Protein Biosynthesis

PROTEIN BIOSYNTHESIS No part of this digital document may be reproduced, stored in a retrieval system or transmitted in any form or by any means. The publisher has taken reasonable care in the preparation of this digital document, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained herein. This digital document is sold with the clear understanding that the publisher is not engaged in rendering legal, medical or any other professional services.

PROTEIN BIOSYNTHESIS

TOMA E. ESTERHOUSE AND

LADO B. PETRINOS EDITORS

Nova Biomedical Books New York

Copyright © 2009 by Nova Science Publishers, Inc.

All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers’ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works. Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. LIBRARY OF CONGRESS CATALOGING-IN-PUBLICATION DATA Available Upon Request

ISBN: 978-1-61470-704-2 (eBook)

Published by Nova Science Publishers, Inc.  New York

CONTENTS

Preface

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Chapter I

Protein Synthesis and Ageing Kostoula Troulinaki and Nektarios Tavernarakis

Chapter II

Neuropeptide Biosynthesis in the Nematode Caenorhabditis elegans: From Precursor to Bioactive Peptides Steven J. Husson and Liliane Schoofs

13

Stereochemical Mechanism of Translation Based on Intersubunit Complementarities Kozo Nagano

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Chapter III

1

Chapter IV

Trans-Translation of tmRNA and a Protein Mimicking tRNA and mRNA Hyouta Himeno, Daisuke Kurita, and Akira Muto

Chapter V

Modification of mRNA Translation Initiation to Stimulate Protein Synthesis in Sepsis Thomas C. Vary

Chapter VI

Protein Synthesis in Hepatocytes of Mice as Revealed by Electron Microscopic Radioautography Tetsuji Nagata

Chapter VII

Recent Advances in Label-free BiosensorsApplications in Protein Biosynthesis and HTS Screening Shawn O’Malley

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Translating Memories: The Role of Protein Biosynthesis in Synpatic Plasticity Cara J. Westmark and James S. Malter

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Chapter VIII

vi Chapter IX

Contents Secreted Protein and Peptide Biosynthesis: Precursor Structures and Processing Mechanisms Sergey A. Kozlov, Alexander A. Vassilevski and Eugene V. Grishin

Chapter X

The effects of temperature on ectotherm protein metabolism Nia M. Whiteley and Keiron P. P. Fraser

Chapter XI

Protein Biosynthesis:A new Method for Functional Expression of Sodium-Dependent Glucose Transporter (SGLT) to Study Inhibition of Transport Activity and Drug Discovery Francisco Castaneda

Chapter XII

Chapter XIII

Index

Effect of Hypoxic Conditions on Translational Control of Gene Expression Ota Fuchs The Role of Eukaryotic Translation Intiation Factor 4E and its Binding Factors 4E-BP1 and 4E-BP2 in Body Weight Regulation, Ageing and Tumorigenesis Ota Fuchs

225 249

267

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307 331

PREFACE Protein biosynthesis (synthesis) is the process in which cells build proteins. The term is sometimes used to refer only to protein translation but more often it refers to a multi-step process, beginning with amino acid synthesis and transcription which are then used for translation. Protein biosynthesis, although very similar, differs between prokaryotes and eukaryotes. This new book presents important research in the field from around the globe. Chapter I - Protein synthesis is an essential cellular process affecting growth, reproduction and survival in response to both intrinsic and extrinsic cues such as nutrient availability and energy levels. Studies in many organisms, including humans, have revealed that during ageing, the rate of global protein synthesis declines, indicating a link between the regulation of protein synthesis and the ageing process. Recent studies in C. elegans demonstrate that depletion of specific translation initiation factors, such as eIF4G, eIF2B and eIF4E increases lifespan. Similarly, depletion of specific ribosomal proteins increases lifespan both in yeast and worms. In all cases, these manipulations reduce the rate of general protein synthesis. Why does attenuation of protein synthesis promote longevity? The process of mRNA translation is one of the most energy consuming cellular processes, requiring, depending on growth conditions, up to 50% of the total energy generated by the cell. A reduction of protein synthesis would moderate this energy load, generating an energy surplus, which can be channeled to mechanisms of damage repair and cellular maintenance, thus, extending lifespan. In addition, lowering protein synthesis may be beneficial during ageing by reducing the accumulation of altered, misfolded, aggregated or damaged proteins, as it occurs in many age-related pathologies, such as Alzheimer’s and Parkinson’s disease. The recent experimental findings reveal a key role for protein synthesis in ageing and suggest that perturbation of mRNA translation provides an effective approach for interventions aiming to modulate ageing and senescent decline. Chapter II - Endogenous neuropeptides are small signaling molecules that occur in all metazoan species. They function as neurotransmitters or hormones and orchestrate a wide variety of physiological processes by binding G protein-coupled receptors upon initiation of diverse signaling pathways. Over 115 neuropeptide-encoding genes appear to be present in the soil nematode Caenorhabditis elegans. The biosynthesis of these endogenous biologically active peptides involves a series of enzymatic processing steps, starting from a preproprotein. First, the signal peptide is removed and the remaining part of the precursor will be cleft by

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proprotein convertases at defined motifs displaying basic amino acids. Next, carboxyterminal basic amino acids are removed by the action of a carboxypeptidase. Finally, the carboxyterminal glycine (if present) will be transformed into an amide. A peptidomics platform, which uses state-of-the-art liquid chromatography combined with mass spectrometry, allows us to biochemically identify endogenous peptides present in any tissue or organism. This technology was used to compare the peptide profiles of C. elegans strains having mutations in the presumed peptide precursor processing enzymes. Doing so, we were able to characterize the major processing enzymes KPC-2/EGL-3, CPE/EGL-21 and the chaperone protein 7B2/SBT-1. Chapter III - A universal rule is found in nucleotide sequence complementarities between the regions 2653-2666 in the GTPase binding site of 23S rRNA and 1064-1077 of 16S rRNA as well as between the region 1103-1107 of 16S rRNA and GUUCG (or GUUCA) of tRNAs. This means that there are two extreme cases of conformational states between the above regions. One is responsible for GTP hydrolysis, and the other plays an important role in the structural transitions, particularly for activation of three tRNAs bound to A, P, and E sites in the proofreading step of codon recognition, and in the process of translocation. In order to understand the mechanism of the conformational change, the present author assumed that four kinds of GTPases, viz. aminoacyl-tRNA•EF-Tu•GTP ternary complex, EF-G•GDP complex, peptide chain release factor 3 (RF-3), and initiation factor 2 (IF-2), first binds to the cavity region on the 30S ribosomal subunits, where it is known that an antibiotic spectinomycin binds. Amino acid sequence comparison of effector region for the above four GTPases has shown why the first three GTPases unfolds the region around helix 35 of 16S rRNA and goes to the GTPase- associated region on the 50S subunit, while IF-2 promotes binding of initiator tRNA to the 30S subunit. On the other hand, the crystal structure of the whole ribosome binding three deacylated tRNAs (PDB accession number 1GIX) has shown that the three elbow regions of the tRNAs are distantly separated from each other, resulting in difficulty in explaining the negative cooperativity between A- and E-site tRNAs. Moreover, the existence of a barrier region at the nucleotides G1338 and A1339 of 16S rRNA presents a difficulty in understanding how codon recognition and translocation could occur. The present author explains on the basis of the universal rule of intersubunit complementarities, 1) how the barrier melts before movements of tRNAs, 2) how the cooperative phenomenon in the proofreading is brought about, and 3) how translocation occurs and results in expelling E-site tRNA from the decoding centre. Such a mechanism was reasonably explained by a series of three-dimensional models of rRNAs and proteins that have already been deposited in the PDB. Chapter IV - Usually, a single polypeptide or even multiple polypeptides is produced from a single mRNA. Trans-translation is an irregular translation system in eubacteria, in which a single polypeptide is synthesized from two separate molecules of coding RNAs, mRNA and tmRNA. It rescues a stalled translation on the ribosome and provides a peptide tag for degradation to the C-terminus of the nascent polypeptide to enable recycling of ribosomes, promote degradation of truncated mRNA and prevent accumulation of abortively synthesized polypeptides in the cell. Trans-translation is involved not only in a quality control system in the cell but also in various kinds of cellular functions. During transtranslation, tmRNA plays a dual function both as a tRNA and as an mRNA. Alanyl-tmRNA

Preface

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somehow enters the vacant A-site of the stalled ribosome like aminoacyl-tRNA but without a codon-anticodon interaction, and thereafter the coding region of tmRNA is substituted for mRNA. As a consequence, alanine encoded nowhere but aminoacylated to tmRNA serves as the connector between the truncated peptide encoded by mRNA and the C-terminal tagpeptide encoded by tmRNA. Such an acrobatic feat is accomplished by elaborate interplay between the tRNA and mRNA functions of tmRNA with the help of a protein factor SmpB. Our recent study has suggested that both tmRNA and SmpB mimic the structures and functions of tRNA and mRNA during trans-translation, addressing how tmRNA preferentially recognizes the stalled ribosome, and what substitutes for a codon-anticodon interaction. Chapter V - Sepsis, the systemic manifestation to bacterial infection, induces profound alterations in whole-body protein metabolism. Nitrogen losses up to 17% of total body protein may be observed in septic patients despite aggressive nutritional support. Organ system dysfunction and, eventually, organ failure can result from the persistent loss protein in sepsis. Sustained muscle protein catabolism continues to complicate recovery in septic patients. This review will illuminate potential molecular mechanisms responsible for increasing mRNA translation initiation in striated muscle. Emphasis will be placed on the role of growth hormones and nutrients in promoting rates of protein synthesis during sepsis. In this regard, elevating amino acids and IGF-I both interact to maximally enhance rates of protein synthesis acutely during sepsis through an acceleration of the mRNA translation initiation. IGF-I appears unique in accelerating protein synthesis during sepsis as growth hormone appears to enhance mortality while muscle shows a general resistance to the anabolic actions of insulin. Like IGF-I, amino acids and leucine in particular stimulate mRNA translation initiation by targeting specific signal transduction pathways. The hastening of mRNA translation initiation most likely results from a stimulation of mammalian target of rapamycin (mTOR) acting through its downstream effector proteins to enhance assembly of eIF4G with eIF4E via 4E-BP1 and eIF4G phosphorylation and to increase S6K1 phosphorylation. The physiologic importance lies in the potential ability of IGF-I and amino acids as specific nutrients designed to counteract the accelerated host protein wasting in septic patients and improve nutrition to maintain muscle mass. Chapter VI - For the purpose of studying the aging changes of protein synthesis in mouse hepatocytes, 20 groups of aging mice during development and senescence, each consisting of 3 individuals of both sexes, total 60, from fetal day 19 to postnatal day 1, 3, 9 and 14, month 1, 2 and 6, and year 1 and 2, were injected with RI-labeled amino acids, such as 3H-leucine, 3H-glycine, 3H-proline or 3H-hydroxyproline, which are the protein precursors, sacrificed 1 hr later and the liver tissues were fixed, sectioned and processed for electron microscopic radioautography. On electron microscopic radioautograms obtained from each animal, the localization of silver grains due to 3H-amino acids incorporations showing protein biosynthesis in respective cell organelles, the nucleus, Golgi apparatus, endoplasmic reticulum, mitochondrion and cytoplasmic matrix was qualitatively observed. On the other hand, the numbers of silver grains localizing over respective cell organelles were counted and analyzed quantitatively. The numbers of silver grains localized over the nuclei, Golgi apparatus, endoplasmic reticulum and cytoplasmic matrices increased from perinatal stages to the young adult stage at postnatal month 1, reaching the maximum, then decreased to year 2

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due to aging and senescence. However, the number of silver grains localizing over the mitochondria, i. e. the number of labeled mitochondria per cell and the mitochondrial labeling index in each hepatocyte revealed that the numbers of mitochondria increased gradually from perinatal stages to the postnatal year 2, while the numbers of labeled mitochondria and the labeling indices of intramitochondrial protein syntheses in hepatocytes of mice at various ages increased from embryonic day 19 to postnatal month 6, reaching the maxima, then decreased to year 2 due to aging of animals. Chapter VII- The enzymatic maintenance of biopolymeric structures within a cell is widely known to play a critical role in the regulation of numerous bio-processes from activation, cell signaling and metabolic pathways. This chapter examines the recent advances in label-free biosensing and describes how these technologies have been used to examine protein biosynthesis and protein degradation. Label free biosensing has matured through the years into a powerful technique for examining these processes with quantitative metrology. The emergence of imaging tools with microarray technology in these label free platforms will greatly expand the throughput of these assays thus enabling the user to study globally biosynthetic reactions. Label-free functional enzymatic biosensor assays have recently been applied toward the development of a new generation of protein biosynthesis assays. Labelfree biosensors enable the study of real time biosynthetic and biodegradation reactions while maintaining an open format for exploring modulation factors. When these surface based synthetic and degradation assays are applied in high throughput platforms they provide yet a new screening tool for drug discovery. Chapter VIII - The 1990s, “The Decade of the Brain”, resulted in major scientific advances involving brain imaging, gene therapy, brain/robotic interfacing and the neurobiology and molecular biology of learning and memory. However, despite these critical insights, we still do not know exactly how thoughts or memories are formed or stored in the brain, which leaves much exciting research for the twenty-first and probably centuries to come. This review will elaborate on recent advances in the field of protein biosynthesis as related to synaptic plasticity. We will discuss the molecular players (RNA binding proteins and neuronal mRNAs), the signal transduction pathways that have been implicated in learning and memory and how localized translation of selected mRNAs is involved in synaptic plasticity. We will also discuss the pathology of human diseases including Alzheimer’s disease, Fragile X syndrome, autism and Down syndrome, which show altered or diminished protein synthesis dependent synaptic plasticity. Learning and memory are manifested in their highest form in humans and allow for the retrieval of and action on past events. Understanding the pathology of these neurological disorders will elucidate the normal mechanisms of memory formation and storage. Chapter IX - Protein biosynthesis is rarely restricted to mRNA translation into an amino acid sequence. To yield the mature form, most proteins undergo various posttranslational modifications due to the action of different enzymes. Certain combinations of amino acid residues (primary structure motifs) have been defined to guide the sequence of modifications during the process of precursor protein maturation into the final product. In this chapter, we specifically focus on the secreted polypeptide maturation. For a number of precursor sequences retrieved from UniProt databank, complete sets of enzymes have been identified that execute processing of secreted polypeptides. This finds reflection in the amino acid

Preface

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sequences of the corresponding protein precursors that carry information about the queue of posttranslational events in the form of specific motifs arrangement. Extensive data analysis allowed us to propose a simple set of principals that facilitate effective sequence information handling. Utilization of the proposed principals significantly improves mature protein sequence prediction from available gene structures. We also address the problem of known motif identification and novel motif prediction from large sets of data. A number of proteins are considered in greater detail as examples of the proposed principals utilization conveniences. Chapter X - Protein metabolism in ectotherms is strongly influenced by body temperature and thermal history. In many species, rates of protein synthesis increase with temperature up to a species-specific, thermal optimum. Temperature effects protein synthesis by directly influencing the rates of specific biochemical processes involved in the synthesis of proteins, and also by effecting food consumption. In turn, an increase in food consumption will elevate rates of protein synthesis. Animals have evolved the ability to at least partially compensate rates of protein synthesis as ambient temperatures change, by increasing or decreasing tissue RNA concentrations (RNA to protein) and RNA activity (kRNA). However, at polar temperatures, full compensation of protein synthesis does not appear to occur, and ectotherms are only capable of very low rates of protein synthesis. Temperature also has a direct effect on the proportion of synthesised protein that is degraded, with approximately twice as much protein degraded in polar ectotherms as tropical ectotherms. The result of this is that protein growth at temperatures near the lower limits of life is considerably less efficient than at warmer temperatures. The aim of this review is to examine the effects of temperature on protein metabolism in ectotherms from stable thermal environments and those from more variable thermal regimes. Examples mainly from aquatic environments, will be considered at different levels of biological organisation. In this way, the review will cover both temporal and spatial changes in protein metabolism in ectotherms, with particular interest in those organisms living at thermal extremes. Chapter XI - The sodium-dependent D-glucose transporter (SGLT) family is involved in glucose uptake via intestinal absorption (SGLT1) or renal reabsorption (SGLT1 and SGLT2). SGLT plays an important role in the regulation of glucose blood levels. As a result, increasing attention is being focused on SGLT as a drug target for the therapy of diabetes. Therefore, a selective and specific technique for the study of different potential SGLT inhibitors is mandatory. The expression of functional SGLT is regulated by a complex mechanism involving changes in transcription, mRNA stability, and amount of transporter within the plasma membrane. In addition, SGLT expression depends on the state of cellular differentiation of epithelial cells, which can be observed by confluent cell monolayer growth. Therefore, the use of differentiated epithelial cells represents a unique factor required to obtain functional recombinant SGLT protein that can not be reproduced in other cell systems, such as the brush border membrane, oocytes or liposomes. We found that differentiated Chinese hamster ovary (CHO) cells, either stable or transient transfected with a eukaryotic expression vector containing human SGLT1 or SGLT2 gene, expressed functional SGLT in the cell membrane. The extent of hSGLT1 and hSGLT2 expression was evaluated by relative real-time reverse transcription-polymerase chain reaction (RT-PCR) quantification, Western blotting, and immunocytochemical analysis. Moreover, functional activity of hSGLT1 and

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hSGLT2 activity was determined by measuring the sodium-dependent uptake of α-methyl [14C]-D-glycoside. The advantage of the 96-well method we developed is the low amount of radioactive compounds and inhibitory substances required, and its reproducibility. This method represents an initial approach in the development of transport-based high-throughput screening in the search for inhibitors of glucose transport and the development of new antidiabetic drugs. Chapter XII - Hypoxic conditions, found in limb ischemia, aortic aneurysms, myocardial ischemia and in tumors as well as during normal embryogenesis activate a transcriptional response that promotes vascular development and the formation of red blood cells. The master transcriptional regulator of oxygen-controlled gene expression is the hypoxiainducible factor HIF. Many of the proangiogenic and antiangiogenic factors are directly or indirectly regulated by transcription factor HIF. Hypoxia suppresses protein synthesis at the level of mRNA translation initiation in many nontransformed cells whereas highly transformed cells are largely resistant. Two different pathways are involved in response of mRNA translation initiation in cells to hypoxia and it results in biphasic inhibition of translation. The first pathway is associated with endoplasmic reticulum (ER) stress and through it with activation of unfolded protein response. Transient phosphorylation of eukaryotic initiation factor (eIF) 2 alpha in the first phase by double-stranded RNA-activated protein kinase-like endoplasmic reticulum kinase PERK inhibits mRNA translation initiation. The second phase occurs more slowly and is independent on eIF2alpha and is connected with eIF4F (containing the cap- binding protein eIF4E, the scaffold protein eIF4G and the RNA helicase eIF4A) disruption and with inactivation of the ternary complex (eIF2/MettRNA/GTP). The availability of the cap-binding protein eIF4E is rate-limiting under normal conditions. During tumorigenesis eIF4E is often over expressed because eIF4E-binding proteins sequester eIF4E in hypophosphorylated form. Hyperphosphorylation of the eIF4Ebinding proteins lowers their affinity for eIF4E, resulting in an increased interaction between eIF4E and eIF4G and stimulation of translation. Phosphorylation of eIF4E-binding proteins is largely controlled by the mammalian target of rapamycin (mTOR) kinase. Hypoxia inhibits the activation of kinase mTOR and results in hypophosphorylated eIF4E-binding proteins and in increased their affinity for eIF4E and in decreased association between eIF4E and eIF4G necessary for eIF4F disruption. Inhibition of the kinase mTOR suppresses mRNA translation also through a novel mechanism mitigated in transformed cells. This mechanism is based on disruption of proteasome-targeted degradation of eukaryotic elongation factor 2 (eEF2) kinase. However, regulation of translation also results in a specific increase of the synthesis of a subset of hypoxia-induced proteins as activating transcription factor 4 (ATF4), CCAAT/enhancer binding protein homologous protein (CHOP, also named GADD153), growth arrest and DNA damage inducible protein (GADD34), hypoxia inducible factor (HIF1alpha), vascular endothelial growth factor (VEGF), platelet-derived growth factor (PDGF) and immunoglobulin heavy chain binding protein (BiP). Translation in these cases is often cap independent due to the presence of an internal ribosome entry site (IRES) in the 5´ noncoding region of some these mRNAs. Hypoxic tumor cells that are target of tumor therapy are exposed to additional endoplasmic reticulum stress by using proteasome inhibitors, such as PS-341 (Velcade), which is Food and Drug Administration approved for

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treating human malignancies. This therapy may be selectively cytotoxic to hypoxic tumor cells. Chapter XIII - Translation in eukaryotes is usually regulated at the level of initiation. Translation initiation on the majority of eukaryotic cellular mRNAs is mediated by a cap deperndent mechanism. Eukaryotic translation initiation factor 4E (eIF4E) binds to the mRNA cap structure and interacts with the RNA helicase eIF4A and a large scaffold protein eIF4G to create the eIF4F complex. Binding of eIF4E to eIF4G can be blocked by eIF4Ebinding proteins (4E-BP1 and 4E-BP2) acting as competitive inhibitors of eIF4E-eIF4G interaction. Scaffold protein eIF4G brings the mRNA to the 40 S small ribosomal subunit in a complex with eIF2, GTP and the initiator methionine-transfer RNA by its interaction with eIF3 and forms the 48 S preinitiation complex on the mRNA. Scanning of the mRNA with the aid of eIF4A and recognition of initiation AUG start codon in an optimal context is required and other factors as well as the 60S ribosomal subunit are then recruited and polypeptide chain elongation begins. 4E-BP1 and 4E-BP2 double knockout (DKO) mice were used to determine the physiologic functions of these factors. 4E-BP1 and 4E-BP2 DKO mice had a significant increase in both body weight and fat content. The obese phenotype was caused by reduced energy expenditure and reduced lipolysis. Both embryonic fibroblasts and preadipocytes from these 4E-BP1 and 4E-BP2 DKO mice had an increased expression of CCAAT/enhancer-binding proteins and of peroxisome proliferator-activated receptor (PPAR) gamma, essential regulators of adipogenesis. Specific knockdown of the main eIF4E isoform in Caenorhabditis elegans resulted in an increased lifespan of the organism. Because eIF4E is the least abundant among translation initiation factors, changes in the levels of this translation initiation factor affect translation rates, preferentially of a subset of mRNAs with strong secondary structure in the 5´ untranslated region encoding proteins such as Myc, fibroblast growth factor, ornithine decarboxylase, cyclin D1, survivin, Bcl-2, matrix metalloprotease 9 and vascular endothelial growth factor. These mRNAs play important roles in the control of cell growth, proliferation, angiogenesis, survival and malignancy. In experimental models, eIF4E overexpression induces cellular transformation, tumorigenesis, invasion, and metastasis, notably cancers as lymphomas, lung and prostatic adenocarcinomas, bladder cancers, cervical cancers, hepatomas, breast tumorigenesis, head and neck cancers, colorectal cancers and angiosarcomas. A better understanding of the role of eIF4E and its binding proteins (eIF4E-BP) in regulating the translation of the diverse gene products involved in tumorigenesis will improve the capacity to exploit eIF4E and eIF4E-BP as therapeutic targets and as markers for human cancer progression. Modulators of the eIF4E activity by peptides containing an eIF4E-binding site, RNA aptamers which bind eIF4E, eIF4E-specific antisense oligonucleotides and small molecule inhibitors of the eIF4E-eIF4G interaction are tested and may be in future used in therapy for the treatment of cancer.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter I

PROTEIN SYNTHESIS AND AGEING Kostoula Troulinaki and Nektarios Tavernarakis∗ Institute of Molecular Biology and Biotechnology, Foundation for Research and Technology, Heraklion 71110, Crete, Greece.

ABSTRACT Protein synthesis is an essential cellular process affecting growth, reproduction and survival in response to both intrinsic and extrinsic cues such as nutrient availability and energy levels. Studies in many organisms, including humans, have revealed that during ageing, the rate of global protein synthesis declines, indicating a link between the regulation of protein synthesis and the ageing process. Recent studies in C. elegans demonstrate that depletion of specific translation initiation factors, such as eIF4G, eIF2B and eIF4E increases lifespan. Similarly, depletion of specific ribosomal proteins increases lifespan both in yeast and worms. In all cases, these manipulations reduce the rate of general protein synthesis. Why does attenuation of protein synthesis promote longevity? The process of mRNA translation is one of the most energy consuming cellular processes, requiring, depending on growth conditions, up to 50% of the total energy generated by the cell. A reduction of protein synthesis would moderate this energy load, generating an energy surplus, which can be channeled to mechanisms of damage repair and cellular maintenance, thus, extending lifespan. In addition, lowering protein synthesis may be beneficial during ageing by reducing the accumulation of altered, misfolded, aggregated or damaged proteins, as it occurs in many age-related pathologies, such as Alzheimer’s and Parkinson’s disease. The recent experimental findings reveal a key role for protein synthesis in ageing and suggest that perturbation of mRNA translation provides an effective approach for interventions aiming to modulate ageing and senescent decline.

∗

Correspondence concerning this article should be addressed to: Nektarios Tavernarakis, Institute of Molecular Biology and Biotechnology, Foundation for Research and Technology, Vassilika Vouton, P.O.Box 1527, Heraklion 71110, Crete, GREECE. tel: +30 2810 391066; fax: +30 2810 391067; email: [email protected].

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Kostoula Troulinaki and Nektarios Tavernarakis∗

Keywords: Ageing, Caenorhabditis elegans, Lifespan, mRNA translation regulation, Protein modification

INTRODUCTION To maintain their homeostasis and function, cells must continuously synthesize and degrade proteins in a highly regulated manner. The mechanisms of transcription and mRNA translation regulate the synthesis of new proteins. Damaged or aggregated proteins are removed by specialized and selective protein degradation mechanisms (Ding et al., 2007). Studies in various organisms, ranging from yeast to humans, revealed that both protein synthesis and protein degradation rates change during ageing (Makrides, 1983; Partridge and Gems, 2002; Rattan, 1996). The activity of key translation factors appears to decline with age, resulting in reduction of protein synthesis rates (Kimball et al., 1992; Moldave et al., 1979; Takahashi et al., 1985; Vargas and Castaneda, 1981, 1983; Webster and Webster, 1983). However, it was unclear whether these changes were simply a consequence of the general deterioration of the cellular functions that characterize ageing or they had a causative role in the process. Recent studies in the nematode C. elegans revealed that impeding mRNA translation significantly affects longevity, indicating that the levels of protein synthesis may affect ageing (Hansen et al., 2007; Kaeberlein and Kennedy, 2007; Pan et al., 2007; Syntichaki et al., 2007).

DOWN-REGULATION OF PROTEIN SYNTHESIS EXTENDS LIFESPAN Protein synthesis or the translation of mRNA is a conserved process involving three main steps: initiation, elongation and termination. Initiation of translation requires the concerted action of a large number of proteins known as translation initiation factors (eIFs). These factors recognize the cap structure at the 5’ end of mRNA and allow the binding of the 40S ribosome subunit that scans downstream for the initiation codon. The next step is the elongation, during which the fully assembled ribosome reads the transcript and uses amino acid-charged tRNAs to synthesize the peptide chain, with the participation of elongation factors. Finally, the process is terminated when a release factor binds to the final (STOP) codon and releases the complete polypeptide from the ribosome that breaks apart into its two subunits (Gebauer and Hentze, 2004; Proud, 2007). All steps of protein synthesis are tightly regulated and are carried out with superb precision, ensuring the fidelity of the proteins. Many studies have demonstrated that mRNA translation fidelity does not change during ageing (Filion and Laughrea, 1985). However, it has been shown that the rate of protein synthesis declines with age in a variety of organisms (Makrides, 1983; Partridge and Gems, 2002; Rattan, 1996). Initiation, the first step of mRNA translation, is a rate-limiting process and the most common target of protein synthesis control. The initiation factor eIF4E plays a key role in the

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process, by recognizing the 5’-end cap structure of most eukaryotic mRNAs and facilitating their recruitment to the ribosomes (Gingras et al., 1999). In the nematode C. elegans, five eIF4E isoforms, termed IFE-1 - IFE-5, are encoded in the genome (Keiper et al., 2000). IFE1, IFE-3 and IFE-5 are expressed in the germline, whereas IFE-2 and IF-4 are expressed specifically in somatic cells (Keiper et al., 2000). Depletion of the isoforms that are expressed in the germline, or of the somatic isoform IFE-4, does not affect lifespan (Syntichaki et al., 2007). However, loss of IFE-2, which is the most abundant in the soma causes significant extension of lifespan (Syntichaki et al., 2007). This result indicates that down-regulation of protein synthesis specifically in the soma extends lifespan. Interestingly, the observed lifespan extension does not require a functional germline, since lack of germline does not suppress the effect of IFE-2 deficiency (Syntichaki et al., 2007). Notably, depletion of the IFE-1 during adulthood leads to moderate adult lifespan extension, indicating that IFE-1 also modulates longevity (Pan et al., 2007). Other studies in the nematode have further revealed that elimination of other translation initiation factors or their regulators, after completion of the development of the organism, results in similar effects on adult lifespan. For example, knocking down of eIF4G (ifg-1) by RNAi or the eIF2 beta subunit (iftb-1) during adulthood results in 30% increase of lifespan (Chen et al., 2007; Hansen et al., 2007; Pan et al., 2007). In addition, RNAi with several ribosomal proteins or the ribosomal-protein S6 kinase (S6K) –after completion of development- leads to increased lifespan. In all cases, the rate of protein synthesis in “longlived” animals is reduced, compared to wild type control animals (Hansen et al., 2007; Pan et al., 2007). Furthermore, an RNAi screen for essential genes that extend lifespan when inactivated post-developmentally has revealed many genes encoding for several components of the eIF complex and the ribosome. These include C. elegans homologs of eIF2G, eIF3F and eIF4A (Chen et al., 2007; Curran and Ruvkun, 2007).

SIGNALING PATHWAYS THAT REGULATE PROTEIN SYNTHESIS AND AGEING Translation of mRNA is a highly regulated process that enables the cell to fine-tune gene expression by stimulating or repressing translation of specific mRNAs, usually through the reversible phosphorylation of mRNA translation factors. Various signaling pathways, activated by hormones, growth factors and nutrients regulate protein synthesis (Figure 1). For example, the insulin-like pathway, the TOR pathway and the MAPK pathway are key signal transduction pathways implicated in ageing that also modulate protein synthesis (Gingras et al., 2004; Proud, 2007). The insulin-IGF-1 pathway plays a vital role in the regulation of somatic growth and cellular proliferation and in parallel is a key modulator of ageing in various organisms (Guarente and Kenyon, 2000). The pathway is engaged by the insulin receptor that binds the insulin ligand and phosphorylates the phosphatidylinositol 3 kinase (PI3K) that generates phosphatidylinositol-3,4,5-trisphosphate (PtdIns(3,4,5)P3) or phosphatidylinositol-3,4biphosphate (PtdIns(3,4)P2), which in turn activate the 3-phosphoinositide-dependent protein kinase 1 (PDK1). Subsequently, PDK1 activates the serine-threonine protein kinase

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Kostoula Troulinaki and Nektarios Tavernarakis∗

AKT/protein kinase B (PKB). These kinases target the FOXO transcription factor DAF-16 and block its transcriptional activity. Mutations that down-regulate this signaling cascade- for example in the insulin/IGF-like receptor DAF-2 or the phosphatidylonisol-3-OH kinase (PI3K) AGE-1 have been found to extend nematode life span. The extension is dependent on the activity of DAF-16, a forkhead (FOXO) transcription factor that controls the expression of a variety of genes involved in stress resistance (superoxide dismutase, catalase, glutathione, heat-shock protein 16 and others), metabolism (apolipoproteins, glyoxylate cycle and cytochrome P450s) fat accumulation and fertility (Gems and Partridge, 2001; Kenyon, 2005). Similarly, mutations in the insulin-IGF-1 pathway in Drosophila (in the insulin-like receptor), also increase the lifespan of flies (Clancy et al., 2001; Tatar et al., 2001). For example, a mild reduction of the insulin-like receptor (Inr) increases mean female lifespan by up to 85% (Partridge and Gems, 2002). In mammals, different receptors for insulin and IGF-1 participate in divergent pathways in different tissues (Yang et al., 2005). Mutations that down-regulate either the insulin pathway or the IGF pathway, result in prolonged lifespan (Bluher et al., 2003; Flurkey et al., 2001; Holzenberger et al., 2003). In addition, such long lived mice have decreased rate of protein synthesis compared with the control animals (Hsieh and Papaconstantinou, 2004). TOR (target of rapamycin) signaling is stimulated by serum, insulin and growth factors, and promotes protein synthesis through multiple outputs (Gingras et al., 2004; Proud, 2007). The most characterized effector is the ribosomal S6 kinase (S6K), which induces mRNA translation through phosphorylation of ribosomal protein S6 and through regulation of translation initiation factors, such as eukaryotic initiation factor 4B (eIF4B) (Proud, 2004). In addition, TOR signaling promotes translation by regulating the activity of the initiation factor eIF4E. More specifically, TOR phosphorylates the eIF4E–binding protein (4E-BP) and liberates the eIF4E, enabling it to interact with eIF4G and to form a complex competent to mediate cap-dependent translation. Moreover, it has been found that the TOR pathway promotes transcription of genes encoding for ribosomal proteins. In many cell types, the TOR pathway is impaired under amino acid starvation. In this case many of the above proteins undergo dephosphorylation. Studies in C. elegans revealed that TOR deficiency, which dampens the rate of translation, extends lifespan (Vellai et al., 2003). MAPK signaling pathway is also clearly linked with the control of the protein synthesis, by affecting a number of mRNA translational machinery components to promote the assembly of initiation factor complexes and the activation of the elongation machinery (Hsieh and Papaconstantinou, 2004; Proud, 2007). It contains several modules of which the best understood are the classical MAPK (ERK), p38 MAPK α/β and JNK (c-Jun N-terminal kinase) pathways. Each involves kinases that phosphorylate components of the translational machinery and/or other proteins that regulate mRNA translation. ERK activates the protein kinases RSKs (or p90RSKs) that phosphorylate other kinases leading to activation of the TOR pathway. Moreover, RSKs phosphorylate at least two other proteins involved in translational control: the eEF2 kinase and eIF4B, promoting its association with eIF3. p38 MAPK α/β activates MK-2, which regulates the stability of mRNAs, probably through the phosphorylation of ARE-binding proteins. In addition, the MAPK signaling pathway leads to the phosphorylation and activation of the kinases Mnk1 and 2. Mnks bind to eIF4G and mediate eIF4E phosphorylation (Waskiewicz et al., 1999). The physiological significance of

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Figure 1. Signaling pathways that modulate protein synthesis by regulating the activity of specific translation factors. Insulin-insulin growth factor 1 (INS-IGF-1) signaling is activated by the binding of insulin or IGF-1 to the insulin receptor and results in the activation of the phosphatidylinositol 3 kinase (PI3K). PI3K converts phosphatidylinositol (4,5) –bisphosphate (PIP2) to phosphatidylinositol (1,4,5)trisphosphate (PIP3). PIP3 in turn activates the serine threonine protein kinase Akt (also known as protein kinase B; Akt/PKB) which phosphorylates and activates the S6 kinase (S6K), while it suppresses the serine threonine protein kinase GSK3 (glycogen synthase 3). The GSK3 kinase regulates the activity of the eukaryotic translation initiation factor 2B (eIF2B). S6K phosphorylates the small ribosomal subunit S6 and the eukaryotic initiation factor 4B (eIF4B). S6K can also be activated by the target of rapamycin (TOR) signaling pathway. In addition to S6K, the TOR pathway leads to the phosphorylation of the eukaryotic initiation factor 4E-binding protein (4E-BP), which inhibits protein synthesis by blocking the eukaryotic translation initiation factor 4E (eIF4E). In addition, TOR signaling can be activated by the mitogen activated protein kinase (MAPK) signaling cascade. The MAPK pathway stimulates the RSKs or p90RSKs kinases that phosphorylate the eukaryotic translation initiation factor 4B (eIF4B) and the kinase of the eukaryotic elongation factor 2 (eEF2K). Moreover, MAPK signaling activates the mitogen-activated protein kinase-interacting kinases (Mnks) that in turn phosphorylate the eukaryotic translation initiation factor 4E (eIF4E) and the eukaryotic translation initiation factor 4G (eIF4G). Arrows indicate positive regulation events whereas the bar lines indicative negative regulation events.

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this phosphorylation is still unclear. However, it is notable that such phosphorylation is generally mediated by mitogen- or stress- and cytokine-activated signaling. In addition to eIF4E, Mnks also phosphorylate eIF4G.

PROTEIN DAMAGE DURING AGEING One of the most common symptoms of ageing at the molecular level is the accumulation of altered proteins both within the cells and extracellularly (Hipkiss, 2006; Rothstein, 1975, 1979, 1989). Protein damage may result in the loss of protein function or it can also lead to protein aggregation. In the latter case, the interaction of damaged proteins with normal cellular proteins may cause sequestration and inhibition of key molecules, like transcription factors, cytoskeletal proteins, molecular chaperones and hydrolytic enzymes (Hipkiss, 2006). Although protein modifications are continuously generated, through a variety of processes, cellular homeostatic mechanisms either suppress the formation of altered proteins, or enhance their destruction, thereby preventing their accumulation. Such protective mechanisms are the lysosomal and proteasomal pathways. Many studies, involving both biochemical and micro array expression assays, have shown that proteolytic activity decreases with age in many cell types (Gems and McElwee, 2003; Makrides, 1983; Martinez-Vicente et al., 2005; Sarkis et al., 1988; Szweda et al., 2002). Certain pathways of lysosomal protein degradation, such as macroautophagy and chaperone-mediated autophagy, exhibit age-dependent decline in function (Arslan et al., 2006). In addition, alterations in the activity of certain lysosomal enzymes, including cathepsins, have been reported to occur during ageing (Sarkis et al., 1988). Moreover, decline in the function of the proteasome during ageing has been observed in cultured cells and in tissues from various organisms, resulting in an increased half-life of oxidized proteins (Sitte et al., 2000a; Sitte et al., 2000b). This can be attributed to down regulation of genes that encode proteasome subunits and the accumulation of proteasome inhibitory proteins as a function of ageing. Interestingly, many age-related pathologies such as Alzheimer’s disease, Parkinson’s disease and atherosclerosis are characterized by increased levels of altered proteins, which are thought to be the cause of aged-related pathology (Hipkiss, 2006).

WHY IS LIFESPAN EXTENDED WHEN PROTEIN SYNTHESIS IS REDUCED? It is well known that mRNA translation is one of the most energy-consuming cellular processes (Proud, 2002). The addition of a single amino acid in the polypeptide chain during mRNA translation requires the energy released by the hydrolysis of four ATP molecules. The amount of total cellular energy that is devoted to translation varies between different cell types and growth states. For example, dividing cells spend more energy on mRNA translation than post-mitotic cells, given their higher requirement for protein synthesis. In all cases, a high proportion of cellular metabolic energy is used in translation and almost all is consumed

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during the elongation phase. Therefore, reduction of protein synthesis would lead to significant conservation of cellular energy. Extra energy could be channeled towards mechanisms of maintenance and repair, contributing to cell survival under stress conditions, such as oxidative stress (Figure 2). Interestingly, the basic concept of the “disposable soma” theory of ageing is that soma is mortal and frail because fails to divert energy towards repairing stochastic damage that accumulates during life (Kirkwood, 1977; Kirkwood and Austad, 2000). By contrast, the germ line may achieve immortality by investing most of the energy to mechanisms of repair.

Figure 2. A working model linking down regulation of protein synthesis to prolonged lifespan. A) Normal lifespan: Cellular energy is distributed between both protein synthesis and repair. However, since protein synthesis is a highly energy-consuming process, the energy remaining for mechanisms of repair and maintenance is limited, resulting in progressive structural and functional deterioration and ageing. B) Increased lifespan: Down-regulation of protein synthesis may lead to prolonged lifespan by allowing for more cellular energy to be channeled towards repair mechanisms, enabling the cell to better withstand stress.

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In support of this hypothesis, it has been found that ife-2 C. elegans mutants, which are defective for a somatic isoform of the translation initiation factor eIF4E and have reduced rate of protein synthesis, are more resistant than wild type animals to oxidative stress induced by the herbicide paraquat and/or sodium azide, an inhibitor of the respiratory chain cytochrome c oxidase, (Syntichaki et al., 2007). Moreover, IFE-2 deficiency increases oxidative stress resistance and extends lifespan of mev-1 mutants that are continuously under oxidative stress due to their lack of the cytochrome b large subunit in complex II of the mitochondrial electron transport chain (Ishii et al., 1998; Syntichaki et al., 2007). Resistance to oxidative stress corresponds to increased capability for detoxification and repair, indicating a higher capacity for damage repair in these mutants. Thus, down regulation of protein synthesis in the soma, due to elimination of a specific initiation factor of translation (eIF4E) leads to increased oxidative stress resistance and increased lifespan.

CONCLUDING REMARKS AND FUTURE PROSPECTS Protein synthesis and protein degradation are the two essential processes that determine the rate of cellular protein turnover. The recent finding that down-regulation of mRNA translation leads to increase of lifespan establishes a direct link between protein synthesis and ageing (Hansen et al., 2007; Pan et al., 2007; Syntichaki et al., 2007). However, the exact mechanism through which protein biosynthesis affects ageing still remains unknown. Given that mRNA translation is one of the most energy consuming processes; its reduction would result in notable energy savings. This energy could be diverted to cellular repair and maintenance processes, thus contributing to longevity. Moreover, reduction of mRNA translation may prevent the synthesis of unwanted proteins that could interfere with the cellular stress response. Interestingly, under stress, global mRNA translation is attenuated, while there is a switch to selective translation of proteins that are required for cell survival under stress (Clemens, 2001; Holcik and Sonenberg, 2005). The mechanisms that regulate this switch are poorly understood. Mild stress is known to stimulate maintenance and repair mechanisms, a phenomenon known as “hormesis” (Mattson, 2008). Hormesis is associated with reduced accumulation of damaged proteins, stimulation of proteasomal activity and increased cellular resistance to toxic agents (Cypser and Johnson, 2002; Rattan, 2004). Hormesis has also been found to prolong lifespan. It is possible that hormesis depends on lowering protein synthesis to levels that increase energy availability but enable production of essential and vital proteins. In this context, it remains to be seen whether reducing protein synthesis is part of hormetic effects on ageing.

ACKNOWLEDGEMENTS Work in the authors’ laboratory is funded by grants from the European Union 6th Framework Programme, the European Molecular Biology Organization (EMBO) and the Institute of Molecular Biology and Biotechnology.

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REFERENCES Arslan, M.A., Csermely, P., and Soti, C. (2006). Protein homeostasis and molecular chaperones in aging. Biogerontology 7, 383-389. Bluher, M., Kahn, B.B., and Kahn, C.R. (2003). Extended longevity in mice lacking the insulin receptor in adipose tissue. Science 299, 572-574. Chen, D., Pan, K.Z., Palter, J.E., and Kapahi, P. (2007). Longevity determined by developmental arrest genes in Caenorhabditis elegans. Aging Cell 6, 525-533. Clancy, D.J., Gems, D., Harshman, L.G., Oldham, S., Stocker, H., Hafen, E., Leevers, S.J., and Partridge, L. (2001). Extension of life-span by loss of CHICO, a Drosophila insulin receptor substrate protein. Science 292, 104-106. Clemens, M.J. (2001). Translational regulation in cell stress and apoptosis. Roles of the eIF4E binding proteins. J Cell Mol Med 5, 221-239. Curran, S.P., and Ruvkun, G. (2007). Lifespan regulation by evolutionarily conserved genes essential for viability. PLoS Genet 3, e56. Cypser, J.R., and Johnson, T.E. (2002). Multiple stressors in Caenorhabditis elegans induce stress hormesis and extended longevity. J Gerontol A Biol Sci Med Sci 57, B109-114. Ding, Q., Cecarini, V., and Keller, J.N. (2007). Interplay between protein synthesis and degradation in the CNS: physiological and pathological implications. Trends Neurosci 30, 31-36. Filion, A.M., and Laughrea, M. (1985). Translation fidelity in the aging mammal: studies with an accurate in vitro system on aged rats. Mech Ageing Dev 29, 125-142. Flurkey, K., Papaconstantinou, J., Miller, R.A., and Harrison, D.E. (2001). Lifespan extension and delayed immune and collagen aging in mutant mice with defects in growth hormone production. Proc Natl Acad Sci U S A 98, 6736-6741. Gebauer, F., and Hentze, M.W. (2004). Molecular mechanisms of translational control. Nat Rev Mol Cell Biol 5, 827-835. Gems, D., and McElwee, J.J. (2003). Ageing: Microarraying mortality. Nature 424, 259-261. Gems, D., and Partridge, L. (2001). Insulin/IGF signalling and ageing: seeing the bigger picture. Curr Opin Genet Dev 11, 287-292. Gingras, A.C., Raught, B., and Sonenberg, N. (1999). eIF4 initiation factors: effectors of mRNA recruitment to ribosomes and regulators of translation. Annu Rev Biochem 68, 913-963. Gingras, A.C., Raught, B., and Sonenberg, N. (2004). mTOR signaling to translation. Curr Top Microbiol Immunol 279, 169-197. Guarente, L., and Kenyon, C. (2000). Genetic pathways that regulate ageing in model organisms. Nature 408, 255-262. Hansen, M., Taubert, S., Crawford, D., Libina, N., Lee, S.J., and Kenyon, C. (2007). Lifespan extension by conditions that inhibit translation in Caenorhabditis elegans. Aging Cell 6, 95-110. Hipkiss, A.R. (2006). Accumulation of altered proteins and ageing: causes and effects. Exp Gerontol 41, 464-473. Holcik, M., and Sonenberg, N. (2005). Translational control in stress and apoptosis. Nat Rev Mol Cell Biol 6, 318-327.

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Holzenberger, M., Dupont, J., Ducos, B., Leneuve, P., Geloen, A., Even, P.C., Cervera, P., and Le Bouc, Y. (2003). IGF-1 receptor regulates lifespan and resistance to oxidative stress in mice. Nature 421, 182-187. Hsieh, C.C., and Papaconstantinou, J. (2004). Akt/PKB and p38 MAPK signaling, translational initiation and longevity in Snell dwarf mouse livers. Mech Ageing Dev 125, 785-798. Ishii, N., Fujii, M., Hartman, P.S., Tsuda, M., Yasuda, K., Senoo-Matsuda, N., Yanase, S., Ayusawa, D., and Suzuki, K. (1998). A mutation in succinate dehydrogenase cytochrome b causes oxidative stress and ageing in nematodes. Nature 394, 694-697. Kaeberlein, M., and Kennedy, B.K. (2007). Protein translation, 2007. Aging Cell 6, 731-734. Keiper, B.D., Lamphear, B.J., Deshpande, A.M., Jankowska-Anyszka, M., Aamodt, E.J., Blumenthal, T., and Rhoads, R.E. (2000). Functional characterization of five eIF4E isoforms in Caenorhabditis elegans. J Biol Chem 275, 10590-10596. Kenyon, C. (2005). The plasticity of aging: insights from long-lived mutants. Cell 120, 449460. Kimball, S.R., Vary, T.C., and Jefferson, L.S. (1992). Age-dependent decrease in the amount of eukaryotic initiation factor 2 in various rat tissues. Biochem J 286 ( Pt 1), 263-268. Kirkwood, T.B. (1977). Evolution of ageing. Nature 270, 301-304. Kirkwood, T.B., and Austad, S.N. (2000). Why do we age? Nature 408, 233-238. Makrides, S.C. (1983). Protein synthesis and degradation during aging and senescence. Biol Rev Camb Philos Soc 58, 343-422. Martinez-Vicente, M., Sovak, G., and Cuervo, A.M. (2005). Protein degradation and aging. Exp Gerontol 40, 622-633. Mattson, M.P. (2008). Hormesis defined. Ageing Res Rev 7, 1-7. Moldave, K., Harris, J., Sabo, W., and Sadnik, I. (1979). Protein synthesis and aging: studies with cell-free mammalian systems. Fed Proc 38, 1979-1983. Pan, K.Z., Palter, J.E., Rogers, A.N., Olsen, A., Chen, D., Lithgow, G.J., and Kapahi, P. (2007). Inhibition of mRNA translation extends lifespan in Caenorhabditis elegans. Aging Cell 6, 111-119. Partridge, L., and Gems, D. (2002). Mechanisms of ageing: public or private? Nat Rev Genet 3, 165-175. Proud, C.G. (2002). Regulation of mammalian translation factors by nutrients. Eur J Biochem 269, 5338-5349. Proud, C.G. (2004). Role of mTOR signalling in the control of translation initiation and elongation by nutrients. Curr Top Microbiol Immunol 279, 215-244. Proud, C.G. (2007). Signalling to translation: how signal transduction pathways control the protein synthetic machinery. Biochem J 403, 217-234. Rattan, S.I. (1996). Synthesis, modifications, and turnover of proteins during aging. Exp Gerontol 31, 33-47. Rattan, S.I. (2004). Aging, anti-aging, and hormesis. Mech Ageing Dev 125, 285-289. Rothstein, M. (1975). Aging and the alteration of enzymes: a review. Mech Ageing Dev 4, 325-338. Rothstein, M. (1979). The formation of altered enzymes in aging animals. Mech Ageing Dev 9, 197-202.

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Rothstein, M. (1989). An overview of age-related changes in proteins. Prog Clin Biol Res 287, 259-267. Sarkis, G.J., Ashcom, J.D., Hawdon, J.M., and Jacobson, L.A. (1988). Decline in protease activities with age in the nematode Caenorhabditis elegans. Mech Ageing Dev 45, 191201. Sitte, N., Huber, M., Grune, T., Ladhoff, A., Doecke, W.D., Von Zglinicki, T., and Davies, K.J. (2000a). Proteasome inhibition by lipofuscin/ceroid during postmitotic aging of fibroblasts. Faseb J 14, 1490-1498. Sitte, N., Merker, K., von Zglinicki, T., and Grune, T. (2000b). Protein oxidation and degradation during proliferative senescence of human MRC-5 fibroblasts. Free Radic Biol Med 28, 701-708. Syntichaki, P., Troulinaki, K., and Tavernarakis, N. (2007). eIF4E function in somatic cells modulates ageing in Caenorhabditis elegans. Nature 445, 922-926. Szweda, P.A., Friguet, B., and Szweda, L.I. (2002). Proteolysis, free radicals, and aging. Free Radic Biol Med 33, 29-36. Takahashi, R., Mori, N., and Goto, S. (1985). Accumulation of heat-labile elongation factor 2 in the liver of mice and rats. Exp Gerontol 20, 325-331. Tatar, M., Kopelman, A., Epstein, D., Tu, M.P., Yin, C.M., and Garofalo, R.S. (2001). A mutant Drosophila insulin receptor homolog that extends life-span and impairs neuroendocrine function. Science 292, 107-110. Vargas, R., and Castaneda, M. (1981). Role of elongation factor 1 in the translational control of rodent brain protein synthesis. J Neurochem 37, 687-694. Vargas, R., and Castaneda, M. (1983). Age-dependent decrease in the activity of proteinsynthesis initiation factors in rat brain. Mech Ageing Dev 21, 183-191. Vellai, T., Takacs-Vellai, K., Zhang, Y., Kovacs, A.L., Orosz, L., and Muller, F. (2003). Genetics: influence of TOR kinase on lifespan in C. elegans. Nature 426, 620. Waskiewicz, A.J., Johnson, J.C., Penn, B., Mahalingam, M., Kimball, S.R., and Cooper, J.A. (1999). Phosphorylation of the cap-binding protein eukaryotic translation initiation factor 4E by protein kinase Mnk1 in vivo. Mol Cell Biol 19, 1871-1880. Webster, G.C., and Webster, S.L. (1983). Decline in synthesis of elongation factor one (EF1) precedes the decreased synthesis of total protein in aging Drosophila melanogaster. Mech Ageing Dev 22, 121-128. Yang, J., Anzo, M., and Cohen, P. (2005). Control of aging and longevity by IGF-I signaling. Exp Gerontol 40, 867-872.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter II

NEUROPEPTIDE BIOSYNTHESIS IN THE NEMATODE CAENORHABDITIS ELEGANS: FROM PRECURSOR TO BIOACTIVE PEPTIDES Steven J. Husson∗ and Liliane Schoofs Functional Genomics and Proteomics, Department of Biology, K.U.Leuven, Naamsestraat 59, B-3000 Leuven, Belgium.

ABSTRACT Endogenous neuropeptides are small signaling molecules that occur in all metazoan species. They function as neurotransmitters or hormones and orchestrate a wide variety of physiological processes by binding G protein-coupled receptors upon initiation of diverse signaling pathways. Over 115 neuropeptide-encoding genes appear to be present in the soil nematode Caenorhabditis elegans. The biosynthesis of these endogenous biologically active peptides involves a series of enzymatic processing steps, starting from a preproprotein. First, the signal peptide is removed and the remaining part of the precursor will be cleft by proprotein convertases at defined motifs displaying basic amino acids. Next, carboxyterminal basic amino acids are removed by the action of a carboxypeptidase. Finally, the carboxyterminal glycine (if present) will be transformed into an amide. A peptidomics platform, which uses state-of-the-art liquid chromatography combined with mass spectrometry, allows us to biochemically identify endogenous peptides present in any tissue or organism. This technology was used to compare the peptide profiles of C. elegans strains having mutations in the presumed peptide precursor processing enzymes. Doing so, we were able to characterize the major processing enzymes KPC-2/EGL-3, CPE/EGL-21 and the chaperone protein 7B2/SBT-1.

∗

Correspondence concerning this article should be addressed to: Steven J. Husson, Tel.: 0032 16324260, Fax: 0032 16323902; [email protected].

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Keywords: Neuropeptide, peptidomics, mass spectrometry, MALDI-TOF MS, proprotein convertase, carboxypeptidase, egl-3, egl-21.

ABBREVIATIONS: CPE, flp, KPC, m/z, MALDI-TOF, MS, MS/MS, nlp, PC, Q-TOF,

carboxypeptidase E; FMRFamide-like peptide; kex2/subtilisin-like proprotein convertase; mass to charge ratio; matrix-assisted laser desorption ionization time-of-flight; mass spectrometry; tandem mass spectrometry; neuropeptide-like protein; proprotein convertase; quadrupole time-of-flight

INTRODUCTION Neuropeptides are important signaling molecules which are contained in inactive preproproteins or peptide precursors that need to be processed in order to yield the bioactive entities (Figure 1). After removal of the signal peptide, the peptide precursor is processed by proprotein convertases (PCs) at defined cleavage places that display basic amino acids like lysines and arginines. In mammals, seven PCs (furin, PC1/3, PC2, PC4, PC5/6, PC7/8/LPC and PACE4) have been described [1-3]. They all show similarity with the prototype member of the family, kex2 from yeast [4]. PC1 (also named PC3) and PC2 specifically recognize substrates that contain pairs of basic residues, reflecting their role in the processing of neuropeptide precursors. Searching the genomic sequence of C. elegans revealed the presence of four genes that encode for kex2/subtilisin-like proprotein convertases (KPC) [510]. As kpc-2 (also named egl-3) is exclusively expressed in the nervous system and appears to modulate mechanosensory responses in C. elegans, it is considered as the major PC needed for the processing of neuropeptides in this nematode [7]. Proprotein convertases are expressed as inactive proproteins that need to be activated. As extensively studied in mice, the neuroendocrine chaperone 7B2 appears to be responsible for the proteolytical activation of proPC2 [11]. Interestingly, the carboxyterminal part of 7B2 regulates the enzymatic activity of mature PC2 [12]. The orthologous 7B2 gene (sbt-1) from C. elegans has already been cloned in 1998 [13], but no biochemical data about the effects of 7B2 on neuropeptide processing in C. elegans was available in pre-peptidomics times. In the next step of biological peptide synthesis, a carboxypeptidase that is encoded by egl-21 in C. elegans will remove the basic residues from the carboxyterminal end of the intermediate peptides. It has been shown that this carboxypeptidase facilitates acetylcholine release at neuromuscular junctions in C. elegans [14]. Finally, a carboxyterminal amidation reaction can occur. This common post-translational modification involves an oxydation of the carboxyterminally

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located glycine residue, followed by a lyase reaction in which the amidated peptide is formed in additon to a glyoxylate molecule.

Figure 1. The neuropeptide processing pathway in C. elegans. All bioactive neuropeptides are derived from inactive preproproteins or peptide precursors that require several highly regulated posttranslational processing reactions to yield the endogenous neuropeptides. A typical peptide precursor contains an aminoterminal signal peptide that is cleaved off upon entry into the secretory pathway by a signal peptidase. Next, a proprotein convertase (EGL-3) cleaves the remaining part of the precursor at specific motifs containing basic amino acids (K, R, KR, RR, RK and KK). The neuroendocrine chaperone protein 7B2 (SBT-1) is needed for the proteolytical activation of pro-proprotein convertases, while the carboxyterminal part of 7B2 is involved in the regulation of the enzymatic activity of the mature proprotein convertase. The resulting intermediate peptides that are formed after cleavage of the neuropeptide precursor all contain carboxyterminal basic amino acids that are removed by a specific carboxypeptidase. Finally, the carboxyterminal glycine residue, if present, is transformed into an amide.

As neuropeptidergic signaling underlies many behaviors in C. elegans, knowledge about the biologically active peptides and their processing enzymes is inevitable for further functional research. Using genomic and biochemical techniques in addition to bioinformatics, a total of 33 flp (FMRFamide-like peptide) genes, 45 nlp (neuropeptide-like protein) genes and 40 insulin-like peptide (INS) genes could be found in C. elegans [15-21]. In the past, biochemical analyses of neuropeptides and their processing enzymes have been hampered by difficulties in purification and characterization of the peptides. As a consequence, only 12 FLP neuropeptides could be biochemically characterized in the pre-peptidomics era [22-27]. Elucidation of peptide sequences required laborious efforts for tissue collection and multiple chromatographic separations in order to isolate and characterize one active signaling molecule. The characterization of their processing enzymes relied on the generation of specific antibodies for a wide variety of neuropeptides and subsequent radioimmuno assays to determine potential substrates. In contrast, the peptidomics technology aims to biochemically visualize and identify endogenous peptides present in a cell, tissue or organism using high-throughput liquid chromatographic techniques and mass spectrometry [28-30]. Using a differential peptidomics approach, in which HPLC fractions are monitored with a matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) instrument, we compared the peptide profiles of various mutant animals, among which mutants of the presumed neuropeptide precursor

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processing enzymes. This way, we were able to characterize the major processing enzymes kpc-2/egl-3 [31], egl-21 [32] and the neuroendocrine chaperone protein 7B2 [33] of the nematode C. elegans. Here, we briefly report on the overall workflow used and discuss the obtained results.

Figure 2. Differential Peptidomics Workflow. Peptide extracts form different C. elegans strains that were made in exactly the same way are separated by reversed phase high performance liquid chromatography (RP-HPLC) generating a chromatogram as indicated. Each fraction from each mutant strain is analyzed by a matrix-assisted laser desorption ionization time-of-flight mass spectrometer (MALDI-TOF MS) of which a schematic representation is shown. All samples are deposited on a steel target plate (TP), together with a UV-absorbing matrix. By firing a focussed laser beam onto the target plate, an ion plume is generated. Individual ions are accelerated by an electric field that is applied on the acceleration plates (Acc) before entering the field-free flight tube of the time-of flight (TOF) analyzer. The main principle of the TOF analyzer is to measure the time an ion needs to reach the detector (DL) at the end of the field-free flight tube. Since the acquired velocity is a function of its mass (m) and charge (z), m/z ratios can easily be determined by measuring the time needed to cross the flight tube. To increase the resolution and mass accuracy of the instrument, a reflectron at the end of the flight tube acts as an electrostatic mirror to reflect the ions that are now detected by a second detector (DR).

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DIFFERENTIAL PEPTIDOMICS: WORKFLOW All C. elegans strains were kindly provided by the Caenorhabditis Genetics Centre (CGC) and were cultured on conventional nematode growth media (NGM) plates containing E. coli OP50 bacterial lawns, at 20°C. Peptide extracts of the wild type C. elegans strain N2 and kpc-2/egl-3, egl-21 and sbt-1 (7B2) mutants were made in exactly the same way, as previously described [15]. Briefly, mixed stage nematodes from twelve to fifteen fully grown Petri dishes, having a diameter 90 mm, were placed in 10 to 20 mL of an ice cold extraction solvent containing methanol/water/acetic acid (90/9/1). Worms were homogenized and the resulting solution was sonicated prior to centrifugation. The pellet was discarded and the methanol was evaporated. The remaining aqueous residue was delipidated by re-extraction with an equal volume of ethyl acetate or n-hexane. Finally, the peptide sample was desalted by solid phase extraction using a C18 cartridge and stored at 4°C. Each peptide extract was subjected to HPLC analyses on a Symmetry C18 column (5 µm, 4.6 x 250 mm, Waters). After injection of the sample, a wash step for ten minutes using 2% CH3CN in 0.1% aqueous TFA was initiated, followed by a linear three-step gradient of 60 minutes at a flow-rate of 1 mL per minute. The gradient was as follows: from 2% CH3CN to 22% CH3CN in 0.1% aqueous TFA in 20 minutes, from 22% CH3CN to 37% CH3CN in 0.1% aqueous TFA in 30 minutes and from 37% CH3CN to 50% CH3CN in 0.1% aqueous TFA in 10 minutes. Obviously, other HPLC columns and gradients can be used. Sixty fractions were automatically collected from the beginning of the three-step gradient. All the generated HPLC fractions were monitored by MALDI-TOF MS in positive ion, reflectron mode. Ion peaks were compared with the theoretical masses of predicted and previously identified neuropeptides from C. elegans to generate peptide profiles of the different strains. This overall differential peptidomics workflow is indicated in figure 2.

RESULTS In our first peptidomics experiments using two dimensional nanoscale liquid chromatography and quadrupole time-of-flight tandem mass spectrometry (2D-nanoLC QTOF MS/MS), we were able to sequence 60 naturally occurring FLP and NLP neuropeptides from C. elegans [15]. Here we report on the use of an off-line approach combining HPLC separation with mass spectrometric detection of peptides using a MALDI-TOF mass spectrometer. This approach allows a robust comparison of neuropeptide profiles from mutant strains. Doing so, analysis of the wild type N2 strain yielded the identification of 91 neuropeptides while only 41 peptides could be monitored in the 7B2 mutant [33]. As an example, mass read-outs of fraction 30 are shown in figure 3 (other fractions not shown). The potential role of the four kpc genes in the processing of endogenous neuropeptides was also assessed by this differential peptidomics approach. Animals mutant in kpc-2/egl-3 were found to be severely depleted in both FLP and NLP neuropeptides [31]. Only a few peptides (or even no peptides) could be monitored in the C. elegans strains carrying different alleles of the kpc-2/egl-3 gene. Our results provide biochemical evidence that KPC-2/EGL-3 is indeed the essential proprotein convertase in C. elegans [31]. In contrast, peptide profiles from kpc-1

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and kpc-4/bli-4 mutants resembled that of wild type animals, while the kpc-3/aex-5 mutant shows slightly decreased peptide content [31]. Finally, 115 carboxyterminally extended intermediate peptides could be found in the egl-21 mutans, in combination with 2D-nanoLC Q-TOF MS/MS [32]. Interestingly, about 20 fully processed neuropeptides could also be measured. These ion peaks, however, all showed a strongly reduced intensity in contrast to the wild type peptides. As an example, fraction 30 (Figure 3C) shows two fully processed peptides, (NGAPQPFVRFamide from the FLP-11 precursor and ASYDYIRFamide from FLP-25) in addition to five peptides that all contain KR residues at their carboxyterminus. These incompletely processed peptides are completely absent in wild type N2 (Figure 3A).

Figure 3. Comparison of MALDI-TOF MS spectra. Mass read-outs from wild type (A), egl-3 (B), egl21 (C) and sbt-1 (D) mutants are shown. Peptide extracts form wild type C. elegans and peptide processing mutants were subjected to a HPLC separation generating 60 fractions which were analyzed by MALDI-TOF MS. As an example, zoom regions from m/z 1000 to m/z 2000 of fraction 30 are shown.

CONCLUSION A peptidomics approach aims to biochemically visualize and identify all endogenous peptides present in a cell, tissue or organism, with a greater resolution and specificity than by traditional techniques like e.g. SDS-PAGE, Western blots and immunocytochemistry [34]. Using liquid chromatography and mass spectrometry, bioactive peptides can now be isolated

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and identified from small tissue extracts [28]. This allows us to construct and compare the neuropeptide profiles of various mutant C. elegans strains. While 41 out of 91 observed peptides from wild type N2 were unequivocally present in the 7B2 mutant, a clear absence of some defined ion peaks could be monitored in the latter strain [33]. Interestingly, highly abundant ion peaks that were present in the wild type strain were totally absent in the 7B2 mutant. This specific and determined absence or presence of a subset of neuropeptides was clearly not observed in strains mutant for kpc-2/egl-3 that only revealed the presence of a few peptides [31]. Our results indicate that KPC-2/EGL-3 is the major proprotein convertase in C. elegans, while the 7B2 chaperone protein is also (indirectly) involved in this processing pathway. Identification of 115 carboxyterminally extended intermediate peptides in the egl21 mutant animals establishes EGL-21 as the major carboxypeptidase for the processing of neuropeptides in C. elegans [32]. As the name implies, both egl-3 and egl-21 were initially identified in a genetic screen for egg laying defective mutants [35]. They also show a mild coiling phenotype and display altered defecation [14,35]. Recently, we found that both of these mutants show a widening of the intestinal lumen, particularly towards the anterior end [32]. Moreover, the egl-3 strain showed reduced intestinal fat accumulation [32]. In conclusion, our differential peptidomics approach provides a biochemical characterization of the major neuropeptide processing enzymes in the nematode C. elegans. Moreover, the altered neuropeptide processing in egl-3, egl-21 and sbt-1 mutant animals is reflected in diverse physiological processes including egg-laying, defecation rhythms, fat storage and locomotion. Our studies therefore establish the basis for identifying specific bioactive peptides that modulate these biological processes.

ACKNOWLEDGEMENTS Financial support from the Research Foundation Flanders (FWO) is greatly acknowledged (FWO-Vlaanderen grants G.0434.07 and 1.5.137.06). The authors want to thank the Interfacultary Centre for Proteomics and Metabolomics “ProMeta”, K.U.Leuven and appreciate the kind gifts of the Caenorhabditis Genetics Centre (CGC). S.J. Husson is a postdoctoral fellow of the Research Foundation Flanders.

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[4] Julius,D., Brake,A., Blair,L., Kunisawa,R., & Thorner,J. (1984) Isolation of the putative structural gene for the lysine-arginine-cleaving endopeptidase required for processing of yeast prepro-alpha-factor. Cell 37, 1075-1089. [5] Gomez-Saladin,E., Wilson,D.L., & Dickerson,I.M. (1994) Isolation and in situ localization of a cDNA encoding a Kex2-like prohormone convertase in the nematode Caenorhabditis elegans. Cell Mol. Neurobiol. 14, 9-25. [6] Gomez-Saladin,E., Luebke,A.E., Wilson,D.L., & Dickerson,I.M. (1997) Isolation of a cDNA encoding a Kex2-like endoprotease with homology to furin from the nematode Caenorhabditis elegans. DNA Cell Biol. 16, 663-669. [7] Kass,J., Jacob,T.C., Kim,P., & Kaplan,J.M. (2001) The EGL-3 proprotein convertase regulates mechanosensory responses of Caenorhabditis elegans. J. Neurosci. 21, 92659272. [8] Thacker,C., Peters,K., Srayko,M., & Rose,A.M. (1995) The bli-4 locus of Caenorhabditis elegans encodes structurally distinct kex2/subtilisin-like endoproteases essential for early development and adult morphology. Genes Dev. 9, 956-971. [9] Thacker,C. & Rose,A.M. (2000) A look at the Caenorhabditis elegans Kex2/Subtilisinlike proprotein convertase family. Bioessays 22, 545-553. [10] The C.elegans Sequencing Consortium (1998) Genome sequence of the nematode C. elegans: a platform for investigating biology. Science 282, 2012-2018. [11] Muller,L., Zhu,X., & Lindberg,I. (1997) Mechanism of the facilitation of PC2 maturation by 7B2: involvement in ProPC2 transport and activation but not folding. J. Cell Biol. 139, 625-638. [12] Van Horssen,A.M., van den Hurk,W.H., Bailyes,E.M., Hutton,J.C., Martens,G.J., & Lindberg,I. (1995) Identification of the region within the neuroendocrine polypeptide 7B2 responsible for the inhibition of prohormone convertase PC2. J. Biol. Chem. 270, 14292-14296. [13] Lindberg,I., Tu,B., Muller,L., & Dickerson,I.M. (1998) Cloning and functional analysis of C. elegans 7B2. DNA Cell Biol. 17, 727-734. [14] Jacob,T.C. & Kaplan,J.M. (2003) The EGL-21 carboxypeptidase E facilitates acetylcholine release at Caenorhabditis elegans neuromuscular junctions. J. Neurosci. 23, 2122-2130. [15] Husson,S.J., Clynen,E., Baggerman,G., De Loof,A., & Schoofs,L. (2005) Discovering neuropeptides in Caenorhabditis elegans by two dimensional liquid chromatography and mass spectrometry. Biochem. Biophys. Res. Commun. 335, 76-86. [16] Husson,S.J., Mertens,I., Janssen,T., Lindemans,M., & Schoofs,L. (2007) Neuropeptidergic signaling in the nematode Caenorhabditis elegans. Prog. Neurobiol. 82, 33-55. [17] Kim,K. & Li,C. (2004) Expression and regulation of an FMRFamide-related neuropeptide gene family in Caenorhabditis elegans. J. Comp. Biol. 475, 540-550. [18] Li,C., Kim,K., & Nelson,L.S. (1999) FMRFamide-related neuropeptide gene family in Caenorhabditis elegans. Brain Res. 848, 26-34. [19] Li,C. (2005) The ever-expanding neuropeptide gene families in the nematode Caenorhabditis elegans. Parasitology 131 Suppl, S109-S127.

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[20] Nathoo,A.N., Moeller,R.A., Westlund,B.A., & Hart,A.C. (2001) Identification of neuropeptide-like protein gene families in Caenorhabditis elegans and other species. Proc. Natl. Acad. Sci. U. S. A 98, 14000-14005. [21] Pierce,S.B., Costa,M., Wisotzkey,R., Devadhar,S., Homburger,S.A., Buchman,A.R., Ferguson,K.C., Heller,J., Platt,D.M., Pasquinelli,A.A., Liu,L.X., Doberstein,S.K., & Ruvkun,G. (2001) Regulation of DAF-2 receptor signaling by human insulin and ins-1, a member of the unusually large and diverse C. elegans insulin gene family. Genes Dev. 15, 672-686. [22] Marks,N.J., Shaw,C., Maule,A.G., Davis,J.P., Halton,D.W., Verhaert,P., Geary,T.G., & Thompson,D.P. (1995) Isolation of AF2 (KHEYLRFamide) from Caenorhabditis elegans: evidence for the presence of more than one FMRFamide-related peptideencoding gene. Biochem. Biophys. Res. Commun. 217, 845-851. [23] Marks,N.J., Maule,A.G., Geary,T.G., Thompson,D.P., Davis,J.P., Halton,D.W., Verhaert,P., & Shaw,C. (1997) APEASPFIRFamide, a novel FMRFamide-related decapeptide from Caenorhabditis elegans: structure and myoactivity. Biochem. Biophys. Res. Commun. 231, 591-595. [24] Marks,N.J., Maule,A.G., Geary,T.G., Thompson,D.P., Li,C., Halton,D.W., & Shaw,C. (1998) KSAYMRFamide (PF3/AF8) is present in the free-living nematode, Caenorhabditis elegans. Biochem. Biophys. Res. Commun. 248, 422-425. [25] Marks,N.J., Maule,A.G., Li,C., Nelson,L.S., Thompson,D.P., Alexander-Bowman,S., Geary,T.G., Halton,D.W., Verhaert,P., & Shaw,C. (1999) Isolation, pharmacology and gene organization of KPSFVRFamide: a neuropeptide from Caenorhabditis elegans. Biochem. Biophys. Res. Commun. 254, 222-230. [26] Marks,N.J., Shaw,C., Halton,D.W., Thompson,D.P., Geary,T.G., Li,C., & Maule,A.G. (2001) Isolation and preliminary biological assessment of AADGAPLIRFamide and SVPGVLRFamide from Caenorhabditis elegans. Biochem. Biophys. Res. Commun. 286, 1170-1176. [27] Rosoff,M.L., Doble,K.E., Price,D.A., & Li,C. (1993) The flp-1 propeptide is processed into multiple, highly similar FMRFamide-like peptides in Caenorhabditis elegans. Peptides 14, 331-338. [28] Baggerman,G., Verleyen,P., Clynen,E., Huybrechts,J., De Loof,A., & Schoofs,L. (2004) Peptidomics. J. Chromatogr. B 803, 3-16. [29] Clynen,E., De Loof,A., & Schoofs,L. (2003) The use of peptidomics in endocrine research. Gen. Comp Endocrinol. 132, 1-9. [30] Husson,S.J., Baggerman,G., Clynen,E., Boonen,K., & Schoofs,L. Peptidomics: the search for endogenous neuropeptides by mass spectrometry is coming of age. In: Farley E.P. Progress in Neuropeptide Research. New York: Nova Science Publishers Inc. 2007, 95-123. [31] Husson,S.J., Clynen,E., Baggerman,G., Janssen,T., & Schoofs,L. (2006) Defective processing of neuropeptide precursors in Caenorhabditis elegans lacking proprotein convertase 2 (KPC-2/EGL-3): mutant analysis by mass spectrometry. J. Neurochem. 98, 1999-2012. [32] Husson,S.J., Janssen,T., Baggerman,G., Bogert,B., Kahn-Kirby,A.H., Ashrafi,K., & Schoofs,L. (2007) Impaired processing of FLP and NLP peptides in carboxypeptidase E

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(EGL-21)-deficient Caenorhabditis elegans as analysed by mass spectrometry. J. Neurochem. 102, 246-260. [33] Husson,S.J. & Schoofs,L. (2007) Altered neuropeptide profile of Caenorhabditis elegans lacking the chaperone protein 7B2 as analyzed by mass spectrometry. FEBS Lett. 581, 4288-4292. [34] Clynen,E., Baggerman,G., Veelaert,D., Cerstiaens,A., Van Der Horst,D., Harthoorn,L., Derua,R., Waelkens,E., De Loof,A., & Schoofs,L. (2001) Peptidomics of the pars intercerebralis-corpus cardiacum complex of the migratory locust, Locusta migratoria. Eur. J. Biochem. 268, 1929-1939. [35] Trent,C., Tsuing,N., & Horvitz,H.R. (1983) Egg-laying defective mutants of the nematode Caenorhabditis elegans. Genetics 104, 619-647.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter III

STEREOCHEMICAL MECHANISM OF TRANSLATION BASED ON INTERSUBUNIT COMPLEMENTARITIES Kozo Nagano Nagano Research Institute of Molecular Biology, 4-8-24 Higiriyama, Kohnan-ku, Yokohama 233-0015 Japan

ABSTRACT A universal rule is found in nucleotide sequence complementarities between the regions 2653-2666 in the GTPase binding site of 23S rRNA and 1064-1077 of 16S rRNA as well as between the region 1103-1107 of 16S rRNA and GUUCG (or GUUCA) of tRNAs. This means that there are two extreme cases of conformational states between the above regions. One is responsible for GTP hydrolysis, and the other plays an important role in the structural transitions, particularly for activation of three tRNAs bound to A, P, and E sites in the proofreading step of codon recognition, and in the process of translocation. In order to understand the mechanism of the conformational change, the present author assumed that four kinds of GTPases, viz. aminoacyltRNA•EF-Tu•GTP ternary complex, EF-G•GDP complex, peptide chain release factor 3 (RF-3), and initiation factor 2 (IF-2), first binds to the cavity region on the 30S ribosomal subunits, where it is known that an antibiotic spectinomycin binds. Amino acid sequence comparison of effector region for the above four GTPases has shown why the first three GTPases unfolds the region around helix 35 of 16S rRNA and goes to the GTPaseassociated region on the 50S subunit, while IF-2 promotes binding of initiator tRNA to the 30S subunit. On the other hand, the crystal structure of the whole ribosome binding three deacylated tRNAs (PDB accession number 1GIX) has shown that the three elbow regions of the tRNAs are distantly separated from each other, resulting in difficulty in explaining the negative cooperativity between A- and E-site tRNAs. Moreover, the existence of a barrier region at the nucleotides G1338 and A1339 of 16S rRNA presents a difficulty in understanding how codon recognition and translocation could occur. The present author explains on the basis of the universal rule of intersubunit

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Kozo Nagano complementarities, 1) how the barrier melts before movements of tRNAs, 2) how the cooperative phenomenon in the proofreading is brought about, and 3) how translocation occurs and results in expelling E-site tRNA from the decoding centre. Such a mechanism was reasonably explained by a series of three-dimensional models of rRNAs and proteins that have already been deposited in the PDB.

INTRODUCTION Biological translation is performed by the following four processes: (1) initiation, (2) elongation, (3) polypeptide chain termination, and (4) recycling. From a viewpoint of evolution of life, the second process could have been developed first, while the others could have been spontaneous at first, and then developed later on by modifications of the first essential process. The second one consists of codon recognition and translocation, the former which requires aminoacyl-tRNA (aa-tRNA) and peptidyl-tRNA (pep-tRNA), and the latter preceeded by transpeptidation. Bacterial ribosomes require two protein factors for the elongation cycle: elongation factor Tu (EF-Tu) for accommodating a cognate aa-tRNA with the A site, and elongation factor G (EF-G) for translocation. The site near A2660 in 23S rRNA is called α–sarcin/ricin stem-loop (SRL), which is essential for binding the two GTPases, and for GTP hydrolysis. The present author has found the nucleotide sequence complementarities as a universal rule for all living kingdoms between the SRL and 5’-side of helix 35 of 16S rRNA (abbreviated as h35 hereafter) and between the other side of h35 and the GTΨC-sequence of almost all tRNAs (Nagano and Nagano, 2007). This finding has led to an explanation of how the driving force of translocation and codon recognition can be produced. Although the 3D structure of ribosome containing three deacylated tRNAs at A, P, and E sites (Yusupov et al., 2001) showed the distance of 100A°≈120A° between the above two regions of rRNAs, inspection of the structure around the region of 16S rRNA binding proteins, S2, S3 and S5, suggested a course of transformation that allows the ribosome to perform translocation and binding of an aa-tRNA•EF-Tu•GTP ternary complex according to the universal rule of the intersubunit rRNA complementarities. The purpose of the present paper is to show a series of reasonable predicted structures of the ribosome and tRNAs around the decoding site and the tRNA binding sites, and to explain how translocation occurs, how stop signals can be recognized, and also how discrimination of cognate aa-tRNA from noncognate and near-cognate ones as a proofreading mechanism can be made from the viewpoint of 3D structures.

RESULTS AND DISCUSSIONS 1. Nucleotide Sequence Complementarities and Possibility of Such Interactions in the 3D Structure The secondary structures of E. coli 16S rRNA, and 5’-half and 3’-half of 23S rRNA are shown in Figures 1a, b and c by following the layout style of Gutell (1993). A large part of

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these secondary structures are the same as the ones presented before (Brimacombe, 1995). E. coli sequence is a representative of eubacteria and its helix numbering is used as a standard for all kinds of living species. The regions of particular concern in the present paper are near h35 and h37, α-sarcin/ricin binding loop (SRL) in helix 95 of 23S rRNA (denoted as H95 hereafter), and L11 binding region in H43 and H44. The T-loop of tRNA is also added in Figure 1a so that it may implicate a possible interaction with one side of h35. Figure 2 shows that the region of nucleotides, 1064-1077 of 16S rRNA, is complementary with the region of SRL in the GTPase-binding site on 23S rRNA. The 3’ side of h35, CGAGC, is also complementary to the GTΨCG sequence on the T-loop of all tRNAs. This is a universal rule that holds for all living organisms (Nagano and Nagano, 2007). This rule implies that there are two extreme cases of conformational states for the nucleotides drawn in red and blue in Figure 2. The left-most structure seems to play an important role in GTP hydrolysis. Another structure is a long helix, which is shown on the right-hand side in Figure 2. This seems to be a transition-state structure, that could be observed neither by X-ray crystallography nor by cryo-electron microscopy. In order to understand the functional role of the structural change, it was assumed that both aa-tRNA•EF-Tu•GTP ternary complex and EF-G•GDP complex first interact with an exposed location on the small subunit. The nucleotide exchange from GDP to GTP could occur on EF-G shortly before GTP hydrolysis (Zavialov et al., 2005). The locations of both complexes to be touched with the small subunit are protruded portions such as T-loop-D-loop contact region of aa-tRNA of the ternary complex and domain IV-V bridge region of EF-G•GDP complex. The anticodon of aa-tRNA can find its A-site codon very easily. Near the binding site of the 3’ end adenine of aa-tRNA on EF-Tu, there exists a sequence of aminoacid residues that is called the effector region. Figure 3 shows a comparison of aminoacid sequences of the effector region of 7GTPases, EF-Tu, eukaryotic EF-1 (eEF-1), EF-G, eEF-2, peptide chain release factor 3 (RF-3), initiation factor 2 (IF-2), and archeal initiation factor 2 (aIF-2). The right-most residue of EF-Tu, Asn64, is known as the binding site of the 3’ end adenine of aa-tRNA. Thr62 is known to bind to two oxygen atoms of Pγ phosphate of GTP (Berchtold et al., 1993). Either tyrosine or phenylalanine at the 47th position of EF-Tu is known to bind to guanine base of GTP. Although KAR59 is a part of short α helix in the crystal structure of EF-Tu with a uncleavable GTP derivative (Berchtold et al., 1993), it is easy for us to imagine a binding mode to some phosphate groups of rRNA, probably a part of double-stranded RNA helix. It is also important to note that the acidic residues drawn in red in Figure 3, such as Asp51 and Glu56 of EF-Tu, are highly conserved in 5 GTPases, EF-Tu, eEF-1, EF-G, eEF-2, and RF-3, but not conserved in IF-2 and aIF-2. It is known that IF-2 promotes binding of fMet-tRNA to the 30S ribosomal subunit. Accordingly, the pair of acidic aminoacid residues could probably play a role in unfolding a specific part of double-stranded rRNA helix. When h35 and h37 are unfolded, the CGAGC nucleotides on the 3’ side of h35 would be taken by the T-loop of aa-tRNA, and its D-loop moves toward the T-loop of P-site tRNA, which is confirmed by the finding that the fluorescence signal of proflavin at D16 or D17 in the D-loop has shown a small initial increase even in the case of noncognate tRNA recognition (Rodnina et al., 1994). On the other hand, the 5’ side of h35 can go to the location of the SRL of 23S rRNA, resulting in formation of the long helix, as shown in Figure 2.

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a

Figure 1a.

Kozo Nagano

Stereochemical Mechanism of Translation...

b

Figure 1b.

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28 C

Figure 1. Secondary structure of E. coli rRNAs following the layout style of Gutell. (a) 16S rRNA along with TΨC-stem-loop of yeast Phe-tRNAPhe. 5’, C, 3’M, and 3’m represent 5’-domain for the region of 1-556, central domain for the region of 557-918, 3’-major domain for the region of 919-1396, and 3’-minor domain for the region of 1397-1542, respectively. Small numerals denote the residue numbers of the nucleotides indicated by the respective lines. The helices are numbered by large numerals as Brimacombe (1995). It is predicted that the T-loop of tRNA interacts with the 3’ side of h35. (b) 5’ half of 23S rRNA. I, II, and III represent domain numbers for the regions of 1-561, 5621269, and 1270-1646, respectively. A-site finger is the nucleotide range from 879 to 898. (c) 3’ half of 23S rRNA. IV, V, and VI represent domain numbers for the regions of 1647-2014, 2015-2627, and 2628-2904, respectively. The α–sarcin/ricin stem-loop region (SRL), P-loop, and A-loop are the nucleotide ranges from 2653 to 2667, from 2251 to 2253, and from 2552 to 2556, respectively. Numerals are as in (a).

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Figure 2. Schematic diagram showing a possibility of interactions between the α-sarcin/ricin loop (SRL) of 23S rRNA and one side of another highly conserved region (mostly helix 35) of 16S rRNA, using E. coli sequence as a representative example, as well as between the other side of h35 and GTΨC-region (T-loop) of tRNA. This figure is obtained from Figure 1 of Nagano and Nagano (2007). The sequence of SRL is shown mostly in red. UAGUACGAG (in red), AG (in gray) and GAC (in red). 5’ and 3’ represent that the adjacent nucleotides are 5’- and 3’-ends, respectively, of the nucleotide fragment. H95 designates the helix number in E. coli 23S rRNA secondary structure. The letters, h35, h36, and h37, represent the helix numbers of the nearby helices in E. coli 16S rRNA secondary structure. SpcF denotes spectinomycin footprint site. The nucleotides 1064-1077, displayed in light blue and gray is hypothesized to make base pairs with the red and gray region 2653-2666 of 23S rRNA, while the nucleotides in green are proposed to interact with the sequence GUUCG or GUUCA (in purple) in tRNA. The sequence UUA in brown is also highly conserved. The nucleotide positions of tRNA sequence are shown in a standard form using two-letter codes. The numbers 1, 18, 33, 37, 56, 61, and 76 are the nucleotide residue numbers in yeast tRNAPhe for the residues indicated by the connecting lines. N at position 33 represents that the residue is pyrimidine (viz. U or C), while R at position 37 is purine (viz. A or G or their modified bases). The nucleotide at position o5 or 61 is in most cases C and highly conserved. In exceptional cases, when the nucleotide at position n5 is A, the nucleotide at o5 could be U. When d4-e4 base pair is neither Watson-Crick nor wobble type, G·A pair occurs most frequently as a favorable noncanonical base pair and A··A and U·U pairs appear less frequently.

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Figure 3. Comparison of amino acid sequences from the GTP binding region to the effector region of the GTPases, EF-Tu, eEF-1α, EF-G, eEF-2, RF-3, IF-2, and aIF-2. Red letters, blue letters, and green letters denote the highly conserved acidic amino acid residues such as aspartic acid and glutamic acid, the highly conserved basic residues such as arginine and lysine, and the other highly conserved residues particularly around the GTP binding and effector regions of EF-Tu, respectively, while the other amino acid residues are represented in black. The abbreviated species name, the species name, the accession file name, and their residue number range of the aminoacid sequence are obtained from Swiss Protein Sequence Data Bank (PIR): (1) eubacterial elongation factor EF-Tu; Ecoli, Escherichia coli EFECT 1765. Taqua, Thermus aquaticus S00229 17-66. Hinfl, Haemophilus influenzae E64078 17-65. Pfalc, Plasmodium falciparum plastid S72277 17-65. clEgrat, Euglena gracilis chloroplast EFEGT 17-66. clPpurp, Porphyra purpurea chloroplast S73208 17-65. mtCeleg, Caenorhabditis elegans mitochondria T37273 50-98. mtScere, Saccharomyces cerevisiae mitochondria EFBYT 53-101. (2) eukaryotic elongation factor eEF-1α; Scere, S. cerevisiae EFBY1A 12-75. Tbruc, Trypanosoma brucei A54760 1275. Pfalc, P. falciparum S21909 12-75. Zmays, Zea mays S66339 12-75. Tpyri, Tetrahymena pyriformis A49171 13-76. Hsapi, Homo sapiens EFHU1 12-75. (3) eubacterial elongation factor EF-G; Ecoli, E. coli EFECG 15-65. S6803, Synechocystis sp. PCC 6803 S75863 15-65. Taqua, T. aquaticus

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EFTWG 17-67. Dradi, Deinococcus radiodurans E75536 18-68. clGmax, Glycine max chloroplast S35701 105-155. mtScere, S. cerevisiae mitochondria S43748 46-96. mtRnorv, Rattus norvegicus mitochondria S40780 52-105. (4) eukaryotic elongation factor eEF-2; Scere, S. cerevisiae A41778 2472. Bvulg, Beta vulgaris T14579 24-72. Athal, Arabidopsis thaliana A96602 27-75. Celeg, C. elegans A40411 24-72. Dmela, Drosophila melanogaster S05988 24-72. Hsapi, H. sapiens EFHU2 24-72. (5) eubacterial peptide chain release factor RF-3; Ecoli, E. coli E91295 18-72. Sente, Salmonella enterica AD1072 18-72. Paeru, Pseudomonas aeruginosa B83159 16-70. B.APS, Buchnera sp. APS C84993 1872. Linno, Listeria innocua AB1556 17-71. Llact, Lactococcus lactis E86668 15-69. Spneu, Streptococcus pneumoniae D97921 15-69. N7120, Nostoc sp. PCC 7120 AC2353 21-75. Saure, Staphylococcus aureus D89870 15-69. Atume, Agrobacterium tumefaciens AC2615 17-71. Bmeli, Brucella melitensis AB3539 15-69. Ccres, Caulobacter crescentus B87382 17-71. (6) eubacterial initiation factor IF-2; Ecoli, E. coli FIEC2 396-463. Bsubt, Bacillus subtilis A35269 224-291. Saure, S. aureus H89900 214-281. clPpurp, P. purpurea chloroplast S73178 268-335. mtScere, S. cerevisiae mitochondria S66706 150-219. mtSpombe, Schizosaccharomyces pombe mitochondria T39351 176245. (7) archeal initiation factor aIF-2; Mther, Methanobacterium thermoautotropicum E69132 10-95. Afulg, Archaeoglobus fulgidus H69345 18-100. Hsali, Halobacterium salinarum T43849 20-103.

2. Transition-state Conformation of three tRNA Molecules in Codon Recognition A crystal structure of the whole ribosome has been presented at 5.5 Å resolution for Thermus thermophilus 70S ribosome binding three deacylated tRNAs at A-, P- and E-sites and mRNA of A- and P-site codons (Yusupov et al., 2001), the atomic coordinates of which can be obtained by two PDB data files, 1GIX and 1GIY. If we assume that the locations of Patoms of the middle nucleotide of P-site codon and A422 of 23S rRNA at the foot of H22 are fixed, the distance between the two P-atoms is calculated as 138.95 Å, as shown in Figure 4. Figure 4 also shows those from the three 3’ end adenines of A-, P-, and E-tRNAs to the middle P-nucleotide, 74.47, 76.30, and 91.56 Å, respectively. In contrast to these values, those from the A-site 3’ end and P-site 3’ end to the middle P-nucleotide for A-P tRNA docking pair (Nagano and Nagano, 1997) in the PDB data file 1IP8 are 74.07 and 85.83 Å, respectively (not shown). In the next predicted model of Nagano and Nagano (1997) for entrance of T-site tRNA (in the recognition mode of A-site) to the two tRNAs in the posttranslocational state, PDB data file 1IPM, those from the T-site 3’ end, P-site 3’ end, and E-site 3’ end to the same position on mRNA are 72.29, 81.94, and 77.26 Å, respectively (not shown). If we concentrate our attention on a transition-state conformation of the three tRNA molecules in codon recognition, such a model can be presented as PDB data file 1IPU, and the corresponding values, 48.08, 79.31, and 75.60 Å, respectively, as shown in Figure 4. This should be an energetically elevated or activated substate between the two states of postribosome and preribosome. Since it is energetically elevated than the other two states, an important question has to be asked beforehand as to what is the source of energy for such an elevation of potential chemical energy. One of the basic hypotheses in the present paper is that this is also the result of the long helix formation, as shown in Figure 2. It has been shown by X-ray crystallography that the relative orientation of small and large ribosomal subunits is open in postribosome and closed in preribosome (Ogle and Ramakrishnan, 2005). We have already explained how it occurs in translation (Nagano and Nagano, 2007). The long helix formation brings about a counterclockwise rotation to the small subunit with respect to the

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large subunit viewing from the mRNA. On the other hand, the conformational change of large-subunit rRNA (LSU-rRNA) binding the 3’ end adenines of A- and P-site tRNAs would occur through a clockwise rotation of H95, that is accompanied by rotational movements about the locations of H94, H96, and H97. Since the P-P distance between C2394 at the foot of H88 and A422 at the foot of H22 of 23S rRNA in 1GIY is 62.02 Å, summation of 62.02 and 75.60, 137.62 Å, becomes almost equal to the P-P distance between the middle of P-site codon and A422 in 1GIY, 138.95 Å, as shown in Figure 4. This means that the C2394 bound to the 3’ end adenine of E-tRNA could move toward the P-site codon by rotation of about 32˚around the pivotal position at A422. Considering that the peptidyl end of P-tRNA is approximately fixed in postribosome, the 3’ end nucleotide of the E-tRNA would have moved by the effect of binding the aa-tRNA•EF-Tu•GTP ternary complex to the 30S ribosomal subunit, similarly to the amount of the movement of C2394 (and G2421) in translocation prior to the movement of 30S subunit by the effect of EF-G•GDP attachment to the small subunit for formation of the hybrid P/E state (Nagano and Nagano, 2007). This could be an important preparatory step for the movements of various associated regions for codon recognition. The putative first interaction site of the elongation factors could be assigned as the location where h34, h35, and h36 are folded back toward the neck of 30S subunit along the cleft side of its head, that is exactly the spectinomycin binding site, as shown in Figure 4c of Carter et al. (2000). It is known that spectinomycin inhibits EF-G dependent translocation (Bollen et al., 1968; Sigmund et al., 1984). Figure 4b of Carter et al. (2000) also showed how G1064, C1066 and G1068 are associated with the binding of spectinomycin. The shape of the cleft of 30S subunit fits very well for that of either T-loop-D-loop contact region of aa-tRNA of the EF-Tu•GTP ternary complex or the domain IV-V bridge region of EF-G•GDP. In the X-ray structures for various forms of ribosomes (Yusupov et al., 2001; Schuwirth et al., 2005; Korostelev et al., 2006; Selmer et al., 2006), a barrier between P- and E-tRNAs is observed as a ridge containing G1338 and A1339 and the 790 loop of 16S rRNA. It would also be important to note that G942 and U1341 form always a G-U wobble pair, indicationg a possibility of a functionally meaningful conformational change. The interatomic P-P distance between G1064 and U1341 is found to be 22.5 Å, which is an ideal distance for formation of a G-U wobble pair. Inspection of the P-P atomic distances in the neighbourhood of A1339 has shown that the important part of the barrier could be removed as a result of formation of a transient short double-stranded helix, GUCA1067/GAAU1341. This step will be referred to as barrier melting mechanism hereafter. Two anticodon stems of both P- and E-tRNAs are rotated, twisted and induce two kinds of conformational changes at T-loop-D-loop (elbow) regions and anticodon loops. Then, the 5’ end of the 5’ side of h35, G1068, remains at the location of U1341, while its 3’ end nucleotides, UGUUG1077, go closer to the SRL of 23S rRNA, resulting in the first contact between G2655 of 23S rRNA and U1075 of 16S rRNA, that was suggested by the result of mutant ribosome experiment with modification of G2655 of 23S rRNA (Leonov et al., 2003; Nagano and Nagano, 2007).

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Figure 4. Schematic representation of P-P atomic distances for two kinds of tRNA binding models on the 70S ribosome. The three tRNAs on the left-hand side are those of the X-ray structure of the whole ribosome binding three deacylated tRNAs (Yusupov et al., 2001; PDB file 1GIX and 1GIY) and their elbow regions are separated from each other, in which the P-P distances from C56 of E-tRNA to G18 of P-tRNA and from C56 of P-tRNA to G18 of A-tRNA are 47.43 Å and 34.04 Å, respectively, while those on the right-hand side are predicted as a transition-state conformation of three tRNAs in the proofreading mechanism and deposited in PDB with an accession number 1IPU, in which the P-P distances from C56 of E-tRNA to G18 of P-tRNA and from C56 of P-tRNA to G18 of A-tRNA are 21.11 Å and 20.57 Å, respectively. The P-P distances between the middle nucleotide of the P-site codon and the other nucleotide positions are as follows: 1) for 1GIX, A422 of 23S rRNA (1GIY), 138.95 Å; A76 of E-tRNA, 91.56 Å; A76 of P-tRNA, 76.30 Å; A76 of A-tRNA, 74.47 Å; 2) for 1IPU, A76 of EtRNA, 75.60 Å; A76 of P-tRNA, 79.31 Å; A76 of A-tRNA, 48.08 Å. The P-P distance between A422 of 23S rRNA and C2394 of 23S rRNA is 62.02 Å for 1GIY. If we assume that the nucleotide A422 and H14 of 23S rRNA in domain I (see Figure 1b) are fixed, the location of A76 of E-tRNA in 1IPU can be obtained by rotating the helical rod (H88, H22 and a pseudoknot helix CCA415/UGG2410) clockwise by about 32◦ around an axis passing through the P atom of A422. A part of barrier region, G1338 and A1339, exists between two anticodon stems of E- and P-tRNAs. GTP hydrolysis occurs near the A76 of A-tRNA (or T-tRNA) in the transition-state structure, 1IPU, in the proofreading mechanism.

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b

c

d

Figure 5. Stereo pictures showing the transition-state conformation of three tRNAs in the proofreading mechanism. The models are viewed from the small subunit to the large subunit, but rotated 30◦ around X-axis to see the nascent polypeptide chain, cysteinyl residue of aa-tRNA, and GTP. The models of

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tRNAs on the right-hand side, the middle one, and the one on the left-hand side are T-tRNA in red, PtRNA in green, and E-tRNA in violet, respectively. (a) Wire model representations of P atoms (which is called P-atom model hereafter) of three whole tRNA molecules without large variable loops are shown along with Cα atoms of the nascent polypeptide chain as α-helical conformation in greenish yellow, Cα atoms of EF-Tu with green small dots, Cα atom of cystenyl residue of aa-tRNA with large red ball, and P atoms of GTP with large golden balls. mRNA is drawn in dark blue. 16S rRNA from G1064 to G1077 are drawn in grey, and 23S rRNA from U2653 to C2666 are in black. (b) P-atom models from D17 to C60 of E-tRNA, from D17 to C60 of P-tRNA, and from D17 to the end nucleotide of the large variable loop (V16) of A-tRNA. mRNA is as in (a). The possibility of base-pairing from G18 of PtRNA to C56 of E-tRNA as well as from G18 of A-tRNA to C56 of P-tRNA is shown. E-site codon (m3, m-2, m-1), P-site codon (m 1, m 2, m 3), and A-site codon (m 4, m 5, m 6) are shown. The locations of three U33 positions of the anticodon loops are also shown. (c) P-atom models of h44 and part of h28 of 16S rRNA in grey and a helical rod through H88 and H22 connected with each other by CCA415/UGG2410 of 23S rRNA along with some of their adjacent nucleotides in black, both atomic coordinates are taken from those of 1GIX and 1GIY, are fitted to the cavity of the three codonanticodon base pairs, shown in (a) and (b). The location of C2394 of 23S rRNA is fitted to that of the 3’ end (A76) of E-tRNA. See text for the spatial relationship of h44 and H88 with mRNA and the three tRNA models. The conformations of the three tRNAs and mRNA are as in (a) and (b). (d) P-atom model of the joint region of h44 and part of h28 of the model in (c). In order to obtain an ideal basepairing capability with A1503-U33(E), A1398-U33(P), and A1396-U33(T), a G1504-C1399 base pair (see Figure 1a) should be broken. See text for further explanation. The locations of G920 as starting nucleotide of h28, A1492 and A1493, that interact with A-site codon-anticodon base pairs as A-minor motifs (Nissen et al., 2001), as well as A1396, A1398 and A1503 are shown by numerals.

It can be pointed out that the two elbow regions of the two tRNAs come close together to make a G-C base pair between G18 of P-tRNA and C56 of E-tRNA, that allows an exposed C56 of P-tRNA towards an incoming aa-tRNA. In the X-ray structure of the whole ribosome binding three deacylated tRNAs, the 3 tRNA molecules are all isolated and compactly folded with the elbow regions. Accordingly, the P-P distances between C56(E) and G18(P) and between C56(P) and G18(A) are 47.4 Å and 34.0 Å, respectively. In the present predicted model of transition-state conformation of three tRNA molecules in codon recognition (PDB data file 1IPU), those between C56(E) and G18(P) and between C56(P) and G18(A) are 21.1 Å and 20.6 Å, respectively. Both are capable of forming a stable single G-C base pair. Figure 5a shows such a model of three tRNAs as well as some of their associated regions displayed by a computer program RasMol V2.7.3 (Sayle and Milner-White, 1995; Bernstein, 2000). Here mRNA, E-tRNA and P-tRNA are drawn in dark blue, violet, and green, respectively. The T-tRNA is also drawn in red. We can see α–helical form of nascent polypeptide chain in brownish yellow, and SRL region of 23S rRNA in black. The grey strands at the lower part on the right-hand side of Figure 5a are the 14 nucleotides from C1064 to G1077 around the 5’ side of h35 and the 5 nucleotides of CGAGC1107 of 16S rRNA. The space between h35 of 16S rRNA and the SRL as well as A-tRNA is occupied by a molecule of EF-Tu, Cα atoms of which are shown by small dots in light green. We can see the triphosphate group of GTP (three P atoms) and the cysteinyl residue (Cα atom) at the 3’ end of aa-tRNA at the T site. In this model, the codon-anticodon base pairings are all made of cognate tRNAs. This state just corresponds to that when GTP hydrolysis is about to occur. GTP hydrolysis does not occur before reaching this state because of a slight distortion of GTP binding region on the long helix. It should also be important to note that the three tRNA molecules drawn in Figure 5b contain the largest variable loop of tRNA3Ser from E. coli (Yamada and Ishikura, 1973) as a

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hybrid molecule with tRNAPhe from yeast (Nagano and Nagano, 1997), but that it is omitted for the three tRNAs in Figure 5a for simplicity of the model. In Figure 5b the base-pairing possibilities of G18(A)-C56(P) and G18(P)-C56(E) can be seen.

3. Discrimination of Cognate, Noncognate and Near-cognate tRNAs and Proofreading Another conformational change accompanied by the barrier melting mentioned above occurs at the anticodon loops of A-, P-, and E-tRNAs. Such a conformation is shown in Figure 5b, in which mRNA and only three anticodon stem-loop of A-tRNA, P-tRNA, and EtRNA are drawn in dark blue, red, green, and violet, respectively. Here, we can see that three anticodon loops are all sharply pointed at U33. The codon-anticodons are nearly vertical and closely resemble an ideal A-form RNA helix structure. Accordingly, the base pairings of codon-anticodons are more stable than those observed by X-ray crystallography, but the shape of anticodon loops around U33 looks rather unstable unless it is supported by some conserved bases, such as adenines of 16S rRNA. It is well known that the nucleotides at the positions 32 and 33 are usually pyrimidines, and that U33 is highly conserved except for C33 of fMet-tRNA (Sprinzl et al., 1998). This implies a possibility of conformational change in the 7-nucleotide anticodon loop by exposing U33. It loses one hydrogen bond formed between N3 atom of the base and O5’ atom of A36 of the loop, and gains two hydrogen bonds if the uracil base could find an unpaired and conserved adenyl base in its neighbourhood. Such an adenine can be found as a highly conserved one at A1396. Accordingly, if a strong right-handed twist movement is given between the first and second anticodons by a cognate codon-anticodon base pairs, such a flip-flop motion of the loop could easily occur. This is exactly an induced fit that is expected to enhance the accuracy in translation. Since it is known that C1400 can be crosslinked to the first anticodon base (mG34 in ASLPhe) (Prince et al., 1982), it is possible for another highly conserved nucleotide, A1398, to base-pair with U33 of P-site tRNA. Both C1399 and C1397 are also highly conserved but are most weakly base-paired to the U33s of tRNAs at the A and P sites. In order to play a similar role for a deacylated tRNA at the E site in the poststate, A1503 of 16S rRNA could be the likeliest candidate. Although the P-P distance of U33(E)-A1503(16S) is 34.19 Å in the PDB data file 1GIX (Yusupov et al., 2001) (U33(A)-A1396(16S); 26.49Å, and U33(P)-A1398(16S); 23.48Å), the conformational twist of the transition-state three tRNA molecules, discussed in the preceding section, would make it much closer under a condition that both U33 nucleotides of P- and E-tRNAs are fully exposed. In this connection, it would be informative for us to know that the P-P distances of A1398(16S)-A1503(16S), A1396(16S)-A1503(16S), and A1398(16S)-A1396(16S) in 1GIX are only 16.69 Å, 15.71 Å, and 8.13 Å, respectively. Figure 5c shows that the cavity of three codon-anticodon helices surrounded by three exposed U33 nucleotides of the anticodon loops of T-, P-, and E-tRNAs is fitted to the joint region of h28 and h44 of 16S rRNA, viz. the region of nucleotides, 1393-1410 and 14901506, and that the 3’ end A76 of E-tRNA is taken by C2394 of 23S rRNA, as observed crystallographically by Schmeing et al. (2003) and Yusupov et al. (2001). The atomic

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coordinates of h28 and h44 of 16S rRNA and H88 and H22 of 23S rRNA are obtained from the PDB files 1GIX and 1GIY (Yusupov et al., 2001), but fitted to the coordinate system of 1IPU, in which the three tRNA molecules are viewed down along its Z-axis, as shown by Figures 6, 8, 11, and 12. However, the orientations of both helices of 16S and 23S rRNAs are modified by keeping only the distance between A1503 of 16S rRNA and C2394 of 23S rRNA. The surface of the nucleotides, 1393-1410 and 1490-1506, are fitted to the mRNA, and then, H88 and H22 along with their neighbouring nucleotides are rotated around the Zaxis, Y-axis, and X-axis, by -110.0◦, -11.0◦, and -4.1◦, respectively, with the A1503 fixed at the origin. Since this operation of the relative orientational change of the two helices h44 and H88 makes C2422 instead of C2394 closer to the 3’ end of E-tRNA, the rotation of 150.0◦ around the Y-axis, -38.8◦ around the X-axis, and -58.8◦ around the Z-axis with the C2394 fixed as the origin of the coordinate system. This implies that a considerable degree of rotational freedom could exist for both large helices of the small and large ribosomal subunits. Figure 5c also shows that a rotation of the helical rod of H88, H22, and a pseudo knot CCA415/UGG2410 around an axis passing through A422 and A423 would allow the C2394 to approach the 3’ end of P-tRNA very easily. After the E-tRNA leaves the large subunit E site near C2394, it would move in such a direction particularly when P-tRNA is deacylated in the ribosomal prestate, even if the conformation of the mRNA and the ASLs of A- and P-tRNAs still remain at the previous positions. This state could be exactly what we call the P/E and A/P hybrid sites, if both P-loop could also shift simultaneously toward the CCA end of A-tRNA by expelling A-loop. It can be achieved by binding of EF-G•GTP complex at the GTPase associated centre. Explanations for such a movement of LSU-rRNA will be given for Figure 8c in section 4. Specific interactions of A2433 and A2434 at the base of H74 with 2’-hydroxyl groups of the residues 71 and 76 of deacylated P-tRNA (Feinberg and Joseph, 2001) could play a decisive role in shifting the P-loop toward the old A-tRNA for formation of the hybrid sites, although the effect of C1892 of H68, as observed by Feinberg and Joseph (2001), cannot be explained by this model. On the other hand, this model explains very well the negative cooperativity between the T and E sites (Nierhaus, 1990), because EtRNA resists the entrance of T-tRNA into the A site by its steric hindrance, and because the T-tRNA, once it gets into the A site, expels the E-tRNA with exchange of two G-C pairs, as described by tRNA docking pair model of Nagano and Nagano (1997). In order to stabilize the transition-state three tRNA binding structure, three A-U pairs with the exposed U33 of E, P-, and T-tRNAs would be required under the auspices of the region of the nucleotides, 1393-1410 and 1490-1506. Figure 5d shows more detailed locations of A1503, A1398, and A1396 of 16S rRNA, assuming that the conformation of the region are the same as that of Yusupov et al. (2001) and just fitted to the cavity of mRNA and three ASLs shown in Figure 5b. It is clear that G1504-C1399 pair should be broken in order to obtain ideal base-pairings with A1503-U33(E-t), A1398-U33(P-t), and A1396-U33(T-t). This seems to be a kind of cooperative phenomenon that produce 6 hydrogen bonds of 3 A-U pairs at the expence of 3 hydrogen bonds of G1504-C1399 and 3 hydrogen bonds between N3 atom of U33 and O5’ atom of A36 of the three anticodon loops. This site is very close to h29, where the trgger of the barrier melting occurs as a result of h35 unfolding. A-minor motif interactions (Nissen et al., 2001) of A1492 and A1493 to the A-site codon-anticodon helix could be much more facilitated without help of paromomycin (Ogle et al., 2001).

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4. Predicted Models of two tRNAs and their Surrounding Regions in the Preribosome (1IP8) and Postribosomes (1IPM) Four types of experimental results have shown that conformational differences around the elbow region of tRNAs bound to A and P sites as well as to P and E sites; 1) G18 and G19 in the D-loop of the tRNA specifically bound to the A site are protected from kethoxal modification in contrast with those in the free state as well as those in the tRNA bound to the P site (Bertram et al., 1983). 2) Footprinting studies revealed striking differences in the Tand D-loops of tRNAs bound to the P and A sites (Jørgensen et al., 1985). They observed a more open structure for the tRNA in the A site. 3) G18 of A-site tRNA and C56 of P-site tRNA was found to crosslink to protein L27 (Abdurashidova et al., 1990). 4) Cleavage patterns of thioated tRNAs by iodine (I2) in the P site of preribosome and in the E site of postribosome showed that their T-loop (G53-C61) are strongly protected in contrast with the unprotected nucleotides in the D-loop (G15-A21) (Nierhaus et al., 1995). Furthermore, the patterns of A- and P-site tRNAs in the preribosome are very similar to those of P- and E-site tRNAs in the postribosome, respectively (Nierhaus et al., 2000). They proposed an α–ε domain carrying the two tRNAs from the A and P sites to the P and E sites (Spahn and Nierhaus, 1998). On the basis of the above findings, the present author partly followed the idea of Nierhaus and his coworkers, because their conveyor domain in the large subunit could not be assigned confidently, but presented different models of two tRNAs, A-P docking pair and P-E docking pair, in which the angle between the ASLs of A- and P-site tRNAs (or Pand E-site tRNAs) is 50° (Nagano and Nagano, 1997) under a requirement that the conformation of P-site ASL is common for both pre- and postribosome. Crystal structures of L11•rRNA complexes have been presented with and without thiostrepton binding N-terminal region of the protein, respectively (Conn et al., 1999; Wimberly et al., 1999). The above regions from G1051 to U1108 of 23S rRNA is called GTPase associated centre, while the SRL region is called GTPase binding site. The crystallographic analysis of 50S subunit of bacterial ribosome at 5 Å resolution (Ban et al., 1999) identified the locations of the L11•rRNA complex and the SRL on their electron density map. SRL was almost fully exposed on the surface of the 50S subunit, and its conformation was almost the same as that obtained by NMR study (Szewczak et al., 1993). The tentative model of EF-G was presented so that its location should be consistent with the EF-G binding site of SRL but should not contradict the assigned locations of H96 and H97 (Ban et al., 1999), resulting in dissatisfaction of crosslinking distance between EFG•GDPCH2P complex and the region of nucleotides 1055-1081 of 23S rRNA (Sköld, 1983). It was shown by crosslinking experiments that the regions of nucleotides 877-913 of H38 (Asite finger) of 23S rRNA are found near the A-site tRNA (Rinke-Appel et al., 1995). The nucleotide 885 at the top of H38 of the A-site finger is close to the position 20:1 of A-site tRNA, while 2475 to the position 47 of A-site tRNA (Rinke-Appel et al., 1995). (The position 20:1 of lupin tRNAmMet (Rinke-Appel et al., 1995) is an extra nucleotide residue between G20 and A21.) Such a model is obtained in the present work, as shown in Figure 6a, in which the structure of GTP is fitted to the G domain of EF-G, and also makes a close contact with A2660 that can be depurinated by ricin to inactivate the ribosome (Marchant and Hartley, 1995). (The atomic coordinates for Figure 6a are presented in the PDB file 1IP8.)

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This orientation of EF-G was determined so that its domain III would make a contact with the region of nucleotides 1055-1081 of 23S rRNA, as observed by crosslinking study (Sköld, 1983), and that the tip of domain IV would be directed toward the region around C1920 in H69, as suggested by directed hydroxyl radical probing (Wilson and Noller, 1998), although it was observed to occupy the location of A-site codon-anticodon helix after completion of translocation in the cryo-EM study of yeast 80S•eEF2•sordarin complex (Spahn et al., 2004). Figure 6a also shows the conformations of two tRNAs, mRNA, and helices h34, h35, h36, h37, h38 and h39 in the prestate. This substate is obtained after the putative first interaction of EF-G•GDP complex at the cavity of 16S rRNA, that is the binding site of spectinomycin which inhibits EF-G-dependent translocation, as already discussed in the section.2. Before reaching this substate, the barrier region of G1338 and A1339 as well as AU1341 interacted with the region of GUCA1067, rotated around the axis of the four base-paired helix, and unfolded again. This is also shortly before the long helix formation, that could induce a large conformational change in the central loop region of the 50S subunit, resulting in the reversible transition from the ‘relaxed state’ of preribosome, in which the 3’ end of A-site tRNA is peptidylated, the 3’ end of P-site tRNA is deacylated and does not lock the translocation to the A/P and P/E hybrid state (Valle et al., 2003; Zavialov et al., 2005), as described by Nagano and Nagano (2007). The structure of EF-G in Figure 6a is the same as the crystal structure of apo-EF-G (or nucleotide-free EF-G) (Ævarsson et al, 1994), that was found to be very similar to that of EF-G•GDP complex (Czworkowski et al., 1994). GTP model was fitted to the EF-G following the corresponding structure of EF-Tu•GDPNP (Berchtold et al., 1993). It was found that a pep-tRNA in the P site regulates the GTPase activity of EF-G and RF-3 in a similar manner by decreasing the binding affinity of the respective factors in the GTP form to the ribosome complex for both translocation and peptide release (Zavialov and Ehrenberg, 2003). A guanine nucleotide exchange was observed for both termination (Zavialov et al., 2001) and translocation (Zavialov et al., 2005). As described in the section.2, the region around h35 could unfold when the GTPases first interact with the cavity region, where it is known as the spectinomycin binding site. In such an instant, a certain region on the structures of EF-G and RF-3 could also be influenced so that they would lose the GDP molecule. Figure 3 suggests that Trp50 and Met51 in EF-G and Trp58 and Met59 in RF-3 on the amino acid sequences of the effector region of both GTPases are typical and could be responsible for instability of GDP binding. Then, it would be quite natural to think that the binding of GTP into the vacant G domain of both GTPases could occur when they collide with either SRL or L11 binding region of the GTPase associated centre of 23S rRNA. Figure 3b of Berchtold et al. (1993) suggests that hydroxo group of Tyr47 of EF-Tu is bound to an oxygen atom of Pα phosphate of GTP. Since the residues at the position of EF-Tu is either Tyr or Phe, as shown in Figure 3, the stacking effect of the aromatic ring could be the cause of the stability of GDP in EF-Tu. Along with the effect of either Thr60 in EF-G or Ser68 in RF-3 for binding to two oxygen atoms of Pγ phosphate of GTP, the conformational change at the instant of the collision to the SRL and GTPase associated centre would facilitate the GTP molecule to get into the G domain. It is, however, important to note that the recovery of GTP does not immediately result in its hydrolysis. Some kind of distortion must be effected to inactivate the G domain for a while. It

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could be due to the formation of the long helix around A2660 of SRL near the GTP binding site of the GTPases.

a

b

Figure 6. Stereo pictures showing the predicted 3D structures of A-tRNA and the regions around the GTPase associated centre. The main chains (P-O5’-C5’-C4’-C3’-O3’-) of RNAs and (N-C-C-O-) of proteins are drawn in colours by WebLabViewer Pro 3.0, and the labels and numerals are added by Adobe Illustrator 7.0J. (a) A- and P-tRNAs in the A-P tRNA docking pair model (Nagano and Nagano, 1997) are shown in magenta and green, respectively. The mark 76A represents the location of the position 76 of A-tRNA. The other numbers ending with A correspond to the residue numbers in yeast tRNAPhe at the A site, while those ending with P denote them at the P site. mRNA from the first nucleotide of P-site codon, which is denoted by m 1, to 6th nucleotide count from m 1, which is the third nucleotide of A-site codon and denoted by m 6, is shown in greenish yellow. The regions of nucleotides of 16S rRNA, 1069-1075 and 1103-1107, capable of base-pairing with SRL and GTΨCG sequence of almost all tRNAs, respectively, are drawn in light blue and grey, respectively, while the

Stereochemical Mechanism of Translation...

41

region 1090-1092 capable of binding stop signal is in pink. The other nucleotides from 1055 to 1202 are shown in blue. It can be seen that the interaction between SRL and the nucleotides 1069-1075 is possible. A purple letter U indicates the location of nucleotide U1075 of 16S rRNA, while a purple letter G shows that of nucleotide G2655 in the SRL region of 23S rRNA. This figure represents a substate of two tRNAs and their surrounding regions along with EF-G•GTP complex shortly after the first interaction between the complex and h35, and also after the melting of the barrier, G1338 and A1339. The distance between the two letter positions, G and U, is 58.7A°, but could be much closer, considering that the secondary structure around h35 is totally unfolded. See text for explanation about the interaction between U1075 and G2655. Large characters of A and P indicate the locations of the anticodon stem-loop of tRNAs at the A and P sites, respectively. (b). The main chains of the regions of nucleotides surrounding the 3’-end of P-tRNA in (a). P-tRNA in green, and the domains IV and V of EF-G in bluish olive green, and the domain III of EF-G in pink are the same as in (a), while H89 is in light blue. The nucleotides from 2246 to 2258 of H80 are drawn in red, while those from 2547 to 2561 of H92 are in magenta. The nucleotides from 2057 to 2085, from 2234 to 2240, from 2434 to 2454, and from 2607 to 2611 of 23S rRNA are drawn in brown, those from 2497 to 2546, and from 2562 to 2585 (including H90 and H91) are in blue, and those from 2586 to 2606 (H93) are in dark blue, and those from 1350 to 1378 (h43) are in grey. The nucleotides from 2299 to 2317 (H84) are in olive green. The nucleotides from 1860 to 1882 (H68) are in dark brown. The nascent polypeptide in α-helical form is shown in violet and denoted by a small letter p at its N-terminus. Helix numbers and the other symbols are as in (a).

The location of two highly conserved G18 and G19, near the position 20:1 in A-tRNA is facing toward the T-loop of P-tRNA. This is because the position 20:1 is accessible to h23 of 16S rRNA (Rinke-Appel et al., 1995). Their variable loops are replaced by the largest ones observed so far and situated remote from the two codons on the mRNA (Nagano and Nagano, 1997). The location of A1196 in h34 is closest to m8 and m9 positions of mRNA to represent a crosslinking distance between the two nucleotides (Sergiev et al., 1997). The closeness between h37 and h40 meets the requirement from a crosslink between the nucleotides 10901094 and 1161-1164 (Mueller and Brimacombe, 1997), while the location of U1126 in h39 is to satisfy a crosslink between the nucleotides 1125-1127 and 1280-1281 in h41 (not shown here but contained in 1IP8) (Mueller and Brimacombe, 1997). One of the purposes of Figure 6a is to show a possibility of SRL in red interacting with its complementary nucleotides 1069-1075 in light blue before the formation of the long helix, when domain G of EF-G enters into the interface of the two subunits and collides with the SRL. The nucleotide U1075 of 16S rRNA is shown by a purple letter U in Figure 6a, in which G2655 of 23S rRNA is also shown by another purple letter G. It was found that a G2655C mutation in the SRL region in E. coli sequence showed a lethal effect on cell growth (Macbeth and Wool, 1999). A more detailed analysis about the mutation of G2655 on EF-G-dependent translocation has shown the following effects: G2655A showed a little difference from the wild type, G2655C was the most defective, and G2655U was intermediate (Leonov et al., 2003). These results as well as no effect on EF-G-independent spontaneous translocation were reasonably explained by the base-pairing capabilities, as described above. Figure 6a can explain that the G2655 is the most probable target for triggering the EF-G-dependent translocation by an attack of U1075 of 16S rRNA under a condition that the h36 and a part of h35 could be considerably fragile. The distance between G2655 of 23S rRNA and U1075 of 16S rRNA in the X-ray structure of ribosome is 126.4A° and too far to allow the hypothetical base-pairing mechanism, while it is 58.7 A° in the present model. The shortest distance between the two regions is 47.6 A° for A2657 of 23S rRNA and U1073 of 16S rRNA, that would be close enough to make a

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Kozo Nagano

distance of about 20 A° for a base pair in the A-form helix upon conformational fluctuation. The distance could be much smaller when the region around h35 is totally unfolded shortly after the barrier melting, as discussed before. In the case of EF-G-independent spontaneous translocation, such a specific mutational effect was not detected (Leonov et al., 2003). It would probably mean that the collision between h35 and SRL is more randomly fluctuated without EF-G•GTP complex. A2309 at the loophead of H84 of 23S rRNA crosslinks strongly to the position 20:1 of A-site tRNA in the prestate and also to the same position of P-site tRNA (Rinke-Appel, 1995) probably in the pre- and poststates, as shown in Figure 6b. (The atomic coordinates for Figure 6b are the same as those for Figure 6a, viz. the PDB file 1IP8.) Since the same nucleotide can crosslink to the position 47 and the nucleotide C2306 to the position 8 of P-site tRNA (Rinke-Appel, 1995), this must be situated on the left-hand side to the position 20:1 of AtRNA, but closer to the nucleotide G693 of 16S rRNA. The location of the loophead of H84 in the crystal structure of 70S ribosome (see Figure 6E of Yusupov et al., 2001) does not seem to satisfy such a crosslinking condition. U2586 at the feet of both H90 and H93 also crosslinks to the same position of A-site tRNA, while U2584 to the aminoacyl residue (Rinke-Appel, 1995). On the other hand, A2602 is a footprint site of both A- and P-site tRNAs (Moazed and Noller, 1989). One more strong crosslinking nucleotide C885 at the loophead of H38 to the same position of A-site tRNA must be situated nearer to G693 of 16S rRNA. This location of H38 is not shown in Figures 6a and b, but in Figure 8a. H74 and H75 are positioned behind P-tRNA, because both helices seem indispensable for all living species but H76, H77, H78, and H79 are missing in animalia and trypanosome mitochondrial LSUrRNA (see Figures 10B and C of Raué et al., 1988). Such an arrangement of the abovementioned nucleotides are shown in Figure 6b. A stereochemically sound connection between H74 and H80 through the foot of H75 is made in order to achieve a G-C pair between G2252 in the P-loop of 23S rRNA and C74 of P-site tRNA (Samaha et al., 1995). Another direct base pair to C75 of A-site tRNA from G2553 of 23S rRNA (which is called Aloop) was found (Kim and Green, 1999) and was confirmed by use of a puromycin derivative in the difference fourier analysis of the 50S ribosome (Nissen et al., 2000). The puromycin crosslinked to G2553 could still form a peptide bond (Green et al., 1998). Figure 6b shows such a placement of helices, H74 and H80 as well as H90, H91 and H92. Under this condition the most reasonable arrangements of H73, H93 and H94 were explored. In order to connect the H73 with the SRL in H95, H73 must occupy a considerably large space below both tRNAs at the A and P sites, reserving rather small space for connecting with H90 and H93. The nascent polypeptide chain in α–helical form is also shown in Figure 6b, although it might not be α–helical. The locations of H89 and the domains IV and V of EF-G are quite the same as those in Figure 6a. Besides, the location of h43 of 16S rRNA in the prestate is also shown, at the foot of which the nucleotide C1378 crosslinks to position 32 of A-site tRNA (Rinke-Appel et al., 1995). The nucleotides U1376 and C1378 are observed to crosslink to the same position of E-site tRNA instead of P-site tRNA in the poststate (Rinke-Appel et al., 1995). (The distances discussed above are listed in Table 1.) This means that a considerably drastic conformational change would be involved in the 3D structure of 16S rRNA during translocation. A694 at the loophead of h23 is a strong crosslinking nucleotide for the 20:1 position of both A- and P-site tRNA (Mueller et al., 1997). When we restrict the crosslinking

Stereochemical Mechanism of Translation...

43

condition to the prestate, the A694 can approach to the position 20:1 of A-tRNA from the left-hand side. It can also crosslink to the same position of the P-tRNA after translocation. The adjacent nucleotide G693 is also a strong crosslinking candidate for the position 32 of both P- and E-tRNAs. This is the main reason why h23 must be situated on the left-hand side in both pre- and poststates, as shown in Figures 7a and b. (The atomic coordinates of the models in Figures 7a and b are given by 1IP8 for prestate and 1IPM for poststate, respectively). The location of A694 in Figure 7b explains well the crosslinking to the mC32 of E-tRNA but does not seem to be good for that to the A20:1 position of P-tRNA (Mueller et al., 1997). If it is situated at the same position as in Figure 7a, such a crosslinking condition would be satisfied. The location of the loophead of h23 in the crystal structure of the 70S ribosome binding three deacylated tRNAs is found near E-tRNA (see Figure 8A of Yusupov et al. (2001)), which seems to be the structure in the poststate.

a

b

Figure 7. Stereo pictures showing the locations of the nucleotides A694, U1376, and C1378 of 16S rRNA and some of the other nucleotides. The models are viewed as in Figure 5. (a) P-atom model of Aand P-tRNAs in the preribosome. A694 is situated near A20:1 which is adjacent to G19 of A-tRNA. (b) P-atom of P- and E-tRNAs in the postribosome. This model shows that G693 occupis a favourable location for explaining the crosslinking possibility of mC32 of E-tRNA to G693, but is not suitable for explaing a crosslinking of A694 to A20:1 in poststate. If the location of h23 is almost the same as in (a), A694 could be crosslinked to A20:1 of P-tRNA.

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Table 1. Comparison of the distances of interest among two different ribosome structures, X-ray structure of three tRNA binding 70S ribosome (1GIX, 1GIY), and the predicted models in the present work (1IP8, 1IPM, 1IPU) distance 1GIX-1GIY G2655(23S)-U1075(16S) U2656(23S)-G1074(16S) A2657(23S)-U1073(16S) C2658(23S)-G1072(16S) G2659(23S)-C1071(16S) A2660(23S)-U1070(16S) G2661(23S)-C1069(16S) A2662(23S)-U1068(16S) A1196(16S)-m61 A1196(16S)-m8 A1196(16S)-m9 U or A1197(16S)-m6 U1052(16S)-m6 G1053(16S)-m6 C1054(16S)-m6 C1054(16S)-Ala163(S3) A1055(16S)-m6 U1090(16S)-G1164(16S) U1091(16S)-G1164(16S) U1126(16S)-U1281(16S) G1127(16S)-A1280(16S) Pro 92(L11)-A 896(23S) Pro 92(L11)-C2474(23S) Thr 19(L11)-A2660(23S) C 885(23S)-G20(A-t) C 885(23S)-A21(A-t) C2475(23S)-U47(A-t) U1065(23S)-Gly502(EF-G) U1065(23S)-Gly503(EF-G) G1074(16S)-Asp393(EF-G) G1920(23S)-Gly520(EF-G) G1920(23S)-Gly520(EF-G) D16(A-t)-U47(P-t) A694(16S)-G20(A-t) A694(16S)-A21(A-t) A694(16S)-G20(P-t) A694(16S)-A21(P-t) G693(16S)-omC32(P-t) G693(16S)-omC32(E-t)

(Å) 126.40 125.46 123.71 117.40 109.45 102.20 105.56 106.66 18.91

20.72 27.88 24.05 21.88 29.98 23.64 19.62 19.54 8.97 16.64 26.69 22.95 33.64 20.90 22.70 29.98

11.90 69.52 69.07 50.96 48.82 35.07 26.43

distance 1IP8

distance 1IPM

distance 1IPU

(Å) 58.68 49.53 47.64 49.28 55.51 60.51 62.58 65.31 20.59 11.90 17.82 17.65 32.17 34.02 33.68

(Å) 18.53 20.05 19.49 18.96 18.46 18.29 20.66 23.54 37.24 30.81 29.02 39.95 27.32 30.73 35.78

(Å) 91.49 86.28 90.99 86.46 80.33 75.69 76.15 77.33 30.45 22.15 24.93 27.45

29.26 9.62 13.15 23.67 13.98 59.62 31.54 39.06 22.57 20.23 39.88

32.98 66.19 71.03 74.81 71.56

38.85 9.62 13.15

68.32 9.45 9.72 72.67 51.92 51.51

77.64

16.71 29.14 29.36 35.52 17.60 12.40 55.50 54.15 52.80

34.00

65.92 64.73 53.66 22.23

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45

Table 1. (Continued) distance 1GIX-1GIY C1378(16S)-omC32(A-t) C1378(16S)-omC32(E-t) U1376(16S)-omC32(E-t) A2309(23S)-G20(A-t) A2309(23S)-A21(A-t) A2309(23S)-G20(P-t) A2309(23S)-A21(P-t) A2309(23S)-G47(P-t) C2306(23S)-U 8(P-t) U2586(23S)-G20(A-t) U2586(23S)-A21(A-t) U2584(23S)-aa(A-t)2 A2602(23S)-A73(A-t) A2602(23S)-A76(A-t) A2602(23S)-U47(P-t) A2602(23S)-A76(P-t) A2451(23S)-aa(A-t) A2062(23S)-aa(A-t) G2252(23S)-C74(P-t) G2553(23S)-C75(A-t) C421(23S)-A76(A-t) C421(23S)-A76(P-t) C421(23S)-A76(E-t) C421(23S)-C2394(23S) C421(23S)-m2 G2394(23S)-A76(A-t) G2394(23S)-A76(P-t) G2394(23S)-A76(E-t) G2394(23S)-m2 1

(Å) 54.72 19.63 18.82 34.67 38.84 19.92 20.49 26.01 44.14 71.71 73.06 10.95 15.14 14.57 48.34 7.81 8.88 20.37 15.46 17.48 102.14 91.20 57.39 62.02 126.40 59.25 53.06 16.32 91.72

distance 1IP8 (Å) 13.52

distance 1IPM

distance 1IPU

(Å)

(Å)

6.83 11.36 15.19 12.96 43.19 42.38 46.81 51.50 9.03 13.96 31.68 32.48 33.36 12.57 35.77 29.47 13.08 17.55 16.39 74.42 60.17 35.72 118.12 58.18 48.04 112.58

10.87 15.55 38.42 23.23

67.64

66.45 46.62 48.70 32.70 18.50

89.93 64.14 36.39 132.64 61.29 63.39 118.72

42.06 38.77 17.35 12.55 51.13 36.49 54.53 55.32 81.03 38.67 51.18 54.33 17.53 107.62 51.18 18.50 79.95 128.57 91.11 64.14 55.86 138.38 120.57 63.28 46.56 110.00

m1 m9 represent P-atom positions of respective nucleotides on mRNA upstream count from m1 as the first P-site codon base-pairing with the 36th nucleotide of P-tRNA. m6 denote the third A-site codon base-pairing with the 34th nucleotide of A-tRNA. 2 aa denotes the aminoacyl-residue of the ternary complex of EF-Tu.GTP. The atomic coordinate of aminoacyl-residue of the ternary complex of EF-Tu.GTP is represented by that of O3* atom of A-tRNA in the crystal structure 1GIX. For the predicted models, 1IP8, 1IPM, and 1IPU, aa is represented by the P-atom coordinates of A76(A-t), A76(P-t), and A76(E-t), respectively. (A-t) denotes A-tRNA in the crystal structure, 1GIX, and A-tRNA in the predicted models, 1IP8 and 1IPU. (A-tRNA in 1IPU is the same as T-tRNA.) (E-t) denotes E-tRNA in the predicted models, 1IPM and 1IPU, while (P-t) represents three different P-tRNA models in 1IP8, 1IPM, and 1IPU.

Kozo Nagano

46

a

b

c

Figure 8. Stereo pictures showing the connectivity between H89 and H95. (a) The main chains of 23S rRNA in the preribosome corresponding to Figures 6a and b. H95 in red and H89 in light blue are the same as those in Figure 6. The region of nucleotides 862 to 915 of H38, the tip of which 881-895 is Asite finger, is drawn in orange. H92 and H93 as in Figure 6b. The nucleotides from 2246 to 2258 (H80) are in pink. The region from 2043 to 2057 and from 2611 to 2625 (H73) is in green, the region from 2626 to 2645 and from 2770 to 2788 (H94) in violet, the region from 2675 to 2731 (H96) in olive green, and the region from 2732 to 2769 (H97) in blue. The other regions are from 2023 to 2042 (H72), from 2058 to 2079 (H74 and H75), from 2241 to 2245 (H75), from 2434 to 2454 (H75), from 2497 to 2585, from 2607 to 2610 are in grey. The locations of four crosslinking nucleotides C885, A2309, and U2586 to the position 20:1 of A-tRNA and A2451 to aminoacyl residue, as well as two guanines G2252 base-pairing to C74 of P-tRNA and G2553 to C75 of A-tRNA are denoted by numerals. (b) The main chains of 23S rRNA in the postribosome. The regions drawn in this figure are almost the same as in (a) except the A-site finger region of H38. This structure represents the changes in the positions of the regions after the long helix formation, in which recognition of either stop signal or sense codon is about to occur. The colours and other symbols are as in (a). The location of U2586 is shown by a numeral, although it is not favourable for crosslinking to any particular nucleotides. (c) The main chains of the regions as in (b) in the midway substate during translocation. This structure represents the substate in which the tRNA docking pair (not shown) is rotated by 25° from the prestate around the symmetry axis of the ASLs of A-tRNA in the prestate and E-tRNA in the poststate (angle difference between A- and P-tRNAs and between P- and E-tRNAs is 50°). The colours and other symbols are as in (a). It is noticeable that H83 and H84 are behind the decoding region so that they do not obstruct the tRNA movement.

Before GTP hydrolysis of the EF-G•GTP complex, the domains IV and V of EF-G in the orientation, as shown in Figure 6a, would not only push A-tRNA toward P site, but also touch somewhere of 16S rRNA that could influence the h43 for inducing the necessary conformational change as a device for triggering in translocation. This might be associated with the movement of a part of the barrier, G1338 and A1339, as discussed in the section.2. Another device for triggering seems to be the orientation of H89 of 23S rRNA, which is locked by H38 in the prestate, as shown in Figure 8a. (The atomic coordinates for Figure 8a are the same as those for Figure 6a, viz. the PDB file 1IP8.) In Figure 8a, H89 is located on

Stereochemical Mechanism of Translation...

47

the right-hand side of H38. The reason for this is that C2475 at the tip of H89 is found to crosslink to the position 47 of A-site tRNA, while C885 at the tip of H38 (A-site finger) to the position 20:1 of A-site tRNA (Rinke-Appel et al., 1995). It was also shown by the method of directed hydroxyl radical probing that the 92nd residue of L11 is close to U896 at the stem of H38 (A-site finger) and U2474 at the tip of H89, while that its 19th residue is near to A2660 at the loophead of SRL (Holmberg and Noller, 1999). As to the placement of H89, G2455 must be situated near the 3’-end of A-tRNA, because both A2451 and A2062 at the foot of H74 crosslink to the aminoacyl residue attached to the 3’-end of A-site tRNA (RinkeAppel, 1995). Moreover, it was found that A2451 plays an important role in forming the peptide bond being synthesized (Nissen et al., 2000). (The distances between the residues discussed above are listed in Table 1.) After the binding of EFG•GTP complex the H89 is pushed down by the domains IV and V of EF-G, giving rise to opening the block of H93 against spontaneous translocation. H89 moves below the new P-tRNA (which was at the A site in Figure 6a), as shown in Figure 8b, although the P-tRNA is not shown. (The atomic coordinates for Figure 8b are given in 1IPM.) When translocation starts, H93 would move along with the old P-tRNA in the direction of E site, and work as a stopper of the new P-tRNA. Therefore, the orientation of EF-G in Figure 6a seems to be consistent with the well-known functions of the ribosome in translation. In Figure 8b, it is assumed that SRL is interacting with the region 1069-1107 of 16S rRNA in preparing for the interaction with the GTΨCG sequence of a new aa-tRNA by the region 1103-1107 of 16S rRNA. H38 would not appear because A-tRNA does not exist in the poststate. The location of H38 would be the same as that observed in the crystal structure of 70S ribosome (Yusupov et al., 2001). On the other hand, an intermediate substate model between the pre- and the poststates was explored as shown in Figure 8c, in which the location of a docking pair tRNA model (Nagano and Nagano, 1997) was rotated by 25˚ around the symmetry axis between the two tRNA pairs in the two stable elongation states. (The atomic coordinates for Figure 8c are given in the PDB file 1IPL.) The locations of H83 and H84 are quite different from the corresponding locations in Figures 8a and b, because base pairs of the pseudo knots, CCA415/UGG2410, AUG2330/CAU2387, and G2282/C2427 (Gutell, 1993) are kept, although their locations are also changed drastically. The SRL interacting with the region 1064-1107 of 16S rRNA gives a rotational twist about the relative locations of H94, H96 and H97 resulting in passing H73 and H94 across the base pair between the A-loop and the position 75 of A-tRNA and putting off the brake (H38) against the spontaneous translocation of the tRNA docking pair, while the interaction of the region 1103-1107 of 16S rRNA with the GTΨCG of A-tRNA produces a rotational twist around the symmetry axis of the two elongation states. This movement makes H93 collide with H80, disrupts the base pair between the P-loop and the position 74 of P-tRNA, and makes it possible to form new base pairs between the two consecutive guanines 2251 and 2252 and the two consecutive cytidines 74 and 75 of the old A-site tRNA at the P site. This seems to be the essential point of the mechanism of translocation.

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Kozo Nagano

5. Estimation of a Degree of Correctness in the Predicted Models Presented Above Moazed and Noller (1989) first presented by chemical probing the evidence of the nucleotide C2394 of 23S rRNA being in contact with E-site tRNA. Direct contact of this nucleotide with the A76 of the E-site tRNA was confirmed by X-ray analysis (Schmeing et al., 2003). It was also observed in the X-ray results of the whole ribosome (Yusupov et al., 2001). It was a little before the paper of Yusupov et al. (2001) appeared when the present author had deposited the models of 1IP8, 1IPU, and so on, to the Protein Data Bank without seriously considering the importance of the direct contact of C2394 with A76 of E-site tRNA. Figure 6a presents a model of two tRNA molecules bound to the A and P sites, in which the nascent polypeptide is just bridging between the two 3’-oxy groups of the A76 nucleotides in transpeptidation. After finishing the transpeptidation, the deacylated 3’-end nucleotide of PtRNA is open for accepting C2394 at the foot of H88 in preparing for the hybrid P/E site on the side of large subunit. Figures 9a and b shows the locations of the C2394 in the models of 1IP8 and 1IPU, just behind and below the A76 at the P and E sites, respectively. These models were built following the knowledge of secondary structure of 23S rRNA (Gutell, 1993; Brimacombe, 1995), the reasonability of experimental facts and the stereochemical consistency. As a result, the distances between the locations of A76 at the P site in Figure 9a (1IP8) as well as at the E site in Figure 9b (1IPU) and C2394 are quite reasonable considering the flexibility of the regions around the nucleotides 2388-2393 and 2422-2426 of 23S rRNA and the rather arbitrary positioning of the 3’-end of H22. The distance between the location of A421 of 23S rRNA in Figure 5c and that in Figures 9a and b is 26.0 A°. The 3’-end CCA conformation of the deacylated P-tRNA in Figure 9a and E-tRNA in Figure 9b can be directed toward the position of C2394. Figure 9a also shows that the spatial relationship between H88 and H22 could be rather flexible, unless the position of C2394 is fixed. This seems to show that a degree of correctness in the present prediction is rather satisfactory. When the long helix is made immediately after the binding of EF-G•GTP, the binding of C2394 to the A76 at the P site would affect the orientations of helices H74 and H80, and the changes of the locations of large subunit helices, described concerning to Figure 8b, would precede the rotation of the decoding site on the small subunit. Although the present author did not think about such a time lag at that time, the formation of the intermediate ‘twisted‘ hybrid state of P/E and A/P sites, that is still reversible (Moazed and Noller, 1989; Zavialov et al. 2005) before completion of translocation, can be explained from this model building work rather without difficulty. Kajiro (1978) first insisted that the binding of EF-G•GTP to ribosomes triggers for translocation, because EF-G complexed with an uncleavable GTP derivative promoted one cycle of translocation. The results of cryo-EM study of Spahn et al. (2004) on yeast 80S eEF2•sordarin complex supported the above view. In their structure, the tip of domain IV of eEF2 occupied the location of ASL of the A-site tRNA after completion of translocation despite no GTP hydrolysis. The main body of the domain IV was found near h34 and h33, while its G domain made a contact with the SRL. Experimental technique such as cryo-EM was not able to detect the structure of the whole ribosome shortly before translocation started, as suggested by the predicted model of the preribosome in Figure 6a. At that time the present

Stereochemical Mechanism of Translation...

49

author thought that the conformational change of EF-G between the GTP and GDP forms could be enough for pushing the A-site ASL toward the P site. In contrast, the nucleotides 1103-1107 could interact with the T-loop of A-site tRNA for pushing it toward P site in the case of EF-G independent spontaneous translocation. The potential energy of the preribosome binding eEF2•GTP (or EF-G•GTP for prokaryote) complex would be considerably high compared to that of the postribosome with the factor•GDP complex. Spahn et al. (2004), however, did not mention the conformational change between the two forms of the eEF2. GTP hydrolysis would be required for releasing the eEF2•GDP (EF-G•GDP) and restoring the open form of the small subunit (Ogle and Ramakrishnan, 2005). a

b

Figure 9. Stereo pictures showing a degree of correctness of the present predicted models. The models are viewed as in Figure 5. It is worthy to note that the C2394 in both pre- and poststates is at the reachable range from the 3’ end nucleotide A76 of both E- and P-tRNAs, even though the models were built without knowing the tight binding as a large subunit E site (Yusupov et al., 2001; Schmeing et al., 2003). (a) P-atom model of A- and P-tRNAs in the prestate and the H88 and H22 bound together by a pseudoknot helix CCA415/UGG2410. (b) P-atom model of T-, P-, and E-tRNAs in the transition-state in the proofreading mechanism and the H88 and H22 bound together.

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50

Before closing the foregoing sections on the problems of translocation, it must be clarified that the E site of the P-E tRNA docking pair model is consistent with the E site of Nierhaus et al. (2000) and the E’ site of Paulsen and Wintermyer (1986) in the sense that three nucleotides of the anticodon of the deacylated tRNA bound to this site are base-paired with the E-site codon on the mRNA, while the anticodons of the E2 site of the former and the E site of the latter are separated from the E-site codon. Nierhaus and his coworkers insist that such a situation of the three base pairings of E-site codon with the anticodon of the tRNA bound to only one defined E site can be achieved in poststate under the conditions of polyamine buffer and in native polyribosomes (Nierhaus et al., 2000).

a

b

Figure 10. Stereo pictures showing the all-atom representation of the conformation of the decoding site when a stop-signal recognizing tripeptide, Pro-Ala-Thr, is bound to the stop codon, UAA. This model is viewed as in Figure 5. The tripeptide is drawn in red, mRNA in light blue, and the anticodon region from A31 to A36 of P-tRNA in green. Some of the residue numbers are shown by numerals. (a) Part of the long helix is formed between the SRL nucleotides, G2655-A2662, of 23S rRNA in magenta and those of G1068-U1075 of 16S rRNA in grey. The other related regions of 16S rRNA are also shown in grey. UUA indicates the P-atom position of U1090 of 16S rRNA. (b) A more detailed model of the tripeptide and the UAA/UUA helix at the loophead of h37 of 16S rRNA, which is drawn in dark blue.

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6. Recognition of Stop Signal after Translocation Shortly after the step of translocation, described above, the next codon for a coming new aa-tRNA is checked whether it is one of the three stop signals, UAA, UAG, and UGA or not. Three kinds of class-1 release factors are known to recognize stop signals as follows: RF1 has the conserved tripeptide motif Pro-Ala-Thr for recognition of UAG and UAA, RF2 in prokaryotes has the motif Ser-Pro-Phe for UGA and UAA, and a single eukaryotic one eRF1 has the tetrapeptide motif Asn-Ile-Lys-Ser for UAG, UGA, and UAA (Kisselev and Buckingham, 2000). It was insisted that a tripeptide Pro-Ala-Thr in prokaryotic RF1 can recognize the stop signals specifically (Ito et al., 2000). Furthermore, the above three motifs of class-1 RFs have been believed to recognize stop signals directly as if they are anticodons, and are recently confirmed for RF2 by cryo-EM experiments (Rawat et al., 2003; Klaholz et al., 2003), and for eRF1 by analogy from its tRNA-like shape of the crystal structure (Song et al., 2000). However, a big question still remains unsolved as to how the first base of stop signals, U, is recognized (Kisselev and Buckingham, 2000). Here, it is explained by use of base pairing with the UUA sequence at the loop of h37, as a transient precursor before replacement with the stop-signal recognizing tripeptides. Such a model of tripeptide Pro-AlaThr binding to UAA/UUA helix is found stereochemically reasonable and shown in Figure 10a. (Its atomic coordinates for Figure 10a are given in the PDB file 1IPN.) A more detailed picture around the tripeptide Pro-Ala-Thr of the model is presented in Figure 10b. (Its coordinates are the same as those for Figure 10a.) If the codon is UGG, the UGG/UUA base pairs would not be stable enough because of two consecutive wobble pairs. If the stop signal is UAA, any one of RF1, RF2, and eukaryotic eRF1 will work for termination. If the stop signal is either UGA or UAG, the stability of the stop-codon/UUA helix would be a little weaker. Based on the effects of mutations of a conserved C1054 in h34 of small-subunit rRNA (SSU-rRNA) (C1054A, C1054U, and C1054G caused a suppression of UGA, UGA and UAG, and UAA and UAG, respectively), and the other two conserved nucleotides at the GTPase associated centre in H43 and H44 of LSU-rRNA (A1067 deletion, A1067C, and A1067U as well as G1093A caused a UGA-specific suppression), interactions of the rRNAs either with mRNA or with a stop-codon-RF complex have been proposed (Arkov and Murgola, 1999). On the other hand, a direct interaction between stop signals and RF2 has been proposed on the basis of crosslinking data suggesting that the fourth base of the stop signal interacts RF2 as well as the decoding site of SSU-rRNA, the nucleotides 1385-1420 in h28 and h44 (Brown and Tate, 1994). The two results of the above different approaches can be confirmed by the recent result of cryo-EM structures (Rawat et al., 2003; Klaholz et al., 2003). Particularly important is the result of mutations suggesting that the regions of both SSU- and LSU-rRNAs considerably far from the stop signal itself (Arkov and Murgola, 1999) actually affects the stability of the stop-codon-RF2 complex and that UGA makes it most unstable in contrast to the most stable complex of UAA. The other result of mutations on stop codon UGA and UAG readthrough in E. coli (O’Connor and Dahlberg, 1995) suggests that the change in the nucleotides only in contact with the body of RFs (not directly with the stop codon) could affect the stability of the stop-codon-RF complex. Ivanov et al. (2001) insisted that the hairpin loops of H69 and H89 of LSU-rRNA provide anticodons of UAG and UGA, respectively, as if both helices play a role of a stop signal recognizing tRNA.

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Although both loopheads were found in the proximity of the tripeptide motif Ser-Pro-Phe in the cryo-EM structures of the termination ribosome complex (Rawat et al., 2003; Klaholz et al., 2003), their locations were not close enough to make a complete helix in the presence of solid and voluminous structures of mRNA and tRNAs bound to P and E sites. The loophead of H89, in particular, seems to be located fairly far from the stop signal. Besides, what would happen to them when the A-site codon is either UAA or UGG. The codon for tryptophanyltRNA, UGG, would make more stable codon-hairpin helices with the hairpin loops of H69 and H89, CUA and UCA, respectively, than those of UAA/CUA and UAA/UCA. If one of the hairpin loops binds to the stop signal as if they were a kind of tRNA, there would be no more enough space for release factors. It is known that the neighborhood of mRNA in the A site is almost occupied by h18, h34, and h44 of SSU-rRNA. Accordingly, it would be more favourable for the UUA at the loophead of h37 of SSU-rRNA to make a first contact with the stop signal, to allow one of the tri- or tetrapeptide motifs to come close enough, and to withdraw immediately either after the recognition of a RF (or eRF1) for the purpose of receiving an aa-tRNA•EF-Tu(or eEF1)•GTP ternary complex. It seems to be a common mechanism to all living kingdoms except for mitochondria of some protista (T. brucei and P. falciparum) and animalia. An evolutionary trend can be observed that the number of nucleotides involved in the region of h34 – h37 has been larger and larger from mitochondria to eubacteria and eukarya. It is known that stop signals for vertebrate mitochondria are UAA, UAG, and instead of UGA, AGA and AGG (see Tables 10 and 12 of Osawa et al., 1992). Since mitochondrial tRNAs of animalia and some of the above protista are so simple, the first release factors might have evolved with a help of some regions of LSU-rRNA. This problem, however, remains still vague and uncertain. The codon for tryptophan is UGA for animalia and fungi mitochondria (see Table 12 of Osawa et al., 1992), UGG for plantae mitochondria as well as for archeae, chloroplast and eukarya, and either UGA or UGG for protista mitochondria and eubacteria (Sprinzl et al., 1998). The evolution of Trp-tRNA seems to have been developing in relation with that of stop signals (and protein factors, RF1, RF2, eRF1, and so on) as well as suppressor-tRNAs (Hirsh and Gold, 1971) and selenocysteine-tRNA (Hatfield and Diamond, 1993). Here the present author does not go into the problem of peptRNA hydrolysis. Rather interesting as to the present topic is the mechanism of removing class-1 release factors from the ribosome that requires GTP hydrolysis (Kisselev and Buckingham, 2000). If it is a process independent of translocation, the stability of the stopcodon/peptide-motif would not be strong enough compared to that of stop-codon/UUA helix. It is known that the X-ray structure of prokaryotic release factor does not resemble the shape of tRNA (Vestergaard et al., 2001). It is still enigmatic in what shape the RFs enter the decoding centre when a stop signal is detected, and retire from the location when GTP is hydrolyzed. When RF1 comes deep in touch with a stop signal UGA and finds itself unsatisfactory, how does RF2 take over the position by expelling the former occupant RF1 without GTP hydrolysis? The UUA in the loop of h37 could stay at the stop signal until a correct RF (either RF2 or eRF1 in this case) finds its binding site. When the UUA once leaves the position without finding any suitable RF, the stop signal could be open for either some suppressor-tRNAs or selenocysteine-tRNA, as described in the next section. If the stop signal is suppressed without having any frameshift, some role of the UUA could be important.

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7. Recognition of Sense Codon after Translocation When the A-site codon does not turn out to be a stop signal, the structures around the Asite tRNA as well as both the SRL region of 23S rRNA and the 5’-side of h35 of 16S rRNA return to those of poststate. When a newcomer of aa-tRNA•EF-Tu•GTP ternary complex comes at first to the strands 1103-1107 of 16S rRNA, its anticodon must be checked in the second place. It would be, however, almost without a break between translocation and codon recognition in the sense that the latter process could start before the elongation factor•GDP complex of the former process leaves the ribosome. More than 20 types of aa-tRNA complexed with EF-Tu•GTP could visit the A-site codon without need of making contact of its G domain with the GTPase associated centre. It would be a more efficient way of searching a cognate aa-tRNA if the GTPase associated centre around the SRL is still occupied by EF-G•GDP, although such an orientation of the ternary complex could not be confirmed by cryo-EM reconstruction of kirromycin-stalled ternary complex in the A site of the 70S ribosome at a resolution of ~ 20Å (Stark et al., 1997; Valle et al., 2002). It is known that peptidyl transferase reaction does not occur if aa-tRNA is bound to ribosome with EF-Tu and a nonhydrolyzable analogue of GTP (Skogerson and Moldave, 1968). Only when the GTPase associated centre is needed by a cognate (or near-cognate) ternary complex, the space would be open for the EF-Tu•GTP. On the other hand, EF-Tu•GDP must leave the place as quickly as possible after GTP hydrolysis, because EF-G•GTP comes to take over for accomplishing translocation. This could be the reason why EF-Tu•GDP (Kjeldgaard and Nyborg, 1992) makes a drastic conformational change with respect to the conformation of EF-Tu•GTP (Berchtold et al., 1993; Kjeldgaard et al., 1993). It seems possible for the newcomer ternary complex to approach the T site before EF-G•GDP leaves the GTPase associated centre of the ribosome completely. Figure 11a shows the model of the ternary complex of Cys-tRNA•EF-Tu•GDPNP (Nissen et al., 1995) at the instant of entering by the right-handed twist of the long helix into the GTPase binding site, the most of which is still occupied by EF-G•GDP complex. (The atomic coordinates for Figure 11a are given in the PDB file 1IPO.) It can be seen that the G base of position 53 of T-tRNA (designated as 53R) is closest to the C base of the nucleotide 1107 of 16S rRNA and that the anticodon of the TtRNA (from 34R to 36R) is already base-paired with the A-site codon (from m4 to m6). Most of the T-tRNA would be rejected from the ribosome without having GTP hydrolysis. Some of the noncognate newcomer bound to the T site might have good base pairs with a certain frame shift, yet the base pairing of its GTΨCG strand with the region 1103-1107 of 16S rRNA would not be satisfactory. Accordingly, only cognate and near-cognate newcomer would be successful in forming the base pairs. When the newcomer is either cognate or nearcognate, both anticodon and T-loop of the aa-tRNA would have close contacts with mRNA and the region 1103-1107 of 16S rRNA, respectively, so that its D-loop could also make a contact close enough to make two G-C pairs between G18 and G19 of the T-tRNA and C56 and C61 of P-tRNA. Sprinzl et al. (1976) found that various tRNA fragments containing Dloop inhibited the aa-tRNA binding at the A site. This is one of the main reasons why we adopt a tRNA docking pair model (Nagano and Nagano, 1997) for explaining the mechanism of translocation and codon recognition. Such a stereo picture is shown in Figure 11b. (The atomic coordinates for Figure 11b are given in 1IPQ.) In this model, its GTΨCG strand forms

Kozo Nagano

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a

b

Figure 11. Stereo pictures showing the behavior of aa-tRNA in the mechanism of sense codon recognition. (a) A ternary complex of Cys-tRNA•EF-Tu•GTP (Nissen et al., 1995) enters the decoding site to make the codon-anticodon base pairs in the most favourable orientation for its GTΨCG sequence (53R-57R) in light blue closest to the nucleotides 1103-1107 of 16S rRNA in light blue. This site is a recognition mode of A site, which is refered to as T site, and abbreviated as R in these figures. The TtRNA in the ternary complex is drawn in magenta, while its G domain of EF-Tu in violet (and designated as EF-Tu-G), its GTP in brown, and domain II of EF-Tu (EF-Tu-II) in pink. P-tRNA, mRNA, 16S rRNA and SRL are a little different from those in Figure 6a. SRL as in Figure 8b. P-V denotes the largest variable loop chimerically introduced onto the 3D structure of yeast tRNAPhe. This type of interaction between the A-site codon and 20 kinds of aa-tRNAs seems possible shortly after GTP hydrolysis of EF-G•GTP and translocation, even if EF-G•GDP still occupies a large space near the GTPase associated centre. The numerals ending with R are residue numbers of T-tRNA. (b) The region surrounding an aa-tRNA brought by a ternary complex with EF-Tu•GTP. T-tRNA is drawn in magenta. This picture represents base pairs between GTΨCG sequence of aa-tRNA and nucleotides 1103-1107 of 16S rRNA in light blue and also between the region 2655-2661 of 23S rRNA (SRL) in red and the region 1069-1075 of 16S rRNA in light blue. This arrangement of tRNA facilitates fitting of its anticodon with A-site codon and also its D-loop with the T-loop of P-tRNA. The large R denotes the location of the anticodon loop of T-tRNA. P-T is the T-loop of P-tRNA, while R-V is the chimerically introduced largest variable loop of T-tRNA. The other labels are as in Figures 6a and 8a.

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an A-form helix with the strand 1103-1107 of 16S rRNA in light blue. The SRL is still basepairing with the region 1069-1075 of 16S rRNA. EF-Tu•GTP is far from the viewer without disturbing both P-tRNA and 16S rRNA. This is a putative transition-substate model of mRNA•tRNA•16S rRNA complex in the mechanism of codon recognition. The conformation of the anticodon loop of the T-tRNA is the same as that of yeast tRNAPhe (Holbrook et al., 1978), in which a hydrogen-bond between N3 atom of the base of U33 and O5’ atom of A36 of T-tRNA is formed (see Figure 5c of Holbrook et al., 1978). When the long helix is unfolded in the next step, the competition for survival between the T- and E-tRNAs is going to be held through the trasition-state three tRNA binding conformation, as already described in the section 2 and shown in detail by Figures 5a, b, c, and d. In the case of noncognate tRNA recognition, the base-pairing between the A site codon and the anticodon of the T-tRNA does not fit very well. This discrepancy at the anticodon loop of the T-tRNA is amplified at the R-P contact region under the condition of the basepairing between the GTΨCG of T-tRNA and the strand 1103-1107 of 16S rRNA and leads to a failure in forming two G-C pairs between the P- and T-tRNAs. As its result, the ternary complex of such a noncognate tRNA could be quickly discarded, as visualized in Figure 12a. (The atomic coordinates for Figure 12a are given in the PDB file 1IPR.) This structure minus leaving noncognate aa-tRNA•EF-Tu•GTP and EF-G•GDP complexes is the structure closest to that of poststate ribosome, although the region around the spectinomycin binding site is still interacting with the SRL. When the T-tRNA is either cognate or near-cognate, the conformational change at the anticodon loop can be expected. More precisely speaking, the codon-anticodon base pairs have a tendency to become more stabilized to form an ideal Aform with a right-handed twist between the first and second Watson-Crick base pairs. This would break a hydrogen-bond between N3 atom of U33 and O5’ atom of A36 of T-tRNA and force the U33 base to stick out. This conformational change of the anticodon loop at the A site seems to affect the increase in the proflavin fluorescence from wybutine at position 37, which monitors the environment of the anticodons with cognate and near-cognate tRNAs, but is not observed with noncognate tRNA (Rodnina et al., 1995), while that of proflavin at the D-loop showed a small initial increase on binding with all tRNAs including noncognate aatRNA (Rodnina et al., 1994). The present author assumes that the U33 is base-paired with an invariant adenine base of 16S rRNA for much better stability, as already described in the section 3. As a result, rotation at the middle part of the T-tRNA occurs and gives rise to a collision of D-loop of T-tRNA with T-loop of P-tRNA, resulting in formation of two G-C pairs between the consecutive invariant G18 and G19 of T-tRNA and the other invariant C56 and C61 of P-tRNA at the expense of one G-C pair between G53 and C61 of P-tRNA (viz. a gain of three hydrogen bonds). If the T-loop of P-tRNA is blocked by the D-loop in the various tRNA fragments (Sprinzl et al., 1976), such a recognition would end in failure. Valle et al. (2002) showed by cryo-EM that the T-loop side of cognate aa-tRNA•EF-Tu•GDP ternary complex bound to kirromycin-stalled ribosome interacts with the L11 binding region of the GTPase associated centre. When the T-tRNA is either cognate or near-cognate, the conformational change at the anticodon loop leads to formation of two G-C pairs between the consecutive invariant G18 and G19 of T-tRNA and the other invariant C56 and C61 of PtRNA. In this case, the U33 base at the anticodon loophead of the T-tRNA is half exposed, but the conformation of 16S rRNA concerned is pushed out by a pressure exerted from the

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triangular RNA helices to restore the conformation as shown in Figure 6a. Thus, the SRL of 23S rRNA restores the GTPase-active conformation. Such a model is shown in Figure 12b. (The atomic coordinates for Figure 12b are given in the PDB file 1IPU.) The G domain of EF-Tu intervenes between the 16S rRNA and the SRL at the GTPase associated centre, although the location and the orientation of the SRL is a little different from the model that is shown in Figure 6a. The spatial relationship between the T-tRNA and EF-Tu•GTP in Figure 12b seems to be exactly what we see in Figure 1b of Stark et al. (1997). This might be a reasonable explanation for why thiostrepton does not inhibit binding of the ternary complex of aa-tRNA•EF-Tu•GTP on the ribosome. We can understand also why the 3’-end of the tRNA does not reach the site of puromycin when GTP is replaced by a noncleavable derivative such as GDPNP (Skogerson and Moldave, 1968). In order to achieve a new AtRNA with its 3’-end at the peptidyl transferase centre, the highest barrier to be overcome is to bring the G-C pairs between the D-loop of T-tRNA (R-D in Figure 12b) and the T-loop of P-tRNA (P-T in Figure 12b) much closer to the axis of rotational movement of tRNA docking pair in translocation to break the G-C pairs between the D-loop of P-tRNA (P-D) and the Tloop of E-tRNA (not shown in Figure 12b) as well as the codon-anticodon base pairs at the E site. Since the near-cognate tRNA is characterized by an excess of GTP cleavages, most of near-cognate ternary complex would be discarded after GTP hydrolysis. Only very few nearcognate and most of cognate ones go to the structure shown in Figures 6a and b (except for EF-G•GDP structure). In the case of a near-cognate aa-tRNA, particularly when it is discarded, the conformation of 16S rRNA should come back to that of Figure 12a (without EF-G•GDP), and further return to the structure of poststate ribosome. The functional role of H96 and H97 would be important in switching the location and orientation of the SRL back and forth. H97 could make a direct contact with the tip of H84 in this sub-state, as shown in Figure 12c. (The atomic coordinates for Figure 12c are the same as those for Figure 12b, viz. the PDB file 1IPU.) The locations of H84 and H97 are just below the E- and P-tRNAs and help the two G-C pairs of P-E tRNA docking pair resist against the invasion of the newly formed two G-C pairs between the T- and P-tRNAs. H93 is directed toward the SRL and located at the intermediate position between the pre- and poststates. This situation is just like two poises on a balance, and the shock transmitted from the conformational change of EFTu•GDP breaks the weaker side of the two codon-anticodon base pairs between the E and T sites. If the codon-anticodon interaction at the E site is either weaker or as strong as that of the T site, the H84 would move upward and breaks the P-E G-C pairs and expel the E-tRNA from the ribosome. T-tRNA occupies the A site, and H93 goes to the location of the prestate. On the contrary, if the T-site base pairs are weaker than those of the E site, H84, H93 and H97 return to their respective locations in the poststate, as shown in Figure 8b. This is exactly what we call proofreading mechanism. It would be important to realize that the above discussion does not hold at all for the most primitive ribosome structure of Trypanosoma brucei mitochondria, because H84, H96, and H97 are all missing. Accordingly, the functional role of the helices, H84, H96, and H97, could be that of enhancing the efficiency in the negative cooperativity between the T and E sites (Nierhaus, 1990), that is immediately related with enhancing the accuracy in codon recognition.

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a

b

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c

Figure 12. Stereo pictures showing the behavior of aa-tRNA in the mechanism of sense codon recognition as well as the regions surrounding the decoding site in the transition substate in with aatRNA•EF-Tu•GTP ternary complex. (a) When the T-tRNA in magenta is noncognate, it is discarded in the opposite direction from the GTPase associated centre near SRL in red. The ternary complex would shift in this direction under a pressure of the 16S rRNA involved in the model in Figure 8b when it’s Aform helix between the GTΨCG sequence of T-tRNA and the nucleotides 1103-1107 of 16S rRNA is unfolded. EF-G•GDP still occupies the same space as in Figure 6a and Figure 11a. Another ternary complex can approach the A-site codon by thrusting with its anticodon loop into the space between the discarded ternary complex and the EF-G•GDP complex and by restoring the 3D structure shown in Figure 6a. The symbols and colours are as in Figures 6a and b. (b) When the T-tRNA in magenta is either cognate or near-cognate, the 16S rRNA involved in the model in Figure 6a pushes the body of EF-Tu•GTP complex toward the SRL at the GTPase associated centre so that it could start GTP hydrolysis. Both base-pairings of codon-anticodon and between D-loop of T-tRNA and T-loop of PtRNA are kept. The A-form helices between the GTΨCG sequence of T-tRNA and the nucleotides 1103-1107 of 16S rRNA as well as between 1069-1075 of 16S rRNA and SRL are unfolded. G domain of EF-Tu shifts downward, and SRL also shifts below the G domain with its active A2660 facing to the GTP, which is not visible in this figure. In order for the aminoacyl group to reach the peptidyl transferase centre, EF-Tu•GDP must be released after GTP hydrolysis. A pressure due to the EFTu•GDP release would push the T-tRNA toward the axis of rotational movement in translocation, resulting in expelling E-tRNA, which is bound to the D-loop of P-tRNA at the location designated as PD in the case of cognate T-tRNA. On the contrary, a near-cognate codon-anticodon base pairs at the T site would be broken instead of breaking the codon-anticodon base pairs at the E site. R-D and R-T are the D- and T-loops of T-tRNA, respectively. It is clear from this picture that EF-G•GDP can no more occupy the space as in Figsures 6a and 11a. The other symbols and colours as in (a). (c) The regions surrounding the decoding regions in the transition substate of GTP hydrolysis in the aa-tRNA•EFTu•GTP ternary complex. It is important to note that the tip of H84 makes direct contact with H97 under the two G-C pairs of P-E tRNA docking pair (not shown in this figure), resulting in formation of a strong support against a pressure exerted from the new G-C pairs of R-P tRNA pair in the case of near-cognate T-tRNA but breaks the two G-C pairs of P-E tRNA pair in the case of cognate T-tRNA. See text for further explanation about the proofreading mechanism. The other labels and colours are as in Figures 6a and b as well as in Figures 8a, b and c.

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When the anticodon of T-tRNA is near-cognate, in contrast to the cognate codonanticodon base pairings at the P and E sites, the base pair, U33(A)-A1396(16S), would be finally broken, even if it could be once formed, because of an impulse caused by a conformational change of EF-Tu due to GTP hydrolysis and break the A-site codonanticodon base pairings as well as the G-C base pair between T- and P-tRNAs. In the case of cognate T-tRNA, on the other hand, those base pairs between P- and E-tRNAs would be broken faster by the reverse mechanism of barrier melting after the long helix formation is over. The codon-anticodon base pairings as well as the base pairs, U33(A)-A1396(16S) and G18(A)-C56(P), would be kept and induce another G-C pair, G19(A)-C61(P), that is exclusive against G18(P)-C56(E), resulting in formation of crystallographically observed Tloop-D-loop contact region. Such an elbow region of E-tRNA would be caught at the plane of its G19-C56 by a pseudoknot helix GUAGGAUA2119/ GACCUUGA2169, as observed by Korostelev et al. (2006), and brought away by a movement of L1 stalk as a result of 30S subunit closure (Ogle et al., 2002). The total of the above described conformational changes of the tRNAs and the surrounding decoding site on the small and large ribosomal subunits before and after GTP hydrolysis is usually referred to as proofreading mechanism. The 3’ end of aa-tRNA at the T site goes to the peptidyl transferase centre at the A site and becomes AtRNA, giving rise to transpeptidation with the 3’ end of P-tRNA. The final structure is ready for the next step of the elongation cycle in cooperation with EF-G•GTP complex at the GTPase associated centre of the large subunit ribosome.

8. Comparison of the Present Proofreading Mechanism with the other Proposed Ones In protein biosynthesis, mostly cognate and very few near-cognate aa-tRNAs can be allowed to enter the A site under the structural control of both EF-Tu•GTP and ribosome with an error rate of about 1 per 103 ~ 104 amino acid residues incorporated (Fersht, 1985). Such a high specificity of codon recognition is far beyond the thermodynamically expectable limit if the codon recognition occurs only in a single step. Even if a kind of induced-fit mechanism is introduced, only a simple conformational change would not be enough to explain such a low error rate of recognition (Hershlag, 1988). Accordingly, the two-step tRNA selection mechanism has been proposed. The first step before GTP hydrolysis is called initial selection, while the second after GTP hydrolysis is proofreading (Rodnina and Wintermeyer, 2001). Various steps more than two were detected using single-molecule fluorescence resonance energy transfer (smFRET) (Blanchard et al., 2004b), the schematic representation of which was shown as seven steps in total (Blanchard et al., 2004a). The measurements were made for the time-dependent distances between donor and acceptor fluorophores attached to the elbow region of tRNA molecules on the ribosome. The transition of FRET = 0.74 to 0.45 implies that the distance between tRNA elbows increases by about 10 Å (from 44.3 Å in the crystal structure of Yusupov et al. (2001) to 56.5 Å in our two tRNA docking model (Nagano and Nagano, 1997), as an example, although Blanchard et al. (2004b) insisted the presence of a hybrid state structure). In the seven-step mechanism of Blanchard et al. (2004a), step 0 binds only one tRNA at the P site, step 1 with 0.0 FRET has one aa-tRNA•EF-Tu•GTP ternary

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complex, step 2 with 0.35 FRET has the above complex making contact with the A-site codon, step 3 with 0.5 FRET has the above complex at the A site, step 4 with 0.5 FRET, in which GTP is hydrolyzed, step 5 with 0.5 FRET, in which some of the tRNA at the A site is rejected, step 6 with 0.75 FRET has the aa-tRNA•EF-Tu•GDP complex at the A site, and step 7 with 0.75 FRET, in which peptidyl transfer is completed but EF-Tu•GDP still remains at the A site. In these steps, steps 2 and 3 are considered to be the initial selection, while steps 5 and 6 are thought to be proofreading (Blanchard et al., 2004a). The step 1 of their mechanism could be the same as what we call the putative first interaction of the ternary complex with the cavity region of the small subunit in the present mechanism, as described in section 2, while the steps 3, 4, and 5 could correspond to unfolding of 16S rRNA, formation of the long helix between 16S and 23S rRNAs, and unfolding of the long helix, respectively, in the present mechanism. In order to achieve the high overall selectivity, the initial selection step must be about as precise as proofreading (Gromadski and Rodnina, 2004). During proofreading alone, about 1 amino acid out of 100 near-cognate aa-tRNAs is incorporated into peptide (Pape et al., 1999). On the other hand, early estimations for the efficiency of initial selection suggested the value of not more than 1 out of 10 ~ 100 near-cognate ternary complexes (Bilgin and Ehrenberg, 1994). There are two ways for enhancing efficiency of initial selection as follows: (1) by selective stabilization of cognate, but not near-cognate, codon-anticodon helices on the ribosome, and (2) by modulating the rates of forward reactions depending on the structure of the codon-anticodon helix by means of an induced fit (Pape et al., 1999) or ‘domain closure’ mechanism (Ogle et al., 2002). The structure of the 30S subunit with A-site codon and cognate tRNA ASL showed conformational changes in the 30S subunit, in which A1492, A1493, and G530 interact with the minor groove of the first two codon-anticodon base pairs (Ogle et al., 2001). In order to find the structural differences caused by binding one cognate and two near-cognate anticodons to the above, Ogle et al. (2002) used the sequences of the following three ASLs: (1) cognate ASLPhe GGGGAUUGAAAAUCCCC; (2) near-cognateASLLeu2 CUACCUUGAGGΨGGUAG; (3) near-cognate ASLSer CACGCCUGGAAAGΨGUG. Here, as (4), a hairpin loop of Thermus thermophilus helix 6 (so-called spur region) in a neighbouring 30S molecule having a following sequence was used as a mimic of P-site ASL: …GGCCGCGGGGUUUUACUCCGUGGUC… In the above four sequences, single underlines, double underlines, and broken underlines indicate anticodons, conserved U33s, and base-paired nucleotides, respectively. (It is important to note that the sequence (4) is not like the ASL of a real tRNA, in the sence that the base of U33 in (4) would not stick out because of a strong base pair G32-C38, which seems fatal in our proofreading mechanism.) As mRNAs, either …UUUAAA or …UUCAAA is used. The ‘domain closure’ is observed in the presence of cognate ASLPhe and involves rotations of the head toward the shoulder and the subunit interface and of the shoulder (S4, G530 loop with surrounding regions of 16S rRNA and S12) toward the intersubunit space and the h44/h27/platform region (Ogle et al., 2002). This ‘domain closure’ is enhanced in the additional presence of paromomycin and induces the decoding error. The NMR structural study of paromomycin bound to a fragment of A-site 16S rRNA suggested a conformational stabilization of the decoding site that specifically recognizes the codon-

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anticodon helix (Fourmy et al., 1996). With near-cognate ASLs, the closed 30S domain is not observed except for the cases in the presence of paromomycin. This is a kind of paradox, suggesting that stabilization does not necessarily contribute to enhancing the accuracy in translation. The ‘domain closure’ should still have a possibility of dynamic behavior that is called an induced fit. The structural role of A1492 and A1493 is hydrogen bonding to the minor groove of codon-anticodon helix at the A site, in which A1493 and A1492 interact with the first and second bases, respectively, as typical cases of A-minor motif (Nissen et al., 2001). However, the mode of interaction is rather vague (Ogle et al., 2001). It does not explain why both adenines are highly conserved, because substitution for either adenine by guanine could work about the same. In the case of cognate ASLPhe with paromomycin, the mode of interaction can be seen more clearly (Ogle et al., 2002). Even in the case of nearcognate ASLLeu2 with paromomycin, the N3 atom of A1493 is found to hydrogen-bond to N2 amino group of guanine base in the G-U mismatch at the first codon position as is the case for a Watson-Crick pair of either G-C or C-G. The hydrogen bond cannot be seen very well between the N3 of A1492 and the N2 amino group of guanine at the second codon position even in the presence of paromomycin, although it does not look like a wobble base pair (Ogle et al., 2002). Considering that both first and second codon positions take various base pairs, even though they are all Watson-Crick type, the hydrogen bonding mode of both A1492 and A1493 should be variable. Fixing their position in the presence of paromomycin does not seem to contribute to enhancing the accuracy in translation. The mechanism of translation described in the preceding sections has started from a consideration that, at the beginning of long evolutionary events of life, a process of noncognate aa-tRNA discarding without GTP hydrolysis could have been more important than the process of discriminating cognate one from near-cognate ones by means of GTP hydrolysis. Although it is well known that an aa-tRNA•EF-Tu•GTP ternary complex comes to the L7/L12 stalk shortly before GTP hydrolysis, it does not necessarily mean that the stalk is the first interaction site for the ternary complex, because most of them, such as noncognate ones, are discarded without GTP hydrolysis. Thiostrepton, which inhibits binding of EFG•GTP at the stalk, does not inhibit binding of an aa-tRNA•EF-Tu•GTP ternary complex. Another related basic question is how the next codon can be found without having a frame shift, if the stalk is the first interaction site for the elongation factors to the ribosome. The interaction between a GTΨC-loop of tRNA and the 3’-side of h35 seemed important for that purpose. The present hypothetical mechanism of the long helix formation between small and large ribosomal subunits was introduced in this way. Based on the smFRET method, described above, Blanchard et al. (2004a) proposed a kinetic proofreading mechanism, in which ribosomal recognition of correct codon-anticodon helix drives rotational movement of the incoming complex of aa-tRNA•EF-Tu•GTP toward pep-tRNA during selection on the ribosome, although their experiment completely ignores the significance of E-site tRNA. They found that the formation of the stable 0.5 FRET state occurs after initial selection is completed. The step of the ternary complex weakly bound to the ribosome before GTP hydrolysis could be considered to be the stage in our mechanism, at which a long base-pairing helix between the 5’-side of h35 of 16S rRNA and the SRL region of 23S rRNA is being unfolded, and the final match of the codon-anticodon helices between A and E sites is coming to a climax, as shown by the 3D structures in Figures 5 and 12.

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CONCLUSION The mechanism of translation has been studied for more than 40 years. First approaches were mostly based on chemical analyses such as sequencing of proteins (Giri et al., 1984) and rRNAs (Brosius et al., 1978; 1980), footprinting (Moazed and Noller, 1989), crosslinking (Brimacombe, 1995), and so on. At that time, the ribosome looked like an elephant elegantly moving its nose, while a mass of researchers were analogous to people watching and analyzing its functional movements. Electron microscopy (Stöffler and Wittmann, 1977) was like photography that allowed us to have an idea that the elephant has also large ears and feet. X-ray analysis of the whole ribosome (Yusupov et al., 2001) gave us a first picture of the roentgenograms of the elephant that had thick bones and internal organs. People first thought that the elephant was quite static, but finally realized that it was quite dynamic, when it was alive. This is the present situation regarding the image of the whole ribosome after the appearance of four more X-ray structures (Bashan et al., 2003; Schuwirth et al., 2005; Korostelev et al., 2006; Selmer et al., 2006). When the present author first submitted a paper on the mechanism based on the intersubunit long helix formation nearly ten years ago, it was rejected by saying that the mechanism would be soon denied by a forthcoming result of X-ray analysis. After the appearance of Yusupov et al. (2001), the distance between SRL and h35 larger than 100Å was the main reason for the critical objections. Despite the dynamic character of the X-ray results, however, the existence of the barrier between P- and E-tRNAs bothers crystallographers in explaining the first step of translocation. What the present author insists in this chapter is that such a phenomenon could not be explained based on the frozen results of X-ray structures, but by a chain reaction of conformational changes, which could allow the functional interactions of the regions widely separated by more than 100Å on the X-ray structure. Almost all enigmatic steps of the mechanism of translation could be clarified by this type of approach.

METHODS Programs and Coordinates The programs for manipulating the 3D models in the present work are the same as those used in the modelling of tRNA docking pair (Nagano and Nagano, 1997). The 3D structures of the region of the nucleotides 2646-2674 of 23S rRNA domain were built with reference to the NMR structure (Szewczak et al., 1993). The atomic coordinates of EF-G (Ævarsson et al, 1994) and the ternary complex of Cys-tRNA•EF-Tu•GDPNP (Nissen et al., 1995) are obtained from the Protein Data Bank. GTP model was fitted to EF-G following the corresponding structure of EF-Tu•GDPNP (Berchtold et al., 1993). In order to draw coloured stereo pictures and to edit them as we like, a program called RasMol (Sayle, 1992-1999; Bernstein, 1995-2000) and a program from Molecular Simulations Inc., San Diego, U.S.A., called WebLabViewer Pro 3.0, were used. The latter models were edited by adding labels and numerals by use of a software Adobe Illustrator 10.0J from Adobe Systems Inc., San Jose, U.S.A.

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Sprinzl, M., Horn, C., Brown, M., Ioudovitch, A. & Steinberg, S. (1998). Compilation of tRNA sequences and sequences of tRNA genes. Nucl. Acids Res., 26, 148-153. The database is available at www.uni-bayreuth.de/departments/biochemie/sprinzl/trna/ Sprinzl, M., Wagner, T., Lorenz, S. & Erdmann, V. A. (1976). Regions of tRNA important for binding to the ribosomal A and P sites. Biochemistry, 15, 3031-3039. Stark, H., Rodnina, M. V., Rinke-Appel, J., Brimacombe, R., Wintermeyer, W. & van Heel, M. (1997). Visualization of elongation factor Tu on the Escherichia coli ribosome. Nature, 389, 403-406. Stöffler, G. & Wittmann, H. G. (1977). Primary structure and three-dimensional arrangement of proteins within the Escherichia coli ribosome. In: Molecular mechanism of protein biosynthesis. Academic Press, Inc., New York, San Francisco, London. pp. 117-202. Szewczak, A. A., Moore, P. B., Chan, Y. L. & Wool, I. G. (1993). The conformation of the sarcin/ricin loop from 28S ribosomal RNA. Proc. Natl. Acad. Sci. USA, 90, 9581-9585. Valle, M., Sengupta, J., Swami, N. K., Grassucci, R. A., Burkhardt, N., Nierhaus, K.H., Agrawal, R. K. & Frank, J. (2002). Cryo-EM reveals an active role for aminoacyl-tRNA in the accommodation process. EMBO J., 21, 3557-3567. Vestergaard, B., Van, L. B., Andersen, G. R., Nyborg, J., Buckingham, R. H. & Kjeldgaard, M. (2001). Bacterial polypeptide release factor RF2 is structurally distinct from eukaryotic eRF1. Mol. Cell, 8, 1375-1382. Wilson, K. S. & Noller, H. F. (1998). Mapping the position of translational elongation factor EF-G in the ribosome by direct hydroxyl radical probing. Cell, 92, 131-139. Wimberly, B. T., Guymon, R., McCutcheon, J. P., White, S. W. & Ramakrishnan, V. (1999). A detailed view of a ribosomal active site: The structure of the L11-RNA complex. Cell, 97, 491-502. Yamada, Y. & Ishikura, H. (1973). Nucleotide sequence of tRNASer3 from Escherichia coli. FEBS Lett., 29, 231-234. Yusupov, M. M., Yusupova, G. Zh., Baucom, A., Lieberman, K., Earnest, T. N., Cate, J. H. D. & Noller, H. F. (2001). Crystal structure of the ribosome at 5.5 Å resolution. Science, 292, 883-896. Zavialov, A. V., Buckingham, R. H. & Ehrenberg, M. (2001). A posttermination ribosomal complex is the guanine nucleotide exchange factor for peptide release factor RF3. Cell, 107, 115-124. Zavialov, A. V. & Ehrenberg, M. (2003). Peptidyl-tRNA regulates the GTPase activity of translocation factors. Cell, 114, 113-122. Zavialov, A. V., Haurylink, V. V. & Ehrenberg, M. (2005). Guanine-nucleotide exchange on ribosome-bound elongation factor G initiates the translocation of tRNAs. J. Biol., 4(9), 119.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter IV

TRANS-TRANSLATION BY TMRNA AND A PROTEIN MIMICKING TRNA AND MRNA Hyouta Himeno, Daisuke Kurita, Akira Muto Department of Biochemistry and Molecular Biology, Faculty of Agriculture and Life Science, Hirosaki University, Hirosaki 036-8561, The United Graduate School of Agricultural Sciences, Iwate University, Morioka 020-8551 and RNA Research Center, Hirosaki University, Hirosaki 036-8561, Japan

ABSTRACT Usually, a single polypeptide or even multiple polypeptides is produced from a single mRNA. Trans-translation is an irregular translation system in eubacteria, in which a single polypeptide is synthesized from two separate molecules of coding RNAs, mRNA and tmRNA. It rescues a stalled translation on the ribosome and provides a peptide tag for degradation to the C-terminus of the nascent polypeptide to enable recycling of ribosomes, promote degradation of truncated mRNA and prevent accumulation of abortively synthesized polypeptides in the cell. Trans-translation is involved not only in a quality control system in the cell but also in various kinds of cellular functions. During trans-translation, tmRNA plays a dual function both as a tRNA and as an mRNA. Alanyl-tmRNA somehow enters the vacant A-site of the stalled ribosome like aminoacyltRNA but without a codon-anticodon interaction, and thereafter the coding region of tmRNA is substituted for mRNA. As a consequence, alanine encoded nowhere but aminoacylated to tmRNA serves as the connector between the truncated peptide encoded by mRNA and the C-terminal tag-peptide encoded by tmRNA. Such an acrobatic feat is accomplished by elaborate interplay between the tRNA and mRNA functions of tmRNA with the help of a protein factor SmpB. Our recent study has suggested that both tmRNA and SmpB mimic the structures and functions of tRNA and mRNA during transtranslation, addressing how tmRNA preferentially recognizes the stalled ribosome, and what substitutes for a codon-anticodon interaction.

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INTRODUCTION Recent progress in molecular biology has brought a new concept that a vast number of noncoding RNAs play diverse roles in the eukaryotic and prokaryotic cells (Zamore & Haley, 2005, Wasserman, 2002). In 1980th, however, most of the textbooks of molecular biology referred to only tRNA, mRNA and rRNA. TmRNA was initially found from Escherichia coli at 1979 (Ray & Apirion, 1979). Until the middle of the 1990th, it has been regarded merely as a small stable and presumably noncoding RNA of an unknown function. Later findings of the tRNA-like structure, a tRNA-like function and an mRNA function, have opened a door towards a novel molecular mechanism and a novel cellular system. TmRNA shares two different kinds of properties, a carrier of genetic information and a device for decoding genetic information, which are usually carried in separated RNA molecules. TmRNA facilitates an irregular translation named trans-translation, in which a single polypeptide is produced from two coding RNA molecules (Figure 1). Trans-translation rescues the ribosome stalled at the 3’ end of truncated mRNA lacking a termination codon and provides the incomplete nascent polypeptide from a truncated mRNA with a tag for degradation. This promotes recycling of the stalled ribosomes in the cell. As these marvellous functions were revealed, new questions arose in the light of the recent progress in our understanding about the processes of translation on the ribosome at the atomic level (Ramakrishnan, 2002) and the unexpectedly diverse functions of small noncoding RNAs in the control of gene expression (Zamore & Haley, 2005, Wasserman, 2002). What is the higher order structure of tmRNA? How does tmRNA elaborately coordinate the tRNA and mRNA functions in the limited space of the ribosome to accomplish trans-translation? How does tmRNA find a stalled translation? Are there any other factors involved in trans-translation? What is the fate of the aberrant polypeptide from tuncated mRNA? Does trans-translation serve merely as a surveillance system in the cell? Here, we review the history, the molecular mechanism and the cellular functions of trans-translation including our recent model.

TmRNA About thirty years ago, Ray & Apirion (1979) found a stable RNA in the fraction of sedimentation coefficient 10S from E. coli cell extracts and it was named 10S RNA. Later, 10S RNA has been found as a mixture of two RNA species that have been renamed 10Sa RNA and 10Sb RNA (Jain et al., 1982), the latter being a component of RNase P. After ten years from the finding, the primary structure of 10Sa RNA was determined (Chauhan & Apirion, 1989). As its novel functions have been revealed, its name has gradually been replaced from 10Sa RNA to tmRNA (transfer-messenger RNA), although some groups still call it SsrA from the name of its gene, ssrA (small stable RNA A).

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Figure 1. Schematic representation of trans-translation. Alanine aminoacylated to tmRNA (shown in white with a red background) serves as the connector between the truncated peptide encoded by truncated mRNA (shown in grey) and the C-terminal residues encoded by tmRNA (shown in red).

TmRNA is widely or perhaps almost exclusively distributed among the eubacterial kingdom. Their primary structures are available from two databases of tmRNA, the tmRDB (http://psyche.uthct.edu/dbs/tmRDB/tmRDB.html) (Zwieb et al., 2003; Andersen et al., 2006) and the tmRNA Website (http://www.indiana.edu/~tmrna/) (Williams, 2000). TmRNA is also present in chloroplasts or mitochondria of some lineages of lower eukaryotes. However, tmRNA or its gene has never been found in the eukaryotic cytoplasms or archaebacteria.

STRUCURE OF TMRNA A tRNA-like secondary structure has been found by our and Inokuchi’s groups (Ushida et al., 1994; Komine et al., 1994). It can be formed by the 5’- and 3’-terminal sequences of tmRNA from Bacillus subtilis, Mycoplasma capricolum and E. coli. A 5-base-pair stem with a 7-base loop, a stem of 7 base pairs, a discriminator base and the CCA-3’ sequence are arranged just like the TΨC-arm, the amino acid acceptor stem, a discriminator base and the CCA-3’ sequence in the upper right half of the canonical secondary structure of tRNA (Figure 2). Consensus sequences in tRNA, such as UUCPuAPyU sequence in the TΨC-loop and the CCA-3’ sequence, are present in tmRNA. In addition, two major modified bases in typical tRNA, 5’-methyl cytosine (T or thymine) and pseudouridine (Ψ), have been identified in the TΨC-loop in E. coli tmRNA (Felden et al., 1998) (Figure 2). TmRNA has a noncanonical D-arm apparently lacking a stem structure, which is often found in some animal mitochondrial tRNA (Himeno et al., 1987), while it contains a GG sequence that is a consensus sequence in the D-loop of tRNA (Figure 2). The tertiary interaction between the D-arm and the TΨC-loop has been suggested by a cross-linking study (Zwieb et al., 2001). It has also been suggested that the GG sequence in the D-arm has a functional interaction with

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Figure 2. The secondary structure of E. coli tmRNA. The tag-encoded sequence is highlighted in white with a black background. The region required for recognition by alanyl-tRNA synthetase is designated with a blue background. Several base substitutions affect the efficiency and fidelity of trans-translation in vitro (Lee et al., 2001), designated by closed and open arrowheads in red, respectively.

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the TΨC-loop, like in the canonical tRNA structure (Barends et al., 2002). The finding of the tRNA-like secondary structure in tmRNA has brought the first breakthrough in its function, described below. In contrast to high conservation of both terminal regions comprising a tRNA-like secondary structure, the primary structure of the remaining 90% of total approximately 350 nucleotides are much less conserved, making difficult to predict the entire secondary structure of tmRNA. Sequence comparisons (Williams & Bartel, 1996; Felden et al., 1997) and chemical and enzymatic structural probings (Felden et al., 1996, 1997; Hickerson et al., 1998) have enabled to build a secondary structure model of this molecule (Figure 2). The central two third of this molecule forms a large loop comprised of four pseudoknot structures (PK1 to PK4) and a few stem and loops, being connected to the terminal tRNA-like domain by a long stem with a few bulges or loops. This secondary structure model has been validated by mutational analyses (Nameki et al., 1999a, 1999b) and by later accumulation of the tmRNA sequences. The secondary structure of tmRNA is widely conserved, although with some variation in the pseudoknot-rich region. For example, the fifth pseudoknot is found in tmRNA from some cyanobacteria (Williams, 2002). As an extreme example, a tmRNA molecule from some lineages of alphaproteobacteria, betaproteobacteria, cyanobacteria or mitochondria of lower eukaryotes is comprized of two separated chains, a 5’-coding piece and a 3’-amino acid acceptor piece (Keiler et al., 2000; Williams, 2002; Jacob et al., 2004; Sharkady &Williams, 2004). 2-piece tmRNA shares similar structural features such as the tRNA-like structure with one-piece tmRNA, although possessing fewer pseudoknots (Keiler et al., 2000; Gaudin et al., 2002; Sharkady & Williams, 2004). Indeed, the functional redundancy of pseudoknot structures except PK1 in E. coli tmRNA has been exemplified in vitro (Nameki et al., 2000), although they participate in the proper folding and processing of 1-piece tmRNA (Wower er al., 2004). The tertiary structure of tmRNA has been presumed to be difficult to be determined, since it is large and flexible as compared to tRNA (Felden et al., 1997). Models of PK2 and PK4 structures have been proposed from a comparative study (Zwieb et al., 1999). A model of the tRNA-like structure has been built based on the angle between two helices obtained by transient electric birefringence measurement (Stagg et al., 2001). NMR studies have revealed the structure of PK1 from Aquifex aeolicus (Nonin-Lecomte et al., 2006). A crystal structure of a fragment of the tRNA-like domain in complex with a binding protein, SmpB, from A. aeolicus (Gutmann et al., 2003) or Thermus thermophilus (Bessho et al., 2007) has been shown. According to the latter crystal structure, the long stem with a few bulges or loops leading to PK1 is thought to generate from the tRNA-like domain in a direction similar to that of the long variable arm in class II tRNA (Bessho et al., 2007). The entire 3D structure models of tmRNA from several eubacterial species have been built in consideration of the functional distance between the tRNA-like domain and the coding region (Burks et al., 2005). The entire structure of T. thermophilus tmRNA in complex with the ribosome and a few binding proteins has been visualized by cryo-electron microscopy, although with low resolution (Valle et al., 2003a; Kaur et al., 2006).

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PROCESSING OF TMRNA E. coli genome has a single gene for tmRNA (ssrA) under a putative sigma 70 promoter sequence (Oh et al., 1990; Komine et al., 1994). The 457-nucleotide primary transcript is processed into 363-nucleotide mature tmRNA (Komine et al., 1994). The tRNA-like structure allows removal of 7 nucleotides from the 5’-end of the 457-nucleotide precursor by RNase P that usually functions in the 5’ processing of tRNA (Inokuchi et al., 1994). The 3’ processing involves endoribonucleases, RNase III, RNase E (Srivastava et al., 1990, 1992; Lin-Chao et al., 1999), and exoribonucleases, RNase T, RNase PH and RNase R (Li et al., 1998; Cairrão et al., 2003). Like tRNA, tmRNA terminates with a CCA-3' sequence. The CCA-3' sequence of tmRNA in some eubacteria such as B. subtilis is not encoded on the gene and is added enzymatically (Ushida et al., 1994). The gene for 2-piece tmRNA in Caulobacter crescentus or cyanobacteria is circularly permuted (Keiler et al., 2000; Williams, 2002). The gene for the 3’-amino acid acceptor piece, an intervening segment and the gene for the 5’-amino acid acceptor piece are aligned in the genome in the 5’ to 3’ direction. The permuted precursor might be processed into a mature 2-piece tmRNA with the help of RNase P and exonucleases. A similar strategy has recently been found in the processing of tRNA in Cyanidioschyzon merolae (Soma et al., 2007). The pseudoknot structures are involved in the maturation of tmRNA, although they are apart from the tRNA-like structure in the secondary structure (Wower et al., 2004). It has also been shown that SmpB, a tmRNA binding protein, enhances the maturation of tmRNA (Wower et al., 2004).

FUNCTION AS A TRNA Unlike most other aminoacyl-tRNA synthetases, alanyl-tRNA synthetase does not require the anticodon sequence upon recongnition of tRNA, and instead it recognizes the upper half structure of the L-shaped tRNAAla molecule, especially G-U base pair at the third position of the amino acid acceptor stem (McClain & Foss, 1988; Hou & Schimmel, 1988) and A at the fourth position from the 3' end (discriminator base) (Tamura et al., 1991). Thus a model RNA fragment consisting of only an acceptor stem and a TΨC -arm of tRNAAla can be aminoacylated with alanine (Francklyn & Schimmel, 1989). A G-U base pair and an adenine residue are found at the corresponding positions in tmRNA (Figure 2). As expected, tmRNA from E. coli or B. subtilis is aminoacylated with alanine by alanyl-tRNA synthetase in vitro (Komine et al., 1994; Ushida et al., 1994). The tRNAAla-like features are exclusively conserved among tmRNA from various sources, suggesting the significance of aminoacylation with alanine for the function of tmRNA. TmRNA associates with 70S ribosomes but not with the dissociated ribosomal subunits in vivo (Ushida et al., 1994; Tadaki et al., 1996; Komine et al., 1996) and in vitro (Himeno et al., 1997). A mutant tmRNA having an A-U base pair instead of the G-U base pair at the third position of the amino acid acceptor stem, which fails to accept alanine, does not bind to the

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70S ribosomes (Tadaki et al., 1996; Himeno et al., 1997), indicating that aminoacylation with alanine is responsible for binding of tmRNA to the ribosome. Since the secondary structure of tmRNA fulfills the minimum requirements for the formation of a ternary complex of aminoacyl-tRNA, EF-Tu and GTP, it is conceivable that alanyl-tmRNA enters the A-site of the ribosome as a ternary complex. However, the absence of tmRNA in the polysome fraction has questioned the involvement in the canonical protein synthesis (Tadaki et al., 1996).

Function as an mRNA Tu et al. (1995) have found a strange fraction upon purification of overexpressed mouse interleukin-6 (IL-6) from the extracts of E. coli cells. It contains a mixture of polypeptides in that an AANDENYALAA sequence is connected to the IL-6 sequences that are C-terminally truncated from random positions. The last 10 of this 11-amino acid sequence called tagpeptide is identical to the last 10 amino-acid sequence of a putative peptide encoded in the middle of the tmRNA gene. Since the tag-peptide addition depends on the presence of the tmRNA gene on the genome, the last 10 amino acids are likely to be encoded by the gene for tmRNA, although the first alanine is of an unassigned origin. An identical 11 amino-acid tag sequence appears at the C-terminus of λ cI repressor or cytochrome b-562 that is translated in vivo from mRNA desinged to lack a termination codon (Keiler et al., 1996). The possibility that tmRNA itself acts as an mRNA for the tag-peptide has been raised (Figure 1). Direct evidence for the function as an mRNA for the tag-peptide was obtained from in vitro translation experiments (Muto et al., 1996; Himeno et al., 1997). E. coli tmRNA faciltates an incorporation of tag-specific amino acids into the growing polypeptide in a stoichiometrical fashion, only when poly (U)-directed poly-phenylalanine synthesis occurs. The tag-specific amino acid incorporation is not facilitated by a single-point mutant tmRNA that fails to accept alanine. These results have revealed the dual function of tmRNA both as an mRNA and as a tRNA and that the mRNA function requires the tRNA function in addition to an additional protein synthesis.

Trans-translation Based on the dual function of tmRNA, the novel suquence of tag-peptide and accumulation of truncated proteins with a tag-peptide from truncated mRNA, a transtranslation model has been proposed (Figure 1) (Keiler et al., 1996; Atkins & Gesteland, 1996; Muto et al., 1996, 1998). In this model, when a ribosome is stalled on a truncated 3' terminal of mRNA, alanyl-tmRNA, instead of aminoacyl-tRNA, is recruited to the vacant Asite of the ribosome, followed by a peptidyl transfer from peptidyl-tRNA in the P-site to alanyl-tmRNA in the A-site. Then translocation replaces truncated mRNA by the tagencoding region of tmRNA on the decoding region of the small subunit to allow translation of tmRNA-encoded tag-peptide. As a consequence, a chimeric protein of the truncated peptide and the tag-peptide connected by alanine is released from the ribosome using the inframe stop codon, allowing the ribosome to recycle. During these processes, tmRNA plays a

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dual function both as a tRNA and as an mRNA, and therefore tmRNA can be categorized as a new type of RNA, termed tmRNA. It should be noted that the connector alanine is encoded nowhere but was once aminoacylated to tmRNA. Alanine aminoacylated to tmRNA is actually incorporated into the growing polypeptide in the poly (U)-dependent tag-peptide synthesis system in vitro (Nameki et al., 1999c), strongly supporting the trans-translation model. The model of trans-translation addresses the question as to why tmRNA is not detected in the polysome fraction (Tadaki et al., 1996). All the ribosomes but the last one on a truncated mRNA are released from tmRNA upon translocation, and thereafter the translation of the last 10 amino acid residues of the tagpeptide encoded by tmRNA occurs on a single ribosome.

EVOLUTIONAL IMPLICATION FOR THE UNIQUE RECOGNITION MANNER OF ALANYL-TRNA SYNTHETASE It is generally believed that tmRNA universally adopts alanine as an amino acid charged, because of the exclusive conservation of G-U at the third base-pair position in the acceptor stem and A at the discriminator base position (Williams, 2000; Zwieb et al., 1999). Interestingly, trans-translation still occurs in vitro even when mutations are introduced so that tmRNA is aminoacylated with histidine instead of alanine (Nameki et al., 1999c), indicating that aminoacylation of tmRNA rather than alanylation is required for trans-translation. At the birth of tmRNA, some aminoacyl-tRNA synthetase should have been selected as the enzyme that catalyzes aminoacylation of tmRNA. TmRNA has no apparent anticodon, whereas up to 17 of 20 aminoacyl-tRNA synthetases except for alanyl-, seryl- and leucyltRNA synthetases require the anticodon sequence for recognition of the cognate tRNA. Since seryl- and leucyl-tRNA synthetases may be less appropriate for recognition of tmRNA, since they require the characteristic tertiary structures of their corresponding tRNA (Himeno et al., 1990; Asahara et al., 1993,1994; Soma & Himeno, 1998). Because alanyl-tRNA synthetase is unique in that the recognition area is unusually biased to the upper half of the tRNA molecule (Figure 2) (McClain & Foss, 1988; Hou & Schimmel, 1988; Tamura et al., 1991), it should have been the best one to recognize tmRNA. Eubacterial alanyl-tRNA synthetase has undergone co-evolution with two different kinds of molecules, tRNAAla and tmRNA. This has obliged the recognition area of alanyl-tRNA synthetase to be concentrated in their common structure including the acceptor stem and the TΨC-arm, which are also involved in recognitions by RNase P and EF-Tu. This might be the reason why alanyl-tRNA synthetase adopts the unique recognition manner unusually biased to the acceptor stem of tRNA.

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Figure 3. Trans-translation mediated by tmRNA and its binding proteins. By virture of transtranslation, the ribosome is recycled and the truncated mRNA and the resulting tagged proteins are degraded. The requirement of S1 is still controversial.

TmRNA Binding Factors Several protein factors, such as EF-Tu, S1 and SmpB, interact with tmRNA in vivo and in vitro (Figure 3). EF-Tu•GTP binds to the tRNA-like structure of alanyl-tmRNA (Rudinger-Thirion et al., 1999; Barends et al., 2000). It is thus conceivable that an alanyl-tmRNA, just like an aminoacyl-tRNA, enters the A-site as a ternary complex with EF-Tu and GTP. However, some reports have shown that EF-Tu is not essential for trans-translation in vitro (Hallier et al., 2004; Shimizu & Ueda, 2006). Additional EF-Tu binding sites outside the tRNA-domain have also been suggested (Zvereva et al., 2001; Stepanov & Nyborg, 2003). S1 protein is a component of the small subunit of the ribosome, although some population of S1 seems to be apart from ribosomes in the cell (Subramanian, 1983). S1 is thought to help the association of mRNA having a weak SD-sequence with the ribosome (Tzareva et al., 1994; Tedin et al., 1997). E. coli S1 binds tmRNA with high affinity (Wower et al., 2000; Hanawa-Suetsugu et al., 2001). It has also been suggested that more than one S1 can bind to one molecule of tmRNA in vitro (Bordeau & Felden, 2002). E. coli S1 has six repeating homologous oligonucleotide/oligosaccharide binding folds (OB-folds), of which the second OB-fold is essential for binding to tmRNA (Okada et al., 2004). S1 cross-links to PK2, PK3 and the upstream region of the tag-encoding sequence of tmRNA (Wower et al., 2000) to induce a conformational change of tmRNA (Bordeau & Felden, 2002; Valle et al., 2003a; Gillet et al., 2006). Thus, it is presumed that S1 guides tmRNA to the ribosome,

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anchors the upstream region of the resume codon on the ribosome and destabilizes the surrounding structured region to precisely set the resume codon at the A site. However, the role of S1 in trans-translation is still controversial. S1 is absent in the low-G+C group of Gram-positive bacteria, and indeed tmRNA from B. subtilis that lacks S1 can facilitate transtranslation in E. coli that has S1 (Ito et al., 2002). Note that B. subtilis has an S1 homologue, although it has no interaction with E. coli tmRNA probably due to the lack of the second OBfold (Okada et al., 2004). Overproduction of the N-terminal fragment of E. coli S1 inhibits general translation but not trans-translation, also suggesting the lack of significance for trans-translation (McGinness & Sauer, 2004). It has been shown that S1 is not essential for trans-translation in vitro, at least for the early stage (Takada et al., 2007; Qi et al., 2007; Saguy et al., 2007). This is consistent with a cryo-electron microscopic map of the ribosomal small subunit in complex with S1, in which S1 binds to the mRNA exit tunnel formed by the junction of the head, platform and main body of the small subunit of the ribosome (Sengupta et al., 2001), rather than to the shoulder side where tmRNA initially binds for transtranslation (Valle et al., 2003a; Kaur et al., 2006). A report has shown that S1 is involved in a later stage of trans-translation (Saguy et al., 2007), while another report has shown that it is not involved throughout the trans-translation processes (Qi et al., 2007). SmpB (small protein B) has been identified in E. coli as a tmRNA binding protein (Karzai et al., 1999). SmpB is widely conserved among eubacteria and chloroplasts, but not in mitochondria (Andersen et al., 2006). The gene for SmpB from many but not all eubacteria is located immediately upstream of the tmRNA gene on the genome, although the E. coli SmpB gene has an independent promoter sequence. NMR studies have revealed that SmpB is comprised of an antiparallel ß barrel core with three alpha helices and C-terminal basic residues that are disordered in solution (Dong et al., 2002; Someya et al., 2003). The primary binding site of SmpB of biological significance is in the TΨC-arm and the D-loop equivalents in the tRNA-like domain (Hanawa-Suetsugu et al., 2002; Gutmann et al., 2003; Nameki et al., 2005; Bessho et al., 2007). It has also been reported that total three molecules of SmpB can bind around the tRNA-like domain (Wower et al., 2002). Another report has also shown three SmpB binding sites in tmRNA, one of which is located in the lower half of tmRNA including the coding region and its surrounding pseudoknots (Metzinger et al., 2005). As described later, SmpB has crucial roles in trans-translation both inside and outside the ribosome. The level of SmpB in the cell changes in concert with that of tmRNA, probably because the complex formation between them protects not only tmRNA from nucleases but also SmpB from proteases (Hallier et al., 2004; Hong et al., 2005; Sundermeier & Karzai, 2007). Several proteins, phosphoribosyl pyrophosphate synthetase, RNase R and YfbG as well as S1 were copurified with tmRNA and SmpB from the E. coli cells in that both tmRNA and SmpB were overexpressed (Karzai & Sauer, 2001). A genetic interaction between phosphoribosyl pyrophosphate synthetase and tmRNA has been suggested by an observation that the slow growth of a temperature-sensitive mutant of phosphoribosyl pyrophosphate synthetase is suppressed by tmRNA (Ando et al., 1996). RNase R, a 3’ to 5’ exoribonuclease, is involved in tmRNA-mediated degradation of non-stop mRNA (Oussenko et al., 2005; Richards et al., 2006; Mehta et al., 2006), maturation of tmRNA in E. coli (Cairrão et al., 2003) and in cell cycle-regulated degradation of tmRNA in Caulobacter crescentus (Hong et

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al., 2005). YfbG is a protein of an unidentified function with sequence similarities to methionine tRNA formyl transferase and to sugar dehydratases and epimerases. A recent proteomic study has identified several tmRNA-binding proteins from Streptomyces aureofaciens, which includes DNA dependent RNA polymerase, polyribonucleotide nucleotidyltransferase (PNPase), SmpB, EF-Tu and a few ribosomal proteins (Mikulík et al., 2008). E. coli tmRNA has affinity to tRNAAla (Gillet & Felden, 2001a). Since the first codon for the tag-peptide on tmRNA is GCA designating alanine, the possibility of the involvement of this interaction in the trans-translation mechanism has been raised (Gillet & Felden, 2001b).

Cell-free trans-translation Systems As described above, a cell-free trans-translation system coupled with poly (U)-dependent polyphenylalanine syntheisis has initially been established using E. coli crude cell extracts (Himeno et al., 1997). This has contributed not only to providing the trans-translation model with evidence (Himeno et al., 1997; Nameki et al., 1999a) but also to identifying the nucleotides on tmRNA required for efficient and precise trans-translation (Figure 2) (Nameki et al., 1999a; Lee et al., 2001). Later, several trans-translation systems using purified factors from E. coli (Shimizu & Ueda, 2002; Ivanova et al., 2004; Asano et al., 2005; Konno et al., 2007) or from T. thermophilus (Takada et al., 2007) have been developed. These cell-free trans-translation systems using purified factors have revealed the minimum requirement for the first few steps of trans-translation including the binding of tmRNA to the ribosome, peptidyl-transfer from peptidyl-tRNA to alanyl-tRNA and decoding of the first codon on tmRNA for the tag-peptide. In addition to an mRNA lacking a termination codon and general translation factors, EF-Tu, EF-G, ribosome and aminoacyltRNAs, only two trans-translation-specific factors, alanyl-tmRNA and SmpB, are sufficient (Figure 3). The efficiency of trans-translation can be evaluated either by monitoring the incorporation of radio active amino acid attached to tmRNA or tRNA into the growing peptide or by monitoring the chimera polypeptide using antibody raised against the tagpeptide. For example, two steps of trans-translation, binding of Ala-tmRNA to the ribosome to undergo a peptide transfer reaction (1st step) and setting of the resume alanine codon to the A-site to be translated (2nd step), can independently be evaluated by monitoring the synthesis of peptidyl-Ala-tmRNA from peptidyl-tRNA and Ala-tmRNA and the synthesis of peptidylAla-Ala-tRNAAla from peptidyl-Ala-tmRNA and Ala-tRNAAla, respectively (Takada et al., 2007; Konno et al., 2007). Using a cell-free T. thermophilus trans-translation system containing S1-free ribosomes, S1 has been shown to be required for neither of these two steps (Takada et al., 2007). Essentially the same conclusion has been obtained using the cell-free E. coli trans-translation systems containing S1-free ribosomes (Qi et al., 2007; Saguy et al., 2007). These in vitro trans-translation systems have greatly contributed to clarifying the molecular mechanism of trans-translation, described below.

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MULTIPLE FUNCTIONS OF SMPB OUSIDE THE RIBOSOME SmpB has been suggested to play multiple roles outside the ribosome by binding to the tRNA domain of tmRNA. SmpB enhances the maturation of tmRNA (Wower et al., 2004), enhances the aminoacylation of tmRNA (Barends et al., 2001; Hanawa-Suetsugu et al., 2002; Shimizu & Ueda, 2002), protects tmRNA from degradation in the cell (Hanawa-Suetsugu et al., 2002; Hong et al., 2005) and recruits tmRNA in complex with EF-Tu and GTP to the stalled ribosome (Karzai et al., 1999; Hanawa-Suetsugu et al., 2002). Thus a single-point mutation in the tRNA domain of tmRNA, which seriously affects the interaction with SmpB, affects the latter three functions of SmpB (Hanawa-Suetsugu et al., 2001, 2002). Likewise, a single-point mutation in SmpB, which affects the interaction with tmRNA, also affects the recruitment of tmRNA to the stalled ribosome (Dulebohn et al., 2006). SmpB is essential for trans-translation in vitro and in vivo but not for canonical translation (Karzai et al., 1999; Hanawa-Suetsugu et al., 2002). Although SmpB has diverse roles in trans-translation outside the ribosome, its roles inside the ribosome are more crucial.

The Role of SmpB upon Entrance of tmRNA to the Ribosome Since Ala-tmRNA forms a complex with SmpB, EF-Tu and GTP in vitro (Barends et al., 2001; Hanawa-Suetsugu et al., 2001), this quarternary complex is likely to enter the empty Asite of the stalled ribosome for trans-translation. It has been demonstrated in vitro that E. coli SmpB can bind to the ribosome in the absence of tmRNA and that a stalled ribosome to that SmpB is pre-bound can trigger trans-translation in vitro, leading to the proposal of an alternative pathway in that SmpB pre-binds the ribosome to recruit a ternary complex of AlatmRNA, EF-Tu and GTP (Hallier et al., 2004). However, the alternative pathway does not seem to be consistent with another observation that SmpB is fewer than the ribosomes in the cell (Moore & Sauer, 2006). SmpB would not be enough for all the ribosomes, consequently for the stalled ribosomes of emergency, in the cell. A recent report has shown that SmpB binds the stalled ribosome in the presence of tmRNA more tightly than it does in the absence of tmRNA and that SmpB is enriched in the stalled ribosome only in the presence of tmRNA, making the alternative pathway unlikely to occur in the cell (Sundermeier & Karzai, 2007). Total two SmpB molecules can bind to a 70S ribosome, one to the small subunit and the other to the large subunit (Ivanova et al., 2005; Hallier et al., 2006). SmpB footprints nucleotides in the vicinity of the P-site facing the E-site in the small subunit and below the L7/L12 stalk in the large subunit (Ivanova et al., 2005). This is in agreement with a cryoelectron microscopic map of the putative pre-accommodated state complex of T. thermophilus ribosome•Ala-tmRNA•SmpB•EF-Tu•GDP in the presence of kirromycin that prevents EF-Tu from leaving the ribosome after hydrolysis of GTP, in which two molecules of SmpB bind both of the 70S ribosome and the tRNA-like domain of tmRNA (Kaur et al., 2006). One SmpB binding to the TΨC-arm equivalent of tmRNA contacts with the A-site on the small subunit and the other binding to the D-loop equivalent of tmRNA contacts with the vicinity of the GTP associated center on the large subunit. The latter SmpB is not visible in a complex of ribosome•Ala-tmRNA•SmpB representing the accommodated state of tmRNA.

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Thus the following model has been proposed: two molecules of SmpB are required upon binding of Ala-tmRNA to the ribosome and one of them is released from the ribosome concomitant with the release of EF-Tu after hydrolysis of GTP, so that the 3’-terminal of tmRNA is oriented toward the peptidyl-transferase center. This is also supported by a lead (II) probing of tmRNA in a putative pre-accommodated state (Ivanova et al., 2007). However, the 1:1 stoichiometry of tmRNA to SmpB in the cell (Sundermeier & Karzai, 2007) as well as only one molecule of SmpB binding to the tRNA-like domain with high affinity (Gutmann et al., 2003; Nameki et al., 2005; Bessho et al., 2007) does not appear to be consistent with the involvement of the sencond SmpB in trans-translation. Further studies are required to assess the validity of the above hypothesis.

STRUCTURAL AND FUNCTIONAL MIMICRY OF THE LOWER HALF OF TRNA In a crystal structure of a fragment of the tRNA-like domain (TLD) in complex with SmpB, SmpB binds the TΨC-arm and D-loop equivalents inside the opened L-form structure of TLD (Figure 4A,B) (Gutmann et al., 2003; Bessho et al., 2007). If TLD is fixed on the amino acid acceptor stem and the TΨC-arm, SmpB can be superimposed on the anticodon stem and loop of tRNA, indicating that SmpB structurally mimics the anticodon arm. This raises the possibility that SmpB acts as the functional mimicry of the anticodon arm during the trans-translation processes (Haebel et al., 2004). The functional mimicry is suggested by some studies showing the occupation of SmpB in the tRNA binding sites. In a cryo-electron microscopic map, T. thermophilus SmpB occupies the A-site in the pre-accommodated or accommodated state (Kaur et al., 2006). A study of site-directed hydroxyl radical probing using Fe(II)-BABE shows that two molecules of E. coli SmpB occupy the A-site and the Psite and that each SmpB can be superimposed on the lower half of tRNA behaving in translation (Kurita et al., 2007).

The C-terminal Tail of SmpB According to NMR studies, the C-terminal 1/5 of SmpB (comprised of about 30 amino acid residues in E. coli) is disordered in solution (Dong et al., 2002; Someya et al., 2003). Although the functional significance of the C-terminal tail has been exemplified in vivo and in vitro by its truncation or mutation (Sundermeier et al., 2005; Jacob et al., 2005; Konno et al., 2007; Kurita et al., 2007), cryo-electron microscopic studies fail to identify its location in the ribosome due to the lack of resolution (Kaur et al., 2006; Gillet et al., 2006). Instead, a site-directed hydroxyl radical probing study has succeeded in identifying the locations of two molecules of SmpB including its C-terminal tail in the ribosome from E. coli (Kurita et al., 2007). They can be superimposed on the lower halves of tRNA molecules behaving in translation at the A-site and the P-site. The C-terminal residues of A-site SmpB are aligned along the mRNA path towards the downstream tunnel, while those of P-site SmpB are located almost exclusively around the region of the codon-anticodon interaction in the P-site.

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Figure 4. Proteins structurally mimicking tRNA. (A) Saccharomyces cerevisiae tRNAPhe (PDB: 1EHZ), (B) T. thermophilus SmpB•TLD (PDB: 2CZJ), (C) Thermus aquaticus tRNAPhe•EF-Tu•GTP (PDB: 2EFG), (D) T. thermophilus EF-G•GDP (PDB: 2EFG), (E) Thermotoga maritima RRF (PDB: 1DD5) and (F) T. thermophilus EF-P (PDB: 1UEB) are shown. RNA and protein are designated in blue.

Figure 5. A model of the early stage of trans-translation based on a directed hydroxyl radical probing study. Upon or before entrance of tmRNA to the stalled ribosome, the C-terminal tail of SmpB may recognize the vacant A-site free of mRNA to lie along the mRNA path towards the downstream tunnel. After peptidyl-transfer to tmRNA, translocation of peptidyl-tmRNA/SmpB from the A-site to the P-site, possibly with the help of EF-G, occurs to drive out mRNA from the ribosome. During this event, the extended C-terminal tail somehow folds to substitute for the codon-anticodon interaction in the P-site. Then the resume codon of tmRNA is accommodated in the decoding region. SmpB and the tagencoding region are shown by red and blue, respectively.

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The probing signals appear at interval of 3 residues of the latter half of the C-terminal tail of A-site SmpB, suggesting the alpha helix structure, which has been predicted from the periodical occurrence of positively charged residues (Jacob et al., 2005).

A NEW MODEL OF TRANS-TRANSLATION SmpB molecules bound to the A-site and P-site shown by site-directed hydroxyl radical probing appears to reflect the pre- and post-translocation steps of trans-translation, respectively. Consequently, we have proposed the following model (Figure 5) (Kurita et al., 2007). The C-terminal tail of SmpB mimics mRNA and/or the codon-anticodon interaction and the main body of SmpB mimics the lower half of tRNA both before translocation in the A-site and after translocation in the P-site, while the upper half of tRNA is mimicked by TLD. Upon entrance of tmRNA to the stalled ribosome, the C-terminal tail of SmpB may recognize the vacant A-site free of mRNA to trigger trans-translation. After peptidyl-transfer to Ala-tmRNA, translocation of peptidyl-Ala-tmRNA·SmpB from the A-site to the P-site may occur. During this event, the extended C-terminal tail folds around the region of the codonanticodon interaction in the P-site, which drives out mRNA from the P-site, and consequently from the ribosome. Just after the movement of SmpB to the P-site, the A-site becomes free so that the resume codon of tmRNA can be accommodated. In this model, the truncated mRNA is pushed from the P-site to the E-site by the C-terminal tail of SmpB rather than by tmRNA, leading to the spontaneous dissociation from the E-site. This model can provide several insights into the yet-unidentified mechanism of transtranslation; how the stalled ribosome free of tRNA and mRNA is preferentially recognized, and what substitutes for a codon-anticodon interaction during trans-translation. It is quite reasonable that tmRNA preferentially recognizes the stalled ribosome free of tRNA and mRNA in the A-site. TLD and SmpB mimic the upper and lower halves of a tRNA molecule, respectively. TLD together with SmpB mimics an entire tRNA molecule. The C-terminal tail of SmpB mimics mRNA and/or the codon-anticodon interaction both before and after translocation, during which it undergoes a drastic conformational change. Consequently, SmpB•TLD behaves as mRNA+tRNA so that SmpB•tmRNA might decode the first alanine encoded by the C-terminal tail of SmpB. It has been shown that trans-translation can occur in the middle of a synthetic mRNA in vitro (Ivanova et al., 2004; Asano et al., 2005). The efficiency of trans-translation is inversely correlated with the length of the 3’-extension from the decoding region. This might be a result of competition in the mRNA path between the 3’-extension of mRNA and the Cterminal of A-site SmpB. Messenger RNA of a short 3’-extension occupies the mRNA path in the downstream of the decoding center with lower affinity than that of a long 3’-extension, and thereby tmRNA•SmpB prefers the stalled ribosome with a shorter 3’-extension.

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A NEW TYPE OF MOLECULAR MIMICRY The concept of “molecular mimicry” emerged, when the structure of EF-Tu in complex with Phe-tRNAPhe was found to be very much similar to that of EF-G (Figure 4C,D) (Nissen et al., 1995). The structural mimicry of the tRNA portion by a part of EF-G might reflect the common structure required for the entrance to the A-site. A ternary complex of aa-tRNA•EFTu•GTP initially binds the vacant A-site in the A/T hybrid state, and subsequent GTP hydrolysis at the GTPase-associated center in the large subunit promotes the dissociation of EF-Tu•GDP to allow aa-tRNA to be accommodated in the A/A state. EF-G•GTP binds to the A-site occupied by a peptidyl-tRNA to move it from the A/P state to the P/P state, and subsequent GTP hydrolysis at the same GTPase-associated center promotes the dissociation of EF-G•GDP from the ribosome. The concept of “molecular mimicry” has been extended to other translation factors. Since the structure of ribosome recycling factor (RRF) is very much similar to the L-shape of tRNA (Figure 4E) (Selmer et al., 1999), it has been predicted to enter the vacant A-site of the posttermination complex by mimicking tRNA to move deacylated tRNA from the P-site to the Esite. However, the structure of RRF in complex with the ribosome, which has been revealed by hydroxyl radical probing, cryo-electron microscopic and X-ray crystallographic studies (Lancaster et al., 2002; Agrawal et al., 2004; Wilson et al., 2005), does not support the functional aspect of “molecular mimicry”. In this complex structure, RRF binds the ribosome in an orientation dissimilar from that of tRNA relative to the ribosome: the longer arm of RRF is oriented to the P-site when the shorter arm is around the decoding region. EF-G moves RRF from the initial binding site to the P-site to dissociate the small subunit from the post-termination complex, although the mode of interaction of RRF with the P-site is different from that of tRNA in the P/P or P/E states (Barat et al., 2007). Release factor, RF1 or RF2, has been predicted to enter the vacant A-site by mimicking aminoacyl-tRNA (Nakamura et al., 1996), whereas the crystal structures of release factors are far from similar to the L-shape of tRNA (Vestergaard et al., 2001). Cryo-electron microscopic studies have suggested that RF1 or RF2 undergoes an extensive conformational change upon entrance to the ribosome, allowing the distance between two functional domains of a release factor, the GGQ domain that functions as the catalytic site for hydrolysis of peptide-tRNA in the P-site and the SPF domain that acts as an anticodon for the termination codon, quite similar to that between two functional domains of tRNA, the CCA-3’ end and the anticodon (Rawat et al., 2003; Vestergaard et al., 2005). This could be categorized into a kind of the functional mimicry rather than the typical structural mimicry. As far as we know, EF-P is the protein of that the structure is the most similar to the L-shape structure of tRNA (Figure 4F) (HanawaSuetsugu et al., 2004), although its function remains equivocal (Ganoza et al., 2002). TmRNA itself functionally and structurally mimics both tRNA and mRNA. These two different kinds of molecular mimicries are in collaboration with each other for transtranslation. In this case, a functional RNA (tRNA or mRNA) is mimicked by an RNA molecule rather than by a protein. The IRES（internal ribosomal entry site）5’-upstream of the coding sequence of Dicistroviridae virus RNA has a base pair with the immediately preceding the initiation codon at the P-site of the ribosome (Costantino et al., 2008). This

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structure apparently mimics a set of tRNA and mRNA with the codon-anticodon interaction for the 5’-cap-independent initiation of translation. This RNA is similar to tmRNA in that an RNA molecule acts both as tRNA and mRNA and that two functions are in collaboration with each other. The above trans-translation model provides a new concept of molecular mimicry. A protein molecule mimics both tRNA and mRNA. A wide variety of functional meanings of the tRNA and mRNA mimicry by SmpB•TLD are considered: (1) a function as the structural unit for the entrance to the A-site of the ribosome, (2) a function as the structural unit from the A-site to the P-site during translocation, (3) a function as the mRNA for the first alanine residue of the tag-peptide, (4) a function to find the target ribosome lacking mRNA downstream of the decoding region, (5) a function as the codon-anticodon interaction and etc. It has been reported that SmpB with a TLD fragment facilitates polyalanine synthesis in vitro without temperate mRNA (Shimizu & Ueda, 2006), functionally supporting the molecular mimicry of a set of tRNA and mRNA by SmpB•TLD. Unlike other proteins mimicking tRNA listed above, SmpB mimics not only the shape of tRNA but also the tRNA-like movement from the A-site to the P-site and presumably to the E-site in the ribosome.

TRANS-TRANSLATION PROCESSES IN THE RIBOSOME The functional mimicry by SmpB•TLD assumes the same behavior of SmpB•TLD as that of canonical tRNA+mRNA in the ribosome through several hybrid states, A/T, A/A, A/P, P/P and P/E. Although it is still controversial as already stated, the pre-accommodated state of SmpB•TLD in trans-translation could be in a somewhat different situation from the canonical A/T hybrid state; the second SmpB is required (Hallier et al., 2006; Kaur et al., 2006) and EF-Tu is not essential (Hallier et al., 2004; Shimizu & Ueda, 2006). In the canonical translation, EF-Tu contributes to proofreading to eliminate incorrect tRNA by recognizing a near-cognate codon-anticodon interaction (Valle et al., 2003b). This process might not be involved in the initial step of trans-translation in that no codon-anticodon interaction is assumed. A cryo-electron microscopic map has shown the location of the complex of tmRNA with the main body of SmpB in the A/A state (Kaur et al., 2006), and a directed hydroxy radical probing has revealed the positions of SmpB probably in the A/A and P/P states (Kurita et al., 2007). How the movement of the tRNA-like domain of tmRNA from the A/A state to the A/P state modulates the conformation and positioning of SmpB remains unknown. The existence of stable SmpB binding sites in the A-site and the P-site suggests the requirement of translocation, like in the canonical translation. It might possibly involve EFG•GTP, although the thermodynamic stability of the A/P state is yet unknown. Perhaps concomitantly with translocation, mRNA should be released from the ribosome. In fact, the release of mRNA as well as of deacylated tRNA is stimulated by EF-G•GTP (Ivanova et al., 2005). The next translocation is thought to move SmpB•TLD to the E-site, followed by the release from the ribosome. These ribosomal processes should involve extensive changes in the conformation of tmRNA (Wower et al., 2005) as well as in the modes of interactions of tmRNA with SmpB and the ribosome (Shpanchenko et al., 2005; Ivanova et al., 2007). According to a chemical probing study, SmpB remains bound to tmRNA at least in the initial

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few steps of trans-translation (Ivanova et al., 2007), although the SmpB binding site has not yet been found around the E-site. Another study has suggested the 1:1 stoichiometry of tmRNA to SmpB from the initiation to the termination of translation of the tag-peptide (Bugaeva et al., 2008).

Determination of the Resume Codon The coding region for the tag-peptide starts from the position about ten nucleotides downstream of PK1. In E. coli, it starts with G at position 90, 12 nucleotides downstream of PK1. The absence of an SD-like squence in the upstream region as well as of an apparent codon-anticodon interaction before the first translocation event of trans-translation makes the mechanism by which tmRNA resumes translation from the first codon mysterious. It should require an elaborate coordination between the two functional domains apparently distant from each other on the secondary structure (Hanawa-Suetsugu et al., 2001). It seems reasonable to assume that some structural unit on tmRNA fits elsewhere on the ribosome to set the first codon on the decoding center after the first translocation event. Indeed, PK1 is important for efficiency of trans-translation (Nameki et al., 1999a), whereas changing the span between PK1 and the coding region does not affect the start point of tag-translation, indicating the lack of contribution of PK1 to the determination of the initiation point of the tag-encoding sequence (Lee et al., 2001). A genetic selection of molecules active in trans-translation from a pool of E. coli tmRNAs having randomized sequences around the tag-initiation point has revealed strong base preference in the single-stranded region between PK1 and the tag-encoding region, especially at positions -4 and +1 (position 90) (Williams et al., 1999). The importance of this region for trans-translation has also been shown by an in vitro study (Lee et al., 2001). Some point mutations in the upstream sequence encompassing –6 to -1 decrease the efficiency of tag-translation, while some of them shift the tag-initiation point by –1 or +1 to a considerable extent (Figure 2) (Lee et al., 2001; Konno et al., 2007), indicating that the upstream sequence contains not only the enhancer of trans-translation but also the determinant for the taginitiation point. Thus the importance of an interaction of the upstream sequence with the ribosome either directly or via a trans-acting factor in determination of the tag-initiation point should be focused. The involvements of RF-1 and S1 as the trans-acting factor have been proposed based on the phylogenetically high conservation of –4 to –2 as UAA or UAG (Williams et al., 1999) and the cross-linking of uridine residue at position –5 to S1 (Wower et al., 2000), respectively. However, neither RF-1 nor S1 is essential for trans-translation in vitro (Takada et al., 2007; Qi et al., 2007; Saguy et al., 2007). A potential interaction between the upstream sequence and the putative anti-downstream box sequence on the decoding helix (44th helix) of the small subunit of the ribosome has been proposed based on the sequence complementarity (Muto et al., 1998), whereas it is not supported by a mutational study of the small subunit (O’Connor et al., 2000). Another proposal that the recognition of the triplet immediately preceding the resume codon by the decoding center is important for the entrance of tmRNA to the stalled ribosome (Lim & Garber, 2005) is not consistent with recent

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findings showing the occupation of A-site SmpB in the decoding center (Kaur et al., 2006; Kurita et al., 2007). Evidence for the interaction between the upstream region and SmpB has been obtained from a study of chemical modification (Konno et al., 2007). E. coli SmpB protects U at position –5 (85) from chemical modification. The main body of SmpB rather than the Cterminal tail is involved in this interaction. The protection at -5 was suppressed by a point mutation in the tRNA-like domain critical to SmpB binding, suggesting that SmpB serves to bridge two separate domains of tmRNA to determine the initial codon for tag-translation. Mutations that induce –1 and +1 shifts of the start point of tag-translation also shift the site of protection at -5 from chemical modification by –1 and +1, respectively (Konno et al., 2007), indicating the significance of the fixed span between the site of interaction on tmRNA with SmpB and the resume point of translation: translation of the tag-peptide starts from the position 5-nucleotides downstream of the site of interaction with SmpB (Figure 6). Thus the interaction between tmRNA and P-site SmpB as well as between the ribosome and P-site SmpB rather than that between tmRNA and the ribosome might be responsible for the determination of the resume codon on tmRNA. The interaction between the position at –5 from the intiation point of tag-translation and P-site SmpB would possibly be a substitute for the codon-anticodon interaction between intiator methionine tRNA and the initiation codon in the P-site. The initiation shift of tag-translation can also be induced by the addition of 4,5- or 4,6disubstituted class of aminoglycosides (Takahashi et al., 2003; Konno et al., 2004) that usually cause miscoding of translation by binding to the decoding center to induce a conformational change in its surroundings (Carter et al., 2000). This suggests the importance of the interaction of the decoding center with any portion of SmpB or tmRNA for precise tagtranslation.

Figure 6. The mechanism of the determination of the resume point of translation. The translation of the tag-peptide starts from the position 5-nucleotides downstream of the site of interaction with SmpB. Mutations that induce –1 and +1 shifts of the start point of tag-translation also shift the site of by –1 and +1, respectively. The tag-encoding sequence is designated in blue. The resuming point of translation is shown by red arrowhead. The interaction of SmpB and tmRNA is highlighted by red square.

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Degradation of Truncated mRNA Several kinds of endoribonucleases and exoribonucleases participate in the mRNA decay in the eubacterial cell (Condon et al., 2007). The ribosome translating on a truncated mRNA produced by an endonucleolytic cleavage would stall at and thus mask its 3’-end. It has been shown that trans-translation promotes decay of truncated mRNA in E. coli cell (Yamamoto et al., 2003; Sunohara et al., 2004a,b). Trans-translation would remove the stalled ribosome from truncated mRNA to allow the access of 3’ to 5’ exonucleases to its 3’-end. This had shed light on the significance of trans-translation as the quality control system of mRNA. It is a crucial role of trans-translation to prevent accumulation of aberrant mRNAs in the cell. Mutation introduced to the stem-loop structure of E. coli tmRNA including the last 1/3 of the tag-encoding sequence with the termination codon affects tmRNA-mediated decay of truncated mRNA without affecting the trans-translation activity (Mehta et al., 2006). This indicates that trans-translation is not enough for tmRNA-mediated decay of truncated mRNA, even though it unveils the 3’-end of truncated mRNA. Some kind of exoribonuclease associated with this stem-loop structure of tmRNA might somehow access the 3’-end of truncated mRNA on the ribosome before the dissociation of mRNA from the ribosome. RNase R, a 3’ to 5’ exoribonuclease, has been shown to be involved in tmRNA-mediated decay of truncated mRNA in B. subtilis and E. coli (Oussenko et al., 2005; Richards et al., 2006). Indeed, RNase R has been copurified with a complex of tmRNA and SmpB from E. coli cell (Karzai & Sauer, 2001).

Targets of Trans-translation Trans-translation occurs on stalled ribosomes at the 3’-end of mRNA, when mRNA has no stop codon (Keiler et al., 1996) or when the normal termination codon is read through by a nonsense suppressor tRNA (Ueda et al., 2002) or a miscoding drug (Abo et al., 2002). This system also operates upon translational pausings generated by rare codons (Roche & Sauer, 1999), inefficient termination codons (Roche & Sauer, 2001; Hayes & Sauer, 2002; Sunohara et al., 2002), programmed stalling sequences (Nakatogawa & Ito, 2002) and limited tRNA or release factors (Ivanova et al., 2004; Asano et al., 2005; Li et al., 2007). Efficiency of transtranslation is enhanced, when a proline codon or a minor arginine codon is present just preceding the termination codon (Hayes et al., 2002a,b). Eubacterial mRNAs rapidly turnover in the cells. Since mRNA degradation occurs by endonucleolytic cleavage followed by exonucleolytic degradation, most mRNAs might once be turned into a truncated mRNA lacking a stop codon, which can be a target of transtranslation. On the other hand, analyses of trans-translation products using a tmRNA variant with an undegradable tag-peptide sequence, terminating with a DD sequence instead of the AA sequence (tmRNA-DD), revealed that trans-translation preferentially occurs at specific sites of specific mRNAs. Six E. coli mRNAs (LacI, RbsK, GalE, YbeL, PhoP and RpsG) (Roche & Sauer, 2001; Collier et al, 2002) and eight B.subtilis mRNAs (TreP, PerR, EF-Tu, FolA, GsiG, YqaP, YtoQ and YloN) (Fujihara et al., 2002) have so far been identified as tmRNA substrates. These are no common substrates between the two species. Another recent

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proteomic study has identified at least 73 trans-translation products in C. crescentus (Hong et al., 2007). Only EF-Tu was common in two of three species. Thus most target proteins seem to be different in different species. The target proteins within a cell are also variable depending on the culture conditions (Fujihara et al., 2002). In C. crescentus, proteins involved in DNA replication, recombination and repair are overpresented among the target proteins identified.

Does Trans-translation Occur when Translation Stalled in the Middle of mRNA? Classically, tmRNA has been assumed to target a ribosome stalled at the 3’-end of mRNA. Later, it has been shown that trans-translation also occurs in the middle of mRNA, for example that at a tandem rare arginine (AGA) codons (Roche & Sauer, 1999) or at an inefficient termination codon (Roche & Sauer, 2001; Hayes & Sauer, 2002; Sunohara et al., 2002). This raises the question as to whether tmRNA actually targets the ribosome stalled in the middle of mRNA prior to cleavage. The apparent absence of accumulation of truncated mRNA in the cell suggests that trans-translation can occur in the middle of mRNA without cleavage of mRNA (Roche & Sauer, 1999). However, accumulation of truncated mRNA has been successfully detected in a tmRNA-deleted cell, arguing against this possibility (Li et al., 2006). Indeed, trans-translation by tmRNA promotes the degradation of truncated mRNA, making the detection of truncated mRNA in the normal cell difficult (Yamamoto et al., 2003; Sunohara et al., 2004a,b). Trans-translation can occur in the middle of mRNA in vitro, although less efficiently than it does in the 3’-end of mRNA (Ivanova et al., 2004; Asano et al., 2005). The efficiency is dramatically decreased with increase of the length of the 3’-extension from the decoding region. It has been found that bacterial toxins such as RelE (Pedersen et al., 2003) or ChpAK/MazF (Christensen et al., 2003) cleave an mRNA specifically at the A-site in the stalled ribosome, providing a new concept that mRNA of stalled translation is targeted initially by such A-site-specific endoribonucleases and subsequently by tmRNA for transtranslation. Perhaps, tmRNA would be recruited to a ribosome stalled in the middle of mRNA, but preferentially or exclusively after the cleavage around the rare codon. Several recent studies have suggested that the ribosome pausing induces endonucleotytic cleavage of mRNA around the A-site (Li et al., 2007; Garza-Sánchez et al., 2008). Cleavage of mRNA at the A-site has been detected even in cells lacking several bacterial toxins and some cellular ribonucleases (Hayes & Sauer, 2003; Sunohara et al., 2004a,b), raising the possibility that the ribosome itself is directly involved in the cleavage of mRNA at the A-site. However, trans-translation occurs without cleavage of mRNA in vitro (Asano et al., 2005).

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Competition between Trans-translation and Translation Several in vivo and in vitro studies have indicated that competition occurs at a nonsense codon between trans-translation and the termination of translation (Collier et al., 2002; Ivanova et al., 2004; Asano et al., 2005; Li et al., 2007) or at a sense codon between transtranslation and the elongation of translation (Asano et al., 2005). It has been shown in vitro that trans-translation can occur at either the nonsense codon (Ivanova et al., 2004; Asano et al., 2005) or the sense codon (Asano et al., 2005) without cleavage of mRNA at the A-site, although a shorter 3’-extention of mRNA from the A-site is preferable. The preference of a shorter 3’-extention of mRNA suggests the competition between the 3’-extention of mRNA and alanyl-tmRNA•SmpB. Perhaps, the mRNA path encompassing the decoding region to the downstream tunnel required for interaction with alanyl-tmRNA SmpB might be sequestered extensively by mRNA with a long 3’-extension but only weakly by mRNA with a short 3’extension. This is consistent with a directed hydroxyl radical probing study showing that the C-terminal tail of SmpB occupies the mRNA path downstream of the decoding region probably upon entrance of tmRNA to the stalled ribosome (Kurita et al., 2007), described in a previous session. Collectively, aminoacyl-tRNAs, release factors, alanyl-tmRNA and A-site-specific ribonucleases would always compete with one another for the A-site in the ribosome stalled on the intact mRNA. Alanyl-tmRNA might not substantially win the competition before the cleavage of mRNA by an A-site-specific ribonuclease (Ivanova et al., 2004). The competition should significantly be influenced by the activities or levels of these molecules in the cell, which fluctuate with change in the physiological conditions. For example, bacterial toxins are activated by degradation of antitoxins upon starvation of amino acids (Pedersen et al., 2003; Christensen et al., 2003), tmRNA is induced under some stressful conditions in B. subtilis (Muto et al., 2000) and tmRNA is cell cycle-dependently induced and degarded in C. crescentus (Keiler & Shapiro, 2003a; Hong et al., 2005).

ALTERNATIVE PATHWAYS TO RESCUE A RIBOSOME PAUSING The eubacterial cell has an alternative mechanism to rescue the stalled ribosome; a peptidyl-tRNA is dropped-off from the stalled ribosome, and then it is hydrolyzed by peptidyl-tRNA hydrolase (Pth) (Das & Varshney, 2006). Unlike tmRNA or SmpB, Pth is essential and it is distributed not only among eubacteria but also among eukaryotes and archaebacteria. It has been shown that the slow growth phenotype of a temperature-sensitive mutant of Pth is suppressed by overexpression of tmRNA (Singh & Varshney, 2004), suggesting that a single stalled ribosome can be rescued both by trans-translation and by drop-off. In fact, both trans-translation product with a tag-sequence and drop-off product without a tag-sequence probably produced from the same truncated mRNA have been identified in a cell (Williams et al., 1999; Ito et al., 2002). Perhaps, the drop-off mechanism would allow the trans-translation mechanism to be nonessential in some eubacteria including E. coli. Drop-off is enhanced by RRF alone or RRF together with RF3 (Heurgue-Hamard et

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al., 1998; Herr et al., 2001; Gong et al., 2007). By contranst, high concentration of RRF inhibits the trans-translation by tmRNA in vitro (Asano et al., 2005). There might be a wide variety of ribosome stallings, which are caused by a rare codon, a downstream pseudokonot or stem-loop structure or a too strong interaction between the nascent peptide and the peptide tunnel. They might be rescued by trans-translation, drop-off or sometimes LepA-dependent back-translocation (Qin et al., 2006). Some kinds of ribosome stallings may be programed for regulation of gene expression, e.g., frameshifting or translational bypassing, which may be a result from a temporary drop-off and immediate resumption of translation at another site prior to hydrolysis by Pth (Herr et al., 2000). As in the case of drop-off, translational bypassing is also enhanced by RRF (Herr et al., 2001). E. coli tryptophanase (tna) operon is induced by tryptophan via the translation arrest of the leader peptide (TnaC) due to the inhibition of the hydrolysis of peptidyl-tRNA (TnaCtRNATrp) by RF2 (Yanofsky, 2007). This stalled ribosome is rescued slowly by RRF and RF3 leading to drop-off, whereas it is not rescued by tmRNA-mediated trans-translation in the presence of tryptophan (Gong et al., 2007). The ribosome is also stalled at an internal proline codon of E. coli secM mRNA, which up-regulates the translation of the downstream secA mRNA presumably by disrupting the secondary structure that sequesters the ribosome binding site for translation of secA mRNA (Muto et al., 2006). This translational arrest is caused by the inefficient peptidyl-transfer of Prolyl-tRNAPro in the A-site, which inhibits the entrance of Ala-tmRNA to the A-site and the A-site specific cleavage of mRNA (GarzaSánchez et al., 2006).

mRNA Surveillance Systems in Eukaryotes Because of the absence of tmRNA and SmpB, trans-translation is not believed to occur in eukaryotes with the exception of some chloroplasts or mitochondria. Thus the question arises as to how the stalled translation is rescued in eukaryotes. Recently, two novel systems termed non-stop mRNA decay (NSD) and no-go mRNA decay (NGD) have been revealed. When translation stops at the 3’-end of a truncated mRNA having no termination codon, Ski7p, a GTPase having sequence similarity to EF1A or eRF3, enters the empty A-site to promote dissociation of the stalled complex, thereafter allowing 3’-to-5’ degradation of truncated mRNA by exosome (NSD) (Frischmeyer et al., 2002; van Hoof et al., 2002). When translation stops in the middle of mRNA, Dom34p and Hps1p, eRF1 and eRF3 homologues, respectively, enters the empty A-site to trigger cleavage of mRNA at the A-site by an A-site specific endonuclease that has not been identified (NGD) (Doma & Parker, 2006; Clement & Lykke-Andersen, 2006). The resulting 5’-half of mRNA is degraded by exosome in the 3’ to 5’ direction, while the 3’-half is degraded by Xrn1p in the reversed direction. The stalled ribosome due to the presence of a stem-loop structure is a preferential target of NGD, and that due to the presence of a pseudoknot, rare codon or even a premature stop codon is also a target. Neither tmRNA nor SmpB homologue is involved in these eukaryotic mRNA surveillance systems. The stalled translation is rescued only by protein factors, but not by any RNA molecule. On the other hand, it has never been found that any release factor homologue

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is involved in the rescue of the stalled translation in eubacteria. Therefore, the eukaryotic and eubacterial systems appear to have no evolutionary relationship with each other. Many components of exosome, a multi-protein complex including ribonucleases and helicases, are conserved among eukaryotes and archaebacteria (Büttner et al., 2006), and some of them have weak sequence similarity to those included in degradosome in eubacteria (Symmons et al., 2002).

DEGRADATION OF TAGGED PROTEINS Several proteases including ClpXP, ClpAP, FtsH, Tsp and Lon are involved in degradation of tmRNA-tagged proteins in E. coli. ClpXP and ClpAP are typical cytoplasmic ATP-dependent proteases, consisting of ClpP peptidase and hexamers of ClpX and ClpA, respectively. ClpX or ClpA recognizes the C-terminal ALAA sequence of the tagged proteins to unfold them for degradation by ClpP (Gottesman et al., 1998). SspB, a ribosomeassociated protein, specifically binds to the N-terminal AAND sequence of the tag-peptide to increase the affinity of ClpX to the tagged proteins, and consequently ClpXP serves as the major degradation machinery of tagged proteins in the cytoplasm (Flynn et al., 2001; Lessner et al., 2007). Lon, another cytoplasmic ATP-dependent protease, is also involved in the degradation of tagged proteins under stressful conditions (Choy et al., 2007). FtsH, an ATPdependent hexameric protease, anchored to the cytoplasmic side of the inner membrane, degrades the tagged proteins in the inner membrane (Herman et al., 1998). Tsp is involved in the degradation of the tagged proteins in the periplasm (Keiler et al., 1996). Although the Cterminal AA sequence of the tag-peptide is highly conserved, it terminates with FA in Mycoplasma, which lacks ClpXP, ClpAP and Tsp (Karzai et al., 2000).

Biological Function The trans-translation system seems to be present in all eubacteria, suggesting that it emerged at a very early stage of the eubacterial evolution. The high sequence conservation of tmRNA, together with that of SmpB, suggests crucial roles of this system in the cell growth. In some eubacteria, such as Neisseria gonorrhoeae (Huang et al., 2000), Haemophilis influenzae (Akerley et al., 2002) and Mycoplasma genitalium (Hutchison et al., 1999), tmRNA is essential for growth. In Salmonella enterica and Yersinia pseudotuberculosiss, tmRNA-deficiency leads to the loss of pathogenesity, probably due to an inability of the cells to survive in macrophages (Bäumler et al., 1994; Julio et al., 2000; Okan et al., 2006). The growths of immP22, a and P22 hybrid phage, in E. coli and of P22 in S. enterica cells also require tmRNA (Withey & Friedman, 1999; Julio et al., 2000). Mu prophage is induced in E. coli in a tmRNA-dependent manner (Ranquet et al., 2001). Although with such instances, in most eubacteria including E. coli and B. subtilis, the growths under normal nutrient culture conditions are not seriously affected by the deletion of tmRNA. However, the requirement of tmRNA is increased under several stressful conditions. A high temperature decreases the growth rate of tmRNA-deleted mutant of E. coli (Oh &

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Apirion, 1991) or B. subtilis (Muto et al., 2000). The heat shock response is constitutively induced in a tmRNA-deleted mutant cell (Munavar et al., 2005), and the recovery from carbon starvation is slower in E. coli (Oh & Apirion, 1991) and in Yersinia pseudotuberclosis (Okan et al., 2006). The expression level of a stress-response sigma factor RpoS, which controls the expression of many genes for the survival of the cell under stressful conditions, is positively controled by tmRNA (Ranque & Gottesman, 2007). In B. subtilis, several stresses, such as ethanol, cadmium and low temperature (Muto et al., 2000; Shin & Price, 2007), also seriously affect the growth of the cells lacking tmRNA, and the requirement of tmRNA increases as the strength of stresses increases (Muto et al., 2000). The spore formation of B. subtilis caused by nutrient starvation is significantly impaired by tmRNAdeficiency (Abe et al., submitted). Moreover, the amount of tmRNA in B. subtilis cells increases with the increase of stresses (Muto et al., 2000). Concomitantly, total transtranslation products increase under stresses. Stresses might increase aberrant translation events in the cells, which can be rescued by trans-translation. This may explain why tmRNA genes are conserved among eubacterial species, as bacteria must have been exposed to various environmental stresses during evolution. Many other phenotypes of tmRNA-deletion cells are summarized in the review article of Keiler (2007). The stall of translation should decrease the amount of free ribosomes for a new round of translation to impair the overall translation, leading to a decrease of the amount of the normal translation products in the cell. In particular, the amount and/or function of the protein translated from a very low level of mRNA would be seriously affected. This would be the case especially under insufficient growth conditions, such as under stressful conditions. Most of the defective phenotypes by the depletion of tmRNA can be complemented by an introduction of tmRNA-DD or tmRNA-His6, which is active in trans-translation reaction but defective in proteolytic degradation of tagged proteins, into the cells. N. gonorrhoeae and H. influenzae can grow with tmRNA-DD instead of wild-type tmRNA. The defective growth of immP22 and other phages in the cell lacking tmRNA is also complemented by tmRNADD. Thus it is generally accepted that the primary function of tmRNA-mediated transtranslation is rescuing stalled ribosomes rather than degrading the trans-translation products (for reviews, Withey & Friedman, 2003; Moore & Sauer, 2007; Keiler, 2007). The degradation of the trans-translation products is not important for the growth of many eubacteria in normal nutrient medium. Perhaps, the accumulation of nonfunctional proteins would not be serious for the cell. However, upon starvation of amino acids, the supply of amino acids from tagged proteins should be critical for a new protein synthesis in the cell (Pedersen et al., 2003; Li et al., 2008). Toxins such as RelE would accelerate the supply of amino acids from the nascent polypeptides on the stalled ribosomes via trans-translation. Trans-translation is sometimes involved in the regulation of the gene expression such as in the stress-responsive events described above. Trans-translation occurs at specific sites of specific mRNAs and it occurs on different mRNAs with the difference of culture conditions (Fujihara et al., 2002), supporting this concept. Several other examples have also been exemplified. The tagging of E. coli LacI, a repressor protein of the lac operon, occurs near the C-terminal, and the cells lacking tmRNA exhibit slower induction of -galactosidase (Abo et al., 2000). Degradation of LacI mRNA via tmRNA-mediated trans-translation might accelerate the derepression of the lac operon. In C. crescentus, the cell cycle (Keiler &

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Shapiro, 2003a) and the initiation of DNA replication (Keiler & Shapiro, 2003b) are controled by trans-translation. Indeed, the proteins involved in DNA replication have been overpresented in a proteomic study (Hong et al., 2007). Among eight target proteins identified in B. subtilis, the genes for three proteins, TreP, YtoQ and FolA, contain cre sequences (catabolite responsive element) within the protein-encoding regions around the presumed tagging sites (Fujihara et al., 2002). The genes involved in the carbon catabolite repression in B. subtilis are negatively regulated by binding of a repressor protein CcpA to the cre sequence (Fujita et al., 1995), which leads to a transcriptional roadblock upstream of the authentic stop codon (Miwa et al., 2000). Indeed, trans-translation occurs on TreP mRNA truncated at about 8-9 bp upstream of the cre sequence (Ujiie et al., unpublished). In consideration of the limitation of the detection of regulatory protreins of a low expression level, trans-translation would be much more deeply involved in the regulation of the gene expression than we have expected.

CONCLUSION Translating ribosomes often stall during the canonical mRNA turnover, upon stressdependent degradation of mRNA or under some special contexts of the sequence or structure of mRNA. Sometimes, translation is programmed to stall at a specific site on mRNA. Although with alternative pathways, trans-translation is a main pathway in the eubacterial cell to rescue the stalled translation for recycling of the ribosomes and dagradation of the truncated mRNA. Based on a directed hydroxyl radical probing study, we have proposed a novel molecular mechanism of trans-translation. In this model, a collaboration of a hybrid RNA molecule of tRNA and mRNA (tmRNA) and a protein mimicking a set of tRNA and mRNA (SmpB) facilitates trans-translaion. Upon entrance to the stalled ribosome, alanylTLD of tmRNA mimicking the upper-half of aminoacyl-tRNA and the main body of SmpB recognizes the A-site free of tRNA, and the C-terminal of SmpB mimicking mRNA interacts with the decoding region and the downstream mRNA path free of mRNA to recognize the stalled ribosome. While several proteins including SmpB mimic tRNA or its portion, SmpB is the first protein that mimics mRNA. When the tRNA domain of tmRNA•the main body of SmpB moves from the A-site to the P-site and then to the E-site like tRNA, a truncated mRNA, the C-terminal tail of SmpB encoding the first alanine residue of the tag-peptide and tmRNA encoding the last 10 amino acid residues of the tag-peptide pass through the decoding region in order. Our model depicts the outline of the trans-translaion processes in the ribosome. However, not only many enigmas remain unsolved but also the model poses new questions. Does SmpB•TLD move from the P-site to the E-site, and then release from the ribosome like deacylated tRNA, although no such protein has been found? How do the intermolecular interactions between tmRNA and ribosome, between tmRNA and SmpB and between ribosome and SmpB as well as the intramolecular interactions within tmRNA and within SmpB change during the course of the trans-translaion processes? Whether is EF-G required for translocation of SmpB•TLD from the A-site to the P-site? If so, how does it promote

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translocation? How deeply is trans-translaion involved in the regulations of gene expressions in the cell? How had tmRNA, SmpB and the trans-translaion system appeared and evolved?

ACKNOWLEDGEMENTS Authors are grateful to all those who have been involved in this work. This work was supported by Grants-in-Aid for scientific research from the Ministry of Education, Science, Sports and Culture, Japan to H.H. and A.M., Grants-in-Aid for scientific research from the Japan Society for the Promotion of Science to H.H. and A.M. and the 21th COE program of Iwate University for D.K.

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Valle, M., Zavialov, A., Li, W., Stagg, S.M., Sengupta, J., Nielsen, R.C., Nissen, P., Harvey, S.C., Ehrenberg, M., & Frank, J. (2003b). Incorporation of aminoacyl-tRNA into the ribosome as seen by cryo-electron microscopy. Nat. Struct. Biol., 10, 899-906. van Hoof, A., Frischmeyer, P.A., Dietz, H.C., & Parker, R. (2002). Exosome-mediated recognition and degradation of mRNAs lacking a termination codon Science, 295, 22622264. Vestergaard, B., Van, L.B., Andersen, G.R., Nyborg, J., Buckingham, R.H., & Kjeldgaard, M. (2001). Bacterial polypeptide release factor is structurally distinct from eukaryotic eRF1. Mol Cell, 8, 1375-1382. Vestergaard, B., Sanyal, S., Roessle, M., Mora, L., Buckingham, R.H., Kastrup, J.S., Gajhede, M., Svergun, D.I., & Ehrenberg, M. (2005). The SAXS solution structure of RF1 differs from its crystal structure and is similar to its ribosome bound cryo-EM structure. Mol. Cell, 20, 929-938. Wasserman, K.M. (2002). Small RNAs in bacteria: Diverse regulators of gene expression in response to environmental changes. Cell, 109, 141-144. Williams, K.P. (2000). The tmRNA website. Nucleic Acids Res, 28, 168. Williams, K.P. (2002). Descent of a split RNA. Nucleic Acids Res, 30, 2025-2030. Williams, K.P., & Bartel, D.P. (1996). Phylogenetic analysis of tmRNA secondary structures. RNA, 2, 1306-1310. Williams, K.P., Martindale, K.A., & Bartel, D.P. (1999). Resuming translation on tmRNA: a unique mode of determining a reading frame. EMBO J., 18, 5423-5433. Wilson, D.N., Schluenzen, F., Harms, J.M., Yoshida, T., Ohkubo, T., Albrecht, R., Buerger, J., Kobayashi, Y., & Fucini, P. (2005). X-ray crystallography study on ribosome recycling: the mechanism of binding and action of RRF on the 50S ribosomal subunit. EMBO J., 24, 251-260. Withey, J.H., & Friedman, D.I. (1999). Analysis of the role of trans-translation in the requirement of tmRNA for immP22 growth in Eschericia coli. J. Bacteriol., 187, 21482157. Withey, J.H., & Friedman, D.I. (2003). A salvage pathway for protein structures: tmRNA and trans-translation. Annu. Rev. Microbiol., 57, 101-123. Wower, I.K., Zwieb, C.W., Guven, S.A., & Wower, J. (2000). Binding and cross-linking of tmRNA to ribosomal protein S1, on and off the Escherichia coli ribosome. EMBO J., 19, 6612-6621. Wower, J., Zwieb, C.W., Hoffman, D.W., & Wower, I.K. (2002). SmpB: a protein that binds to double-stranded segments in tmRNA and tRNA. Biochemistry, 41, 8826-8836. Wower, I.K., Zwieb, C., & Wower, J. (2004). Contributions of pseudoknots and protein SmpB to the structure and function of tmRNA in trans-translation. J. Biol. Chem. 279, 54202-54209. Wower, I.K., Zwieb, C., & Wower, J. (2005). Transfer-messenger RNA unfolds as it transits the ribosome. RNA, 11, 668-673. Yamamoto, Y., Sunohara, T., Jojima, K., Inada, T., & Aiba, H. (2003). SsrA-mediated transtranslation plays a role in mRNA quality control by facilitating degradation of truncated mRNAs. RNA, 9, 408-418.

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In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter V

MODIFICATION OF MRNA TRANSLATION INITIATION TO STIMULATE PROTEIN SYNTHESIS IN SEPSIS Thomas C. Vary* Department of Cellular and Molecular Physiology, The Pennsylvania State University College of Medicine, Hershey, PA 17033, USA

ABSTRACT Sepsis, the systemic manifestation to bacterial infection, induces profound alterations in whole-body protein metabolism. Nitrogen losses up to 17% of total body protein may be observed in septic patients despite aggressive nutritional support. Organ system dysfunction and, eventually, organ failure can result from the persistent loss protein in sepsis. Sustained muscle protein catabolism continues to complicate recovery in septic patients. This review will illuminate potential molecular mechanisms responsible for increasing mRNA translation initiation in striated muscle. Emphasis will be placed on the role of growth hormones and nutrients in promoting rates of protein synthesis during sepsis. In this regard, elevating amino acids and IGF-I both interact to maximally enhance rates of protein synthesis acutely during sepsis through an acceleration of the mRNA translation initiation. IGF-I appears unique in accelerating protein synthesis during sepsis as growth hormone appears to enhance mortality while muscle shows a general resistance to the anabolic actions of insulin. Like IGF-I, amino acids and leucine in particular stimulate mRNA translation initiation by targeting specific signal transduction pathways. The hastening of mRNA translation initiation most likely results from a stimulation of mammalian target of rapamycin (mTOR) acting through its downstream effector proteins to enhance assembly of eIF4G with eIF4E via 4E-BP1 and eIF4G phosphorylation and to increase S6K1 phosphorylation. The physiologic *

Corresponding Author: Dr. Thomas C. Vary. Department of Cellular and Molecular Physiology, Rm C4718, Penn State University College of Medicine, H166, 500 University Drive, Hershey, PA 17033, Telephone: 717-5315014. Fax: 717-531-7667; Email: [email protected]

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INTRODUCTION Sepsis represents the host’s response to systemic infection. Despite the availability of a new generation of antibiotics and intensive supportive care, the overall hospital mortality rate ranges from 25-40% in septic patients making sepsis the leading cause of death in noncoronary intensive care units [23, 59] and the 12th leading cause of death [60, 72] accounting for nearly 10% (215,000 deaths) of all deaths in The United States (1). Mortality, morbidity, and cost remain high, despite many advances in the care for critically ill patients with severe injury or sepsis. The duration and extent of the metabolic changes seen in response to critical surgical illness and intensive care treatments have become better characterized. Although some of the changes in body water and fat are modifiable, loss of large amounts of (functional) protein has been resistant to various strategies so far studied. A major complication contributing to the morbidity and mortality in septic patients is the development of diffuse tissue injury referred to as multiple organ system dysfunction (MODS). MODS can lead to the failure of major organ systems of the body to maintain homeostasis. The septic process spares no organ system of the body. The hallmark of host’s response to systemic infection is the dyshomeostasis in protein metabolism that manifests itself in a severe loss of urea nitrogen. The net effect of these alterations is an overall catabolic condition, which seriously compromises recovery. These alterations lead to a functional redistribution of nitrogen (amino acids and proteins) and substrate metabolism among wounded tissues and major body organs. The redistribution of amino acids and proteins results in a quantitative reordering of the usual pathways of carbon and nitrogen flow within and among tissues of the body with resultant depletion of structural and functional proteins responsible for maintaining cellular homeostasis in important organs. Nitrogen losses up to 17% of total body protein may be observed in septic patients despite aggressive nutritional support (see for example [73]). Because skeletal muscle comprises approximately 45% of body weight, whole-body nitrogen balance reflects changes in protein turnover in muscle during sepsis. As such, approximately 70% of the septic-induced whole-body protein loss comes from erosion of skeletal muscle [73]. The persistent loss of large amounts of protein in sepsis leads to organ system dysfunction and, eventually, organ failure [46]. The clinical implications of continued loss of skeletal muscle protein in septic patients include poor wound healing [13], loss of muscle strength, diminished muscle activity [9, 11, 115], and, if severe enough, death [46]. Muscle weakness in septic patients contributes to a continued dependence on mechanical respirators, an increased risk of pneumonia, and the complications associated with extended periods of bed rest. These complications prolong hospitalization and convalescence, thereby increasing health-care costs [114]. Much effort has focused to modulate the overall amount of nutrients given to septic patients in a hope to improve efficiencies in utilization and nitrogen economies, rather than support of specific end-organ targets. Indeed, early enteral pharmaconutrition formula results in significantly

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faster recovery of organ function in medical patients with sepsis [7]. This review examines current understanding of the processes affected by sepsis and testable therapies (leucine and/or IGF-1) to circumvent the sepsis-induced defects in protein synthesis in skeletal muscle. In a healthy individual, nutrition involves an alternating system of feeding and fasting, with periods of fasting longer than 72 hr inducing a state of starvation. The hormonal response to nutrition is substrate controlled with glucose and amino acids, during the fed state, stimulating insulin secretion and decreasing glucagon secretion. Glycogen reserves and protein synthesis increase and the excess carbohydrate, amino acids and fats are stored as lipid. During the fasted state, plasma levels of glucose and amino acids fall, reducing insulin secretion and increasing glucagon secretion, stimulating gluconeogenesis and glycogenolysis. During injury and sepsis the nutritional hormones are no longer substrate controlled. In the septic state, in addition to the hormonal stress response, polypeptide mediators of tumor necrosis factor (TNF-alpha) and interleukin-1 are liberated, accelerating net skeletal muscle protein catabolism determined by the balance between protein synthesis and degradation. Emphasis will be placed on the role of selective nutrients in promoting rates of protein synthesis during the septic episode to limit net catbolism. In this regard, elevating amino acids or IGF-1 maximally enhance rates of protein synthesis acutely during sepsis through an acceleration of the mRNA translation initiation. IGF-I appears unique among peptide hormones in accelerating protein synthesis during sepsis as skeletal muscle shows a general resistance to the anabolic actions of insulin or growth hormone. Like IGF-I, amino acids stimulate mRNA translation initiation by targeting specific signal transduction pathways. For example, branched-chain amino acids (BCAA: leucine, isoleucine and valine) are not just structural constituents of proteins, but have ''pharmacologic'' properties that enhance protein synthesis through enhancing mRNA translation initiation. The hastening of mRNA translation initiation most likely results from a stimulation of mammalian target of rapamycin (mTOR) acting through its downstream effector proteins to enhance at the molecular level the assembly of eIF4G with eIF4E via 4E-BP1 and eIF4G phosphorylation and to increase S6K1 phosphorylation. The physiologic importance lies in the potential ability of IGF-I and amino acids as specific stimulators of mRNA translation designed to counteract the accelerated host protein wasting in septic patients and maintain muscle mass.

Skeletal Muscle Protein Metabolism Dyshomeostasis during Inflammation and Sepsis The protein wasting observed in skeletal muscle during sepsis results from an imbalance between rates of protein synthesis and protein degradation. In sepsis, an inhibition of protein synthesis and/or an acceleration of protein degradation leads to the loss of muscle protein during sepsis. The severity of the septic insult determines the relative contribution of slowing in protein synthesis and acceleration of proteolysis to the overall net catabolic state in muscle. Protein degradation varies, increasing as the septic episode worsens [10] and appears independent of the fiber composition of the muscle. In contrast, rates of protein synthesis are reduced to a similar extent regardless of the severity of the septic insult. Furthermore,

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inhibition of protein synthesis occurs preferentially in muscles composed of fast-twitch fibers [14, 51, 100]. Sepsis affects the overall process of protein synthesis. The synthesis of both myofibrillar and sarcoplasmic proteins is diminished to the same extent, indicating that some global mechanism controls protein synthesis during sepsis [106]. Indeed skeletal muscle mRNA content of myosin and actin are unaffected by the septic process. As the septic process wanes, muscle protein mass and protein synthesis return toward control values [43]. Non-septic trauma can also sometimes results in a decrease in protein synthesis and increase in proteolysis in skeletal muscle. In contrast to severe infection, changes in proteolysis and protein synthesis are short-lived in non-septic, trauma patients. With adequate nutritional support, restoration of positive nitrogen balance and lean body mass occurs within days of the injury. In agreement with studies in well-nourished non-septic trauma patients, skeletal muscle protein synthesis is not decreased by a wide-range of non-septic inflammatory insults [5, 39, 47, 50, 100, 108].

REGULATION OF PROTEIN SYNTHESIS Protein synthesis is a multistep, highly regulated process that includes cellular transport of amino acids, activation of protein factors, transcription, and translation. The process involves the association of the 40S and 60S ribosomal subunits, messenger RNA (mRNA), initiator methionyl-tRNA (met-tRNAimet), other amino acyl-tRNAs, cofactors (i.e. GTP; ATP), and protein factors, collectively known as eukaryotic initiation factors (eIF), elongation factors (eEF), and releasing factors (RF), through a series of discrete reactions that lead to translation of mRNA into proteins. Translation of mRNA is comprised of three phases: (a) initiation, whereby met-tRNAimet and mRNA bind to 40S ribosomal subunits, and subsequent binding of the 40S ribosomal subunit to the 60S subunit to form a ribosome complex capable of translation; (b) elongation, by which tRNA-bound amino acids are incorporated into growing polypeptide chains according to the mRNA template; and (c) termination, where the completed protein is released from the ribosome. Sepsis primarily affects the mRNA translation initiation step of protein synthesis. Impediments in mRNA translation initiation occur predominately through reductions in the abundance of ribosomes and/or translational efficiency. Sepsis does not diminish the relative abundance of ribosomes [14, 43, 100], but rather reduces the translational efficiency 50% in the skeletal muscle [14, 43, 82, 100]. The efficiency of translation, calculated by dividing the protein synthetic rates by the total RNA (or mRNA) content, provides an index of how rapidly the existing ribosomes synthesize protein. The decline in translational efficiency does not result from a decreased abundance of mRNA in muscles from septic rats [106]. Inhibition of the translational efficiency occurs by retarding either peptide-chain initiation whereas the rate of peptide-chain elongation is unaffected in skeletal muscle [100]. Therefore, inhibitory effects of sepsis on protein synthesis are related to derangements in mRNA translation initiation.

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Steps in Translation Initiation Inhibited during Sepsis In the first step, the 80S ribosome dissociates into 40S and 60S ribosomal subunits. In the second step, binding of met-tRNAimet to the 40S subunit forms the 43S preinitiation complex. The binding of met-tRNAimet to the 40S ribosomal subunit to form the 43S preinitiation complex is mediated by eukaryotic initiation factor 2 (eIF2) and is regulated by the activity of another eukaryotic initiation factor, eIF2B. In the third step, mRNA binds to the 43S preinitiation complex and from 48S preinitiation complex. The binding of mRNA to the 43S preinitiation complex is mediated by eIF4F. In the fourth step, the 60S ribosomal subunit associates with the 48S preinitiation complex leading to the formation of an 80S ribosome capable of carrying out the elongation phase of protein synthesis. The second and third steps in the process of mRNA translation initiation control the overall rate of protein synthesis (Figure 1). Sepsis inhibits both these regulatory steps involved in mRNA translation initiation [14, 43, 82, 98, 100, 109, 110].

REDUCED EIF2B ACTIVITY IN SKELETAL MUSCLE DURING SEPSIS Two mechanisms regulate the assembly of a 43S preinitiation complex catalyzed by eIF2. First, the amount of eIF2 in the cell may be altered. However, the cellular content of eIF2 is not significantly decreased in skeletal muscle from septic rats [97, 109]. Second, restraining recycling of GTP for GDP on eIF2 by lowered eIF2B activity restricts eIF2.GTP availability to form the ternary complex, thereby curbing translation initiation. Sepsis diminishes the activity of eIF2B in skeletal muscle where protein synthesis is inhibited [97, 98]. The best-characterized mechanism controlling eIF2B is through phosphorylation of eIF2 on its α subunit [eIF2(αP)] [54]. Phosphorylation of eIF2  converts the protein from a substrate into a competitive inhibitor of eIF2B [24, 54, 77], thereby limiting the ability of eIF2B to exchange GDP [24, 54, 77]. While this mechanism is important in regulating protein synthesis in other conditions, skeletal muscle from septic rats does not show elevations in eIF2α phosphorylation [97]. eIF2B is undergoes allosteric regulation by NAD+ and NADP+ [24, 45, 66, 79]. However, the NADPH/NADP+ concentration ratio is not significantly altered in gastrocnemius of septic rats [98]. Thus, diminished eIF2B activity does not relate to changes in the redox state during sepsis. Instead, the cellular content of eIF2B is reduced in chronic sepsis [109, 110]. However, eIF2B protein content is not increased in studies where protein synthesis in gastrocnemius from septic rats is acutely augmented [42, 44]. Thus, reduced eIF2B protein content can be overridden and, thus, other mechanisms may be involved.

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INCREASED PHOSPHORYLATION OF EIF2B DURING SEPSIS In addition to the above mechanisms for regulating eIF2B activity, the catalytic, εsubunit of eIF2B (eIF2Bε) undergoes reversible phosphorylation. An increase in the state of eIF2Bε phosphorylation (catalyzed by glycogen synthase kinase-3 (GSK-3)) inactivates the guanine nucleotide exchange activity [40, 111-113] Phosphorylation of eIF2Bε was significantly augmented over 2-fold and 2.5-fold after 3 and 5 days and returned to control after 10 days of sepsis when the septic process wanes. Sepsis augmented eIF2Bε phosphorylation by enhancing eIF2B kinase activity rather than reducing phosphatase activity [93]. Phosphorylation of glycogen synthase kinase-3 (GSK-3), a potential upstream kinase responsible for the elevated phosphorylation of eIF2Bε, was significantly reduced over 36 and 41% after 3 and 5 days and returned to control values after 10 days of sepsis. PKB has emerged as the most likely candidate to phosphorylate and inactivate GSK-3 [17]. PKB undergoes reversible phosphorylation with high levels of PKB activity associated with an increased GSK-3 phosphorylation. A disconnect between the phosphorylation of PKB and GSK-3 exists over the course of the septic episode. Initially, the extent of PKB phosphorylation becomes depressed during abscess formation stage, but beyond day 3 postinfection, the phosphorylation of PKB returns to basal values. Hence, reductions in PKB phosphorylation may explain the decrease in GSK-3 phosphorylation on day 3 postinfection. However, PKB phosphorylation is not altered in septic rats on day 5 postinfection, a time when GSK-3 phosphorylation remains diminished compared with sterile inflammatory rats. Treatment of septic rats with TNF-binding protein prevented the sepsis-induced changes in eIF2Bε and GSK-3 phosphorylation, implicating TNF in mediating the effects of sepsis. However infusion of TNF was without effects on eF2B phosphorylation [49]. The reasons for this dichotomy remain obscure. Thus increased phosphorylation of eIF2Bε via activation of GSK-3 is an important mechanism to account for the inhibition of skeletal muscle protein synthesis during sepsis. The dynamics of this signal transduction pathway over the course of the septic process requires additional explanation.

MRNA

RECRUITMENT TO RIBOSOME BY EIF4E IS IMPAIRED IN SKELETAL MUSCLE DURING SEPSIS

Another rate-controlling step in the process of peptide-chain initiation involves the recognition, unwinding and binding of mRNA to the 40S ribosomal subunit. The multisubunit complex of eukaryotic factors eIF4F catalyzes this step [81, 104]. eIF4F is composed of 1) eIF4A (a RNA helicase that functions with eIF4B to unwind secondary structure in 5’untranslated region of mRNA), 2) eIF4E (a protein that binds directly to the m7GTP cap structure present at the 5’-end of most eukaryotic mRNAs), and 3) eIF4G (a protein that functions as a scaffold for eIF4E, eIF4A and the mRNA and the ribosome). eIF4G appears to be the nucleus around which the initiation complex forms, because it has binding sites not only for eIF4E but also for eIF4A and eIF3 [48]. eIF4E is the least abundant initiation factor in most cells. eIF4E activity plays a critical role in determining global rates of mRNA

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translation, because essentially all mammalian mRNAs contain the m7GTP cap structure at their 5'-ends. eIF4E is regulated by alterations in either its availability or phosphorylation. Decreasing eIF4E content through transfection with anti-sense RNA causes an inhibition of protein synthesis at the level of translation initiation. However, the cellular content of eIF4E is not decreased in gastrocnemius of septic rats [99]. eIF4E is also regulated by reversible phosphorylation following activation of the MNK1 kinase. Phosphorylation of eIF4E enhances the affinity of the factor for m7GTP cap on mRNA and for eIF4G and eIF4A and correlates with enhanced rates of protein synthesis in cells in culture stimulated with mitogens, growth factors or serum, or transformed with ras or src oncogenes [61-63]. Conversely, reduced phosphorylation of eIF4E correlates with an inhibition of protein synthesis with serum depletion [26]. Sepsis does not modify the extent of eIF4E phosphorylation in skeletal muscle. Translation initiation may also be regulated through the formation of the eIF4E·eIF4G complex. A positive linear relationship between rates of protein synthesis and amount of eIF4G associated with eIF4E in muscle is observed in vivo [96]. Although this correlation does not prove cause and effect, the relationship between protein synthesis and amount of eIF4G associated with eIF4E is consistent with the proposed role of eIF4G·eIF4E complex in the overall regulation of protein synthesis. The assembly of the eIF4E·eIF4G complex is significantly diminished in skeletal muscle from septic rats [92, 94, 96, 99]. The diminished assembly of eIF4E·eIF4G complex is not the result of a reduced amount of eIF4G in the muscles from septic rats. Reduced amounts of eIF4E associated with eIF4G following chronic sepsis would be expected diminish the association of mRNA with the ribosome, and hence limit protein synthesis. The recruitment of the translational machinery to the 5’end of mRNA can be modulated by alterations in the phosphorylation of eIF4G. In support of this suggestion, Increased phosphorylation of eIF4G correlates with accelerated rates of protein synthesis in cell extracts [63]. Sepsis, but not sterile inflammation, diminishes phosphorylation of eIF4G(Ser1108), and are consistent with a role for reduced phosphorylation of eIF4G in mediating the diminution in formation of eIF4E·eIF4G complex during sepsis [94]. Phosphorylation of eIF4G is dependent upon the cytokine response. Infusion of tumor necrosis factor (TNF)- for 24 h in non-septic rats resulted in a 70% decrease in the phosphorylation of eIF4G. The biologic activity of TNF is modulated in vivo by the proteolytic shedding of the extracellular domain of the p55 and p75 TNF receptors. An increase in soluble TNF receptors in the bloodstream neutralizes circulating TNF , thereby lowering the biologically active concentration of TNF in the plasma. Treatment of septic animals with a specific TNF binding protein (TNFbp) completely attenuated the septicinduced decrease in eIF4G phosphorylation. Howevr, it is not possible to ascribe the metabolic effects of TNFbp on phosphorylation of eIF4G in gastrocnemius during hypermetabolic sepsis to a primary effect of TNF , or a secondary effect mediated through another cytokine or inflammatory mediator whose expression is dependent upon TNF . TNF is known to stimulate the secretion of other cytokines, including IL-1 and IL-6, as well as other inflammatory mediators. Furthermore, TNF often acts in synergy with other cytokines. TNFbp lowers plasma IL-1 following an E. coli or endotoxin challenge [75, 80, 88]. Consistent with a potential role of IL-1 in regulation protein metabolism, inhibition of

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IL-1 bioactivity with a specific IL-1 receptor antagonist (IL-1ra) abates the reduction in muscle loss and protein synthesis during sepsis through preventing the inhibition in peptidechain inhibition [16, 109]. Like TNFbp, IL-1ra reversed the sepsis-induced decrease in the phosphorylation of eIF4G. These results suggest a role of IL-1 in mediating the effects of sepsis on eIF4G phosphorylation. Thus, cytokines modulate skeletal muscle signal transduction pathways leading to phosphorylation of eIF4G. The anabolic and/or anticatabolic properties of branched-chain amino acids (leucine, valine, isoleucine)(BCAA) or their ketoacid derivatives have been known since the 1970's. This led to several clinical studies in the late 1970s and early 1980s aimed at evaluating the potential benefits of BCAA supplementation in nutritional support of the critically ill. The data on the efficacy of BCAAs in burn, trauma, and septic patients were far from definitive. In some studies the plasma concentration of leucine may not have raised to a sufficiently high enough concentration to evoke stimulate of protein synthesis [58]. Indeed, Cerra observed that the ability of BCAAs to lower nitrogen loss was proportional to the BCAA load [12]. In addition, many of these studies used very small numbers and there were problems with study design such as studying patients with an expected high mortality [89]. For example, while leucine is effective in stimulating protein synthesis whereas isoleucine and valine are much less efficacious [3, 56, 57]. In many studies, BCAA-supplemented nutrition very frequently consisted of virtually equivalent amounts of all three BCAAs. Moreover, several studies were performed without adequate basal nutritional support, which most probably hampered the correct metabolic utilization of these amino acids. More recently, acute oral leucine administration stimulates protein synthesis in an experimental model of sepsis [91]. BCAA supplementation in septic patients also demonstrated an improvement in patients' nutritional status and outcome [33, 34, 41, 64]. The binding of eIF4E with eIF4G occurs at a site that also binds the translation repressor protein 4E-BP1. Hence, the association of eIF4E with 4E-BP1 in an inactive eIF4E·4E-BP1 complex is thought to prevent binding to eIF4G and, hence limit translation initiation. The association of eIF4E with 4E-BP1 is regulated in part by the phosphorylation of 4E-BP1. Phosphorylation of 4E-BP1 lowers the affinity of 4E-BP1 for eIF4E and allows for the release of eIF4E from eIF4E·4E-BP1 complex and eIF4E reciprocal binding to eIF4G. This simplistic inverse relationship between formation of active eIF4E·eIF4G complex and release of eIF4E from inactive eIF4E·4EBP1 complex through phosphorylation of 4EBP1 represents an attractive hypothesis to account to changes observed. However, a disconnect exists in the proposed relationship between the formation of active eIF4E·eIF4G complex and inactive eIF4E·4EBP1 complex in response to infection or treatment with IGF-1/IGFBP-3 binary complex [82]. Despite the lack of effect on the abundance of inactive eIF4E·4EBP1 complex, the formation of active eIF4E·eIF4G was modulated. Likewise, infusion of TNF reduces the assembly of eIF4G·eIF4E complex by approximately 80% [49]. One mechanism to account for this change would be a decreased availability of eIF4E via sequestration in an inactive 4EBP1·eIF4E complex. However, the amount of 4EBP1 associated with eIF4E only rose by 60% indicating other factors may play a role in the assembly of eIF4G·eIF4E complex following infusion with TNF [94]. The availability of eIF4E for binding to eIF4G is regulated, in part, through the association of eIF4E with a family of translational repressor proteins (4E-BPs) and through

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phosphorylation of eIF4G. In skeletal muscle, 4E-BP1 is the predominant form of he 4E-BPs. When eIF4E is bound to 4E-BP1, eIF4E cannot bind to eIF4G. Consequently, the mRNA cannot bind to the ribosome [35], thereby inhibiting cap-dependent translation of mRNA by physically sequestering eIF4E into an inactive 4E-BP1·eIF4E complex. In muscles from control rats, phosphorylation of 4EBP-1 releases eIF4E from 4E-BP1·eIF4E complex and allows the eIF4E·mRNA complex to bind to eIF4G and, then to the 40S ribosome [55]. Unlike muscles from controls, increased phosphorylation of 4E-BP1 and decreased eIF4E·4E-BP1 complex formation is not associated with an enhanced formation of eIF4E·eIF4G complex in muscles from septic rats [96]. This observation indicates that sepsis alters the formation of eIF4E·eIF4G complex through mechanisms other than eIF4E availability.

MEAL FEEDING Feeding promotes protein accretion in skeletal muscle through a stimulation of the mRNA translation initiation phase of protein synthesis either secondary to nutrient-induced rises in insulin or owing to direct effects of the nutrients (amino acids including leucine) themselves. The initial rise in insulin causes stimulation of PI3-kinase-PKB signal pathway leading to enhanced mTOR phosphorylation and subsequent stimulation of phosphorylation of S6K1 and 4E-BP1 [102, 103]. However, stimulation of PKB is not maintained throughout the feeding period. Whereas PKB phosphorylation returns to baseline values, mTOR activation continues presumably because of the elevation of plasma amino acid concentrations. The branched chain amino acids are the most robust of the plasma amino acids in their ability to cause increased phosphorylation of mTOR [56, 57]. Meal feedinginduced activation of mTOR phosphorylation is maintained as long as the food is present. With removal of food, the assembly of eIF4G·eIF4E returns to levels observed prior to feeding. The reversal of the effects of meal feeding correlate with a fall in both plasma insulin and amino acid concentrations. Thus, acute leucine-induced stimulation of protein synthesis and the phosphorylation states of 4E-BP1 and S6K1 are facilitated by the transient increases in serum insulin concentrations [2, 4]. Hence, meal feeding stimulates assembly active eIF4F complex of through both insulin-dependent and -independent mechanisms involving phosphorylation of 4E-BP1, eIF4G and S6K1.

LEUCINE STIMULATES SKELETAL MUSCLE PROTEIN SYNTHESIS Amino acids and leucine in particular, serve not only as substrates for protein synthesis but regulators of protein synthesis as well. In healthy men is postabsorptive condition, infusing a solution containing mixed amino acids reverses whole-body protein balance from negative to positive and a major component of this is the increase in muscle protein synthesis [8]. Extracellular amino acid concentrations determine amino acid balance across peripheral

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tissues independently of non-protein calories, insulin and IGF-I [83]. The balance of globular proteins improved due to the stimulation of synthesis and attenuation of degradation across arm and leg tissues, despite insignificant uptake of tyrosine, tryptophan, and cysteine. Provision of amino acids did not attenuate the degradation of myofibrillar proteins. Neither insulin nor circulating IGF-I explained improved protein balance in skeletal muscles after elevation of plasma amino acids [84]. The effect of leucine to stimulate protein synthesis does not depend upon its metabolism [78]. Supraphysiological doses of amino acids stimulate protein synthesis through accelerating translation initiation in skeletal muscle [42, 95]. This is achieved by enhancing mRNA translation initiation independent of the activity of eIF2B, in the amount of eIF4E associated with the eIF4E-binding protein (4E-BP1), or in the phosphorylation of 4E-BP1. The order of potency of amino acids to stimulate this process was leucine > norleucine > threo-L-beta-hydroxyleucine approximately = Ile > Met approximately = Val. Other structural analogues of leucine, such as H-alpha-methyl-D/Lleucine, S-(-)-2-amino-4-pentenoic acid, and 3-amino-4-methylpentanoic acid, possessed only weak agonist activity [57]. The stimulatory effect of leucine on protein synthesis in skeletal muscle is unaffected by a specific inhibitor of PI3-kinase (LY 294002). Moreover, signaling through mTOR, as monitored by the phosphorylation status of 4E-BP1 or S6K1, was not further enhanced raising the leucine concentration [95]. However, binding of eIF4E to eIF4G and eIF4G(Ser1108) phosphorylation is enhanced when leucine concentration is elevated. Collectively, these observations illustrate an experimental model whereby leucine in the absence of other regulatory agents stimulates eIF4E·eIF4G assembly and protein synthesis directly in skeletal muscle, possibly by augmenting phosphorylation of eIF4G through a signaling pathway independent of mTOR. Hence formation of the active eIF4E·eIF4G complex controls protein synthesis in skeletal muscle when the amino acid concentration is raised above the physiological range. In using leucine or amino acids a nutritional supplement, the effects of chronic administration have not been well defined. Leucine or norleucine supplementation of drinking water for 12 days is accompanied by increased rates of protein synthesis in adipose tissue, liver, and skeletal muscle, but not in heart or kidney. Supplementation is not associated with increases in the anabolic hormones insulin or insulin-like growth factor I. Chronic supplementation did not cause apparent adaptation in either components of the mTOR cell-signaling pathway that respond to leucine (mTOR, ribosomal protein S6 kinase, and 4E-BP1) or the first two steps in leucine metabolism (the mitochondrial isoform of branched-chain amino acid transaminase, branched-chain keto acid dehydrogenase, and branched-chain keto acid dehydrogenase kinase), which may be involved in terminating the signal from leucine. These results suggest that provision of leucine or norleucine supplementation via the drinking water results in stimulation of postprandial protein synthesis in adipose tissue, skeletal muscle, and liver without notable adaptive changes in signaling proteins or metabolic enzymes. However, sepsis decreases plasma concentrations of valine, leucine, isoleucine, alanine, serine, glutamic acid, histidine, proline and glycine; while the concentrations of threonine, cysteine, the ratio of phenylalanine and tyrosine (Phe/Tyr) were elevated [29, 107]. Indeed, lowering the plasma leucine concentrations decreases the rate of protein synthesis in muscle. The inhibition of protein synthesis was associated with a 40% decrease in eIF2B activity and

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an 80% fall in the abundance of eIF4E·eIF4G complex. The fall in eIF4G binding to eIF4E was associated with increased 4E-BP1 bound to eIF4E and a reduced phosphorylation of 4EBP1. In contrast, the extent of phosphorylation of eIF4E was unaffected [95]. Hence, removal of leucine reduces protein synthesis through changes in both eIF2B and eIF4E. Based on the aforementioned studies, restoring and further elevating plasma amino acids represent one approach to augment protein synthesis in sepsis. Indeed, infusion of IL-1ra in septic patients elevates the plasma concentration of eleven amino acids and total amino acid concentration was increased by 50% within 70 hrs [15] and is associated with a lower nitrogen excretion. Concentrations of several amino acids including leucine were increased up to two-fold by infusion of IL-1ra. Hence IL-1ra may be a useful adjunct to promote muscle accretion during sepsis. Amino acids and leucine in particular stimulate mRNA translation initiation. Gavage with leucine stimulated protein synthesis and enhanced the assembly of the active eIF4G·eIF4E complex [91]. Increased assembly of the active eIF4GeIF4E complex was associated with a robust rise in phosphorylation of eIF4G(Ser1108) and a decreased assembly of inactive 4E-BP1-eIF4E complex in both sterile inflammatory and septic rats. The reduced assembly of 4E-BP1-eIF4E complexes was associated with an increase in phosphorylation of 4E-BP1 in the gamma-form following oral leucine gavage. Phosphorylation of 70-kDa ribosomal protein S6 kinase on Thr389 was also increased following oral leucine gavage, as well as the phosphorylation of mammalian target of rapamycin on Ser2448 or Ser2481. In contrast, phosphorylation of protein kinase B (PKB) on Thr308 or Ser473 was not augmented following oral leucine gavage in septic rats. Hence, leucine stimulates a PKB-independent signal pathway elevating the eIF4G-eIF4E complex assembly through increased phosphorylation of eIF4G and decreased association of 4E-BP1 with eIF4E in skeletal muscle during sepsis [91]. Likewise, amino acids stimulate skeletal muscle protein synthesis during acute endotoxemia via mTOR-dependent ribosomal assembly despite reduced basal protein synthesis rates in neonatal pigs [67]. There appears to be a threshold effect with regard to effects of branched chain amino acid effects on protein synthesis [18]. Phosphorylation of S6K1 is maximal at an oral dose of leucine that increases plasma leucine concentrations approximately threefold [58]. This may account for the failure of branched chain amino acids to augment protein synthesis in muscle during sepsis [38]. In contrast, changes in phosphorylation state of the branched chain -keto acid dehydrogenase (the enzyme responsible for oxidative decarboxylation of branched chain keto acids) require higher plasma leucine concentrations than that observed for activation of S6k1. The results seem more consistent with a role for BCKD and BCKD kinase in the activation of leucine metabolism/oxidation and removal of leucine signal than in the activation of the leucine signal to mTOR. Enteral leucine administration increases the formation of active eIF4G·eIF4E complex and activation of S6K1 pathway both of which are associated with accelerated rates of protein synthesis to values observed in rats with a sterile, non-septic abscess. Enteral leucine treatment modality appears to overcome the inhibition of protein synthetic process through acutely augmenting eIF4G·eIF4E complex formation during chronic sepsis. In summary, raising the plasma leucine concentration via oral administration of leucine to septic rats stimulated phosphorylation of both 4E-BP1 and eIF4G, maximizing the assembly of active eIF4G·eIF4E complex. The increase in formation of active eIF4G·eIF4E

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complex and activation of S6K1 pathway is associated with accelerated rates of protein synthesis. The enteral leucine treatment modality appears overcome the sepsis-induced inhibition of protein synthetic process through acutely augmenting eIF4G·eIF4E during chronic sepsis. Hence leucine appears to belong to a class of pharmaconutrients that act through modulating cell signaling, exert anabolic/anticatabolic functions when provided in sufficient amounts, with these effects occurring independently of the nutritional value of the supplement. This means that leucine supplementation should be given in addition to, and not as a replacement for, sufficient and balanced nutritional support [19]. How human muscle cells sense an increase in leucine and/or essential amino acids to activate mammalian target of rapamycin signaling is currently unknown.

INCREASING IGF-1 BIOAVILABILITY STIMULATES OF PROTEIN SYNTHESIS IN STRIATED MUSCLE DURING SEPSIS Protein synthesis in muscles from septic rats was unresponsive to stimulation by insulin [42, 96]. Insulin induced hyperphosphorylation of 4E-BP1 and of S6K1, two targets of insulin action on mRNA translation in gastrocnemius of septic rats. Hyperphosphorylation of 4E-BP1 in response to insulin resulted in its dissociation from the inactive eIF4E. 4E-BP1 complexes. However, assembly of the active eIF4F complex as assessed by the association of eIF4E with eIF4G did not follow the pattern predicted by the increased availability of eIF4E secondary to enhanced phosphorylation of 4E-BP1. Indeed, sepsis caused a dramatic reduction in the amount of eIF4G associated with eIF4E in the presence or absence of insulin [96]. Thus the inability of insulin to stimulate protein synthesis during sepsis may be related to a defect in signaling to a step in translation initiation involved in assembly of an active eIF4F complex. In contrast to adults, plasma levels of insulin are increased, and glucose and amino acids decreased, suggesting the absence of insulin resistance in neonatal pigs [70, 71]. Furthermore, high rates of neonatal muscle protein synthesis remain largely maintained as long as adequate substrate supply is present during sustained endotoxemia (20h) [68]. Maintenance of an anabolic response to the feeding-induced rise in insulin likely exerts a protective effect for the neonate to the catabolic processes induced by endotoxin [69]. Hence there may be fundamental differences in the response of skeletal muscle from neonates compared with adults. Aside from insulin, growth hormone promotes nitrogen retention and improves nitrogen balance in a variety of catabolic conditions. Septic patients have inappropriately high growth hormone concentrations indicative of a severe growth hormone resistance. Growth hormone has a reduced effectiveness to limit protein catabolism in septic patients [20, 74]. More problematic is the association of growth hormone administration with an increased morbidity and mortality in critically ill patients [85]. Therefore, growth hormone administration may have limited usefulness in the treatment of septic patients. IGF-I is believed to mediate the anabolic action of growth hormone in muscle. Consequently IGF-I may be of more importance than growth hormone in improving nitrogen balance in skeletal muscle during sepsis. IGF-I is a circulating hormone synthesized

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predominantly by the liver. It shares structural and functional similarities with insulin, and like insulin, IGF-I plays a distinct role in the regulation of whole body protein metabolism. More importantly IGH-1 preferentially stimulates protein synthesis in skeletal muscle [6]. Likewise, intravenous infusion of IGF-I directly increases protein synthesis in skeletal muscle provided plasma amino acid concentrations are maintained in humans [32, 76]. The importance of IGF-1 in muscle protein metabolism is shown by the positive linear relationship between IGF-I and protein synthesis in skeletal muscle [51]. IGF-1 accelerates protein synthesis by stimulating PI3 kinase and enhancing S6k1 phosphorylation [22]. In addition, IGF-I promotes assembly of the translationally active eukaryotic initiation factor (eIF)4G·eIF4E complex. The increased assembly of eIF4G·eIF4E is associated with an enhanced eIF4G phosphorylation and increased availability of eIF4E. Enhanced availability of eIF4E occurs as a consequence of diminished abundance of the inactive 4E-BP1·eIF4E complex following IGF-I administration. The assembly of the 4EBP1·eIF4E complexes diminishes through an IGF-I-induced phosphorylation of 4E-BP1. Activation of the potential upstream regulators of 4E-BP1 and S6K1 phosphorylation via PKB and mTOR is also observed. In contrast, IGF-I has no effect on phosphorylation of elongation factor 2. Thus, the major impact of IGF-I in striated muscle takes place via stimulation of translation initiation rather than elongation [101]. Furthermore, the assembly of active eIF4G·eIF4E complex and activation of S6K1 mediates the stimulation of mRNA translation initiation by IGF-I partially through a PKB/mTOR signaling pathway. However, the IGF-I effect was only partially inhibited by rapamycin, indicating a non-mTOR mediated pathway exists for IGF-I [22]. The mTOR-independent pathway remains obscure. Systemically administered IGF-I results in weight gain in normal rats, reduces weight loss in during starvation [65] or diabetes [87], and attenuates protein loss during glucocorticoid-induced cachetic states [21, 86], pediatric burn patients [36], chronic sepsis [44, 90], acute peritonitis [37], and endotoxin administration [25]. In contrast to insulin [105], the plasma concentration of IGF-I is depressed during endotoxemia [28], cytokine infusion [27] or following infusion of growth hormone in septic patients [20]. Based on the responsiveness of skeletal muscle to IGF-I in vitro and the decreased plasma concentrations of IGF-I during severe trauma [116] and sepsis, alterations in the bioavailability of IGF-I would be expected to have profound effects on rates of protein synthesis in skeletal muscles. Whereas, neutralization of muscle IGF-I by the addition of IGF-I antibody to the incubation medium reduced protein synthesis an average of 22%, rates of protein degradation were not affected [30]. The neutralizing effects of anti-IGF-1 on protein synthesis can be overcome by elevating amino acids [30]. The IGF-I system is composed of the hormone as well as at least six different IGF binding proteins (IGFBP). IGFBPs carry the IGF-I in the blood and modulate its bioavailability, thereby either inhibiting or potentiating the interaction of IGF-I with its receptor. In this regard a common feature of catabolic conditions including sepsis is an elevation in the plasma IGFBP-1 and IGFBP-2 concentration [27, 28, 51]. Indeed, IGF-BP-1 concentrations are elevated more than fivefold 2 h after LPS injection, and thereafter levels gradually returned toward baseline. IGFBP-2 concentration also increased after LPS injection, but the maximal increase (approximately 50% above basal) is observed during the final 2 h of the protocol. In contrast, IGFBP-3 levels do not vary over the period examined in

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response to LPS [52]. An increase in the plasma IGFBP-1 concentration would be expected to sequester IGF-I lowering the effective IGF-I concentration in plasma of septic rats. Acute in vivo elevation in IGFBP-1, of the magnitude observed in various catabolic conditions, is capable of inhibiting protein synthesis in fast-twitch skeletal muscle and up-regulating the hepatic and renal syntheses of IGFBP-1 per se [53]. Similarly, IGFBP-1 inhibits IGF-Imediated protein synthesis by binding to IGF-I in human skeletal muscle cells in culture [31]. IGFBP-1, acting independently of IGF-I, inhibits protein degradation. The IGF-independent response occurs via beta1 integrin binding and stimulation of a rapamycin-sensitive signal transduction pathway. Hence, elevations in circulating and tissue levels of IGFBP-1 may be an important mediator for the muscle catabolism observed in various stress conditions. Unlike IGFBP-1, IGFBP-3 is reduced in critically ill humans, but similar reductions in plasma IGFBP-3 are not observed in rodent models of sepsis. Infusion of IL-1ra completely prevented the rise in IGFBP-1, but was without effect on plasma IGFBP-2 or IGFBP-3 concentrations [51]. Inclusion of IGF-I (1 or 10 nM) in the isolated perfused hindlimb or incubated muscles stimulated protein synthesis in gastrocnemius of septic rats 2.5-fold, and restored rates of protein synthesis to those observed in control rats [44]. The stimulation of protein synthesis did not result from an increase in the RNA content, but was correlated with an increase in the translational efficiency. The enhanced translational efficiency was accompanied by a 33% and 55% decrease in the abundance of free 40S and 60S ribosomal subunits, respectively, indicating IGF-I accelerated peptide-chain initiation relative to elongation/termination. These studies provide evidence that IGF-I can accelerate protein synthesis in gastrocnemius during chronic sepsis by reversing the sepsis-induced inhibition of peptide-chain initiation. A limitation to infusing free IGF-1 is its rapid elimination from the circulation. rhIGFI/rhIGFBP-3 (SomatoKine™) stimulates muscle protein synthesis in chronically semi-starved animals whereas IGF-I alone failed to increase protein synthesis during the same experimental conditions. This stimulation was because of increased translation initiation of translation, likely induced by more physiologic concentrations/kinetics of plasma IGF-I and amino acids following rhIGF-I/rhIGFBP-3 treatment, compared to IGF-I in its free form [82]. Enhancing the bioavailability of IGF-I through administration of IGF-I/GFBP-3 complex stimulates the rate of protein synthesis in gastrocnemius by enhancing the translational efficiency in vivo [82]. The delivery of rhIGF-I/rhIGFBP-3 does not inhibit endogenously produced IGF-1 restores plasma IGF-1 concentrations in face of continuing septic challenge.

SUMMARY Sepsis rudely unhinges nitrogen metabolism homeostasis and profoundly disturbs the integration of inter-organ cooperatively in overall nitrogen and energy economy of the host. The result of such dyshomeostasis is a nitrogen catabolic state, which seriously compromises recovery and renders the septic patient semi-refractory to current therapeutic modalities. These alterations lead to a functional redistribution of nitrogen (amino acids and proteins) and substrate metabolism among major body organs. The metabolic response to sepsis is a highly integrated, complex series of reactions that manifest itself as excessive urea nitrogen

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excretion. To understand the regulation of the host's response to sepsis, a comprehensive, integrated analysis of the fundamental physiological relationships of key metabolic pathways and mechanisms is essential. Skeletal muscles possess the body's largest pool of nitrogen. The catabolism of skeletal muscles during sepsis, which is manifested by acceleration in protein degradation and inhibition in protein synthesis, persists despite state-of-the-art nutritional care. Much effort has focused on the overall amount of nutrients given to septic patients in a hope to improve efficiencies in utilization and nitrogen economies, rather than support of specific end-organ targets. Elevating leucine concentrations or enhancing IGF-1 bioavailability show promise as potential therapeutic modalities to improve skeletal muscle nitrogen balance during sepsis. The rationale for these two compounds appears to have the ability to reverse the septic-induced reduction in formation of active eIF4G·eIF4E. Unlike other therapeutic modalities, these approaches have been shown to improve protein metabolism after initiation of the septic insult rather than treating prophylactically.

ACKNOWLEDGEMENTS This work was supported in part by a National Institute of Health General Medical Sciences Award GM39277.

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Thomas C. Vary Oldfield S and Proud CG. Purification, phosphorylation, and control of guanine nucleotide exchange factor from rabbit reticulocyte lysate. Eur J Biochem 208: 73-81, 1992. Orellana RA, Jeyapalan A, Escobar J, Frank JW, Nguyen HV, Suryawan A, and Davis TA. Amino acids augment muscle protein synthesis in neonatal pigs during acute endotoxemia by stimulating mTOR-dependent translation initiation. Am J Physiol Endocrinol Metab 293: E1416-1425, 2007. Orellana RA, Kimball SR, Nguyen HV, Bush JA, Suryawan A, Thivierge MC, Jefferson LS, and Davis TA. Regulation of muscle protein synthesis in neonatal pigs during prolonged endotoxemia. Pediatr Res 55: 442-449, 2004. Orellana RA, Kimball SR, Suryawan A, Escobar J, Nguyen HV, Jefferson LS, and Davis TA. Insulin stimulates muscle protein synthesis in neonates during endotoxemia despite repression of translation initiation. Am J Physiol Endocrinol Metab 292: E629-636, 2007. Orellana RA, O'Connor PM, Bush JA, Suryawan A, Thivierge MC, Nguyen HV, Fiorotto ML, and Davis TA. Modulation of muscle protein synthesis by insulin is maintained during neonatal endotoxemia. Am J Physiol Endocrinol Metab 291: E159166, 2006. Orellana RA, O'Connor PM, Nguyen HV, Bush JA, Suryawan A, Thivierge MC, Fiorotto ML, and Davis TA. Endotoxemia reduces skeletal muscle protein synthesis in neonates. Am J Physiol Endocrinol Metab 283: E909-916, 2002. Peters KD, Kochanek KD, and Murphy SL. Deaths: final data for 1996. National Vital Statistics Reports 47: 1-100, 1998. Plank LD, Connolly AB, and Hill AG. Sequential changes in the metabolic response in severely septic patients duirng the first 23 days after onset of peritonitis. Ann Surg 228: 146-158, 1998. Ross RJ, Miell JP, Freeman E, Jones J, Matthews D, Preece M, and Buchanan C. Critically ill patients have high basal growth hormone levels with attenuated oscillatory activity assocaited with low levels of insulin-like growth factor-I. Clin Endocrinol (Oxf) 35: 47-54, 1991. Roth J, Martin D, Storr B, and Zeisberger E. Neutralization of pyrogen-induced tumour necrosis factor by its type 1 soluble receptor in guinea-pigs: effects on fever and interleukin-6 release. J Physiol 509: 267-275, 1990. Russell-Jones DL, Umpleby A, Hennessey T, Bowes S, Shojaee-Moradies F, Hopkins K, Jackson N, Kelly J, Jones R, and Sonksen P. Use of a leucine clamp to demostrate that IGF-I actively stimulates protein synthesis in normal humans. Am J Physiol Endocrinol Metabol 267: E591-E598, 1994. Scorsone KA, Panniers R, Rowland AG, and Henshaw EC. Phosphorylation of eukaryotic initiation factor 2 during physiological stresses which affect protein synthesis. J Biol Chem 262: 14538-14543, 1987. She P, Reid TM, Bronson SK, Vary TC, Hajnal A, Lynch CJ, and Hutson SM. Disruption of BCATm in mice leads to increased energy expenditure associated with the activation of a futile protein turnover cycle. Cell Metab 6: 181-194, 2007.

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Thomas C. Vary Vary TC. IGF-I stimulates protein synthesis in skeletal muscle through multiple signaling pathways during sepsis. Am J Physiol Regul Integr Comp Physiol 290: R313-R321, 2006. Vary TC, Deiter G, and Kimball SR. Phosphorylation of eukaryotic initiation factor eIF2Be in skeletal muscle during sepsis. Am J Physiol Endocrinol Metabol 283: E1032-E1039, 2002. Vary TC, Deiter G, and Lang CH. Cytokine-triggered decreases in levels of phosphorylated eukaryotic initiation factor 4G in skeletal muscle during sepsis. Shock 26: 631-636, 2006. Vary TC, Jefferson LS, and Kimball SR. Amino acid-induced stimulation of translation initiation in rat skeletal muscle. Am J Physiol Endocrinol Metabol 277: E1077-E1086, 1999. Vary TC, Jefferson LS, and Kimball SR. Insulin fails to stimulate muscle protein synthesis in sepsis despite unimpaired signaling to 4E-BP1 and S6K1. Am J Physiol Endocrinol Metab 281: E1045-E1053, 2001. Vary TC, Jurasinski CV, Karinch AM, and Kimball SR. Regulation of eukaryotic initiation factor 2 expression during sepsis. Am J Physiol Endocrinol Metab) 266: E193-E201, 1994. Vary TC, Jurasinski CV, and Kimball SR. Reduced 40S initiation complex formation in skeletal muscle during sepsis. Mol Cell Biochem 178: 81-86, 1998. Vary TC and Kimball SR. Effect of sepsis on eIF4E availability in skeletal muscle. Am J Physiol Endocrinol Metabol 279: E1178-E1184, 2000. Vary TC and Kimball SR. Sepsis-induced changes in protein synthesis: differential effects on fast- and slow-twitch muscles.. Am J Physiol Cell Physiol 262: C1513C1519, 1992. Vary TC and Lang CH. IGF-I activates the eIF4F system in cardiac muscle in vivo. Mol Cell Biochem 272: 209-220, 2005. Vary TC and Lynch CJ. Meal feeding enhances formation of eIF4F in skeletal muscle: role of increased eIF4E availability and eIF4G phosphorylation. Am J Physiol Endocrinol Metabol 290: E631-642, 2006. Vary TC and Lynch CJ. Meal feeding stimulates phosphorylation of multiple effector proteins regulating protein synthetic processes in rat hearts. J Nutr 136: 2284-2290, 2006. Vary TC and Lynch CJ. Nutrient signaling components controlling protein synthesis in striated muscle. J Nutr 137: 1835-1843, 2007. Vary TC and Murphy JM. Role of extrasplanchnic organs in the metabolic response to sepsis: Effect of insulin. Circ Shock 29:: 41-57, 1989. Vary TC, Owens E, Beers JK, Verner K, and Cooney R. Sepsis inhibits synthesis of myofibrillar and sarcoplasmic proteins: Modulation by interleukin-1 receptor antagonist. Shock 6: 13-18, 1996. Vary TC and Siegel JH. Sepsis, abnormal metabolic control and the multiple organ failure syndrome. In: In Trauma: Emergency Surgery and Critical Care. edited by (ed) JH Siegel. New York, NY: Churchill Livingstone, 1987, p. 411-502.

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[108] Vary TC, Siegel JH, Tall DH, Morris JG, and Smith JA. Inhibition of skeletal muscle protein synthesis in septic intraabominal abscess. J Trauma 28: 981-988, 1988. [109] Vary TC, Voisin L, and Cooney RN. Regulation of peptide-chain initiation during sepsis by interleukin-1 receptor antagonist. Am J Physiol Endocrinol Metabol 271: E309-E316, 1996. [110] Voisin L, Gray K, Flowers KM, Kimball SR, Jefferson LS, and Vary TC. Altered expression of eukaryotic initiation factor 2B in skeletal muscle during sepsis. Am J Physiol Endocrinol Metabol 270: E43-E50, 1996. [111] Welsh GI, Miller CM, Loughlin AJ, Price NT, and Proud CG. Regulation of eukaryotic initiation factor eIF2B: glycogen synthase kinase-3 phosphorylates a conserved serine which undergoes dephosphorylation in response to insulin. FEBS Lett 421: 125-130, 1998. [112] Welsh GI and Proud CG. Glycogen synthase kinase-3 is rapidly inactivated in response to insulin and phosphorylates eukaryotic initiation factor eIF-2B. Biochem J 294: 625-629, 1993. [113] Welsh GI, Wilson C, and Proud CG. GSK3: a shaggy frog story. Trends in Cell Biology 6: 274-279, 1996. [114] Wilmore DW. Metabolic response to severe surgical illness: Overview. World J Surg 24: 705-711, 2000. [115] Windsor JA and Hill GL. Grip strength: a measure of the proportion of protein loss in surgical patients. Br J Surg 75: 880-882, 1988. [116] Wojnar MM, Fan J, Frost RA, Gelato MC, and Lang CH. Alterations in the insulinlike growth factor system in trauma patients. Am J Physiol Regul Comp Physiol 268: R970-R977, 1995.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter VI

PROTEIN SYNTHESIS IN HEPATOCYTES OF MICE AS REVEALED BY ELECTRON MICROSCOPIC RADIOAUTOGRAPHY Tetsuji Nagata Department of Anatomy and Cell Biology, Shinshu University School of Medicine, Matsumoto 390-8621, Japan, and Department of Anatomy, Shinshu Institute of Alternative Medicine and Welfare, Nagano 380-0816, Japan

ABSTRACT For the purpose of studying the aging changes of protein synthesis in mouse hepatocytes, 20 groups of aging mice during development and senescence, each consisting of 3 individuals of both sexes, total 60, from fetal day 19 to postnatal day 1, 3, 9 and 14, month 1, 2 and 6, and year 1 and 2, were injected with RI-labeled amino acids, such as 3H-leucine, 3H-glycine, 3H-proline or 3H-hydroxyproline, which are the protein precursors, sacrificed 1 hr later and the liver tissues were fixed, sectioned and processed for electron microscopic radioautography. On electron microscopic radioautograms obtained from each animal, the localization of silver grains due to 3H-amino acids incorporations showing protein biosynthesis in respective cell organelles, the nucleus, Golgi apparatus, endoplasmic reticulum, mitochondrion and cytoplasmic matrix was qualitatively observed. On the other hand, the numbers of silver grains localizing over respective cell organelles were counted and analyzed quantitatively. The numbers of silver grains localized over the nuclei, Golgi apparatus, endoplasmic reticulum and cytoplasmic matrices increased from perinatal stages to the young adult stage at postnatal month 1, reaching the maximum, then decreased to year 2 due to aging and senescence. However, the number of silver grains localizing over the mitochondria, i. e. the number of labeled mitochondria per cell and the mitochondrial labeling index in each hepatocyte revealed that the numbers of mitochondria increased gradually from perinatal stages to the postnatal year 2, while the numbers of labeled mitochondria and the labeling indices of intramitochondrial protein syntheses in hepatocytes of mice at various ages increased

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Keywords: mitochondria, EM radioautography, protein syntheses, mouse hepatocytes, aging

1. INTRODUCTION Macromolecular synthesis such as nucleic acids (DNA and RNA), proteins, glucides or lipids in nuclei and cell bodies of various kinds of cells in various organs of experimental animals has been extensively studied since many years by both biochemical and morphological approaches [1-18]. Among of these studies, we first demonstrated the intranuclear and cytoplasmic nucleic acid syntheses, both DNA and RNA, in mammalian and avian cells morphologically by means of electron microscopic radioautography with accurate localization in cell organelles such as mitochondria, endoplasmic reticulum and Golgi apparatus in primary cultured cells of the livers and kidneys of mice and chickens in vitro [1, 2] as well as some other established cell lines such as HeLa cells [3-15] or mitochondrial fractions prepared from in vivo cells [5]. These phenomena were later commonly found in various cells and tissues not only in vitro cells obtained from various organs in vivo [16-24], but also in vivo cells of such various organs taken out from experimental animals as the salivary gland [25], the liver [26-43], the pancreas [44-45], the trachea [46], the kidney [47], the testis [48,49], the uterus [50-53], the spleen [54,55], the adrenal gland [56,57], the brain [58] and of mice and the eyes of chickens [59-63] and mice [64,65] in our laboratory. The relationship between the intramitochondrial DNA and RNA syntheses and cell cycle was formerly studied and it was clarified that the intramitochondrial DNA and RNA syntheses were performed without any nuclear involvement [5]. This paper reviews the relationship between the protein synthesis in hepatocyte nuclei and cytoplasm including cell organelles especially mitochondria and aging of mice in vivo at various ages by means of electron microscopic radioautography which was developed and carried out in our laboratory [1-65]. These studies supply additional data to the serial studies on special cytochemistry [66] and radioautographology [67].

2. RADIOAUTOGRAPHIC PROCEDURES Four groups of normal adult ddY strain mice, aged at postnatal month 1, each consisting of 3 litter mates, total 12, were housed under conventional conditions and bred with normal diet (mouse chow Clea EC2, Clea Co., Tokyo, Japan) with access to water ad libitum in our laboratory. The animals of 4 groups were injected with one of the protein precursors, 3Hglycine (Amersham, England, specific activity 1002 GBq/mM), 3H-leucine (Amersham, England, specific activity 1002 GBq/mM), 3H-tryptophane (Amersham, England, specific activity 877 GBq/mM), 3H-proline (Amersham, England, specific activity 877 GBq/mM) and 3 H-hydroxyproline (Amersham, England, specific activity 877 GBq/mM), at 9:00 a.m. simultaneously at a dosage of 370 KBq/gm body in saline. They were perfused at 10 a.m.,

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one hr. after the injections, via the left ventricles of the hearts with 0.1 M cacodylate-buffered 2.5% glutaraldehyde under Nembutal (Abbott Laboratories, Chicago, Ill., USA) anesthesia, then the liver tissues were taken out, cut into small tissue blocks, prefixed in 0.1 M cacodylate-buffered 2.5% glutaraldehyde, postfixed in buffered 1.0% osmium tetroxide, dehydrated, embedded in epoxy resin, sectioned and processed for electron microscopic radioautography according to the conventional wet-mounting procedure to demonstrate insoluble macromolecular compounds [67]. On the contrary, in order to demonstrate soluble small molecular compounds, some liver tissues were taken out, trimmed into small tissue blocks (1.0 x 0.5 x 0.5 mm) with two pieces of razor blades on a cold plate cooled to 0˚C immediately after the tissues were taken out and the tissue pieces were attached to small pieces of aluminum foils, 5 x 5 mm in size. The tissue pieces were quickly plunged into cooling agents (isopentane and propane mixture) at -161˚C cooled in quenching fluid (liquid nitrogen at -196˚C) for rapid freezing, then either freeze-dried at -80˚C, embedded in epoxy resin and dry-sectioned or cryo-sectioned with an LKB ultrotome 4800 equipped with an LKB cyrokit 14800 or LKB-NOVA (LKB, Bromma, Sweden) at -100˚C [68]. After the cryotechniques, all the frozen sections were processed for dry-mounting radioautography in order to demonstrate the soluble radiolabeled compounds [68]. Other normal ddY strain mice of 8 aging groups, each consisting of 6 litter mates of male animals, aged from fetal day 19 to postnatal day 1, 3, 9, and 14, month 1, 2 and 12, each group consisting of 3 litter mates respectively, total 48, were housed under conventional conditions and bred with normal diet (mouse chow Clea EC2, Clea Co., Tokyo, Japan) with access to water ad libitum in our laboratory. They were divided into 2 sub-groups, each 8 aging groups and administered with one of the 2 RI-labeled macromolecular precursors, 3H4,5-leucine (Amersham, England, specific activity 1002 GBq/mM) and 3H-proline (Amersham, England, specific activity 877 GBq/mM), at 9:00 a.m. simultaneously at a dosage of 370 KBq/gm body in saline. The animals were perfused at 10 a.m., one hr. after the injections, via the left ventricles of the hearts with 0.1 M cacodylate-buffered 2.5% glutaraldehyde under Nembutal (Abbott Laboratories, Chicago, Ill., USA) anesthesia, then the liver tissues were taken out, postfixed in buffered 1.0% osmium tetroxide, dehydrated, embedded in epoxy resin, sectioned and processed for electron microscopic radioautography. All the procedures used in this study concerning the animal experiments were in accordance with the guidelines of the animal research committee of Shinshu University School of Medicine as well as the principles of laboratory animal care in NIH publication No. 86-23 (revised 1985). For light microscopy, thick sections at 1 µm thickness from respective specimens were cut in sequence and processed for radioautography with Konica NR-M2 radioautographic emulsion (Konica, Tokyo, Japan) by a dipping method [19,21,42,64,65]. For electron microscopy, semithin sections at 0.2µm thickness were cut in sequence, collected on collodion coated copper grid meshes and coated with Konica NR-H2 radioautographic emulsion (Konica, Tokyo, Japan) by a wire-loop method [19, 20, 21, 42, 64,65]. The electron microscopic radioautograms were examined in either a Hitachi H-700 electron microscope (Hitachi, Tokyo, Japan) at an accelerating voltage of 200 kV or a JEOL JEM-4000EX electron microscope (JEOL, Tokyo, Japan) at accelerating voltages of 300-400 kV for observing thick specimens [42,64,65].

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For quantitative analysis of electron micrographs, twenty EM radioautograms showing cross sections of mononucleate hepatocytes from each group, based on the electron microscopic photographs taken after observation on at least 100 hepatocytes from respective animals, were analyzed to calculate the total number of mitochondria in each hepatocyte and the number of labeled mitochondria covered with silver grains by visual grain counting. On the other hand, the number of silver grains in 10 circles with the same area size as a mitochondrion outside cells was also calculated in respective specimens as background fog. The average number of silver grains per mitochondrial area was 0.01-0.03/area in the respective groups, which resulted in less than 1 silver grain per area. Therefore, the grain count in each specimen was not corrected with background fog. Thus, the mitochondrion that was labeled with more than one silver grain was defined as labeled. The data were stochastically analyzed using variance and Student's t-test. The differences were considered to be significant at P value methylated BSA. The percent conversion to digested product when averaged for 2 digestion trials was as followed: 21.2 ± 9.3 % acetylated BSA > 17.2 ± 5.3 % BSA> 11.8 ± 2.3 % H.S.A.> 9.2 ± 2.3 % carbonic anhydrase > 4.1± 0.4 % methylated BSA. Surprisingly, the acetylated BSA and normal fraction V BSA appear closely matched. Acetylation of BSA should eliminate lysine cleavage sites leaving only 26 arginine cleavage sites. The human serum albumin gave a slightly lower extent of digestion relative to normal BSA perhaps due to reflecting subtle differences in conformation. The digestion rate and extent of digestion for methylated bovine serum albumin was dramatically reduced relative to normal BSA fraction V. These profiles observed were very reproducible and clearly suggested that the digestion signal from the RWG biosensor does convey substrate specific digestion properties. Figure 4D depicts the digestion profile for three proteases using a fixed concentration of carbonic anhydrase II. All three proteases differ in their cleavage sites as well as in their mechanism. The typical proteolytic signature for three proteases (at 0.5 units/µL) for the immobilized substrate carbonic anhydrase II was measured. The observed hierarchy of proteolytic digestion rate of carbonic anhydrase II was as follows; Thermolysin> Chymotrypsin> trypsin. The thermolysin digestion resulted in a significantly higher degree of proteolysis which is likely due to the fact that thermolysin has more than two times the number of cleavage sites over either trypsin or chymotrypsin.

Figure 4. Real-time functional proteolytic assay on an HTS resonant waveguide (RWG) Epic® system biosensor microplate. A. Schematic illustration of how a protease removes immobilized proteins off of a sensor. B. Spectral profile of light selectively reflected off of the resonant waveguide. C. Trypsin digestion profiles for five different proteins immobilized on the RWG sensor. Trace 1) acetylated BSA, Trace 2) BSA fraction V, Trace 3) carbonic anhydrase II, Trace 4) human serum albumin (HAS), Trace 5) methylated BSA and trace6) PEG-amine control well. D. Effect of proteolytic enzyme digestion on carbonic anhydrase II. Trace 1) thermolysin, trace 2) chymotrypsin and 3) trypsin. All enzymes were at 0.5 units/μl. Reprinted here by permission from SAGE publishers.

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Figure 5. Protease inhibitor drug screens on an HTS label-free biosensor. A. Effect of titrating increasing amounts of leupeptin inhibitor on carbonic anhydrase II digestion from trypsin. B. Digestion response versus Log -molar concentration of Phosphoramidon . C. Direct protease inhibitor specificity profiling of 3 different classes of proteases obtained on an HTS biosensor. Reprinted here by permission from SAGE publishers.

The digestion technique was extended further into a drug screening approach. The ability of known inhibitors to be assayed using proteases and a dextranase over a 20 minutes time regime which is in keeping with standard high through put screening. Proteases from three different protease classes (Serine protease; trypsin, cysteine protease; papain and metalloproteinase; thermolysin) were examined. Carbonic anhydrase II was used as the common substrate for all protease variable screens. The effect of various concentrations of leupeptin on the trypsin-carbonic anhydrase II digestion time course is shown in Figure 5 A. As can be seen from figure 5A most of the inhibition has occurred prior to 6.5 µM leupeptin. In addition the digestion signal taken at the 20 minute time point appears to provide a reliable measure of inhibitor effect. These digestion profiles were found to be representative of the other protease inhibitor screens. The experimentally measured EC 50 values compared very closely with those reported EC 50 values (O’Malley et al., 2007 [93]). This agreement provided sufficient confidence that the surface based digestion profiles were capable of reliably detecting inhibitor effects. A representative response versus Log [Dose] curve for thermolysin-phosphoramidon is shown in figure 5 B. The EC 50 determinations were obtained from fitted curves for the enzyme-inhibitor pairs using a 2 log concentration range. The sigmoidal fitted data was obtained using Prism GraphPad. The work in O’Malley et al., 2007 [93] also demonstrated that the lytic enzymes were shown to be differentially

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susceptible to various inhibitors. Figure 5 C depicts the protease inhibitor specificity using three classes of proteases for various protease class inhibitors. The percent normal lytic activity was obtained by normalization to the digestion signals in 1x PBS. The inhibitor selectivity undoubtedly reflected the difference between the various hydrolytic mechanisms between the three enzymes. As a screening tool biosensors clearly appear to be highly effective tools at distinguishing between a non-inhibitor and an inhibitor A future prospect is that these surface-based hydrolytic assays could be used in highly parallel microarray formats. Microarray biosensors have provided the ability to look at global protein interactions (Yuk and Ha, 2005 [103]). Conceivable, the lytic assays from O’Malley et. al., 2007 [93] could also be applied toward global degradomic studies. In some applications one may wish to look at the off target specificity of a various hydrolytic enzyme inhibitors. In other cases, one may wish to examine the viability of peptide based therapeutics under in vivo like conditions. Whole proteomic digestion profiles may also be possible with this technology and will likely provide useful insights into a poorly understood cellular phenotype.

Atypical Proteolytic Assays In the previous section we focused on using label free detection as a tool for examining the inhibition of proteolytic enzymes. Here, we turn our attentions to the effects that ligands can have on substrates tethered to a biosensor with regard to their conformational state. The conformational state of the protein targets in the previous section was accepted as is. It was a given that the conformational state was intrinsic and specific to each of the protein targets examined. Yet, just as each protease has its own specific amino acid preference for cleavage it is also true that each protein target has its own amino acid sequence displayed in its own conformational state. However, molecular interactions such as small molecule ligand-ligate binding or macromolecular complex formation (e.g. protein-protein, nucleic acid-protein association) have been known to result in dramatic conformational changes. One anticipates that for some molecular interactions a conformational change might result which could either enhance or reduce proteolysis by changing the availability or degree of exposure of the cleavage sites. Many traditional biophysical methods have been used to study localized conformational changes such as fluorescence anisotropy, Stern-Volmer fluorescence quenching, FRET, circular dichroism, UV-Vi difference spectroscopy, Raman, NMR and so on. In each of these methods a specific structural signature is utilized to track changes in conformational state. For fluorescence assays either intrinsic fluorescent amino acids (e.g. tryptophan) or extrinsic fluorescent tags are used. The ability to use proteolytic signatures as a means to track conformational transitions for RNA-protein complex formation using mass spectrometry and gel electrophoresis profiling has been shown previously (O’Malley et al, 1995 [104]). However, a direct label free high through put technique for measuring changes in conformational state has been lacking. In recent years several publications using label free optical biosensing have been directed toward detection of conformational changes. One of the earliest efforts to use optical biosensing to detect conformational changes was reported by Sota et al, 1998 [105]. In this

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paper, a pH specific signal was measured for immobilized dihydrofolate reductase (DHFRASC) relative to blank sensors. The use of this comparative difference between bulk and surface pH to explain hydrodynamic modulation of protein structure was nonetheless not widely adopted. Similarly, Gestwicki et al., 2001 [106] later reported the use of negative refractive index changes via hydrodynamic -fluidic shifts to infer conformational changes between a small molecular weight binder (maltose) and its receptor maltose binding protein. Of course the absence or presence of affinity between two binder pairs by itself can be taken as evidence for a conformational state (Honjo et al., 2002 [107] and Robelek et al, 2007 [108]). Conformational diversity in some SPR studies has been inferred through the failure of an association and dissociation model to be adequately fitted to an observed kinetic reaction rate via global fitting routines (Von Der Haar et. al., 2006 [109]). However, despite these efforts the field of optical biosensing is still in need of specific assay tools which can be employed toward assaying conformational changes robustly. In this next section, it is shown that small molecular weight binding agents can result in a modulation (reduction) of a proteolytic signature. Also in this section we demonstrate the use of catalytic agents which can facilitate proteolysis degradation. One model system which conveniently demonstrates a dramatic conformational transition upon small ligand binding is heme insertion. Heme proteins are evolutionarily designed to stereo-specifically retain the heme moiety. Hemin or heme is a planar 20 carbon iron containing macro-cyclic compound (651.96 Dalton). The insertion of heme into the heme pocket of most heme-proteins is known to be extremely stable and requires special extraction methods. Removal of heme from human hemoglobin has been measured to cause a 50% loss in alpha helical content (Waks et. al., 1973 [110]). Perhaps the most compelling reason for this dramatic transition lies in the fact that both the alpha and beta protein subunits of the hemoglobin tetramer contain 18 to 20 atomic contacts (respectively) with the heme molecule (Fermi and Perutz, 1984 [111]). The results of the experiment shown in Figure 6 were done in order to demonstrate the effect ligand induced conformational changes on proteolytic cleavage. Horse apomyoglobin used in this experiment was prepared using the acid-acetone method of Rossi-Fanelli et al., 1958 [112] with modifications described in Ascoli et. al., 1981 [113]. The extent of heme removal was confirmed spectrophotometrically to be greater than 90%. Figure 6 A illustrates how heme insertion can result in a “protection” of cleavage sites on the surface of immobilized apomyoglobin. The red triangle on the protein surface in figure 6 A represents a trypsin cleavage site. Prior to heme insertion the structure is fairly lose and the cleavage site is fully available. Upon heme insertion the structure changes and now lacks complete solvent accessibility to trypsin. Figure 6 B depicts the sensogram recorded on the Corning Epic® system for the effect of heme insertion into apomyoglobin. Each trace represents an average of six kinetic time traces for trypsin digestion of myoglobin (3), apomyoglobin (1) and hemin inserted apomyoglobin (2). Traces 4 &5 are control wells for trypsin digests of blank wells coated with PEG only. Not surprisingly, the apomyoglobin exhibited the largest degree of digestion due to collapsed secondary structure while myoglobin (3) and hemin reconstituted myoglobin (2) both demonstrated a significant protection by maintenance of the occupied heme pocket structure. This experiment illustrates

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how proteolytic digestion can be used to provide a profile of ligand induced conformational changes. It is contemplated that these two ligand dependent effects on proteolysis can be extended further in two specific ways. First, for some specific ligand- ligate systems the interaction may be sufficiently robust to allow label free HTS drug screens. In such as case, one would first screen various enzymes and under various buffer conditions using a known ligand ligate binding pairs. Those conditions which give a ligand specific digestion signature are then used to screen for other putative ligands which yield a similar or matching substrate/target digestion signature. Controls would then be required to verify that the effect was restricted to the immobilized ligand and not a protease modulating drug. The second prospective expansion of these atypical label-free digestion assays is to use as a tool for comparing native state via an expected digestion signature to a mis-folded state. If the mis-folded state is validated to cause a detectable change in the digestion profile then these assays should provide a HTS-screens capability.

Figure 6. Effect of ligand induced protease protection. A. Schematic of heme insertion into apomyoglobin B. Kinetic time course for a trypsin digest of three forms of myoglobin on an RWG biosensor. Trace 1) Apomyoglobin, trace 2) Heme reconstituted myoglobin (Heme inserted), trace 3) native myoglobin and traces 4/5 are PEG-amine control wells. Note the reconstituted apomyoglobin trace 2 is almost as protected as the native myoglobin.

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Figure 7. Catalytic proteolysis probed by label-free biosensing. A. Schematic of how dimeric porphyrins invades an immobilized protein making its buried residues exposed to proteolytic cleavage. B. Kinetic time course for trypsin digest of cyctochrome C in the presence and absence of compound 2b (proteolytic catalyst). Note the dramatic increase in digestion when the catalyst is present.

Catalytic Digestion Assays Catalytic digestion involves the use of a chemical agent to alter the digestion activity of a digesting entity. The alteration to digestion can be an increase or a decrease. The catalytic description applies to this system wherein the chemical agent used to modulate the digestion process is not consumed in the reaction. The chemical agent can be organic, inorganic, polymeric and biopolymeric, organometalic, biologically derived phage peptide, a synthetic peptide, and aptamer, and DNA or any combination thereof. Groves et al., 2004, [114] describe a class of copper porphyrin dimmers which are examples of catalytic digestion agents. These agents when added to a digestion system can result in an enhancement or actual enablement of the digestion process. Below is a description of the Groves et. al. 2004 [113] based catalytic digestion system. Figure 7 A illustrative depiction of how a dimeric porphyrin physically invades a protein structure causing an unfolding event which then exposes amino acid residues within the biopolymer to be cleaved by a proteolytic. The authors demonstrate both circular dichroism and SDS PAGE gel electrophoresis of the protein. This paper limits itself to just protein targets yet in theory it should be extendable to any macromolecular structure such as a carbohydrate, protein, peptide, RNA, DNA, membrane protein and the like. The catalytic

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digestion should also not be limited to just copper II containing porphyrins. Chemical agents may be used to modulate label-free digestion signals on a biosensor. As in the previous section, these catalytic agents could be used to study biopolymer stability, detection of intermolecular interactions such a ligand-ligate systems, or to enable or aid in the formulation of a digestion assay when the immobilized agent is difficult to digest. The experiment in Groves et. al., 2004 [114] was reconstituted and is shown in Figure 7B. The catalytic agent described in Groves et.al. 2004 [114] that was used in this study was copper (II) tetra-(4-carboxyphenyl) phorine AKA “compound 2b”. The procedure for making compound 2b was as follows; (1) A 10-fold excess (w/w) of copper II acetate was added to a solution of the free base tetra-(4-carboxyphenyl_porphine in methanol. After 16 hours at room temperature, the insoluble copper porphine was then filtered off and washed with 1mM HCl, distilled water, ethanol and then dried by nitrogen and heat in a speed vac. The copper porphine extracts from the water phase wash were sufficient to use in the digestion assay. Equine cytochrome C was chemically immobilized onto the amine reactive surface of an Epic® System sensor microplate. The comparative trypsin digestion on the Epic® system was measured. The catalyst arrow points to the digestion trace (red/yellow) for the copper II porphine and demonstrate a significant enhancement of digestion. In these assays 15 units of trypsin/100 μl were used in each well. As can be seen from figure 7B the addition of catalyst does in deed enhance the rate and extent of catalysis.

CONCLUSION In this chapter recent advances in label free technology were reviewed with special focus on high throughput screens and protein biosynthesis. The biosensor field has matured to the point where label free detection can be applied in either high through put screens or in multiplexed microarray formats. The application space depending on the method of label free detection can be applied to high content cell based screens, conventional direct bind assays and functional cell free protein synthesis and protein degradation assays. As the application space for label-free detection continues to grow inevitably so will the user base. We have also seen that these biosensors can be used in conjunction with other formats to provide even more unique data. As seen in the union of mass spectrometry with SPR. As sensor designs improve, shrink and become free standing it is possible that we may see future applications wherein transfectable biosensors are deployed directly into cells for real time assay of in vivo biological events. The interaction partner pairs for protein biosynthesis (Table III) may now become components for high throughput drug screens using optical biosensors are microplate formats are now available. The last section on protein degradation offers the possibility that HTS lytic inhibitors can be rapidly and reliably screened. The dependence of the functional lytic screen on substrate structure may also avail itself toward the study of mis-folded proteins

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[120] Ptushkina M, Berthelot K, von der Haar T, Geffers L, Warwicker J, McCarthy JE. A second eIF4E protein in Schizosaccharomyces pombe has distinct eIF4G-binding properties. Nucleic Acids Res. 2001. 15;29(22):4561-4569. [121] Iwasaki K, Kikukawa S, Kawamura S, Kouzuma Y, Tanaka I, Kimura M. On the interaction of ribosomal protein L5 with 5S rRNA. Biosci Biotechnol Biochem. 2002. 66(1):103-9. [122] Petrushenko ZM, Budkevich TV, Shalak VF, Negrutskii BS, El'skaya AV. Novel complexes of mammalian translation elongation factor eEF1A.GDP with uncharged tRNA and aminoacyl-tRNA synthetase. Implications for tRNA channeling. Eur J Biochem. 2002. 269(19):4811-8. [123] Honjo E, Watanabe K, Tsukamoto T. Real-time kinetic analyses of the interaction of ricin toxin A-chain with ribosomes prove a conformational change involved in complex formation. J Biochem. 2002. 131(2):267-75. [124] Mandon EC, Jiang Y, Gilmore R. Dual recognition of the ribosome and the signal recognition particle by the SRP receptor during protein targeting to the endoplasmic reticulum. J Cell Biol. 2003. 162(4):575-85. [125] von der Haar T, Hughes JM, Manjarul Karim M, Ptushkina M, McCarthy JE. Translation initiation and surface plasmon resonance: new technology applied to old questions. Biochem Soc Trans. 2002. 30(2):155-162. [126] Nakano H, Yoshida T, Uchiyama S, Kawachi M, Matsuo H, Kato T, Ohshima A, Yamaichi Y, Honda T, Kato H, Yamagata Y, Ohkubo T, Kobayashi Y. Structure and binding mode of a ribosome recycling factor (RRF) from mesophilic bacterium. J Biol Chem. 2003. 278(5):3427-36. [127] Komarova AV, Real E, Borman AM, Brocard M, England P, Tordo N, Hershey JW, Kean KM, Jacob Y. Rabies virus matrix protein interplay with eIF3, new insights into rabies virus pathogenesis. Nucleic Acids Res. 2007. 35(5):1522-32. [128] von der Haar T, Oku Y, Ptushkina M, Moerke N, Wagner G, Gross JD, McCarthy JE. Folding transitions during assembly of the eukaryotic mRNA cap-binding complex. J Mo.l Biol. 2006. 356(4):982-92 [129] Lee KH, Joung HA, Ahn JH, Kim KO, Oh IS, Shin YB, Kim MG, Kim DM. Real-time monitoring of cell-free protein synthesis on a surface plasmon resonance chip. Anal. Biochem. 2007. 366(2):170-4.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter VIII

TRANSLATING MEMORIES: THE ROLE OF PROTEIN BIOSYNTHESIS IN SYNPATIC PLASTICITY Cara J. Westmark and James S. Malter Department of Pathology & Laboratory Medicine and Waisman Center for Developmental Disabilities, University of Wisconsin, Madison, WI 53705, USA

ABSTRACT The 1990s, “The Decade of the Brain”, resulted in major scientific advances involving brain imaging, gene therapy, brain/robotic interfacing and the neurobiology and molecular biology of learning and memory. However, despite these critical insights, we still do not know exactly how thoughts or memories are formed or stored in the brain, which leaves much exciting research for the twenty-first and probably centuries to come. This review will elaborate on recent advances in the field of protein biosynthesis as related to synaptic plasticity. We will discuss the molecular players (RNA binding proteins and neuronal mRNAs), the signal transduction pathways that have been implicated in learning and memory and how localized translation of selected mRNAs is involved in synaptic plasticity. We will also discuss the pathology of human diseases including Alzheimer’s disease, Fragile X syndrome, autism and Down syndrome, which show altered or diminished protein synthesis dependent synaptic plasticity. Learning and memory are manifested in their highest form in humans and allow for the retrieval of and action on past events. Understanding the pathology of these neurological disorders will elucidate the normal mechanisms of memory formation and storage.

INTRODUCTION Synapses are the junctions between the axon of one neuron and the dendritic spine of another neuron. Electrical signals traveling down axons are converted to chemical

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information and released as neurotransmitters into the synaptic cleft to activate adjacent dendrites. Synaptic plasticity refers to changes in transmission frequency or strength in response to neurotransmitter stimuli. Donald Hebb proposed in 1949 that, “When an axon of cell A is near enough to excite cell B or repeatedly or persistently takes part in firing it, some growth process or metabolic change takes place in one or both cells such that A's efficiency, as one of the cells firing B, is increased [1].” If true, changes in synaptic plasticity would be the biochemical basis for learning and memory. Exactly how are experiences, knowledge and skills formed, stored and recalled in the brain? Recent neuroscience research has established an essential role for biochemical changes in the structure of neurons and their synaptic connections. Santiago Ramon y Cajal described dendritic spines in the late 1800s as thorn-like structures that could provide the basis for learning and memory [2]. Dendritic spines project from the dendritic shaft and vary in shape and size. Mature, stable spines have a squat, short mushroom shape and reflect strong synaptic connections whereas immature spines are long and thin and form weaker synaptic connections. Thus, dendritic spines are “dynamic” or “plastic” in nature as their shape, size and number change in response to synaptic activity. A single neuron in the mammalian nervous system may contain ten thousand synapses. Dendritic spines regulate the movement of calcium into the dendritic shaft and organize the complex protein mesh adjacent to the synaptic cleft. This meshwork contains cytoskeleton and hundreds of proteins, and due to its electron dense appearance under electron microscopy, is called the post-synaptic density (PSD). Chemical and electrical stimuli can be transformed into biomolecules by de novo protein biosynthesis at dendritic spines. Activitydependent translation can then alter synapse morphology and synaptic strength. The morphology (shape and size) and density of dendritic spines thus reflects synaptic plasticity and depends on the organization of proteins at the PSD. The PSD is composed of approximately 100 different proteins including neurotransmitter receptors, scaffolding proteins and signal transduction molecules [3,4]. Scaffolding proteins, such as Homer, PSD95 and Shank, connect membrane receptors to downstream signaling molecules and regulate their number, distribution and subcellular location. Glutamate is the primary neurotransmitter at excitatory synapses, which through N-methyl-D-aspartic acid (NMDARs), alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPARs) and metabotropic glutamate (mGluRs) receptors contributes to dendritic protein synthesis [5]. The major downstream signaling pathways that bridge glutamate receptor activation with translation initiation include protein kinase A (PKA), extracellular-regulated kinase of the mitogen-activated protein kinase family (MAPKERK) and mammalian target of rapamycin (mTOR) [6,7]. Thus, interactions between membrane receptors, scaffolding proteins and signal transduction molecules in dendritic spines and spine heads underlie activity-dependent protein synthesis and ultimately the composition of the dendritic spines, generally, and the PSD in particular. Synapse remodeling occurs in response to learning [8] and the addition of new synapses is required for long-term information storage in the brain [9]. The role of protein biosynthesis in learning and memory has been studied in both invertebrate and vertebrate systems [10,11]. The marine mollusk Aplysia californica is a model system to study synaptic function. Aplysia has an intact central nervous system and local protein synthesis in presynaptic sensory

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neurons, but not postsynaptic cells, is required to induce long-term branch facilitation [12]. In mammals, long-term potentiation (LTP) and long-term depression (LTD) are electrophysiological correlates reflecting the cellular basis for learning and memory. LTP is the persistent increase in synaptic strength following high-frequency stimulation of a chemical synapse. LTP is composed of two forms, so-called “early” and “late-phase”. Earlyphase LTP models short-term memory, which lasts minutes to hours and is protein synthesis independent, whereas late-phase LTP (L-LTP) models long term memory (LTM), which lasts from days to a lifetime and requires de novo protein synthesis, gene transcription and the growth of new synaptic connections. Several processes may contribute to LTM, including (1) the sequestration of specific proteins at activated synapses, ie. synaptic tagging [13], (2) local protein synthesis at dendrites, and (3) protein trafficking. LTD is the weakening of a neuronal synapse resulting from strong synaptic stimulation or persistent weak synaptic stimulation. LTD can be elicited through the activation of NMDAR or mGluR. Translation, but not transcription, is required for the maintenance of LTD in the CA1 region of the hippocampus [14]. Thus, de novo protein synthesis is essential for L-LTP as well as LTD, suggesting that increases or decreases in synaptic strength reflect alterations in dendritic protein composition, amount or both. The transport and localization of mRNAs to synaptic sites have been recently reviewed [15-17]. Hence in this review, we will highlight the mechanisms that control the translation of dendritically localized mRNAs. A large number of dendritic mRNAs and mRNA binding proteins (RBPs) have been identified that contribute to these regulatory events. Finally, we will extrapolate to the pathology of neurological disease and in particular to the abnormal synaptic plasticity that occurs in Fragile X syndrome (FXS). We will discuss the role(s) of cis-elements and trans-factors in the translation of amyloid precursor protein (APP) mRNA, which is likely involved in the pathogenesis of Alzheimer’s disease (AD), Down syndrome (DS), FXS, autism and seizures.

TRANSLATIONAL MACHINERY AT DENDRITIC SPINES Dendrites have specific sites specialized for rapid translation [18]. These translational “hot spots” contain all of the required molecules, such as mRNAs, ribosomes, tRNAs, and translation initiation, elongation and release factors (eIF2α, eIF4E, eIF4G, eIF5, eIF6, eEF1α, eEF2, and ERF1), required for protein biosynthesis [19,20]. Neuronal stimulation increases the percentage of spines containing polyribosomes, which precedes enlargement of the PSD [21]. Similarly during LTP, polyribosomes move from dendritic shafts into dendritic spines [21] where they are preferentially located under the base of the spines [22]. Translational machinery and mRNAs localized near synapses allow for modulation of synaptic strength through local protein synthesis. The rate and efficiency of translation is regulated by phosphorylation. The initiation step of translation is the rate-limiting step and is controlled by the initiation factor eIF4E, which is a component of the cap binding complex, eIF4F. eIF4F is a binding partner of 4E-BP1/2 [23]. Stimulation with neurotrophic agonists, for example, brain-derived neurotrophic factor (BDNF), facilitates the translocation of eIF4E into dendritic spines [24]. The cap structure

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(eIF4F) facilitates the attachment of the 40S ribosomal subunit to mRNA. 4E-BPs inhibit translation by hindering eIF4F complex assembly through repressive interactions with eIF4E [25]. The translational repressor 4E-BP2 is necessary for synpatic plasticity and memory in the hippocampus [26]. The phosphorylation of eIF2α regulates the switch from short- to long-term synaptic plasticity and memory [27]. In addition to the translational machinery, dendritic spines contain the organelles necessary for protein processing and membrane insertion. The translation and processing of integral membrane proteins requires organelles of the secretory pathway to provide for proper synthesis, glycosylation, membrane targeting and insertion. Dendritic shafts and spines contain rough endoplasmic reticulum (RER), Golgi apparatus (GA) and associated vesicular transport machinery [28-30]. Hence, all of the mRNA and protein components as well as secretory organelles required for protein biosynthesis, modification and membrane insertion are locally available at dendritic spines. The key questions then are, “what mRNAs and proteins constitute the biological basis of memory,” and “how do their respective cis- and trans-elements interact to mediate mRNA transport and localization to and translation at dendrites?”

DENDRITICALLY LOCALIZED RNAS AND THEIR CIS-ELEMENTS Hippocampal dendrites contain approximately 400 distinct mRNAs coding for membrane receptors and channels, signaling molecules, cytoskeletal and adhesion molecules and proteins involved in membrane trafficking, translation, post-translational protein modification and protein degradation [31-34]. The immediate early gene Arc/Arg3.1, which is an activityregulated cytoskeleton-associated protein, selectively localizes to postsynaptic sites on activated dendrites [35,36]. Several neuronal mRNAs, such as the α-subunit of calcium/calmodulin-dependent kinase II (CaM-KIIα), MAP2 and β-actin, contain dendritic targeting signals in their 3’-untranslated regions (UTRs) [37-39]. While these and other mRNAs localize to dendritic spines, that does not necessarily mean that they are translated in response to synaptic signals [15,32]. Likewise, a wide range of proteins with varied functions are found at dendrites and include MAP2, dendrin, CaM-KIIα, Arc/Arg3.1, G-protein gamma subunit, calmodulin, NMDAR1, glycine receptor alpha subunit, vasopressin, neurofilament protein 68, inositol 3 phosphate (InsP3) receptor, ribosomal protein L7 and PEP1. Protein accumulation could reflect dendritic transport [16,17], dendritic translation [40-43] or both [15,44-47]. The “mRNA targeting hypothesis” suggests that plasticity-related mRNAs are localized/targeted to activated synapses, which allows for local protein synthesis as seen with Arc/Arg3.1 mRNA and protein [35,36]. To discriminate between transport to versus de novo protein synthesis at dendritic sites, the translation of specific dendritic mRNAs has been assessed in synaptoneurosome (SN) preparations. SN are sealed pre- and post-synaptic membranes formed after homogenization of brain tissue in sucrose solution. It is worth noting, however, that SN are not entirely pure dendritic preparations. CaM-KIIα is a dendritically localized mRNA, and if disrupted by 3’-UTR mutation, knock-in mice show

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reduced L-LTP and impairments in long-term spatial memory, associative fear conditioning and object recognition memory [48]. The fraction of CaM-KIIα mRNA associated with polyribosomes in WT SN as well as protein in the PSD is increased after neurotransmitter activation [49]. Therefore, the majority of CaM-KIIα in the PSD originates from local protein synthesis, and not transport from the cell body. The stability, transport and translation of mRNAs are often regulated through protein/mRNA interactions. Cis-elements often found in the 3’-UTR are bound by RBPs, which mediate variable stability, degradation, localization, docking and/or translation. Examples include AU-rich elements (AREs) [50], zip code elements [51], internal ribosome entry sites (IRES) [52] and cytoplasmic polyadenylation elements (CPEs) [53]. Almost 10% of mammalian coding mRNAs contain 3’-UTR AREs, which are composed of reiterated pentamers of AUUUA or simpler U-rich domains. These are commonly found in rapidly degraded mRNAs coding for cytokine, transcription factor, proto-oncogene and receptor mRNAs. HuR/ELAV family members, AUF1, TTP and KSRP (KH-type splicing regulatory protein) among others bind to AREs [54-57]. The HuR/ELAV proteins protect messages from degradation [58-60] whereas AUF1, TTP and KSRP can destabilize mRNAs [56,57,61]. ARE binding proteins (AUBPs) can also regulate translation. For example, competitive binding between AUF1 and TIAR determine translational efficiency of Myc mRNA [62]. RNA zip code elements route mRNAs to specific subcellular sites. These elements are bound by zip code binding proteins, such as ZBP2 whose mutation dysregulates mRNA localization and hence synthesis [63]. Internal ribosome entry sites (IRES) are found in several dendritically localized mRNAs including Arc/Arg3.1, CaM-KIIα, dendrin, MAP2, neurogranin and Fmr-1 [64,65]. The 5’-UTR IRES of Arc/Arg3.1, CaM-KIIα and MAP2 mRNAs enhanced translational efficiency [64]. Multiple cis-elements may co-exist in a single mRNA. For example, the 3’-UTR of PSD95 mRNA has 3 potential translational control elements including a differentially controlled element (DICE), a putative CPE involved in mRNA localization and translation and a G-quartet [66,67]. While a definitive understanding of how information from multiple and varied cis-elements are integrated is not known, the number, location and type of these elements clearly determines the stability, localization and translation of the messages. The function of some of these RBPs, such as AUF1, have not been specifically studied in dendritic locations, but others including the KSRP orthologue MARTA1 and CPEB are active in dendrites [68,69]. RBPs bind to the newly synthesized mRNAs in the nucleus and commence nuclear export to the cytoplasm via ribonucleoprotein complexes (RNPs). Exported RNPs may be translated in the perikaryal somatic region or travel to distant extrasomatic destinations for local on-site translation. Proteins and mRNAs travel along a network of microtubules/microfilaments composed predominantly of polymerized tubulin in cells and actin in dendrites [70-72]. Dendritic spines contain a network of dense, highly branched short actin filaments. RNA binding proteins bind to cis-acting elements in the mRNAs as well as to the cytoskeletal network to mediate transport. A well-studied example is the formation of an RNP including β-actin mRNA, ZBP1 and staufen2, which localizes to dendrites [73-76]. Subcellular mRNA localization and protein targeting and anchoring mechanisms play an important role in the functional organization of the dendrite and are dependent on ciselements, trans-factors and signaling pathways. However, communication between the cell

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body and synapses through RNP trafficking on the microtubule network is not required for protein synthesis dependent L-LTP [77]. Along with mRNA stability and localization, the timing and degree of polyadenylation is an important translational control mechanism. The translation of maternal mRNAs in maturing oocytes can be modulated by CPE-mediated polyadenylation [78]. Not all CPEcontaining mRNAs are polyadenylated at the same time, however. Polyadenylation may occur early in prophase or later at metaphase I. Early cytoplasmic polyadenylation requires CPE and Hex elements, which do not overlap, and late polyadenylation requires at least 2 CPEs with one of them overlapping the Hex element(s) [79]. Twenty-four different configurations of the basic CPE, Hex and PBE cis-elements define different modes of translational behavior with the number and relative position of the elements determining the timing and efficiency of translation [79]. Other RBPs, such as Xenopus Pumilio (Pum), interact with CPEB and are also involved in translational activation and repression. Neuroguidin binds to eIF4E and to CPEB and represses translation in a CPE-dependent manner [80]. Many mRNAs undergo activity-dependent polyadenylation in neurons, which coincides with enhanced translation in synaptodendritic compartments [81]. In addition to mRNA, microRNAs (miRNAs), are found at dendrites. miRNAs are small, non-coding RNAs that copurify with polyribosomes. After binding to complementary sequences, translation is suppressed by DICER-induced cleavage of the target mRNA [82,83]. Eighty-six miRNAs have been identified in mammalian neurons [84]. One example is miRNA-134, which is localized to synaptic sites in hippocampal neurons, and through translational repression of Limk1 mRNA, negatively regulates the size of dendritic spines. BDNF overrides miRNA-134-mediated inhibition of Limk1 synthesis and thus promotes synaptic plasticity [85]. RBPs, such as FMRP interact with miRNAs and components of the miRNA pathway, DICER and argonaute [86]. Thus, miRNAs and the mRNA sequences that they target play an important role in message degradation and translation.

RNA BINDING PROTEINS AND |MRNP COMPLEXES AT DENDRITES mRNA transport and translational repression are tightly coupled. Transport ribonucleoprotein particles (RNPs), RNA granules and processing bodies (P bodies) all contain mRNAs and RBPs that can repress translation [16,45,87]. RNPs function to transport mRNAs from the soma to dendrites in the absence of ribosomes. RNA granules are large assemblies of mRNAs, protein components of the translational machinery and RBPs involved in localization, stabilization and/or translational repression. In RNA granules, translation is repressed but ribosomes are present [47]. RNA granules are found in dendrites and neuronal somata, but formation likely begins in the nucleus because hnRNPs are present [47,88,89]. The best-characterized components include ZBP, FMRP, the double-stranded RNA binding protein staufen, the ELAV-like protein HuR and the DEAD-box RNA helicase RCK/Me31B/Dhh1p [16,90]. The RNA granule protein 105 (RGN105) represses translation, which can be reversed by BDNF [91]. P bodies are ribosome-free, cytoplasmic sites for mRNA decapping and turnover and transient storage of translationally repressed mRNAs

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[45,92]. Drosophila neuronal mRNPs are structurally and functionally related to P bodies and both types of granules contain staufen, FMRP, the non-sense mediated decay protein Upf1p, the RNA-degradative enzymes Dcp1p and Xrn1p/Pacman, the miRNA protein argonaute and the translation repression protein Dhh1p/Me31B [93]. Staufen1 regulates protein synthesisdependent LTP. Decreased levels of staufen1 impaired L-LTP and modified dendritic spine shape from regular to elongated spines, but did not change spine density [94]. Both Me31B/RCK and FMRP are core P-body components [95]. We would like to discuss in more detail the function of two well-studied RBPs, cytoplasmic polyadenylation element binding protein (CPEB) and FMRP. CPEB plays an important role in synaptic plasticity, learning and memory [96]. It contains an RNArecognition motif (RRM) and a zinc finger domain and recognizes cytoplasmic polyadenylation elements (CPEs) in the 3’-UTRs of mRNAs, such as maskin, and thereby modulates translation [96]. CPEB interacts with other RNA binding proteins, including p54 and Pum [97,98]. The Drosophila homologs of CPEB and FMRP interact [99]. CPEB has a prion-like form that has the greatest capacity to stimulate translation of mRNA suggesting that long-term synaptic changes are maintained by this physiological conversion [100]. Thus, CPEB may serve as a tag to mark active synapses. Over-expressed CPEB1 phosphorylation site mutants, Thr171 and Ser177, bind to CPE-containing mRNAs but do not stimulate translation and hence attenuate protein synthesis-dependent L-LTD [101]. This is accompanied by increased spine number and spine length in cerebellar neurons and suggests that CPEB1 is required for activity-driven protein synthesis and synaptic maturation [101]. FMRP is an mRNA binding protein that is absent in individuals with FXS. FMRP is ubiquitously expressed throughout the body, but with highest levels in the brain, gonads and young animals [102,103]. The protein has two heterogeneous nuclear ribonucleoprotein (hnRNP) K homology domains and one RGG box as well as nuclear localization and export signals [104,105]. FMRP is found in polyribosomes, transport particles, RNA granules, stress granules, P bodies and RISC complexes [106-108] where it functions as a repressor of translation [107,109-113]. The hypothesis that the mistranslation of one or multiple target mRNAs causes FXS has driven identification of the protein and mRNA binding partners of FMRP. To date, these include FXR1P, FXR2P, nucleolin, YB1/p50, Purα, staufen, NUFIP1, CYFIP1 and CYFIP2 as well as over 500 mRNAs with the potential to influence synaptic plasticity [109,114-116]. However, as some or even many of these mRNAs have not yet been defined by CLIIP [117], the true number may be lower. FMRP is locally translated at synapses in response to group 1 metabotropic glutamate receptor (mGluR) stimulation [118] or primary cortical neurons [67]. RNA targets of FMRP, such as postsynaptic density 95 (PSD95) and APP, are also rapidly translated in response to the group 1 mGluR agonist, DHPG [67,119]. PSD-95 is a scaffolding protein and its expression increases the number, size and maturation of dendritic spines [120]. APP is an integral membrane protein and the precursor of beta-amyloid (Aβ). FMRP is found in dendrites, dendritic branch points, the origins of spine necks, in spine heads, as well as in all known locations of neuronal polyribosomes where it associates with actively translating polyribosomes [121]. A missense mutation at I304N in the second KH domain of FMRP prevents polyribosome association [122]. FMRP represses the translation of microtubule associated protein 1B (Map1B) [123]. The lack of FMRP in Fmr-1 KO brain results in elevated Map1B levels and abnormally

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increased microtubule stability. Thus, FMRP plays a critical role in cytoskeleton organization. FMRP can function in the translational repression of mRNAs through protein/mRNA interactions [124]. Thus, FMRP plays important roles in mRNP complex formation, translation, dendritic spine structure and synaptic plasticity [125,126].

SIGNAL TRANSDUCTION PATHWAYS THAT MEDIATE MRNA/RBP INTERACTIONS AND PROTEIN BIOSYNTHESIS AT DENDRITES Cell signaling induces a variety of post-translational modifications to RBPs, which could provide a mechanism of epigenetic tagging of the mRNA pool. Epigenetic tagging of the genome is implicated in the formation of long-term memories [127], and it is likely that tagging of neuronal mRNAs and proteins contributes to synaptic plasticity. The pattern of DNA methylation and acetylation in conjunction with alterations in chromatin structure constitutes epigenetic modification [127]. Thus, the DNA/protein interactions involved in chromatin formation, which are regulated post-translationally, constitute an “epigenetic memory”. There are many structural and functional similarities between the nucleic acid/protein interactions important for transcription and translation. For example, the formation of transcription complexes on the promoters of genes targeted for active transcription allows for the recruitment of RNA polymerases and requires non-compacted DNA. Similarly, RBPs bind to and protect mRNAs from degradation as well as prevent translation as part of large mRNPs. Post-translational modifications of those RBPs in response to cell signaling alters the stability or constituents of those mRNPs such that rapid translational activation ensues. As some RBPs compact and translationally repress RNA while other RBPs form active complexes that recruit the translational machinery and polyribosomes to specific mRNAs, RBPs may provide similar functionality as DNA BPs. The stimulation of ion channels on the post-synaptic membrane by neurotransmitters and neurotrophic factors and the downstream involvement and activation of multiple scaffolding proteins and protein kinase pathways constitutes cell signaling and leads to alterations in cellular functions such as de novo protein synthesis [10]. Noradrenaline is an example of a neurotransmitter that is involved in memory [128]. Noradrenaline enhances the expression of monocarboxylate transporter MCT2 through translational activation via PI3K/Akt and mTOR/S6K [129]. The neurotrophic factor NT-3 regulates short-term plasticity through a de novo protein synthesis-dependent pathway at lateral perforant path-dentate granule cell synapses [130]. Glutamate stimulation rapidly increases translation of tissue plasminogen activator (tPA) [131]. Intracellular signaling molecules, such as cAMP, also contribute to synaptic plasticity and de novo protein synthesis. cAMP mediates LTP and LTD at CA3-CA1 synapses [132]. Brain-derived neurotrophic factor (BDNF) is the most well studied neurotrophic factor involved in neuronal cell signaling and LTP. BDNF produces long-lasting enhancement of synaptic transmission in the hippocampus [133]. Dendritic protein synthesis is stimulated by BDNF and completely blocked by anisomycin [134]. BDNF up-regulates 230 proteins in SN

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derived from cultured cortical neurons including those involved in translation [135]. De novo protein synthesis of FMRP, CaM-KIIα and Arc/Arg3.1 are induced in SNs by BDNF or glutamate receptor agonists [118,136,137]. BDNF initiates neuronal cell signaling that results in increased de novo protein synthesis through multiple pathways. BDNF modulates LTP through activation of TrkB receptor tyrosine kinases [138,139]. LTP induced by BDNF causes a rapid, transient phosphorylation of eIF4E and eEF2 and increased expression of eIF4E, which can be blocked by inhibition of MAPKERK [140]. Homer2 is localized to dendrites and locally translated at synapses in response to BDNF [141]. The activation of distinct voltage-gated ion channels by neurotransmitters, neurotrophins and ions are essential for the induction of various forms of LTP and LTD. NMDAR and AMPAR are upstream of LTP while mGluRs play a comparable role in LTD [142-144]. Stimulation of these membrane receptors results in conformational changes, which impact associated scaffolding proteins or kinases. Stimulation of NMDARs activates Src kinase signaling [145]. The Src family tyrosine kinases (Src, Fyn, Lyn, Lyk and Yes) are expressed in the central nervous system and bind to scaffolding proteins of the NMDA receptor complex. Src couples G-protein coupled receptors with downstream signaling molecules via the intermediary cell-adhesion kinase β (CAKβ), which activates Src thereby upregulating NMDAR in CA1 hippocampal neurons [146]. LTP induction can be blocked by inhibiting CAKβ [146]. NMDA receptor activation mediates phosphorylation and activation of CPEB [147]. The group 1 mGluRs (mGluR1 and mGluR5) are localized on the postsynaptic membrane and are excitatory, while group 2 and 3 mGluRs are primarily presynaptic and tend to suppress neurotransmission [148,149]. The group 1 mGluRs that mediate LTD signal through phosphoinositide 3-kinase, mTOR, MAPKERK and PKC [150-153]. Other receptors besides NMDAR, AMPAR and mGluRs are involved in neurotransmission. Stimulation of the μ-opioid receptor induces phosphorylation and activation of Akt and p70S6k and phosphorylation and inactivation of 4E-BP1 and 4E-BP2 [154]. The Akt signaling pathway is associated with neuronal survival while p70S6k, 4E-BP1 and 4E-BP2 are associated with translational control. M1 muscarinic acetylcholine receptor (mAChR) are involved in LTD in the CA1 region of the hippocampus, which is dependent on rapid protein synthesis as well as MAPKERK and mTOR [155]. Stimulation of the retinoic acid receptor (RAR) with all-trans-retinoic acid increases dendritic growth by approximately 2-fold within 30 min of stimulation through MAPK and mTOR pathways and increased dendritic translation [156]. Membrane receptors interact with scaffolding proteins on the inner side of the plasma membrane. RACK1 is an inhibitory scaffolding protein for the phosphorylation and function of NMDAR [157]. PSD-95 is a major scaffolding protein of the post-synaptic density at excitatory synapses, which is enriched in synaptically active neurons and reduced in inhibited neurons, in contrast to other PSD proteins, such as SAP102, Shank and GKAP [158]. PSD-95 can mediate the interaction of NMDAR with downstream signaling molecules. Homer interactions with mGluR are necessary for mGluR-induced LTD and translational activation [159]. The signaling pathways that regulate translation of mRNAs at dendrites has been an area of intense study. Signal transduction pathways that regulate LTP also regulate the translational machinery [160]. The mammalian target of rapamycin (mTOR) kinase

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contributes to long-term fear memory formation in the amygdala [161] and regulates the translation of many key proteins involved in synaptic plasticity. mTOR is a serine/threonine kinase that activates translation by phosphorylating 4E-BP1, which is eIF4E-binding protein [162-164]. Hyperphosphorylation causes dissociation of eIF4E and subsequent initiation of translation [165,166]. mTOR-mediated translation is dependent on PI3K and MAPKERK [167]. Several components of the ribosome recruitment machinery are targets of mTOR, including eIF4B, eIF4G, eIF4E, 4E-BPs, S6K, ribosomal protein S6 and eEF2. Components of the mToR pathway are present in dendrites [168]. Disruption of mTOR signaling by rapamycin reduces hippocampal late-phase LTP induced by high-frequency stimulation or BDNF, both of which require protein synthesis [168]. BDNF regulates the translation of Homer2 in the synaptodendritic compartment through activation of translation initiation via the mTOR-PI3K pathway [141]. PKA, PKC, CaM-KIIα and MAPKERK are intracellular kinases involved in LTP [169,170]. The early and late phases of LTP require activation of cAMP-dependent PKA, but only the late phase requires protein synthesis [171]. Serotonin regulates transcriptionindependent translation in Aplysia through protein kinase C, cAMP-dependent protein kinase (PKA), and a tyrosine kinase [172]. PKMζ is a form of PKC that is necessary and sufficient for maintaining LTP in the hippocampus [173,174]. The expression of pCaM-KII, BDNF, PSD-95 and zif268/egr-1 are significantly increased in the hippocampus of FVB/N mice after behavioral training, which supports the importance of these proteins for neuronal information storage [175]. zif268/egr-1 is necessary for protein synthesis-dependent late-phase LTP [176]. Rapid ocular dominance plasticity requires cortical protein synthesis [177]. Visual experience induces the translation of CaM-KIIα in the visual cortex, which is dependent on NMDAR [178]. CaM-KIIα contains CPEs, which are involved in glutamate-induced translation in cultured hippocampal cells [178]. CaM-KII regulates the synthesis and the phosphorylation of CPEB [179]. Co-activation of both MAPKERK and PI3K-Akt-mTOR after β1-adrenergic stimulation are required for the translation of striatal-enriched protein tyrosine phosphatase (STEP) [180]. Activation of β-adrenergic receptors facilitates the maintenance of LTP through MAPKERK and mTOR at the level of translation initiation [181]. MAPKERK inhibition blocks neuronal activity-induced translation and phosphorylation of eIF4E, 4EBP1 and ribosomal protein S6 [182]. Inhibition of MAPKERK blocks NMDA-induced protein synthesis in dendrites [183]. Group 1 mGluR activation causes phosphorylation of MAPKERK in WT SN and dephosphorylation in Fmr-1 KO SN [184]. Other kinases have also been implicated in synaptic plasticity. For example, p21activated kinase 3 (PAK3) knockout mice exhibit learning and memory deficiencies and impaired late-phase LTP. These mice have a large reduction in the active, phosphorylated form of cAMP-responsive element-binding protein (CREB) [185]. The expression of constitutively active CREB facilitates the late-phase of LTP [186]. Another example is the protein kinase GCN2, which regulates the initiation of translation. GCN2 regulates synaptic plasticity through modulation of the ATF4/CREB pathway [187]. Activated GCN2 induces translation of ATF4, which is an antagonist of CREB. Under basal conditions, GCN2 and eIF2α are phosphorylated, ATF4 levels are high and CREB-dependent transcription is repressed. Phosphorylation of eIF2α selectively enhances translation of ATF4 while inhibiting overall translation [188]. Changes in the phosphorylation status at a single site

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(serine 51) on eIF2α modulates synaptic plasticity and memory storage [27]. Decreased eIF2α phosphorylation facilitates while increased phosphorylation impairs the induction of L-LTP and LTM [27]. Adenyl cyclase activation is involved in LTP and translational regulation at the elongation step [189]. Thus, many proteins are involved in translational regulation and synaptic plasticity with CaM-KII, PKMζ, MAPKERK, Fos, C/EBP and CREB implicated as memory master control molecules [190].

PATHOLOGY OF SYNAPTIC PLASTICITY The morphology and density of dendritic spines is pivotal to synaptic plasticity. In FXS the loss of a single protein, FMRP, results in altered dendritic spine morphology, density and plasticity [191,192]. FXS is the most prevalent form of inherited mental retardation, affecting 1 in 4,000 males and 1 in 8,000 females. This X-chromosome-linked disorder is characterized by moderate to severe mental retardation (overall IQ2-fold more secreted APPα than children without autism [219]. Individuals with trisomy 21/DS overexpress APP mRNA by four- to five-fold, deposit extracellular amyloid at a greatly accelerated rate [220], and develop senile plaques twenty to thirty years earlier than normal individuals [221]. Increased translation of APP provides more targets for cleavage by β- and γ-secretases and hence increased Aβ accumulation. A century after the discovery of Aβ, what do we know about the RNA/protein interactions that regulate APP mRNA stability, decay and translation and hence contribute to APP and Aβ production? APP mRNA decay and translation are post-transcriptionally regulated through ciselements in the 5’-UTR, coding region and 3’-UTR of the mRNA and mediated by RBPs that bind to these elements in response to cell signaling. APP mRNAs (70% of APP695 and 50% of APP 751/770) are associated with polyribosomes in rat brain [222], suggesting that translational regulation could play an important role in APP production. There is an average of 8 ribosomes per polyribosome with 1-2 polyribosomes per spine synapse in the hippocampus and gentate gyrus [21] with polyribosomes predominantly located at the bases of spines [21,22]. The length of APP mRNA can be estimated at 1.07 μm (mouse APP695 mRNA) suggesting that only 1 mRNA at a synapse is translated at a time and that there is competition between mRNAs for the translational machinery [223]. Translational regulation of APP mRNA occurs through at least two identified cis-elements. The first identified was a 90-nucleotide cis-element in the 5’-UTR mapping from +55 to +144 from the 5’-cap site. This translational control element is homologous to the iron-response elements (IRES) found in the light and heavy ferritin genes. APP translation is also responsive to IL-1α and IL-1β, but does not alter steady state levels of APP mRNA [224]. Translational control of APP mRNA also occurs through FMRP [119]. FMRP expression increases in the barrel cortex of the rat after unilateral whisker stimulation, a model of experience dependent plasticity [225]. FMRP is phosphorylated N-terminal to the RGG box and phosphorylation/dephosphorylation status of the protein correlates with binding to stalled versus active polyribosomes [226]. The RGG box of FMRP binds to intramolecular G-quartet sequences while the KH2 domain has been proposed to bind to kissing complex RNAs based on in vitro selection assays [116,227]. APP mRNA contains a putative G-quartet cis-element in its coding region (position 825-846 of the mouse sequence) embedded within a guanine-rich domain (nucleotides 694-846). FMRP binds directly to the 5’ end of this guanine-rich region prior to the putative G-quartet sequence [119]. FMRP also protects a 29 base instability element in the 3’-UTR of APP mRNA from ribonuclease digestion [119]. FMRP binds to uridine-rich mRNAs [228,229], and the 29 base element is uridine rich, but FMRP likely protects this region through protein/protein interactions with nucleolin and not by direct binding to the mRNA. No

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kissing complex-like elements have been identified in APP mRNA. Stimulation of WT SN with the mGluR agonist DHPG rapidly releases FMRP from APP mRNA and de-represses translation, an effect not seen in Fmr-1 KO neurons [119]. In KO SN or neurons, APP is constitutively elevated. This data suggests that FMRP represses the translation of APP through mGluR-dependent interactions with APP mRNA. Consistent with constitutively elevated APP levels, there are increased Aβ1-40 and Aβ1-42 in Fmr-1 KO mouse brain. Increased APP and/or APP proteolytic products could provide a common denominator at the molecular level for the impaired synaptic plasticity observed in AD, FXS, autism, DS and epilepsy. FMRP mRNA and protein expression are down regulated as a function of aging in the mouse brain [230], suggesting that repressed transcripts, such as APP, would be upregulated, a well-known phenomenon in animals and humans. APP over-production favors the β-secretase pathway. and alterations in APP processing have a detrimental effect on synaptic plasticity. Expression of the 104-amino acid carboxy-terminal fragment of APP, which contains the Aβ region, impairs spatial learning and maintenance of LTP in mice [231].A recent report indicates that soluble APPα is elevated in the sera of severely autistic children with the highest levels in two children who were also FXS [219]. In addition, all of these neurological disorders are associated with an increased prevalence of seizures. The underlying molecular mechanism(s) that cause seizures are not well understood. Epilepsy in the aging population is a significant, but often overlooked, clinical problem [232], and recent studies indicate that “silent seizures” are a characteristic of AD [233]. Seizures are 6-10X higher AD than in an age-matched population [234,235] with incidence increased in mild-tomoderate AD [236] and in early-onset AD with familial presenilin mutations [237]. 10-22% of AD patients have at least 1 unprovoked seizure [237]. The cumulative lifetime risk of unprovoked seizures is 4.1% in the general population with a higher incidence in the elderly [238]. FXS mice have a high propensity for audiogenic seizures [239], and two mouse models for AD have an increased susceptibility to both chemically induced and audiogenic seizures (AGS) [240,241]. Thus, APP or a proteolytic product of APP could contribute to seizure induction and severity.

CONCLUSION Localized protein synthesis in dendrites is essential for learning and memory. Understanding the mechanisms of protein/mRNA interactions in and localization to dendrites and their roles in selective protein biosynthesis are challenges for the 21st century. Refining our understanding of the mechanism of these pivotal interactions will define the molecular basis of learning and memory. This knowledge will also form the basis for the rational design of therapeutics to treat neurological disorders that result from aberrant protein expression and/or activity at synapses. Synaptic plasticity is required for normal learning and memory and is impaired in disorders such as FXS. Defining the role(s) and regulation of key dendritic proteins, such as FMRP, will be necessary to designing therapies for FXS and related disorders. Localized protein synthesis within individual dendrites regulates synaptic function and neurite outgrowth and guidance. FMRP is synthesized locally in response to synaptic activity and is likely a major player in the selective translation of dendritic mRNAs. In FXS,

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mRNAs that are normally repressed by FMRP, for example APP and Map1B, lose pulsatile expression and are constitutively elevated, resulting in loss of synaptic plasticity. Unraveling the mystery of the “RNA epigenetic memory” as regards FMRP/APP mRNA interactions will likely provide therapeutic targets for the treatment of AD, FXS, DS, autism and seizures as well as a better understanding of the normal processes that contribute to protein biosynthesis in learning and memory.

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[239] Chen L, Toth M. (2001) Fragile X mice develop sensory hyperreactivity to auditory stimuli. Neuroscience 103(4): 1043-1050. [240] Westmark CJ, Westmar PR, Beard AM, Hildebrandt SM, Malter JS. (2007) Seizure susceptibility and mortality in mice that over-express amyloid precursor protein. IJCEP 1(2): 157. [241] Westmark CJ, Westmark PR, Malter JS. (2008) Manuscript in preparation.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter IX

SECRETED PROTEIN AND PEPTIDE BIOSYNTHESIS: PRECURSOR STRUCTURES AND PROCESSING MECHANISMS Sergey A. Kozlov∗, Alexander A. Vassilevski and Eugene V. Grishin Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, 117997 Moscow, Russia.

ABSTRACT Protein biosynthesis is rarely restricted to mRNA translation into an amino acid sequence. To yield the mature form, most proteins undergo various posttranslational modifications due to the action of different enzymes. Certain combinations of amino acid residues (primary structure motifs) have been defined to guide the sequence of modifications during the process of precursor protein maturation into the final product. In this chapter, we specifically focus on the secreted polypeptide maturation. For a number of precursor sequences retrieved from UniProt databank, complete sets of enzymes have been identified that execute processing of secreted polypeptides. This finds reflection in the amino acid sequences of the corresponding protein precursors that carry information about the queue of posttranslational events in the form of specific motifs arrangement. Extensive data analysis allowed us to propose a simple set of principals that facilitate effective sequence information handling. Utilization of the proposed principals significantly improves mature protein sequence prediction from available gene structures. We also address the problem of known motif identification and novel motif prediction from large sets of data. A number of proteins are considered in greater detail as examples of the proposed principals utilization conveniences.

∗

Correspondence concerning this article should be addressed to: S.A. Kozlov, [email protected]; tel.: +74953366540; fax: +74953307301.

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I. MAIN STAGES OF POLYPEPTIDE MATURATION PROCESS A wealth of different secreted proteins apart from signal peptide removal upon entry into the endoplasmic reticulum (ER) are subjected to further modifications along the secretory pathway. These modifications may imply a single or multiple steps of the polypeptide chain fragmentation, glycosylation, phosphorylation, acylation as well as other more or less common posttranslational events. In most cases, however, the so-called limited proteolysis due to the action of specific endopeptidases constitutes the crucial step of active polypeptide formation [1-5]. Polypeptides carry important functions in metabolism, immunity, development, signaling, etc., therefore, strict control over their genes’ expression level and maturation process is critical. This “quality control” function is in part performed by proteolytic enzymes [6-11]. Cell surface receptors, serum proteins, enzymes, neuropeptides, hormones, growth factors, and cytokines are among the natural “substrates” of limited proteolysis [12-16]. Accuracy of the enzymes involved in polypeptide maturation is strictly necessary for the organism survival, and flaws in their activity may lead to different pathologies including cancer [9,10,17,18]. Protein precursor amino acid sequence analysis in many cases permits identification of a doublet of positively charged residues exactly preceding the site of polypeptide chain degradation. In 1984, the first enzyme was shown to cleave this kind of site in yeast when αpheromone processing was addressed [1]. The new enzyme was named kexin and was found able to cleave large precursor molecules not only from yeast but also from mammals in vitro. Furin belonging to the same family of subtilisin-like serine endoproteinases became the first discovered mammalian processing enzyme [19]. A large number of related proteins were later found in mammals and named subtilisin-like proprotein convertases (SPC). Several types of SPC that differ both in size and subcellular localization have been described [20]. SPC are irregularly distributed in tissues; the most widespread and best studied remains furin or SPC1 that is found inside cells in the Golgi network, secretory vesicles, endosomes as well as on the cell surface. Throughout, the nomenclature of Schechter and Berger [21] will be followed in designating the cleavage sites of substrates as …P4-P3-P2-P1 ↓ P1’-P2’-P3’-P4’…, etc., with the scissile bond between P1 and P1’ and the C terminus of the substrate on the prime site. In case of SPC, the P2 position is usually represented by positively charged residues Arg or Lys, although Pro was revealed effective by phage display screening of substrates for SPC1 [22]. The P1 position is the most conserved and is most often occupied by Arg, although Lys is also possible [23]. The necessary conditions of proteolysis also include occurrence of another positively charged residue in position P6 or P4 [24,25]. Besides subtilisin-like proprotein convertases specifically recognizing doublets of positively charged residues, a number of other enzyme types are known to participate in limited proteolysis of precursor proteins. In these cases, Arg does not form the scissile bond. Pyrolysin-type subtilisin/kexin isozyme 1 (SKI-1) also known as site 1 protease (S1P) predominantly cleaves proproteins at Leu residues (P1) with Arg in position P4 [26,27]. Neural apoptosis-regulated convertase 1 (NARC-1) also known as SPC9 has not been

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assigned defined substrate specificity, yet a polar residue in position P1 is preferable and a hydrophobic stretch in the P5-P2 region is thought essential [28,29]. Specific enzymes of plant origin that perform limited proteolysis have also been described. These enzymes have been shown to play important roles in processes of plant development and storage protein hydrolysis. Vacuolar processing enzymes (VPE) are cysteine proteinases that have been found to selectively cleave vacuolar proteins such as legumin-type globulins (12S globulins) and napin-type albumins (2S albumins) at Asn residues (P1) in different plants [30,31]. A number of processing subtilases (subtilisin-like serine proteases) have been described: LeSBT1 from tomato [32], C1 from soybean [33] and two highly related saspases from oat [34]. Saspases just like VPE cleave scissile bonds formed by Asn residues and have been implicated in the proteolytic cascade of programmed cell death [34]. Subtilases are utilized by plants on the primary stages of storage protein digestion. C1 protease cleaves Glu-Glu and Glu-Gln scissile bonds [33], whereas LeSBT1 cleaves at Gln residues (P1), although its specificity has not been studied in detail and the recognized motif is thought to include other regulatory residues [32]. Table 1 lists motifs recognized by some extensively studied enzymes that selectively cleave precursor proteins. Table 1. Motifs recognized by processing enzymes Substrate position Origin

Enzyme P6

fungi

P5

K/R

X

SPC2, SPC3, SPC4, SPC6

P2

P1

↓

K/R

R

↓

R

X

K/R/P

R

↓

X

X

K/R

R

↓

R

X

K/R

R

↓

K

X

X

R

↓

SPC5 K

X

K

X

X

R

↓

R

X

X

X

K/R

R

↓

R

X

L/I/V

L/T/K

↓

L

V

F

A

Q

↓

Y/I

V

V

V/L

L/M

↓

VPE

N

↓

saspases

N

↓

subtilase C1

E

↓

subtilase LeSBT1

Q

↓

SPC7 SKI-1 (S1P) NARC1 (SPC9)

plants

P3

kexin furin (SPC1)

animals

P4

P1’

E/N

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Precursor protein polypeptide chain cleavage by convertases usually does not represent the final stage of active molecule maturation. The newly formed fragments in most cases carry excess positive charge at their C-termini that may interfere with the mature molecule function and, moreover, inhibits the process of limited proteolysis by SPC itself. This negative effect is removed due to C-terminal trimming of the fragments by carboxypeptidases (CP) specific to positively charged residues, carboxypeptidase D (CPD) or E (CPE) in most cases [35-37]; the productivity of convertases increases accordingly as was shown in vitro for SPC2 + CPE [38] and in vivo in CPE-defective knockout mice [39]. Besides C-terminal trimming, gradual stepwise fragmentation of the N-terminal sequence often occurs during polypeptide maturation. One of the most widespread enzymes that catalyzes this process is dipeptidyl-peptidase IV (DPP IV/CD26) that plays a role in regulation (inactivation) of biologically active peptides as well as in polypeptide maturation [40]. The enzyme cleaves off a pair of N-terminal residues Xaa-Pro or Xaa-Ala and the corresponding maturation motifs are usually represented by several tandem dipeptide repeats. Another important process in active molecule maturation is cleavage of the C-terminal Gly residue with simultaneous amidation of the preceding residue. This often leads to formation of a more stable and/or active form of the target molecule. The process is catalyzed by peptidylglycine α-amidating monooxygenase (PAM), a complex of two enzymes that act successively: peptidylglycine α-hydroxylating monooxygenase (PHM) that converts Gly residue into an α-hydroxylated intermediary product, and peptidyl-α-hydroxyglycine αamidating lyase (PAL) that catalyzes glyoxylate cleavage and amide formation [41-44]. As stated above, positively charged polypeptides are inclinable to inhibit proprotein convertases. Many highly basic peptides, for example olygoarginine (5-9 residues), represent a special class of SPC inhibitors [45,46]. Histidine-rich antibacterial peptides from human saliva histatin 3 (32 residues, 22% His) and histatin 5 (24 residues, 33% His) are able to inhibit convertases like furin and SPC7 whereas serve substrates for SPC3 [47]. Nevertheless, a great number of biologically active polypeptides are known to carry large positive charge with positively charged residues being indispensable for their function. These polypeptides are mainly represented by antimicrobial peptides and toxins. Due to their high activity and possible toxicity for the producing cells these molecules are usually produced in the form of inactive precursors with auxiliary stabilizing and protective fragments. Specialized convertases are needed to correctly cleave such Arg/Lys-rich polypeptide chains without unwanted fragmentation of the mature active molecules. In certain cases subtilisin-like convertases are utilized for the cause. For example, some bacterial and viral protein toxins such as anthrax toxin protective antigen, diphtheria toxin, clostridial α-toxin and hemagglutinin of a virulent avian influenza virus are known to get activated by SPC after their interaction with the target cell and/or endocytosis [6,48-50].

II. PRECURSOR AMINO ACID SEQUENCE ENCODES THE MATURATION PATHWAY The advent of powerful tools of bioinformatics and public databases of nucleotide and protein sequences forms the modern basis of protein maturation pathways investigation.

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Comparison between mature polypeptide sequences obtained experimentally and the corresponding genes enables identification of all stages of posttranslational modification. Conversely, given that all possible maturation events are known and the corresponding motifs of amino acid sequence established, prediction of active molecule structures from gene becomes reliable. This approach is illustrated by two examples – pro-opiomelanocortin (POMC, P01189), precursor of a number of hormones in humans [51], and prepromelittin (P01501), precursor of melittin, the principal component of the honeybee venom [52] (Figure 1). The primary cleavage of a secreted polypeptide precursor takes place cotranslationally. The leader peptide is cleaved off by the signal peptidase associated with the ER translocon. The widely used SignalP 3.0 program (available at www.cbs.dtu.dk/services/SignalP/) offers a robust method of signal peptide prediction. The next step of limited proteolysis is performed by different convertases that are characterized by a common recognition motif (K/R)-Xn-(K/R), where X – any residue except Cys; n = 0-4. A strong preference for Arg in the last position and n = 0 give the more common doublet motif (K/R)-R. In the given example, POMC has 9 such motifs (the identified cleavage sites are marked with red vertical arrows in Figure 1). Due to differential convertase specificity, however, formation of longer fragments corticotropin and β-lipotropin as well as shorter fragments (α-melanotropin, corticotropin-like intermediary peptide, γ-lipotropin, β-melanotropin, β-endorphin) is achieved. The newly formed molecules that still carry the convertase recognition motifs are subsequently trimmed C-terminally due to the action of CP (basic residues are cleaved off, blue left arrows in Figure 1). The peptides are also trimmed N-terminally by DPP (shown for melittin precursor, blue right arrows in Figure 1). Finally, PAM converts exposed C-terminal Gly residues into amide groups (red left arrows in Figure 1).

Figure 1. Precursor primary structure defines the maturation process. Amino acid sequences of proopiomelanocortin (P01189) and prepromelittin (P01501) are shown. Mature active molecules are marked with rectangles; convertase recognition motifs are shown in bold and the corresponding cleavage sites are marked with red vertical arrows. Residues cleaved off by dipeptidyl-peptidase are marked with blue right arrows; by carboxypeptidases, blue left arrows; by peptidylglycine α-amidating monooxygenase, red left arrows.

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To sum up, the primary structure of precursors not only specifies the set of processing enzymes but also defines their order of action.

III. NOVEL MOTIFS OF LIMITED PROTEOLYSIS In a large number of deduced amino acid sequences of precursor proteins the classical convertase-recognition motifs cannot be identified, although the N-terminal residue of mature molecules is known from experiment. For example, this is true for many toxins and antimicrobial peptides. It is obvious that processing is carried out by some specific endoproteolytic enzymes that have not been identified yet. The large information content, however, indicates a wide distribution of novel processing motifs and therefore suggests existence of the predicted enzymes in different organisms. In most precursors of spider toxins, the P1 residue is Arg, and one or more Glu residues are found in the P4-P2 region. This motif was named Processing Quadruplet Motif (PQM) [53]. It was later discovered that the same motif is found in other unrelated polypeptide precursors and should be extended to further include positions up to P6. A second similar motif was identified that also specifies polypeptide cleavage at Arg (P1) but has one or more Glu residues in the P1’-P5’ region. By analogy, this motif was named inversed Processing Quadruplet Motif (iPQM) [54]. Examples of precursors processed at PQM and iPQM are shown in Figure 2.

Figure 2. Novel motifs PQM and iPQM in polypeptide precursors. Amino acid sequences of precursors of ω-agatoxin IA (P15969) and acrorhagin II (Q3C256), and maximins-S type A precursor (Q5GC94) are shown. Mature active molecules are marked with rectangles; PQM and iPQM are marked with large rose right arrows and large blue left arrows, respectively, the corresponding cleavage sites are marked with red vertical arrows. Residues cleaved off by carboxypeptidases are marked with blue left arrows; by peptidylglycine α-amidating monooxygenase, red left arrows.

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Protein precursor cleavage at PQM/iPQM can result either in mere pro-sequence separation, or in extended polypeptide fragmentation. The latter case includes formation of a number of mature chains that can either be held together by disulfide bridges (ω-agatoxin IA in Figure 2) or represent independent functional molecules (maximins in Figure 2). In a number of cases, the PQM/iPQM contains a basic residue in position P2 that corresponds to the kexin recognition site. However, the lack of additional basic residues in the P4/P6 positions (Table 1) and the episodic character of such dibasic motif occurrence render processing of the corresponding precursors by SPC of a known type highly improbable. The common feature of the new motifs and the conventional convertase recognition sites is the high selectivity for Arg in the P1 position. It could be proposed that the predicted new types of processing enzymes are structurally related to subtilisin-type convertases. The name protoxin convertases (PTC) is suggested to describe the new type enzymes since the new motifs most often occur in precursors of molecules that exhibit toxic functions (neurotoxins, cytolytic peptides, etc.). Although a slim possibility exists that the same enzyme recognizes both PQM and iPQM, joint operation of a pair of enzymes – PTC “direct” and PTC “inversed” – seems more reasonable. The subcellular localization of PTC might differ from that of SPC. It is probable that toxin processing takes place in specific vesicles, or even extracellularly. In the latter case, both PTC and toxin precursors could be secreted into the gland duct. As discussed above, excess positive charge of the polypeptide chain has a negative effect on SPC activity. However, many active molecules rely on this feature and their highly basic nature is indispensable for their function. Therefore, the predicted PTC bridge the gap and are suggested to play a leading role in processing of precursors of basic molecules.

Figure 3. Complex precursor processing. The chart shows processing of two types of precursors: those containing the conventional R(K)toR motifs by SPC and those with the symmetrical EtoR/EafterR motifs by predicted PTC.

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IV. REGULAR AND SYMMETRICAL MOTIFS OF LIMITED PROTEOLYSIS Most processing motifs identified to date have an Arg in the P1 position. In case of SPC, another basic residue, Arg or Lys, is located upstream up to position P6. In order to simplify primary structure analysis, the term “arginine cleavage by basic control” or R(K)toR in abbreviated form was introduced to describe this set of motifs [54]. Following this principle, processing of precursors of a number of signaling molecules, neuropeptides and hormones, such as insulin, gastrin, endorphins, enkephalins, corticotropins, etc. occurs [55-58]. In case of the predicted PTC, the correct Arg-Xaa scissile bond is defined by a Glu residue located upstream up to position P6 for PQM and downstream up to position P5’ for iPQM. By analogy, these motifs were termed “arginine cleavage by glutamic acid control” [54]. Since these motifs are actually symmetrical with respect to the P1 residue, they have the short names EtoR and EafterR. The symmetrical nature of the EtoR/EafterR motifs has interesting consequences that are most evident in the structure of the so called complex precursor proteins [59] that get processed into a number of active molecules (Figure 3). The resultant mature polypeptides are usually relatively small and may be functionally similar as well as have distinct functional properties. For example, the precursor of the neurotensin-like octapeptide xenopsin is also processed into the linear antimicrobial peptide XPF (xenopsin precursor fragment) [60]; another example is the maximins-S type A precursor shown in Figure 2. By convention, the amino acid sequence of polypeptide precursors can be split into two parts, the target mature functional part and the “ballast” auxiliary part. In many cases the biological role of the auxiliary part is equally important and can be regulatory, inhibitory, structural, etc. One of the most important functions of the auxiliary part is to ensure correct folding, trafficking and processing of the functional part. Limited proteolysis motifs are usually located in the “ballast” part of the precursor polypeptide chain. In case of the R(K)toR motif recognition, the corresponding amino acid residues are removed from the N-terminus of the newly formed mature chain, but can be (sometimes partially) retained at the C-terminus. The symmetrical EtoR/EafterR motifs are typically present in pairs: the EtoR motif is always found in the N-terminal part of the precursor with respect to the mature sequence, whereas the EafterR motif always resides in the C-terminal part. This characteristic motif arrangement secures removal of the auxiliary amino acid residues within the “ballast” fragments. Following the final stage of maturation, the active molecules contain no additional “unwanted” residues (Figure 3).

V. PROCESSING ENZYME TYPE CAN BE PREDICTED FROM THE PRECURSOR PRIMARY STRUCTURE Precursor amino acid sequence analysis to locate the motifs of limited proteolysis and divide the functional and auxiliary parts does not always unambiguously determine the type of the processing enzyme. For example, the P6-P5’ region can simultaneously contain both

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the R(K)toR and EtoR/EafterR motifs (Figure 4). In other words, both Arg/Lys and Glu residues can be found in close proximity to the Arg residue in the P1 position. If it is a complex precursor with several processing sites then it is logical to assume that the same type of enzyme is utilized to cleave all scissile bonds. If all of the processing motifs are ambiguous, the amino acid sequence of the mature product should be addressed. Whereas the “ballast” auxiliary part can contain any number of processing sites and get cleaved into any number of fragments, the target mature functional part should be virtually devoid of such motifs in order to escape possible undesirable proteolysis and retain structural integrity.

Figure 4. Primary structure analysis of polypeptide precursors. Pro-opiomelanocortin (P01189) and precursors of PBAN (P09971), insulin (P01308), gomesin (P82358), apamin (P01500), huwentoxin III (P61103), imperatoxin I (P59888), latarcin 4a (Q1ELU5), magainin (P11006) and toxin Am 1 (P69929) are considered. In each case, partial amino acid sequences that contain the indicated processing sites and the motifs of limited proteolysis are shown. The split sites located in the analyzed sequences are shown inside the corresponding mature chains. Compact fold stands for mature polypeptide sequence containing a number of Cys residues and able to form a closely packed structure. Ballast peptides represent the auxiliary part of the precursors removed during maturation.

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Mature polypeptide sequence analysis reveals the presence of “unwanted” convertase recognition motifs that could be cleaved during biosynthesis; these motifs were named “split sites” [54]. The final choice of the processing enzyme type should be made on the basis of absence of the corresponding split sites in the mature chain(s). Quite often, however, when the mature product represents a closely packed structure, stabilized by disulfide bridges for instance, the possible processing motifs – split sites – are buried and protected from cleavage. One of the most evident examples is the structure of polypeptide neurotoxins. The closely packed structure named “compact fold” is most probably stable to limited proteolysis even if it contains a split site. Moreover, incorporation of split sites into the mature sequence might have a certain “quality control” function: incorrectly folded mature products will have such sites exposed and accessible to proteolysis. A totally different situation is seen in case of short linear (no Cys residues) peptides or molecules having extended loops: the split sites are always exposed and the type of the processing enzyme can be deduced directly. Assigning the correct type of limited proteolysis as a result of the known precursor sequences analysis has practical value and can be utilized to predict mature sequences. A closer attention should be paid to the possibility of occurrence of multiple mature chains in a single precursor. The evolutionary rationale often leads to complex precursor design where all mature sequences are difficult to infer. In this case the regular R(K)toR and symmetrical EtoR/EafterR motifs location allows a detailed analysis to predict mature polypeptide sequences. An interesting family of single-chain linear insectotoxins was discovered, each member consisting of a pair of typical short linear cytolytic peptides linked together by a mutated EtoR motif [61]. The lack of the key Arg residue in the P1 position abolishes the polypeptide chain fragmentation and the toxins are suggested to represent an example of complex precursor evolution towards a more simple structure with novel functions.

VI. POLYPEPTIDE PRECURSOR PRIMARY STRUCTURE ANALYSIS Polypeptide amino acid sequences were retrieved from the fully annotated entries in the Swiss-Prot section of the UniProt Knowledgebase. In total, 39,903 sequences of polypeptide precursors from different organisms were found in UniProtKB/Swiss-Prot (as of January 2008). Among these, 6,571 sequences from multicellular organisms were preproproteins (or prepropeptides) with one or more pro-sequences and were further inspected to locate the motifs of limited proteolysis recognized by furin-type convertases SPC and predicted PTC, as well as DPP, CP and PAM. 5,168 sequences were identified as simple precursors with just one mature chain and at least one pro-sequence (“ballast” peptide). The auxiliary “ballast” part may be located both N- and C-terminally. 770 and 109 precursors were found to contain the R(K)toR and EtoR motifs, respectively, and 407 sequences contained both motifs. These 407 sequences were inspected in greater detail. The split site and compact fold analysis assigned 102 sequences to the R(K)toR motif, and 65 sequences to the EtoR motif. In summary, primary structure

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analysis of simple precursors identified 872 R(K)toR motif-containing, 174 EtoR motifcontaining sequences; 240 sequences could not be assigned a preferable motif. 908 sequences were found to contain two mature polypeptide chains (heavy + light chains of a single molecule, active component + peptide of unknown function, or two active components) and were named “binary” precursors. In each sequence, four possible cleavage sites were examined: N-terminal with respect to the first mature chain, both N- and Cterminal in the fragment between the mature chains, and C-terminal with respect to the second mature chain. The EtoR/EafterR symmetrical motifs were additionally checked to reside inside the “ballast” part; as described above, this was not needed in case of the regular R(K)toR motifs. The processing type was reliably assigned to the sequences with all four sites suiting the described conditions. In total, 446 “binary” precursors were identified as R(K)toR motif-containing, 19 as EtoR/EafterR symmetrical motif-containing and 69 sequences could not be assigned a preferable processing type. 495 sequences were complex precursors processed into three or more mature polypeptide molecules. Among these, the ice-structuring glycoprotein precursor (P24856) contains 46 annotated mature chains, whereas the most composite precursor processed by convertases – LWamide neuropeptides precursor from a sea anemone (Q16992) – contains 36 annotated mature peptides. The sequence analysis was carried out as described above for the “binary” precursors, the total number of the analyzed sites equaled the number of mature chains multiplied by 2 (to inspect both N- and C-terminal processing events). A fairly common feature of such extended precursors was that although a great majority of mature chains were flanked by specific processing motifs, one or several mature sequences were not. One explanation is that some of the annotated mature molecules were only predicted from similarity and not isolated experimentally. Another possibility is that additional processing events under the action of unknown enzymes are operating in these cases. As a result, 203 complex precursors were found to get processed at R(K)toR motifs, 9 sequences at EtoR/EafterR motifs, and 39 sequences were ambiguous. The primary structure analysis revealed that for the majority of known active mature polypeptides the corresponding precursors are processed by SPC at the R(K)toR motifs (further referred to as type 1 maturation). Another group is formed by precursors that are processed by predicted PTC at the EtoR/EafterR motifs (type 2 maturation). A number of members from the second group are listed in Table 2. Uncommon maturation motifs have been noticed earlier for some precursor polypeptide sequences. Thorough investigation of the type 2 maturation was performed for the first time in the present work. It was not noticed before that the specific EtoR/EafterR motifs were present not only in toxin precursors. The analysis revealed that apart from neurotoxin precursors (51 spider toxin precursors and 10 conotoxin precursors annotated in UniProt), diverse precursors of antimicrobial peptides (37 precursors from spiders, insects, fish, amphibians and mammals), different types of enzymes (41 proenzymes mainly from spiders, insects and mammals), neuropeptides and hormones (48 proproteins mainly from mammals) and some pore-forming toxins from marine organisms (4 from sea anemones and 1 from cone snails) all carry the EtoR/EafterR maturation motifs.

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Table 2. Some polypeptide precursors containing EtoR/EafterR motifs Source organism taxonomy

Mature polypeptide

Reference

Activity

Name

Amphibia

antimicrobial peptides

brevinin, dermaseptins, magainins, ranatuerin

[62-65]

Anthozoa

phospholipase A2

AcPLA2

[68]

Anthozoa

pore-forming agents

Cytolysins, equinatoxins

[66,67]

Arachnida

antimicrobial peptides

cyto-insectotoxins, latarcins

[59,61]

Arachnida

neurotoxins

omega-agatoxin, BmCa-1, huwentoxins, neurotoxins Ph3X, plectoxins

[98,102105]

Arachnida

phosphodiesterases

dermonecrotic proteins

[106]

Arachnida

phospholipase A2

phaiodactylipin

[97]

Aves

regulatory protein

inhibin beta A

[73]

Aves

structural protein

fibrinogen

[72]

Crustacea

neurohormone

mandibular organ-inhibiting hormone

[74]

Echinoidea

neuropeptides

exogastrula-inducing peptides

[99]

Gastropoda

neurotoxin

T-1-conotoxin

[94]

Gastropoda

pore-forming agents

echotoxins

[95]

Insecta

antimicrobial peptide

lebocin

[100]

Insecta

neuropeptides

AeaHP, leucokinin

[79,80]

Insecta

non-toxic compound

secapin

[71]

Insecta

phenoloxidase

PO 2

[101]

Insecta

phospholipase A2

Api m I

[70]

Insecta

protease

trypsin iota

[78]

Mammalia

antimicrobial peptide

defensin-related cryptdin-6/12

[93]

Mammalia

GDNF family receptor

GFR-alpha-4

[92]

Mammalia

hormone

insulin-like growth factor II

[86]

Mammalia

interleukin

IL-1 alpha

[87]

Mammalia

neuropeptides

galanin-like peptide, neuropeptide S

[90,91]

Mammalia

proteases

acrosins, elastase, granzyme A

[82-85]

Mammalia

regulatory proteins

ghrelin, osteocrin, obestatin

[88,89]

Mollusca

pheromones component

enticin

[69]

Pisces

antimicrobial peptides

moronecidins

[75]

Pisces

hormone

C-type natriuretic peptide 1

[76]

Pisces

kininogenases

natterins

[77]

Polychaeta

antimicrobial peptides

arenicins

[81]

Trematoda

peptidase

hemoglobinase

[96]

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Figure 5. Maturation motifs in precursors grouped according to mature molecule functions. The numbers of precursors with the R(K)toR maturation motifs (green bars), the EtoR/EafterR maturation motifs (blue bars) and ambiguous (cyan bars) are indicated above the corresponding bars. AMP – antimicrobial peptides.

As was noted above, type 1 fragmentation and the corresponding R(K)toR motifs vividly prevail. The analyzed sequences were grouped according to the mature product function. Figure 5 shows the occurrence of the type 1 vs. type 2 maturation motifs inside six groups of precursors of molecules with certain functions. Type 1 R(K)toR motif is predominant in each group. The type 2 EtoR/EafterR motif-containing sequences constitute a large part of precursors for antimicrobial peptides and cytolytic toxins only; notably, together with the unassigned sequences they outnumber the conventional R(K)toR-containing precursors. It was also found that no more than one third of known toxins and even a smaller fraction of enzymes and regulators (neuropeptides and hormones) follow type 2 maturation. For four groups of precursors with a certain mature product function diagrams showing maturation motifs distribution among taxonomic groups of organisms were constructed (Figure 6). In precursors of antimicrobial peptides and cytolytic toxins, type 2 fragmentation prevails in most animals with the exception of insects and mammals with the classical type 1 maturation. To date, most antimicrobial peptide precursors have been described from amphibians, numerous active mature molecules are found in skin secretions [107,108]. Both simple and complex precursors have been annotated, and it seems more likely that processing is carried out at type 2 EtoR/EafterR motifs. The same stands for precursors from sea anemones, fish and gastropods, although current available information for these animals is scarce. Certain differentiation occurs in arachnids with most known precursors from spiders

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carrying type 2 motifs, whereas precursors of scorpion antimicrobial peptides and varisin A1 from acarian hemolymph are processed at conventional type 1 motifs [109-111].

Figure 6. Maturation motifs in animals of different taxonomy. Precursors of (A) antimicrobial peptides and cytotoxins, (B) neurotoxins, (C) enzymes, and (D) regulatory polypeptides were considered. The numbers of precursors with the R(K)toR maturation motifs (green bars), the EtoR/EafterR maturation motifs (blue bars) and ambiguous (cyan bars) are indicated on the Y-axis. Anem. – anemones, Arachn. – arachnids, Amphib. – amphibians, Mamm. – mammals, Gastrop. – gastropods.

In neurotoxin precursors, type 2 maturation is a characteristic feature of polypeptides from spiders, although some protoxins such as magi-4 precursor lack the EtoR/EafterR motifs [112]. Precursors of neurotoxins from scorpions, insects, sea anemones and snakes are all processed at the R(K)toR motif. The most numerous neurotoxins described to date originate from cone snails and according to the performed analysis many precursors follow type 1 maturation. A lot of conotoxin precursors are known to be processed under the action of a different type of enzymes. As an example, Tex31 recognizing a motif of four residues has been isolated and characterized [113]. Precursors of hormones, neuropeptides and other regulatory polypeptides are processed by SPC. Most secreted enzymes also feature type 1 maturation with the exception of those from arachnids. Just as neurotoxin precursors, arachnid proenzymes most often carry the EtoR/EafterR motifs. Two major results were obtained from motif distribution analysis. First, the EtoR/EafterR motifs are characteristic of polypeptide precursors from spiders, and not only of protoxins as suggested earlier [53]. Spiders seem to have deviated from other animals in terms of protein

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processing machinery due to some evolutionary reasons. Second, the type 2 maturation motifs are present in most antimicrobial and cytolytic peptide precursors. Selection for this type of motifs is due to the highly basic nature of the mature products: utilization of the EtoR/EafterR motifs as opposed to the R(K)toR motif allows any combinations of Arg/Lys residues inside the mature chain.

VII. MATURE CHAIN PREDICTION ALGORITHM The set of amino acid sequences analyzed to date is tiny compared to the expected number of polypeptides awaiting discovery and/or investigation and biased towards molecules with certain activity (the more intensively studied and with a higher medical impact) and source organisms of certain taxonomy (those more intensively studied and those with a full genome sequenced). For these reasons novel maturation motifs might be identified in the near future, and many more sequences with the motifs already described will appear. A detailed scheme for analysis of polypeptide precursor primary structure was developed to help identify mature product sequences (Figure 7). This procedure is thought to be effective in terms of mature polypeptide structure prediction from in silico translated nucleotide sequences. The presented scheme summarizes all the described approaches to precursor primary structure analysis. Four consecutive steps of precursor limited proteolysis (both in vivo and in silico) can be distinguished. On the first step, the signal peptide is identified. Next, on the second step, the R(K)toR and EtoR/EafterR motifs are located. If the sequence contains both type 1 and type 2 maturation motifs, a closer inspection of the sequence with identification of cysteine-rich regions (presumably forming a compact fold) and possible split sites is needed. The final decision between SPC or PTC type of fragmentation is then made and the probable processing sites are located (it is worth remembering that complex precursors with type 2 maturation carry symmetrical EtoR/EafterR motifs). As a result, on the second step the mature active and auxiliary “ballast” parts are separated. On the third step, N- and C-terminal trimming of the target mature sequences occurs due to the action of DPP and CP. As a rule, basic residues are removed from the C-terminus and repetitive dipeptides Xaa-Pro and XaaAla are cleaved off from the N-terminus. On the last stage, a C-terminal Gly residue (if present) is converted to an amide group by PAM. In the end of such an analysis according to the proposed scheme, the mature polypeptide sequences designed by nature are retrieved. Nevertheless, experimental “wet” verification of the acquired in silico data is always invaluable.

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Figure 7. Secreted protein and peptide maturation in silico. A detailed chart showing four major steps of secreted polypeptide maturation due to precursor proteolytic cleavage and trimming. Step 1, signal peptide removal. Step 2, adequate convertase processing. Step 3, N- and/or C-terminal trimming. Step 4, C-terminal amidation. For each step, sequence motifs that guide the corresponding processing events are indicated.

ACKNOWLEGMENTS This work was supported in part by the Russian Foundation for Basic Research (grant no. 08-04-00454), and the Program of Cell and Molecular Biology of Russian Academy of Sciences.

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[101] Kawabata, T; Yasuhara, Y; Ochiai, M; Matsuura, S; Ashida, M. Molecular cloning of insect pro-phenol oxidase: a copper-containing protein homologous to arthropod hemocyanin. Proc. Natl. Acad. Sci. U.S.A., 1995, 92, 7774-7778. [102] Cardoso, FC; Pacifico, LG; Carvalho, DC; Victoria, JM; Neves, AL; Chavez-Olortegui, C; Gomez, MV; Kalapothakis, E. Molecular cloning and characterization of Phoneutria nigriventer toxins active on calcium channels. Toxicon, 2003, 41, 755-763. [103] Quistad, GB; Skinner, WS. Isolation and sequencing of insecticidal peptides from the primitive hunting spider, Plectreurys tristis (Simon). J. Biol. Chem., 1994, 269, 1109811101. [104] Heck, SD; Siok, CJ; Krapcho, KJ; Kelbaugh, PR; Thadeio, PF; Welch, MJ; Williams, RD; Ganong, AH; Kelly, ME; Lanzetti, AJ; et al. Functional consequences of posttranslational isomerization of Ser46 in a calcium channel toxin. Science, 1994, 266, 1065-1068. [105] Diao, J; Lin, Y; Tang, J; Liang, S. cDNA sequence analysis of seven peptide toxins from the spider Selenocosmia huwena. Toxicon, 2003, 42, 715-723. [106] Tambourgi, DV; de F Fernandes Pedrosa, M; van den Berg, CW; Goncalves-deAndrade, RM; Ferracini, M; Paixao-Cavalcante, D; Morgan, BP; Rushmere, NK. Molecular cloning, expression, function and immunoreactivities of members of a gene family of sphingomyelinases from Loxosceles venom glands. Mol. Immunol., 2004, 41, 831-840. [107] Nicolas, P; Vanhoye, D; Amiche, M. Molecular strategies in biological evolution of antimicrobial peptides. Peptides, 2003, 24, 1669-1680. [108] Simmaco, M; Mignogna, G; Barra, D. Antimicrobial peptides from amphibian skin: what do they tell us? Biopolymers, 1998, 47, 435-450. [109] Johns, R; Sonenshine, DE; Hynes, WL. Identification of a defensin from the hemolymph of the American dog tick, Dermacentor variabilis. Insect Biochem. Mol. Biol., 2001, 31, 857-865. [110] Zeng, XC; Wang, SX; Zhu, Y; Zhu, SY; Li, WX. Identification and functional characterization of novel scorpion venom peptides with no disulfide bridge from Buthus martensii Karsch. Peptides, 2004, 25, 143-150. [111] Dai, L; Corzo, G; Naoki, H; Andriantsiferana, M; Nakajima, T. Purification, structurefunction analysis, and molecular characterization of novel linear peptides from scorpion Opisthacanthus madagascariensis. Biochem. Biophys. Res. Commun., 2002, 293, 15141522. [112] Corzo, G; Gilles, N; Satake, H; Villegas, E; Dai, L; Nakajima, T; Haupt, J. Distinct primary structures of the major peptide toxins from the venom of the spider Macrothele gigas that bind to sites 3 and 4 in the sodium channel. FEBS Lett., 2003, 547, 43-50. [113] Milne, TJ; Abbenante, G; Tyndall, JD; Halliday, J; Lewis, RJ. Isolation and characterization of a cone snail protease with homology to CRISP proteins of the pathogenesis-related protein superfamily. J Biol Chem., 2003, 278, 31105-31110.

In: Protein Biosynthesis Editors: Toma E. Esterhouse and Lado B. Petrinos

ISBN 978-1-60692-156-2 © 2009 Nova Science Publishers, Inc.

Chapter X

THE EFFECTS OF TEMPERATURE ON ECTOTHERM PROTEIN METABOLISM Nia M. Whiteley1,* and Keiron P. P. Fraser2 1

Bangor University, School of Biological Sciences, Bangor, Gwynedd, LL57 2UW, USA 2 British Antarctic Survey, Natural Environment Research Council, High Cross, Madingley road, Cambridge, CB3 OET, USA

ABSTRACT Protein metabolism in ectotherms is strongly influenced by body temperature and thermal history. In many species, rates of protein synthesis increase with temperature up to a species-specific, thermal optimum. Temperature effects protein synthesis by directly influencing the rates of specific biochemical processes involved in the synthesis of proteins, and also by effecting food consumption. In turn, an increase in food consumption will elevate rates of protein synthesis. Animals have evolved the ability to at least partially compensate rates of protein synthesis as ambient temperatures change, by increasing or decreasing tissue RNA concentrations (RNA to protein) and RNA activity (kRNA). However, at polar temperatures, full compensation of protein synthesis does not appear to occur, and ectotherms are only capable of very low rates of protein synthesis. Temperature also has a direct effect on the proportion of synthesised protein that is degraded, with approximately twice as much protein degraded in polar ectotherms as tropical ectotherms. The result of this is that protein growth at temperatures near the lower limits of life is considerably less efficient than at warmer temperatures. The aim of this review is to examine the effects of temperature on protein metabolism in ectotherms from stable thermal environments and those from more variable thermal regimes. Examples mainly from aquatic environments, will be considered at different levels of biological organisation. In this way, the review will cover both temporal and spatial

*

Mailing Address: N. M. Whiteley. School of Biological Sciences. Bangor University, Bangor, Gwynedd, LL57 2UW, UK. Tel.:+44 1248 388080; Fax: +44 1248 370371. Email: [email protected]

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Nia M. Whiteley and Keiron P. P. Fraser changes in protein metabolism in ectotherms, with particular interest in those organisms living at thermal extremes.

INTRODUCTION Temperature is an important environmental variable that has a marked effect on ectothermic animals (invertebrates and lower vertebrates) at all levels of biological organisation. As biological rate processes are influenced by temperature, thermal fluctuations are a particular problem for ectotherms because their body temperatures generally vary with environmental temperature. All metazoan life is restricted to a relatively narrow band of thermal conditions imposed by the freezing point of seawater and the thermal sensitivity of proteins (-1.86 to 40ºC). Within these physical limits, ectotherms experience a tremendous variety of temperature ranges, dictated by the physical properties of their thermal surroundings and their thermal habitats. Differences in sensitivity to temperature are described as variations in thermal tolerances or thermal windows (Pörtner, 2002). The upper and lower critical limits of which are determined by genotype but are subject to phenotypic plasticity. Consequently, ectotherms are adapted by genetic selection to function optimally in their particular thermal niche (Hochachka & Somero, 2002). This means that ectotherms experiencing relatively stable temperatures in their natural environments (stenotherms) are less likely to tolerate temperature fluctuations, for example polar and tropical aquatic ectotherms. Those that are more tolerant of temperature change tend to be temperate aquatic ectotherms or terrestrial ectotherms that naturally encounter variations in temperature (eurytherms), either on a seasonal or diurnal scale. In order to maintain adequate function and activity during temperature changes many ectotherms can compensate biological functions within limits, and acclimate to a range of temperatures in the laboratory, or acclimatize in the wild under the influence of many different factors. The degree of thermal compensation, however, varies tremendously and can differ between species and also between tissues of the same species, also one biochemical pathway may show compensation while others do not (Hochachka & Somero, 2002). The variations in body temperatures experienced by ectotherms and the variability in thermal experiences and responses, provides thermal biologists, attempting to understand the relationship between temperature and biological rate processes, with an almost infinitely variable and highly complex biological system. Protein metabolism is the continuous synthesis and degradation of an animal’s total protein pool, and as such encompasses two important biological rate processes. Soft tissue growth is the result of protein that is retained when synthesis rates exceed degradation rates. As protein degradation is so difficult to measure (cf. Fraser and Rogers, 2007), the majority of ectothermic studies depend on protein synthesis estimates. The main methodology used is the flooding-dose technique which introduces a single flooding dose of labelled and unlabelled amino acids into the intracellular free-pools (Garlick et al., 1980; Martinez, 1987). Changes in protein synthesis rates are estimated from the specific activities of radio-labelled amino acids in both the protein-bound and intracellular free-pools. Temperature is one of the most important factors controlling protein synthesis rates, but much of the protein synthesis work has been carried out on aquatic ectotherms at temperatures raging from 0 to 27ºC

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(McCarthy & Houlihan, 1997; Fraser & Rogers, 2007; Lewis & Driedzic, 2007). Only limited data has been collected from tropical species, and very little is available from terrestrial ectotherms. Consequently most of the information available comes from polar and temperate species where body temperatures are 10 to 37ºC lower than the body temperatures of mammals. Rates of protein metabolism in ectotherms are temperature dependent at a variety of levels of biological organisation, from single cells to tissues and whole organisms, and from single celled organisms to metazoans. The relationship, however, is complex, and it is important to note that there are distinct differences in examining the effect of a range of temperatures on a physiological process in a single species and in making the same measurements across a range of species adapted to differing temperatures (Fraser & Rogers, 2007). For example, the slopes describing metabolic rates of a single species exposed to a range of temperatures are likely to be significantly steeper than a slope fitted to the metabolic rates of a range of species adapted to differing temperatures (Clark & Fraser, 2004). A similar response occurs for whole-organism protein synthesis rates. In the ectotherms studied to date, there is some compensation for low temperatures in the medium to long-term but the relationship varies within and between species, and tissue responses do not always follow that of the whole-organism. Thermal histories (natural thermal environment) and thermal experiences (acclimation versus acclimatization) also make a difference, as does the timecourse of the temperature change (Loughna & Goldspink, 1985). The increase in food consumption with temperature is an added complication because the associated increase in amino acid uptake increases protein synthesis rates at constant temperature (McCarthy et al., 1993; 1994). Consequently it is difficult to determine the effects of temperature in isolation, especially during seasonal acclimatization where temperature doesn’t act alone but interacts with a large number of abiotic and biotic factors. Individual variation can also occur due to variations in size, stage of development and age. This chapter aims to summarise all of this information, to present our current understanding of the effects of temperature on protein metabolism in ectotherms at different levels of biological organisation. We will include examples from ectotherms exposed to a variety of thermal experiences, both in the laboratory and in the field, and the emphasis throughout will be on overall rates of synthesis rather than the synthesis of specific proteins. As ectothermic animals make up the majority of life on earth in terms of biomass and number of species, and as ectotherms are important sources of food for many higher vertebrates, it is important that we understand the fundamental process affecting the ability of organisms to cope with temperature fluctuations. Changes in protein metabolism influence growth rates and ultimately both individual and population performance. Understanding the effects of temperature on protein metabolism is of great interest, in particular with regard to understanding how ectotherms will continue to survive further changes in global temperatures.

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FUNDAMENTAL EFFECTS OF TEMPERATURE ON PROTEIN METABOLISM Rates of protein synthesis in ectotherms generally increase with temperature up to a species-specific thermal maxima, after which they will rapidly decrease (Haschemeyer et al., 1979; Haschemeyer & Mathews, 1983; McCarthy et al., 1999). The rate at which an animal synthesises protein can be regulated in a variety of ways, including; transcriptional control and the concentration and activity of the RNA (Fraser et al., 2007; Hofmann et al., 2005; Mathers et al., 1993; Storch et al., 2005; McCarthy & Houlihan, 1997). As we are focussing on the effect of temperature on overall protein synthesis rates, rather than specific proteins in this chapter, our discussion will concentrate on the regulation of global protein synthesis rates via RNA concentration and activity. RNA concentrations are sometimes expressed relative to fresh weight of tissue (μg RNA.mg-1 fresh weight) or more commonly as RNA to proteins ratios (μg RNA.mg-1 protein), while RNA activity (kRNA) is expressed as mg protein synthesised per mg RNA per day. Measurements of the total RNA concentrations in an organism will include not only ribosomal RNA, but also messenger (mRNA) and transfer RNA (tRNA). With regard to the overall control of protein synthesis rates we are really interested in the concentration and activity of ribosomes, however, mRNA and tRNA only account for ~ 13% of the total RNA measured, therefore total RNA concentrations provide a reasonable proxy estimate of ribosomal concentrations (Sugden & Fuller, 1991). If RNA to protein ratios are compared in ectotherms living at a range of water temperatures the ratio decreases with increasing water temperature (Figure 1; Fraser et al., 2002; Fraser & Rogers, 2007). Likewise if either whole animal, tissue or cellular RNA to protein ratios are compared in individuals of the same species living at differing water temperatures the ratios will tend to be elevated at lower temperatures (Foster et al., 1992; McCarthy et al., 1999; Wagner et al., 2001). It should be noted, however, that food consumption rates in ectotherms, which will also effect RNA concentrations, are also elevated at higher temperatures (McMillan & Houlihan, 1988; Houlihan et al., 1990). It is therefore impossible to fully separate the effects of food consumption and temperature per se on RNA concentrations in ectotherms. Body mass also has a significant effect on RNA to protein ratios, therefore to allow valid comparisons of RNA to protein ratios between species it is important that data are mass standardised as in Figure 1 (Fraser & Rogers, 2007). In bacteria the relationship between ambient temperature and RNA to protein ratios appears less clear. In E. coli there was no significant change in RNA to protein ratios between 25 and 37oC (Farewell & Neidhardt, 1998). RNA activity is temperature dependent and increases with temperature (McCarthy & Houlihan, 1997; Fraser et al., 2002; Fraser & Rogers, 2007). To compensate for the decrease in RNA activity at low temperatures, higher tissue RNA concentrations are maintained. This mechanism appears to be widespread across ectotherms studied to date, although in the Antarctic zoarcid, Pachycara brachycephalum thermal compensation of protein synthesis maybe achieved primarily via an increase in the RNA activity rather than elevated levels of RNA (Storch et al., 2005). However, it appears that full compensation of protein synthesis

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Table 1. Sources of RNA to protein data used to plot Figure 1 Data point label 1 2 3 4 5 6 7 8 9 10 11 12 13

Species

Group

Reference

Limanda limanda Clupea harengus Oncorhynchus mykiss Ctenopharyngodon idella Gadus morhua Dicentrarchus labrax Anarhichas lupus Nacella concinna Saduria entomon Glyptonotus antarcticus Idotea resecata Homarus gammarus Rattus norvegicus

Teleost Teleost Teleost Teleost Teleost Teleost Teleost Gastropod Isopod Isopod Isopod Decapod Mammal

Houlihan et al., 1994 Mathers et al., 1994 McCarthy et al., 1994 Carter et al., 1993 Lyndon et al., 1992 Langar and Guillaume, 1994 McCarthy et al., 1999 Fraser et al., 2002 Robertson et al., 2001a Robertson et al. 2001b Whiteley et al., 1996 Mente et al., 2001 Goldspink and Kelly, 1984

RNA to protein ratio (μg.mg-1)

16 14

8

12

1 5

10

10

7

8

11

6

3 9

4

12 13

2

6

4

2

0

3.2

3.3

3.4

3.5

3.6

3.7

Temperature (1000/K) Figure 1. Mass standardised whole-body RNA to protein ratios plotted against the reciprocal of temperature. All values are standardised to a mass of 160g, the mean body mass of the animals used in the analysis, using a scaling coefficient of -0.26. The scaling coefficient was calculated by fitting a least-squares regression model to the natural log-transformed body mass and RNA to protein ratio data for all species. The plotted regression line was fitted using least-squares regression analysis ( y = -81.5 + 25.0x, r2 = 49%, p