2009). RNase III site specifically cleaves stem structures that are formed by the pairing of complementary sequences that flank the 16S and 23S rRNAs. The stems are highly conserved structures (Saito et al. 2000), and can be prepared as small hairpins that are site-specifically cleaved by purified enzyme in vitro (Nathania and Nicholson 2010; Shi et al. 2010). The positions of the cleavage sites are consistent with the creation of the immediate precursors to the mature rRNAs (Nathania and Nicholson 2010; Shi et al. 2010). In E. coli, other ribonucleases that normally function downstream of RNase III in the maturation pathway can act in the absence of RNase III to provide the mature 16S rRNA, while the 23S rRNA retains the unprocessed stem, but is incorporated into a functional 50S subunit (King et al. 1984) In certain bacteria, RNase III also cleaves double-stranded structures within the rRNAs, providing fragmented, albeit functional species (Pronk and Sanderson 2001). Mini-III of B. subtilis(Bs) creates the mature termini of 23S rRNA (Redko et al. 2008). In the rRNA maturation pathway, Bs-RNase III cleaves the 23S and 16S processing stems, followed by Mini-III action on the 23S rRNA precursor. The latter reaction occurs in the 50S subunit and is dependent upon ribosomal protein L3. L3 binds near the 23S rRNA 50 and 30 termini, and may support Mini-III action through local alteration of rRNA structure. In fact, the stimulatory effect of L3 can be mimicked by dimethylsulfoxide, which is known to perturb RNA structure (Redko and Condon 2009). Mini-RNase III is not an essential enzyme, and in its absence the mature 23S rRNA 50 and 30 termini are created by the action of uncharacterized activities (Redko and Condon 2010).

11.4.3

RNase III and Small Noncoding RNA Function

sRNAs have important functions in regulating diverse processes in bacterial cells, such as virulence factor production and response to stress. sRNAs are complementary to sequences in target RNAs, and their binding creates base-paired structures that regulate RNA function and/or stability (reviewed by Wagner 2009; Waters and Storz 2009). However, while the binding event per se can be sufficient to confer regulation, there are an increasing number of examples where RNase III participates in sRNA function by providing irreversible hydrolytic cleavage. Summarized here are examples of the ability of RNase III to function in concert with sRNAs to regulate gene expression. The E. coli ss transcription factor is encoded by the rpoS gene and participates in the cellular response to stress. The

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expression of ss is stringently controlled by a variety of cis- and trans-acting factors, including RNase III (Freire et al. 2006). The rpoS mRNA contains a hairpin element in the 50 -leader that hinders ribosome access to the translation initiation region. The hairpin is cleaved by RNase III, causing an accelerated degradation of the rpoS mRNA that holds ss production in check under normal growth conditions (Resch et al. 2008; McCullen et al. 2010) (Fig. 11.5). However, several sRNAs promote ss production by altering rpoS mRNA structure. The sRNA named DsrA is synthesized during cold shock and binds to the rpoS 50 -UTR, forming a duplex structure in place of the hairpin, and which allows 30S subunits to bind to the translation initiation region (Resch et al. 2008; McCullen et al. 2010). The interaction of DsrA with rpoS mRNA is dependent on Hfq (Sledjeski et al. 2001; Soper and Woodson 2008), but does not require ribosome participation (Vecerek et al. 2010). The alternative duplex structure also is cleaved by RNase III to provide a processed, translationally active mRNA that is more stable, presumably due to the presence of translating ribosomes (Resch et al. 2008) (Fig. 11.5). Thus, RNase III not only functions to keep ss levels low in the absence of DsrA, it also reinforces the action of DsrA. DsrA function at lower temperatures is also dependent on the DEAD-box helicase CsdA, which may destabilize the rpoS 50 -leader hairpin structure and facilitate DsrA binding (Resch et al. 2010). At least two additional sRNAs control rpoS mRNA function. RprA sRNA is induced by cell envelope stress, and interacts with the rpoS 50 -UTR in an Hfq-dependent manner, similar to that seen with DsrA (McCullen et al. 2010; Updegrove et al. 2008; Soper et al. 2010). Also, OxyR sRNA, induced by oxidative stress, is proposed to interact with the rpoS 50 -leader (Zhang et al. 2002; Basineni et al. 2009). RNAIII of Staphylococcus aureus is a trans-acting RNA that also functions as an mRNA. RNAIII encodes the toxin D-Hemolysin, and also binds to and regulates

RN

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as e

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RNase III DsrA-Hfq TIR

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rpoS mRNA

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Fig. 11.5 RNase III involvement in DsrA regulation of rpoS expression (Resch et al. 2008). The left side of the diagram shows the processing of rpoS mRNA by RNase III in a DsrA-independent manner, thereby limiting rpoS expression. DsrA in association with Hfq associates with rpoS mRNA, creating an alternative duplex structure that is processed by RNase III, leading to a translationally competent form of rpoS mRNA, and also a cleaved form of DsrA. The RNase III cleavage sites are shown by the scissors, and the ribosomes are shown in yellow-orange

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Fig. 11.6 Staphylococcus aureus RNAIII regulation of coa mRNA is RNase III-dependent (Chevalier et al. 2010). Panel (a). (left to right): (i) secondary structure of RNAIII, showing helices H7 and H13 (shaded); (ii) initial interaction of RNAIII involving H13 recognition of the Shine-Dalgarno (SD) sequence of the coa mRNA ribosome-binding site (rbs) (I), and H7 recognition of a downstream hairpin element (II); (iii) conformational change to form a sequestered rbs that blocks 30 S subunit binding; (iv) RNase III cleavage of coa mRNA and RNAIII at the two duplex regions (cleavage sites indicated by yellow arrows). Panel (b). Model of the H7-Hairpin II interaction, highlighting the dependence of substrate formation on coaxial stacking

the stability of several mRNAs that encode virulence factors including surface protein A (spa) and staphylocoagulase (coa) (Novick 2003; Huntzinger et al. 2005). RNAIII action is linked to quorum sensing, such that when the cells reach a certain density, the newly synthesized RNAIII binds to the coa mRNA rbs, as well as to a second site within the coding region. The duplexes formed at both sites are targets for RNase III, whose action is required for rapid coa mRNA degradation (Chevalier et al. 2010) (Fig. 11.6). RNase III binding to the upstream site also blocks 30S ribosomal subunit binding and inhibits staphylocoagulase production. The repression of coa mRNA translation and its accelerated turnover provides an effective block to staphylocoagulase synthesis upon entry into stationary phase. RNAIII also controls surface adhesin protein A (Spa) production by a similar mechanism. A 30 -proximal segment of RNAIII binds to the spa mRNA rbs and inhibits translation initiation. The duplex region is cleaved by RNase III and accelerates the decay of the otherwise long-lived mRNA (Huntzinger et al. 2005). The same study also detected an interaction of RNase III with RNAIII. Since RNAIII half-life is not affected by RNase III, it was proposed that RNase III binds RNAIII to form a complex that recognizes spa mRNA (Huntzinger et al. 2005; Chevalier et al. 2010). If this model is correct, then it would be an example of RNase III acting in a noncatalytic manner (see also below). A third example of RNAIII control is provided by regulation of the transcriptional regulatory protein Rot. A reduction in Rot levels by RNAIII allows expression of S. aureus exoproteases and toxins (Boisset et al. 2007). Here, the 30 -domain of RNAIII represses rot mRNA translation through two pairing interactions: one involving the rbs, and the other with a site within the 50 -leader. The 50 -leader interaction may be necessary for specificity, since targeting only the rbs may cause inappropriate

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repression of other mRNAs with accessible rbs (Boisset et al. 2007). RNase IIIdependent Rot mRNA downregulation is not efficient, compared to spa and coa mRNA regulation, and may reflect the need for providing a dynamic range appropriate to a transcriptional regulatory protein. RNase III is not essential for S. aureus viability and the rRNA precursor is still efficiently processed in the absence of the enzyme (Chevalier et al. 2008). Thus, for this organism, RNase III has a prominent function as a higher-order regulator of virulence. The E. coli gadX and gadW genes each encode transcription factors involved in the acid stress response. GadX protein levels are positively regulated by GadY sRNA, which is encoded by the gadY gene. The gadY locus is adjacent to the gadX gene and is on the opposite strand (Opdyke et al. 2004). GadY binds to the 30 -region of the GadX mRNA, forming a duplex structure that is cleaved by RNase III. This reaction creates a shortened mRNA with increased stability, thus providing sustained production of GadX (Opdyke et al. 2010). In the absence of RNase III, GadX mRNA is processed by uncharacterized activities, also in a GadY-dependent manner. A similarity has been noted between this pathway and that of lambda OOP sRNA, which in conjunction with RNase III can positively control lambda cII protein production (Krinke and Wulff 1990). In some instances, RNase III cleaves the sRNAs when they are bound to their targets. This reaction results in stoichiometric rather than catalytic (multiple turnover) sRNA action of the sRNA, providing an extra level of control of sRNA function (Masse´ et al. 2003; Viegas et al. 2010). The E. coli IstR-1 sRNA negatively regulates the production of the toxic peptide TisB during normal cell growth (Vogel et al. 2004). IstR-1 limits TisB production by binding to a specific site in the tisAB mRNA, which contains an extra (“standby”) ribosome binding site upstream of the TisB translation initiation region. The standby site is required for efficient translation of the TisB cistron, and the IstR-1 binding site overlaps the standby site (Darfeuille et al. 2007).The presence of IstR-1 inhibits ribosome loading and prevents TisB translation (Darfeuille et al. 2007). The tisAB mRNA is processed by RNase III at the site of IstR-1 binding, permanently inactivating the mRNA (Vogel et al. 2004), and IstR-1 is cleaved as well (Vogel et al. 2004). Another example is provided by E. coli RyhB, an sRNA that negatively regulates the production of iron superoxide dismutase (SOD) by binding to the ribosome-binding site in the sodB mRNA (Afonyushkin et al. 2005; Pre´vost et al. 2011). RyhB binding blocks ribosome binding and simultaneously promotes RNase E-dependent cleavage of the mRNA at a downstream site (Pre´vost et al. 2011). RyhB turnover is RNase III-dependent and depends upon RyhB binding to sodB mRNA (Afonyushkin et al. 2005). Thus, as with IstR-1, RNase III establishes the stoichiometric action of RyhB. A third example is Salmonella typhimurium MicA, an sRNA that negatively regulates the expression of outer membrane protein A (OmpA). MicA directs RNase III cleavage of a complementary sequence within the ompA mRNA, thus irreversibly inactivating the mRNA. The ompA mRNA can also be cleaved by RNase III in a MicA-independent manner (Viegas et al. 2010). MicA itself is cleaved by RNase III when bound to its target site (Viegas et al. 2010), and also

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by RNase E in a pathway that is independent of MicA binding to ompA mRNA or its other targets (Viegas et al. 2010). In summary, these sRNA-dependent systems underscore the multiple levels of control exerted by sRNAs in conjunction with RNase III, which also can act on the sRNAs.

11.5

On the Site-Specificity of RNase III Action

The functional diversity of RNase III-dependent reactions and associated substrates not only underscores a broad underlying role for dsRNA processing in posttranscriptional regulation of bacterial gene expression, but also brings into focus the critical requirement for the site-specificity of RNase III action. Studies using purified enzyme showed that RNase III can accurately process substrate in vitro (Dunn 1976). In contrast, the eukaryotic ortholog Drosha requires a protein cofactor (DGCR8 in mammals) for activity (Han et al. 2004). Specific structural features in RNase III substrates are necessary for reactivity, and to establish the specific pattern of cleavage. An ~11 bp helix is the minimum length necessary for reactivity of EcRNase III substrates, and reflects the presence of protein-RNA contacts that span this interval and provide binding energy. Additional structural features such as internal loops or bulges can limit cleavage to a single strand, or suppress cleavage without affecting RNA binding (Calin-Jageman and Nicholson 2003a, b) (Fig. 11.7). Although helix length and local secondary structural elements establish basic reactivity and the pattern of cleavage, respectively, neither feature identifies the target site. Instead, base-pair sequences within discrete double-helical regions function as positive determinants of cleavage site selection (Zhang and Nicholson 1997; Pertzev and Nicholson 2006). For Ec-RNase III substrates, the 3 bp proximal box (pb) and the 2 bp distal box (db) function in concert to determine the cleavage site (Fig. 11.7). Thus, duplication of a pb/db pair in a substrate creates an additional cleavage site that is appropriately positioned with respect to the two duplicated elements (Pertzev and Nicholson 2006). The db is centered 11 bp from the target site and directly interacts with a peptide segment (RBM4) in the NucD of AaRNase III (Gan et al. 2006, 2008). Deletion of the db causes a drop in substrate reactivity due to a loss of binding affinity (Zhang and Nicholson 1997). Specific bp substitutions in the db inhibit binding of Ec-RNase III (Zhang and Nicholson 1997; Pertzev and Nicholson 2006), but bp substitutions in the db of Aa- or T. maritima (Tm) RNase III substrates have minimal effect on reactivity (Shi et al. 2010; Nathania and Nicholson 2010). This suggests species-specific variation in the dbRNase III interaction, which is consistent with the variability in both the length and sequence of RBM4. RNase III binding also is sensitive to bp substitutions at each position in the pb, which is located immediately adjacent to the target site, but is particularly sensitive to specific substitutions at pb position 2. Here, a CG or GC substitution in place of the canonical AU or UA bp inhibits the binding of Ec-, Aa-, and Tm-RNase III (Pertzev and Nicholson 2006; Shi et al. 2010; Nathania and Nicholson 2010). Crystallographic studies of Aa-RNase III show that the pb

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Fig. 11.7 Substrates recognized by RNase III. RNase III cleavage sites are indicated by arrows. (a) R1.1[WC-L] RNA (Li and Nicholson 1996). The RNAs are drawn with the main secondary structural features. The rectangles indicate the positions of the distal box (db), middle box (mb), and proximal box (pb). The boxes below the cleavage sites indicate the symmetry-related pb, mb, and db. The individual pb positions are numbered (1,2,3). The RNase III domains (RBM1, RBM2, RBM3, and RBM4) that interact with the boxes are indicated on the left. (b) R1.1 RNA (Chelladurai et al. 1993). The rectangle indicates the position of the proximal box, and the single RNase III cleavage site is shown by the arrow. (c) R1.1[CL3B] RNA (Calin-Jageman and Nicholson 2003a). This substrate is bound by RNase III, but is resistant to cleavage. (d) Neisseria Mini-26/26a RNA (left) and Mini-26/27 RNA (right) are CREE element RNA hairpins that are recognized by RNase III (De Gregorio et al. 2003). Mini-26/26a RNA is cleaved by RNase III, while Mini-26/27 RNA is bound by RNase III, but resistant to cleavage. Since the cleavage sites have not been mapped, assignment of the pb (indicated by the rectangle) in Mini-26/26a is tentative

engages in several contacts with residues in the N-terminal portion of the first a helix in the dsRBD (termed RBM1) (Gan et al. 2006, 2008). This interaction is consistent with the functional requirement of the dsRBD in substrate binding (Sun et al. 2001). Alanine substitution of an invariant Glutamine in the Aa-RNase III RBM1 strongly weakens substrate binding (Shi et al. 2010). This mutation is expected to cause the loss of two hydrogen bonds with the bp at pb position 2. The proximal box position 2 also is the site of strong inhibition by the CG/GC bp substitution (see above). One continuing puzzle is the retention of site-specificity of substrate cleavage by the NucD in the absence of the dsRBD (Sun et al. 2001). This suggests that a specific feature(s) of the NucD interacts with the pb or with a yet uncharacterized feature of the substrate to establish site-specificity of action.

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The inhibitory GC/CG substitution at pb position 2 provides a mechanism by which RNase III action can be modulated by RNA sequence. For example, cellular dsRNA structures of 11 bp, whose cleavage would otherwise be detrimental, can be protected by a single bp substitution (CG or GC) at a site corresponding to pb position 2. The inhibitory bp is proposed to function as an antideterminant, to borrow a term that describes tRNA sequence elements that prohibit recognition by noncognate aminoacyl-tRNA synthetases (Rudinger et al. 1996). Thus, the dsRBD not only provides binding energy, but also is responsive to antideterminants and can act to restrict RNase III access to only bona fide substrates.

11.6

Noncatalytic Functions of RNase III

Several lines of evidence suggest that RNase III can function by binding RNA in a noncatalytic manner. RNase III binding could control mRNA translation or stability by a steric effect, or by inducing RNA conformational changes. An sRNA could be involved in either mechanism, wherein RNase III stabilizes the duplex structure without cleavage. Genetic studies suggest that expression of the phage lambda cIII and Int proteins may be controlled in part by RNase III acting in a noncatalytic manner (reviewed by Court 1993), and it was also shown that a catalytically inactive, binding-competent Ec-RNase III mutant expressed in vivo can interfere with endogenous RNase III action in a dominant negative manner (Dasgupta et al. 1998). There is no conceptual barrier in considering RNA structures or additional factors that can selectively suppress cleavage without affecting enzyme binding. The intercalating agent ethidium bromide binds to the internal loop-containing substrate, R1.1 RNA, and blocks Ec-RNase III cleavage of the single scissile bond without inhibiting binding (Calin-Jageman et al. 2001). Two ethidium-binding sites were identified, with one adjacent to the cleavage site, and it was proposed that occupancy of the latter site may cause a local structural change that prohibits cleavage. RNAs have been isolated by in vitro selection techniques that bind Ec-RNase III in vitro, but are resistant to cleavage (Calin-Jageman and Nicholson 2003a, b) (Fig. 11.7). There has been some success in the identification of cellular RNAs that function in this manner. Transcripts of a subset of the Correia Repeat Enclosed Elements (CREE) (see above) form irregular RNA hairpins that resist cleavage, yet are able to bind RNase III in vitro (De Gregorio et al. 2003) (Fig. 11.7). Given the proximity of these hairpins to genes, these elements may regulate mRNA stability or translation in a (noncatalytic) RNase III-dependent manner. How might a dsRNA resist cleavage by RNase III? Crystallographic studies show that RNase III can engage dsRNA by the dsRBD alone (Blaszczyk et al. 2004; Gan et al. 2005), and the absence of dsRNA-NucD contacts in these structures can rationalize the noncatalytic action of RNase III (Pastore 2004; Ji 2006). However, the NucD may be able to engage dsRNA without concomitant cleavage. The NucD of Ec-RNase III appears to recognize a cleavage-resistant R1.1 RNA variant, since bp substitutions in the db – an element which interacts with the NucD – inhibit

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binding of the RNA (Calin-Jageman and Nicholson 2003a). If the RNA were bound only to the dsRBD, then these mutations would not be expected to affect binding. The cleavage-resistant R1.1 variant may exhibit structural features at or near the target site that prevent placement of the scissile phosphodiester in a catalytic site. Finally, current notions of how RNase III functions have been based on an either/or model, wherein substrate either is cleaved, or binds enzyme without cleavage. However, it is possible that RNase III may regulate RNA function and stability in a mixed mode. Here, the enzyme acts in an immediate manner as an RNAbinding protein, then cleaves the target after a lag time, providing irreversibility to the initial regulatory event, and perhaps also initiate turnover. This model can be compared to sRNA action where a rapidly formed duplex exerts regulation, and is then cleaved by RNase III at a rate that is dependent upon a slower RNA conformational change.

11.7

Regulation of RNase III

E. coli RNase III regulates its levels by cleaving a base-paired structure within the 50 -leader of the mRNA, which initiates rapid decay (Matsunaga et al. 1996). RNase III autoregulation has been demonstrated in Streptomyces. The Sc-RNase III gene (absB) is transcribed as one of three cistrons within the operon transcript that is cleaved by Sc-RNase III at two sites, one of which is within the RNase III coding sequence (Xu et al. 2008). Eukaryotic RNase III orthologs exhibit autoregulation, with some variation on the theme. Ascoviruses express an RNase III-like protein that probably acts to subvert the host RNAi response by cleaving viral dsRNAs to products that are too short to enter the RNAi pathway (Hussain et al. 2010). The Ascovirus RNase III also cleaves its own mRNA, which limits enzyme production as infection proceeds (Hussain et al. 2010). Dicer orthologs regulate their expression with the help of a miRNA. A global analysis of miRNA target sites within human protein coding sequences identified three let-7 miRNA consensus sites within the Dicer mRNA (Forman et al. 2008). Dicer expression is reduced when let-7 levels increase, and mutations that disrupt pairing of let-7 with Dicer mRNA abrogate the reduction (Forman et al. 2008). A separate investigation showed that antisense-imposed decrease in let-7 levels enhances Dicer expression in mammalian cells, while let-7 overexpression reduced the level of the enzyme (Tokumaru et al. 2008). Since Dicer participates in let-7 maturation, a negative autoregulatory loop is established. Dicer-Like protein 1 (DCL1) of Arabidopsis thaliana regulates its expression with the participation of a miRNA (miR162) that has a target site in the DCL1 mRNA. The reduction of miR162 activity either by mutation or by expression of trans-acting factors increases the levels of DCL1 (Xie et al. 2003). Drosha and the RNA-binding protein DGCR8 are the primary components of the nuclear microprocessor complex that converts primary miRNA transcripts (primRNAs) to pre-miRNAs, which are then exported to the cytoplasm for final maturation by Dicer. Drosha cleavage of pri-miRNAs requires DGCR8, which

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recognizes specific structural features of the substrates (Han et al. 2004). A hairpin within the 50 -UTR of the DGCR8 mRNA is a microprocessor target. Cleavage of this structure downregulates DGCR8 expression, thereby reducing microprocessor levels (Triboulet et al. 2009; Han et al. 2009). The hairpin substrate also can regulate the expression of fused reporter genes in a microprocessor-dependent manner (Triboulet et al. 2009; Han et al. 2009). The involvement of the microprocessor in mRNA turnover appears to be limited thus far to DGCR8 mRNA (Shenoy and Blelloch, 2009). These examples underscore the conserved function of RNase III family members in autoregulation. The E. coli cell contains ~500 molecules of RNase III, with the enzyme levels increasing with higher growth rates (Court 1993; Britton et al. 1998). The growth rate dependence may be a consequence of rRNA gene transcription rate, in that the higher amounts of the rRNA precursors would titrate the limited amount of RNase III. This in turn would decrease the amount of enzyme available for autoregulation, thereby providing higher RNase III levels. However, the precise mechanism of growth rate control of RNase III levels is likely to be more complex (Britton et al. 1998). Ec-RNase III also is regulated by trans-acting factors. Cohen and coworkers (Kim et al. 2008) used a novel genetic screen to identify a protein, YmdB, that downregulates Ec-RNase III activity in vivo. YmdB is produced during cold shock or entry into stationary phase, and its expression is dependent on the alternative sigma factor ss. Cross-linking studies indicate that YmdB interacts with the NucD, and that the cleavage step is inhibited without affecting RNA binding. It was proposed that YmdB binding perturbs NucD structure without affecting the ability of the dsRBD to bind dsRNA (Kim et al. 2008). However, the in vitro system may not precisely recapitulate the inhibition observed in vivo, since inhibition in vitro required a high amount of YmdB relative to RNase III, and also required Mn2+ rather than Mg2+. An additional factor downregulates Ec-RNase III activity during osmotic stress (Sim et al. 2010). This factor has not been characterized, but functions to increase the levels of the bdm mRNA, which is a substrate for RNase III and encodes a protein that promotes biofilm formation. The general requirement for inhibition of Ec-RNase III by YmdB or other factors during cold stress, osmotic stress, or entry into stationary phase (see above) is unclear, but at least one posttranscriptional pathway is suggested. As discussed above, EcRNase III cleaves a structure within the rpoS mRNA 50 -leader and reduces ss production (Resch et al. 2008). The presence of YmdB would limit this reaction and, as a consequence, elevate ss levels and support entry into stationary phase. YmdB involvement in response to cold stress may involve a similar need to suppress cleavage of specific substrates. However, one must include the inhibition of cleavage of the rnc mRNA 50 -leader as a necessary consequence of inhibiting RNase III, which would increase RNase III levels. While the complex features of gene regulatory networks involving dsRNA processing remain to be fully characterized, it is clear that trans-acting factors can provide rapid control of RNase III activity in response to diverse changes in cellular environment and physiology (Kim et al. 2008). Ec-RNase III is regulated by covalent modification. Bacteriophage T7 expresses a serine/threonine-specific protein kinase (T7PK) that phosphorylates ~100 proteins

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in the infected cell (Rahmsdorf et al. 1974; Robertson et al. 1994). RNase III undergoes phosphorylation on serine in the T7-infected cell, which stimulates its catalytic activity ~3–5 fold (Mayer and Schweiger 1983). Phosphorylation may enhance the action of a limited pool of RNase III in order to process in a more efficient manner the high levels of T7 pre-mRNAs with multiple processing sites. T7PK supports T7 growth at elevated temperatures or in poor carbon/energy sources (Hirsch-Kauffmann et al. 1975). This may reflect the phosphorylation of additional proteins that function in other steps in the T7 replication pathway. There is no evidence for a protein phosphatase that could reverse the reaction, nor is there evidence for a cellular counterpart of T7PK that has a similar function. Another type of covalent regulation is supported by studies of Sc-RNase III. As discussed above, Sc-RNase III levels are reduced by a proteolytic activity that is stimulated during S. coelicolor sporulation (Xu et al. 2010). Whether proteolytic regulation of RNase III occurs in other bacterial systems remains to be determined.

11.8

Summary and Prospects

This review has attempted to summarize and analyze the recent advances in our understanding of RNase III structure and function. These studies reveal an enzyme that can also act as an RNA-binding protein and is responsive to multiple types of regulation. These findings are now informing systems biology based investigations that are revealing complex posttranslational regulatory networks involving RNase III, along with other nucleases, proteins, and sRNAs. These RNA-based regulatory networks provide the cell with the ability to quickly adapt to changes in environment, and undergo developmental programs such as sporulation, antibiotic production, or biofilm formation. Early speculations on a host defense function for RNase III (discussed by Robertson 1982) have been reinvigorated by the demonstration that RNase III of Streptococcus pyogenes, in conjunction with Csn1 protein and the trans-encoded small RNA, tracrRNA, provide a maturation pathway for antiphage and antiviral cRNAs of the CRISPR/Cas host defense system (Deltcheva et al. 2011). Here, duplex structures that are formed by tracrRNA binding to repetitive sequence elements in the pre-cRNAs are cleaved by RNase III (Deltcheva et al. 2011). Recent practical applications of RNase III and specific mutant versions include the production of siRNAs from dsRNA in vitro (Yang et al. 2002; Xiao et al. 2009). Also, knowledge of the structures and reactivities of RNase III substrates are informing the de novo design of regulated metabolic pathways (Babiskin and Smolke 2011). To conclude, from its modest debut as a novel activity in cell-free extracts, RNase III now occupies center stage as a functionally versatile nuclease with diverse roles in dsRNA-dependent gene expression and regulation. Acknowledgments The author thanks Dr. Zhongjie Shi and Dr. Lilian Nathania and for assistance in figure preparation, Dr. Pascale Romby for providing a high resolution version of Fig. 11.6, and Dr. Rhonda H. Nicholson for comments on the manuscript.

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Chapter 12

Structure and Function of RNase H Enzymes Thomas Hollis and Nadine M. Shaban

Contents 12.1 12.2 12.3 12.4 12.5

Introduction to RNase H Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Overall RNase H Structure and Catalytic Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RNase H1 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RNase H2 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Other RNase H Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.5.1 RNase HIII Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.5.2 Viral RNase H Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

300 301 303 306 309 309 310 312 312

Abstract RNase H enzymes are endonucleases that specifically cleave ribonucleotides within an RNA:DNA duplex. RNase H proteins are divided into type 1 and type 2 enzymes based on amino acid sequence similarities, substrate specificity, and structure. Both RNase H1 and RNase H2 enzymes play important roles in DNA replication, repair and transcription, and at least one type of RNase H is found in most organisms. Both RNase H1 and RNase H2 enzymes share a common structural fold of mixed b-sheets surrounded by several helices at their catalytic core. The enzymes utilize a two-metal-ion mechanism of phosphoryl hydrolysis mediated by divalent Mg2+ ions. RNase H1 enzymes are single polypeptide proteins with eukaryotic members containing an additional hybrid-binding domain (HBD). Eukaryotic RNase H2 is a heterotrimeric complex of the RNase H2A, RNase H2B, and RNase H2C proteins that are all necessary for enzymatic activity. Mutations in the human RNase H2 complex result in immune dysfunction.

T. Hollis (*) • N.M. Shaban Department of Biochemistry, Center for Structural Biology, Wake Forest School of Medicine, Winston-Salem, NC 27157, USA e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_12, # Springer-Verlag Berlin Heidelberg 2011

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12.1

T. Hollis and N.M. Shaban

Introduction to RNase H Enzymes

RNase H enzymes specifically recognize ribonucleotides hybridized to DNA within an RNA:DNA duplex and catalyze the hydrolysis of the phosphodiester bond on the 50 side of the ribonucleotides to create a single-strand nick within the duplex. RNase H enzymatic activity was first identified in calf thymus by Peter Hausen’s group in 1969 (Stein and Hausen 1969; Hausen and Stein 1970). Later, the first genes encoding bacterial RNase HI and RNase HII enzymes were identified in Escherichia coli (Kanaya and Crouch 1983; Itaya 1990), and as genome sequences became increasingly available it became apparent that most organisms contain at least one RNase H enzyme. RNase H enzymes are classified into Type 1 and Type 2 according to amino acid sequence similarity, structure, and substrate specificity (Ohtani et al. 1999a; Crouch et al. 2001). The current nomenclature for RNase H was adopted to allow for facile comparison of enzymes from different organisms and utilizes Roman numerals to denote RNase H proteins from prokaryotes (I/II) and Arabic numerals for eukaryotes (1/2). RNase H1 and RNase H2 enzymes each display distinct cleavage patterns on RNA:DNA hybrid substrates (Eder and Walder 1991; Frank et al. 1994; Pileur et al. 2000; Haruki et al. 2002). While both types of enzymes will cleave an RNA:DNA hybrid duplex, the major distinctions in substrate specificity are that RNase H2 preferentially cleaves on the 50 -side of ribonucleotides at an RNA-DNA junction, and will also catalyze the hydrolysis of single ribonucleotides within a DNA duplex, while RNase H1 requires a minimum of four ribonucleotides for recognition (Fig. 12.1). The precise physiological roles for RNase H enzymes are unclear but evidence for RNase H function in DNA replication and repair, viral replication, and processing of R-loops has been demonstrated (Eder and Walder 1991; Eder et al. 1993;

Fig. 12.1 Substrate specificities of RNase H enzymes. RNase H enzymes are differentiated into Type 1 and Type 2 with distinct differences in substrate specificity. RNase H1 enzymes readily hydrolyze ribonucleotides within an RNA:DNA duplex while RNase H2 preferentially cleave on the 50 side of a ribonucleotide at an RNA-DNA junction within the hybrid substrate. Another significant distinction between the two types is that RNase H2 enzymes are able to hydrolyze a single ribonucleotide within a DNA duplex while RNase H1 requires a minimum of four ribonucleotides for recognition

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Structure and Function of RNase H Enzymes

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Qiu et al. 1999; Arudchandran et al. 2000; Chai et al. 2001; Chapados et al. 2001; Rydberg and Game 2002; Cerritelli et al. 2003; Lin et al. 2010). While RNase H activity is not essential for bacterial growth, deletions of RNase H1 display embryonic lethality in mice due to a failure to replicate mitochondrial DNA (Kanaya and Crouch 1984; Cerritelli et al. 2003). Deletions of RNase H1 and RNase H2 genes in Saccharomyces cerevisiae result in an increased sensitivity to hydroxyurea, caffeine, and ethyl methanesulfonate (EMS) (Arudchandran et al. 2000). More recently, mutations in the genes encoding for the human RNase H2 enzyme have been linked to the autoimmune disease, Aicardi–Goutie`res syndrome (AGS), supporting a role for this enzyme in processing cellular RNA:DNA intermediates to avoid immune dysfunction (Crow et al. 2006b; Ramantani et al. 2010). While many organisms contain both RNase H1 and RNase H2 genes, some contain only a single RNase H gene. Most archaeal genomes and some plants lack genes coding for RNase H1 and have only a gene for RNase H2 (Shultz et al. 2007; Tadokoro and Kanaya 2009). The biological significance of having a single versus multiple RNase H genes is unclear. Functional similarities between the two enzymes may indicate redundancy in an essential biological system. However, deletions of both RNase HI and HII from bacteria are not lethal, but rather how a temperature sensitive growth phenotype. Additionally, the ability of RNase H2 to hydrolyze a single ribonucleotide embedded in DNA is an activity not present in RNase H1, and supports a model for differing biological functions between the two enzymes.

12.2

Overall RNase H Structure and Catalytic Mechanism

Crystal structures of both RNase H1 and RNase H2 enzymes from many species have been determined, and reveal they share a common structural architecture. The RNase H fold consists of a five-stranded mixed central beta sheet flanked by several alpha helixes that has become defined as the archetypal structure of the well-conserved ribonuclease H-like superfamily (Pfam CL0219, http://pfam. sanger.ac.uk (Finn et al. 2010)). This large structural superfamily contains at least 25 other nucleases including 30 -exonucleases, transposases, and RuvC resolvase. The highly conserved fold suggests it has evolved as an effective scaffold for recognition and hydrolysis of DNA and RNA substrates. The conserved structure of these enzymes indicates that they utilize a common catalytic mechanism, which has been confirmed by structural and biochemical studies of RNase H proteins from a number of species (Chapados et al. 2001; Nowotny et al. 2005, 2007; Nowotny and Yang 2006; De Vivo et al. 2008; Rychlik et al. 2010; Shaban et al. 2010). RNase H enzymes utilize a two-metal-ion-mediated mechanism of phosphoryl hydrolysis (Fig. 12.2) (Nowotny et al. 2005; Yang et al. 2006). This mechanism is facilitated by four positionally conserved acidic residues within the active site that form the catalytic core of the enzyme (Fig. 12.2). In enzyme-substrate complexes these acidic amino acids along with the oxygen atoms

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Fig. 12.2 Two-metal-ion catalytic mechanism of RNase H enzymes. Structure of Bacillus halodurans RNase H1 in complex with RNA:DNA hybrid (pdbid: 1ZBL) shows four conserved acidic residues in the active site coordinating two divalent Mg2+ ions (blue spheres) (Nowotny et al. 2005). The metal ions also ligand water molecules (red spheres) and the phosphate oxygen of the nucleic acid substrate. The canonical labeling of metal ions A and B are shown. A water molecule above metal A is likely activated by the metal and the Rp oxygen of the adjacent phosphate for in-line attack of the scissile phosphate (red arrow)

from the scissile phosphate on the ribonucleotide substrate provide coordination ligands to the two Mg2+ ions. There is experimental evidence that catalytically relevant binding of the two metal ions in the active site occurs only in the presence of substrate (Goedken and Marqusee 2001; Nowotny et al. 2005). The positions of the two divalent ions are canonically labeled as A and B (Fig. 12.2). Hydrolysis is thought to occur via deprotonation of a water molecule located above metal A to form a nucleophile for in-line attack of the target phosphate. In many two-metal-ion mechanisms metal ion A is proposed to play a contributing role in generation of the water nucleophile. Structural and biochemical evidence from the bacterial RNase HI and archaeal RNase H2 enzymes however support a model for the pro-Rp oxygen of the phosphate immediately to the 30 side of the scissile bond in orienting the water for nucleophilic attack (Haruki et al. 2000; Nowotny et al. 2005; Rychlik et al. 2010). The role for metal B is in stabilizing the transition state intermediate ˚ of one another. during which the two metal ions are thought to move to within 3.5 A Finally, after hydrolysis the cleaved phosphate can no longer simultaneously coordinate the two metal ions, and likely one of the metal ions is displaced from the active site leading to the release of cleavage product (Nowotny and Yang 2006). Several RNase H enzymes, such as the human RNase H1, E. coli RNase HI, and HIV reverse transcriptase, have a conserved histidine residue located on a mobile loop C-terminal to the active site residues that is postulated to contribute to substrate release by dislodging the 50 phosphate product from the active site (Nowotny et al. 2007).

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12.3

303

RNase H1 Enzymes

The prokaryotic RNase HI enzymes are the best structurally and biochemically characterized members of the RNase H family (Fig. 12.3). The first RNase H structures from E. coli defined the steric configuration of the catalytic residues within the active site cleft and identified regions of the protein likely involved in RNA:DNA hybrid interactions (Katayanagi et al. 1990; Yang et al. 1990). Although an accumulation of experimental information has provided significant comprehension as to the mechanistic aspects of the enzymes, the exact cellular roles and physiological function of RNase H1 enzymes remain in question.

Fig. 12.3 RNase H1 enzymes. Structures of the (a) Bh-RNase HI and (b) human RNase H1 proteins show the common RNase H architecture among the enzyme family. The four conserved acid residues in the active site of each enzyme are highlighted in orange. The basic protrusion present in the human and Escherichia coli RNase H1 enzymes that contributes to substrate binding and specificity is labeled. (c) The complex of Bh-RNase H1 and RNA:DNA hybrid reveals that the protein utilizes two grooves on the surface that are spaced to recognize the spacing of the RNA: DNA hybrid (pdbid: 1ZBL). (d) Structure of the hybrid-binding domain (HBD) from human RNase H1 in complex with RNA:DNA (pdbid:3BSU); a common feature of eukaryotic and some prokaryotic RNase H1 enzymes which anchors the protein to the nucleic acid substrate and likely contributes to processivity. The protein domain interacts with the RNA strand through a protein loop that forms hydrogen bonds with the 20 -OH groups and an aromatic patch (residues shown) that appears selective for binding deoxyribonucleotides on the DNA strand

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The RNase H1 enzymes require a minimum of four ribonucleotides within an RNA:DNA hybrid for substrate recognition (Ohtani et al. 1999a) suggesting that the enzyme recognizes a unique feature of the hybrid not present with fewer ribonucleotides. The structure of the Bacillus halodurans RNase H1 in complex with an RNA:DNA hybrid revealed that the protein specifically recognizes the mixed A and B form of the nucleic acid duplex (Nowotny et al. 2005). The RNA strand is in the A form and the DNA strand is in B form and together create a hybrid with intermediate A/B character. As a consequence of this, the minor groove width ˚ , larger than the average 4 A ˚ for the RNA:DNA hybrid is narrowed to about 8.4 A ˚ for A form RNA and significantly less than 11.5 A for the average of B form DNA helices. Other work has shown that altering the conformation and flexibility of the RNA:DNA hybrid reduces the cleavage activity of RNase H1 (Lima et al. 2004, 2007a, b), supporting the model that the enzyme must make contact with both the RNA and DNA strands. The active site face of the RNase H1 protein has two ˚ into which the backbones of the RNA:DNA grooves separated by about 8.5 A hybrid fit (Fig. 12.3). Specificity for the RNA strand is imparted by direct contacts between the active site groove and the consecutive 20 -OH groups on the substrate and explains the need for two ribonucleotides on each side of the scissile bond (Nowotny et al. 2005). A variation of this substrate recognition theme is seen in the structurally similar human, E. coli and Moloney murine leukemia virus (M-MLV) RNase H1 enzymes (Katayanagi et al. 1990; Yang et al. 1990; Lim et al. 2006; Nowotny et al. 2007). In addition to the two surface grooves for RNA:DNA hybrid binding, these proteins have a basic protrusion that contributes to the formation of the DNA-binding channel (Fig. 12.3). The basic protrusion provides additional van der Waals and hydrogen bond interactions with the DNA strand that contribute to substrate specificity. In addition to the RNase H domains, all eukaryotic and some bacterial RNase H1 enzymes have highly conserved regions at their N-terminal region that were first described as a dsRNA and RNA:DNA-binding domain with high specificity for RNA:DNA hybrids (Cerritelli and Crouch 1995; Cerritelli et al. 2003; Gaidamakov et al. 2005). This N-terminal region is called the hybrid-binding domain (HBD), and it has about 25 fold higher affinity for an RNA:DNA duplex over dsRNA (Nowotny et al. 2008). Although S. cerevisiae contains two such domains at its N-terminus, only one seems to be able to form stable complexes with duplex RNAs alone. The second copy of the HBD does contribute to the overall binding affinity when in combination with the first HBD (Cerritelli and Crouch 1995; Cerritelli et al. 1998). In other eukaryotic RNase H1 enzymes a single HBD followed by a linker region and the RNase H domain is the most commonly observed physical arrangement (Cerritelli and Crouch 1998). Deletion of the HBD in the human and mouse RNase H1 proteins severely reduces RNase H activity and decreases processivity when compared the wild-type enzyme (Gaidamakov et al. 2005; Nowotny et al. 2008). These experiments predict a model for RNase H1–nucleic acid interaction where the HBD first approaches the RNA:DNA substrate to facilitate initial binding. The HBD then acts as an anchor to allow the RNase H domain to perform

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multiple interactions at nearby sites of the hybrid, thereby allowing several rounds of RNA hydrolysis per HBD-binding event and contributing to processivity. The length of the linker region between the HBD and RNase H domain may also contribute to the avidity of the complex. The shorter linker of ~10 amino acids in bacterial versus the longer ~60 amino acids in mammalian RNase H proteins appear to provide increased binding affinity, but at the expense of processivity (Nowotny et al. 2008). Alternatively, the increased length of the connection region in eukaryotes presents the possibility that this region of the protein may have evolved for additional protein–protein interactions in higher organisms. Structural and mutagenesis studies of the HBD have provided insight into substrate specificity and binding. The structure of the human HBD in complex with a 12 base pair RNA:DNA hybrid shows that the protein interacts with the hybrid mostly through phosphodiester backbone interactions along the minor groove (Fig. 12.3). Specificity for the RNA strand is provided by a protein loop that forms hydrogen bonds with the 20 -OH groups, while the HBD protein also has an aromatic patch that appears selective for binding deoxyribonucleotides on the DNA strand (Nowotny et al. 2008). The precise physiological roles of RNase H1 enzyme in the cell are still under investigation. RNase H1 has been suggested to participate in several biological functions including DNA repair and replication. During DNA replication, synthesis of the lagging strand occurs through extension of discontinuous segments primed with RNA primers known as Okazaki fragments. Removal and processing of the RNA primers is essential for the proper joining of the fragments to form an intact DNA strand. While early work supported a role for RNase H in processing of Okazaki fragments (Huang et al. 1994; Kogoma and Foster 1998; Murante et al. 1998), more recently, other mechanisms more likely to be involved in this process have been elucidated (Kao and Bambara 2003; Rossi et al. 2006). RNase HI is also unable to remove the last ribonucleotide at the RNA-DNA junction, a step that would be necessary for the complete processing of Okazaki fragments. Nonetheless, a role for RNase H in DNA replication is still likely as Rnaseh1 knockouts are lethal in mice embryo due to a failure to replicate mitochondrial DNA. Another possible in vivo substrate for RNase H1 could be extended stretches of RNA:DNA hybrids that form during transcription, and are known as R-loops. In E. coli, replication of ColE1-type plasmids can be initiated by RNA primers and DNA polymerase I (Itoh and Tomizawa 1979, 1980, 1982). In this case, an RNA transcript of the plasmid binds to the DNA and is processed by RNase H to generate the primers. Persistence of R-loops can also have deleterious effects on transcription and DNA repair (Bentin et al. 2005; Lin et al. 2010). RNase HI activity is required for efficient production of full-length RNA synthesis in E. coli cells deficient in topoisomerase I (Baaklini et al. 2004; Usongo et al. 2008). Recently, R-loops were also shown to stimulate genetic instability of G/C-rich repeats in DNA through a process requiring active transcription. This transcription dependent repeat instability in human cells is stimulated by knockdown of the RNase H1 and RNase H2 genes (Lin et al. 2010).

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12.4


RNase H2 Enzymes

For several years after the discovery of the RNase HI gene in E. coli, it was thought that the organism contained only a single RNase H enzyme. In 1990, identification of a second RNase H protein from the bacteria, named RNase HII, was reported (Itaya 1990). The RNase HII protein turned out to be present in most organisms including bacteria, archaea, and eukaryotes. RNase HII enzymes in prokaryotes have only limited amino acid sequence similarity to their RNase HI counterparts, although they share a similar three-dimensional structure and two-metal-ion mechanism of catalysis (Lai et al. 2000; Chapados et al. 2001; Muroya et al. 2001). The bacterial and archaeal RNase HII enzymes are single subunit proteins possessing the RNase H-like fold and four conserved acidic residues in the active site for coordination of Mg2+ ions (Fig. 12.4). The structural nature and composition of the eukaryotic RNase H2 enzyme remained enigmatic for several years following its discovery in 1969. Identification and purification of RNase H2 homologues from human and S. cerevisiae cells allowed the initial characterization of enzymatic activity, but attempts to clone and express the genes encoding the enzyme in E. coli produced only inactive protein (Frank et al. 1994, 1998a, b; Qiu et al. 1999; Arudchandran et al. 2000). In contrast, RNase HII enzymes from bacterial and archaeal genes produced active enzyme when expressed in E. coli, providing the first hint that eukaryotic RNase H2 may be posttranslationally modified or composed of multiple subunits. Using

Fig. 12.4 RNase H2 enzymes. The structure of (a) the single-domain Methanocaldococcus jannaschii RNase HII shows the prokaryotic RNase HII enzymes share significant structural homology to the RNase HI enzymes, and utilize the two-metal-ion mechanism for phosphoryl hydrolysis (pdbid: 1EKE). (b) Eukaryotic RNase H2 enzymes are a heterotrimeric complex of the RNase H2A, RNase H2B, and RNase H2C proteins (labeled). The catalytic RNase H2A protein has the conserved RNase H-like fold, while the H2B and H2C proteins interweave to form a triple barrel motif that structurally resembles the general transcription factor TFIIF. The eukaryotic RNase H2 complex interacts with the PCNA protein through a motif in the C-terminus of the H2B protein

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affinity tagged protein from S. cerevisiae, Jeong et al. were able to show that eukaryotic RNase H2 is a complex of three proteins that together are necessary and sufficient for activity (Jeong et al. 2004). Deletions of the gene encoding any of the three proteins in yeast resulted in a loss of RNase H2 activity, and overexpression of the three proteins in bacteria produced an active RNase H2 enzyme. These individual proteins are now named RNase H2A, RNase H2B, and RNase H2C in which RNase H2A is the catalytic subunit. Despite the identification of the heterotrimeric complex in yeast, the amino acid similarity to the human proteins was not strong enough to allow for their immediate identification. Finally, in 2006, 37 years after the first description of RNase H activity, Crow et al. identified the genes encoding the individual subunits for the human RNase H2 complex. Coincident with this, they also showed that specific mutations in the three individual genes or in the gene coding for the 30 exonuclease, TREX1, cause the recessive genetic disorder, Aicardi– Goutie`res syndrome (AGS) (Crow et al. 2006a, b). AGS is a neurological disease that mimics congenital viral infection with accompanying increase in interferon a and lymphocytes in the cerebrospinal fluid of affected individuals. It is thought that the underlying pathogenesis of this disease is autoimmune in nature and involves the accumulation of nonprocessed DNA and RNA intermediates after cell death, which leads to the activation of innate immunity (Rice et al. 2007; Crow and Rehwinkel 2009). The available evidence indicates that the nucleolytic action of the RNase H2 complex is necessary to degrade nucleic acids during cell death in order to avoid autoimmune and inflammatory responses. The identification of the three protein components comprising eukaryotic RNase H2 raised immediate structural and biological questions as to the roles for the additional subunits and their contributions to substrate recognition, binding, and catalysis. The RNase H2B and RNase H2C proteins shared little amino acid sequence with other known proteins, had no obvious homologues in bacteria, and it was unclear how they might combine with the RNase H2A catalytic subunit to create an active enzyme. The first information regarding a possible function for these additional protein components came when human RNase H2 was shown to physically interact with proliferating cell nuclear antigen (PCNA) through a short region in the C-terminus of the RNase H2B protein (Chon et al. 2008). PCNA is a protein heavily involved in chromosomal DNA replication and repair, reinforcing the idea of a role for RNase H2 in these biological processes. Although the interaction between the RNase H2 and PCNA proteins appears to be evolutionarily conserved, the interaction seems to have little effect on the activity of the human RNase H2 enzyme (Chon et al. 2008). The crystal structure of the heterotrimeric mouse RNase H2 complex provided the most significant insight into the potential function of the RNase H2B and H2C proteins (Shaban et al. 2010). The overall structure reveals an elongated arrangement of the interacting subunits with the H2C protein in the middle flanked by the H2A and H2B proteins on the ends (Fig. 12.4). The H2C protein consists of 11 b-strands and 2 a-helices and appears to be largely a structural domain that facilitates cohesion of the complex. It weaves together with the N-terminal region (amino acids 1–92) of the H2B protein to form three b-barrels in a motif known as

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a “triple-barrel” (Fig. 12.4) (Gaiser et al. 2000). The triple barrel is formed from a total of 18 parallel and antiparallel b-strands and produces a pseudo twofold axis of symmetry along the central barrel. The intricate interweaving of the b-strands suggests a co-folding of the H2B and H2C proteins rather than a docking of the two, and may explain the necessity for all three subunits to generate a stable, active enzyme. Interestingly, the expression of the RNase H2B and H2C subcomplex is possible in the absence of the catalytic RNase HA subunit (Chon et al. 2008). Whether this happens in vivo and if it means the RNase H2B/H2C subcomplex can act as scaffold for interacting with proteins other than RNase H2A has yet to be determined. The triple-barrel arrangement of the H2B-H2C subcomplex in the mammalian RNase H2 has a very similar structural architecture to that of the RAP34-RAP70 protein subdomains of the general eukaryotic transcription factor IIF (TFIIF) complex (Gaiser et al. 2000). The TFIIF protein complex has a role in transcription initiation and elongation by stabilizing a short RNA:DNA hybrid in the RNA Pol II enzyme active site, inducing promoter DNA wrapping around the polymerase initiation complex (PIC), and mediating protein–protein interactions within the PIC (Robert et al. 1998; Robert and Coulombe 2001; Khaperskyy et al. 2008). The smaller RAP34 (240 amino acids) protein forms the triple-barrel interaction with the N-terminal region of the larger RAP70 (517 amino acids) protein and the mostly a-helical C-terminal domain of RAP70 functions in protein–protein interactions and DNA binding (Kamada et al. 2003). By analogy, the smaller mammalian RNase H2C protein interacts with the N-terminal region of H2B, leaving the mostly a-helical C-terminal region of H2B available for potential interactions with other proteins or to facilitate bending of longer polynucleotide molecules. The striking structural similarity between the RNase H2 and the TFIIF complexes suggests a possible role for RNase H2 in transcription or in interacting with RNA polymerase to process R-loops. The catalytic component of the mammalian RNase H2 complex, RNase H2A, contains an RNase H-like fold and has significant structural homology to the archaeal RNase H2 enzymes and the four acidic residues for metal coordination, and catalysis have been identified (Shaban et al. 2010). Unlike the RNase H1 enzymes that have dual grooves on the protein surface for RNA:DNA hybrid recognition, the mammalian RNase H2 contains only a single cleft that is the active site for substrate binding. It is hypothesized that this structural difference between the RNase H1 and RNase H2 proteins may account for the ability of RNase H2 to recognize single ribonucleotides within a DNA duplex that have mainly B-form helical structure, whereas longer RNA:DNA hybrids adopt intermediate A/B form structure (Shaban et al. 2010). Mutations in human RNase H2 that cause immune dysfunction are found in the genes for all three subunits, suggesting that all components of the complex may contribute to substrate recognition or catalysis (Crow et al. 2006b; Ramantani et al. 2010). Activity assays on purified recombinant proteins containing the most frequently identified mutations in patients showed nearly wild-type activity when tested on a substrate RNA:DNA duplex mimicking an Okazaki fragment

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(Chon et al. 2008; Perrino et al. 2009). Interestingly, a reduction in RNase H activity has been observed in the G37S mutant of the RNase H2A protein on a substrate containing a single ribonucleotide (Shaban et al. 2010). Residue G37 is located in the active site of the enzyme and contributes to a tight turn at the end of one of the b-strands containing two of the catalytic residues. The turn appears to require the flexibility of a glycine residue, and mutation to serine would not permit the torsion angles required to maintain proper active site geometry, thus reducing activity. Whether this selective reduction in catalytic activity between the two substrates is a reflection of the difference in cleaving a ribose–deoxyribose bond versus a ribose–ribose bond or a difference in structural recognition of the two types of substrates currently remains in question. A major distinguishing characteristic in RNase H activities is the ability of RNase H2 to recognize single ribonucleotides in a DNA duplex as a substrate (Eder et al. 1993). This activity may provide another significant clue to the biological function of the enzyme, and it has been proposed that RNase H2 acts to remove single ribonucleotides misinserted during DNA replication. Although it has been generally thought that DNA polymerases are very efficient at discriminating between ribo- and deoxyribonucleotides, recent evidence reveals that the major replicative DNA polymerases in yeast incorporate ribonucleotides into DNA with a significant frequency both in vitro and in vivo (Nick McElhinny et al. 2010a, b). Yeast lacking RNase H2 activity retain higher levels of the incorporated ribonucleotides in their genomic DNA; strengthening the idea that they are removed by an RNase H2-dependent repair mechanism. Additionally, using a yeast strain with a mutated DNA polymerase e that incorporated higher levels of ribonucleotides, it was shown that specific mutations accumulated in the genome in the absence of RNase H2 activity. This data supports a model that single ribonucleotides incorporated during DNA replication may play a role in some cellular signaling or epigenetic process and are metabolized by RNase H2.

12.5 12.5.1

Other RNase H Enzymes RNase HIII Enzymes

A second RNase HII-like enzyme termed RNase HIII has been identified in several bacteria such as B. subtilis, Streptococcus pneumonia, and Aquifex aeolicus (Ohtani et al. 1999a, b). Although RNase HIII is closely related to RNase HII enzymes the distinction in terminology exists because while RNase HII is widely present in most organisms, several bacteria have two genes coding for RNase HII. The protein with the highest amino acid sequence similarity to E. coli RNase HII is given the name RNase HII and the other is called RNase HIII. Biochemical properties of RNase HIII are also more similar to RNase HII than to RNase HI, in that RNase HIII is able to recognize single ribonucleotides in DNA as a substrate and cannot serve as a substitute for E. coli RNase HI (Hou et al. 2007; Liang et al. 2007).

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Fig. 12.5 RNase HIII and viral RNase H. (a) The Bacillus stearothermophilus RNase HIII protein (pdbid: 2D0B) is structurally similar to RNase HII proteins but contains an extended N-terminal domain that has a TBP fold. The TBP domain functions in substrate binding. (b) Viral RNase is part of reverse transcriptase (RT) as in the HIV-1 RT protein (pdbid: 1HYS) which has an RNase HI domain (shown in red and blue) adjacent to the polymerase domain (gray)

Although functionally similar to archaeal RNase HII, RNase HIII proteins are characterized by an extended N-terminal domain that adopts a fold similar to TATAbox-binding proteins (TBP) (Fig. 12.5) (Chon et al. 2006). Biochemical evidence from deletion mutations supports a role in substrate binding for the TBP-like domain, probably through an interaction with the flat surface face, similar to the mechanism for TBP–DNA interactions (Chon et al. 2006).

12.5.2

Viral RNase H Enzymes

In 1970, Baltimore and Temin described the first RNA-dependent DNA polymerases (Baltimore 1970; Temin and Mizutani 1970). Further discoveries revealed that reverse transcriptase-mediated replication of the single-stranded retroviral RNA proceeds via an RNA/DNA hybrid intermediate. This hybrid is processed by viral RNase H into single-stranded DNA (minus strand) to provide a template for the synthesis of double-stranded DNA (plus strand) that is then incorporated into the host genome (Telesnitsky and Goff 1997). In 1971, Peter Hausen’s group discovered that the RNase H activity is contained within the C-terminal end of the viral reverse transcriptase (Molling et al. 1971). The reverse transcriptases of several viruses are therefore bifunctional enzymes containing both polymerase and RNase H activities (Fig. 12.5). This tightly associated RNase H activity plays essential roles in viral replication, as mutations that inactivate the RNase H domain inhibit reverse transcription and therefore viral replication (Repaske et al. 1989; Tisdale et al. 1991). Several HIV virus strains contain acquired mutations in and near the RNase H domain that can increase drug resistance (Delviks-Frankenberry et al. 2010). Consequently, anti-HIV drug targets against the RNase H domain of

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reverse transcriptase are being continuously pursued (for reviews see (Yu et al. 2008; Tramontano and Di Santo 2010)). The reverse transcriptase enzymes from human, murine, and avian viruses have different subunit structures but a conserved orientation of polymerase and RNase H domains. The Moloney murine leukemia virus (M-MLV) enzyme is a single 80 kD polypeptide with N-terminal polymerase and C-terminal RNase H domains. HIV-1 reverse transcriptase is a heterodimer consisting of a p66 domain (66 kDa) and p51 domain (51 kDa). The p66 domain contains an N-terminal polymerase domain, connecting domain, and the C-terminal RNase H domain. These viral RNase H domains share strong structural homology to the bacterial RNase HI and human RNase H1 enzymes. Structural and biochemical studies with bacterial and M-MLV enzymes suggest a role for the positively charged C-terminal helix in substrate recognition (Kanaya et al. 1991; Lim et al. 2002). Although the corresponding helix is absent from the HIV-1 RNase H domain, a region of positively charged residues in the p66 connection domain may perform similar function (Lim et al. 2006). Another important and unique feature defined for the viral RNase H domains is the primer grip region near the active site that contacts the nucleotides in the DNA strand of the RNA:DNA hybrid substrate (Sarafianos et al. 2001; Lim et al. 2006). Mutations in the primer grip region reduce both the enzymes specificity and RNase H activity (Julias et al. 2002; Zhang et al. 2002; Mbisa et al. 2005; McWilliams et al. 2006). Structural similarities between the bacterial and viral RNase H domains support the model for a two-metal-ion-mediated mechanism of catalysis (Schultz and Champoux 2008; Champoux and Schultz 2009). As in the bacterial protein, RNase H domains of HIV-1 and M-MLV reverse transcriptase contain four highly conserved acidic residues (DEDD) within their active sites to coordinate the binding of two divalent Mg2+ ions. Hydrolysis of ribonucleotide substrates likely proceeds through a mechanism similar to other RNase H domains by activation of a water nucleophile through the divalent metal ion in position A and subsequent stabilization of the transition state by both Mg2+ ions (Yang et al. 2006; Nowotny et al. 2007). Although RNase H proteins are generally recognized as sequence-independent endonucleases, viral RNase H domains demonstrate some sequence preferences, likely due to their multiple roles in reverse transcription (Champoux and Schultz 2009). RNase H activity is required to degrade genomic RNA, generate polypurine tract (PPT) primers, and to remove both the PPT and tRNA primers for continued plus strand synthesis during retroviral replication. Viral RNase H has two enddirected modes of cleavage that are unique to itself in addition to the typical endoribonucleolytic activity on RNA:DNA hybrids. The DNA 30 end-directed and the RNA 50 end-directed activities arise as a consequence of association with the polymerase domain. A recessed 30 end of DNA within an RNA:DNA hybrid is recognized as a substrate for the reverse transcriptase polymerase domain. When bound to the polymerase, the RNase H domain is positioned about 15–20 nucleotides downstream of the DNA terminus. Under conditions where the polymerase pauses or fails to extend, the RNase H domain is able to cleave the RNA strand (Furfine and Reardon 1991; Gopalakrishnan et al. 1992; DeStefano et al. 1994).

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Reverse transcriptase also binds a recessed RNA 50 end of an RNA:DNA duplex and cleaves the RNA 13–19 nucleotides from the end (Schatz et al. 1990; Gotte et al. 1995). The shorter spacing in 50 end cleavage has been attributed to a feature of the polymerase active site that interacts with the recessed RNA end.

12.6

Conclusions

RNase H proteins are found in most species, underscoring their biological significance. Since the discovery of RNase H activity in 1969 we have learned much about the structure and mechanism of these proteins, yet we still have little comprehension of their precise biological functions in cells. The major outstanding questions regarding RNase H are currently what the in vivo substrates for these enzymes are, and how do mutations in the human RNase H2 complex lead to disease. In vitro activity assays show that the enzymes are active on a wide variety of possible RNA:DNA hybrids that are likely present in cells including Okazaki fragments, R-loops, single ribonucleotides in DNA, and viral replication intermediates. Mutations in any of the subunits of the human RNase H2 result in severe and chronic autoimmune activation. Interestingly, most of the identified disease-causing mutations seem to have little effect on the catalytic competency of the enzyme. These results might be an indication that the true in vivo substrate has yet to be identified or that RNase H2 protein interactions within an even larger protein complex play an important role in regulation of enzymatic activity. Future work in this area will be critical for understanding the connection between nucleic acid processing and immune dysfunction.

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Chapter 13

Ribonucleoprotein Ribonucleases P and MRP Andrey S. Krasilnikov

Contents 13.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2 RNase P Is a Universal RNA-Based Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.3 Bacterial RNase P . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.4 Archaeal RNase P . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.5 Eukaryotic RNase P . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.6 RNase MRP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.7 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Ribonucleoprotein Ribonuclease (RNase) P and RNase MRP consist of a large RNA component and an essential protein part. RNases P/MRP differ from all other known ribonucleases in that it is their RNA component, not protein that is responsible for the endonucleolytic cleavage of substrates. RNase P is universally essential in all three domains of life; the closely related RNase MRP is a ubiquitous eukaryotic enzyme. RNase P is primarily responsible for the maturation of the 50 -ends of tRNA, whereas RNase MRP is known to be involved in the maturation of eukaryotic rRNA and the degradation of specific mRNAs. Here we discuss available information on functions, structural organization, and mechanisms of substrate recognition and catalysis of RNases P/MRP.

A.S. Krasilnikov Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park, PA 16802, USA e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_13, # Springer-Verlag Berlin Heidelberg 2011

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13.1

A.S. Krasilnikov

Introduction

The enzymes of the RNase P/MRP family constitute a unique class of ribonucleases. While all other known ribonucleases (and indeed the vast majority of enzymes in general) use protein-based catalysis, RNases P/MRP are ribonucleoprotein complexes that rely on their RNA moieties to perform endonucleolytic cleavages of their substrates. RNase P is the best-known member of the RNase P/MRP family. It was first identified as the catalytic entity responsible for the maturation of the 50 -ends of tRNA (Altman and Smith 1971; Robertson et al. 1972). Further characterization of bacterial RNase P revealed that it contained a large RNA molecule and a small protein (Stark et al. 1978; Kole and Altman 1979; Kole et al. 1980; Kole and Altman 1981). Surprisingly, it was the RNA moiety, not the protein that turned out to be responsible for the catalytic activity of the enzyme (Guerrier-Takada et al. 1983). In 1989, Sidney Altman, jointly with Thomas Cech, was awarded the Nobel Prize for their discovery of the catalytic properties of RNA. In this chapter, we briefly review available information on biological roles, structural organization, and mechanisms of substrate recognition and catalysis of RNA-based RNase P, as well as the closely related universal eukaryotic enzyme RNase MRP.

13.2

RNase P Is a Universal RNA-Based Enzyme

tRNAs are initially produced as longer transcripts that must undergo trimming and chemical modifications to form mature molecules (reviewed in (Phizicky and Hopper 2010)). RNase P is universally responsible for the formation of the mature 50 -ends of tRNA, and is essential in all three domains of life (Bacteria, Archaea, and Eukarya) (Altman 2010). While most other enzymes utilize protein-based catalysis, RNase P is practically universally an RNA-based enzyme. There are, however, several known cases of endosymbiotic organelles (in particular, human mitochondria, as well as mitochondria and chloroplasts in at least some plants) in which RNA-based RNase P is replaced by protein-only isozymes (“proteinaceous” RNase P) (Gegenheimer 1996; Thomas et al. 2000a; Holzmann et al. 2008; Holzmann and Rossmanith 2009; Rossmanith and Holzmann 2009; Gobert et al. 2010) and one can expect additional examples of such “unorthodox” RNases P in endosymbiotic organelles. The existence of proteinaceous RNase P isozymes in endosymbiotic organelles highlights one of the major RNase P mysteries: considering that RNase P activity can easily be performed by relatively simple proteins or protein complexes, why, in the vast majority of cases throughout all domains of life, is RNase P conserved as a ribonucleoprotein complex with a catalytic RNA component? In bacteria, the RNase P holoenzyme consists of an RNA component (Fig. 13.1a) and a single small protein (Stark et al. 1978; Kole and Altman 1979, 1981; Kole

13

Ribonucleoprotein Ribonucleases P and MRP

a

CA G A C G G C C G C G C G G UA G C AU A GC G C G G UA C GA A A C G GU

321

b

P12

C

A

CR-II GAG A J11/12 G

S-domain

P12

J12/11 G A CR-III A

A G GG UGC G GU C CA A G CG C A A C C G A GA C GG C A G G GC UG G G U G A C U C C CA G A A U G UG C C A G C U A A C C A GA U CC AA A C AA UA UA G CAC G GG G A A G C C G G A G C A A G G C CA G GG UUC G G C GU GU G G G G CC U C UCGG CC CA AG U AC A U GG A A U AC G G G CA G C CG U G C U G CG GG A G C G U G G CC U C GA G C C A G U G A G C C U C AA UG CR-I G U CC G G U C G U U A AA A A G G G A A U G G G C G G G G A G A CG GC G G AG G G G A U C U C CU CU G C U G C U U C G C C A U G GC G U U A G C A U C G A U G C C A CG AC 5 G A AG C U G AC C A A G 3 U C C A C U U U G A C U G G UA C A U U CG G CCC A

P10/11 P9

CR-II J11/12

P13

A C

C-domain

P14 P7 P5

P15

P16

P4

P4

CR-II

P4

P1

CR-V

Archaeal RNase P RNA

d

J12/11

Domain 2

CR-III

C-domain

P10/11

eP9

CR-IV

P3 P2

S-domain

J11/12

eP15

P7

eP8

P6

CR-I

CR-V

P12

P16

P4

P2

UA C G C G G U G C C G U G U A G C A U G C G C AC G A UC U U G A A G A G G A A C A UA C AA U G G U G G CC G C G G AG U A U U A C UC G C A U C G U A U A UU C C G A C G U A A U A GUUCCG U U A G G AG A AG C G AG GU UC C U C G U C C G C U U C ACA G C C U C C UG C C G GG G A U UU AGG U A U AA C G G U G G G UC UU C U U U CG A A A U A AU U U A G G UU A U G CAGGU G UG G G CG G G A U C A C C AC C C U A G GUCC A U C U G AA U AA AG G G A C AU U C AAU A U C C U

P5 P15

P8

P6

Bacterial RNase P RNA

c

J12/11

P7

P18

P1

C-domain

CR-III

P10/11

CR-IV

P3

S-domain

P9

P17

P8

GU G G A U C GC C U G C G G G G A A A G G U G C A U G C GCGG U G A A C G CC G C A G G AA UU G AG U G G CC A G C C U C C G C GA C C GG G A C A CG GA C AC G U A G A U G G A A C A A C CA G GG G C C C A G A C U G C G AGG G AGG C A AG UU AU G A G G CC CGC CU A G U C G G G G U G C A A G G C CG GGGCC U C G CC A G C G CC C C A C CCGG CC CG G G A C GU G GC A C G G U G GU G G GGCCAC C G U UG C G G CC G A U C A CG U UC U A AG G A U G G C A G C U CC CG A G U G G G G CU A U C CC G G C G C G C G C G C GU A G U A C 5 U A G G C G A G G G GG C U A G 3 A UCC GC U CC CCC G A U A AU C G G G C G G A G A UG A

A U G A C U U G U C G C C C G CU C C U G A A C U G C G A G U G C GG U G G U U G A G U G C A A U C G U A UU G G CA A A GU UG C U GG 5 G U G G A A C A G U GG U A A GC A AC G A C C U U G U C G U C G UC C C C A 3 U U A G C C A U AG

CR-I

P4

CR-IV

P3

P2

P1

P4

eP19

CR-V

S. cerevisiae RNase P RNA

AU U U UCU CG U GG A CC G CC G A U G C G AU GU UG UA U A C UG CA G C C G U GU G UG G A U G CU C UG G UU A U UU A C U U C G G U A C U G G A U U C C G U UU U 5’-GARAG-3’ U A U G AU A U C U A A G G U A UG U C A G C AA U U A A A A GCG C A G C C AG AAUU A U G C A CA A C AA A C U UU C C G UU AU C U A A UU A UC U A A AA G A U A AA U UU A G AAU G U G A U U A U U U U GCU UUGG G U UUUUACUC A U U A A A AU G A G CGA AGC U U U G A U UA AC G A A C C AA A A U A C G UG G C CC C G C UU U U A GU A U CUC AG A UG U A G C A 5 A AUCC AU G A CC A A G UG A AC CU U A G G U A CCU G G U U CU CC A CA A UCU CC AU U C G AG G U A 3

ymP7

Domain 1

ymP6

eP15

ymP5

mCR-I P4

mCR-IV

P3

P2

P1

eP19

P4 mCR-V

S. cerevisiae RNase MRP RNA

Fig. 13.1 Secondary structure diagrams for bacterial, archaeal, and eukaryotic RNase P and RNase MRP RNAs. (a) Bacterial RNase P RNA (A-type, E. coli). (b) Archaeal RNase P RNA (A-type, P. horikoshii). (c) Eukaryotic RNase P RNA (S. cerevisiae). (d) RNase MRP RNA (S. cerevisiae). The diagrams and nomenclature of the structural elements are based on (Haas et al. 1991, 1996a, b; Brown et al. 1996; Frank et al. 2000; Li et al. 2002; Walker and Avis 2004; Torres-Larios et al. 2006; Esakova et al. 2008; Esakova and Krasilnikov 2010). Dotted lines separate the specificity (S-) and the catalytic (C-) domains. Solid lines represent tertiary interactions. Red arrows point to the position of the conserved uridine that is involved in the coordination of a catalytic metal ion in bacterial RNase P (Reiter et al. 2010)

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et al. 1980). Both the RNA and the protein are essential in vivo (Kirsebom et al. 1988; Gossringer et al. 2006 and references therein) and are required for RNase P activity under physiological conditions (Kole and Altman 1979). The RNA component – the catalytic moiety of bacterial RNase P – can perform proper substrate cleavage without the protein component in vitro, but only under conditions of elevated ionic strength or in the presence of polyamines; divalent metal ions (normally Mg2+) are absolutely required for RNase P activity (Guerrier-Takada et al. 1983). It should be noted that the size of the RNA component of bacterial RNase P, while considerable (several hundred nucleotides), corresponds to the size of the mRNA of a relatively small (10–20 kDa) protein. Thus, bacterial RNase P represents a relatively “cheap” way of making an enzyme as it comprises a small protein component plus an RNA molecule that is equivalent to a small mRNA, but does not even require subsequent translation. The relative ease of producing bacterial RNase P could be a possible explanation of the conservation of RNase P as an RNA-based enzyme in bacteria. However, this argument does not apply to the more complex RNases P from higher organisms. Archaeal RNase P contains more constituents than its bacterial counterpart: it is comprised of an RNA component resembling that of the bacterial enzyme (Fig. 13.1b) (Brown 1999) and four or five protein components (Hall and Brown 2002; Hartmann and Hartmann 2003; Kouzuma et al. 2003; Fukuhara et al. 2006; Pulukkunat and Gopalan 2008; Cho et al. 2010). RNA components of some archaeal RNases P were shown to be capable of catalysis without proteins in vitro, but only under conditions of extremely high ionic strengths (Pannucci et al. 1999; Kouzuma et al. 2003; Tsai et al. 2006). Eukaryotic RNase P (Walker et al. 2010) is a large ribonucleoprotein complex containing an RNA component that resembles the bacterial and archaeal RNase P RNAs (Chen and Pace 1997) (Fig. 13.1c) and multiple (nine in Saccharomyces cerevisiae [Esakova and Krasilnikov 2010 and references therein])) essential protein components which constitute the bulk of the enzyme. The RNA component of eukaryotic RNase P is the enzyme’s catalytic moiety (Thomas et al. 2000b; Kikovska et al. 2007), although the level of the catalytic activity demonstrated by the RNA component of eukaryotic RNase P in the absence of proteins is extremely low (five or six orders of magnitude lower than that for bacterial RNase P RNA). The reasons for the dramatically increased complexity of the protein part of eukaryotic RNase P are not clear. The presence of RNA-based RNase P in practically all organisms and the similarity of its RNA components throughout all three domains of life indicate that RNase P is an ancient enzyme and possibly a remnant of the hypothetical RNA World (Gilbert 1986). The major activity of RNase P – cleavage of tRNA precursors – is the only known RNase P function conserved in all three domains of life. In addition to tRNA precursors, RNase P cleaves multiple other naturally occurring substrates that typically (but not always) resemble tRNA. Among such substrates in bacteria are 4.5S RNA (Guerrier-Takada and Altman 1984a; Peck-Miller and Altman 1991),

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tmRNA (Komine et al. 1994), operon mRNAs (Alifano et al. 1994; Li and Altman 2003, 2004), OLE RNA from extremophilic bacteria (Ko and Altman 2007), and transient structures in riboswitches (Altman et al. 2005; Seif and Altman 2008). Human RNase P was shown to cleave off a tRNA-like segment in the 30 -end region of the nascent metastasis-associated lung adenocarcinoma transcript 1 (MALAT1) (Wilusz et al. 2008). In yeast, the role of RNase P in the splicing-independent maturation of intron encoded box C/D small nucleolar RNAs was suggested (Coughlin et al. 2008). In addition, recent reports suggest that human RNase P (or at least several of its components) may be involved in transcription by RNA polymerases I and III (Reiner et al. 2006; Jarrous and Reiner 2007; Reiner et al. 2008), a surprising and very interesting observation that needs further investigation. It seems very likely that additional roles for RNase P in both bacteria and eukaryotes are yet to be identified and characterized. The existence of multiple and diverse essential roles for RNase P in the cell could provide a very plausible answer to the question of why this enzyme has retained its ancient RNA-based form in practically all organisms.

13.3

Bacterial RNase P

Compared to proteins, RNA is made up of a very limited number of building blocks. However, the ability of RNA to form a wide variety of hydrogen bonds, a multitude of non-Watson–Crick base pairs and base triplets, combined with the inherent backbone flexibility, allow RNA to fold into structures that are sufficiently complex to perform substrate recognition and catalysis. The RNA component of bacterial RNase P can adopt and maintain a well-defined and highly ordered threedimensional conformation that allows it to perform substrate recognition and cleavage even in the absence of the protein component. The RNA component of bacterial RNase P folds into a complex threedimensional structure that includes functionally important regions responsible for substrate recognition and catalysis. These regions are stabilized by an extensive network of interactions between variable auxiliary elements serving as structural buttresses (Krasilnikov et al. 2003, 2004; Torres-Larios et al. 2005; Kazantsev et al. 2005; Reiter et al. 2010), reviewed in (Mondragon 2010). Sequences of the RNA components of bacterial RNases P are very diverse (Brown 1999). However, the secondary structures of bacterial RNase P RNAs are very well conserved and almost all known bacterial RNase P RNAs are essentially variants of the two major types: the most common A-type (Ancestral, Fig. 13.1a) and the B-type (Bacillus, found in low G+C Gram-positive bacteria) (GuerrierTakada and Altman 1984b; James et al. 1988; Brown et al. 1991; Haas et al. 1991; Brown and Pace 1992; Haas et al. 1994, 1996a). The major difference between the two types of bacterial RNase P is in the arrangements of the auxiliary elements that stabilize the functionally important and structurally conserved core regions (Krasilnikov et al. 2004; Torres-Larios et al. 2006). Individual auxiliary structural

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elements within each of the two types of RNase P demonstrate significant variation in their sizes, or can be completely missing; however, the general arrangement of the structural buttresses appears to be well conserved within each type. Despite differences in their structural organizations, the RNA components of the two types of bacterial RNase P can be interchangeable (Guerrier-Takada et al. 1983; Waugh and Pace 1990; Wegscheid et al. 2006). While there is significant variability in RNase P RNA sequences, five regions in RNase P RNA show a high degree of conservation (Chen and Pace 1997). These regions are termed Conserved Regions (CR-I, CR-II, CR-III, CR-IV, and CR-V) (Fig. 13.1); several nucleotides found there are conserved not only in bacteria, but in all three domains of life (Chen and Pace 1997; Brown 1999). Not unlike large proteins, large RNA molecules can often be divided into structural domains. There are two independently folding structural domains in bacterial RNase P RNA (Pan 1995; Loria and Pan 1996, 1999): the specificity domain (S-domain) and the catalytic domain (C-domain) (Fig. 13.1a). When combined, individually produced and folded S- and C-domains can form a bimolecular complex that is capable of specific substrate cleavage (Pan 1995; Loria and Pan 1996). The catalytic domain contains the active site and binds the protein component, but by itself has little affinity for RNase P substrates (Odell et al. 1998; Loria and Pan 1999, 2001; Mobley and Pan 1999). The specificity domain confers the specificity for pre-tRNA substrates: it is responsible for the recognition of the TCC and D-loops in pre-tRNA and serves to bind and position pre-tRNA for cleavage by the catalytic domain (Loria and Pan 1997; Odell et al. 1998; Mobley and Pan 1999; Qin et al. 2001; Reiter et al. 2010). The recognition of the pre-tRNA by the specificity domain of bacterial RNase P RNA relies on the overall shape of the substrate molecule, as opposed to its sequence. The non-helical module containing phylogenetically conserved regions CR-II and CR-III (Fig. 13.1a) folds into a complex structure comprised of two interweaving T-loop motifs (Krasilnikov et al. 2003; Krasilnikov and Mondragon 2003) and plays a key role in the substrate recognition. This non-helical module participates in the formation of a phylogenetically and structurally conserved opening (Krasilnikov et al. 2003, 2004; Reiter et al. 2010). The substrate enters this opening, and the complementation of the shape of the opening and the shape of the substrate (mostly the TCC-loop), along with the stacking of nucleobases of the tRNA’s D- and TCC-loops with unstacked nucleobases in the S-domain, serve to provide specificity of the pre-tRNA recognition (Reiter et al. 2010). It is highly likely that the general mode of the D- and TCC-loop recognition by the S-domain is conserved throughout the three domains of life (Esakova and Krasilnikov 2010 and references therein). The catalytic domain of bacterial RNase P RNA also participates in the recognition of the substrate. The recognition of the pre-tRNA substrate by the C-domain involves base pairing between phylogenetically conserved nucleotides of the P15 loop (Fig. 13.1a) and the 30 -CCA motif found in the acceptor stem of most of the bacterial tRNA transcripts (Guerrier-Takada et al. 1984; McClain et al. 1987;

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Kirsebom and Svard 1994; Oh and Pace 1994; LaGrandeur et al. 1994; Svard et al. 1996; Oh et al. 1998; Heide et al. 1999; Busch et al. 2000; Wegscheid and Hartmann 2006; Reiter et al. 2010). As opposed to the recognition of the universal tRNA features by the S-domain, 30 -CCA recognition by the C-domain of bacterial RNase P RNA does not appear to be conserved in higher organisms where the 30 CCA is added posttranscriptionally. The mutual orientation of the specificity and catalytic RNA domains in bacterial RNase P is stabilized by tertiary interactions. The well-defined juxtaposition of the two RNA domains, combined with the specificity of the two domains’ interactions with pre-tRNA, ensures the required degree of substrate selectivity, and provides a “measuring mechanism” that allows for the proper cleavage of pre-tRNA by RNase P RNA even in the absence of the protein component found in the bacterial RNase P holoenzyme. While the RNA component of bacterial RNase P is capable of proper cleavage of pre-tRNA substrates in the absence of any proteins, such cleavage occurs only under conditions of elevated ionic strength or in the presence of polyamines (Guerrier-Takada et al. 1983). The activity of bacterial RNase P under physiological conditions and in vivo requires the presence of a small basic protein, the protein component of bacterial RNase P (Schedl and Primakoff 1973; Stark et al. 1978; Kole and Altman 1979; Kole et al. 1980; Kole and Altman 1981; Guerrier-Takada et al. 1983; Reich et al. 1988; Kirsebom et al. 1988; Gossringer et al. 2006 and references therein). The protein component of bacterial RNase P is well conserved phylogenetically and forms a globular abbbaba fold containing a rare left-handed bab crossover (Stams et al. 1998; Spitzfaden et al. 2000; Kazantsev et al. 2003). The protein component is involved in extensive interactions with the catalytic (C-) domain of RNase P RNA and with the 50 -leader of pre-tRNA, but only in limited, if any, interactions with the product tRNA (Reiter et al. 2010) (Fig. 13.2). While the protein component is positioned in the general vicinity of the substrate’s scissile bond, the distance between the protein and the cleavage site appears to be too large

a tRNA

P16/P17 P15 3’ 5’ P3

tRNA P1 P4

P13

5’-leader C protein

b

P12 J11/12, J12/11

P4

CR-IV

P2

P1

P14 P8/P9

ader protein

5’-le

P12 P3

J11/12, J12/11 P10/11 P6

P15 P16/17

Fig. 13.2 Crystal structure of bacterial RNase P holoenzyme in complex with product tRNA and 50 -leader (Reiter et al. 2010). Individual elements are color-coded. The nomenclature of RNase P RNA elements matches that in Fig. 13.1a

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for the direct involvement of the protein in catalysis (Reiter et al. 2010). The localization of the protein component obtained using X-ray crystallography (Reiter et al. 2010) is in agreement with its previously suggested roles in helping to overcome electrostatic repulsion between the substrate and the catalytic RNA (Reich et al. 1988) and in aiding substrate recognition and catalysis by discriminating between substrate and product (Kurz et al. 1998; Crary et al. 1998; Niranjanakumari et al. 1998; Rueda et al. 2005; Sun et al. 2006; Hsieh and Fierke 2009; Koutmou et al. 2010). The overall structure of the bacterial RNase P holoenzyme is in a good general agreement with the results of molecular modeling (Masquida et al. 2010 and references therein). While a complete mechanism of RNase P cleavage is yet to be characterized, a body of biochemical data (reviewed in (Kirsebom and Trobro 2009)) and the available crystallographic data (Kazantsev et al. 2005, 2009; Torres-Larios et al. 2005; Reiter et al. 2010) strongly indicate that, similar to other large ribozymes, RNase P utilizes a two-metal-ion catalytic mechanism (Stetz and Steitz 1993; Stahley and Strobel 2005; Toor et al. 2008; Reiter et al. 2010). Several nucleotides in the P4 stem, including the phylogenetically conserved bulged uridine (Fig. 13.1), participate in the formation of the RNase P active site (Reiter et al. 2010); judging by the conservation of this part of RNase P RNA, the active site is likely to be conserved throughout the three domains of life. The putative active site metal ions (normally, Mg2+) appear to be coordinated by phosphate oxygens in the RNase P RNA P4 stem as well as those of the scissile phosphate of the substrate, and – for one of the ions – by the O4 oxygen from the conserved bulged uridine in the P4 stem (Reiter et al. 2010). It appears that RNase P RNA serves as a scaffold for the proper positioning of the substrate and the active site metal ions (Reiter et al. 2010).

13.4

Archaeal RNase P

Archaeal RNase P is more complex and not as well characterized as its bacterial counterpart; however, important information related to its structural organization, roles of individual parts, and its evolutionary relation to the even more complex eukaryotic RNase P is emerging (recently reviewed in (Jarrous and Gopalan 2010)). There are two major types of archaeal RNases P, divided according to the secondary structures of their RNA components: the more common A-type (Ancestral, Fig. 13.1b) and the M-type (represented by Methanococcus RNases P) (Harris et al. 2001 and references therein). The A-type archaeal RNase P RNA resembles that of the A-type bacterial RNase P (Fig. 13.1a, b) and appears to retain many peripheral elements involved in the stabilization of the functionally important core in bacterial enzymes. At the same time, these archaeal RNase P RNAs are typically missing some of the elements serving as structural buttresses in bacteria, while possessing unique structural features (such as elaborate elements incorporated into the P12 stem, Fig. 13.1b). Similar to their bacterial counterparts, A-type archaeal RNase P RNAs can be

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catalytically active in vitro in the absence of proteins, albeit this activity requires extremely high ionic strengths (Pannucci et al. 1999; Kouzuma et al. 2003; Tsai et al. 2006); changes toward the bacterial consensus can significantly improve the level of activity (Li et al. 2009). The M-type archaeal RNase P has lost almost all of the elements that serve as structural buttresses in bacterial RNase P RNA, but still possesses all essential core elements (Harris et al. 2001). While it does not exhibit any noticeable catalytic activity without proteins (Pannucci et al. 1999 and references therein), it can cleave substrates attached in cis; replacement of its specificity domain with a bacterial one also results in substrate cleavage, indicating that the lack of noticeable catalytic activity is mostly due to defects in substrate binding (Pulukkunat and Gopalan 2008). In addition to the two major (A- and M-) types of archaeal RNase P, at least some Pyrobaculum species and the related crenarchaea Caldivirga maquilingensis and Vulcanisaeta distributa possess a “minimized” form of RNase P, where the specificity (S-) domain of the RNA component is replaced with a much more simple, mostly helical structure (Lai et al. 2010). Typical archaeal RNase P holoenzymes contain at least four proteins (Hall and Brown 2002): aPop4, aPop5, aRpp1, and aRpr2. These proteins are homologous to eukaryotic RNase P proteins Pop4, Pop5, Rpp1, and Rpr2 (see next section). Recent data indicate that at least some archaeal RNases P contain a fifth protein, a ribosomal protein L7Ae (related to eukaryotic RNase P protein Pop3) (Fukuhara et al. 2006; Cho et al. 2010). The protein component aRpr2 appears to be absent in the “minimized” RNases P of the Pyrobaculum species (above) (Lai et al. 2010). High resolution structures of all proteins associated with archaeal RNase P have been determined (Boomershine et al. 2003; Sidote and Hoffman 2003; Numata et al. 2004; Takagi et al. 2004; Sidote et al. 2004; Kakuta et al. 2005; Fukuhara et al. 2006; Wilson et al. 2006; Amero et al. 2008) (reviewed in (Esakova and Krasilnikov 2010)). The proteins fold into a diverse set of structures (Fig. 13.3); none of them is directly related to the bacterial RNase P protein. Structural organizations of aPop5 and L7Ae (aPop3) somewhat resemble that of the bacterial protein, but while their secondary structure elements are positioned in a somewhat similar way, the connectivity between them is different, and the proteins do not appear to be evolutionarily related. At the same time, it cannot be excluded that this weak structural similarity may play a role in the reported activation of archaeal RNase P RNA by bacterial RNase P protein (Nieuwlandt et al. 1991; Pannucci et al. 1999). The details of the structural organization of the archaeal RNase P holoenzyme, in particular, the details of RNA-protein interactions, are not yet known. Active (although not necessarily structurally homogeneous) archaeal RNase P can be reconstituted from individual components (Boomershine et al. 2003; Kouzuma et al. 2003; Tsai et al. 2006; Terada et al. 2006; Cho et al. 2010). The four established protein components act in pairs: aPop4 + aRpr2 and aPop5 + aRpp1 (Tsai et al. 2006). The aPop5 + aRpp1 pair appears to be involved in interactions with the catalytic domain of RNase P RNA, playing a role in substrate cleavage/product release, while the aPop4 + aRpr2 pair is likely involved in the stabilization of the structure/orientation of the specificity domain (Tsai et al. 2006;

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C

N

C C N aPop3

N aPop5

aPop4 N C

N C

aRpp1

Zn2+

aRpr2

Fig. 13.3 Crystal structures of proteins associated with archaeal RNase P (PDB ID: 2CZW, 1V76, 2AV5, 1V77, 1X0T). a-helices are shown in red; b-strands in yellow; loops in green; Zn2+ ion is shown in gray

Fukuhara et al. 2006; Pulukkunat and Gopalan 2008; Xu et al. 2009; Honda et al. 2010). The fifth protein component of archaeal RNases P, L7Ae (aPop3), was reported to increase the thermal stability of some RNases P and the efficiency of catalysis (Fukuhara et al. 2006; Terada et al. 2006; Cho et al. 2010). The archaeal RNase P proteins are likely to perform the roles of the single bacterial RNase P protein, and also compensate for the partially (in the A-type RNase P) or almost completely (in the M-type RNase P) missing RNA–RNA interactions that serve to stabilize the three-dimensional structure of RNase P RNA in bacteria; however, further studies are necessary to advance our understanding of the roles of proteins in the structure and function of archaeal RNase P.

13.5

Eukaryotic RNase P

Eukaryotic RNase P is considerably more complex than its bacterial and archaeal counterparts. In addition to the apparently catalytic RNA component that resembles the RNA component in bacterial and archaeal RNases P (Fig. 13.1c), eukaryotic RNase P contains multiple protein subunits, which constitute the bulk of the enzyme (recently reviewed in (Esakova and Krasilnikov 2010)). The best characterized eukaryotic RNase P from S. cerevisiae has nine known protein components: Pop1 (100.5 kDa) (Lygerou et al. 1994), Pop3 (22.6 kDa) (Dichtl and Tollervey 1997), Pop4 (32.9 kDa) (Chu et al. 1997), Pop5 (19.6 kDa), Pop6 (18.2 kDa), Pop7 (15.8 kDa), Pop8 (15.5 kDa) (Chamberlain et al. 1998), Rpp1 (32.2 kDa) (Stolc and Altman 1997), and Rpr2 (16.3 kDa) (Chamberlain et al. 1998). All of these proteins are essential for the viability of yeast (Lygerou et al.

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1994; Dichtl and Tollervey 1997; Chu et al. 1997; Stolc and Altman 1997; Chamberlain et al. 1998). Human RNase P has a similar composition (Jarrous and Reiner 2007; Esakova and Krasilnikov 2010). All known archaeal RNase P proteins have homologues in eukaryotic RNase P (see previous subchapter). Furthermore, the protein Pop8 appears to be evolutionarily related to Pop5 (Rosenblad et al. 2006). In addition to the proteins present in archaeal RNase P and the closely related Pop8, eukaryotic (yeast) RNase P has acquired additional protein components Pop1, Pop6, and Pop7. The RNA component of eukaryotic RNase P has the same major structural elements that are responsible for substrate recognition and catalysis in bacterial and archaeal RNases P, consistent with its catalytic role. However, practically all RNA elements that are involved in tertiary RNA–RNA interactions in bacterial and A-type archaeal RNases P appear to be missing in eukaryotes (Fig. 13.1). The extremely low level of eukaryotic RNase P RNA activity in the absence of proteins (Kikovska et al. 2007) could be explained by this absence of the RNA buttresses, which stabilize bacterial and A-type archaeal RNase P RNAs and allow them to demonstrate an appreciable level of activity even without proteins. Accordingly, one can suggest an obvious role for eukaryotic RNase P proteins in stabilizing the structure of the catalytic RNA component. Indeed, unlike bacterial RNase P RNA, eukaryotic RNase P RNA was shown to be unable to form a compact structure in the presence of Mg2+ (which allows bacterial RNase P to fold properly) without proteins (Marquez et al. 2006), and results of footprinting studies indicate that the presence of proteins is required for the proper folding of key parts of RNase P RNA in eukaryotes (Esakova et al. 2008). However, the number of eukaryotic RNase P proteins would seem to be excessive if they were to serve only this role. Indeed, eukaryotic RNase P RNA is not the only type of RNase P RNA that apparently lacks RNA buttresses: the M-type of archaeal RNase P is similar to eukaryotic RNase P RNA in this respect. Yet the M-type RNase P manages to maintain the structure of its RNA using a much smaller number of proteins than is present in eukaryotic RNase P. In these terms, it is unclear why eukaryotic RNase P (which has homologues of all archaeal proteins) needs multiple extra proteins. Thus, it is quite likely that at least some of the eukaryotic RNase P proteins (particularly the ones that do not have archaeal homologues: Pop1, Pop6 and Pop7) have functions going beyond the simple stabilization of RNA structure. The complex protein part of eukaryotic RNase P was suggested to allow processing of a broader range of substrates (Marvin and Engelke 2009). Indeed, it was suggested that yeast RNase P is involved in the maturation of intron encoded box C/D small nucleolar RNAs (Coughlin et al. 2008) – substrates that are unlikely to resemble the canonical RNase P substrate, pre-tRNA, and additional eukaryotic RNase P substrates are likely to be identified. Another possible explanation for the increased complexity of the protein part of RNase P in eukaryotes is that the proteins serve to fine-tune and improve substrate recognition, which could be essential given the more complex intracellular environment of the eukaryotic cell. An additional (and very likely) function for the proteins might be their involvement in interactions with other parts of the cellular machinery to facilitate the proper

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localization of the RNase P complex or to regulate its activity. Further studies are needed to clarify roles of proteins in eukaryotic RNase P. Available structural information on eukaryotic RNase P is still very limited. Certain assumptions can be inferred from information available on bacterial and archaeal enzymes. While the RNA component of eukaryotic RNase P is noticeably different from its bacterial counterpart, the universally phylogenetically conserved regions in eukaryotic RNase P (CR-I, CR-II, CR-III, CR-IV, and CR-V, Fig. 13.1) are very likely to adopt folds similar to those of corresponding regions observed in crystal structures of bacterial RNase P (Krasilnikov et al. 2003, 2004; Torres-Larios et al. 2005; Kazantsev et al. 2005; Reiter et al. 2010). However, it should be noted that the distinct protein compositions of the eukaryotic and bacterial RNases P, and the diverged auxiliary RNA elements severely limit the predictive power of such an approach. Available structural information on archaeal RNase P proteins aPop3, aPop4, aPop5, aRpp1, and aRpr2 (reviewed in (Esakova and Krasilnikov 2010)) provides a glimpse of the structural organization of their eukaryotic homologues, but does not allow establishing their structural or functional roles. Our current knowledge of protein–protein and protein–RNA interactions in eukaryotic RNase P is primarily based on the results of UV cross-linking (Pluk et al. 1999; Jiang et al. 2001), yeast two- or three-hybrid studies (Jiang and Altman 2001; Jiang et al. 2001; HouserScott et al. 2002), and GST pull-down experiments (Welting et al. 2004; Aspinall et al. 2007). However, the available low-resolution data on protein–protein and protein–RNA interactions is not entirely consistent and does not yet provide a convincing low-resolution map of interactions between the components of eukaryotic RNase P (Esakova and Krasilnikov 2010). High-resolution structural information is currently available only for the P3 subdomain of the RNA component which was recently crystallized in a complex with proteins Pop6 and Pop7 (Perederina et al. 2007, 2010a, b). The helix–loop–helix subdomain P3 is a universal and unique essential feature of all eukaryotic enzymes of the RNase P/MRP family (Esakova and Krasilnikov 2010). It replaces the helical stem P3 found in bacterial and archaeal enzymes (Fig. 13.1). In yeast, this subdomain interacts with protein components Pop6, Pop7 (Perederina et al. 2007) and, likely, Pop1 (Ziehler et al. 2001). Pop1, Pop6, and Pop7 do not have homologues in bacterial or archaeal RNases P; it was suggested that the helixloop-helix P3 subdomain evolved from the P3 helix found in bacterial and archaeal enzymes as a protein-binding hub that allowed binding of additional protein components (Perederina and Krasilnikov 2010). Proteins Pop6 and Pop7 form a heterodimer that is involved in extensive interactions with the P3 subdomain RNA (Perederina et al. 2007, 2010b) (Fig. 13.4); the observed structural organization of this RNA–protein complex is likely to be conserved in the eukaryotic enzymes of the RNase P/MRP family (Perederina et al. 2010b; Hands-Taylor et al. 2010). Despite their very diverse sequences, proteins Pop6 and Pop7 form similar folds (Perederina et al. 2010b) and appear to originate from common archaeal proteins of the Alba family (Wardleworth et al. 2002; Aravind et al. 2003; Perederina et al. 2010b; Perederina and Krasilnikov 2010). In summary, while there is a growing

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P3 RNA 3’ N

N 5’

C C Pop7 Pop6

Fig. 13.4 Crystal structure of the S. cerevisiae P3 RNA subdomain in a complex with proteins Pop6 and Pop7 (Perederina et al. 2010b). RNA is shown in red; Pop6 in light blue; Pop7 in purple. Missing or unresolved regions are shown as dotted lines. In the orientation shown, the distal terminal loop of the P3 subdomain (Fig. 13.1c, d) would be located on the left

body of information on the structural organization of eukaryotic RNase P, additional studies are needed to shed light on its structural organization and the roles of its numerous components.

13.6

RNase MRP

RNase MRP (Mitochondrial RNA Processing) was first identified as an entity involved in the maturation of RNA primers for mitochondrial DNA replication (Chang and Clayton 1987); further studies identified RNase MRP substrates outside the mitochondria and characterized this enzyme as a universal eukaryotic ribonuclease closely related to RNase P (reviewed in (Esakova and Krasilnikov 2010)). Mitochondrial RNase MRP (Chang and Clayton 1987; Stohl and Clayton 1992; Topper et al. 1992), which represents a small fraction of the cellular RNase MRP, has a distinct composition and substrate specificity (Lu et al. 2010), and is not yet well characterized structurally or functionally. This subchapter focuses on the more studied and more abundant nucleolar RNase MRP. RNase MRP, an exclusively eukaryotic ribonucleoprotein complex, was identified in practically all eukaryotes analyzed for its presence (Piccinelli et al. 2005; Rosenblad et al. 2006), and thus appears to be a universal eukaryotic enzyme. RNase MRP is similar and apparently evolutionarily related to eukaryotic RNase P (Piccinelli et al. 2005; Rosenblad et al. 2006; Zhu et al. 2006; Woodhams et al. 2007). Like RNase P, RNase MRP contains a large RNA component and multiple protein subunits. Most of the protein subunits of RNase MRP are also found in RNase P. S. cerevisiae RNase MRP has ten known essential protein components: Pop1, Pop3, Pop4, Pop5, Pop6, Pop7, Pop8, Rpp1, Snm1, and Rmp1; eight of these proteins are also components of RNase P (Chamberlain et al. 1998). RNase MRP

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protein Snm1 (Schmitt and Clayton 1994) has similarities to RNase P protein Rpr2, while RNase MRP protein Rmp1 (Salinas et al. 2005) has no apparent counterpart in RNase P. Most of the human RNase P and RNase MRP proteins are also shared (reviewed in Esakova and Krasilnikov 2010; Jarrous and Gopalan 2010). The RNA components of RNases MRP and P demonstrate clear similarity (Fig. 13.1c, d). The catalytic (C-) domain of RNase P and Domain 1 of RNase MRP share essentially all main secondary structure elements; several nucleotides that are phylogenetically conserved in the C-domain of RNases P (Chen and Pace 1997) are also found in RNases MRP (Lopez et al. 2009). The RNA–protein interactions in the C-domain of RNase P and Domain 1 of RNase MRP appear to be very similar (Esakova et al. 2008); moreover, key subdomains P3 in RNases P and MRP (Fig. 13.1c, d) were shown to be interchangeable (Lindahl et al. 2000). While the catalytic ability of the RNA component of RNase MRP has not yet been demonstrated, the very close similarity between RNase MRP Domain 1 and the catalytic RNA domain of eukaryotic RNase P strongly indicates that RNase MRP uses RNA-based catalysis similar to that of RNase P. Whereas the catalytic RNA domain of RNase P has a clear counterpart in RNase MRP (Domain 1), the specificity (S-) domain of RNase P is very distinct from RNase MRP Domain 2 (Fig. 13.1c, d). While it is not clear whether Domain 2 in RNase MRP plays the substrate recognition role parallel to that of the S-domain in RNase P, the drastic divergence between the RNase P S-domain and RNase MRP Domain 2 immediately suggests that the two related enzymes have distinct substrate specificities. Substrate selection by RNase MRP is not yet understood and contributions of the protein and RNA components to the specificity of this enzyme are not clear. In vitro substrate selection assays indicate that RNase MRP cleaves single-stranded RNA in a sequence-specific manner (Esakova et al. 2011). There is a weak conservation of the sequence in the immediate vicinity of the cleavage site (notably, a cytosine at position +4 appears to be required for RNase MRP cleavage in S. cerevisiae); however, substrate recognition in vivo is likely to require additional, yet unidentified factors such as specific proteins bound to substrate RNA (Esakova et al. 2011). The best characterized RNase MRP function is its participation in the maturation of nuclear rRNA. In yeast, RNase MRP is known to cleave the internal transcribed spacer 1 (ITS1) at the specific site A3 of the pre-rRNA. RNase MRP cleavage at the A3 site leads to subsequent (apparently exonucleolytic) trimming of pre-rRNA, which results in the formation of the predominant form of the mature 50 -end of 5.8S rRNA (Schmitt and Clayton 1993; Chu et al. 1994; Clayton 1994; Lygerou et al. 1994, 1996). In addition to the predominant form of the mature 5.8S rRNA, there exists a fraction of a slightly (seven nucleotides in yeast) longer form, 5.8SL rRNA. The significance of the existence of the two forms of mature 5.8S rRNA is not clear, but it appears to be common in eukaryotes (Bowman et al. 1983; Smith et al. 1984; Henry et al. 1994). RNase MRP cleavage at the A3 site is required for the formation of the more common shorter version of 5.8S rRNA, but not the longer one in S. cerevisiae and D. melanogaster (Schmitt and Clayton 1993; Schneider et al. 2010); the role of RNase MRP in the 5.8S rRNA maturation pathway, which

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involves pre-rRNA cleavage at the A3 site is likely to be universal, but it does not appear to be essential for cell viability. In addition to its relatively well-characterized role in the maturation of 5.8S rRNA, RNase MRP was recently suggested to play a much broader role in the canonical rRNA maturation pathway in general: inactivation of RNase MRP severely affected the abundance of all early intermediates in the canonical rRNA maturation pathway in yeast (Lindahl et al. 2009); a similar effect was observed in D. melanogaster (Schneider et al. 2010). However, the details of the involvement of RNase MRP in the general rRNA maturation are not currently known and additional cleavage sites on pre-rRNA or other RNA molecules involved in rRNA maturation are yet to be identified. Aside from its role in the maturation of rRNA, yeast RNase MRP was shown to be involved in the regulation of the cell cycle: mutations in RNase MRP resulted in missegregation of plasmids (Cai et al. 1999) and caused a cell cycle delay at the end of mitosis (Cai et al. 2002). The cell cycle delay appeared to be caused by the accumulation of cyclin B2, which resulted from an increase in the concentration of cyclin B2 mRNA (Cai et al. 2002). RNase MRP was shown to cleave the 50 -UTR of this mRNA, which allowed its 50 to 30 exonucleolytic degradation by ribonuclease Xrn1 (Gill et al. 2004); a fraction of RNase MRP was shown to transiently (after the initiation of mitosis and until the completion of telophase) accumulate in a discrete cytoplasmic spot, consistent with its role in the degradation of mRNA (Gill et al. 2006). Mutations of RNase MRP in D. melanogaster resulted in growth and developmental defects starting in the early larval period and resulting in larval death during the second instar stage (Schneider et al. 2010). The mechanism responsible for these defects in flies is not clear. RNase MRP RNA was recently demonstrated to form a complex with human telomerase reverse transcriptase (TERT) (Maida et al. 2009); however the meaning of this observation is yet to be established. Defects in the function of RNase MRP in humans result in a variety of recessive pleiotropic diseases, typically involving dwarfism. These diseases include Cartilage Hair Hypoplasia (CHH) – the first identified human disease caused by mutations in a noncoding RNA – (Ridanpaa et al. 2001) and a variety of dysplasias (Ridanpaa et al. 2003; Kuijpers et al. 2003; Thiel et al. 2005). These recessive diseases are usually caused by a combination of two alleles, each carrying mutation(s) in the gene encoding the RNA component of RNase MRP (RMRP), or, alternatively, insertions/deletions in the promoter region of the RMRP gene typically resulting in a reduced/abrogated level of transcription for one allele, combined with mutations in the RMRP gene itself in the other allele (Bonafe et al. 2005; Hermanns et al. 2005; Nakashima et al. 2007). The disease-causing mutations in the RNA component of RNase MRP are typically localized to phylogenetically conserved regions and may include a wide variety of insertions, deletions, substitutions, or duplications (Bonafe et al. 2005; Hirose et al. 2006; Hermanns et al. 2006; Martin and Li 2007 and references therein). These mutations can potentially affect RNA interactions with RNase MRP proteins (which, among other things, may result in reduced RNA stability),

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or change RNase MRP activity/specificity. The effects of mutations on the enzyme should depend on their nature and localization; consistent with this, their clinical manifestations appear to be defined by their nature and localization in the two affected alleles; a possible correlation between clinical manifestations and the general localization of RMRP mutations has been suggested (Thiel et al. 2007). The mechanisms responsible for the development of human pleiotropic diseases caused by defects in RNase MRP are not clear, but it seems to be likely that they may be related to the effects on cell cycle and development observed in yeast and flies (Gill et al. 2004; Schneider et al. 2010).

13.7

Summary

Ribonuclease P is a universal essential enzyme responsible for the maturation of the 50 -ends of tRNA in practically all organisms from all three domains of life; RNase P is likely to play important roles in the processing of additional RNA molecules. RNase MRP is closely related to RNase P, but it is a strictly eukaryotic enzyme involved in the maturation of rRNA as well as the degradation of certain mRNAs. Ribonucleases P and MRP are distinct from all other known ribonucleases as they contain RNA moieties that are responsible for catalysis. The RNA component of RNase P is generally well conserved in all three domains of life, while RNase MRP closely resembles RNase P, but possesses an RNA domain that is unique to this eukaryotic enzyme. In addition to their catalytic RNA, RNases P/MRP contain essential protein parts. The size and complexity of these protein parts grows dramatically in the more evolutionarily advanced organisms. The reasons for this increase in complexity are not clear. Available biochemical and structural information sheds light on the structure and function of bacterial RNase P, but the more complex archaeal and, especially, eukaryotic enzymes of the RNase P/MRP family are much less understood. Acknowledgments I apologize to the authors of works that were not cited in this brief review due to space limitations. I would like to thank Lydia Krasilnikova for her help with the manuscript preparation. This work was supported by NIH grant GM085149 to A.S.K.

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Chapter 14

Nonenzymatic and Metal-Ion-Dependent RNA Cleavage, and RNase Models Harri L€ onnberg

Contents 14.1 14.2 14.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cleavage of Phosphodiester Bonds via a Dianionic Phosphorane Intermediate . . . . . . Cleavage and Isomerization of Phosphodiester Bonds via a Monoanionic Phosphorane Intermediate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.4 Cleavage and Isomerization of Phosphodiester Bonds via a Neutral and Monocationic Phosphorane Intermediate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.5 Cleavage of RNA Phosphodiester Bonds by Multifunctional Organic Agents . . . . . . . . 14.6 Cleavage of RNA Phosphodiester Bonds by Metal Ion Complexes . . . . . . . . . . . . . . . . . . . 14.7 Solvent Effects on the Cleavage Rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.8 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

344 345 347 349 351 356 360 361 362

Abstract Systematic variation of individual amino acid residues within the catalytic core by means of protein engineering has turned out to be a powerful tool for the mechanistic studies of ribonucleases. The results of such studies are, however, open to alternative interpretations, since the replacement of even a single residue may affect chain folding. This, in turn, alters the geometry, non-covalent interactions and mutual orientation of the catalytically active residues to such an extent that identification of the real origin of the observed influence on rate remains uncertain. Unambiguous structure–reactivity correlations based on studies with structurally simplified chemical models may help to distinguish between alternative mechanisms. The present review is aimed at summarizing the results of such model studies. Accordingly, cleavage of RNA phosphodiester bonds by solvent-derived species, general acids and bases, metal ions, and multifunctional small molecular entities is surveyed.

H. L€onnberg Department of Chemistry, University of Turku, FIN-20014 Turku, Finland e-mail: [email protected] A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5_14, # Springer-Verlag Berlin Heidelberg 2011

343

344

14.1

H. L€ onnberg

Introduction

Although the sugar-phosphate backbone of RNA is, owing to the presence of the 20 -hydroxy group, hydrolytically labile compared to DNA, it still is a chemically stable molecule. The half-life for the cleavage of an individual phosphodiester bond is of the order of 100 years under physiological conditions (Emilsson et al. 2003). Br€ onsted or Lewis acids or bases at a high concentration, or elevated temperature, are required for reasonably fast cleavage. The high cleavage rates obtained by ribonucleases are based on synergistic exploitation of several individual rateenhancing factors, such as activation of the entering nucleophile, stabilization of the resulting intermediate (if any), orientation of the entering and departing nucleophile, and stabilization of the departing nucleophile. Although the catalytic mechanisms utilized by different ribonucleases considerably differ in details, the course of the reaction is always the same. The cleavage is initiated by a nucleophilic attack of the neighboring 20 -hydroxy group on the phosphorus atom which leads to departure of the 50 -linked nucleoside and concomitant formation of a 20 ,30 -cyclic phosphate (Scheme 14.1), as known since the early studies of Brown (Brown and Todd 1955). The reaction proceeds via a phosphorane intermediate (or transition state), obeying the rules of Westheimer (1968). According to these rules, the ligands occupy either an apical (a) or equatorial (e) position within a trigonal bipyramidal phosphorane, the P-O(a) bonds being longer and weaker than the P-O(e) bonds. Nucleophiles enter and depart the phosphorane only through apical positions. Electron deficient atoms prefer apical and electron-rich equatorial positions. When two of the ligands are members of a five-membered ring, one of them is apical and the other equatorial. Sufficiently stable phosphorane intermediate may pseudorotate: the apical ligands adopt equatorial positions and two of the equatorial ligands become apical. This leads to isomerization of the 30 ,50 -bond to a 20 ,50 -bond. The cyclic 20 ,30 -phosphate is hydrolytically less stable than 30 ,50 -phosphodiesters, being quite rapidly hydrolyzed to a mixture of 20 -and 30 -phosphates. Usually, the intermediary accumulation of the cyclic phosphate is barely noticeable.

2′ O O H O P O 3′

O 5′

eO P

Oa

O e P

aO

O- e a O HO e

e -O

O

O P

O

O-

e O

2′ 3′ OH O O P O-

a OH

O 5′

HO +

O O P O-

Scheme 14.1 Cleavage and isomerization of RNA phosphodiester bond

OH

OH

OH O +

O P OOH

14

Nonenzymatic and Metal-Ion-Dependent RNA Cleavage, and RNase Models

345

The cleavage of phosphodiester bonds is accompanied by deprotonation of the attacking 20 -OH and protonation of the departing 50 -O. Accordingly, agents that facilitate these proton transfers accelerate the cleavage. Likewise, interactions that stabilize the phosphorane intermediate by reducing the electron density at the non-bridging oxygen atoms are rate enhancing. Finally, constraints that influence the geometry of the phosphorane intermediate (or transition state) are of importance. The constraints which prevent the departing nucleoside to freely take an apical position within the phosphorane intermediate are rate retarding, while an initial state conformation which allows 20 -O to approach the phosphorus atom onnberg and L€onnberg 2005). colinearly to the P-O50 bond is rate enhancing (L€ As mentioned, ribonucleases obviously are able to exploit several of the rateenhancing factors in a cooperative manner. The catalytic repertoire of small molecule RNA-cleaving reagents is more limited; the catalysis is usually based on a synergistic influence of only a couple of the factors listed above. While this shortcoming reduces the efficiency of small molecules as cleaving agents, it simultaneously offers one clear advantage for mechanistic studies. With appropriately designed simple chemical models, the effect of only a single structural parameter on reactivity may be studied and this data may help to distinguish between alternative mechanistic interpretations of enzyme action. The results of such studies are surveyed in the present review.

14.2

Cleavage of Phosphodiester Bonds via a Dianionic Phosphorane Intermediate

The cleavage of RNA phosphodiester bonds is initiated by an attack of the 20 -O on the central atom of the neighboring phosphate group (Scheme 14.1). Deprotonation dramatically increases the nucleophilicity of the attacking oxygen atom. Studies on mutual isomerization of 50 -O-methyluridine 20 - and 30 -dimethylphosphate suggest that 20 -O- is up to 1010 times better nucleophile than 20 -OH (Kosonen et al. 1994). This reaction, which undoubtedly proceeds by an attack of the 20 -oxygen atom on the phosphorus atom, becomes hydroxide ion-catalyzed even at pH 3, although the mole fraction of the 20 -oxyanionic form at pH 3 is only around 1010 (the pKa value of the 20 -OH is around 13). Owing to the high nucleophilicity of 20 -O-, the cleavage of RNA phosphodiester bonds becomes hydroxide ion catalyzed even at pH 6 at 90 C (Fig. 14.1) (J€arvinen et al. 1991). The reaction proceeds by a rapid initial deprotonation of the 20 -OH, followed by an attack of the 20 -O- on the phosphorus atom and breakdown of the resulting dianionic phosphorane intermediate by departure of the 50 -linked nucleoside as an oxyanion (Scheme 14.2a). Studies with RP and SP phosphorothioates have indicated that the reaction proceeds with 100% inversion (Oivanen et al. 1995). The blg value of the cleavage reaction is much more negative with ribonucleoside 30 -alkylphosphates (blg ¼ 1.28; Kosonen et al. 1997) than with their aryl counterparts (blg ¼ 0.59;

346

H. L€ onnberg -1 -2

log (k /s-1)

-3 -4 -5 -6 -7 -8 -2

0 H0

2

4

6

8

10

12

pH

Fig. 14.1 pH-rate profiles for the cleavage of 30 ,50 -UpU to uridine and its 20 ,30 -cyclic phosphate (cleavage, filled circles), and mutual isomerization of 20 ,50 - and 30 ,50 -UpU (open circles). Ho is an acidity function (an extension of pH-scale to concentrated acids) (J€arvinen et al. 1991)

a 3′ 2′ O OH O P OO

–H+

R

-

e O O a P O e RO Oe a

O OO P OO R

O

O P O

+H+ HO

O + O-

b 3′ 2′ O O H O P OO 5′

‡ :B

eO O

Oa

P O O- e e

-

O

O O P O O

+H+

HO

+

Scheme 14.2 Specific base-catalyzed cleavage of ribonucleoside 30 -phosphodiesters (a) and general base-catalyzed cleavage by buffer bases having pKa > 8 (b)

Davis et al. 1988), suggesting that the reaction is stepwise and the dianionic phosphorane has a finite lifetime, although it undoubtedly is only marginally stable (L€ onnberg et al. 2004). The transition state is late, exhibiting a marked buildup of negative charge on the departing 50 -oxygen. This conclusion is corroborated by the values of primary 18O isotope effects and DFT calculations. Studies with 30 ,50 -UpG have shown that the kinetic 18O isotope effect for the attacking 20 -O and departing

14


347

50 -O (0.984 0.004 and 1.034 0.004, respectively) are both large compared to cleavage of phosphodiesters with aryl leaving groups, consistent with a transition state where both the P-O20 bond formation and P-O50 bond fission are far advanced (Harris et al. 2010). An almost equal value (1.0272 0.0001) has been reported for the departure of the 3-nitrobenzyloxy ion from uridine 30 -(3-nitrobenzyl)phosphate (Gerratana et al. 2000). The bnucl value of 0.75 is, in turn, consistent with far advanced P-O20 bond formation in the transition state (Ye et al. 2007). The fact that the attacking 20 -OH is really deprotonated in a pre-equilibrium step, not concerted with changes in heavy atom bonding, is evidenced by the observation that at pH < pKa of the 20 -OH, the kinetic solvent deuterium isotope effect equals the equilibrium isotope effects of alcohol deprotonation (Virtanen et al. 2005), while at pH > pKa, the effect is close to unity (Usher et al. 1970). In sufficiently basic buffers (pKa > 8), the deprotonating agent may be the buffer base instead of hydroxide ion. In other words, the cleavage is subject to general base catalysis (Breslow et al. 1996; Beckmann et al. 1998; Kirby and Marriott 2002). The attacking 20 -OH is deprotonated by the buffer base concerted with the formation and breakdown of the dianionic phosphorane (Scheme 14.2b). DFT calculations lend some support to the mechanism of the specific base catalysis discussed above. Calculations with a continuum solvation model show that the attack of methoxide ion on ethylene phosphate monoanion proceeds by intermediary formation of a marginally stable phosphorane, the formation of which is rate limiting (Liu et al. 2006). The implication to RNA hydrolysis is that a marginally stable dianionic phosphorane is formed in a pre-equilibrium step and the rupture of the P-O50 bond is rate-limiting. Calculations on the cleavage of tetrahydrofuran-3,4-diol cyclic phosphate have led to similar conclusions (Lopez et al. 2006). It is worth noting that hydroxide ion catalyzed isomerization has never been observed. The dianionic phosphorane cannot pseudorotate since the two oxyanions are locked into equatorial positions, and evidently the intermediate is too unstable to pseudorotate via a temporary kinetically invisible protonation, in spite of the fact that protonation to a monoanionic phosphorane is thermodynamically allowed. Estimations for the second pKa value of phosphoranes range from 13.5 to 14.3 (Davies et al. 2002; Lopez et al. 2002).

14.3

Cleavage and Isomerization of Phosphodiester Bonds via a Monoanionic Phosphorane Intermediate

Buffer bases having a pKa value around 7 are too weakly basic to compete with the oxyanions of the developing phosphorane for the 20 -OH proton, but the proton is transferred from the attacking 20 -OH to a phosphorane oxyanion concerted with the P-O20 bond formation (Scheme 14.3). The reaction, hence, proceeds by formation of a monoanionic phosphorane intermediate (Kosonen et al. 1998). This species, in

348

H. L€ onnberg 3′ 2′

O

O H

O P OO

H O H 5′

3′ 2′ eO -

e

O

P

O a/ e

O a

a O -

OH a/e

e

O

Oe

P O a/e a

H O

OH a/e H

O H

O O- P O O

5′

‡ O a/e P – e O O e H O a H O

HO

a/e O

O O

P

O

+

O–

H

Scheme 14.3 pH- and buffer-independent cleavage and isomerization of RNA phosphodiester bonds (Kosonen et al. 1998)

contrast to its dianionic counterpart, is sufficiently stable to pseudorotate, which allows isomerization to compete with the cleavage. In fact, isomerization is even faster than cleavage, indicating the breakdown of the monoanionic phosphorane to be the rate-limiting step of the cleavage reaction. Consistent with this observation, DFT calculations based on polarizable continuum model indicate that the barrier for the exocyclic PO bond fission (14.1 kcal mol1) is considerably higher than that for the endocyclic fission (3.2 kcal mol1) (Lopez et al. 2004). The latter barrier is comparable to the barrier of the pseudorotation of the monoanionic phosphorane and, hence, the pseudorotation may partly limit the rate of isomerization in the pH range 4–6. The reaction is rather insensitive to the polar nature of the esterified alcohol, the blg value being 0.23 0.11 (Kosonen et al. 1998). The susceptibility of the cleavage reaction to the basicity of the leaving group is much lower than in the cleavage via a dianionic phosphorane, blg ¼ 0.59 0.12. Evidently, a proton is transferred from the phosphorane hydroxyl ligand to the departing oxygen concerted with the P-O bond fission, which partly neutralizes the negative charge accumulated on the leaving group. With triester-derived monoanionic phosphoranes, where such proton transfer is not possible, isomerization is up to 105 times faster than cleavage (Kosonen et al. 1999), whereas with the corresponding diester-derived species the isomerization is only one order of magnitude faster. DFT calculations also highlight the importance of proton transfer between the attacking/ departing nucleophile and a non-bridging phosphoryl oxygen atom upon formation/ breakdown of the monoanionic phosphorane (Lopez et al. 2006), but the detailed mechanism still is subject to alternative interpretations. A direct (Yang and Cui 2009) and water-mediated proton transfer (Boero et al. 2002) have both been supported by calculations. The observation that no isomerization takes place in anhydrous tert-butyl alcohol, however, lends support for the essential role of water (Gerratana et al. 2000). The primary 18O isotope effect of the pH-independent cleavage of 30 -(3-nitrobenzyl) phosphate is 1.009 0.001, consistent with a considerably advanced PO bond cleavage in the transition state (Gerratana et al. 2000).

14


a

O –H+ 3′

O

2′

O

OH

O P OO

-

O -

O

P O

P O

349

HO

O O-

O O

P

O

+

O–

O OH

BH+

5′ HO O

b

O

P

O

+

OH –H+

O O

P

O O-

Scheme 14.4 Mechanistic alternatives proposed for the buffer-catalyzed cleavage of RNA phosphodiester bonds around pH 7 (a) (Breslow et al. 1996), (b) (Beckmann et al. 1998)

The cleavage via a monoanionic phosphorane is also subject to catalysis by buffers having a pKa value around 7. Two rather different mechanisms have been proposed. According to Breslow et al. (1996), the monoanionic phosphorane obtained in a pH-independent pre-equilibrium step first undergoes specific base catalyzed deprotonation and the resulting dianionic phosphorane then undergoes general acid catalyzed breakdown to 20 ,30 -cyclic phosphate and alcohol (Scheme 14.4a). In other words, the overall cleavage reaction is argued to be catalyzed by general bases. The group of Kirby (Beckmann et al. 1998) has, in turn, suggested that the breakdown of the monoanionic phosphorane is simply general acid catalyzed, no preceding deprotonation step is required (Scheme 14.4b). The buffer catalysis observed for the isomerization is at most very weak, consistent with the view that pseudorotation is at least partially rate limiting (M€aki et al. 1999).

14.4

Cleavage and Isomerization of Phosphodiester Bonds via a Neutral and Monocationic Phosphorane Intermediate

Besides deprotonation of the 20 -OH, protonation of the phosphodiester linkage markedly accelerates the formation of the phosphorane intermediate. For this reason, the cleavage and isomerization become hydronium ion catalyzed at pH < 5. Rapid pre-equilibrium protonation of the phosphodiester monoanion is followed by an attack of the 20 -oxygen on the phosphorus atom, accompanied by a proton transfer from the 20 -OH group to the developing phosphorane oxyanion ligand (Scheme 14.5) (J€arvinen et al. 1991). According to quantum chemical

350

H. L€ onnberg ‡ a /e O O a/e P HO Oe H e O a H O H

HO O O + P O OH -

H+

O O P O O 3′ 2′

3′ 2′ O OH O P O O 5′

+H+

O O H HO P O H O H O 5′

+H+

eO

Oa P OH a/e HO a/e O a/e

a O Oe P OH a/e HO O a/e a/e

2′, 5′-diester

+H+

-

H+

3′ 2′ O OH O OH +H+ HO P OH+ HO P O O O 5′

eO

OH+ a P HO OH a/e a/e O a /e

a OH+ O e P HO OH a/e O a/e a/e

O a/e P OH a/e HO a/e OH+ a

2′, 5′-diester

a /e O

HO O O + P + HO OH -

H+

O O P O OH

Scheme 14.5 Mechanisms for the hydronium ion catalyzed cleavage and isomerization of RNA phosphodiester bonds via a neutral and monocationic phosphorane (Mikkola et al. 2002)

calculations, the proton transfer is mediated by a water molecule (Uchimaru et al. 1996). The neutral phosphorane obtained is decomposed approximately as readily to a 20 ,5-diester as to a 20 ,30 -cyclic phosphate and the 50 -linked nucleoside (J€arvinen et al. 1991). This reaction mechanism predominates from pH 4 to 2 (at 90 C). Since the leaving group departs as an alcohol, not an alkoxide ion, the susceptibility to the polar nature of the leaving group is low. The blg values for the cleavage and isomerization are: 0.19 0.12 and –0.23 0.11, respectively (Kosonen et al. 1998). The suggested mechanism is consistent with the kinetic 18O isotope effects observed for the cleavage of uridine 30 -(3-nitrobenzyl)phosphate at pH 2.5, viz. 1.0019 and 0.9904 for 18O in the bridging and two non-bridging positions, respectively (Gerratana et al. 2000). At pH < 2, cleavage and isomerization via a monocationic phosphorane becomes predominant. The disappearance of the starting material is second order

14


351

in hydronium ion concentration, insofar as the phosphodiester monoanion is still the predominant ionic form (pH > 1) and exhibits first-order dependence in more acidic solutions (J€arvinen et al. 1991). Most likely, both of the non-bridging phosphoryl oxygens are protonated in a rapid pre-equilibrium step and the 20 -hydroxy group then attacks on the phosphate giving a cationic phosphorane intermediate, which after protolytic rearrangement undergoes breakdown to either cleavage or isomerization products. The cleavage and isomerization rates are again comparable, the relative rate constants for the cleavage, isomerization of a 30 ,50 bond to a 20 ,50 -bond, and its reverse reaction being 1.2, 1.0 and 0.9, respectively. The blg value is low, viz. 0.04 0.04 (Kosonen et al. 1998). The rate-accelerating effect of the protonation of both of the non-bridging phosphoryl oxygens, which might be called double Br€onsted acid activation, is remarkable. The reaction via this monocationic species starts to predominate already at pH 3, although the pKa value most likely is > > > > > > < 2S2G2H ! 3S1G1H þ GSH > = 3S1G1H þ GSH ! 2S2G2H “2S2G2H” > > > > > > 3S1G1H þ GSH ! 3S2H þ GSSG > > : ; 3S2H þ GSSG ! 3S1G1H þ GSH “3S2H” “3S1G1H”

3S2H ! 3S2H 9 8 < 3S1G1H ! 3S1G1H = 3S1G1H ! 4S þ GSH ; : 4S þ GSH ! 3S1G1H

a Each pathway is expressed in terms of the most reduced “Intermediate” involved in the ratelimiting steps. b Possible rate-limiting reactions in each pathway.

Fig. 15.5 Calculated fraction (%) of fully regenerated RNase A produced (through the six pathways of Table 15.1) by reaction of reduced RNase A (7.3 105 M) with initial concentrations of GSH of 4.2 105 – 0.11 M and a fixed concentration of GSSG (2.3 mM) at pH 8.2 and 22 C for 60 min (Konishi et al. 1982a)

380

15.9

H.A. Scheraga

Oxidative Folding of RNase A with a Dithiothreitol Couple

While the number of regeneration pathways with GSSG/GSH has been assigned on the basis of a kinetic analysis (Table 15.1), the determination of the specific disulfide-bonded intermediates that are involved in these pathways has been hampered by the complexity of the problem. Linear disulfide reagents, such as GSSG, can form stable mixed disulfides with protein thiols, as shown in Eq. 15.1. This increases the number of possible disulfide-bonded species of RNase A from 763 to 7,192. In addition, the stability of various mixed disulfides with GSSG strongly determines which pathway is taken. Because mixed disulfides with oxidized dithiothreitol (DTTox) are highly unstable (Creighton 1986), these problems do not exist when DTTox is used as the oxidizing agent. Previously published attempts to use DTTox in studies of the regeneration pathways of RNase A have been unsuccessful because of an apparent inability of DTTox to regenerate RNase A (Creighton 1977, 1979, 1986; Creighton et al. 1980; Wearne and Creighton 1988). Since reduced dithiothreitiol (DTTred) is a potent reducing agent (Cleland 1964), the amount of DTTred produced by the formation of protein disulfides inhibits the rate of regeneration when DTTox is used as the oxidizing agent. Since the thiolate anion is the relevant reduced species responsible for the reducing power of DTTred, a decrease of pH will decrease the concentration of DTTred and increase the oxidative strength of the redox pair significantly. While previous studies (Creighton 1977, 1979, 1986; Creighton et al. 1980; Wearne and Creighton 1988) were conducted at pH 8.7, the majority of our experiments was carried out at pH 8.0 (Rothwarf and Scheraga 1991), the pH close to that of the glutathione studies cited above. Typical results, shown in Fig. 15.6 (Rothwarf and Scheraga 1991), demonstrate the oxidizing ability of DTTox in agreement with our determination (Rothwarf and Scheraga 1992) of the equilibrium and kinetic constants for the thiol-disulfide exchange reaction between GSSG and DTTred. An additional factor in the failure of previous workers to detect native RNase A is the lag time between the start of the oxidation process and the appearance of the native protein. This lag varies from 15 to 90 min and depends on the pH, temperature, and starting concentrations of DTTox, DTTred, and protein (Rothwarf and Scheraga 1991). In the previous regeneration studies of Creighton and coworkers, data were taken only during the first 30 min of the oxidative process. We have observed (Rothwarf and Scheraga 1991) that, even under the less favorable conditions used in those earlier studies of Creighton et al., an apparent half-life following the lag period (t½) of 34 h can be observed. The shortest t½ that we have been able to observe is 128 min. This compares favorably with the t½ of 108 min obtained at the optimal ratio of oxidized and reduced glutathione under similar solution conditions (Konishi et al. 1982a). Because DTTox regeneration is not limited by mixed disulfide formation, higher ratios of DTTox/DTTred will result in regeneration rates greater than those obtainable with glutathione, as we have found (Rothwarf and Scheraga 1991).

15

Ribonucleases as Models for Understanding Protein Folding

381 0.4

0.04

0.2 R

0.02

[NaCI] (M)

Absorbance

I II

N III

0.00

0

20

40 Time (min)

60

0.0

Fig. 15.6 The solid curve represents the chromatographic separation of disulfide intermediates in the regeneration of reduced RNase A on a Bakerbond CBX column at room temperature in 25 mM Hepes, 1 mM EDTA, pH 7.0. Fully reduced RNase A (30 mM) was regenerated for 90 min at 25 C in 100 mM Tris, 2 mM EDTA, pH 8.0. The starting concentration of DTTox was 100 mM. The regeneration process was quenched by addition of an excess of the blocking reagent AEMTS and desalted. R is the fully reduced protein, N is the fully regenerated native protein. The Roman numerals indicate the number of intramolecular disulfide bonds of the protein in the labeled peak. The peaks at the beginning of the chromatogram pertain to buffer salts. The salt gradient used to elute the protein is represented by the dashed line (Rothwarf and Scheraga 1991)

The pathway for reduction of RNase A by DTTred has been shown to have a native-like three-disulfide intermediate lacking the 65–72 disulfide. This work has been reported by Creighton and coworkers to be nonreproducible. We have, however, been able to isolate this species in good yield, greater than 8% of the total protein (Rothwarf and Scheraga 1991). The primary difference between our result and previous results of Creighton and coworkers is the method used to block the free thiols in the protein. The blocking reagent that we used, 2-aminoethyl methanethiosulfonate (AEMTS) (Bruice and Kenyon 1982), is at least five orders of magnitude more reactive with thiols in solution (Rothwarf and Scheraga 1991) than the reagent (iodoacetic acid) used in previous studies. It appears that the accessibility of the Cys-65 and Cys-72 thiols is somewhat restricted. We have found that they are not blocked by iodoacetic acid under the conditions normally employed by Creighton and coworkers. Following these observations, an extensive series of oxidation/reduction studies of RNase A was carried out with DTTox/DTTred (Milburn and Scheraga 1988; Altmann and Scheraga 1990; Rothwarf and Scheraga 1993a, b, c, d; Talluri et al. 1993; Xu et al. 1996; Shimotakahara et al. 1997; Laity et al. 1997; Rothwarf et al. 1998a, b; Iwaoka et al. 1998; Xu and Scheraga 1998; Welker et al. 1999; Volles et al. 1999; Carty et al. 2002). The regeneration of bovine pancreatic ribonuclease

382

H.A. Scheraga

A (RNase A) from the reduced to the native form with mixtures of DTTox and DTTred was studied at 25 C, pH 8.0, by using a variety of experimental techniques, including quenching the regeneration reaction with AEMTS, fractionation of intermediates by HPLC, and analysis by both UV and disulfide-specific detection systems. The disulfide-containing protein intermediates achieve a steady-state distribution after which the native protein regenerates at a rate comparable to the rates observed previously during the regeneration of RNase A with glutathione. Equilibrium constants at 25 C, pH 8.0, for the interconversion of species containing different numbers of disulfide bonds were evaluated from the concentrations in the steady-state distribution. These equilibrium constants were compared to those obtained when native RNase A was regenerated with glutathione. The observed equilibrium constants (with the dithiothreitol system) for the interconversions among all intermediates are very similar once statistical factors arising from the different numbers of disulfide-containing species in each grouping are taken into account. None of the disulfide-containing intermediates has any significant enzymatic activity, in agreement with earlier conclusions that these intermediates are considerably disordered. This is in sharp contrast to disulfide-containing intermediates populated during the regeneration of bovine pancreatic trypsin inhibitor, which have significant native-like structure. Analysis of the experimental data for RNase A revealed that this protein regenerates with DTToxand DTTred through a rearrangement pathway involving one or more three-disulfide species; that is, multiple pathways could be involved (Rothwarf and Scheraga 1993b). These pathways are different from those observed when RNase A is regenerated with oxidized and reduced glutathione (GSSG and GSH, respectively). These differences result primarily from the very different characteristics of the oxidation of thiols with DTTox and GSSG, respectively. In addition, the concept of multiple pathways, as applied to the regeneration of RNase A, was developed. Monothiol reagents such as GSSG and GSH form stable mixed disulfides with protein thiols while dithiol reagents such as DTTox and DTTred do not. This large difference in the stabilities of the mixed disulfides is reflected in much greater rates of formation and reduction of protein disulfide bonds with GSSG/GSH than with DTTox/DTTred. With dithiothreitol, the concentrations of intermediate species in the steady state depend on the redox potential, that is, on the [DTTox]/[DTTred] ratio, and not on the absolute concentrations of these reagents. With glutathione, the redox potential, and hence the concentrations of intermediate species in the steady state, depend on the [GSSG]/[GSH]2 ratio; hence, with glutathione, in contrast to dithiothreitol, the absolute values of the concentrations do affect the steady-state concentrations. Consequently, the regeneration pathways of bovine pancreatic ribonuclease A depend on the nature of the redox reagent as well as the redox potential at which they are used. The use of GSSG/GSH favors multiple regeneration pathways, while the use of DTTox/DTTred favors regeneration through fewer pathways (Rothwarf and Scheraga 1993c). The rate of regeneration of RNase A with DTTox and DTTred decreases by a factor of 10 when the temperature is increased from 25 C to 37 C (Rothwarf and Scheraga 1993d). The rate of

15


383

regeneration of RNase A with oxidized and reduced glutathione increases slightly over this same range of temperature. This suggests that the regeneration processes with the two types of redox reagents proceed through different pathways. There is a significant change in the distribution of three-disulfide intermediates populated during regeneration with DTTox/DTTred over the range of temperature 15–37 C that suggests that the three-disulfide species populated at 15 C are directly involved in the major regeneration pathway observed at 25 C. During the regeneration of RNase A from the reduced to the native form with mixtures of oxidized and reduced dithiothreitol at 25 C, pH 8.0, the disulfidecontaining protein intermediates achieve a steady-state distribution. By manipulating the redox conditions after the attainment of the steady-state condition, it has been possible to kinetically trap and, thereby, isolate and identify the disulfide-bonded species that follow the rate-determining step in the regeneration pathway. Two three-disulfide species have been identified by peptide mapping (Rothwarf et al. 1998a). Both species contain three native disulfide-bond pairings; one lacks the 65–72 disulfide bond (des-[65–72]), and the other lacks the 40–95 disulfide bond (des-[40–95]). These species are the same as those identified during the reduction of RNase A (Li et al. 1995). By restarting the regeneration process from isolated des-[65–72] and des-[40–95], it was shown that both intermediates lie directly on regeneration pathways. The populations of these two three-disulfide species during the regeneration process were monitored directly through the use of a reduction pulse in which the disulfide bonds of nonnative structures are reduced (Rothwarf et al. 1998a; Li et al. 1995). A detailed kinetic analysis of the regeneration process was carried out to obtain rate constants for disulfide interconversion among the various disulfide-bonded intermediates (Rothwarf et al. 1998b). This analysis indicates that these two pathways account for almost 100% of the native protein regenerated and that the amount regenerated by S-S/SH interchange in a ratio of about 4:1 for des-[40–95], relative to des-[65–72], is insensitive to the redox conditions used. These results indicate that the rate-determining step in both pathways involves formation of the native-like three-disulfide species by S-S/SH reshuffling from the 3S ensemble, a step in which most of the conformational folding takes place. The experimentally determined rate constants indicate that these two pathways are sufficient to explain the differences in the temperature dependence of the regeneration rate with different redox reagents. These two resulting major pathways are shown in Fig. 15.7.

RU

1s U

2s U

3s U

4s U

des [65-72] des [40-95]

Fig. 15.7 Major oxidative folding pathways of RNase A (Rothwarf et al. 1998a)

N

384

H.A. Scheraga

NMR studies carried out on analogs of des-[65–72] (Talluri et al. 1994; Shimotakahara et al. 1997), and des-[40–95] (Laity et al. 1997), respectively, indicate that these analogs have a close structural similarity to RNase A in all regions with the only major differences occurring in regions consisting of residues 60–72 and 40–95. This led to the conclusion that, in reduction pathways (Li et al. 1995) that include these analogs (at 25 C, pH 8.0), the rate-determining step corresponds to a partial unfolding event in two separate regions of the protein and not to a global conformational unfolding process (Laity et al. 1997). The results further suggest that, in the regeneration pathways involving these analogs, the regions 60–72 and 40–95 are the last to fold to the native structures. From analogous kinetic studies of the oxidation of analogs of des-[65–72] (Iwaoka et al. 1998) and des-[40–95] (Xu and Scheraga 1998), respectively, with dithiothreitol, it was possible to detect two minor pathways in addition to the major ones that arise from reshuffling of the 3S ensemble to des-[65–72] and des-[40–95], respectively. These minor pathways are illustrated in Fig. 15.8. Both analogs folded through the same pathways, observed for the wild-type protein, but with a different rate-determining step, viz., the oxidation of the two-disulfide ensemble (2S) to the post-transition state des-[65–72] and des[40–95], respectively, which then proceed to the native structure. These two minor pathways constitute only about 5% of the two major pathways, and could be detected only by oxidative kinetic studies of the analogs rather than of the wild-type protein. A two-disulfide mutant, in which the 65–72 and 40–95 disulfide bonds were deleted, was subjected to oxidative folding from the reduced form with DTTox (Lester et al. 1997). All three of the possible two-disulfide pairings, including the native one, formed. One-dimensional 1H NMR spectra demonstrated that the conformations of these three species are similar and are predominately disordered; however, there is evidence of local structure in the vicinity of one histidine residue. It was also shown that disulfide pairing is not completely random and that both entropic factors and enthalpic interactions contribute to the formation of the nativedisulfide bonds. The presence of more than a statistical population of nativedisulfide pairings indicates that specific local interactions present in the reduced protein direct the preferential formation of native-disulfide bonds in the twodisulfide mutant. By carrying out the oxidative folding of reduced wild-type RNase A at 15 C instead of 25 C, two new three-disulfide intermediates were detected (Welker

RU

Fig. 15.8 Minor oxidative folding pathways of RNase A (Iwaoka et al. 1998; Xu and Scheraga 1998)

1SU

2SU

des [65–72]

3SU

des [40–95]

N

4SU

15


385

Fig. 15.9 Schematic representation of the structure of RNase A, showing that disulfide bonds 40–95 and 65–72 are relatively exposed whereas 26–84 and 58–110 are buried (Li et al. 1995)

et al. 1999). These intermediates, des-[26–84] and des-[58–110], possess all but one of the four native disulfide bonds and have stable tertiary structure, similar to the two previously observed intermediates, des-[65–72] and des-[40–95]. While the latter two des species each lack one surface-exposed disulfide bond, the newly discovered intermediates each lack one buried disulfide bond. These two new species are possibly involved in the rate-determining steps during oxidative folding of RNase A. Figure 15.9 shows the relative exposure of the 65–72 and 40–95 disulfide bonds, and the burial of the 26–84 and 58–110 disulfide bonds. The early stages of oxidative folding of RNase A were probed by determining the populations of the possible one-disulfide species in the one-disulfide ensemble (Xu et al. 1996). A total of 24 out of 28 possible one-disulfide intermediates were found to be populated in the one-disulfide mixture. The population of one-disulfide intermediates displays a nonrandom distribution. All four native disulfide pairings have populations greater than those predicted by loop entropy calculations, suggesting the presence of enthalpic contributions stabilizing these species. The one-disulfide intermediate [65–72], containing the disulfide bond between cysteines 65 and 72, comprises 40% of the entire one-disulfide population. The interactions that stabilize this intermediate may play an important role in the regeneration pathways of RNase A. A similar investigation was carried out to characterize the distribution of onedisulfide bonds in the two-disulfide ensemble in the oxidative folding of RNase A (Volles et al. 1999). Again, the relative concentration of each of the 28 possible one-disulfide bonds in the two-disulfide ensemble was determined. Comparison with a statistical mechanical treatment of loop formation showed that the twodisulfide intermediates are probably compact. All 28 disulfide bonds were observed, demonstrating the absence of specific long-range interactions in these intermediates. Thermodynamic arguments suggest that the absence of such specific long-range interactions in the two-disulfide species may elevate the concentration of kinetically important three-disulfide intermediates and thereby increase the folding rate. Disulfide bond [65–72] was found to make up ~27% of the disulfide bonds of the two-disulfide species, significantly more than all other disulfides, because of stabilization by loop entropy factors and an energetically favorable b-turn. This turn may be one of several chain-folding initiation sites (Montelione

386

H.A. Scheraga

and Scheraga 1989), accelerating folding by decreasing the dimensionality of the conformational space that has to be searched. The interactions that favor the early formation of the 65–72 disulfide bond in wild-type RNase A were also found to favor preferential formation of this loop in a Ser-50-Met-79 fragment of RNase A (Milburn and Scheraga 1988). Included in this sequence are residues cysteine-65 and cysteine-72, which form a disulfide bond in the native enzyme, as well as cysteine-58. This molecule may form one of three possible intramolecular disulfide bonds (58–72, 58–65, or 65–72) upon thiol oxidation, viz., 1 loop of 15 and 2 loops of 8 residues each. These isomeric peptides were prepared by oxidation of residual thiols, and fractionation by reverse-phase HPLC. The 26-atom ring incorporating the native disulfide bond between residues 65 and 72 is the dominant product, in a ratio of 4:1 compared to the nonnative 58–65 disulfide bond (as in oxidation of the full protein), after protracted incubation under oxidizing conditions at 25.0 C and pH 8.0. Assuming that equilibrium was established, we inferred that local interactions in the sequence of RNase A significantly stabilize the native 8-residue disulfide loop with respect to the nonnative 8-residue loop (DG ¼ 1.1 0.1 kcal mole1). Two mutations were subsequently introduced separately in this fragment (Lys-66 ! Gln and Asn67 ! Ala) to try to disfavor formation of the 65–72 disulfide bond (Altmann and Scheraga 1990). However, as with the wild-type fragment, for which the equilibrium constant for the two possible isomers containing intramolecular eight-residue cyclic disulfides was 3.58 0.10 in favor of the disulfide bond between Cys-65 and Cys-72 (the disulfide bond present in the native protein), the equilibrium constants for the mutants were similar (3.02 0.17 for the Gln – 66 and 4.49 0.23 for the Ala-67 homolog), demonstrating alternative pathways by which local interactions can stabilize the loop formed by the disulfide bond between Cys-65 and Cys-72. Since both the correctly paired 65–72 loop and the incorrectly paired 58–65 loop contain the same number of amino acid residues, and hence involve the same entropy change in loop formation, the question arose as to whether the native pairing results from interactions within the 58–72 segment that lead to a nativelike structure even in its fully reduced form. Conformational energy calculations (Carty et al. 2002) showed that the native 65–72 loop was favored over the nonnative 58–65 loop. This result suggests that the native pairing of the 65–72 loop arises from energetic determinants (adoption of left-handed single-residue conformations by Gly-68, and side-chain interactions involving Gln-69) contained within this peptide sequence. NMR measurements (Talluri et al. 1993) on a 61–74 fragment of RNase A showed that short-range interactions help to stabilize the 65–72 disulfide bond with the formation of a type-II b-turn involving residues 66–69; this turn is shifted by one residue from the native type-III turn at residues 65–68. Figure 15.10 provides a summary of the essential features of the oxidative folding of RNase A with DTT. In the 28 possible species of the 1S ensemble, the dominant disulfide bond is the native 65–72, which persists in the 2S ensemble, ultimately leading to the native-like dominant des-[40–95] species.

15


387

Fig. 15.10 Summary of oxidative folding pathways of RNase A

15.10

Reductive Unfolding of RNase A

The reductive unfolding of RNase A with dithiothreitol proceeds through parallel pathways with the formation of two well-populated partially unfolded threedisulfide intermediates. This follows from the same type of experiments that were carried out for oxidative folding. Chromatograms obtained at various times of reduction show the accumulation of intermediates des-[65–72] and des-[40–95] at pH 8.0, 15 C. As shown in Fig. 15.9, disulfide bonds 65–72 and 40–95 are very exposed on the surface of the protein whereas the remaining two disulfide bonds of the native protein are quite buried. This exposure of the 65–72 and 40–95 disulfide bonds undoubtedly accounts for the initial preferential reduction of these two disulfide bonds. The mechanism of reductive unfolding is shown in Fig. 15.11. Two distinct local unfolding events, in which des-[65–72] and des-[40–95] are formed, rather than a global one, are involved in the rate-determining steps. These results are contrary to the current view that protein unfolding generally follows an all-or-none mechanism, and that the rate-determining step is controlled by an extensive rearrangement of the native structure. Sequential or simultaneous breakage of disulfide bonds through local unfolding events is energetically more favorable than disruption of the native structure through global unfolding. The results also indicate that, as shown in Sect. 15.9 and in Fig. 15.10, the oxidative refolding of ribonuclease A from the fully reduced form proceeds through parallel conformationally distinct transition states.

388

H.A. Scheraga

Fig. 15.11 Parallel reductive unfolding pathways of RNase A at 15 C and 25 C, pH 8.0 (Li et al. 1995)

Des-[65–72] k1(DTT red) N k2(DTT red)

k3(DTT red)

k5 3S’

very fast

k6

2S

very fast

R

k4(DTT red)

Des-[40–95]

15.11

Ribonuclease B (RNase B)

Bovine pancreatic ribonuclease B (Williams et al. 1987) constitutes ~15% of the total pancreatic ribonuclease (Grafl et al. 1987) and occurs naturally as a mixture of five glycoforms (Fu et al. 1994; Rudd et al. 1994). Its amino acid sequence is identical to that of RNase A, and differs from the A variant by the presence of a single carbohydrate moiety attached to Asn 34 (Williams et al. 1987). A comparison of the crystal structures of the two variants (Wlodawer and Sj€olin 1983; Williams et al. 1987) reveals no significant differences, suggesting that the oligosaccharide moiety has no influence on the static conformation of the protein. A novel method for characterization of the simultaneous reductive unfolding pathways of the five isoforms of RNase B led to results (Xu et al. 2004a) that indicate that each isoform unfolds reductively through two three-disulfide-containing structured intermediates before proceeding to the fully reduced form, as in the reductive unfolding pathways of the A variant lacking the carbohydrate chain. The rates of reduction of RNase A and RNase B and the formation and consumption of their reductive intermediates are identical, indicating that the unfolding events necessary to expose disulfide bonds for reduction are not affected by the oligosaccharide. The method utilizes top-down mass spectrometry and a naturally occurring tag on the protein, viz., the carbohydrate moiety, to obtain unfolding information of an ensemble of protein isoforms and is a generally applicable method for conducting folding studies on mixtures of different proteins (Xu et al. 2005).

15.12

Onconase

Besides RNase B, two other homologs [frog onconase (ONC) and bovine angiogenin (ANG)] of RNase A have been investigated in our laboratory. The motivation for considering these two homologs was primarily that both are missing an analog of the 65–72 disulfide bond, which is the location where folding is initiated in RNase A. So, clearly although they are homologs of RNase A, with similar three-dimensional structure (Pradeep et al. 2006), they must fold/unfold by pathways that differ from those of RNase A. ONC (Mosimann et al. 1994) has four disulfide bonds (C19–C68, C30–C75, C48–C90, C87–C104), three of which are approximately homologous to three of

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RNase A, but its C87–C104 disulfide bond replaces the C65–C72 disulfide bond of RNase A. ANG (Acharya et al. 1995) has only three disulfide bonds (C27–C82, C40–C93, and C58–C108), likewise missing the 65–72 disulfide bond of RNase A. Our studies of ONC and ANG are discussed in Sects. 15.12 and 15.13, respectively. The same methodology that was used for oxidative folding and reductive unfolding of RNase A was used for similar studies of ONC and ANG. In the reductive unfolding of ONC with DTTred, a fast-forming intermediate, des-[30–75], analogous to the des-[40–95] intermediate found in the reductive unfolding of its structural homolog, RNase A, was isolated and characterized (Xu et al. 2004b). The midpoints of the thermal transition and chemical denaturing curves (representing global unfolding) indicate that the conformation of des[30–75] is considerably less stable than that of the parent molecule, suggesting that the (30–75) disulfide bond plays a significant role in the conformational stability of ONC. While des-[30–75] is formed very quickly by a partial reduction of the parent molecule in a local unfolding step, it is not as easily susceptible to further reduction, indicating that its three disulfides are much more buried compared to the (30–75) disulfide bond in the parent protein. The nature of des-[30–75] is similar to that of des-[40–95] RNase A, in that des-[30–75] ONC is also a disulfide-secure species. In addition, based on the resistance to mild reducing conditions, structured des species appear to form in ONC from unstructured three-disulfide-containing ensembles. This step is key in the oxidative folding of RNase A, and is much faster in ONC than the formation of the structured des [40–95] species in RNase A. The reductive unfolding pattern of onconase is consistent with the analysis of the exposed surface area of the cysteine sulfur atoms in the (30–75) disulfide bond, which reveals that these atoms are about four- and sevenfold, respectively, more exposed than those in the two maximally exposed disulfides. By contrast, in the reductive unfolding of the homolog, RNase A, there are two intermediates, arising from the reduction of the (40–95) and (65–72) disulfide bonds, which takes place in parallel, and on a much longer time scale, compared to the initial reduction of onconase; this behavior is consistent with the almost equally exposed surface areas of the cysteine sulfur atoms that form the (40–95) and (65–72) disulfide bonds in RNase A and the fourfold more exposed cysteine sulfur atoms of the (30–75) disulfide bond in onconase. Analysis and in silico mutation of the residues around the (40–95) disulfide bond in RNase A, which is analogous to the (30–75) disulfide bond of onconase, reveal that the side chain of tyrosine 92 of RNase A, a highly conserved residue among mammalian pancreatic ribonucleases, lies atop the (40–95) disulfide bond, resulting in a shielding of the corresponding sulfur atoms from the solvent; such burial of the (30–75) sulfur atoms is absent from onconase, due to the replacement of Tyr92 by Arg73, which points away from the (30–75) disulfide bond and into the solvent, resulting in the large exposed surface-area of the cysteine sulfur atoms forming this bond. Removal of Tyr92 from RNase A resulted in the relatively rapid reduction of the mutant to form a single intermediate (des[40–95] Y92A), that is, it resulted in an onconase-like reductive unfolding behavior. The reduction of the P93A mutant of RNase A proceeds through a single

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intermediate, the des-[40–95] P93A species, as in onconase. Although mutation of Pro93 to Ala does not increase the exposed surface area of the (40–95) cysteine sulfur atoms, structural analysis of the mutant reveals that there is greater flexibility in the (40–95) disulfide bond compared to the (65–72) disulfide bond that may make the (40–95) disulfide bond much easier to expose, consistent with the reductive unfolding pathway and kinetics of P93A. Mutation of Tyr92 to Phe92 in RNase A has no effect on its reductive unfolding pathway, suggesting that the hydrogen bond between the hydroxyl group of Tyr92 and the carbonyl group of Lys37 has no impact on the local unfolding free energy required to expose the (40–95) disulfide bond. Thus, these data shed light on the differences between the reductive unfolding pathways of the two homologous proteins and provide a structural basis for the origin of this difference (Narayan et al. 2004). The oxidative folding mechanism of ONC, illustrated in Fig. 15.12, shows markedly different behavior compared to RNase A under similar conditions (Gahl et al. 2008; Gahl and Scheraga 2009). Differences in the intermediates that are generated during the oxidative folding of the two structural homologs, RNase A and ONC, demonstrate that regenerative pathways are not necessarily influenced by tertiary structure. This indicates that the lack of a disulfide bond in ONC, analogous to the (65–72) disulfide bond in RNase A which plays an important role in its oxidative regeneration, does not adversely affect the oxidative folding of ONC (Gahl et al. 2008; Gahl and Scheraga 2009). ONC is an anticancer chemotherapeutic agent (Leland and Raines 2001), and cyclization of its N-terminal residue to pyroglutamic acid increases the activity and stability of this protein. Consequently, we examined the correlated effects of the folding/unfolding process and the formation of this N-terminal pyroglutamic acid (Welker et al. 2007). The results of this study indicate that cyclization of the N-terminal glutamine has no significant effect on the rate of either reductive unfolding or oxidative folding of the protein. Both the cyclized and uncyclized proteins seem to follow the same oxidative folding pathways; however, cyclization altered the relative flux of the protein in these two pathways by increasing the rate of formation of a kinetically trapped intermediate. Glutaminyl cyclase (QC) catalyzed the cyclization of the unfolded, reduced protein but had no effect on the disulfideintact, uncyclized, folded protein. The structured intermediates of uncyclized onconase were also resistant to QC catalysis, consistent with their having a nativelike fold. These observations suggest that, in vivo, cyclization takes place during the

Fig. 15.12 Oxidative folding pathways of ONC, in which no 3SU form was observed (Gahl and Scheraga 2009)

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initial stages of oxidative folding, specifically, before the formation of structured intermediates. The competition between oxidative folding and QC-mediated cyclization suggests that QC-catalyzed cyclization of the N-terminal glutamine in onconase occurs in the endoplasmic reticulum, probably co-translationally.

15.13

Angiogenin

The ribonuclease A homolog ANG induces the formation of blood vessels, making ANG a possible anticancer target for inhibiting formation of blood vessels in tumors. Studies of ANG initially required the availability of large amounts of this protein. This problem was solved (Jang et al. 2004) by preparing recombinant reduced bovine ANG in Escherichia coli, and then developing a procedure to refold the protein from inclusion bodies in high yield. Subsequently, a procedure involving the use of a protein disulfide isomerase fusion system was used to produce soluble recombinant proteins (including ANG) in E. coli without the formation of inclusion bodies (unpublished results). By examining the cis proline content of RNase A, ONC, and ANG, and the folding kinetics of the disulfide-intact forms of these proteins, it was shown (Pradeep et al. 2006) that there is a direct correlation between the number of cis-prolyl bonds in a native protein and the complexity with which it folds via slow phases. By mutating the amino acid residues in the region of ANG that corresponds to a particular surface loop of RNase A, it was found that this surface loop of ANG plays a critical role in maintaining the stability and enzymatic activity of ANG (Jang et al. 2009). Currently, folding/unfolding kinetic studies of ANG are being carried out in the laboratory of my former coworker, H.-C. Shin.

15.14

Summary and Conclusions

The protein-folding problem involves two aspects: determination of (a) the structure of the native protein and (b) the folding pathways to reach the native structure, given the amino acid sequence. We have adopted a physical chemical experimental and theoretical approach to solve this problem. The experimental approach focused on homologous ribonuclease proteins, examining the folding of both disulfideintact and disulfide-reduced forms. This chapter has focused on the experimental approach, which provided distance constraints, such as three tyrosyl–aspartate hydrogen bonds (Scheraga 1967), that motivated our development of the theoretical approach (Ne´methy and Scheraga 1965; Scheraga 1968, 2008; Scheraga et al. 2002) to make use of these distances as constraints for a conformational-energy determination of, first, the three-dimensional structure and, later, the folding pathways leading to the native structure. The development and application of the theoretical approach is discussed elsewhere (Scheraga et al. 2004).

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Our early work on side-chain hydrogen bonds (Laskowski and Scheraga 1954, 1956) and hydrophobic interactions (Ne´methy and Scheraga 1962; Griffith and Scheraga 2004), and conformational changes induced by either alteration of temperature (Hermans and Scheraga 1961a) or addition of denaturing agents (Houry et al. 1994), led to a procedure to determine distance constraints that limited the number of conformations available to a protein. The focus has been on the identification of inter-residue interactions that determine structure and folding pathways (Scheraga et al. 1984; Rothwarf et al. 1998b). Use of homologous ribonucleases, bovine pancreatic ribonuclease A, frog onconase, and bovine angiogenin enabled us to identify common physical and structural features that ultimately are responsible for the biological functions of these proteins. This research has been an excellent vehicle with which to train numerous undergraduate, graduate, and postdoctoral students in protein physical chemistry. Their important contributions are evident in the numerous cited publications. Finally, this research owes much to the financial support from government agencies, initially the Office of Naval Research, and, later and more extensively, the National Institutes of Health, and the National Science Foundation.

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.

Index

A Addition of a protein translocation domain, 66 Aicardi–Goutie`res syndrome, 301, 307 Alba proteins, 327, 330 Amphibian RNases, 19, 20 Amphinase, 58 Amyloid fibrils, 5 Amyotrophic lateral sclerosis (ALS), 8, 24 Angiogenin, 2, 7–12, 20–24, 45, 118, 125, 126, 128 Antibacterial peptides, 9 Antibacterial RNases, 9, 10 Antibody-RNase fusion protein, 71 Antideterminant, 287 Antimicrobial, 101, 102, 106 Antitumor RNases, 14, 17, 18 Apoptosis, 60 Arenavirus endouclease function, 143 structure, 142 Arenaviruses, 140 3’–5’ exoribonuclease, 143 L-protein, 142 AU-rich element (ARE), 226

B Bacteriophage lambda, 270, 278 Bacteriophage T7, 270, 289 Bifunctional enzymes, 310 Bird RNases, 5, 19 Bovine seminal ribonuclease, 61 B1 site, 97, 98 B2 site, 97, 98 Bunyavirus endonuclease

function, 143 metal ion coordination, 142 mutagenesis analysis, 142 Bunyaviruses, 140

C Cap snatching, 140 Cartilage hair hypoplasia, 333 b-CASP, 246, 248–251, 253–262 Catalysis, 320, 322, 323, 326, 328, 329, 332, 334 Catalysis-independent function, 105–106 Catalytic mechanism, 301–302 Cationization, 66 CCA motif, 251, 252 Cell cycle, 333, 334 Cell cycle arrest, 60 Cellular radiation sensitivity, 61 Clinical trials, 73 Correia Repeat Enclosed Elements (CREE), 280 CPSF, 246, 248, 258–262 CREE elements. See Correia Repeat Enclosed Elements (CREE) CRISPR, 290 Cryptic unstable transcripts (CUTs), 227 Csl4, 226 2’,3’ Cyclic phosphate intermediate, 90, 93, 94 Cyclin B2, 333 Cytotoxic, 100–103 Cytotoxic mechanism, 55

D 3D domain swap, 12 Deadenylases, 213–214

A.W. Nicholson (ed.), Ribonucleases, Nucleic Acids and Molecular Biology 26, DOI 10.1007/978-3-642-21078-5, # Springer-Verlag Berlin Heidelberg 2011

399

400 Decrease its affinity for the RI, 65 Defense, 90, 91, 100–103 Degradosome, 194, 196, 198 DGCR8, 285, 288, 289 Dicer, 270, 271, 275, 276, 288 Dimeric variants, 67 Dis3, 225 Discriminator nucleotide, 250 Dis2L, 225 Disulfide-intact folding/unfolding of ribonuclease, 371–373 DnaG, 225, 230 DNA replication, 300, 305, 307, 309 Double-stranded RNA, 269–290 Drosha, 270, 271, 285, 288 Drug resistance, 310 Drugs with the ability to target nucleic acids, 56 DsrA, 282 dsRBD. See dsRNA-binding domain (dsRBD) dsRNA-binding domain (dsRBD), 272–275, 286–289

E ECP. See Eosinophil cationic protein (ECP) EDN. See Eosinophils-Derived Neurotoxin (EDN) Eosinophil-associated ribonucleases (Ears), 42 Eosinophil cationic protein (ECP), 4, 5, 9, 39, 63 Eosinophils, 39 Eosinophils-derived neurotoxin (EDN), 4, 5, 39 Exoribonucleases, 193, 194, 196, 202, 205, 206, 213, 214 5’–3’-exoribonucleases (XRNs) catalytic activity, 170 cellular localization, 172 inhibition by pAp, 171 Mg2+ dependent enzymes, 169, 170 sequence conservation, 168–170 Exosome, 193, 194, 196, 199–201, 207–209, 212, 224, 226

Index H Herpesvirus vhs protein, 149 function, 152 host shutoff, 150 modulation of RNase activity, 152 RNase activity, 150 specificty for mRNA, 151 Histidine motif, 246 HIV–1, 310, 311 Host defense, 37 Housekeeping, 91, 99, 100, 103–105 HRDC (helicase and RNase-D C-termina) domain, 237 Hybrid binding domain (HBD), 303–305 Hydrogen bond interactions, 304 Hydrolysis, 347

I Immunosuppression, 102 Immunosuppressive action, 15, 16 Immunotoxins, 69 Increase its conformational stability, 66 Influenza A virus, 136 endonuclase, 138 metal ion binding, 138 PB2, 139 Influenza A virus endonuclease function, 140 mutagenesis analysis, 138 RNA substrate, 139 structure, 138 substrate specificity, 139 Influenza virus PA, 138 Innate immunity, 307 Insect feeding, 102 Internalizing cell-binding ligands, 73

K Kinetics, 361

F Fish RNases, 2, 21–23

L b-Lactamase, 245–263 Lassavirus NP exonuclease, 143 L-protein, 141

G Gene sorting, 99 Glycosylation, 91

M Mechanism, 347, 348, 350, 353, 354, 356, 360 Metal ion catalysis, 356, 358

Index Metal ions, 322, 326 Metallo-b-lactamase (MbL), 245–263 microRNA (mRNA), 271, 322, 333, 334 Moloney murine leukemia virus, 304, 311 mRNA. See microRNA Mtr3, 225 Mtr4, 225

N Neisseria, 280, 286 NendoU, 145 activities, 146 caltalytic residues, 146 hexameric structure, 149 metal ions, 148 similarity with RNaseA, 146 N-Hydroxyhomophthalimide, 275 Nidovirus, 144 exoribonuclease, 144 Nidovirus endoribonuclease, 145 Nidovirus ExoN, 144 RNA replication fidelity, 145 Noncoding(nc) RNAs, 270, 271, 278, 281–285 Nuclease hydrolytic, 226 phosphorolytic, 226

O Okazaki fragments, 305, 308, 312 Oligoribonuclease, 195, 210–211, 213 ONC internalization, 59 Onconase, 18, 20, 58 Oxidative folding of ribonuclease, 375–387 Oxidative stress, 8, 18, 103, 105, 106

P PacI, 277 PAP I, 202 poly(A) tails, 195 PD-(D/E)XK family of enzymes, 138 Pestivirus Erns RNase, 153 additional exonucleolytic activity, 155 cleavage product, 155 function, 157 kinetic parameters, 155 protection of the viral genome, 156 role in persistent infection, 158 similariety to RNase T2, 154 substrate specificity, 155

401 Pestiviruses, 153 RNase negative mutants, 156 Phosphate binding pocket, 256 Phosphate scavenging, 91, 103–105 Phosphodiester backbone, 305 Phosphorolytic RNase PH, 224 Phylogenetic analysis, 99, 104 PIN domain, 233 Pi-starvation, 103, 104 PM-Scl complex (polymyositis-scleroderma), 226 PNPase, 195–198 KH domain, 197 SI domain, 197 Polynucleotide phosphorylase (PNPase), 224 Poly(A) polymerase, 235 Poly(A) tail, 234 Preclinical studies, 61 Processing, 193, 194, 198–200, 211, 213, 214 Processivity, 235–236, 303–305 Proliferating cell nuclear antigen (PCNA), 306, 307 Promoter upstream transcripts (PROMTs), 227 Protein K homology (KH) domain, 224 Protein–protein interactions, 305, 308 Protein S1 homology domain, 224 Protein synthesis, 8 P1 site, 96

R Rai1 decapping activity toward unmethylated cap, 183 distinct decapping product compared to Dcp2, 181, 183 Mammalian homolog Dom3Z, 178, 183 Mg2+ dependent active site, 169 pyrophosphohydrolase activity, 183 RNA 5’-end capping quality check point, 184, 186 Yeast homolog Ydr370c, 178 RC-RNase, 61 Redirect the RNase to the nucleus, 65 Reductive unfolding of ribonuclease, 370, 387–390 Reptile RNases, 19–20 Reverse transcriptase, 302, 310–312 Ribonuclease, 119, 120, 123, 125, 127, 128 Ribonuclease 7 (RNase 7), 44 Ribonuclease 8 (RNase 8), 44 Ribonuclease III, 269–290

402 Ribonuclease inhibitor, 119, 125 Ribonuclease inhibitor protein, 58 Ribonucleases, 55 Ribonuclease Xrn1, 333 Ribonucleoprotein complex, 320, 322 Ribosomal RNA, 270, 281 Riboswitch, 323 Ribozyme, 326 RJ-RNase, 61 R-loops, 300, 305, 308, 312 RNA, 116, 117, 119–124, 127, 343–362 RNA-based enzyme, 320, 322 RNA:DNA duplex, 300, 304, 308, 312 RNA hydrolysis, 305 RNAIII, 282, 283 RNA interference (RNAi), 270, 288 RNA polymerase, 323 RNA primers, 305 RNase 1, 4, 6, 17, 18 RNase 2, 4 RNase 3, 4–6, 9 RNase 4, 4, 6–7 RNase 5, 3, 7–10, 12, 22 RNase 6, 9, 10 RNase 7, 9–10 RNase 8, 10 RNase A, 1–3, 7, 10–22, 35 RNase A–1, 46 RNase A–2, 46 RNase D, 195, 210–212, 228 DEDD domain, 212 RNase dimers, 12, 18 RNase-E, 230 RNase evolution, 15 RNase H1, 300–305, 308, 311 RNase H2, 300–302, 305–309, 312 RNase HIII, 309–310 RNase II, 198, 200, 202–205, 207–209, 212, 237 CSD domain, 201, 203 RNB domain, 201, 203 SI domain, 201, 203 RNase III, 270–290 RNase inhibitor, 2, 8, 9, 13–19, 142 RNase J, 247, 248, 253–263 RNase J1, 168, 185–186 RNase MRP, 320, 321, 331–334 domains, 332 function, 332, 333 mitochondrial, 331 nucleolar, 331 protein components, 331–334 RNA component, 331–334

Index secondary structure, 332 substrate, 331, 332 RNase oligomers, 67 RNase P, 321–334 activity, 322, 327, 329 archaeal, 321, 322, 326–330 A-type, 321, 323, 326, 328, 329 bacterial, 320–330, 334 B-type, 323 conservation, 322, 324, 326, 332 core, 323, 326 domains, 324, 325, 332 eukaryotic, 320–322, 326–332, 334 holoenzyme, 320, 325–327 human, 323, 329, 332 M-type, 326–329 proteinaceous, 320 protein component, 322–334 RNA component, 321–334 secondary structure, 323 structural organization, 320, 326–328, 330, 331 substrate, 322, 324–326, 329, 332 yeast, 323, 328, 329 RNase pancreatic family, 57 RNase PH, 195–197, 199, 200 RNase R, 195, 197, 200, 201, 204–207, 209, 227 RNases purified from multiple origins, 64 RNase T, 195, 211, 213 RNase T2 conserved active site, 94 crystallographic analysis, 90, 92 disulfide bridges, 93 molecular weight, 91, 92 pH preference, 90, 98 reaction mechanism, 93, 94 RNase Z, 246–248, 250–254, 256, 257, 260, 262 RNA world, 322 rRNA, 253, 254, 262, 332–334 rRNA biogenesis, 7, 24 rRNA degradation, 104, 106 Rrp4, 226 Rrp17, 168, 176, 185 Rrp40, 226 Rrp41, 226 Rrp42, 226 Rrp43, 226 Rrp44, 200, 201, 207–209, 212, 225 CR3 region, 209 PINc domain, 209 PIN domain, 208, 209 Rrp46, 226

Index S Salmonella, 284 Self-incompatibility, 90, 93, 100–103 Seminal RNase, 12, 14–18 Senescence, 103, 104 Single ribonucleotides, 300, 308, 309, 312 Ski, 226 Ski2, 226 Ski7, 226 Ski8, 226 SKI complex (Superkiller), 225 Smart drugs, 68 sRNA, 270, 281, 282, 284, 285, 287, 288 4.5S RNA, 322 S-RNases, 91, 92, 96, 97, 99–102 5.8S rRNA, 332, 333 Staphylococcus, 282, 283 Strategies to endow an RNase with cytotoxic activity, 65 Streptomyces, 279, 280, 288 Stress, 115–128 Stress-induced ribonuclease, 115–128 Stress response, 115, 116, 119, 121, 123, 125, 126 Structure of ribonuclease, 368–370, 385 Structures of XRNs active site region similar to other nucleases, 184 additional domains in Xrn1, 169–178, 181, 184, 185 molecular basis for exonuclease activity, 184 N-terminus near the active site, 180, 181 Rai1 indirectly contributes to catalysis by Rat1, 171 tower domain in Rat1, 180, 181 Substrate binding, 95, 96, 303, 308, 310

T Takadiastase, 90 Tandems of RNase A, 68 TATA-box binding proteins, 310 Tegument, 150 Telomerase, 333 T-loop motif, 324 tmRNA, 323 TRAMP, 212 poly(A) tails, 202, 212, 213

403 TRAMP complex (Trf4/Air2/Mtr4p polyadenylation complex), 225 TRAMP4, 229 TRAMP5, 229 Transcription factor IIF, 308 Transferase, 93 tRNA, 117, 118, 120–122, 124–126, 128, 246, 247, 251–253, 320, 322, 324, 325, 329, 334 Tumor suppression, 105, 106 Tumor targeting of RNases, 70 Two-metal-ion mechanism, 301, 302, 306, 311

V Vertebrates, 21–24 Virulence, 198, 202, 205 Virus-specific RNases, 136

W Wounding, 102

X Xrn1 function in DNA recombination and chromosome stability, 174–175 function in RNA degradation, 172–174 other functions, 175 protein partners, 178–179 Xrn2/Rat1 complex with Rai1 in yeast, 175–177 function in RNA polymerase transcription termination, 176–177 function in RNA processing, 176 other functions, 177 XRNs in plants and other organisms, 179

Y YmdB, 289

Z Zebrafish, 46 ZR (zinc ribbon) domain, 232

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