The Septins

The Septins The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd...

Author: Peter A. Hall | S. E. Hilary Russell | John R. Pringle

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The Septins

The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

The Septins Edited by

Peter A. Hall Queen’s University Belfast, Belfast, Northern Ireland, UK

S.E. Hilary Russell Queen’s University Belfast, Belfast, Northern Ireland, UK

and

John R. Pringle Stanford University Medical Centre, Stanford, CA, USA

A John Wiley & Sons, Ltd., Publication

This edition first published 2008  2008 by John Wiley & Sons, Ltd. Wiley-Blackwell is an imprint of John Wiley & Sons, formed by the merger of Wiley’s global Scientific, Technical and Medical business with Blackwell Publishing. Registered office: John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Other Editorial Offices: 9600 Garsington Road, Oxford, OX4 2DQ, UK 111 River Street, Hoboken, NJ 07030-5774, USA For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/wiley-blackwell The right of the author to be identified as the author of this work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging-in-Publication Data The septins / edited by Peter A. Hall, S.E. Hilary Russell, and John R. Pringle. p. cm. Includes index. ISBN 978-0-470-51969-1 1. Septins. I. Hall, Peter A. II. Russell, S.E. Hilary. III. Pringle, John R., 1943QP552.S37S47 2009 572 .6 – dc22 2008022154 ISBN: 978-0-470-51969-1 A catalogue record for this book is available from the British Library Typeset in 10.5/12.5pt Times by Laserwords Private Limited, Chennai, India Printed and bound in Great Britain by Antony Rowe Ltd, Chippenham, Wiltshire First impression

2008

Cover images (from left to right): • Localization of the septin Sep1 (green) to the cleavage furrows in dividing cells of a post-gastrulation Drosophila embryo. DNA (stained with propidium iodide) is shown in red. Image kindly provided by Johnny Fares and John Pringle and used with their permission. • Septin (green) in Candida albicans pseudohyphae (wildtype) • Septin (green) in Candida albicans pseudohyphae (rga2/bem3/mutant lacking Cdc42 GAPs). C. albicans images kindly provided by Helen Court and Peter Sudbery and used with their permission.

Contents Authors and Affiliations

vii

An introduction to the septins Peter A. Hall, S. E. Hilary Russell and John R. Pringle

1

Section I

Setting the scene

5

Chapter 1

Origins and development of the septin field John R. Pringle

7

Chapter 2

Evolution and conserved domains of the septins Michelle Momany, Fangfang Pan and Russell L. Malmberg

Section II Septins in model systems Chapter 3

Biochemical properties and supramolecular architecture of septin hetero-oligomers and septin filaments Michael A. McMurray and Jeremy Thorner

Chapter 4 Yeast septins: a cortical organizer Yves Barral Chapter 5

Chapter 6

Septins in four model fungal systems: diversity in form and function Amy S. Gladfelter and Peter Sudbery Septins in the metazoan model systems Drosophila melanogaster and Caenorhabditis elegans Christine M. Field, Amy Shaub Maddox, John R. Pringle and Karen Oegema

Section III Septins in mammals Chapter 7

The genomics and regulation of the human septin genes S.E. Hilary Russell

35

47

49 101

125

147

169 171

CONTENTS

vi

Chapter 8

The functions of septins in mammals Carol D. Froese and William S. Trimble

187

Chapter 9

Septin-interacting proteins in mammals Brandon E. Kremer and Ian G. Macara

211

Chapter 10 Septin functions in the mammalian cytoskeleton Elias T. Spiliotis and W. James Nelson

229

Chapter 11 Septins and the synapse Jing Xue, Victor Anggono and Phillip J. Robinson

247

Chapter 12 Septins and platelets Jerry Ware, Constantino Mart´ınez and Barbara Zieger

269

Chapter 13 Septins and apoptosis Marie-Jeanne Carp and Sarit Larisch

281

Chapter 14 Septins and human disease Peter A. Hall and Fern P. Finger

295

Chapter 15 Insight into septin functions from mouse models Makoto Kinoshita

319

Section IV Envoi

337

Chapter 16 Septins: 2008 and beyond Peter A. Hall, S.E. Hilary Russell and John R. Pringle

339

Appendix A

Septin and septin-like sequences 343 Michelle Momany, Fangfang Pan and Russell L. Malmberg

Appendix B

Mammalian septin nomenclature Peter A. Hall, Elspeth Bruford, Hilary Russell, Ian G. Macara and John R. Pringle

351

Appendix C

Septin meetings and workshops Peter A. Hall and John R. Pringle

355

Index

361

Authors and Affiliations Victor Anggono

Cell Signalling Unit, Children’s Medical Research Institute, Locked Bag 23, Wentworthville, NSW 2145, Australia

Yves Barral

Institute of Biochemistry, ETH Zurich, Schafmattstrasse 18, 8093 Zurich, Switzerland

Elspeth Bruford

Human Genome Nomenclature Committee, European Bioinformatics Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge CB10 1SD, UK

Marie-Jeanne Carp

Apoptosis and Cancer Research Laboratory. Department of Pathology, Rambam Medical Center, Haifa 31096, Israel

Fern P. Finger

Department of Biology and Center for Biotechnology and Interdisciplinary Studies, Rensselaer Polytechnic Institute, 110 Eighth Street, Troy, NY 12180-3590, USA

Carol D. Froese

Program in Cell Biology, Hospital for Sick Children, 555 University, Avenue, Toronto, Ontario M5G 1X8, Canada

Christine Field

Department of Systems Biology, Harvard Medical School, 200 Longwood Avenue, WA 536, Boston, MA 02115, USA

Amy Gladfelter

Department of Biology, Dartmouth College, 409 Gilman Hall, Hanover, NH 03755, USA

Peter A. Hall

Institute of Pathology, School of Medicine, Queen’s University Belfast, Belfast BT12 6BL, Northern Ireland, UK

viii

AUTHORS AND AFFILIATIONS

Makoto Kinoshita

Cell Biology and Biochemistry Unit, Kyoto University Graduate School of Medicine and CREST, Japan Science & Technology Agency, Yoshida Kanoe, Sakyo, Kyoto 606-8501, Japan

Brandon E. Kremer

Department of Microbiology, University of Virginia School of Medicine, Charlottesville, VA 22908-0577, USA

Sarit Larisch

Apoptosis and Cancer Research Laboratory, Department of Pathology, Rambam Medical Center, Haifa 31096, Israel

Ian G. Macara

Department of Microbiology, University of Virginia School of Medicine, Charlottesville, VA 22908-0577, USA

Amy Shaub Maddox

Institute for Research in Immunology and Cancer, Department of Pathology and Cell Biology, University of Montréal, Montréal H3C 3J7, Canada

Russell L. Malmberg

Plant Biology Department, University of Georgia, Athens, GA 30602, USA

Constantino Mart´ınez

Centro Regional de Hemodonación, University of Murcia, Calle de Ronda de Garay S/N, Murcia 30003, Spain

Michael A. McMurray

Department of Molecular and Cell Biology, Division of Biochemistry and Molecular Biology, University of California, Berkeley, CA 94720, USA

Michelle Momany


W. James Nelson

Department of Biological Sciences, The James H. Clark Center, The Bio-X Program, 318 Campus Drive, E-200, Stanford University, Stanford, CA 94305-5430, USA

Karen Oegema

Ludwig Institute for Cancer Research, Department of Cellular & Molecular Medicine, University of California San Diego, CMM East, Room 3053, 9500 Gilman Drive, La Jolla, CA 92093-0653, USA

AUTHORS AND AFFILIATIONS

ix

Fangfang Pan


John R. Pringle

Department of Genetics, MC 5120, 300 Pasteur Drive, M-322 Alway Building, Stanford University Medical Center, Stanford, CA 94305-5120, USA

Phillip J. Robinson

Cell Signalling Unit, Children’s Medical Research Institute, Locked Bag 23, Wentworthville, NSW 2145, Australia

S.E. Hilary Russell

Ovarian Cancer Research Laboratory, Centre for Cancer Research and Cell Biology, Queen’s University Belfast, A floor, Belfast City Hospital, Lisburn Road, Belfast BT9 7AB, Northern Ireland, UK

Elias T. Spiliotis

Department of Biological Sciences, The James H. Clark Center, The Bio-X Program, 318 Campus Drive, E-200, Stanford University, Stanford, CA 94305-5430, USA

Peter Sudbery

Department of Molecular Biology and Biotechnology, Sheffield University, Western Bank, Sheffield S10 2TN, UK

Jeremy Thorner

Department of Molecular and Cell Biology, Division of Biochemistry and Molecular Biology, University of California, Berkeley, CA 94720, USA

William S. Trimble

Program in Cell Biology, Hospital for Sick Children, 555 University Avenue, Toronto, Ontario M5G 1X8, Canada

Jerry Ware

Department of Physiology and Biophysics, University of Arkansas for Medical Sciences, 4301 West Markham Street, Little Rock, AR 72205, USA

Jing Xue

Vascular Biology Centre, Medical College of Georgia, 1459 Laney Walker Boulevard, Room CB 3330, Augusta, GA 30912, USA

Barbara Zieger

Department of Paediatrics and Adolescent Medicine, University Hospital Freiburg, Mathildenstrasse 1, D-79106 Freiburg, Germany

(a)

(b)

(d)

(c)

(e)

Plate 5.1 Septin organisation in C. albicans. (a) A septin ring forms at the bud neck in yeast cells. (b) Septin rings form at the bud necks of pseudohyphal cells, and nuclei divide across the mother-bud necks. (c) Basal septin bars and septin cap in a newly evaginated hyphal germ tube. (d) Septin collars form along the length of hyphal germ tubes. Nuclei migrate out of mother cells and divide across the septin collars. Inset, enlarged view of the septin collar indicated by the arrow. (e) After mitosis, the septin collar separates into two sharply defined rings between which the chitinous primary septum forms. Septin is Cdc10-YFP in all panels. Panels a, c and e are counterstained with Calcofluor white (blue), which stains cell walls and the primary septum in panel c. Panels b and d are counterstained with Concanavaline A – Texas red and the nuclei are stained with 4 ,6-diamidino-2-phenylindole (DAPI) (blue). Scale bars, 5 µm in panels a, b, and d; 1 µm in panels c and e, and in inset d. Images were generated with a DeltaVision RT wide-field epifluorescence microscope and deconvolved with Softworx company software.


S51 W260 G241 G261 H270

c.c. N55

c.c. S31K30

(Y17)

c.c.

(a) c.c.

c.c.

c.c.

c.c.

c.c.

SEPT7-GDP SEPT2-GDP SEPT2-GDP SEPT7-GDP SEPT6-GTP SEPT6-GTP

T228

D185

c.c. c.c.

(b)

Cdc11-GDP Cdc12-GDP Cdc3-GTP Cdc10-GDPCdc10-GDP Cdc3-GTP Cdc12-GDP Cdc11-GDP

c.c.

F156

c.c.

() α0 ) | β1 > () α1 ) Cdc3 70 MGITSSQSEKGQVLPDQPEIKFIRRQINGYVGFANLPKQW-HRRSIKNGFSFNLLCVGPDGIGKTTLMKTLFNNDDIEANLVKDYEEELANDQEEEEGQG 168 Cdc10 1 MDPLSSVQPAS------YVGFDTITNQI-EHRLLKKGFQFNIMVVGQSGLGKSTLINTLFASH--------LIDSATGDD-------- 65 Cdc11 1 MSGIIDASSALRKRKHLKRGITFTVMIVGQSGSGRSTFINTLCGQQ--------VVDTSTTILLP------ 57 Cdc12 1 MSAATATAAPVP-----PPVGISNLPNQR-YKIVNEEGGTFTVMLCGESGLGKTTFINTLFQTV---------LKRADGQQ-------- 66 hsSEPT2 1 MSKQQPTQFIN-PETPGYVGFANLPNQV-HRKSVKKGFEFTLMVVGESGLGKSTLINSLFLTD--------LYPERVIP--------- 69 hsSEPT6 1 MAATDIARQVGEGCRTVPLA-----GHVGFDSLPDQL-VNKSVSQGFCFNILCVGETGLGKSTLMDTLFNTK----------FEGEP---------- 71 hsSEPT7 1 MSVSARSAAAEERSVNSSTMVAQQKNLEGYVGFANLPNQV-YRKSVKRGFEFTLMVVGESGLGKSTLINSLFLTD--------LYSPEYP---------- 81 ident/simil * * *... * * *..* . .* modification P P P

Cdc3 169 Cdc10 66 Cdc11 58 Cdc12 67 hsSEPT2 70 hsSEPT6 72 hsSEPT7 82 ident/simil modification

| β2 > | β3 > () α2 ) | β4 > EGHENQSQEQRHKVKIKSYESVIEEN-GVKLNLNVIDTEGFGDFLNNDQKSWDPIIKEIDSRFDQYLDAENKINR--HSINDKRIHACLYFIEPTGHYLK ----ISALPVTKTTEMKISTHTLVED-RVRLNINVIDTPGFGDFIDN-SKAWEPIVKYIKEQHSQYLRKELTAQRE-RFITDTRVHAILYFLQPNGKELS -----TDTSTEIDLQLREETVELEDDEGVKIQLNIIDTPGFGDSLDN-SPSFEIISDYIRHQYDEILLEESRVRRN-PRFKDGRVHCCLYLINPTGHGLK ----HRQEPIRKTVEIDITRALLEEK-HFELRVNVIDTPGFGDNVNN-NKAWQPLVDFIDDQHDSYMRQEQQPYR--TKKFDLRVHAVLYFIRPTGHGLK ----GAAEKIERTVQIEASTVEIEER-GVKLRLTVVDTPGYGDAINC-RDCFKTIISYIDEQFERYLHDESGLNR--RHIIDNRVHCCFYFISPFGHGLK ------ATHTQPGVQLQSNTYDLQES-NVRLKLTIVSTVGFGDQINK-EDSYKPIVEFIDAQFEAYLQEELKIRRVLHTYHDSRIHVCLYFIAPTGHSLK ----GPSHRIKKTVQVEQSKVLIKEG-GVQLLLTIVDTPGFGDAVDN-SNCWQPVIDYIDSKFEDYLNAESRVNR--RQMPDNRVQCCLYFIAPSGHGLK . . . . .... * *.** . . . * . . * * * *.. * . * *. * P

265 158 150 158 161 163 173


() α3 ) |β5> () α4 ) () α5' ) |β6> PLDLKFMQSVYEKCNLIPVIAKSDILTDEEILSFKKTIMNQLIQSNIELFKPPIYSNDDAEN----------SHLSERLFSSLPYAVIGSNDIVENYS-G RLDVEALKRLTEIANVIPVIGKSDTLTLDERTEFRELIQNEFEKYNFKIYPYDSEELTDEE-----------LELNRSVRSIIPFAVVGSENEIEIN--G EIDVEFIRQLGSLVNIIPVISKSDSLTRDELKLNKKLIMEDIDRWNLPIYNFPFDEDEISDED---------YETNMYLRTLLPFAIIGSNEVYEMGGDV PIDIETMKRLSTRANLIPVIAKADTLTAQELQQFKSRIRQVIEAQEIRIFTPPLDADSKEDAKSGSNPDSAAVEHARQLIEAMPFAIVGSEKKFDNGQ-G PLDVAFMKAIHNKVNIVPVIAKADTLTLKERERLKKRILDEIEEHNIKIYHLPDAESDEDEDF---------KEQTRLLKASIPFSVVGSNQLIEAK--G SLDLVTMKKLDSKVNIIPIIAKADAISKSELTKFKIKITSELVSNGVQIYQFPTDD----ESV---------AEINGTMNAHLPFAVIGSTEELKIG--N PLDIEFMKRLHEKVNIIPLIAKADTLTPEECQQFKKQIMKEIQEHKIKIYEFPETD-DEEE-----------NKLVKKIKDRLPLAVVGSNTIIEVN--G .*. .. . *..*.* *.* .. * . * .. . . .* ...** P P P P

354 245 241 257 250 248 259


| β7> | β8> () α5 ) () α6 ) () α7? ... NQVRGRSYPWGVIEVDNDNHSDFNLLKNLLIKQFMEELKERTSKILYENYRSSKLAKLGIK-QDNSVFKEFDP---ISKQLEEKTLHEAKLAKLE 445 ETFRGRKTRWSAINVEDINQCDFVYLREFLIRTHLQDLIETTSYIHYEGFRARQLIALKENANSRSSAHMSSNAIQR.----------------- 298 GTIRGRKYPWGILDVEDSSISDFVILRNALLISHLHDLKNYTHEILYERYRTEALSGESVAAESIRPNLTKLNGSSSSSTTTRRNTNPFK(8)VL 341 TQVVARKYPWGLVEIENDSHCDFRKLRALLLRTYLLDLISTTQEMHYETYRRLRLEGHENTGEGNE--DFTLPAIAPAR----KLSHNPRYKEEE 346 KKVRGRLYPWGVVEVENPEHNDFLKLRTMLI-THMQDLQEVTQDLHYENFRSERLKRGGRKVENEDMNKD------------------------- 319 KMMRARQYPWGTVQVENEAHCDFVKLREMLIRVNMEDLREQTHTRHYELYRRCKLEEMGFKDTDPDSKPFSLQETYEAK----RNEFLGELQKKE 339 KRVRGRQYPWGVAEVENGEHCDFTILRNMLIRTHMQDLKDVTNNVHYENYRSRKLAAVTYNGVDNNKNKGQLTKSPLAQMEEERREHVAKMKKME 354 * * .. ** *. *. . .* * ** . * P

(c) Plate 3.3 Structural models and structural alignment of the human SEPT2-SEPT6-SEPT7 hexamer and the budding yeast Cdc3-Cdc10-Cdc11-Cdc12 octamer. (a) A ribbon representation of a space-filling depiction of the structure of the human SEPT7-SEPT6-SEPT2-SEPT2-SEPT6SEPT7 heterohexamer (adapted from Figure 4 in Sirajuddin et al., 2007), based on coordinates (PDB accession no. 2QAG) for the asymmetric unit (a SEPT2-SEPT6-SEPT7 trimer) and on the relative order of subunits as determined by EM analysis of a MBP-SEPT2-SEPT6-SEPT7 complex (Sirajuddin et al., 2007). The positions and directions in which the CTEs are predicted to extend from the globular G domains are indicated by the arrows and labeled ‘c.c.’ (for ‘coiled coil’). SEPT2, purple; SEPT6, blue; SEPT7, green; bound guanine nucleotide, red; positions of selected residues from Table 3.1, gold and labeled; disordered residues not represented in the final model, absent or shown in their most likely positions by dashed lines. (b) A ribbon representation of the budding yeast Cdc11-Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12Cdc11 hetero-octamer, based on the relative order of subunits determined by Bertin et al. (2008) [see also Figure 3.1c and modelled on (a)]. Cdc10, purple; Cdc3, blue; Cdc12, green; Cdc11, magenta; bound guanine nucleotide, red. (c) Alignment using the Clustal W algorithm of the G domains and surrounding sequences of the septins shown in (a) and (b), with the elements of secondary structure labeled as for human SEPT2 (Sirajuddin et al., 2007), plus the predicted ‘α7’ helix (based on Versele et al., 2004). Highly conserved positions are indicated below the sequences: ‘∗’, residues identical in all; ‘.’, highly similar residues in all. Residues known to be phosphorylated are in bold and indicated by a ‘P’. ‘(8)’ represents a stretch of eight residues present in Cdc11, but not in the other septins shown.

(a)

(b)

(c)

(d)

(e)

(f)

3.0 µm (g)

(h)

Plate 5.2 Septin organization in A. gossypii. Fluorescence micrographs of cells expressing Sep7-GFP are shown. (a–d) Initial appearance and deposition of septin rings at growing tips. (e– h) Assembly of septin rings at emerging lateral branches. Images prepared by Bradley DeMay.

Sept 3, 5 and 7 Localization in Neurons Septins

Synaptophysin

Merge

Sept3

Sept5

Sept7

Plate 11.3 Localization of septins in primary cultured hippocampal neurons. Hippocampal neurons cultured for 14 days were stained with antibodies against SEPT3 (top), SEPT5 (middle), and SEPT7 (bottom); and they were all co-stained with antibodies against synaptophysin. They show punctate distribution and partial co-localization with the presynaptic nerve terminal marker, synaptophysin. [Figure in the top panel was reproduced with permission from Xue, J., Tsang, C.W., Gai, W.P. et al. (2004b) Septin 3 (G-septin) is a developmentally regulated phosphoprotein enriched in presynaptic nerve terminals. Journal of Neurochemistry, 91, 579–90; figures in the middle and bottom panels were reproduced with permission from Fujishima, K., Kiyonari, H., Kurisu, J. et al. (2007) Targeted disruption of SEPT3, a heteromeric assembly partner of SEPT5 and SEPT7 in axons, has no effect on developing CNS neurons. Journal of Neurochemistry, 102, 77–92].

Dapi

SEPT4_i1

SEPT4_i2

Lymphocytes from healthy donor

Lymphocytes from ALL patient

(c) Plate 13.1 SEPT4 i2 is the only human Septin4 isoform shown to directly induce and promote apoptosis. SEPT4 i2 functions as a tumour suppressor protein. Immunofluorescence staining presents lymphocytes isolated from healthy donor (upper panel) and from acute lymphoblastic leukemia (ALL) patient (lower panel). Staining with Dapi, showing nuclei of cells (blue), the non-apoptotic isoform of SEPT4; SEPT4 i1 (red), and pro-apoptotic protein SEPT4 i2 (green). Whereas all lymphocytes in healthy control contain similar levels of SEPT4 i2 and SEPT4 i1 (upper panel), the sample from leukemia patient exhibits loss of SEPT4 i2 staining in all tumour lymphoblasts (only remaining normal cells are stained), indicating selective loss of the pro-apoptotic protein SEPT4 i2 in leukemia patients (Adopted from Elhasid et al., 2004).

Plate 15.1 Embryonic lethality of Sept9−/− mice. Hematoxylin-eosin (HE) stain indicated that the disorganized histogenesis and/or degeneration involves the neural tube, limb buds, and mesenchyme, where pyknotic nuclei predominate. Unpublished data provided by courtesy of Dr Ernst-Martin Füchtbauer.

(a) Plate 15.2 Sept4 function in dopaminergic nerve terminals in the striatum. A scanning laser confocal microscopic image of a mouse striatal section, immunostained for DNA (blue), the dopamine transporter (green), and tyrosine hydroxylase, a rate-limiting enzyme for dopamine synthesis (red). Note fine network of highly branched axon terminals that synthesize, release, and reuptake dopamine. The nucleated cell bodies belong to the post-synaptic GABAergic neurons.

(a) Plate 15.3 Spermiogenesis defect observed in Sept4 knockout mice. Immunofluorescence signals superimposed on a DIC image of wildtype spermatozoa. Nuclear DNA (blue), Sept4 (green), and mitochondria (red), respectively, are labelled by DAPI, an anti-Sept4 antibody, and MitoTracker Red (Molecular Probes). Note that a septin-based ring (the annulus) is localized to the caudal end of the middle piece, a segment of flagellum covered with a mitochondrial sheath.

An introduction to the septins Peter A. Hall, S. E. Hilary Russell and John R. Pringle

Few modern scientists like specialist monographs. Typically, they are out of date before they are published, and, in fast-moving fields, they show their age quickly! It was therefore with trepidation that we took on the task of planning, commissioning chapters for, and editing a monograph on septins. Why were we persuaded that this was a worthwhile project? The field is still young, yet it has grown quickly in the past few years (see Figure 0.1). Moreover, although it was once largely the preserve of a small band of yeast biologists, this field now encompasses the whole of animal and fungal biology. (Septins do not appear to be present in plants or prokaryotes.) A further stimulus has been the recognition that the septins are involved in a variety of disease processes. It was thus with the anticipation that the septin field would continue to grow rapidly, and that many newcomers would benefit from a comprehensive overview of the state of the field ca. early 2008 (probably the last time that such a thing would be possible), that we embarked on this task. The germ of the idea emerged after the First Septin Workshop in May 2005 (see Appendix C). In the summer of 2006, Andrea Baier of Wileys (now Wiley Blackwell Publishers) approached one of us (Peter Hall) about the idea. A conference call between the three editors followed, and the germ began to grow. Conversations took place with others, including potential authors, and a general view of cautious enthusiasm developed. The original intention was that contributions would be finalized immediately following the Second Septin Workshop (May 2007), but (as in many publishing ventures) reality was different. In this case, a delay was produced not only by the difficulties of some authors in finishing their chapters according to the original schedule, but also by the desirability of including the important structural analyses that were just beginning to emerge in the Summer and Fall of 2007 – so much is thrown into a new light by these important developments! Despite the delay, the contributors have produced a set of chapters that well defines the field as it stands at the end of 2007, not far from our original intention. The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

AN INTRODUCTION TO THE SEPTINS

2

Septin publications 1994 to 2007 500 Cumulative publication 450 400 350 300 250 200 150 Annual publications

100 50 0

1994 1995 1997 1996

1998 1999 2000 2001

2002 2003 2004 2005

2006 2007

Figure 0.1 The increase in publications on septins from October 1994, when the word was first used in the published literature (Sanders and Field, 1994). PubMed was searched using ‘septin*’ as the search string. [Note that the word septin had been employed previously to denote a factor in plasma that opsonizes lipopolysaccharide-bearing particles.]

The septin field now has a solid basis in data from a wide range of systems, providing a foundation on which its future can be built. Furthermore, there are links to a surprising array of physiological processes, not just the role in cytokinesis that for a long time dominated the field. As the contributions here show, we now have a good grip on both the biology and the biochemistry of this fascinating family of proteins. The scene is set by historical perspectives in both recent and geological time. The book begins with a detailed history of the beginnings of the field (including much previously unpublished material) by John Pringle, in whose laboratory most of the early work on the septins took place. The second chapter, by Michelle Momany, Fangfang Pan and Russell L. Malmberg, takes advantage of the many septin sequences now available, from diverse organisms, to describe the evolution of the septin family and the intriguing gain and loss of septin genes during the evolution of different phylogenetic groups. One key point is the evolution of a great diversity of septin genes, which in vertebrates can number as many as 14. A major challenge for the field will be the dissection of the roles of these multiple genes and the evolutionary pressures for their duplication and reduplication. The second section of the book focuses on septins in model systems. Michael McMurray and Jeremy Thorner provide a detailed account of the biochemical properties and supramolecular architecture of septin oligomers and filaments; the


3

work in budding and fission yeast is placed in the context of the important recent work on the structures of the mammalian and C. elegans septin complexes. This contribution dovetails with that of Yves Barral on the function and cell biology of septins in Saccharomyces cerevisiae, where these proteins were first discovered and have since been studied in considerable detail. Amy Gladfelter and Peter Sudbery further develop this theme with a consideration of the septins in other fungi, including Schizosaccharomyces pombe, Candida albicans, Ashbya gossypii and Aspergillus nidulans. Christine Field, Amy Maddox, Karen Oegema and John Pringle complete the section with a discussion of the septins in Drosophila melanogaster and Caenorhabditis elegans. The worm is of particular interest given that it only has two septin genes, unlike the fly with five, yeasts with seven and humans with 14! The third section of the book turns to the mammalian systems. Growth in this area has been particularly dramatic since the recognition by Kinoshita in 1997 that septins are not only essential for cytokinesis, but also seemingly involved in other processes, in mammalian cells. The septin genes in mammals (and particularly in humans) present particular challenges as outlined by Hilary Russell. Not only are there at least 14 distinct human septin genes, but many of these genes show remarkably complex alternative splicing events and regulation by a plethora of strategies. Carol Froese and Bill Trimble provide an overview of the possible functions of mammalian septins, which is complemented by a detailed consideration of the known septin-interacting proteins by Brandon Kremer and Ian Macara. There follows a series of chapters that discuss some specific areas of mammalian septin research including septins and the cytoskeleton (Elias Spiliotis and James Nelson) and the burgeoning area of septins and the nervous system (Jing Xue, Victor Anggono and Philip J. Robinson). Septins have been implicated in the physiology and pathophysiology of platelets, remarkable anucleate ‘cells’ derived from megakaryoctes that have a key role in hemostasis, and Jerry Ware, Constantino Mart´ınez and Barbara Zieger review this area. Marie-Jeanne Carp and Sarit Larisch then cover the controversial area of the role of septins in apoptosis. This section culminates with two chapters that look at septins in disease (Peter Hall and Fern Finger) and at the many insights that we have gained from murine models of septin biology, and, in particular, from knockout mutants (Makoto Kinoshita). The monograph then ends with a speculative overview from the editors of the key questions in the field and of where it may be headed. In addition, several Appendices summarize some important information in a form that may be convenient for those in, or just entering, the field. This monograph could not have been produced without the help and enthusiastic contributions of those in the field. The editors wish to express their sincere thanks to all the authors for their hard work and to the many others in the field whose contributions and comments (particularly at the two Septin Workshops) have helped to push the field, and thus this monograph, forward. We also wish to

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thank Andrea Baier and Fiona Woods from Wiley Blackwell, who have cajoled, persuaded and supported the Editors in this journey! Finally, we wish to note that all royalties from this book have been waived by the Editors and authors and will be used to help support future Septin Workshops.

REFERENCE Sanders, S.L. and Field, C.M. (1994) Cell division. Septins in common? Current Biology, 4, 907–10.

Section I Setting the scene


1 Origins and development of the septin field John R. Pringle Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305, USA

INTRODUCTION In this chapter, I have told the story of the septin field from its conception in Lee Hartwell’s and Breck Byers’s laboratories (1967–1976), through its long gestation, awkward birth, and short childhood, mostly in my own laboratory (1977–1993), and finally through its adolescence in a growing number of laboratories until its coming of age was marked (in 2001) by a session’s being devoted to it at a major meeting (see Appendix C). Although there are many parts of the story that I find embarrassing and guilt-provoking (even many years after the events), I have told the unvarnished truth as best I can reconstruct it. I think that there is value in having an accurate historical record of a scientific field; if nothing else, this account should clarify aspects of the septins’ early history that would otherwise be quite mysterious to a newcomer. I think that this story also illustrates a great and reassuring truth about science: it moves forward even though the people involved, and their behaviour, are often imperfect.

CONCEPTION, GESTATION, AND BIRTH In 1966, Lee Hartwell, then a beginning Assistant Professor at the University of California, Irvine, began isolating a large collection of temperature-sensitive-lethal (ts) mutants of the budding yeast Saccharomyces cerevisiae. His inspiration was the work done on ts mutants of bacteriophage T4 in Bob Edgar’s lab at Caltech, where Lee had done undergraduate research. A publication summarizing The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

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CH01 ORIGINS AND DEVELOPMENT OF THE SEPTIN FIELD

the properties of 400 such mutants appeared soon (Hartwell, 1967); it included brief descriptions of two of the mutants that would later define the septins, namely ts310 and ts471 (later cdc10 and cdc12 ), described at this time as defective ‘in cell-wall formation’. In 1967, another new faculty member at Irvine, Cal McLaughlin, teamed up with Lee, and the two spent the next several years characterizing mutants with defects in RNA and protein synthesis (e.g. Hartwell et al., 1970b); this collaboration continued after Genetics Department Chairman Herschel Roman lured Lee to the University of Washington in 1968. Meanwhile, I had been diverted from my intended graduate study of population ecology by fabulous courses in genetics (Matt Meselson) and cell biology (Keith Porter) that I was required to take as a first-year student. I had also fallen in love with yeast as an experimental organism on the basis of a lab course taught by Nick Gillham, a Chlamydomonas geneticist who had learned about yeast through some exposure as a postdoc. Given my somewhat contrary personality, the fact that no one else in the Boston area was working on yeast at that time only added to its appeal. I solved my immediate problem by working with Guido Guidotti, a protein chemist who was willing to sponsor my study of some yeast proteins (‘so long as they’re interesting’), but I soon also began planning to do a postdoc in Seattle, which at the time had the only concentration of yeast geneticists (three labs) in the country. In November 1968, I approached Herschel at a meeting and soon had arranged to join his Department as a postdoc when I finished my PhD; he kindly agreed to sponsor me although my proposed project (a genetic analysis of the enzymes that I was studying as a graduate student) was unrelated to his own research. In subsequent correspondence, Herschel soon began pushing me toward the laboratory of his new recruit, but I resisted because I wasn’t excited by the papers on RNA and protein synthesis. However, when I visited Seattle in September 1969, Lee told me how he, undergraduate Brian Reid, and first-year graduate student Joe Culotti had recently realized that because of yeast’s budding mode of reproduction, microscopic inspection of the ts mutants after temperature shift would allow recognition of mutants with a variety of specific cell-division-cycle (cdc) defects. Luckily, I had the wit required (it didn’t take much) to see that this project was more exciting than my original plan, and Lee and I soon agreed that I would join his lab. Thus, in July 1970, I happily became Lee’s first postdoc, just a month after the first report on the cdc mutants was published (Hartwell, Culotti and Reid, 1970a). This paper included a description of a third mutant that would later help to define the septins; it was named cdc3 and described as defective in ‘cell separation’. During the next three exhilarating years, I worked mostly on the nutritional and growth control of cell-cycle initiation, but I used the cdc mutants in many of my experiments, and we talked endlessly about them in the lab (often to the short-term detriment of our experiments!). I was particularly captivated by the mutants with grossly abnormal cell morphologies that indicated defects in cytoplasmic rather than nuclear processes (Figure 1.1). These mutants included one that could continue growth and the nuclear cycle but not make buds (cdc24 : Hartwell

CONCEPTION, GESTATION, AND BIRTH

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Figure 1.1 The first attempt to organize the events of the yeast cell cycle into dependent and independent pathways (Figure 3 of Hartwell et al., 1974, a now-famous paper that was originally rejected without review by Nature). Events defined by particular cdc mutants are indicated by the CDC gene number. iDS and DS, initiation and continuation of DNA synthesis; mND and lND, medial and late nuclear division; BE and NM, bud emergence and nuclear migration; CK and CS, cytokinesis and cell separation; HU and TR, the DNA-synthesis inhibitors hydroxyurea and trenimon; MF, the mating pheromone α factor. Note that cdc12 was not included among the cytokinesis mutants because the alleles available at this time were too leaky to allow full characterization. Reproduced with permission from Hartwell et al ., (1974) Science, 183, 46–51, Copyright  1974 Elsevier

et al., 1974) and four that made abnormally elongated buds with multiple nuclei but could not complete cytokinesis (cdc3, cdc10, cdc11 and cdc12 : Hartwell, 1971; Figure 1.2). When I began my own lab at The University of Michigan in 1975 (after a second postdoc in Zürich that focused further on nutritional control), I soon decided to concentrate on these mutants, a decision that was the more

Figure 1.2 Images from the first systematic description of the ts mutants that would later define the septins (Hartwell, 1971); the cells had been incubated for several hours at restrictive temperature and then stained with Giemsa to reveal their nuclei. The defect in cytokinesis was clearly recognized at this time, using a newly developed assay to discriminate cytokinesis from the somewhat later process of separation of the daughter cell walls. (b) cdc3 ; (c) cdc11 . Similar images were also presented for cdc10 and cdc12 mutants. Reproduced with permission from Hartwell, L.H. (1971) Exp. Cell Res., 69, 265–76, Copyright  1971 Elsevier

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appealing to me because it seemed that everyone else (including Lee) was more interested in the mutants with defects in DNA replication and/or nuclear division. My early years at Michigan were dominated by the struggle to get a lab up and running on a miniscule budget, cope with a heavy teaching load, and finish the projects related to nutritional and growth control of the cell cycle. The time and resources available for the morphogenesis mutants were put into further characterizing cdc24 (Sloat and Pringle, 1978; Sloat, Adams and Pringle, 1981) and screening for more mutants with defects in bud emergence. These efforts were quite rewarding (as reviewed elsewhere: Pringle et al., 1995; Pringle, 2006), but they delayed work on cdc3, 10, 11, and 12, and thus the birth of the septins. However, the mutant screen did yield, as a by-product, 28 new mutants with the same phenotype as cdc3, 10, 11, and 12. We were disappointed when all 28 proved to have mutations in the four known genes, but this did help to convince us that the phenotype was highly specific, so that our intended studies would be of manageable scope. Also, we recovered cdc11 and cdc12 alleles (including the now widely used cdc12-6 : Adams and Pringle, 1984) that were tighter, and thus more useful in temperature-shift experiments, than those available previously. During this period, there were also other developments important for what would become the septin field. First, Breck Byers and his outstanding technician Loretta Goetsch, working a few doors down from the Hartwell lab, undertook systematic electron-microscopic (EM) analyses of both wild-type yeast and the various cdc mutants. In wild-type cells, they observed seemingly filamentous structures in close apposition to the plasma membrane in the mother-bud neck (Byers and Goetsch, 1976a; Byers, 1981; Figure 1.3a and b; see also Figure 3.4 of Chapter 3 by McMurray and Thorner). In addition, in the cdc3, 10, 11, and 12 (a)

(c)

(b)

(d)

(e)

Figure 1.3 (a and b) Conventional EM images of the neck filaments (Byers, 1981); these images have better contrast than those published earlier (Byers and Goetsch, 1976a). (c–e) ImmunoEM images obtained using antibodies against (c) Cdc3p and (d and e) Cdc10p (J. Mulholland, B. Haarer, S. Ketcham, D. Preuss, J. Pringle and D. Botstein, unpublished results). (a) and (b) reproduced from Byers, B. (1981) with permission from Cold Spring Harbor Laboratory Press (c–e) ImmunoEM images obtained using antibodies against (c) Cdc3p and (d and e) Cdc10p (Reproduced with permission from Mulholland J., Haarer B., Ketcham S., Preuss D., Pringle J. and Botstein D., unpublished results)


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Figure 1.4 The abstract that reported that certain cdc mutants lacked the ‘neck filaments’ (Byers and Goetsch, 1976b); note that the names of these mutants are not actually mentioned in the abstract, although this information was provided when the poster was presented at the International Congress on Cell Biology (B. Byers, personal communication). Reproduced from Byers, B. and Goetsch, L. (1976a) J. Cell Biol., 69, 717–21, with permission from Rockefeller University Press

mutants, but not in 20 other cdc mutants, they observed that these filament-like structures were lost after temperature shift and that the rates of filament loss in the various mutants correlated with the rates at which cytokinesis ability was lost in the same mutants (Figure 1.4; this is an interesting example of a highly influential study that was never published in detail). Second, my own group’s interest in cell polarization led us to study the actin and microtubule cytoskeletons; this in turn led to the adaptation of immunofluorescence methods to yeast by Alison Adams in collaboration with John Kilmartin in England (Adams and Pringle, 1984; Kilmartin and Adams, 1984). In these studies, we also exploited the elongated-bud phenotype of the cdc3, 10, 11, and 12 mutants to demonstrate the highly polarized distribution of actin structures that first suggested their role in the polarized delivery of secretory vesicles to the tip of the bud. Finally, other yeast geneticists discovered that genes could be cloned by plasmid rescue of the corresponding mutants, and this approach was used to clone CDC10 as a step toward the analysis of the nearby centromere (Clarke and Carbon, 1980). The availability of this CDC10 clone allowed Kaback and Feldberg (1985) to observe that this gene was among those strongly induced during sporulation, a harbinger of later studies showing a role for the septins in this process (see below). In 1983, Sue Lillie and Brian Haarer in my lab finally began systematic molecular analysis of CDC3, 10, 11, and 12, and they were soon joined by several others. Such work was still fairly challenging in those days, and we encountered a number of confusing complications in the course of cloning the genes and establishing that the clones were correctly identified and faithfully represented what was present in the genome. For example, because the common phenomenon of dosage suppression had not been recognized previously, it took us months to

12


realize that the several plasmids isolated by rescue of a cdc11 mutant all actually contained CDC12 (Pringle et al., 1986)! However, we had the clones sorted out by 1986, and Brian had been able to fuse portions of CDC12 in frame to E. coli lacZ and trpE (because we didn’t yet have sequence information, he cloned random fragments of CDC12 into the fusion vectors and screened for clones that produced a fusion protein), use the fusion proteins to make Cdc12p-specific antibodies, and use these antibodies to demonstrate that Cdc12p localized to the mother-bud neck (Haarer and Pringle, 1987). This demonstration required Brian to overcome another confusing artefact that resulted from the presence in most rabbit sera of antibodies that recognized a yeast cell-wall component (probably chitin) that is also localized to the neck and often remains there during the preparation of cells for immunofluorescence. Thus, we had two moments of high excitement in this project: first, when the unpurified antiserum (but not a control serum) gave us staining at the neck, the exact place where we expected to find Cdc12p, and second, when affinity-purified antibodies also stained the neck. Of course, these highs were separated by the profound low of realizing that the first results were an artefact! Before long, we had also obtained good antibodies to Cdc3p, 10p, and 11p, and shown that these proteins also localized to the neck (Figure 1.5a and b; Kim, Haarer and Pringle, 1991; Ford and Pringle, 1991). Importantly, we demonstrated that all four proteins were lost simultaneously from the neck when any one of the four ts mutants was shifted to restrictive temperature; taken together with the results of Byers and Goetsch (see above), this suggested that Cdc3p, 10p, 11p, and 12p formed a complex that contributed to the filament-like structures seen by EM. Further support for this conclusion was obtained in 1991, when Jon Mulholland and Daphne Preuss in David Botstein’s lab obtained some beautiful immunogold images using the antibodies that we provided (Figure 1.3c–e). Although we unfortunately did not find a vehicle for publication of these images, they were shown widely and mentioned in a review (Longtine et al., 1996), and they helped establish two points that were not clear from fluorescence images, namely that the proteins of interest, like the filaments described by Byers and Goetsch, are closely associated with the plasma membrane and are present throughout the neck region (so that the structure in which they are found during most of the cell cycle is an hourglass-shaped band and not a pair of discrete rings, despite the common appearance of fluorescence images resulting from the geometry of the neck). Meanwhile, in the summer of 1986, Brian and others in the lab had begun sequencing the four genes, another process that was rather laborious in those days. In Brian’s report to his thesis committee in December 1986 (which I still have), all four genes were described as partially sequenced, but there was no hint that any sequence similarities had been found. But in early February 1987, I wrote letters of recommendation for Brian that described the ‘very interesting result’ that Cdc3p, 10p, 11p, and 12p formed a family of related proteins. So although I have no record of the precise date, the ‘Eureka moment’ must have been in January 1987. It took quite a bit longer actually to finish the sequencing of both strands and to feel confident that we had all the bugs out of both this and the original


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(a)

(b)

(c)

Figure 1.5 Excerpts from abstracts for talks at the biannual Cold Spring Harbor Yeast Cell Biology Meetings of 1987 (a), 1989 (b) and 1991 (c). In (c), cdc103+ was the gene later named spn1 (Longtine et al., 1996); the Drosophila gene was sep1 (Fares, Peifer and Pringle, 1995); and the mammalian (mouse) genes were Sept1 (Nottenburg, Gallatin and St. John, 1990) and Sept4 (Kato, 1990), originally called DIFF6 and H5 . For a Cold Spring Harbor cytoskeleton meeting in April 1991, we submitted an abstract nearly identical to (c), except that we did not yet know about the mouse genes. Although we were very slow to publish our results, we were not secretive about them! Reproduced with permission from the authors

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characterization of the clones, but we were comfortable with presenting the central results at a major meeting in August 1987 (Figure 1.5a). (After this presentation, Jeremy Thorner referred, prophetically, to the project as a ‘gold mine’, but most other people did not seem very interested in this out-of-the-mainstream work.) It also took many months of hard work to write it all up: the work had been done by seven people (most of whom had already left the lab) over a five-year period; the story had some intrinsic complications; and methods had evolved rapidly during the years that the work was being done (so that the ways we had actually done things were not always how we would have done them at the time of writing). However, by July 1990, we thought that we had satisfactorily addressed every complication and produced a pair of tightly written manuscripts, which we submitted with great pride to Genetics (Figure 1.6). In a telephone conversation on about August 25, the editor indicated that the reviews were generally positive (as indeed they proved to be), and on this basis we described the papers as ‘in press’ when we submitted the Cdc3p-localization manuscript (Kim, Haarer and Pringle, 1991) on August 27. Thus, it was with shock bordering on disbelief that I received an editorial decision letter on August 29 that said, among other unpleasant things, that the manuscripts were ‘intensely irritating to read’ and that I should have known that the two manuscripts would need to be combined into a single manuscript. (Actually, this thought had never even crossed my mind.) The letter was the more shocking in that it came from an editor with whom I had worked on other projects and thought of as someone who shared my deeply held beliefs about the importance of addressing, rather than glossing over, the technical details and complications of a study. I couldn’t really see how to re-write the material as

Figure 1.6 The two manuscripts in which we attempted to report the cloning and sequencing of the original four septin genes were submitted to Genetics in July 1990. The postcards acknowledging receipt are shown. Authors of the first paper were Sue Lillie, Brian Haarer, Laird Bloom, Kevin Coleman, and myself; authors of the second paper were Brian Haarer, Stuart Ketcham, Susan Ford, David Ashcroft, and myself

CHILDHOOD

15

a single paper of reasonable length, and although it was years before I gave up on publishing these papers, both life (I moved from Michigan to North Carolina in 1991, and none of the people involved in this phase of the project moved with me) and science (see below) moved on without its ever getting done. We did talk widely about the results, refer to them in the 1991 protein-localization papers, and provide details and reagents to the few who were interested. We also entered the sequences into the public databases, although, embarrassingly, this didn’t happen until May 1993 (as a result of a miscommunication and a lack of follow-up by me, rather than any attempt to be secretive), with some consequences as described below. At some point in 1989 or 1990, we began in the lab to refer to the protein family as ‘septins’ (for their role in septation), but we were too diffident to apply this term in our 1991 publications, a 1992 paper in which we proposed to introduce the term was aborted (see below), and we published nothing more on septins until 1995. Thus, when publication of the Neufeld and Rubin (1994) Pnut paper (see below) elicited the minireview by Sanders and Field (1994), Sylvia and Chris had to call to ask if we had yet given the proteins a name. I told them, and they kindly used it (with attribution) in their review, thus introducing this convenient term into the literature.

CHILDHOOD As the S. cerevisiae septin picture was coming into focus in the late 1980s, we naturally began to wonder whether such proteins were also found in other organisms. EM observations by Soll and Mitchell (1983) had shown that Candida albicans also had ‘neck filaments’, but even if these proved to contain septins (as indeed found later: see Chapter 5 by Gladfelter and Sudbery), this didn’t seem to expand our horizons very much. Thus, we turned to the fission yeast Schizosaccharomyces pombe, which is more distant phylogenetically from S. cerevisiae and also distinct morphologically in that it divides by medial fission rather than by budding. I thought that we would be competent to handle another yeast in the lab, and this proved more-or-less true, although we also got a huge assist when S. pombe expert Peter Fantes came from Edinburgh on sabbatical early in the project. I also thought that the comparison of septin function in these two yeasts might be highly informative, just as the comparative analysis of cell-cycle control had been (Hartwell, 2002; Nurse, 2002); this has arguably been the case, albeit in the rather perverse way of revealing to us mostly the complexity of septin biology and the major gaps in our understanding of cytokinesis (see below and Chapter 5)! Our initial approach, in 1988, was to screen our antibodies against the S. cerevisiae proteins for cross-reaction with S. pombe proteins; we found a plausible potential homologue only with an anti-Cdc3p antibody (Figure 1.7a). A long struggle by Annette Healy and Hyong Kim with the λgt11 expression-vector system then led eventually to the cloning of the corresponding S. pombe gene (spn1 ),

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(a)

(b)

(c)

Figure 1.7 Initial identification and localization of an S. pombe septin (H. Kim, A. Healy, T. Pugh, P. Fantes, and J. Pringle, unpublished results). (a and b) Recognition of Spn1p in immunoblots. (a) Antibodies raised against Cdc3p (Kim, Haarer and Pringle, 1991) recognized Cdc3p in a blot of S. cerevisiae proteins (lane 1) and a protein of similar molecular weight in a blot of S. pombe proteins (lane 2). As Cdc3p and Spn1p are only ∼45% identical in amino-acid sequence, this cross-reaction sounds a cautionary note for investigators working in organisms with multiple septins. (b) After cloning spn1 , Spn1p-LacZ and Spn1p-TrpE fusion proteins were used to raise and affinity purify antibodies specific for Spn1p, as shown by staining blots of proteins from wild-type cells (lane 1), an spn1 deletion strain (lane 2), and a strain carrying a high-copy spn1 plasmid (lane 3). (c) The antibodies used in (b) were also used for immunofluorescence staining of wild-type S. pombe cells; the cells shown were fixed at the time of cytokinesis. Reproduced with permission from Kim, H.B., Haarer, B.K. and Pringle, J.R. (1991) J. Cell Biol., 112, 535–44, Copyright  1991 Elsevier

and we had enough sequence information to report the result at 1989 summer meetings (Figure 1.5b). Tom Pugh joined the project at this point, and he, Hyong, and Peter (and later Omayma Al-Awar) proceeded both to investigate Spn1p function and to isolate four additional genes (spn2-spn5 ) by a combination of PCR (new to the lab at this time) with degenerate primers and rescue of the S. cerevisiae septin mutants using a library of S. pombe cDNAs in an appropriate expression vector. They found that Spn1p localized to the division site (Figure 1.7b and c) but that knockout of spn1 did not block cytokinesis or septum formation, although it did produce a delay in the separation of the daughter cells. We reported our progress at 1991 summer meetings (Figure 1.5c) and began drafting a manuscript, but our zeal to finish it was reduced by the supposition that the mild phenotype of the spn1 mutant probably resulted from redundancy in function with one or more of the other septins. At this critical juncture came the move to North Carolina and the dispersal of the S. pombe group, of whom only Omayma moved with me. She was joined in a few months by Maria Valencik, and the two of them began to look for additional genes (finding spn6 ) and to construct the required single, double, and multiple mutants (which was challenging in those days because of a paucity of selectable markers for S. pombe gene knockouts and the lack, at the time, of a PCR method for generating the knockout constructs). Discouragingly, even the multiple mutants still displayed only the mild phenotype of cell-separation delay, leaving the puzzle, which endures to this day (see below and Chapter 5), as to why the seemingly conserved septin array at the division site is essential for cytokinesis in some cell types (such as S. cerevisiae) and non-essential in others.

CHILDHOOD

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Maria understandably became discouraged and eventually abandoned the project for a look at the mouse septins (see below); Omayma bravely soldiered on and completed what became half of her thesis, but it was not until 2001 that work by Jürg Bähler and Jian-Qiu Wu had filled in all the gaps (including the identification of spn7 based on a partial sequence released by the S. pombe genome project). In the meantime, we had described many of the results at the 1993 Cold Spring Harbor Yeast Meeting, deposited the sequences in the public databases (although not until August 1995), and described some of the results in our 1996 review (Longtine et al., 1996). Although other groups have now begun to work on the S. pombe septins (see Chapter 5), and we have provided them with information and reagents, we still have considerable material that really ought to be published, and I still have hopes, but with a trail so long and complicated it is not a simple matter. Meanwhile, armed with degenerate PCR and the knowledge that the septins were evolutionarily ancient at least in the fungi, we had been emboldened to look farther afield, and by mid-1990 Johnny Fares had identified a Drosophila gene (sep1 ), which we reported (eliciting no great interest) at meetings in April and August 1991 (Figure 1.5c). Functional analysis of the Drosophila septin was stymied for some time because we had no experience with flies and no appropriate collaborator at Michigan, Johnny’s first attempts to generate antibodies failed, and Johnny himself also had several yeast projects that took much of his time. Then, in May 1991, there was an exciting development: after four years of checking the sequence databases intermittently without success, we noticed a mouse sequence that was clearly homologous to the yeast and fly septins. Nottenburg, Gallatin and St. John (1990) had been trying to clone the gene for a glycoprotein thought to be involved in the binding of lymphocytes to endothelial layers. They recovered a cDNA (‘DIFF6 ’) whose expression levels in different cell lines matched those of the glycoprotein of interest, but, to their disappointment, the clone did not encode that protein, and they found no DIFF6 homologues when they searched the databases (for the reasons described above). Thus, DIFF6 (now SEPT1) has the honour of being the first septin whose sequence was published and entered in the public databases. [Kato (1990) described H5 (now Sept4 ) as one of a set of mouse cDNAs isolated on the basis of their interesting expression patterns in the brain, but the sequence was not entered into the databases until August 1991; Steensma and van der Aart (1991) sequenced a chromosome region that included CDC10 as an early step in the S. cerevisiae genome project, but their sequence (checked against ours, which they knew about from our meeting presentations) wasn’t deposited until May 1992.] Immediately after seeing the DIFF6 sequence, I called Tom St. John (whom I knew slightly from his earlier work on yeast) and told him the septin story, which he was pleased to hear. Remarkably, he had also recently been contacted by George Miklos, a Drosophila geneticist in Australia who had also sequenced a gene encoding a DIFF6 homologue. George’s group was interested in several genes defined by mutations that produced behavioural and/or neurological phenotypes

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and mapped near the centromere of the X chromosome. To identify the genes, they had cloned this chromosome region and were characterizing all the transcription units. Between the genes that proved to correspond to the mutations small optic lobes (Delaney et al., 1991) and sluggish (Hayward et al., 1993) were two other transcribed regions that did not appear to correspond to any of the mutations of interest; one of these had shown the homology to DIFF6 that prompted the contact with the St. John group. On May 14, I sent large packets of information on the septins (including various sequence alignments) to both the St. John and Miklos groups; arriving at the latter without prior warning, I suppose that it produced a stir, especially given that our sep1 proved to be identical to their gene (whose sequence we had not yet seen when I sent our packet). In any case, a cordial and informative letter arrived by fax on May 22 and was followed by a phone call, and we soon agreed to publish a joint paper reporting that proteins homologous to the ‘neck-filament-associated proteins’ in yeast were present in both flies and mammals. I was excited, because I thought that this little paper would markedly raise the level of interest in this family of proteins. I proposed that because St. John and co-workers had not been able to say anything interesting about the DIFF6 sequence when they published it, but had put out the sequence that allowed us all to make contact, they should now be included as co-authors, and this proposal appeared to be accepted (albeit with some reluctance). Work began on the paper but proceeded rather slowly (too slowly for George’s satisfaction) because of the disruption caused by my move in August to North Carolina and the other pressures (notably the editing of the Cold Spring Harbor yeast monograph) that I was facing at that time. George also objected to several aspects of my proposed organization of the paper, as well as to my rather hesitant proposal to call the proteins ‘septins’ (arguing that this was premature without knowledge of their function in animal cells). Nonetheless, by April 1992, after lots of hard work and patient negotiation, I thought that we had a manuscript that all could agree on, and I shipped it off to Australia. Several weeks later, I received a fax announcing George’s intention to withdraw his group from the paper; the main sticking point was the inclusion of St. John and co-workers as co-authors, which was now described (to my astonishment) as ‘outrageous’. This was the last straw for me, and I made no further attempt to rescue the joint publication. The Miklos group entered their sequence into the public databases in July 1992 but never published any of their other information on sep1 , which is a shame because they apparently had genetic data (based on deletion analysis of the region) that would have shed valuable light on Sep1 function. George apparently also considered himself and his group to be the injured parties (which I could never understand), and even after we had finally obtained a good antibody and an appropriate collaborator and published our paper on Sep1 localization and possible functions (Fares, Peifer and Pringle 1995), they declined to acknowledge the septin name or the progress in the field and used their own terminology (‘innocent bystander’ for sep1 and the ‘innocent bystander family’ for the septins) on through their last publication in the area (Maleszka, De Couet and Miklos, 1998).

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During the early 1990s, several other groups encountered mammalian septin genes during studies of other topics (Kumar, Tomooka and Noda, 1992; Nakatsuru, Sudo and Nakamura, 1994; H. Zogbhi, personal communication), but except for an unsuccessful attempt by Makoto Noda’s group (in collaboration with yeast expert Kuni Matsumoto) to rescue the yeast septin mutants using their mouse Nedd5 (now Sept2 ) clone (we sent strains in October 1993, along with lots of information about the yeast and fly septins, and learned of the negative results in December), there was little indication of interest in actually studying septin function. (Although the December letter mentioned that ‘we are raising antibody’, we didn’t learn until much later that Makoto Kinoshita in the Noda lab was leading a serious effort to investigate NEDD5 function, as described below.) Thus, when Maria Valencik became frustrated with the S. pombe project (see above), I encouraged her proposal to begin looking at mammalian septin localization even though our lab was not well equipped for studies of either mice or cultured mammalian cells. We obtained the Diff6 cDNA from Tom St. John and an H5 cDNA from Huda Zoghbi in November 1993 and began trying to make antibodies using fusion proteins, eventually succeeding with both the amino- and carboxyl-terminal halves of DIFF6. (The eventual success was with rabbit antibodies, but this did not happen until after we had first followed the enthusiastic advice of a colleague and tried to raise antibodies in chickens, yielding about half a coldroom full of eggs but no useful antibodies!) Maria used the cDNAs and antibodies to get a variety of interesting results (Figure 1.8), but we still had only fragments of a story when she left the lab in August 1995 (for family reasons and an opportunity to learn mouse methods in a more appropriate environment). Thus, although we briefly described the principal results in our 1996 review (Longtine et al., 1996), no full account of this work was ever published.

EARLY ADOLESCENCE Although a few other groups had happened upon septin genes, as described above, attempts to explore septin function were essentially confined to my laboratory for about 10 years, which of course meant that progress was rather slow and there was little interest by the outside world. Although a paper on cdc10 mutations and yeast budding patterns (Flescher, Madden and Snyder, 1993) was more important than was apparent at the time (see below), I think that the maturation of the septin field really began with the analysis of Drosophila septin function by Neufeld and Rubin (1994). In a complex screen for mutations affecting eye development, Tom Neufeld had recovered a mutation that he named pnut, following the traditional (and maddening to outsiders) practice in Drosophila genetics of giving whimsical names to genes; in this case, the name was based on the superficial resemblance of some cells that had failed cytokinesis, and thus had several nuclei, to peanuts in the shell (T. Neufeld, personal communication). When the gene was sequenced in early 1992, database searches revealed only the DIFF6 and H5 sequences


20 (a)

(b)

(c)

(d)

Figure 1.8 Some early observations on mammalian septins (M. Valencik and J. Pringle, unpublished; Longtine et al., 1996). (a) SEPT1 concentrates in the cleavage furrow (arrow) of at least some dividing mammalian cells (an EL4 lymphoid cell is shown; similar observations were made on NB41A3 neuroblastoma cells). Note that in the coloured originals, the propidium iodide-stained telophase chromosomes (*) were clearly distinguished from the Bodipy-labelled secondary antibody used in the SEPT1 staining. (b) SEPT1 is particularly concentrated in cells of the central nervous system. SDS-PAGE and immunoblotting were performed on protein extracts from various organs of a dissected mouse (1, thymus; 2, spleen; 3, cerebrum; 4, cerebellum; 5, spinal cord; 6, lung; 7, stomach; 8, liver; 9, pancreas; 10, kidney; 11, heart). (c) SEPT1 is concentrated in the growth cones of differentiating PC-12 neural cells in culture. As in (a), the red-fluorescent nucleus (*) and green-fluorescent antibody staining (around the tips of the multiple growth cones) were distinct in the original. (d) Evidence for differential expression and differential splicing of Sept1 and Sept4 in different tissues. A commercially obtained blot of poly A+ RNAs from various mouse organs (shown are 1, heart; 2, brain; 3, spleen; 4, lung; 5, testis) was hybridized to radiolabelled Sept1 (left) and Sept4 (right) cDNAs. Reproduced with permission from Longtine, M.L. et al . (1996) Curr. Opin. Cell Biol., 8, 106–19

and no connection to the yeast proteins or any biological function (see above). Nonetheless, Tom and Gerry were soon in contact with me. (We have been unable to reconstruct exactly how this happened; the path presumably led through Nottenburg and St. John, although George Miklos – who visited the Rubin lab at about this time – may also have been involved.) In March and April 1992 (by phone and mail, and when I visited Berkeley for a seminar that had been arranged months earlier for other reasons), I passed on everything that we knew at that point about the septins, including our progress with Drosophila Sep1. Additional exchanges of information and reagents followed, so that by the time the Pnut paper was submitted in early 1994, Tom and Gerry were able to provide extensive context that added to the impact of their own highly interesting results. The Pnut paper was important for several reasons. First, it was the first conspicuous public announcement that this family of proteins existed in animals and not

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just in budding yeast. [As described above, we had presented this fact at meetings but not in any publication. And although Flescher, Madden and Snyder (1993) had been able to point out that Cdc10p-like proteins existed in flies and mammals, this paragraph was buried within their paper and unlikely to have been noticed by many.] Second, it presented both protein-localization and mutant-phenotype data that were highly informative about septin function in flies. In particular, it showed that the septins were involved in cytokinesis in animal cells as well as in yeast, while also suggesting strongly that they had other roles, notably in the nervous system. Third, it helped to get Chris Field and Tim Mitchison interested in the septins (Sanders and Field, 1994), paving the way for their later seminal contributions to the field (see below). The Rubin lab did not continue studies of the fly septins after Tom Neufeld’s graduation, but my lab carried on with the indispensable collaboration of Mark Peifer, a card-carrying fly geneticist who had joined UNC as an Assistant Professor within a few months of my own move there from Michigan. A combination of genetic, protein-localization, and biochemical studies (Fares, Peifer and Pringle, 1995; Adam, Pringle and Peifer, 2000; Shih et al., 2002; see Chapter 6 by Field et al.) produced additional important information, including (i) evidence for differentiation of function among the fly septins; (ii) evidence that the septins may be non-essential for cytokinesis in some cell types even though they are essential in others; (iii) evidence that Sep2, unlike Pnut, is dispensable for development to adulthood [although this issue is complicated by the possibility of functional redundancy with the very similar (73 % sequence identity) Sep5]; and (iv) evidence that Pnut, Sep1, and Sep2 are all particularly highly concentrated in non-dividing cells of the embryonic central nervous system, suggesting strongly (as did the preliminary data for mouse; see Figure 1.8) that the septins must have important roles in at least some cell types that are unrelated to cytokinesis. Unfortunately, despite this progress and the promise of much more, the fly septin project waned at UNC: Mark lost interest as the other projects in his lab gained momentum; Johnny Fares, Jenny Adam, and Hsin Shih all graduated and were not replaced by other students; and postdoc Karen Hales received an irresistible offer of a faculty position while she still had a year left on her fellowship (and when she was just on the verge of getting the sep5 mutant that would have allowed the critical sep2 sep5 double mutant to be constructed). [We never attempted to generate a sep1 mutant because of the (unrealized) expectation that the Miklos group would eventually publish something on this topic (see above).] I thought that the fly septin project might revive when I moved from UNC to Stanford in 2005, but with more and more of my own attention devoted to our new study of the dinoflagellate-cnidarian symbiosis (we have cloned two anemone septin genes, but it is unlikely that they will become a major focus of the project), I now think that this is unlikely. Thus, it seems to me that study of the septins in the genetically tractable Drosophila system, where septin mutations even give some strong phenotypes, represents an extraordinary opportunity for an enterprising investigator who wants to do something different and important.

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Meanwhile, our labours in Drosophila had also allowed us to contribute in a small but important way (the anti-Sep1 and anti-Sep2 antibodies) to the first serious and successful effort at a biochemical analysis of the long-presumed septin complexes, the breakthrough study by Chris Field, Tim Mitchison, and co-workers (Field et al., 1996). As a research associate in Bruce Alberts’s lab, Chris had been using biochemical approaches to the study of Drosophila cytoskeletal proteins (discovering, among other things, anillin: Field and Alberts, 1995; and see Chapter 6), so that she was well positioned to take a similar approach to the fly septins once her (and husband Tim’s) attention had been drawn to these proteins. This study established or solidified the evidence for multiple important points, including (i) the several septins in a given cell type (in this case, at least Pnut, Sep1, and Sep2) do form a physical complex with each other; (ii) as suggested by their sequence motifs, the septins can bind and hydrolyse GTP; and (iii) the septin complexes can form at least short filaments in vitro (a point of particular interest given the paucity of evidence for septin filaments in vivo other than in S. cerevisiae). With Tim’s student Jen Frazier leading the way, the approach was soon extended to the S. cerevisiae septins (Frazier et al., 1998), leading to similar conclusions (except that much longer filaments could be formed in vitro) and a solid beginning for yeast septin biochemistry (for further discussion, see Chapters 3 and 6). Field et al. (1996) also extrapolated from their structural observations to propose a new model for septin-filament organization in yeast, in which the filaments would run longitudinally along the mother-bud axis rather than in a helix around the neck as Byers and Goetsch (1976a) had proposed and we all had thought ever since. At the time, I thought (and politely mentioned to Chris and Tim) that this model was not justified by the data and indeed was almost certainly totally wrong. However, I soon changed my tune when Mark Longtine showed that cells defective in the protein kinase Gin4p displayed a reorganization of the septins (into thick bars running through the neck) that I found (and, indeed, still find) easy to explain if the septin filaments are longitudinal and hard to explain if they are helical (Longtine, Fares and Pringle 1998). Although the jury is still out on the ultimate validity of this model (see the detailed discussion in Chapter 3), there can be no doubt that it, and the experiments and discussions it has spawned, have contributed greatly to the development of our understanding of septin organization. There were several other important developments in the septin field during the mid-1990s. First, studies by Beth DiDomenico and Yigal Koltin in Candida albicans (first communicated to me in late 1991; later published by DiDomenico et al., 1994), by Michelle Momany and co-workers in Aspergillus nidulans (Momany et al., 1995; Momany and Hamer, 1997), and by Michael Glotzer and Tony Hyman in Xenopus laevis [first communicated to me in October, 1994; abstract and poster presented in 1996 (Glotzer and Hyman, 1996); eventually published in part in 2002 (Mendoza, Hyman and Glotzer, 2002)], taken together with the S. pombe and Drosophila data, made it seem certain that the septins would prove to be ubiquitously present in both fungi and animals.

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Second, Fatima Curcková in Kim Nasmyth’s lab performed a genetic screen to identify yeast proteins that were critical in cells deficient in G1 cyclins; she identified mutations both in CDC12 and in the novel gene CLA4 and showed that cla4 mutations affected septin organization (Cvrcková et al., 1995). Because Cla4p proved to be a protein kinase in the PAK family, activated by Cdc42p and/or other Rho-type GTPases, this was the first step in trying to understand how Cdc42p controls the spatially polarized localization of the septins (among other things), an effort that continues into the present (see Iwase et al., 2006, and references cited therein). Third, in mid-1995 I received an invitation to write a short review on ‘recent progress’ with the septins for Current Opinion in Cell Biology. Recognizing that enough other people were becoming interested that a true septin field was beginning to emerge, and acknowledging (even if tacitly and grudgingly) that some of our early work would probably never be published in detail, I decided that we should try to put the new field on a sound footing by providing a comprehensive review of what was known about the septins to that point (including our own unpublished results and personal communications from others as needed to make the state of the art clear). The resulting paper (Longtine et al., 1996) was rather different from (and about twice as long as!) what the commissioning editors had envisaged, but after some discussion they agreed to publish it anyway. In re-reading this review a dozen years after it was written, it still seems to me that it did quite well in summarizing both what was known at the time and the challenges that lay ahead. Fourth, some important general points about septin function were revealed by studies of the rather specialized process of yeast ascospore formation. To ask if there were additional septins in S. cerevisiae beyond Cdc3p, 10p, 11p, and 12p, Johnny Fares had used degenerate PCR. He recovered and began to study a fifth septin gene. The knockout strain had no detectable vegetative phenotype but had a defect in spore formation, which caused me to start calling friends in the sporulation field. From Mary Clancy [from whose lab Tom Pugh (see above) had come to mine], we learned that our gene was identical to the previously named SPR3 (Ozsarac et al., 1995; Fares, Goetsch and Pringle, 1996), which had been identified much earlier by Mary and Pete Magee (Clancy et al., 1983) as one of a set of genes with SPorulation Regulated patterns of gene expression. Because Mary’s (and collaborator Ian Dawes’s) focus was on the mechanisms of transcriptional regulation (Kao et al., 1989; Ozsarac et al., 1997), there was little information about the function of Spr3p. However, an important role for the septins during sporulation was suggested strongly by the very strong transcriptional induction of SPR3, CDC10 (Kaback and Feldberg, 1985; see above – previously these data had appeared to make no sense), and a sixth septin gene that was revealed by the yeast genome project and named SPR28 (to connect it to SPR3 and minimize the number of gene-name acronyms) when it also proved to be induced strongly during sporulation (De Virgilio, DeMarini and Pringle, 1996). Although the fascinating process of ascospore formation, and the role of the septins in this

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process, remain poorly understood (Fares, Goetsch and Pringle, 1996; De Virgilio, DeMarini and Pringle, 1996; Moreno-Borchart et al., 2001; Neiman, 2005), these early studies provided the first solid evidence for two important general points, namely that an organism can express different subsets of its septin genes, and form complexes containing different sets of septin proteins, in different cell types. Because Maria Valencik’s early studies of the mouse septins had also pointed to at least the first of these conclusions (Figure 1.8), we felt comfortable in making this point quite strongly in Longtine et al. (1996). Fifth, several different lines of investigation led to development of the scaffold model for septin function. In retrospect, the first intimations of this mode of septin action had come much earlier. In the course of their long-running studies of septum formation, Enrico Cabib and his co-workers had reported, based on EM observations that I myself found rather difficult to evaluate, that ‘septum-like structures’ formed at ectopic locations in the cdc3, 10, 11, and 12 mutants (Slater, Bowers and Cabib, 1985). At about the same time, Alison Adams observed by fluorescence microscopy (using a specific dye) that these mutants displayed a diffuse deposition of chitin, in contrast to the normal tight localization of this cell-wall component to a ring around the neck and the primary layer of the septum (Adams, 1984). This observation suggested that Cdc3p, 10p, 11p, and 12p were involved in the localization of the chitin synthase(s), an hypothesis that was reinforced when we later observed that in cells changing shape in response to mating pheromone, the diffuse deposition of chitin (Schekman and Brawley, 1979) was correlated with the presence of the septins in a zone that was much more diffuse than the tight band seen in vegetative cells (Ford and Pringle, 1991; Kim, Haarer and Pringle, 1991). Alison had also observed that the normal concentration of actin at the neck during cytokinesis failed to occur in the cdc3, 10, 11, and 12 mutants (Adams and Pringle, 1984). However, this observation was difficult to interpret at the time (and indeed remained so until 1997, when we finally began to understand the various actin structures that form at the neck during cytokinesis). Thus, in the early 1990s, we had three dots, but two of them were a little fuzzy, and so far as I can reconstruct, we had not connected them into any general model for septin function. This situation might have changed when Mike Snyder and his co-workers analysed a cdc10 mutant that they had isolated in a screen for genes whose products interacted with Spa2p, which they were studying intensively at the time (Flescher, Madden and Snyder, 1993). On the basis of the mutant’s abnormal budding pattern, they proposed (correctly as it turned out) that the septins functioned to localize to the division site some protein(s) that marked future budding sites. However, at the time this paper had little impact (at least on me), partly because the putative marker protein(s) were purely hypothetical at this point and partly because the cell-polarization side of my lab (and of my own brain) felt that the methods used in Mike’s lab to score budding patterns made their conclusions difficult to evaluate. In any event, it was not long before work in my own and Ira Herskowitz’s laboratories identified some of the proteins involved in marking

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potential budding sites and showed that they indeed localized to the division site in a septin-dependent manner (Chant et al., 1995; Sanders and Herskowitz, 1996). In addition, at about this time, Jamie Konopka found that Afr1p, a protein induced by mating pheromone and involved in producing the change of shape in responding cells, interacts with Cdc12p and co-localizes with the septins both in mating cells and (when ectopically expressed) in vegetative cells (Konopka, DeMattei and Davis, 1995), providing another example of a protein whose localization appeared to depend upon the septins. Thus, by 1996, there were lots of dots. However, from re-reading our 1996 review, it does not appear that we had connected them into a coherent general model until a combination of genetic and protein-localization experiments by Doug DeMarini made a persuasive case that the chitin synthase Chs3p was anchored at the neck by a hierarchical scaffold in which Chs3p bound its activator Chs4p, which bound the newly identified Bni4p (necessary for Chs3p localization but not for its activity), which bound the septin complex through Cdc10p (DeMarini et al., 1997). Additional support for the scaffold model soon came from other studies, such as the demonstration by Erfei Bi (Bi et al., 1998) and by Lippincott and Li (1998) that the actin structures at the division site include an actomyosin contractile ring and that the myosin component of this ring is recruited to the budding site early in the cell cycle in a septin-dependent manner. However, I think that the result that really crystallized the scaffold model was the demonstration by Mark Longtine that when the septins reorganize in the absence of Gin4p (see above), the various proteins that display septin-dependent localization undergo a parallel reorganization (Longtine, Fares and Pringle, 1998). From this time forward, additional examples of septin-dependent localization accumulated rapidly, so that by the time Amy Gladfelter bravely led an effort to summarize the known information, there were at least 22 proteins that were known to localize to the neck, in a variety of temporal and spatial patterns, and with a very wide variety of functions, in a septin-dependent manner (Gladfelter, Pringle and Lew, 2001). A similar effort today would certainly at least triple this number, so I think it is well established that at least one role of the septins is to provide a scaffold for the recruitment of other proteins and probably also for their organization at the site to which they have been recruited. However, I also think that it remains a little embarrassing that we still know so little about the molecular details of this recruitment and about which proteins actually interact directly with the septins! (See additional discussion in Chapters 8 and 9).

LATE ADOLESCENCE Although new areas of cell biology are often pioneered by studies in model organisms, they are not usually considered mature (unless they are plant or microbe-specific) until they have captured the attention of mammalian cell biologists. For the septins, the first big step in this direction was the publication by

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Makoto Kinoshita and his co-workers (Kinoshita et al., 1997); this first detailed exploration of mammalian septin function had required more than three years of work since the Noda laboratory began raising antibodies to ‘NEDD5’ (now SEPT2) in 1993 (see above). Among other things, the data presented indicated that septins are important for cytokinesis in mammalian cells as well as in yeast and flies, that GTP binding and perhaps hydrolysis are important for septin function, and that the septins have distinct functions in interphase cells that involve interactions with the actin cytoskeleton. (See further discussion in Chapters 8 and 10.) Other important developments soon followed and included the following. 1. The first links of the septins to human disease (another step toward the full maturity – and funding! – of a field) appeared. The first publication appears to have been the report that a translocation implicated in the genesis of acute myeloid leukaemia fuses the MLL gene to SEPT5 (Megonigal et al., 1998). However, other studies suggesting oncogenic roles also for SEPT9 and SEPT6 soon appeared (Osaka, Rowley and Zeleznik-Le, 1999; Kalikin, Sims and Petty, 2000; Russell et al., 2000; Sorensen et al., 2000; Borkhardt et al., 2001), as did a report that the septins are concentrated in tangled fibres in the senile plaques in the brains of Alzheimer’s disease victims (Kinoshita et al., 1998), suggesting a septin role in this disease. Meanwhile, Barbara Zieger, Jerry Ware, and their co-workers observed that SEPT5 was expressed at high levels in platelets, suggesting a possible role in haemostasis and hence in bleeding disorders (Zieger, Hashimoto and Ware, 1997; Yagi et al., 1998), a hypothesis that was later supported when alterations of secretory function were observed in platelets from a Sept5 knockout mouse (Dent et al., 2002). Relationships between the septins and human diseases are discussed further in Chapters 9, 11, 12, 14, and 15. 2. The very high concentrations of septins in the central nervous system cells of Drosophila (Neufeld and Rubin, 1994; Fares, Peifer and Pringle, 1995) had suggested strongly that the septins had roles unrelated to cytokinesis (which these cells do not do) and hinted that they might have roles in vesicle trafficking (of which these cells do a lot). These ideas gained strong support as papers began to appear documenting both the high concentrations of various septins in non-dividing cells of the mammalian brain and the association of these septins with structures and proteins involved in vesicle trafficking (Caltagarone et al., 1998; Hsu et al., 1998; Kinoshita et al., 1998; Yagi et al., 1998; Beites et al., 1999; Kinoshita, Noda and Kinoshita, 2000; Xue et al., 2000). Understanding exactly what the septins are doing in neurons and in relation to vesicle trafficking there and in other types of cells remain major challenges for the field, as discussed further in Chapters 8, 11, and 15. 3. Bill Trimble, a highly accomplished investigator of vesicle trafficking, became the first major recruit to the septin field from mainstream

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mammalian cell biology when he leaped in with both feet (and four papers) in 1999 (Beites et al., 1999; Trimble, 1999; Xie et al., 1999; Zhang et al., 1999). This initial burst of activity from Bill’s lab provided, among other things, evidence for differentiation of mammalian septin function in different tissues, high levels of expression of some septins in brain, involvement of the septins in vesicle trafficking and secretion, and direct binding of the septins to phospholipids as at least part of the mechanism for septin-membrane association. 4. Ian Macara, a highly accomplished investigator of the many functions of small GTPases, was led to study the septins when they were found as binding partners of the Borg proteins, which his lab had identified previously as apparent effectors of Cdc42 function during cell polarization (Joberty et al., 2001). Ian’s entry into the field had significance well beyond the direct (and substantial) impact of the research contributions from his lab. First, it further legitimized the septins as an object of study by mainstream cell biologists. Second, it was Ian, after he participated in the 2001 ASCB Meeting workshop (see below and Appendix C), who said ‘enough is enough’ to the nightmare of mammalian septin names and led the successful effort to rationalize and simplify the taxonomy and nomenclature of the genes and proteins (Macara et al., 2002). It would be difficult to overstate the value that this contribution has already had, and will continue to have, to the field, and it should be stressed that Ian’s leadership could not have been successful without the cooperation of essentially everyone in the field, all of whom should feel proud of their participation in this community-spirited action. An update on this nomenclature can be found in Appendix B. As the mammalian septin field was taking off, there were also important developments on several other fronts. First, Tri Nguyen, John White, and their coworkers discovered that the Caenorhabditis elegans unc-59 and unc-61 genes, which John had identified many years earlier on the basis of mutants with uncoordinated movements (White, Horvitz and Sulston, 1982), encoded the worm’s two and only two septins and that even a mutant carrying null mutations of both genes produced viable adult worms (Nguyen and White, 1996; Nguyen et al., 2000). This work raised important questions about septin evolution (why do yeasts, flies, and mammals have multiple septins and the worm just two?), structure (do these two septins form assemblies similar to those of the multi-protein assemblies in other organisms, and, if so, how?), and function (why are the worm septins non-essential for cytokinesis in most cells even though they localize to the cleavage furrows as in other cells?) that still engage the field. Second, studies by Yves Barral, Mike Snyder, and co-workers (Barral et al., 2000) and by Peter Takizawa, Ron Vale, and co-workers (Takizawa et al., 2000) established the important point that, at least in yeast, the septins function to restrict the mobility of integral membrane proteins and other cortical components and thus

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allow the cell to maintain distinct mother-cell and bud polarized cortical domains. Whether the septins form the diffusion barrier themselves or by recruiting other proteins that form the actual barrier is still unclear, although some arguments favour the former model (see further discussion in Chapter 4). Third, as in so many other areas of cell biology, the ability to view the behaviour of the septins in real time in living cells by using GFP and other fluorescence tags opened up the study of septin dynamics (Cid et al., 2001; Lippincott et al., 2001). Continuation of these studies in many laboratories, and in other organisms as well as yeast, has provided many insights into septin assembly, organization, and function, as discussed further in various other chapters in this book.

CONCLUDING REMARKS Most of the program for the American Society for Cell Biology’s Annual Meeting is set by the society president and program committee, but a modest number of ‘member-initiated’ workshops can be organized for the Saturday afternoon at the beginning of the meeting. In the summer of 2001, Chris Field called me to suggest that there was enough activity and interest in the emerging septin field to justify organizing such a workshop for the December 2001 meeting. I agreed with enthusiasm, and the workshop was duly proposed to the ASCB, approved, and organized (with Chris doing most of the work). As the program shows (Appendix C), it brought together many of the people who had done interesting work on the septins. Not many things could have caused me to miss this event, but the invitation to Stockholm for the celebration of Lee Hartwell’s Nobel Prize was one of them. My absence did not appear to detract much from what proved to be a very valuable session with multiple beneficial consequences, including the attraction of other investigators into the field, and I think that this session really marked the emergence of the septins as a mature (if still growing) field. From this point forward, to summarize progress with the septins requires a book (i.e. this one) and not a single chapter, so I end my history at this point. Since the 2001 session, there have been two International Septin Workshops (see Appendix C), each of which has been delightful both for its science and for its camaraderie. The first of these workshops was conceived and catalysed by my two co-editors, Hilary Russell and Peter Hall. These workshops have illustrated the growth of the field, our progress, and also how much there is still to do before we really understand the structure, dynamic assembly, and multiple functions (and dysfunctions) of this class of proteins. The next Septin Workshop should be at least as exciting, and I am looking forward to it keenly. In the meantime, for my own part, I will continue, with my group, to try to understand better the question with which we began, and to which I think we still do not really have a very satisfying answer: what do the septins do in cytokinesis, and why is this role essential in some cell types and not in others?

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ACKNOWLEDGEMENTS It is a pleasure to thank the many members of my own group, our collaborators, and the other interested scientists who have helped to build the septin field over the years. To those who have suffered because various parts of the developing story were not published in a timely way (or at all), I can only apologize and express my relief that the field has developed so well despite these lapses. I would also like to thank the National Institute of General Medical Sciences, which has generously supported our work from a time long before it was clear whether the septins would ever turn out to be relevant to human health.

REFERENCES Adam, J.C., Pringle, J.R. and Peifer, M. (2000) Evidence for functional differentiation among Drosophila septins in cytokinesis and cellularization. Molecular Biology of the Cell , 11, 3123–35. Adams, A.E.M. (1984) Cellular Morphogenesis in the Yeast Saccharomyces cerevisiae. Ph.D. Dissertation, The University of Michigan, Ann Arbor. Adams, A.E.M. and Pringle, J.R. (1984) Relationship of actin and tubulin distribution to bud growth in wild-type and morphogenetic-mutant Saccharomyces cerevisiae. The Journal of Cell Biology, 98, 934–45. Barral, Y., Mermall, V., Mooseker, M.S. and Snyder, M. (2000) Compartmentalization of the cell cortex by septins is required for maintenance of cell polarity in yeast. Molecular Cell , 5, 841–51. Beites, C.L., Xie, H., Bowser, R. and Trimble, W.S. (1999) The septin CDCrel-1 binds syntaxin and inhibits exocytosis. Nature Neuroscience, 2, 434–39. Bi, E. et al. (1998) Involvement of an actomyosin contractile ring in Saccharomyces cerevisiae cytokinesis. The Journal of Cell Biology, 142, 1301–12. Borkhardt, A. et al. (2001) An ins(X;11) (q24;q23) fuses the MLL and the Septin 6/KIAA0128 gene in an infant with AML-M2. Genes, Chromosomes and Cancer, 32, 82–88. Byers, B. (1981) Cytology of the yeast life cycle, in The Molecular Biology of the Yeast Saccharomyces: Life Cycle and Inheritance (eds J.N. Strathern, E.W. Jones and J.R. Broach), Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, pp. 59–96. Byers, B. and Goetsch, L. (1976a) A highly ordered ring of membrane-associated filaments in budding yeast. The Journal of Cell Biology, 69, 717–21. Byers, B. and Goetsch, L. (1976b) Loss of the filamentous ring in cytokinesis-defective mutant of budding yeast. (Abstract). The Journal of Cell Biology, 70, 35a. Caltagarone, J., Rhodes, J., Honer, W.G. and Bowser, R. (1998) Localization of a novel septin protein, hCDCrel-1, in neurons of human brain. Neuroreport, 9, 2907–12. Chant, J. et al. (1995) Role of Bud3p in producing the axial budding pattern of yeast. The Journal of Cell Biology, 129, 767–78. Cid, V.J. et al. (2001) Cell cycle control of septin ring dynamics in the budding yeast. Microbiology, 147, 1437–50.

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2 Evolution and conserved domains of the septins Michelle Momany, Fangfang Pan and Russell L. Malmberg Plant Biology Department, University of Georgia, Athens, GA 30602, USA

SEPTIN EVOLUTION Septins were named for their earliest known role in formation of the yeast septum, the cross-wall that partitions mother and daughter cells. In the years since their discovery septins have been characterized in many other fungi and animals, where they are always members of multi-gene families. Most studies of fungal septins have focused on their roles in cell division, shaping emerging protrusions, forming diffusion barriers between compartments and coordinating nuclear division with cell division (Douglas et al., 2005; Gladfelter, 2006). Studies of animal septins have examined roles in cell and nuclear division; however, in contrast to fungal studies, they have also focused on septin roles in vesicle trafficking, apoptosis, cytoskeletal organization (Martinez and Ware, 2004) and, in the case of mammalian septins, links to cancer and neurodegeneration (Hall and Russell, 2004; Kinoshita, 2006; Spiliotis, Kinoshita and Nelson, 2005). In localization studies fungal septins largely form rings shaping new growth or partitions dividing existing cytoplasm (Lindsey and Momany, 2006). Animal septins also localize to division planes, however, they also are frequently seen as filament-like structures colocalizing with actin or microtubules or as punctate spots thought to be involved in vesicle trafficking (Lindsey and Momany, 2006). Fungal septins have been classified by phylogenetic analysis (Momany et al., 2001) and mammalian septins have been classified by primary sequence similarity (Martinez and Ware, 2004). In cross-kingdom phylogenies using sequences from two fungal yeast species and three animal species (Kinoshita, 2003), concluded that orthologous relationships existed within fungal or animal septins, but not The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

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CH02 EVOLUTION AND CONSERVED DOMAINS OF THE SEPTINS

between fungal and animal septins. With the recent availability of more genome sequences, it is now clear that certain fungal and animal septins are orthologous (Pan, Malmberg and Momany, 2007). It is hoped that a better understanding of shared evolutionary history might allow lessons learned from septins in simple model organisms to be extended to mammalian septins.

SEPTINS FORM A DISTINCT SUBFAMILY OF P LOOP GTPases One of the defining characteristics of septins is the presence of G1, G3 and G4 core GTPase domains in the central region of the protein (Field and Kellogg, 1999; Leipe et al., 2002) (Figure 2.1). These well-defined domains form β strands that interact with the nucleotide triphosphate (Leipe et al., 2002; Saraste, Sibbald and Wittinghofer, 1990). Though myosins, kinesins and ras proteins contain closely related GTPase domains, in phylogenetic analyses septins clearly form their own clade, distinct from other GTPases (Figure 2.2).

Septins are in many eukaryotes, but not protists or plants Septins are found in fungi, microsporidia and animals (opisthokonts) (Pan, Malmberg and Momany, 2007). The basal eukaryote Giardia lamblia contains a septinlike sequence that is missing half of the core GTPase domain. Similarly, searches of the genomes of 22 other recently sequenced protists including Dictyostelium discoidium and Entamoeba histolytica returned septin-like sequences, but no true septins. In phylogenetic analysis with septins, myosins, kinesins and ras

Figure 2.1 Typical septin structure. Septin sequences range from about 300 to 600 amino acids. Septins contain the conserved GTP CDC binding domain with three motifs: G1, GxxxxGK[ST] (amino acids 126–135 in S. cerevisiae Cdc3); G3, DxxG (amino acids 204–209 in S. cerevisiae Cdc3) and G4, xKxD (amino acids 280–289 in S. cerevisiae Cdc3). The previously described polybasic region (amino acids 111–116 in S. cerevisiae Cdc3; Casamayor and Snyder, 2003; Versele et al., 2004) is shown as a black box and the previously described ‘septin unique element’ (amino acids 360–413 in S. cerevisiae Cdc3; Versele et al., 2004) is shown as a grey box. S1–S4 mark positions of new septin motifs (amino acids 237–242, 247–259, 261–268, 364–365 in S. cerevisiae Cdc3) and lines below diagram show conserved single amino acid positions (amino acids 117, 295, 300, 339, 360, 396 in S. cerevisiae Cdc3). Many septins also have a predicted coiled-coil domain at the C terminus (amino acids 476–507 in S. cerevisiae Cdc3; Versele et al., 2004) Pan, Malmberg and Momany (2007). Reproduced from Pan et al . (2007) Analysis of septins across kingdoms reveals orthology and new motifs. BMC Evolutionary Biology, 7, 103. Copyright  2007, BioMedCentral Ltd

SEPTINS FORM A DISTINCT SUBFAMILY OF P LOOP GTPases

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CelMyo6

AniCdc42 SceCdc42p CelCdc42 HsaCdc42 HsaRhoF CelRac2 HsaRhoD CelRac1 AniRac1 AniRho2 HsaRhoG HsaRac3 SceRho2 HsaRhoB HsaRac2 HsaRac1 HsaRhoC HsaRhoA CelRho1 AniRhoA SceRho1 SceRho3

CelMyo3 CelMyoII

CelMyo2

HsaMyoVa

RAS

HsaMyoVb AniMyoV

SceSpr3 SceCdc12

SceMyo4p

AniAspC Hsasep4 Hsasep5 Hsasep1

MYOSIN

SceMyo2p

Hsasep2 HsaSept13 CelUnc59 Hsasep6

Hsasep7

Hsasep11 Hsasep8 Hsasep10

SEPTIN CelKrp95

CelUnc61

AniAspB HsaKif3c HsaKif3a

SceCdc3 CelOsm3 Hsasep12

Hsasep3 Hsasep9 AniAspA AniAspD SceCdc10 SceCdc11

HsaKiaa1590

KINESIN

HsaKif13b

SceShs1 AniKif1 SceSpr28 HsaCra HsaKif18A AniAspE

HsaKif4 HsaKif27

AniKip3 SceKip3 HsaKif19 HsaKif2

HsaKif1b

Figure 2.2 Phylogenetic tree of kinesin, myosin, ras and septin GTPases. Bayesian phylogenetic analysis was performed for 74 kinesin, myosin, ras and septin proteins from Aspergillus nidulans, Saccharomyces cerevisiae, Caenorhabditis elegans and Homo sapiens. Analysis was performed using the Mrbayes program run for 1.5 million generations, discarding 400 000 generations as burn-in. Protein designations and GenBank GI numbers follow. From A. nidulans: AniMyoV (67903750), AniRhoA (67539140), AniRho2 (67537566), AniCdc42 (67901000), AniRac1 (67537146), AniKif1 (67541386), AniKip3 (40741666), AniAspA (13398364), AniAspB (1791305), AniAspC (34811845), AniAspD (34148975), AniAspE (34811843). From Caenorhabditis elegans: CelMyo6 (170726), CelMyo2 (37859222), CelMyoII (15718184), CelMyo3 (127737), CelRho1 (17541992), CelRac1 (156424), CelCdc42 (51704309), CelRac2 (115532882), CelKrp95 (9800183), CelOsm3 (22532874), CelUnc59 (17509405), CelUnc61(32566810). From Homo sapiens: HsaMyoVa (119597854), HsaMyoVb (39932736), HsaRhoA (10835049), HsaRhoC (132543), HsaRhoB (4757764), HsaRac1 (249582), HsaRho (74355286), HsaRhoD (20379120), HsaRac3 (118138331), HsaRac2 (20379104), HsaRhoF (13633711), HsaCdc42 (46397381), HsaKif18A (71051935), HsaCra (119571949), HsaKif19 (126215730), HsaKif27 (30794488), HsaKif3a (116283753), HsaKif13b (46852172), HsaKif3c (120660366), HsaKif4 (29351664), HsaKif1b (66347734), HsaKif2 (33187651), HsaKiaa1590 (27529917), HsaSept1 (16604248), HsaSept2 (4758158), HsaSept3 (22035572), HsaSept4 (4758942), HsaSept5 (9945439), HsaSept6 (22035577), HsaSept7 (4502695), HsaSept8 (41147049), HsaSept9 (6683817), HsaSept10 (18088518), HsaSept11 (8922712), HsaSept12 (23242699), HsaSept13, (113418512). From Saccharomyces cerevisiae: SceMyo2 (6324902), SceMyo4 (6319290), SceRho1 (172420), SceRho3 (218474), SceCdc42 (6323259), SceRho2 (1988087), SceKip3 (1723958), SceCdc3 (6323346), SceCdc10 (6319847), SceCdc11 (6322536), SceCdc12 (6321899), SceShs1 (6319976), SceSpr28 (6320424), SceSpr3 (6321496)

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proteins, the septin-like sequences from protists did not group with other clades (data not shown). In fact, they did not even group with each other suggesting that if these sequences are related they have diverged significantly. Neither septins nor septin-related sequences are found in plants.

Septins form five major groups Phylogenetic analysis of 162 septins using Bayesian and maximum likelihood methods showed that they form five major clades (Figure 2.3) (Pan, Malmberg and Momany, 2007). The Group 1 clade contains fungal and animal septins including Saccharomyces cerevisiae Cdc10 and human SEPT3, SEPT6, SEPT8, SEPT9, SEPT10, SEPT11 and SEPT12. The Group 2 clade contains fungal and animal septins including S. cerevisiae Cdc3 and human SEPT1, SEPT2, SEPT4, SEPT5, SEPT7 and SEPT13. The Group 3 clade contains fungal and microsporidial septins and includes S. cerevisiae Cdc11, Shs1 and Spr28. The Group 4 clade contains fungal and microsporidial septins and includes S. cerevisiae Cdc12 and Spr3. The Group 5 clade contains only fungal septins and includes Aspergillus nidulans AspE.

Septin evolution Based on their relationships with other GTPase families Leipe et al., 2002, suggested that septins likely evolved from a bacterial GTPase that was horizontally transferred from the ancestral pro-mitochondrial endosymbiont to eukaryotes. Based on phylogenetic analysis, the septins might have evolved as follows (Pan, Malmberg and Momany, 2007): Either the GTPase septin ancestor was lost early in the lineage leading to plants and retained in the lineage leading to animals and fungi (Figure 2.4), or the horizontal transfer occurred after the divergence of plants. The ancestral septin was likely duplicated before the animal/fungal split and the resulting duplicates became the founding members of Group 1 and Group 2 septin clades. The ancient Group 1 septin duplicated. One paralog lost its C-terminal extension giving rise to Group 1A septins in animals and fungi. The other paralog retained its C-terminal extension giving rise to Group 1B septins in animals, but was lost from fungi. In animals Group 1 septins have undergone significant duplication giving rise to multiple Group 1 paralogs. In contrast, there is a single Group 1 septin in all fungi so far examined. The ancient Group 2 septin gave rise to Group 2A septins in fungi and Group 2B septins in animals. In animals Group 2 septins have undergone significant duplication giving rise to multiple paralogs while a single Group 2 septin is found in all fungi so far examined. In the lineage leading to fungi and microsporidia, Group 1 and Group 2 septins duplicated giving rise to Group 3 and Group 4 septins. Subsequently, Group 1 and Group 2 septins were lost in the lineage leading to microsporidia. Group 3 and Group 4 septins duplicated and diverged, giving rise to multiple fungal paralogs. Because Group 5 septins are found only

SEPTINS FORM A DISTINCT SUBFAMILY OF P LOOP GTPases

39

Saccharomyces cerevisiae Myo Saccharomyces cerevisiae Cdc10 Candida glabrata Hyp3 Eremothecium gossypii Hyp3 0.96/100 0.99/97 Kluyveromyces lactis Hyp3 Candida albicans Cdc10 1.00/100 0.96/100 Debaryomyces hensenii Hyp3 Yarrowia lipolytica Hyp2 Schizosaccharomyces pombe Spn2 1.00/83 Coccidioides immitis Sep3 1.00/59 Aspergillus nidulans AspD 1.00/100 Gibberella zeae Hyp4 0.76/42 Neurospora crassa Hyp3 1.00/84 0.98/82 Magnaporthe grisea Hyp3 0.96/100 Cryptococcus neoformans Hyp3 1.00/90 Ustilago maydis Cdc10 Mus musculus Sept3 1.00/100 Rattus norvegicus Sept3 0.63 1.00/98 Homo sapiens Sept3 Danio rerio Msf 1.00/98 Homo sapiens Sept9 1.00/100 1.00/100 Mus musculus Sept9 1.00/93 Rattus norvegicus Sept9 Xenopus laevis Hyp1 Homo sapiens Sept12 1.00/100 0.55/85 Mus musculus Sept12 0.80/38 1.00/99 Rattus norvegicus Sept12 Drosophila melanogaster Sep5 0.99/92 Drosophila melanogaster Sep2 Anopheles gambiae Hyp2 1.00/78 1.00/100 Mus musculus Sept10a 1.00 /100 Rattus norvegicus Sept10a 1.00/100 Homo sapiens Sept10 Mus musculus Sept10b 1.00/100 0.57/79 Rattus norvegicus Sept10b 1.00/100 Danio rerio Hyp2 1.00/99 Homo sapiens Sept8 1.00/100 Mus musculus Sept8 0.99/89 Rattus norvegicus Sept8 1.00/92 1.00/100 Danio rerio Sept6 1.00/100 Homo sapiens Sept6 Mus musculus Sept6 1.00/100 1.00/97 Rattus norvegicus Sept6 1.00/93 Homo sapiens Sept11 Rattus norvegicus Sept11 1.00/100 0.74/69 1.00/91 Mus musculus Sept11 1.00/100 Caenorhabditis briggsae Hyp2 Caenorhabditis elegans Unc61 Schizosaccharomyces pombe Spn1 Yarrowia lipolytica Hyp1 Saccharomyces cerevisiae Cdc3 1.00/98 1.00/100 0.79/100 Candida glabrata Hyp1 1.00 Eremothecium gossypii Hyp1 1.00/100 0.83 Kluyveromyces lactis Hyp1 1.00/100 Candida albicans Cdc3 Debaryomyces hansenii Hyp1 1.00/60 Aspergillus nidulans AspB 1.00/100 Coccidioides immitis Sep2 Gibberella zeae Hyp1 0.68/54 Neurospora crassa Hyp1 1.00/99 1.00/52 1.00/97 Magnaporthe grisea Hyp1 Ustilago maydis Hyp1 1.00/96 Cryptococcus neoformans Hyp1 Drysophila melanogaster Pnut 1.00/100 Anopheles gambiae Hyp1 1.00/95 0.78/78 Homo sapiens Sept13 Danio rerio Hyp1 1.00/94 1.00/89 Mus musculus Sept7 0.68/71 Rattus norvegicus Sept7 0.97/70 Bos taurus Cdc10 0.85/95 Homo sapiens Sept7 Drysophila melanogaster Sep1 1.00/100 Anopheles gambiae Hyp3 1.00/100 Xenopus laevis SeptA 1.00/100 Danio rerio Nedd5 0.50/69 Rattus norvegicus Sept2 1.00/95 Mus musculus Sept2 1.00/97 0.65/72 0.99/68 Homo sapiens Sept2 1.00/97 1.00/100 Drysophila melanogaster Sep4 Anopheles gambiae Hyp4 Homo sapiens Sept4 1.00/100 Mus musculus Sept4 1.00/100 1.00/98 0.82/70 Rattus norvegicus Sept4 Danio rerio Hyp4 1.00/99 Homo sapiens Sept5 1.00/100 Macaca fascicularis Hyp1 1.00/99 0.99/58 Rattus norvegicus Sept5 1.00/98 1.00/97 Mus musculus Sept5 1.00/100 Homo sapiens Sept1 Mus musculus Sept1 0.81/38 1.00/100 Rattus norvegicus Sept1 Geodia cydonium Sep1 1.00/100 Suberites domuncula Septl Caenorhabditis briggsae Hyp1 1.00/100 Caenorhabditis briggsae Hyp3 0.77/76 Caenorhabditis elegans Unc59 0.97/98

0.97/100

Group 1A

(CDC10 + M_I) Animals + Fungi

Group 1B (M_II) Animals

Group 2A (CDC3) Fungi

Group 2B (M_III & IV) Animals

Figure 2.3 Overview phylogenetic tree of septin gene family. Half-compat consensus phylogram of 1.5 million generations of the MCMC Montecarlo Markov Chain analysis of the Bayesian phylogenetic analysis, discarding 400 000 generations as burn-in. Nodal numbers in front of the slash are posterior probabilities for Bayesian analysis. At the nodes where the maximum likelihood tree topology agrees with the Bayesian analysis, numbers after the slash are bootstrap percentages from maximum likelihood bootstrap analysis using 1024 replicates. Names in parenthesis under Group names indicate the best characterized fungal septin (CDC10, CDC3, CDC11 and CDC12, ASPE) or the mammalian septin classification of Martinez and Ware (2004) (MI, MII and MIII). Pan, Malmberg and Momany (2007). Reproduced from Pan et al . (2007) Analysis of septins across kingdoms reveals orthology and new motifs. BMC Evolutionary Biology, 7, 103. Copyright  2007, BioMedCentral Ltd

40

CH02 EVOLUTION AND CONSERVED DOMAINS OF THE SEPTINS Schizosaccharomyces pombe Spn5 Yarrowia lipolytica Hyp5 Schizosaccharomyces pombe Spn7 0.98/85 Saccharomyces cerevisiae Cdc11 1.00/100 Candida glabrata Hyp5 Eremothecium gossypii Hyp4 1.00/94 0.92 Kluyveromyces lactis Hyp5 0.85/67 1.00/100 Candida albicans Cdc11 0.66/36 Debaryomyces hansenii Hyp6 Encephalitozoon cuniculi Sep3 0.55/51 Kluyveromyces lactis Hyp6 0.52/47 1.00/100 Eremothecium gossypii Hyp7 Saccharomyces cerevisiae Shs1 0.77/56 1.00/99 0.52 1.00/78 Candida glabrata Hyp6 1.00/100 Candida albicans Sep7 Debaryomyces hansenii Hyp5 Yarrowia lipolytica Hyp6 Yarrowia lipolytica Hyp4 0.85/95 0.84/67 0.97/95 Coccidioides immitis Sep1 Aspergillus nidulans AspA 0.52 0.94/97 Pyrenopeziza brassicae Pbs1 Gibberella zeae Hyp3 0.71/74 Magnaporthe grisea Hyp4 0.63/72 0.85/86 0.61/74 Neurospora crassa Hyp4 Schizosaccharomyces pombe Spn3 Ustilago maydis Hyp3 0.63/48 1.00/95 Cryptococcus neoformans Hyp4 Candida albicans Spr28 1.00/91 Saccharomyces cerevisiae Spr28 1.00/100 1.00/96 Candida glabrata Hyp8 0.95/63 Eremothecium gossypii Hyp6 Kluyveromyces lactis Hyp7 0.53 Eremothecium gossypii Hyp2 Kluyveromyces lactis Hyp2 0.90/100 Candida albicans Spr28 0.88/100 Saccharomyces cerevisiae Cdc12 0.90/90 0.57 Candida glabrata Hyp4 0.99/100 Candida albicans Cdc12 0.81/99 Debaryomyces hansenii Hyp2 Yarrowia lipolytica Hyp3 1.00/100 Aspergillus nidulans AspC Neurospora crassa Hyp2 0.56/80 Magnaporthe grisea Hyp2 0.98/70 1.00/90 Gibberella zeae Hyp2 0.54/66 0.86/76 Mucor circinelloides SepA Ustilago maydis Hyp2 0.76/91 1.00/98 Cryptococcus neoformans Hyp2 0.55/69 Schizosaccharomyces pombe Spn4 0.56/97 Schizosaccharomyces pombe Spn6 Yarrowia lipolytica Hyp7 0.99/68 Candida glabrata Hyp7 0.99/87 Debaryomyces hansenii Hyp4 Kluyveromyces lactis Hyp4 1.00/96 0.65/41 1.00/57 Eremothecium gossypii Hyp5 0.90/52 Saccharomyces cerevisiae Spr3 Encephalitozoon cuniculi Sep1 0.98/85 Candida albicans Spr3 Encephalitozoon cuniculi Sep2 Gibberella zeae Hyp6 1.00/100 Neurospora crassa Hyp6 1.00/99 0.55/49 Magnaporthe grisea Hyp6 Cryptococcus neoformans Hyp5 0.67/92 Neurospora crassa Hyp5 0.89/98 Gibberella zeae Hyp5 1.00/100 0.90 Magnaporthe grisea Hyp5 0.81/76 Coccidioides immitis Sep4 Aspergillus nidulans AspE

Figure 2.3

1 2

+5

+3 +4 −1 −2

Group 3 (CDC11)

Fungi + Microsporidia

Group 4 (CDC12)

Fungi + Microsporidia

Group 5 (ASPE)

Fungi

(continued)

Plants

None

Animals

1A, 1B (M_I & II) 2B (M_III & IV)

Fungi

1A (CDC10) 2A (CDC3) 3 (CDC11) 4 (CDC12) 5 (ASPE)

3 (CDC11) Microsporidia 4 (CDC12)

Figure 2.4 Postulated septin evolution. Group 1 and Group 2 septins likely arose before the animal/fungal split. Group 3 and 4 septins likely arose before the fungal/microsporidial split and Group 5 septins likely arose in fungi after the fungal/microsporidial split. Numbers refer to septin Group and + or − represents their gain or loss. Names in parenthesis under Group names indicate the best characterized fungal septin (CDC10, CDC3, CDC11 and CDC12, ASPE) or the mammalian septin classification of Martinez and Ware (2004) (MI, MII and MIII)

SEPTIN DOMAINS

41

in filamentous fungi, they either arose early in fungal evolution and were lost in yeasts or arose more recently.

SEPTIN DOMAINS Motifs with known function Three septin domains have known functions associated with them, though the relevance of the domains for septin assembly, localization and function in vivo is not always clear. The GTPase domain mediates the binding and/or hydrolysis of GTP. The polybasic region mediates association of the septins with phospholipids found in plasma membranes. In some, but not all cases, the C-terminal coiled-coil domains mediate septin–septin interactions.

GTPase domains The G1 core GTPase domain (GxxxxGK[ST]) forms a flexible loop which interacts with the phosphate group of the nucleotide (Bourne, Sanders and McCormick, 1991; Leipe et al., 2002; Saraste, Sibbald and Wittinghofer, 1990). It is the most conserved motif among septins with the consensus conserved in 98 % of 162 septins examined (Pan, Malmberg and Momany, 2007). The residues immediately following the G1 core GTPase domain are also conserved in the septins with [TS][LF] appearing in 96–97 % of septins. The G3 core GTPase domain (hydrophobic residues followed by DxxG) binds Mg2+ and interacts with the β and γ phosphates of GTP (Bourne, Sanders and McCormick, 1991; Leipe et al., 2002; Saraste, Sibbald and Wittinghofer, 1990; Vetter and Wittinghofer, 1999). The G3 consensus for septins (DT[PV]GxG) is found in over 80 % of 162 septins examined (Pan, Malmberg and Momany, 2007). The G4 core GTPase domain (NKxD) determines GTP binding specificity (Dever, Glynias and Merrick, 1987; Leipe et al., 2002). In septins the first position in the consensus (N) is often replaced by A, S or G. However positions two and four of the G4 consensus (K and D, respectively) are found in over 90 % of 162 septins examined (Pan, Malmberg and Momany, 2007).

Polybasic region Septins associate with the plasma membrane and have been shown to bind the membrane phospholipids phosphatidylinositol 4, 5-bisphosphate and phosphatidylinositol 3, 4, 5-triphosphate (Zhang et al., 1999). Deletion and mutation analysis in mammalian cells and later in yeast identified a six amino acid region N terminal to the G1 core GTPase domain that is responsible for the binding of these membrane phospholipids (Casamayor and Snyder, 2003; Zhang et al., 1999) (Figure 2.1). Though individual residues are not conserved, a basic residue (H,

42

CH02 EVOLUTION AND CONSERVED DOMAINS OF THE SEPTINS LPNPRHRKSVKKG*FFQFNLM VVG+

Figure 2.5 Septin polybasic region consensus. Most common amino acid in polybasic region and surrounding area (corresponds to positions 105–126 in S. cerevisiae Cdc3) among 162 septins examined. Bold designates basic residue (H, K or R) found in 51–90 % of septins examined. Underline designates polybasic region identified by Zhang et al. (1999). Asterisk designates highly conserved glycines found in 90 % of septins examined. Plus sign indicates initial glycine in G1 GTPase core domain

K or R) is present in polybasic region positions 1, 2, 5 and 6 of 60–78 % of the 162 septins examined (Pan, Malmberg and Momany, 2007) (Figure 2.5). Glycine immediately follows the polybasic region in 90 % of the 162 septins examined.

Coiled-coil The coiled-coil motif mediates interactions between proteins. This structural motif forms a super helix with heptad repeats and is found in many proteins (Lupas, 1996; Mason and Arndt, 2004). Septins in Group 1A are truncated at the C terminus and so do not contain the region with the coiled-coil motif. In computational structural predictions all Group 1B, Group 2 and Group 4 septins were strongly predicted to form C-terminal coiled-coils (Pan, Malmberg and Momany, 2007). In contrast, 5 of 29 Group 3 septins and all Group 5 septins were not predicted to form C-terminal coiled-coils. Interestingly, previous work showed that the coiled-coil motifs at the C termini of the S. cerevisiae Group 2 septin Cdc3 and the Group 4 septin Cdc12 were absolutely required for septin association and function, while the C-terminal coiled-coil in Group 3 septin Cdc11 was dispensable (An et al., 2004; Casamayor and Snyder, 2003; Versele and Thorner, 2005). The absolute conservation of coiled-coils in all Group 2 and Group 4 septins suggests that interactions mediated by the C-terminal coiled-coils of these groups might also be essential for function across species.

Motifs with unknown function A number of conserved regions with no known function have been identified in septins. Though the functional significance of these motifs is not known, their conservation suggests important roles.

Septin unique element Versele and Thorner (2005) previously identified a conserved 53 amino acid region C-terminal to the G4 core GTPase domain and designated it the septin unique element or SUE (Versele et al., 2004; Versele and Thorner, 2005) (Figure 2.1). Analysis of the SUE in 162 septins showed that just over half of the residues are conserved across 50–93 % of septins (Figure 2.6). When the consensus for

CONCLUSIONS

43

R*XYPW*G*XXEV ENXXHCDFXX LRXXLIRTHX XDLXXXT*XXX HYEXYRXXXL XXX

Figure 2.6 Septin unique element consensus. Most common amino acids in septin unique element identified by Versele and Thorner (2005) (corresponds to positions 360–413 in S. cerevisiae Cdc3). Residues found in at least 50 % of 162 septins examined are shown. Bold designates residues conserved in at least 75 % of septins examined. Asterisks designate single R and T residues conserved in 93 % of septins examined and WG Sep4 motif found in 92 % of all septins examined (Pan, Malmberg and Momany, 2007). Underline designates regions which returned only septins in GenBank searches as described in text

the entire 53 residue SUE was used to query GenBank, no hits were returned. However, querying with SUE residues 5–11 as bait returned 32 hits, all of them septins, and SUE residues 37–46 returned 41 hits, all of them septins. So while there is variation across species, at least potions of the SUE appear to be unique to septins.

Sep1–4 and conserved single residues Alignments of 162 septins revealed four new highly conserved motifs (Sep1–4) and six highly conserved amino acid positions (Pan, Malmberg and Momany, 2007) (Figure 2.1). Each of the two consensus amino acids in the Sep1 motif (ExxxxR, located between the core G3 and G4 GTPase domains) were conserved in 96–98 % of septins examined. Each consensus amino acid in the Sep2 motif (DxR[VI]Hxxx[YF]F[IL]xP, located between G3 and G4 core GTPase domains) was conserved in 88–96% of septins examined. Each consensus amino acid in the Sep3 motif (GxxLxxxD, between the G3 and G4 core GTPase domains) was conserved in 86–96% of septins examined. Each consensus amino acid in the Sep4 motif (WG within the SUE) was conserved in 92 % of septins examined. When used as queries in searches of GenBank, the Sep1, Sep3 and Sep 4 motifs returned proteins in addition to septins. However the Sep2 motif returned 54 proteins, all of them septins. One of the highly conserved single amino acid positions was a glycine immediately following the PB region (Figure 2.5) and two others were within the SUE (Figure 2.6). The remaining highly conserved residues were between the G4 GTPase core domain and the SUE (Figure 2.1).

CONCLUSIONS Our better understanding of the relationships among septins has raised new questions. Perhaps the most obvious question is: What is the significance of the highly conserved SUE and Sep1–4 motifs? This question could be addressed, at least in part, through site-directed mutagenesis targeting these regions. A more difficult question to answer is: Does the shared evolutionary history of Group 1 and Group 2 septins from mammals and fungi translate into shared function or

44


structure? The answer to this question will likely only emerge over time as more researchers investigate septins in different species.

REFERENCES An, H., Morrell, J.L., Jennings, J.L. et al. (2004) Requirements of fission yeast septins for complex formation, localization, and function. Molecular Biology of the Cell , 15, 5551–64. Bourne, H.R., Sanders, D.A. and McCormick, F. (1991) The GTPase superfamily: conserved structure and molecular mechanism. Nature, 349, 117–27. Casamayor, A. and Snyder, M. (2003) Molecular dissection of a yeast septin: distinct domains are required for septin interaction, localization, and function. Molecular and Cellular Biology, 23, 2762–77. Dever, T.E., Glynias, M.J. and Merrick, W.C. (1987) GTP-binding domain: three consensus sequence elements with distinct spacing. Proceedings of the National Academy of Sciences of the United States of America, 84, 1814–18. Douglas, L.M., Alvarez, F.J., McCreary, C. and Konopka, J.B. (2005) Septin function in yeast model systems and pathogenic fungi. Eukaryotic Cell , 4, 1503–12. Field, C.M. and Kellogg, D. (1999) Septins: cytoskeletal polymers or signalling GTPases. Trends in Cell Biology, 9, 387–94. Gladfelter, A.S. (2006) Control of filamentous fungal cell shape by septins and formins. Nature Reviews Microbiology, 4, 223–29. Hall, P.A. and Russell, S.E. (2004) The pathobiology of the septin gene family. The Journal of Pathology, 204, 489–505. Kinoshita, M. (2003) The septins. Genome Biology, 4, 236. Kinoshita, M. (2006) Diversity of septin scaffolds. Current Opinion in Cell Biology, 18, 54–60. Leipe, D.D., Wolf, Y.I., Koonin, E.V. and Aravind, L. (2002) Classification and evolution of P-loop GTPases and related ATPases. Journal of Molecular Biology, 317, 41–72. Lindsey, R. and Momany, M. (2006) Septin localization across kingdoms: three themes with variations. Current Opinion in Microbiology, 9, 559–65. Lupas, A. (1996) Coiled coils: new structures and new functions. Trends in Biochemical Sciences, 21, 375–82. Martinez, C. and Ware, J. (2004) Mammalian septin function in hemostasis and beyond. Experimental Biology and Medicine (Maywood), 229, 1111–19. Mason, J.M. and Arndt, K.M. (2004) Coiled coil domains: stability, specificity, and biological implications. ChemBioChem, 5, 170–76. Momany, M., Zhao, J., Lindsey, R. and Westfall, P.J. (2001) Characterization of the Aspergillus nidulans septin (asp) gene family. Genetics, 157, 969–77. Pan, F., Malmberg, R.L. and Momany, M. (2007) Analysis of septins across kingdoms reveals orthology and new motifs. BMC Evolutionary Biology, 7, 103. Saraste, M., Sibbald, P.R. and Wittinghofer, A. (1990) The P-loop–a common motif in ATP- and GTP-binding proteins. Trends in Biochemical Sciences, 15, 430–34. Spiliotis, E.T., Kinoshita, M. and Nelson, W.J. (2005) A mitotic septin scaffold required for mammalian chromosome congression and segregation. Science, 307, 1781–85.

REFERENCES

45

Versele, M., Gullbrand, B., Shulewitz, M.J. et al. (2004) Protein-protein interactions governing septin heteropentamer assembly and septin filament organization in Saccharomyces cerevisiae. Molecular Biology of the Cell , 15, 4568–83. Versele, M. and Thorner, J. (2005) Some assembly required: yeast septins provide the instruction manual. Trends in Cell Biology, 15, 414–24. Vetter, I.R. and Wittinghofer, A. (1999) Nucleoside triphosphate-binding proteins: different scaffolds to achieve phosphoryl transfer. Quarterly Reviews of Biophysics, 32, 1–56. Zhang, J., Kong, C., Xie, H. et al. (1999) Phosphatidylinositol polyphosphate binding to the mammalian septin H5 is modulated by GTP. Current Biology, 9, 1458–67.

Section II Septins in model systems


3 Biochemical properties and supramolecular architecture of septin hetero-oligomers and septin filaments Michael A. McMurray and Jeremy Thorner Division of Biochemistry and Molecular Biology, Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3202 USA

IMPETUS AND SCOPE Septins are ‘filament-forming GTPases’, but until very recently little was known about how the organization of septin-containing filaments relates to the structure of each of its constituent subunits. How or even whether guanine nucleotide hydrolysis is involved in assembly of higher-order septin structures has also remained an open question. Now, however, more than 30 years of research have reached a watershed with elucidation of the arrangement of septin monomers within hetero-oligomeric complexes from several different species and of how these complexes are polymerized within filaments. These insights have come from intensive biochemical and ultrastructural analysis, especially high-resolution electron microscopy (EM) and X-ray crystallography. Significant progress has also been made in understanding the roles of particular structural elements within an individual septin, and of nucleotide binding and hydrolysis, in the formation of multi-septin complexes and filaments. Our aim here is to interpret the available data so as to present a unified and coherent picture of the biochemical and ultrastructural features of septin complexes and filaments, and to identify key unresolved issues. The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

50

CH03 BIOCHEMICAL PROPERTIES AND SUPRAMOLECULAR ARCHITECTURE

ARRANGEMENT OF SEPTIN SUBUNITS WITHIN COMPLEXES AND FILAMENTS FROM UNICELLULAR EUKARYOTES Core septins of budding yeast (Saccharomyces cerevisiae) Investigation of septin ultrastructure began with the observation of an array of ordered filaments at the mother-bud neck in S. cerevisiae cells (Byers and Goetsch, 1976a). These 10-nm-diameter filaments appeared to be aligned in parallel at ∼28-nm intervals and orthogonal to the mother-bud axis (Byers and Goetsch, 1976a). Of the temperature-sensitive cell division cycle (cdc) mutants isolated by Hartwell et al. (1974) these neck filaments were completely absent in cells carrying mutations in the CDC3 , CDC10 , CDC11 or CDC12 genes (Byers and Goetsch, 1976b). Antibodies against Cdc3, Cdc10, Cdc11 and Cdc12 (Haarer and Pringle, 1987), and later GFP fusions (Cid et al., 1998), demonstrated that all four proteins are located at the bud neck. Moreover, cdc3ts , cdc10ts , cdc11ts and cdc12ts mutants fail to undergo septation (cell division) at the restrictive temperature (Hartwell, 1971). Three related gene products – Shs1/Sep7, which is expressed in mitotically growing cells, and Spr3 and Spr28, which are expressed only in cells undergoing meiosis and sporulation – were identified via their sequence homology to the core septins (De Virgilio, DeMarini and Pringle, 1996; Fares, Goetsch and Pringle, 1996; Carroll et al., 1998; Mino et al., 1998). However, none of these is required for localization of Cdc3, Cdc10, Cdc11 and Cdc12 to the bud neck of a vegetatively growing cell or, presumably, for assembly of Cdc3, Cdc10, Cdc11 and Cdc12 into filaments (although, to our knowledge, the ultrastructure of the bud necks of shs1, spr3 or spr28 cells at the EM level has not been reported). High-stringency biochemical fractionation of yeast cell extracts typically yields a septin-containing complex composed of roughly equal amounts of Cdc3, Cdc10, Cdc11 and Cdc12, having an apparent molecular mass in high salt consistent with a 2 : 2 : 2 : 2 stoichiometry (Frazier et al., 1998; Mortensen et al., 2002). Under conditions of low ionic strength, these purified native septin complexes polymerize into filaments of somewhat smaller diameter (∼5–7 nm) than the neck filaments (Frazier et al., 1998). Likewise, when co-expressed and purified from bacterial (Escherichia coli ) cells, Cdc3, Cdc10, Cdc11 and Cdc12 are necessary and sufficient to form complexes similar in composition and stoichiometry to the native complexes and that polymerize in low salt into filaments that have virtually identical characteristics to those generated by the purified native septin complexes (Versele et al., 2004; Versele and Thorner, 2004; Farkasovsky et al., 2005). Thus, the 2 : 2 : 2 : 2 complex formed by Cdc3, Cdc10, Cdc11 and Cdc12 is the essential building block of yeast septin filaments. Of course, in the cell, other proteins known to co-localize with the septins at the isthmus between a mother and its bud (Gladfelter, Pringle and Lew, 2001) may be responsible for the somewhat

SEPTIN COMPLEXES IN UNICELLULAR EUKARYOTES

51

greater thickness of the endogenous neck filaments and for dictating their spatial organization.

The yeast septin complex is a linear rod Ultrastructural analysis of the septins has focused on examining the features of purified and recombinant hetero-octameric complexes. In conditions of high ionic strength (e.g. 1 M KCl), septin complexes containing Cdc3, Cdc10, Cdc11 and Cdc12 purified from yeast cells (Frazier et al., 1998), or expressed in and purified from E. coli cells (Sirajuddin et al., 2007; Bertin et al., 2008), appear by negative-stain EM as beaded 32-nm-long rods (Figure 3.1a, top). Decreasing the ionic strength of the buffer (e.g. 50 mM KCl) promotes formation of very long filaments of the same diameter as that of the rod (Frazier et al., 1998; Bertin et al., 2008). Depositing septin complexes onto EM grids immediately after lowering the salt concentration yields apparent intermediates that are twice the length of the rod, or three or four times (Figure 3.1a, middle), consistent with the conclusion that the filaments arise from ionic strength-dependent end-to-end polymerization of the rods. Formation of the filaments seems for several reasons to be a highly cooperative process. First, unlike the basic rod itself, the double- and triple-length intermediates are transient and relatively rare. Second, once nucleated, the filaments that form are very long and invariably align in pairs (separated by a gap of 15–25 nm). Moreover, these paired filaments appear to be in register because the ends of each filament in the pair are flush with each other, not staggered (Figure 3.1a, bottom). Under some conditions, still not well defined (but definitely influenced by pH), yeast filament doublets tend to collapse into very large bundles, which are likely non-physiological aggregates. This behaviour may reflect aggregation that occurs upon partial pH-dependent subunit unfolding because an intermediate in the thermal unfolding of a human septin (SEPT4) forms similar amyloid-like fibrils in vitro (Garcia et al., 2007). This property of septins may have implications for human pathophysiology because septins have been found as components of the characteristic neurofibrillary tangles in the brains of patients suffering from Alzheimer’s disease (Kinoshita et al., 1998). In the brain, septins appear to compartmentalize the dendritic spines on neurons, similar to the way they demarcate the boundary between a mother cell and its bud in yeast (Barral and Mansuy, 2007). Averaging the images of very large numbers of classes of individual rods (Sirajuddin et al., 2007; Bertin et al., 2008) affords enhanced resolution that reveals that each rod is composed of eight globular densities of roughly equivalent size and shape, each about 4–5 nm in diameter (Figure 3.1b). These features and dimensions correspond well with those for small GTP-binding proteins (Stewart, Kent and McCoy, 1998), suggesting that the globular density observed reflects, in large part, the conserved GTP-binding domain common to Cdc3, Cdc10, Cdc11 and Cdc12. Moreover, the calculated molecular mass for eight protein spheres


52

of these measured dimensions agrees well with that determined for native and recombinant septin complexes composed of these same four septins in which the apparent stoichiometry is 2 : 2 : 2 : 2. Thus, the appearance of eight globular densities in the 32-nm-long rod viewed by EM is fully compatible with the conclusion that each rod contains two molecules of each of the four different septin polypeptides.

32 nm

4 nm 4 nm

100 nm

(b)

100 nm

Cdc11

Spn3

Cdc12

Spn4

Cdc3

Spn1

Cdc10

Spn2

Cdc10

Spn2

Cdc3

Spn1

Cdc12

Spn4

Cdc11

Spn3

S. cerevisiae

S. pombe

Pnut

SEPT7

Sep2

SEPT6

UNC-59

Sep1

SEPT2

UNC-61

Sep1

SEPT2

UNC-61

Sep2

SEPT6

UNC-59

Pnut

SEPT7

C. elegans

D. melanogaster

100 nm

(a)

(c)

H. sapiens, R. norveticus, M. musculus


53

Arrangement of subunits within the octameric rod To verify the above conclusions and to pinpoint which septin occupies which position within the rod, various strategies can be applied. Use of the two-hybrid method (Fields and Sternglanz, 1994) to examine the hierarchy of interactions that a given yeast septin may have with the others is fraught with potential problems because septin complexes normally assemble in the cytosol (not in the nucleus) and because apparent associations in vivo might arise indirectly from ‘bridging’ through other members of the complex (Farkasovsky et al., 2005). Likewise, even when individual, bacterially expressed and purified yeast septins were examined via in vitro binding experiments, the results were, in some cases, misleading because apparent self-association of a given septin can occur in the absence of its preferred partner septin and the ability of a given septin to associate with itself or another is influenced by the presence of a third (Versele et al., 2004). Some of the non-physiological one-on-one associations observed might be attributable to the fact that the C termini of Cdc3, Cdc11 and Cdc12 each contain segments with 4-3 (heptad) repeats of hydrophobic residues (Figure 3.2) predicted to have the propensity to form a coiled coil with a partner heptad repeat protein (DeMarini et al., 1997; Versele et al., 2004). An analogy is c-Jun, which by itself readily forms homodimers, but forms only c-Fos-c-Jun heterodimers when both proteins are present (O’Shea et al., 1989). In this context, spurious homo-dimerization of septins with the capacity to form a coiled coil is perhaps not surprising. Such

Figure 3.1 EM images of negatively stained core septin complexes from budding yeast and models of the arrangement of subunits in septin complexes of various species. (a) Appropriate vectors were used to co-express Saccharomyces cerevisiae Cdc3, Cdc10, Cdc11 and (His)6 -Cdc12 in Escherichia coli and the resulting heteromeric multi-septin complex was purified in high-salt buffers to near-homogeneity in three chromatographic steps (immobilized metal ion affinity, size exclusion and ion exchange) (Bertin et al., 2008). The resulting complex contained stoichiometric amounts of all four septins in a particle of apparent molecular mass most compatible with a 2 : 2 : 2 : 2 hetero-octamer, as observed previously (Versele et al., 2004; Versele and Thorner, 2004). Samples of the solution of the purified complex were spread under the different conditions indicated on carbon-coated EM grids, negatively stained with uranyl acetate, and viewed in the EM. Top, 300 mM KCl; Middle, diluted from 300 to 50 mM KCl and then immediately prepared for examination in the EM; Bottom, diluted from 300 to 50 mM KCl, incubated for 2 h, and then prepared for viewing in the EM. (b) Individual particles (‘rods’), as in (a, Top), were classified by their appearance and computationally averaged. Image shown is the class average for 1986 particles of the most common class observed, with the dimensions indicated. (c) Models of the number, identity and order of subunits within the rod-like septin complexes from the indicated species. For S. cerevisiae, C. elegans and H. sapiens, the number, identity and order of subunits shown are based on direct experimental evidence generated as described in detail in the text. For other species, the order of subunits and the sizes of the rods are based on low-resolution EM analysis of purified complexes, subunit–subunit interaction data, sequence homology and/or phylogenetic relationships with respect to the corresponding budding yeast and/or human septins, as described in further detail in the text

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CH03 BIOCHEMICAL PROPERTIES AND SUPRAMOLECULAR ARCHITECTURE +

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Cdc3_CTE Cdc12_CTE Cdc12-6_CTE

LGIKQDNSVFKEFDPISKQLEEKTLHEAKLAKLEIEMKTVFQQKVSEKEKKLQKSETELFARHKEMKEKLTKQLKALEDKK ENTGEGNEDFT--LPAIAP-ARKLSHNPRYKEEENALKKYFTDQVKAEEQRFRQW----EQNIVNERIRLNGDLEEIQGKV ENTGEGNEDFT--LPAIAP-ARKLSHNPRYKEEENALKKYFTDQVKAEEQRFRQW----EQNIVNERIRLNGDLEEIQGKV

Cdc3_CTE Cdc12_CTE Cdc12-6_CTE

KQLELSINSASPNVNHSPVPTKKKGFLR. KKLEEQVKSLQVKKSHLK. KKTRRAGQKLASKKIPFKMMIN.

Cdc11_CTE Shs1_CTE

SVAAESIRPNLTKLNGSSSSSTTTRRNTNPFKQSNNINNDVLNPASDMHGQSTGENNETYMTREEQIRLEEERLKAF VANAEEIGPNSTKRQSNAPSLSNFASLISTGQFNSSQTLANNLRADTPRNQVSGNFKENEY(47)PDLPERTKLRNI

Cdc11_CTE Shs1_CTE

EERVQQELLLKRQELLQREKELREIEARLEKEAKIKQEE. SETV--PYVLRHERILARQQKLEELEAQSAKELQKRIQELERKAHELKLREKLINQNKLNGSSSSINSLQQ(23).

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Figure 3.2 The C-terminal extensions (CTEs) of four mitotically expressed septins from budding yeast. As in Versele et al., 2004, the CTE of a septin was defined as that segment of the primary structure from the residue immediately following the ‘septin unique element or domain’ (SUE or SUD) to the C terminus. Gratifyingly, the C-side boundary of the SUE corresponds to the end of α6 helix (see Plate 3.3c, after p. 246), the last well-ordered structural element found in the crystal structures of all three human septins analysed by X-ray diffraction (Sirajuddin et al., 2007). As judged by sequence conservation, the α6 element is present in every septin known in every species that has been analysed to date (Pan, Malmberg and Momany, 2007). Shown, in the one-letter code, are the CTEs of four of the five yeast mitotic septins (Cdc10 lacks a CTE; see Plate 3.3c). Cdc3 and Cdc12 are grouped together, and separately from Cdc11 and Shs1, because the members of each of these pairs are somewhat more related to each other than they are to the other pair. Underlined residues are predicted by a variety of computational algorithms to have strong α-helix forming propensity. Positions marked above or below with a ‘+’ indicate hydrophobic residues (and other compatible side chains) that fit the 4-3 heptad repeat characteristic of sequences known to form α-helical coiled coils. Bold-face type indicates the amino acid substitutions caused by the frameshift mutation in the cdc12-6 allele; and, ‘–’ highlights the absence of a hydrophobic residue at the indicated position. Numbers in parentheses, ‘(47)’ and ‘(23)’, represent stretches of residues of the indicated length present in Shs1, but not found in Cdc11 (or the other two septins)

promiscuous binding seems to extend to certain septin subunits in other species. For example, it has been observed that individually expressed and purified human SEPT2 or SEPT7 homo-dimerize, whereas they only form heterohexameric complexes when they are co-expressed along with another one of their physiological binding partners, SEPT6 (Low and Macara, 2006; Sirajuddin et al., 2007). The ability to reliably image septin complexes in the EM has provided more incisive solutions to surmount these problems. First, one can determine whether any complex is formed, and the structure of that complex seen in the EM, when it is prepared in the absence of one (or more) of the septin subunits. Second, the rods can be decorated with specific antibodies against particular septins. However, when it binds, the antibody must become sufficiently immobilized; otherwise, its position cannot be visualized in the EM. Of course, the primary antigenic site recognized by the antibody must reside at the surface of the folded protein and must not become buried when contact is made with another subunit. To try to avoid the latter problem, a septin can be extended at its N- or C-terminal end with an epitope tag and, if it is competent for incorporation into complexes, they


55

can then be decorated with an appropriate anti-epitope antibody. To be seen, this antibody must also be immobile once bound. Third, a septin can be enlarged at its N- or C-terminal end by fusion to a small monomeric marker protein, such as GFP or E. coli maltose-binding protein (MalE), and if it is competent for incorporation into complexes and thereby immobilized, the location of the extra protein density can be visualized in the EM. It is impossible to conduct the first approach to examine the position of either Cdc3 or Cdc12 in native septin complexes isolated from yeast cells because cdc3 and cdc12 mutants are inviable in any strain background that has been tested. Thus, in the absence of Cdc3 or of Cdc12, the biological processes that require septin function cannot be properly executed. However, the nature of Cdc3-less and Cdc12-less complexes can be assessed by other means. In particular, complexes lacking Cdc3 or Cdc12 have been examined when the remaining three septins were co-expressed in bacterial cells. When Cdc10, Cdc11 and (His)6 -Cdc12 were co-expressed, stoichiometric (His)6 -Cdc12-Cdc11 binary complexes were recovered, but no significant amount of the Cdc10 was incorporated (Versele et al., 2004). The most parsimonious interpretations of this finding are, first, that Cdc11 interacts directly with Cdc12 and that Cdc3 is needed to recruit Cdc10. However, when (His)6 -Cdc3, Cdc10 and Cdc11 were co-expressed, no protein other than (His)6 -Cdc3 was recovered (Versele et al., 2004). Thus, this result suggests that, in the absence of interaction with Cdc12, Cdc3 is not competent to associate with Cdc10. Indeed, Cdc3 and Cdc12 associate avidly with each other and, when co-expressed, Cdc3, Cdc10 and (His)6 -Cdc12 form stoichiometric ternary complexes (Versele et al., 2004). Taken together, these findings already indicated that there is an intrinsic order to the assembly of the octameric rod, with Cdc12 serving as the linchpin since it associates directly with both Cdc11 and Cdc3. In most strain backgrounds, loss of Cdc11 is also lethal (Lee et al., 2002; Versele et al., 2004). However, two groups were able to isolate cdc11 derivatives, which are quite slow-growing and display grossly distended buds (Frazier et al., 1998; Casamayor and Snyder, 2003). When isolated from cdc11 cells, the Cdc11-less septin complexes form rods of somewhat heterogeneous length, but, in general, these are always detectably shorter than those formed by wild-type complexes (Frazier et al., 1998). Quite similarly, when co-expressed in bacteria in the absence of Cdc11, the remaining three septins form complexes containing stoichiometric amounts of Cdc3, Cdc10 and Cdc12 (Versele et al., 2004; Farkasovsky et al., 2005), as mentioned. In the EM, these complexes appear as rods with six clear-cut globular densities instead of the usual eight (Bertin et al., 2008). Thus, Cdc3, Cdc10 and Cdc12 are able to form a stable hexameric complex when no Cdc11 is present. This finding immediately suggests that Cdc11 normally occupies a terminal position in the rod – either a Cdc11-Cdc11 dimer at one end or, more likely, one Cdc11 at each end. Notably, Cdc11-less rods isolated from cdc11 cells (Frazier et al., 1998) or prepared from bacterial cells (Versele et al., 2004) are unable to polymerize into filaments in low salt (although one group

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reported that their Cdc11-less complexes assembled into filament-like structures, but that these were ‘less ordered’ than those generated by Cdc11-containing complexes (Farkasovsky et al., 2005)). Correspondingly, neck filaments cannot be detected in the EM after fixation and staining of cdc11 cells (Frazier et al., 1998). These observations are generally most consistent with the conclusion that Cdc11 molecules situated at the end(s) of octameric rods are necessary for and may directly mediate the efficient end-to-end polymerization required for normal filament formation. Given that the only direct interaction with another core septin detected for Cdc11 is with Cdc12, and that Cdc12 interacts with Cdc3 and that Cdc3, in turn, is able to interact with Cdc10 (and can do so, even when Cdc11 is absent, as long as Cdc12 is present) (Versele et al., 2004; Farkasovsky et al., 2005), already suggests that the arrangement in the rod must include the order: Cdc11-Cdc12-Cdc3-Cdc10. In most strain backgrounds, a cdc10 mutation also causes inviability; however, such a cell is able to pick up as yet unidentified unlinked suppressors or apparent epigenetic modifiers that allow such cells to survive and grow with increasing robustness as they are serially passaged due, perhaps in part, to observed increases in ploidy (McMurray et al., 2008). Nevertheless, for this reason, it has been possible to isolate native septin complexes from Cdc10-less cells (Frazier et al., 1998). Although the Cdc10-less complexes are also somewhat heterogeneous in length, the available images show that they are much shorter on average than the wild-type octameric rod or even the Cdc11-less hexameric rod (Frazier et al., 1998). Because these complexes were obtained by immunoaffinity enrichment using an anti-Cdc3 antibody (Frazier et al., 1998), and given that roughly stoichiometric amounts of Cdc3, Cdc11 and Cdc12 were obtained, these three subunits must be able to associate stably in vivo in the absence of Cdc10. Likewise, when co-expressed in bacterial cells, (His)6 -Cdc12 co-purifies with stoichiometric amounts of Cdc3 and Cdc11, even when Cdc10 is absent (Versele et al., 2004; Farkasovsky et al., 2005). The majority of the images for native Cdc10-less complexes seen on the EM grid seem to represent a Cdc11-Cdc12-Cdc3 trimer (but, because the presence of Cdc11 was somewhat sub-stoichiometric, the shortest forms visible are presumably Cdc3-Cdc12 dimers). When recombinant Cdc11-Cdc12-Cdc3 complexes are viewed in the EM, these are quite clearly trimeric assemblies (Bertin et al., 2008). Accordingly, these results immediately suggest that a Cdc10 homodimer is situated at the centre of the wild-type octameric rod and plays a central role in linking together two flanking Cdc3-Cdc12-Cdc11 trimers. Consistent with this conclusion, both native (Frazier et al., 1998) and recombinant (Versele et al., 2004) Cdc3-Cdc12-Cdc11 complexes are unable to polymerize into long filaments in low salt and, correspondingly, neck filaments cannot be detected in the EM after fixation and staining of cdc10 cells (Frazier et al., 1998). However, Cdc3-Cdc12Cdc11 complexes do form short forms, which likely represent Cdc11-mediated end-to-end polymerization of these trimers into hexamers, which then have a tendency to aggregate into non-specific bundles rather than into the clear-cut paired filaments formed by the wild-type octameric complex (Versele et al., 2004;


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Farkasovsky et al., 2005). In any event, taken all together, an unambiguous order of septin subunits in the octameric rod could be inferred from these observations, namely Cdc11-Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12-Cdc11 (Figure 3.1c). The results of querying the structure of the octameric rod using the other two strategies mentioned above are fully consistent with this arrangement. When added, an anti-Cdc11 rabbit polyclonal antibody (Carroll et al., 1998) can be found bound to both termini of an individual octameric rod and, given their bivalent nature, such ‘Y’-shaped IgG molecules situated at each end often hold two octameric rods together side-by-side (Bertin et al., 2008). These observations definitively establish that Cdc11 is indeed the septin that occupies the outer-most position at both termini of the linear octameric rod. When complexes containing Cdc3, Cdc10, Cdc11 and (His)6 -Cdc12 are prepared, a commercial anti-(His)6 antibody most frequently decorates the second subunit in from one (or both) ends (Bertin et al., 2008), confirming that Cdc12 is the septin that occupies the penultimate position at each end of the octameric rod. It has also been possible to prepare by expression in bacteria complexes that contain Cdc3, Cdc10-GFP, Cdc11 and (His)6 -Cdc12, in agreement with the fact that expression of Cdc10-GFP in vivo is able to fully complement a cdc10 mutation (Cid et al., 1998). When such octamers are examined in the EM, extra density is readily observed and is juxtaposed to the centre of the rod and projects from the fourth subunit from one end and, occasionally, from both of the fourth subunits (i.e. counting in from each end) (Bertin et al., 2008). The fact that most of the Cdc10-GFP-containing rods yield only one extra density per octamer most likely reflects some degree of flexibility in the joint between GFP and Cdc10. Hence, when EM images of particles are grouped together on the basis of a common position for one GFP ‘blob,’ the second GFP presumably has swivelled through a variety of different angles, which tends to cancel out the second GFP density. Nonetheless, this observation confirms that Cdc10 is at the centre of the octameric rod. Similarly, co-expression in bacteria of Cdc10, Cdc11, (His)6 -Cdc12 and a MalE-Cdc3 fusion yields a stable octameric complex with the same overall structure as observed for complexes containing untagged Cdc3, with the exception of extra density juxtaposed to the third subunit from one end and, occasionally, from both of the third subunits (i.e. counting in from each end) (Bertin et al., 2008). Collectively, these findings made using the EM confirm unequivocally that the core yeast septins associate as a linear hetero-octameric rod with the order: Cdc11Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12-Cdc11 (Figure 3.1c). This arrangement demands that polymerization of octamers into filaments occurs via Cdc11-Cdc11 association. Indeed, both two-hybrid analysis (Lee et al., 2002; Casamayor and Snyder, 2003; Farkasovsky et al., 2005) and in vitro binding studies (Versele et al., 2004) showed that Cdc11 is capable of interacting with itself, consistent with the pivotal role of Cdc11-Cdc11 interaction at the octamer–octamer interface. Homotypic pairing of another septin, Cdc10, in the middle of the octamer is also central to the EM-derived structure. Curiously, however, this predicted Cdc10-Cdc10

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interaction was not detectable using the two-hybrid method (Lee et al., 2002; Farkasovsky et al., 2005) and only weak Cdc10-Cdc10 self-association was observed using purified, differentially tagged Cdc10 derivatives (Versele et al., 2004). Given the EM results and the requirements for hetero-octamer assembly deduced from simultaneous expression of various combinations of septins (already summarized above), the most reasonable explanation is that robust Cdc10-Cdc10 interaction requires a conformational change induced in Cdc10 by its prior binding to Cdc3, or, more likely, to a Cdc3-Cdc12 dimer. Such relationships provide further support for the conclusion that proper spatial organization of septin heterooctamers is achieved as the direct result of an ordered pathway for assembly.

Alternative arrangements and substitute subunits There is an additional mitotically expressed septin, Shs1/Sep7 (Carroll et al., 1998; Mino et al., 1998), and two septins expressed only during meiosis and sporulation, Spr3 and Spr28 (De Virgilio, DeMarini and Pringle, 1996; Fares, Goetsch and Pringle, 1996). Therefore, certain alternative arrangements of subunits within yeast hetero-octamers and/or filaments are likely to exist in vivo. To our knowledge, however, the positions of these three proteins within hetero-oligomeric complexes or in filaments have not been delineated. Shs1 is often considered an accessory septin because even null alleles (shs1) have only a mild phenotype (Mino et al., 1998; Iwase et al., 2007) and because it is recovered in sub-stoichiometric amounts in purified septin complexes under certain conditions. When septin complexes are purified from wild-type yeast cells in buffers containing high salt (0.5–1 M ) by immunoaffinity absorption using either an anti-Cdc3 antibody (Frazier et al., 1998) or Protein A-tagged Cdc10 (Farkasovsky et al., 2005), or by co-immunoprecipitation with Gin4, a septin filament-associated protein kinase (Mortensen et al., 2002), Shs1 is present at ≤15 % of the amount of Cdc3, Cdc10 or any of the other septins. However, under conditions of intermediate ionic strength (100–250 mM salt), the complexes recovered contain higher amounts of Shs1, approaching equimolarity with the other septins, regardless of whether the method of purification involves affinity to Cdc3, Cdc10 or Cdc12 (Vrabioiu et al., 2004; Farkasovsky et al., 2005; Sung et al., 2005). This salt-sensitive association suggests that Shs1 interaction with other septins is mainly electrostatic. Perhaps revealing, even when purification is conducted at 100 mM salt, Shs1 is noticeably sub-stoichiometric when purification relies on affinity to Cdc11 (Sung et al., 2005). Conversely, in complexes prepared by immunoaffinity purification of (HA)3 -Shs1, the other septins are present in near stoichiometric amounts, except Cdc11 (Mortensen et al., 2002). In this regard, it is noteworthy that in native septin complexes purified in high salt from yeast cells (which contain Shs1) using either immunoaffinity adsorption to anti-Cdc3 antibody (Frazier et al., 1998) or via Protein A-tagged Cdc10 (Farkasovsky et al., 2005), Cdc11 is present at approximately half the level of the other three septins,


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whereas when Cdc3, Cdc10, Cdc11 and Cdc12 are co-expressed in and purified from bacterial cells (which lack Shs1), the amount of Cdc11 is stoichiometric with the other three septins (Versele et al., 2004). The most parsimonious explanation to reconcile these observations is that Shs1 and Cdc11 compete for the same location in hetero-octamers. Compatible with this view, Cdc11 and Shs1 are closely related at the primary structure level and together occupy a phylogenetic group clearly distinct from the other mitotic S. cerevisiae septins (Pan, Malmberg and Momany, 2007). These considerations predict that there may be three classes of complexes in mitotic cells: Cdc11-Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12-Cdc11; Shs1-Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12-Shs1; and, the mixed oligomer, Cdc11-Cdc12-Cdc3-Cdc10-Cdc10-Cdc3-Cdc12-Shs1. Indeed, various assays to assess the capacity of Shs1 to interact with other septins demonstrated binding only to Cdc12 and Cdc11 (Mino et al., 1998; Versele et al., 2004; Farkasovsky et al., 2005). These latter observations are consistent with the structures proposed and with the idea that Shs1-capped hetero-octamers can interact with Cdc11-capped hetero-octamers, permitting their end-to-end co-assembly into filaments. Nonetheless, because the globular GTP-binding domain of Shs1 contains a large 30-residue insert that the other four mitotic septins lack, and because the C-terminal segment of Shs1 is nearly 100 residues longer than that of the next largest septin, it is possible that the hetero-octamers that contain Shs1 provide a building block that generates filaments with special features or the ability to recruit unique proteins as a means to fine-tune yeast cell morphogenesis. Furthermore, it is possible that the relative proportion of Shs1- versus Cdc11-containing hetero-octamers is tightly regulated, and coupled to events in the cell division cycle (including the status of septin collar assembly), via phosphorylation. This speculation is supported by the fact that, of all of the septins, Shs1 is the most highly phosphorylated and is the target of multiple protein kinases, including bud neck-associated Gin4 (Mortensen et al., 2002; Dobbelaere et al., 2003) and the CDK Pho85 (Egelhofer et al., 2008). Given the roles of septins in subcellular compartmentation and membraneassociated morphogenesis (reviewed in Longtine and Bi (2003); Douglas et al. (2005); Gladfelter (2006); Kinoshita (2006); Spiliotis and Nelson (2006); Barral and Mansuy (2007)), it is not surprising that formation of the novel membrane envelopes around the four meiotic nuclei during sporulation would involve specialized septins, Spr3 and Spr28, expressed only during this developmental event (De Virgilio, DeMarini and Pringle, 1996; Fares, Goetsch and Pringle, 1996). Global transcriptome profiling confirms that both the SPR3 and SPR28 genes are expressed only after the onset of sporulation, and expression of CDC3 and CDC10 is upregulated (Chu et al., 1998; Friedlander et al., 2006). Furthermore, Spr3 and Spr28 co-localize with a select subset of the mitotic septins (Cdc3, Cdc10 and Cdc11) in novel structures associated with the developing spore coats (De Virgilio, DeMarini and Pringle, 1996; Fares, Goetsch and Pringle, 1996), which are distinct from the filamentous septin collar at the bud neck. Although present and detectable throughout sporulation, Cdc12 and Shs1 are excluded

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from the spore-associated structures (McMurray and Thorner, 2008). Moreover, CDC12 and SHS1 (and also CDC11 ) expression is not upregulated detectably during sporulation. It is tempting to speculate (Versele and Thorner, 2005) that Spr3 replaces Cdc12, and that Spr28 replaces Shs1, given that the two subunits in each of these pairs are more closely related to each other at the sequence level than they are to any other septin (Pan, Malmberg and Momany, 2007). These observations suggest that there may be three classes of hetero-octameric complexes in meiotic cells: Cdc11-Spr3-Cdc3-Cdc10-Cdc10-Cdc3-Spr3-Cdc11; Spr28-Spr3-Cdc3-Cdc10-Cdc10-Cdc3-Spr3-Spr28; and, the mixed oligomer, Cdc11-Spr3-Cdc3-Cdc10-Cdc10-Cdc3-Spr3-Spr28. Furthermore, it has been reported that Spr28 is able to interact both with itself and with Cdc11 (De Virgilio, DeMarini and Pringle, 1996; Uetz et al., 2000; Drees et al., 2001), suggesting that the proposed rods would be able to undergo end-to-end polymerization to form filaments of distinctly different composition from those in mitotic cells. Thus, the sporulation-specific filaments presumably possess unique structural characteristics and the ability to associate uniquely with other proteins required for nuclear engulfment and spore membrane formation. Given that the mitotic septin hetero-octamer, once formed, appears to be exceedingly stable in vitro, resisting dissociation through multiple purification steps even in 1 M salt, how is subunit substitution accomplished in vivo? One possibility is that monomeric septins are subject to rapid proteolytic turnover. However, direct examination of two pivotal mitotic septins, Cdc10 and Cdc12, shows that, once synthesized, these individual subunits are practically indestructible, persisting through numerous successive cell divisions (McMurray and Thorner, 2008). Nonetheless, pulse-chase analysis shows that newly made Cdc10 and Cdc12 subunits can be incorporated into pre-formed octamers during vegetative growth (McMurray and Thorner, 2008). This suggests that there are cellular factors that promote subunit exchange, which presumably also allows for insertion of novel subunits for structure-specific purposes. In this regard, it is noteworthy that, even though they are excluded from the spore-associated structures, Cdc12 molecules persist during sporulation. Moreover, they are also precluded from re-assembling into mitotic structures when spores germinate, because only newly made Cdc12 is found at the necks of the first buds (McMurray and Thorner, 2008). This segregation of Cdc12 may be due to a sporulation-specific post-translational modification that cannot be ‘erased’ even in a mitotic cell.

Septins and septin organization in fission yeast (Schizosaccharomyces pombe) As in S. cerevisiae, the S. pombe genome encodes seven septin genes. Four of these (SPN1 , SPN2 , SPN3 and SPN4 ) are expressed during vegetative growth, whereas the remaining three (SPN5 , SPN6 and SPN7 ) are expressed only during sporulation. Primary sequence similarity (Pan, Malmberg and Momany, 2007)


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and other considerations (expression pattern, mutant phenotype, size, etc.) (An et al., 2004) permit identification of an apparent S. pombe counterpart for each S. cerevisiae septin. For the mitotic septins, clearly Spn1 ≈ Cdc3, Spn2 ≈ Cdc10, Spn3 ≈ Cdc11 and Spn4 ≈ Cdc12 (Pan, Malmberg and Momany, 2007). For the sporulation-specific septins, the relationships are less clear, but it seems that Spn5 ≈ Spr28, Spn6 ≈ Spr3 and Spn7 ≈ Shs1. Given the roles attributed to septins in morphogenesis, the fact that S. pombe is bullet-shaped and forms a linear tetrad of spores, whereas S. cerevisiae is ovoid and forms tetrads with tetrahedral geometry, presumably explains the biological basis of some of these differences. Unlike S. cerevisiae, loss of no single septin renders S. pombe inviable, and fission yeast cells remain viable even in the absence of all four of their mitotic septins (An et al., 2004). Nonetheless, absence of any one of the septins causes a pronounced delay in the completion of cell division, resulting in a distinctive morphological phenotype, namely formation of chains of cells. In any event, because every fission yeast septin is non-essential, the effects on septin complex assembly of deleting individual (or combinations of) subunits could be assessed in vivo. When the ability a tagged septin allele to co-immunoprecipitate each of the other septins was examined (An et al., 2004), the resulting data support with few exceptions (noted below) a model for organization of the mitotic septin complex in S. pombe that is reassuringly similar to that of the S. cerevisiae hetero-octamer (Figure 3.1c). First, salt-stable septin complexes from fission yeast contain two copies of each of the four subunits, and are thus hetero-octamers (An et al., 2004). Second, in the absence of Spn2, a stable Spn1-Spn3-Spn4 complex (presumably a 1 : 1 : 1 trimer) still forms and, in the absence of Spn3, a stable Spn1-Spn2-Spn4 complex (presumably a 2 : 2 : 2 hexamer) still forms; but, in the absence of Spn1, only a Spn3-Spn4 complex (presumably a 1 : 1 dimer) is observed and, in the absence of Spn4, only a Spn1-Spn2 complex (presumably a 2 : 2 tetramer) is found (An et al., 2004). These results are fully compatible with a linear hetero-octamer with the order Spn3-Spn4-Spn1-Spn2-Spn2-Spn1-Spn4-Spn3 (Figure 3.1c). Third, in this arrangement, each fission yeast septin occupies a position congruent with that of its budding yeast ortholog. Fourth, and also consistent with this model, Spn2 self-associates in the absence of the other three septins (An et al., 2004). Homotypic interaction of this sort was also observed for Spn1, as seen in vitro (albeit weakly) for its S. cerevisiae ortholog, Cdc3, when its preferred partners are absent (Versele et al., 2004). Likewise, interaction of differentially tagged Spn4 molecules can still be detected when either Spn3 or Spn2 is absent. Because these experiments were performed under relatively low stringency salt conditions (150 mM ) (An et al., 2004), and because the deduced structure of the S. pombe octamer predicts that polymerization should occur via Spn3-Spn3 association, the co-precipitation of differentially tagged forms of Spn4 when Spn2 is absent could be explained by end-on-end joining of Spn1-Spn4-Spn3 trimers to form Spn1-Spn4-Spn3-Spn3-Spn4-Spn1 hexamers. Indeed, under the same conditions, Spn3-Spn3 co-precipitation required Spn1 and Spn4, but not Spn2 (An et al., 2004). Finally, despite the potential pitfalls of the two-hybrid method, especially

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when conducted in a heterologous host (budding yeast) that contains closely related proteins, the observed interactions between the S. pombe septins assessed by this approach agree well, for the most part, with the order predicted above (Figure 3.1c). Spn3 interacts with Spn4, but not with Spn1 or Spn2; and, Spn1 interacts with Spn2 and Spn4, but not Spn3 (An et al., 2004). The sole departure from the predicted arrangement is that a robust interaction of Spn2 with Spn4 was observed (An et al., 2004). The corresponding S. cerevisiae orthologs, Cdc10 and Cdc12, also displayed a modest ability to associate when Cdc3 (Spn1 ortholog) was absent, but a much more robust interaction when all three septins were present (Versele et al., 2004). Even though it is now abundantly clear that the septin complexes from budding yeast (Frazier et al., 1998; Bertin et al., 2008), a nematode (John et al., 2007), an insect (Field et al., 1996) and human cells (Lukoyanova, Baldwin and Trinick, 2007; Sirajuddin et al., 2007) are linear rods of single-subunit width, this has not yet been formally demonstrated for the fission yeast septin complex. Furthermore, to our knowledge, it has not yet been formally demonstrated either in vitro or in vivo that fission yeast septin complexes can, in fact, polymerize into filaments of any sort.

ARRANGEMENT OF SEPTIN SUBUNITS WITHIN COMPLEXES AND FILAMENTS OF INVERTEBRATES The worm’s turn: subunit organization in a minimal septin complex Of all organisms whose genome has been completely sequenced, the one with the fewest number of recognizable septin genes is the nematode, Caenorhabditis elegans (Pan, Malmberg and Momany, 2007), which encodes only two septins, UNC-59 and UNC-61. The phenotype of worms lacking functional versions of either (or both) of the corresponding genes is virtually the same (Nguyen et al., 2000; Finger, Kopish and White, 2003), suggesting that, together, they represent a minimal septin complex whose function relies on both subunits. Biochemical and structural analysis have been performed on UNC-59-UNC-61 complexes expressed in and purified from bacterial cells (John et al., 2007). These behave in many ways like analogously prepared S. cerevisiae Cdc3-Cdc12 complexes (Versele et al., 2004; Farkasovsky et al., 2005). First, based on primary sequence similarity and size (Pan, Malmberg and Momany, 2007), UNC-59 most resembles Cdc12 and UNC-61 most resembles Cdc3. Second, when expressed individually, worm septins are insoluble, as observed especially for native Cdc12 (Versele and Thorner, 2004), but co-expression of UNC-59 and UNC-61 yields a soluble complex, as observed for co-expression of Cdc3 and Cdc12 (Versele et al., 2004). The UNC-59-UNC-61 complex is an equilibrium mixture of heterodimer and heterotetramer, each of which contains equimolar amounts of each septin and no bound nucleotide (John et al., 2007). However, unlike Cdc3-Cdc12 dimers, the

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UNC-59-UNC-61 complex is able to polymerize in vitro into filaments (often paired) with a diameter of 8–9 nm (John et al., 2007), although these structures are rather rudimentary and disorganized compared to the reproducibly long and luxuriant filaments formed from S. cerevisiae hetero-octamers (Bertin et al., 2008). High-resolution EM analysis reveals that the heterotetrameric UNC-59-UNC-61 complex is a linear 17–19-nm-long rod (John et al., 2007). Complexes composed of GFP-UNC-59 and native UNC-61 display extra density localized to the rod termini, whereas complexes composed of GFP-UNC-61 and native UNC-59 display extra density near the centre of the rod (John et al., 2007). Furthermore, in the GFP-UNC-61-UNC-59 complexes, two extra densities were often found at the centre of and on the same side of the heterotetrameric rod (John et al., 2007). Hence, the subunits in the heterotetrameric rod are arranged in the order UNC-59-UNC-61-UNC-61-UNC-59 (Figure 3.1c). Like Cdc3 and Cdc12 (and most septins), both UNC-59 and UNC-61 contain a C-terminal extension (CTE) that contains at its distal end a segment with predicted coiled coil–forming capacity. These elements from UNC-59 and UNC-61 are able to associate with each other to form a parallel heterodimeric coiled coil, as judged by cysteine cross-linking studies (John et al., 2007). Furthermore, in complexes composed of UNC-61-GFP (that is, marked where the end of the coiled coil presumably is) and native UNC-59, the extra density appears at various positions along an arc centred 10 nm from the core of the heterotetramer (John et al., 2007). This observation, combined with the fact that the N-termini of both UNC-61 molecules in a heterotetramer seem to project off the same side of a rod, suggests that coiled coils formed by the C-terminal tails of UNC-59 and UNC-61 project orthogonally off the opposite side of the rod. Thus, the worm heterotetramer, and presumably the budding and fission yeast hetero-octamers, possess twofold rotational symmetry about an axis that runs between and orthogonal to the central septin pair (Figure 3.1c), the only obvious difference being the number of subunits. How did C. elegans ‘lose’ four subunits per septin complex, relative to the yeast octamers? It seems just as likely that other organisms have gained subunits through gene duplication and divergence, generating derivatives that retained the capacity to be incorporated into the rods. For example, the fact that GTP-binding domains of Cdc10 and Cdc12 are more closely related to each other (40 % identity) than either is to any other mitotic septin suggests that Cdc10 could have arisen from duplication and divergence of an ancestral CDC12 -like gene followed by deletion of its C-terminal end. Since the C-terminal coiled coil of Cdc12 is not essential for Cdc12-Cdc12 interaction (Versele et al., 2004; Farkasovsky et al., 2005) (see below), Cdc10 retained the capacity for self-pairing and association with Cdc3, without interfering with the formation of coiled coil–mediated Cdc3-Cdc12 dimers. Of course, alternative scenarios are also possible. Based on phylogenetic relationships (rather than on one-on-one comparisons), and aside from its substantially longer C-terminal end, UNC-61 appears most related to Cdc10 (Pan,

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Malmberg and Momany, 2007). Thus, the Ur-septin to both Cdc10 and UNC-61 may have lacked a CTE and played the ‘central homodimer’ role, which has been conserved; however, over the course of time, UNC-61, but not Cdc10, underwent an illegitimate recombination event that fused it to a sequence with coiled coil–forming capacity, thereby further ‘accessorizing’ this worm septin, making it more like Cdc3. In the absence of capping by a Cdc11-like septin, the Cdc12-like UNC-59 may have retained an inchoate ability for self-association, allowing for end-to-end polymerization of UNC-59-UNC-61-UNC-61-UNC59 rods into filaments.

A word from the Pnut gallery The first Drosophila melanogaster septin gene identified was called pnut, on the basis of the vestigial peanut-shaped imaginal discs found in larvae defective in the corresponding gene (Neufeld and Rubin, 1994), and the genome of this fly encodes four other septin genes. Native multi-septin complexes have been isolated from D. melanogaster (Field et al., 1996) in a manner similar to that used to enrich for endogenous yeast septin complexes (Frazier et al., 1998). Fly embryo extracts incubated with a Pnut-specific antibody yield a salt-stable complex containing, in addition to Pnut, stoichiometric amounts of Sep1 and Sep2 (Field et al., 1996). When dialysed into low salt and examined in the EM, this complex forms filaments of 7–9-nm-diameter that tend to associate laterally and form large bundles at high concentration (Field et al., 1996). In high salt, the complex seen in the EM is a linear ∼26-nm long rod (Field et al., 1996). Based on the dimension of an individual yeast septin subunit (∼4 nm), this length corresponds well to that expected for a linear heterohexameric rod that presumably contains two molecules of each of the three co-purified Drosophila septins. The correspondence between fly septins and their fungal counterparts are not perfectly obvious from either one-on-one comparisons or phylogenetic analysis. In terms of sequence identity (40 %) and match length (435 residues), Pnut (539 residues) is most similar to Cdc3 (520 residues). For the same reasons (42 % identity over 322 residues), Sep1 (361 residues) most resembles Cdc10 (322 residues). In contrast, Sep2 (419 residues) is much more ambiguous. It bears nearly the same degree of similarity to all of the core mitotic septins of yeast; however, it is slightly closer in both size and identity to Cdc12 (407 residues; 34 %) than it is to Cdc11 (415 residues; 32 %). Thus, it is tempting, on these grounds, to speculate that the Pnut-containing septin complex has the order Sep2-Pnut-Sep1-Sep1-Pnut-Sep2. However, to our knowledge, there are no pair-wise interaction data to support this particular arrangement. A slightly different ordering of the subunits within the purified Drosophila hexamer will be proposed below (see also Figure 3.1c), based on structural information about a hexameric complex of human septin subunits that are perhaps more closely related to the fly septins.

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ARRANGEMENT OF SEPTIN SUBUNITS WITHIN COMPLEXES AND FILAMENTS OF VERTEBRATES Onward and upward in evolution and resolution: mammalian septin complexes The preceding presentation of septin complex organization and ultrastructure in several model eukaryotes has set the stage for discussing mammalian septins, in all their resplendent complexity. As detailed elsewhere in this volume, the Homo sapiens genome encodes 14 septin genes, most of which produce an array of products via differential splicing and/or via use of alternative translation initiation sites (Macara et al., 2002; Kinoshita, 2003; Hall and Russell, 2004; Joo, Tsang and Trimble, 2005). This situation makes the combinatorial complexity of potential multi-septin complexes dauntingly high. Only a few such complexes have been analysed in a way that is informative about their ultrastructure. However, this includes the crystal structure of a heterohexameric complex of human SEPT2, SEPT6 and SEPT7 (Sirajuddin et al., 2007), which has provided new and important insights about the interfaces by which individual septin subunits interact with their neighbours, as discussed in detail below.

Rods are the rule: mammalian septins form linear complexes Immunoaffinity isolation of a given septin and its associated proteins from mammalian cells produces complexes containing different septin members, depending on the cell type and the subunit harbouring the epitope used for purification. In mouse brain lysates, anti-SEPT2 antibodies enrich for complexes containing nine different septins, among which SEPT2, SEPT4, SEPT6 and SEPT7 are the most prominent and, together, represent a complex of roughly 1 : 1 : 2 : 2 stoichiometry (Kinoshita et al., 2002). When extracted from a human cell line (HeLa), anti-SEPT2 antibodies pull down a simpler complex containing, as the other septins, only SEPT6 and SEPT7 (Kinoshita et al., 2002). Using anti-SEPT7 antibodies, lysates of rat brains yield a complex containing SEPT2, SEPT4, SEPT6 and SEPT7, but only three (SEPT4, SEPT6 and SEPT7) appear equimolar (Hsu et al., 1998). When examined in the EM, rods were observed in the preparations from the rat; the rods were ∼8 nm in width and ∼25-nm long with ∼10-nm-long thin ‘strands’ projecting laterally from one side of the rod, and were occasionally seen to be associated end-to-end (Hsu et al., 1998). The strands were evenly spaced ∼16 nm apart, with one or two strands per 25-nm rod (Hsu et al., 1998). The significance of these features was not appreciated at the time, but in light of the preceding discussion, it is clear that the strands represent the coiled coils formed between component subunits in the rod. Which of those subunits in the rod are able to make such associations was revealed by the structure determined by X-ray diffraction, described in detail below.

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In addition to recovery of native complexes, human SEPT2, SEPT6 and SEPT7 have been co-expressed heterologously both in insect cells (Kinoshita et al., 2002) and bacterial cells (Sheffield et al., 2003; Sirajuddin et al., 2007), and yield a stable complex with 1 : 1 : 1 stoichiometry. Notably, each full-length septin was insoluble when expressed individually in E. coli (Sheffield et al., 2003), as has also been observed for SEPT4 (Garcia et al., 2007), whereas any pair was soluble and formed a 1 : 1 stoichiometric complex (Sheffield et al., 2003). As judged by size exclusion chromatography (Kinoshita et al., 2002; Sirajuddin et al., 2007) or dynamic light scattering (Sheffield et al., 2003) in 100–500 mM salt, the apparent molecular size of the purified SEPT2-SEPT6-SEPT7 complex is most compatible with a 2 : 2 : 2 hexamer. When examined in the EM, this complex appears as a linear rod that is 5–10 nm in width (Kinoshita et al., 2002; Sheffield et al., 2003; Sirajuddin et al., 2007) and 250-nm long (Sirajuddin et al., 2007), with a substructure composed of six roughly globular subunits each ∼4 nm in diameter (Sirajuddin et al., 2007). Aside from being two subunits shorter, this ultrastructure is virtually identical to what is observed for the yeast hetero-octamer (Figure 3.1b) (Bertin et al., 2008). Also, its length (25 nm) is nearly identical to that of the Drosophila Sep1-Sep2-Pnut (∼26 nm) and rat SEPT4-SEPT6-SEPT7 (∼25 nm) complexes described earlier, suggesting these non-human septin complexes are also 2 : 2 : 2 hexamers. Decreasing the salt concentration allows the human SEPT-2-SEPT6-SEPT7 rods to self-assemble into a variety of structures, including filaments, filament bundles and even rings (Kinoshita et al., 2002; Sirajuddin et al., 2007).

Crystal structure of the human SEPT2-SEPT6-SEPT7 complex Near atomic level structures for crystals of an individual purified septin (SEPT2) ˚ resolution) and the physiologically relevant SEPT2-SEPT6-SEPT7 complex (3.4 A ˚ resolution) have been determined by X-ray crystallography introduced above (4 A (Sirajuddin et al., 2007). The first striking conclusion to be drawn from these two structures is that, both when alone and in the complex, SEPT2 homodimerizes; but, in the isolated SEPT2-SEPT2 homodimer crystal, one of the interfaces by which monomers are joined is completely different from the interface by which SEPT2 self-associates when it is located at the centre of the SEPT2-SEPT6-SEPT7 hexamer (Sirajuddin et al., 2007). This observation clearly illustrates that any given septin has the ability to interact with other septins in multiple ways that depend on the identity of the other available subunits, and implies that a variety of septin complexes differing in composition and subunit arrangement could be achieved in vivo simply by altering the expression, localization, or modification of the different subunits and/or their isoforms. Second, although the structure of the heterohexamer confirmed many aspects of the ultrastructural model for the yeast complex (Figure 3.1c), it also revealed unforeseen features vital for understanding how these structures assemble. That


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the conformation of each septin subunit is dominated by its GTP-binding domain fold (‘G domain’) was expected, given the sequence conservation of this domain in all septins (Pan, Malmberg and Momany, 2007) and the dimensions (4-nm diameter) of the globular densities viewed in the EM (Figure 3.1b). The G domain in SEPT2 and in the other two human septins is very much like the canonical G domain in Ras, with five α-helices and six β-strands (Sirajuddin et al., 2007) (see Plate Figure 3.3a, after p. 246). However, septins have some important additional accoutrements that are critical for conferring the ability to form heteromeric complexes. At their N termini, septins have an additional helix (dubbed α0 so as to preserve the numbering of the first Ras-like α-helix as α1), which can be further extended by additional non-helical sequence (e.g. in SEPT6). Compared to Ras-like GTPases, the G domain of a septin also has an extra helix situated between α4 and β6, two more β-strands between β6 and helix α5 and another helix (α6) located just preceding the CTE present in most septins (Sirajuddin et al., 2007) (Plate Figures 3.3a and 3.3c). The CTE contains at its distal end the heptad repeat sequences predicted to be able to form an α-helical coiled coil. In fact, the sequence that encompasses the structural elements β7, β8 and α6 was already recognized as diagnostic of septins (as opposed to Ras-like GTPases or any other class of known GTP-binding protein) and was, for that reason, dubbed the ‘septin unique’ domain (SUD) or septin unique element (SUE) (Versele et al., 2004). As is necessary for a linear array of similar subunits that possesses twofold rotational symmetry about the axis running orthogonal to and between the central doublet, there must be two types of interfaces between any given pair of ˚ 2 ) comprises a G domain-G domain interaction subunits. One interface (1851 A dominated by contacts in and around the guanine nucleotide-binding site in each monomer, and was dubbed a G dimer. The other significantly more extensive ˚ 2 ) involves contacts involving the extra N-terminal (α0) and interface (2995 A C-terminal (α6) helices that distinguish septins from Ras-like small GTPases, and was dubbed an NC dimer. Individual residues identified as playing key roles in each of these septin–septin interfaces are, as expected, well conserved (Sirajuddin ˚ et al., 2007), although it should be noted that, at the resolution achieved (4 A), many side chains in the final refined structure were modelled as alanines, and several loops and other segments (most notably the region after α6) were either disordered or deleted in the derivatives used to prepare the crystals (Sirajuddin et al., 2007). The structure of the human septin heterohexamer was obtained in sufficient detail to determine that the order of subunits is SEPT7-SEPT6-SEPT2-SEPT2SEPT6-SEPT7. Crystals of SEPT2 alone show that it is able to self-associate in both the G dimer and NC dimer modes, whereas in the hexamer the SEPT2SEPT2 pair is clearly a dimer in the NC orientation. This arrangement forces the SEPT2-SEPT6 interface to be a G dimer and, as a result, the SEPT6-SEPT7 interface is an NC dimer (Plate Figure 3.3a). Hence, these two kinds of interfaces alternate along the rod. Thus, if the hexamer were extended by an additional subunit on each end, as seen in the yeast hetero-octameric rod, the interface

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between SEPT7 and that other septin (and presumably between Cdc12 and Cdc11; Plate Figure 3.3b) is predicted to be in the G-dimer mode. Therefore, when yeast hetero-octamers polymerize end-on-end, the Cdc11-Cdc11 interface will be an NC dimer, whereas when the human heterohexamer polymerizes end-on-end, the SEPT7-SEPT7 interface will be a G dimer. This latter difference could explain the more robust filament-forming capacity of the yeast complex (and the differences in the resulting structures formed) as compared to the human complex. Given the order of subunits within the human SEPT2-SEPT6-SEPT7 hexamer established by crystallography, the order of the orthologous subunits in the corresponding purified mouse and rat complexes is undoubtedly the same (Figure 3.1c). Phylogenetic analysis suggests that D. melanogaster Sep1 is in the same category as human SEPT2, that fly Sep2 is most related to SEPT6, and that Pnut is grouped with human SEPT7 (Pan, Malmberg and Momany, 2007). Hence, we favour, as a model for the fly hexamer, the order Pnut-Sep2-Sep1-Sep1-Sep2-Pnut (Figure 3.1c). It should be recalled that the rat complex subjected to EM analysis was a SEPT4-SEPT6-SEPT7 complex (Hsu et al., 1998). Given that SEPT4 is significantly more related (61 % identity) to SEPT2 than to either SEPT6 (40 % identity) or SEPT7 (53 % identity), this complex presumably represents a SEPT7-SEPT6-SEPT4-SEPT4-SEPT6-SEPT7 heterohexamer. Similarly, the mouse complex obtained using anti-SEPT2 antibodies had a apparent composition of SEPT2, SEPT4, SEPT6 and SEPT7 of 1 : 1 : 2 : 2 (Kinoshita et al., 2002). Thus, these preparations could represent an equimolar mixture of SEPT7-SEPT6-SEPT2SEPT2-SEPT6-SEPT7 and SEPT7-SEPT6-SEPT4-SEPT4-SEPT6-SEPT7 rods that have associated end-on-end to form mixed polymers.

Cellular asymmetry and septin filament polarity To date, every heteromultimeric septin complex analysed at sufficient resolution bears two-fold rotational symmetry about the axis that runs through and orthogonal to the centre of the rod. Consequently, each rod is non-polar in its long dimension and, thus, the two ends (if intact) present indistinguishable surfaces for end-to-end interaction. Therefore, filament assembly in solution should be able to occur by addition of a second complex to either or both ends of a rod, such that a filament can potentially grow in both directions. To our knowledge, however, kinetic studies of polymerization to confirm this prediction have not been performed (for example, using rods with a recognizable fiduciary mark as the seed to nucleate polymerization of an excess of unmarked rods). It is theoretically possible that addition of a second complex to an initial rod may constitute a ‘priming’ step in filament growth, if complex addition at one end of a rod causes a conformational change that is propagated through the rod to its other end and makes that unoccupied terminal subunit more competent to associate with another rod. In this way, once started, polymerization of rods into filaments would be greater than first order with respect to rod concentration. In principle, however, other


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processes could differentially affect the otherwise identical subunits at each end of a rod and cause one end of a rod to be favoured for polymerization or, conversely, could cap one end of a rod and disfavour polymerization at that end. If so, filaments might be able to grow unidirectionally, especially in the cell, where the nature of the guanine nucleotide bound (see below), the state of post-translational modification (see also below), or the extent of association with anchoring proteins or other cellular factors might differ at each of a rod. If the two ends of a rod and/or a filament are somehow different in vivo, septin-containing cellular structures may acquire inherent asymmetry. Indeed, there are clues that such is the case, at least in S. cerevisiae. In budding yeast, a septin-containing ring assembles at the cell cortex and marks the incipient budding site (Longtine and Bi, 2003). During bud formation, this ring expands into a hollow hourglass-shaped collar of septin filaments that lines the mother-bud neck (Byers and Goetsch, 1976a) (Figure 3.4).The mechanism of this expansion and the direction of growth of the septin filaments in the collar are still matters of some controversy, as discussed further below. Fluorescence Recovery After Photobleaching (FRAP) studies of cells expressing GFP-tagged septins support the idea that the septin complexes in the ring are mobile, whereas those in the collar are fixed (Caviston et al., 2003; Dobbelaere et al., 2003). This result is consistent with the idea that, regardless of the length or orientation of the filaments present in the ring, there are free ends from which rods can be lost and onto which rods can be added, but in the collar there are no free ends, as would be expected if the filaments eventually circle around and form closed hoops, as suggested by the available EM images (Figure 3.4). In any event, many proteins bind asymmetrically to one or the other side of this collar (Gladfelter, Pringle and Lew, 2001). There are many possible mechanisms to explain this bipartite behaviour. However, interestingly, it has been reported that at least one protein (Bni4-CFP) is located exclusively on the exterior of the ring even before budding and remains on the mother side of the collar post-budding, whereas at least one other protein (Kcc4-YFP) associates only with the interior aspect of the ring even before budding and is brought to the bud side of the collar during budding (Kozubowski, Larson and Tatchell, 2005). This kind of observation suggests that, both prior to and following filament assembly, there is some sort of polarity imposed on the seemingly non-polar septin filaments. Contrary to the just stated assertion, it has been concluded that at least in terms of the relative arrangement of Cdc3, Cdc11 or Cdc12, the filamentous septin collar is symmetrical (Vrabioiu and Mitchison, 2007). This conclusion was reached as the result of examining the fluorescence anisotropy of GFP-tagged derivatives of each of these three septins in which the GFP was purportedly fused at a fixed rotational angle with respect to the predicted coiled coils of intact Cdc3, Cdc11, or Cdc12 (Vrabioiu and Mitchison, 2007). One hypothesis proposed by Vrabioiu and Mitchison (2007) to explain the apparent discrepancy between this conclusion and the asymmetric distribution observed for many collar-associated proteins, is that during the filament assembly necessary for collar formation (i) the newly synthesized septin complexes are added unidirectionally along the mother-bud axis

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3

5

12

10

28

(a)

10

5

28

(b)

5


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to protofilaments composed of pre-existing complexes and (ii) Bni4 and Kcc4 are somehow able to bind preferentially to septin complexes of different ‘ages.’ However, this proposal seems unlikely because pulse-chase analysis shows that pre-existing and newly synthesized septins are distributed uniformly and symmetrically within the septin collar, at least at the same level of microscopic resolution at which Bni4 and Kcc4 were found to be differentially localized (McMurray and Thorner, 2008). How, then, is the inherent symmetry of the collar filaments ‘broken’ to permit asymmetric localization of septin-associated factors? In yeast, septin accumulation at specific cortical sites depends on the prior arrival of the Rho-type GTPase Cdc42 (Iwase et al., 2006), but assembly into the ring further requires Cdc42-dependent actin filament assembly (Gladfelter et al., 2002; Kozubowski, Larson and Tatchell, 2005; Iwase et al., 2006). Blocking the latter prevents establishment of the observed asymmetry in the localization of Bni4 and Kcc4 (Kozubowski, Larson and Tatchell, 2005). Thus, the ‘polarity’ of septin-containing structures that is responsible for the asymmetric localization of certain septinassociated proteins likely stems from non-septin factors, such as Cdc42-interacting proteins, polymerization of F-actin (which is a filament with inherent polarity) and/or the directed actin-mediated delivery of other molecules to one side or the other of nascent septin structures. According to this idea, this asymmetry would be established early in the maturation process of septin-containing structures (e.g. in non-filamentous septin rings), but would continue during formation of the collar at the bud neck, as long as the asymmetric association(s)/delivery is maintained. In essence, mother-bud asymmetry at the neck may derive from differential interactions of non-septin proteins with ‘anchored’ or ‘free’ ends of inherently non-polar

Figure 3.4 Structure-based interpretation of the arrangement of the septin filaments in the collar at the S. cerevisiae bud neck. In all the EM images and corresponding schematic depictions shown, the mother bud axis runs right-to-left. Arrows in the images in (a) indicate profiles of a single presumptive filament followed through serial sections; arrows in the right image in (b) indicate regions where the presumed filament density is resolved as two fine lines 5 nm apart. The accompanying schematic illustrations are not necessarily to scale. (a) Top left, a negatively stained thin-section of a sagittal section thorough the mother-bud neck of a mitotic S. cerevisiae cell. Bottom left, a schematic representation of the EM shown above. Top right, higher magnification EM image of the region boxed in the bottom left diagram. Bottom right, schematic representation of the EM image shown above. (b) Top left, a negatively stained thin-section of a grazing section of the mother-bud neck just tangential to the plasma membrane. Bottom left, schematic representation of the EM image shown above. Top right, higher magnification image of the central portion of the image shown in top left of (b). Bottom right, schematic representation of the EM image shown above. EM labels: ‘cw’, cell wall; ‘m’, mitochondrion; ‘f’, 10-nm filament; ‘mc’, membrane connection; ‘lc’, lateral connections. Symbols in schematic diagrams: Septin subunits or their G domains, circles or ovals; CTEs wavy lines; numbers, dimensions (in nm). All EM images taken from Byers and Goetsch (1976). Reproduced with permission of Rockefeller University Press, from The Journal of Cell Biology, 69, 717–21

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septin octamers that polymerize unidirectionally via the mechanisms proposed above.

Orientation and organization of the predicted coiled coil–forming domains Homotypic Cdc12-Cdc12 interaction is reproducibly detected using either the two-hybrid method (Lee et al., 2002; Farkasovsky et al., 2005) or in vitro binding of differentially tagged Cdc12 derivatives (Versele et al., 2004), and is eliminated or greatly diminished when its entire CTE (or the predicted coiled coil–forming region at the distal end of the CTE) is deleted. By contrast, deleting the C-terminal 58 residues of Cdc12, which includes the 40-residue segment with predicted coiled coil–forming capacity (Figure 3.2 and Plate Figure 3.3c), does not prevent interaction of Cdc12 with Cdc3, as judged by the two-hybrid method (Farkasovsky et al., 2005), suggesting that a different region of the CTE of Cdc12 must be responsible for its ability to associate with Cdc3. Indeed, deleting the entire CTE (C-terminal 68 residues of Cdc12), thereby removing a segment (‘α7’) (Figure 3.2 and Plate Figure 3.3c) that is strongly predicted to be α-helical, totally ablates the ability of Cdc12 to interact with Cdc3 (Versele et al., 2004). Moreover, when this exact portion (residues 339–407) of Cdc12 was fused to GST, the resulting non-septin chimera is capable of associating specifically with Cdc3 (Versele et al., 2004). These findings highlight, first, an important distinction between the entire CTE and its more distal portion, which contains the segment with potential coiled coil–forming propensity. Second, these findings also highlight that, at least in certain septins, regions in the CTE other than the α6 element (which was revealed by the crystal structure (Plate Figure 3.3a), is shared by all septins (Plate Figure 3.3c) and involved in the NC dimer interface (Plate Figure 3.3a)) can strongly influence specific septin–septin association. It is unfortunate, therefore, that notably absent from the crystallographic structure of the human septin rod (Sirajuddin et al., 2007) is any consistent electron density beyond α6 that would correspond to any part of the CTEs of these septins. Likewise, aside from the work of Hsu et al. (1998) on a septin complex isolated from rat brain, EM images of other septin rods and filaments from budding yeast (Frazier et al., 1998; Bertin et al., 2008), nematode (John et al., 2007) and fruit fly (Field et al., 1996) do not display any orthogonal strands visible by negative staining. Failure to visualize the CTEs is not due to their proteolytic loss because (i) the input full-length septins were recovered when crystals of the human septin complex were redissolved (Sirajuddin et al., 2007), (ii) full-length septins are the sole species present in the yeast preparations visualized by EM (Bertin et al., 2008) and (iii) the C-terminal GFP tag in the UNC-61-GFP fusion was detectable when a worm septin complex containing it was examined in the EM (John et al., 2007). However, as should be recalled, in the latter analysis, the density corresponding to GFP was at a number of positions distributed in an apparent arc


73

∼10 nm from the globular domain of UNC-61. This suggests that pairing of the CTE of UNC-61 and the CTE of UNC-59 to form the parallel coiled coil occurs, but that this element is sufficiently flexible that it does not assume any fixed position relative to the rod. Hence, the locations of these projections are impossible to discern by either EM or crystallography. Indeed, disorder in the CTE was clearly a source of difficulty in obtaining crystals of SEPT2 that diffracted well; the structure of the isolated SEPT2-SEPT2 homodimer was only solved when its C-terminal 46 residues were deleted (Sirajuddin et al., 2007). Nevertheless, in the crystal structure of the human septin hexamer (Sirajuddin et al., 2007), the α6 helices are well ordered and clearly point in a direction perpendicular to the long axis of the rod (Plate Figure 3.3a). Assuming the path of a CTE beyond that point follows in essentially the same direction, for each septin pair at an NC interface, their CTEs and C-terminal coiled coil–forming segments will project from the same face of the rod and be closely juxtaposed, permitting their interaction. As long as the orthogonal ‘strands’ observed in the EM images of Hsu et al. (1998) were not due to another rat brain protein that associates with the immunopurified septin complex, it seems that, compared to the rods of other species, something about these preparations and/or how they were spread, dehydrated and stained for viewing in the EM resulted in immobilization and accentuation of the CTEs so they could be visualized, even in raw (unaveraged) images. If so, these ∼10-nm-long projections confirm that the CTEs of septins bound in the NC mode do pair with each other. The fact that these appear at 16-nm intervals (i.e. four 4-nm subunits apart) suggests that, at least for the constellation of different septin monomers of which the rat brain complex was composed, the CTEs only pair up (or are only visible) for every other NC pair. Although the crystal structure of the human septin hexamer revealed nothing about the structure of the CTEs beyond α6, it was a surprise that stoichiometric complexes could still be prepared by the same expression and purification method using derivatives of all three septins lacking their CTEs (namely, SEPT2(309-361), SEPT6(311-434) and SEPT7(306–437)) and that the hexameric rods so formed did not look significantly different in the EM from the rods prepared with the full-length version of each protein (Sirajuddin et al., 2007). Thus, for these human septins, the additional contacts provided by CTE–CTE interaction are dispensable for complex formation. Likewise, budding yeast Cdc10 lacks any CTE (Plate Figure 3.3c), and a Cdc10-Cdc10 dimer in the NC orientation presumably occupies the centre of the budding yeast rod (Figure 3.1c and Plate Figure 3.3b). The Cdc3-Cdc12 association in the budding yeast rod is also predicted to be a NC interface (Plate Figure 3.3b); but, removal of the CTE from either protein ablates their interaction and eliminates hetero-octamer formation both in vivo and in vitro (Versele et al., 2004). Thus, at least in budding yeast, and at least for this pair, contacts mediated by the corresponding CTEs are essential for this NC interaction, as mentioned above. The importance of the CTEs in these two septins in vivo is further emphasized by the fact that CTE-less alleles of either Cdc3 or Cdc12 act in a dominant-negative fashion to disrupt assembly

74


and function of the filaments at the septin collar when each is expressed in an otherwise wild-type cell (Versele et al., 2004). How could the CTEs be so important for yeast septin complex formation, when the CTEs of the human septins are dispensable? In this and presumably other cases, the devil is likely in the details. Synthetic peptides corresponding to the heptad repeat segments of the nematode septins that are predicted to have coiled coil–forming capability do associate to form a stable, parallel coiled coil in vitro (John et al., 2007). Using a similar approach, the corresponding regions of Cdc3 and Cdc12 showed no evidence of any stable interaction (Versele et al., 2004). Indeed, as already mentioned above, the preceding helical regions (‘α7’) (Figure 3.2 and Plate Figure 3.3c) in the CTEs of both Cdc3 and Cdc12 must also be present for the interaction of these two septins to occur. Thus, this short extra helix is a key component of the NC interaction surface between Cdc3 and Cdc12. Aside from those septin–septin interactions, like that of Cdc3 with Cdc12, where the CTEs clearly provide contacts needed for forming the NC interface, what else, if anything, might CTE–CTE pairing contribute for septin rod and filament ultrastructure? Based on the orientations of the α6 helices in the human heterohexamer (Sirajuddin et al., 2007), there are, theoretically, three CTE–CTE pairs that could project off the same face of a rod (Plate Figure 3.3a). These elements could themselves act as an interaction surface. Such ‘molecular Velcro’ provides an obvious mechanism for recruitment of other non-septin proteins; in this manner, septin filaments could act as scaffolds to bind and organize other structural and/or enzymic functions at the specific subcellular locations where septin complexes and filaments assemble. In animal cells, the Borg family of small Cdc42-binding adapter proteins binds to the CTE–CTE element formed by SEPT6-SEPT7 in the NC orientation (Joberty et al., 2001; Sheffield et al., 2003). Also, it should be recalled that, regardless of the species, when rods polymerize, the filaments so formed have a strong tendency to form parallel ‘railroad tracks’ (Figure 3.1a, middle and bottom). Clearly, if the CTE–CTE interaction forms a parallel heterodimeric coiled coil, then the ‘ties’ in these ‘railroad tracks’ presumably represent interaction between the heterodimeric coiled coil from one filament with the heterodimeric coiled coil from the other filament, that is, formation of an anti-parallel heterotetrameric coiled coil (a four-helix bundle). Four characteristics of the filament pairs seen in the EM are consistent with the notion that the CTE–CTE elements serve as cross-bridges. First, filaments tend to align almost invariably in pairs (Figure 3.1a, middle), implying that the relevant lateral interactions occur between interfaces on only one side of each filament. Second, paired filaments are in register (Figure 3.1a, bottom), as expected if pairing only occurs via interaction at specific points along the filament. Third, the separation between filaments is reasonably uniform over rather long distances and this spacing is of sufficient width to be mediated by contact between CTE–CTE pairs if they extend orthogonally off each filament. Finally, at higher magnification, density spanning


75

the gap between two paired filament with the appropriate periodicity can sometimes be discerned in negatively stained EM images (Bertin et al., 2008). On the other hand, when separated from the context of their attachment to their cognate G domain and the rest of their CTE, the coiled coil segments of the worm and mammalian septins tested showed no evidence of anti-parallel tetramerization of the parallel heterodimeric coiled coils (Low and Macara, 2006; John et al., 2007). Moreover, the heptad repeats in the corresponding sequences are not of the 3-3-1 type that is typically diagnostic of those able to form four-helix coiled coil bundles (Deng et al., 2006). Thus, the disposition of the CTEs in septin rods and filaments remains ill-defined. The yeast septin octamer, despite having two additional subunits, has fewer possible CTE–CTE pairings than the mammalian and insect hexamers (compare Plate Figure 3.3b with 3.3a) because Cdc10 lacks a CTE, and the CTE of a terminal Cdc11 subunit is presumably unpaired unless and until it associates with its sister on another rod. Thus, the CTE of Cdc11 would seem well-positioned to play an important role in mediating rod polymerization via Cdc11-Cdc11 interaction (and via interaction of Cdc11-capped rods with Shs1-capped rods, given that a CTE-dependent interaction between Cdc11 and Shs1 has been observed between the purified proteins (Versele et al., 2004)). However, yeast cells expressing a CTE-less derivative of Cdc11 as the sole source of this septin are viable, whereas a cdc11 null mutation causes a much more severe phenotype (Casamayor and Snyder, 2003) or is outright lethal (Versele et al., 2004), depending on the strain background. On the other hand, based on arguments presented above (and earlier), these effects could reflect the degree to which Shs1 has the capacity in vivo to fully substitute for Cdc11, in a given strain background. Nonetheless, salt-stable octamers can be prepared when CTE-less Cdc11 is co-expressed with Cdc3, Cdc10 and Cdc12, and these are capable of forming filaments in vitro when the salt concentration is lowered (Bertin et al., 2008). Interestingly, whereas Cdc11, like Cdc3 and Cdc12, possesses in its CTE a stretch of residues preceding its presumptive coiled coil–forming segment that are predicted to have α-helix-forming propensity (Figure 3.2 and Plate Figure 3.3c), Shs1 seems to lack this feature. Instead, Shs1 has a predicted β-strand at this location and, moreover, has a CTE that is, overall, nearly 100 residues longer than that of the mitotic septin with the next longest CTE (Cdc11). Because it lacks the α7 helix, Shs1-Shs1 (or Shs1-Cdc11) association in the NC orientation may require interactions that are unique to its apparently highly specialized CTE. Important evidence for a functional role of the coiled coil–forming region in a CTE in helping to stabilize rod and filament structure in vivo comes from the properties of a particular allele (cdc12-6ts ) in the CDC12 gene. Upon shift to the non-permissive temperature, septin-containing structures in cells carrying the cdc12-6 allele rapidly disassemble, accompanied by complete delocalization of all mitotic septins (Ford and Pringle, 1991; Kim, Haarer and Pringle, 1991). These effects are not due to destruction of the mutant protein at the restrictive temperature because the steady-state level of Cdc12-6 is equivalent to that of wild-type

76


Cdc12 under the same conditions (McMurray and Thorner, unpublished results). By contrast, in cells carrying another temperature-sensitive allele, cdc12-1 , further assembly of new structures does not occur, but preformed septin structures do not unravel, after shift to the non-permissive temperature (Dobbelaere et al., 2003). Revealingly, the cdc12-6 allele arises from a frameshift mutation that results in the last 16 residues at the C terminus of Cdc12 being replaced with twenty residues of abnormal sequence (Longtine et al., 2000). This alteration is located far from α7 and only affects the distal half or so of the predicted coiled coil–forming segment in the Cdc12 CTE, but, fortuitously, disrupts only partially the 4-3 heptad repeat in this region (Figure 3.2). It is very telling that such a seemingly subtle change at the extreme C-terminal end of the protein has such a profound affect on the stability of preformed septin complexes in the cell and, thus, supports the conclusion that the coiled coil between the CTEs of Cdc12 and Cdc3, although not essential for their interaction, nonetheless contributes greatly to the stability of this interface once formed. The mutational alteration in the cdc12-1 allele, by comparison, is a G227E substitution (corresponding to residue 241 in human SEPT2), which should affect contacts important for interaction of the G domain of Cdc12 with the terminal rod subunit, Cdc11, thereby explaining its phenotype.

Guanine nucleotide binding and hydrolysis in septin complex assembly The crystal structure of the human septin heterohexamer was of sufficient resolution to discern that GDP is bound to each SEPT2 monomer in the central SEPT2-SEPT2 N-C dimer, whereas the next subunit associated with SEPT2 via a G domain-G domain interaction, namely SEPT6, has GTP bound and the terminal subunit that interacts with SEPT6 in NC orientation, namely SEPT7, has GDP bound (Plate Figure 3.3a). The observed polymerization of hexamers into filaments is undoubtedly mediated by end-on-end, that is, SEPT7-SEPT7, interaction. Thus, at both homotypic septin–septin junctions (SEPT2-SEPT2 and SEPT7-SEPT7), GDP is the nucleotide present in both partners; but, at the heterotypic septin–septin junction (SEPT2-SEPT6), GTP is bound to one partner. These observations and others summarized below suggest that the nature of the nucleotide bound plays a role in both multi-septin complex formation and filament assembly. When expressed individually in bacterial cells, many purified septins are insoluble, aggregated and nucleotide-free (Versele and Thorner, 2004; Hu et al., 2006; Garcia et al., 2007). By contrast, when purified from their native source or when co-expressed with partner septins known to be components of the same physiological complex, the resulting septins are typically soluble and saturated with bound nucleotide (Farkasovsky et al., 2005; Sirajuddin et al., 2007). At the very least, these and other biochemical findings indicate that the presence of a bound guanine nucleotide stabilizes the folded state of every septin examined to date. However, some of the available biochemical data are conflicting and/or subject to multiple interpretations.


77

In agreement with the crystal structure, native septin complexes contain protein and nucleotide in equimolar amounts and a ratio of GDP-to-GTP of ∼2 : 1 (Field et al., 1996; Vrabioiu et al., 2004). One interpretation of these findings is that SEPT2 and SEPT7 bind GDP preferentially, whereas SEPT6 is selective for GTP. An alternative possibility is that all bind GTP, but SEPT2 and SEPT7 can hydrolyse the bound GTP to GDP, whereas SEPT6 is unable to do so. In this regard, it has been observed that both bacterially expressed Cdc10 (which occupies a position in the yeast hetero-octamer equivalent to that occupied by SEPT2 in the human heterohexamer) and bacterially expressed Cdc12 (which occupies a position in the yeast hetero-octamer akin to that occupied by SEPT7 in the human heterohexamer) are able to bind exogenously provided GTP and hydrolyse it (albeit at a slow rate) to GDP (Versele and Thorner, 2004; Farkasovsky et al., 2005). By contrast, when purified in the same way, neither Cdc11 nor Cdc3 (which occupies a position in the yeast hetero-octamer akin to that occupied by SEPT6 in the human heterohexamer) was able to bind exogenously supplied GTP (Versele and Thorner, 2004; Farkasovsky et al., 2005), presumably because each already contained a non-exchangeable guanine nucleotide bound to their G domains. Moreover, mutants of Cdc10 and Cdc12 unable to bind nucleotide still assemble, albeit much less efficiently (Bertin et al., 2008), into complexes with wild-type Cdc3 and Cdc11 and, in such complexes, GTPase activity is almost totally eliminated (Versele and Thorner, 2004), suggesting that, even though GTP may bind to all four of the core yeast septins initially, Cdc10 and Cdc12 are the only two septins in the yeast rod capable of exchanging and subsequently hydrolysing GTP to GDP. In this same regard, the interaction between SEPT2(GDP) and SEPT6(GTP) in the human heterohexamer is clearly more salt-resistant than the SEPT7(GDP)SEPT7(GDP) interaction required for the end-on-end polymerization of rods into filaments. The SEPT2-SEPT6 interaction occurs via a G domain-G domain interface (Plate Figure 3.3a). The parts of a GTP-binding protein that undergo the greatest conformational changes upon nucleotide binding, and are the most different depending on whether the nucleotide is GDP or GTP, are located in the G domain and are dubbed its ‘switch regions’ (Vetter and Wittinghofer, 2001). It was noted in the crystal structure that the switch regions at the SEPT2-SEPT6 interface are better defined than elsewhere in the human rod (Sirajuddin et al., 2007). Taken together, all of these data argue that the state of the bound nucleotide at a given interface could play a role in stabilizing certain subunit–subunit combinations, and also raise the idea that GTP hydrolysis by certain septins and not others is an important step in ordered complex assembly. There is at least some additional experimental evidence in support of these notions, although each is fraught with a potential caveat(s). Formation of a heterodimer from independently expressed and purified SEPT6 and SEPT7 monomers is associated with both binding and hydrolysis of GTP (Sheffield et al., 2003). These findings are at least compatible with the idea that GTP binding and hydrolysis by at least one of these two septins induces the conformational state that

78


is competent for their interaction, even though isolated SEPT6 and SEPT7 presumably also interact, as they do in the hexamer, via a NC-interface (not via their G domains) (Low and Macara, 2006; Sirajuddin et al., 2007). Somewhat similarly, when expressed in and purified from bacteria, Xenopus laevis Sept2 is soluble and nucleotide-free; however, when provided with GTP (or a non-hydrolysable GTP analog), it polymerizes into filaments (Mendoza, Hyman and Glotzer, 2002). However, this filament assembly was preceded by a significant lag phase. Hence, even though the frog Sept2 prepared by bacterial expression was soluble, it could be argued, as others have (Mitchison and Field, 2002), that it was partially unfolded and that GTP merely promotes its stable folding. In other words, this same argument goes, the nature of the guanine nucleotide doesn’t matter for Sept2-Sept2 interaction and filament formation, but only that Sept2 be in its native conformation. Much evidence does suggest that continuous cycles of GTP binding, hydrolysis, product release and rebinding of fresh GTP by the septins themselves are not involved in either complex formation or filament assembly (or disassembly). For example, using liquid chromatography and tandem mass spectrometry to measure the rate of dilution of the 15 N label in septin-associated guanine nucleotides in vivo, it was concluded that the majority of the septin-bound guanine nucleotides do not turn over during the period of a one yeast cell cycle. On the other hand, rapid exchange into one or two of the subunits within an octamer, or into a small population of the rods in the total cellular pool, could have been missed by this kind of bulk measurement (Vrabioiu et al., 2004). In fact, in these in vivo studies guanine nucleotide exchange was only detected in Cdc10 and Cdc12 (Vrabioiu et al., 2004), in agreement with the aforementioned in vitro studies demonstrating that only Cdc10 and Cdc12 show appreciable GTP binding and hydrolysis (Versele and Thorner, 2004; Farkasovsky et al., 2005). The conclusion that neither hydrolysis nor exchange of bound nucleotide is likely to play a regulatory role in the dynamic changes in septin organization that occur during the cell cycle is also consistent with the very slow rate of GTP hydrolysis displayed by isolated Cdc10 and Cdc12 (or the hetero-octameric complexes that contain them) (Versele and Thorner, 2004; Farkasovsky et al., 2005). Nonetheless, such findings certainly do not rule out, as emphasized earlier in this section, that a one-time GTP-binding event is an important step in ensuring that septins are fully folded, in the correct conformational state for properly ordered septin complex assembly, and competent for rod–rod polymerization. It has been difficult, however, to design experimental strategies to nail down this latter point unequivocally. Unlike isolated native complexes, individual septins or septin sub-complexes expressed in and purified from heterologous systems contain variable amounts of guanine nucleotide (and different ratios of GDP-to-GTP), which do not correlate well with the expression system used or the septins examined. For example, ∼60 % of human SEPT2 molecules produced in bacteria contain bound nucleotide, with GDP-to-GTP in a ∼1 : 1 ratio (Huang et al., 2006), whereas, it should be recalled, bacterially expressed frog Sept2 is nucleotide-free


79

(Mendoza, Hyman and Glotzer, 2002)]. Moreover, when expressed in and purified from insect cells, essentially all the human SEPT2 molecules contain bound nucleotide, with GDP-to-GTP in a ∼10 : 1 ratio (Huang et al., 2006). Likewise, yeast septins expressed in E. coli can display different nucleotide- and septin-binding behaviours when expressed individually versus when they are co-expressed with their partner septins (Versele et al., 2004; Versele and Thorner, 2004; Farkasovsky et al., 2005). Also, as already noted above, measuring the ability of a septin or septin complex to bind and hydrolyse guanine nucleotide requires exchange of any bound nucleotide with the exogenously added labelled nucleotide. Thus, only purified septins that are either nucleotide-free, or have an appreciable rate of nucleotide exchange, can be analysed. The same is true for any given subunit within a complex. Conversely, the subunits that bind nucleotide the tightest are most likely to be already saturated with it, exchange it slowly, and thus be much harder to study with regard to their nucleotide binding and hydrolysis properties. With these considerations in mind, it seems plausible that, at least for the yeast hetero-octamer, Cdc10 and Cdc12 are the most reasonable candidates for subunits in which conversion of bound GTP to GDP may be involved in rod assembly. However, single amino acid substitutions in Cdc10 and Cdc12 that do not prevent GTP binding, but do drastically compromise hydrolysis of the bound GTP in vitro, have absolutely no phenotype when expressed as the sole source of either or both of these septins in vivo (Versele and Thorner, 2004). Nevertheless, when modelled on the state of the bound guanine nucleotides in the subunits of the human hexamer, and considering the >2 : 1 GDP : GTP ratio reportedly found in purified yeast octamers (Farkasovsky et al., 2005), it seems likely that, in a yeast rod, only Cdc3 is present in the GTP-bound state and the other three are GDP bound (Plate Figure 3.3b). Thus, even if Cdc10, Cdc11 and Cdc12 bind and hydrolyse GTP, once incorporated into the rod, these subunits do not readily release the resulting bound GDP. Consistent with the need for a bound nucleotide for optimal septin complex assembly and rod stability, single amino acid substitutions that prevent nucleotide binding to Cdc10 and Cdc12 (in contrast to the mutations that prevent hydrolysis of GTP once bound), interfere with efficient hetero-octamer formation when these Cdc10 and Cdc12 mutants are co-expressed with wild-type Cdc3 and Cdc11 in bacterial cells (Bertin et al., 2008) and show a readily detectable and synergistic temperature-sensitive phenotype when expressed in vivo as the sole source of Cdc10 and Cdc12. Specifically, a double mutant of two GTP binding-defective alleles, cdc10(S46N) cdc12(T48N), grows slowly and has drastically elongated buds at 30◦ and is inviable at 37◦ , whereas a double mutant of two GTP-binding competent, but GTP hydrolysis-defective alleles, cdc10(S41V) cdc12(S43V), grows well and shows no abnormality at either temperature (Versele and Thorner, 2004). Moreover, the recombinant doubly mutant rods that can be isolated, that is, Cdc11-Cdc12(T48N)-Cdc3-Cdc10(S46N)-Cdc10(S46N)-Cdc3-Cdc12(T48N)Cdc11 are much less stable than wild-type rods in solution and incapable of polymerizing into filaments (Versele et al., 2004; Bertin et al., 2008). Thus,

80


optimal rod assembly and rod stability require guanine nucleotide binding to at least Cdc10 and Cdc12. However, the opposite conclusion was reached for mutations that putatively affect nucleotide binding by Cdc11 (Casamayor and Snyder, 2003); cdc11 alleles designed to incapacitate nucleotide binding had little discernible functional consequence in vivo (Casamayor and Snyder, 2003). However, it could not be convincingly verified experimentally that the mutations made had the anticipated effects on GTP binding. When modelled on the crystal structure of the human hexamer, these mutational alterations are predicted to perturb the G-domain interface of Cdc11, which would prevent its association with Cdc12. Cdc11-less rods are unable to polymerize into filaments (Versele et al., 2004; Bertin et al., 2008), probably explaining the phenotype of these mutants in vivo. The G domains of Cdc10 and Cdc12 do not participate directly at the interface required for end-on-end polymerization of rods. Thus, the fact that the Cdc11-Cdc12(T48N)-Cdc3-Cdc10(S46N)-Cdc10(S46N)-Cdc3-Cdc12(T48N)Cdc11 double mutant complex fails to polymerize suggests that guanine nucleotide-dependent conformational changes in Cdc10 and Cdc12 are propagated outward and influence the terminal subunits of the octamer. This does not seem far-fetched, at least for Cdc12, because this subunit directly abuts Cdc11 at each end of the rod. In any event, to address these same issues more incisively in the near future, it should be possible to pinpoint, based on the now available crystal structure, residues that when altered will compromise GTP binding (or hydrolysis) without unduly affecting contacts crucial for the G-domain interface.

Post-translational modifications of septins and identification of other important residues For the reasons discussed in the preceding section, septins are unlikely to behave as ‘molecular oscillators’ driven by repeated cycles of nucleotide binding and hydrolysis. Yet, septin-based structures clearly undergo a series of marked structural rearrangements during progression though each cell cycle that need to be properly regulated both spatially and temporally. What processes control these dynamic transitions? Post-translational modifications of the septins are good candidates to regulate the ultrastructure and biochemical properties of rods and filaments in signal-dependent manner. Indeed, septins are heavily modified, especially by both phosphorylation and SUMOylation (Table 3.1), and the locations of these modifications modelled on the crystal structure of the human hexamer are potentially revealing about their conceivable effects. Some of these positions are indicated in Plate Figure 3.3a, along with the locations of other residues implicated in septin function for other reasons (e.g. a cellular phenotype results from mutation of that residue). A few selected examples are discussed below. Phosphorylation of Cdc10 on Ser256 by the Cdc42-regulated protein kinase Cla4 is thought to promote filament assembly (Kadota et al., 2004; Versele and

α1, near NC interfaceb N/A Exposed disordered loop between α1 and β2

SEPT2 K30

SEPT2 S31

SEPT2 G44, G47, G49, K50

SEPT2 S51

SEPT2 N55

HsSEPT2 V/I67d

ScCdc11 R12, K13, R14, K15, H16

ScCdc3 S113

ScCdc11 G29, G32, G34, R35

ScCdc10 S46, ScCdc12 T48

ScCdc11 N40

RnSEPT3 S91

P-loop, near G interfaceb

P-loop, G interface

α0, NC interfaceb

Phosphorylation (cGMP-dependent protein kinase-I

N/A

Phosphorylation (PAK family kinase Cla4) N/A

Extreme N terminus Phosphorylation (disordered)b (unknown) N/Ac Extreme N terminus (‘polybasic motif’)b

SEPT2 Y17

HsSEPT2 Y17

Modification (modifier)

Equivalent/ Location representative residue(s) in human septin of solved structure

Residue(s)a

Subcellular localization? (phospho-formspecific localization)

(continued overleaf )

(Casamayor and Snyder, 2003) (Xue et al., 2004)

(Versele and Thorner, 2004; Farkasovsky et al., 2005)

(Zhang et al., 1999; Casamayor and Snyder, 2003) (Versele and Thorner, 2004) (Casamayor and Snyder, 2003)

Membrane interaction (phospholipid binding in vitro) Unknown Nucleotide binding/hydrolysis (behaviour of purified mutant) Nucleotide binding, filament assembly (behaviour of purified mutant, cellular phenotype) Unknown

(Rush et al., 2005)

Reference(s)

Unknown

Predicted function (evidence)

Table 3.1 Locations of residues in septin proteins known or predicted to be post-translationally modified, and those with effects on septin structure or function when mutated

SEPTIN COMPLEXES IN VERTEBRATES 81

SEPT2 H140

SEPT2 F156

SEPT2 A182

SEPT2 D185

HsSEPT2 T186, T188

SEPT2 G241

ScCdc3 S244

HsSEPT2 F156

ScCdc10 G179

ScCdc10 D182

ScCdc12 T183, T185; ScShs1 S221

ScCdc11 G230, ScCdc12 G227

N/A

Phosphorylation (Mec1,Tel1, and/or Rad53 for ScShs1 S221) N/A

G interfaceb

Between β5 and α4, G interface

Near G interfaceb

N/A



G interface

Disordered residue near NC interface G interfaceb


(continued )

Residue(s)a

Table 3.1

Hetero-oligomer assembly (chromatographic behaviour of purified mutant) Hetero-oligomer assembly (cellular phenotype)e Hetero-oligomer assembly (cellular phenotype)e DNA damage response for ScShs1 S221 (cellular phenotype) Filament assembly but not stability (cellular phenotype)f

Unknown


(Casamayor and Snyder, 2003; Dobbelaere et al., 2003)

(Smolka et al., 2006; Smolka et al., 2007)

(Hartwell, 1971)

(Cid et al., 1998)

(Versele and Thorner, 2004) (Sirajuddin et al., 2007)

Reference(s)

82 CH03 BIOCHEMICAL PROPERTIES AND SUPRAMOLECULAR ARCHITECTURE

SEPT2 S218

Not conserved, near SEPT2 S218 SEPT2 T228, SEPT7 T228

SEPT2 W260

SEPT2 G261

SEPT2 H270

SEPT7 Y319

SEPT7 S334

HsSEPT2 S218, RnSEPT2 S218, MmSEPT4 S325

ScCdc10 T216

MmSEPT7 T227; RnSEPT2 T228

HsSEPT2 W260

ScCdc10 S256

HsSEPT2 H270

HsSEPT7 Y319

HsSEPT7 S334; MmS333


β6, G interfaceb

Disordered, between Phosphorylation α6 and coiled coil (unknown) Disordered coiled coil Phosphorylation (unknown)

G interfaceb

N/A

G interfaceb

(Hoffert et al., 2006, 103, p. 7159; Dai et al., 2007) (Sirajuddin et al., 2007)

(Beranova-Giorgianni et al., 2006; Hoffert et al., 2006; Huang et al., 2006; Nousiainen et al., 2006; Olsen et al., 2006; Dai et al., 2007; Molina et al., 2007) (Li et al., 2007)

Unknown


(Olsen et al., 2006; Munton et al., 2007)

Hetero-oligomer assembly (chromatographic behaviour of purified mutant) Filament assembly (Versele and Thorner, (cellular phenotype) 2004) Hetero-oligomer (Sirajuddin et al., 2007) assembly (chromatographic behaviour of purified mutant) Unknown (Rush et al., 2005)

Unknown

Unknown

Phosphorylation (casein Nucleotide binding kinase II) (behaviour of purified mutant)

Disordered loop Phosphorylation between α4 and α5’ (unknown) Phosphorylation Exposed α5’, near (unknown) G interfaceb

Disordered loop between α4 and α5’, near NC interface


(continued )

Not conserved

Not conserved

Not conserved

Not conserved

Not conserved

Not conserved

ScCdc3 S503g

ScCdc3 S509g

ScCdc10 S312

ScCdc10 H314

ScCdc10 S316

ScCdc11 S305, S316, S318, S319, S327

Phosphorylation (ATR/ATM?) SUMOylation (SUMO-protein ligase Siz1) Phosphorylation (cyclin-dependent kinase) Phosphorylation (cyclin-dependent kinase) Phosphorylation (PAK family kinase Cla4)

Extreme C terminus, disordered Extreme N terminus

Phosphorylation (unknown) Exposed C terminus Phosphorylation (unknown) Between α6 and Phosphorylation predicted coiled coil (unknown)

Exposed C terminus

Exposed C terminus

Extreme C terminus

Extreme C terminus

Phosphorylation (unknown)


Extreme C terminus, disordered


HsSEPT7 S423, T426; SEPT7 S423, T426 MmSEPT7 T425; RnSEPT7 T425 HsSEPT6 S408, S411, SEPT6 S408, S411, S416, T418; S416, T418 MmSEPT6 S416 ScCdc3 K4, K11, Not conserved K30, K63

Residue(s)a

Table 3.1

(Hoffert et al., 2006; Dai et al., 2007; Molina et al., 2007) (Matsuoka et al., 2007; Molina et al., 2007; Munton et al., 2007) (Johnson and Blobel, 1999)

Reference(s)

Unknown

Unknown

(Chi et al., 2007; Li et al., 2007; Smolka et al., 2007)

(Smolka et al., 2007)

Filament disassem(Tang and Reed, 2002) bly/reorganization (cellular phenotype) Filament disassem(Tang and Reed, 2002) bly/reorganization (cellular phenotype) Unknown (Versele and Thorner, 2004; Smolka et al., 2007) Unknown (Chi et al., 2007)

DNA damage response (role of modifier) Unknown

Unknown



Not conserved

HsSEPT9 S82, S85, S89

Not conserved

Not conserved

Not conserved

HsSEPT9 T42

Not conserved

ScShs1 S519, S525, S539, S545, S548 MmSEPT8 A2; HsSEPT11 A2 MmSEPT8 S10

Not conserved

Extreme N terminus

Not conserved

ScShs1 S408, S416

Not conserved

Extreme N terminus

Not conserved

ScShs1 S447, S460

HsSEPT9 S30; MmSEPT9 S30 RnSEPT9 T31i

Extreme C terminus

Not conserved

ScShs1 K426, K437

Phosphorylation (unknown) Unknown Phosphorylation (unknown) Unknown Phosphorylation (unknown) Unknown Phosphorylation (unknown) Near residues affected Phosphorylation (unknown) in HNj

Extreme C terminus

Extreme C terminus

Extreme C terminus

Extreme C terminus

Not conserved

ScCdc11 K412

SUMOylation (SUMO-protein ligase Siz1) SUMOylation (SUMO-protein ligase Siz1) Phosphorylation (unknown) Phosphorylation (unknown) Phosphorylation (unknown) Acetylation (unknown)

Between α6 and N/A predicted coiled coil

ScCdc12 K351, E368h Not conserved

(Olsen et al., 2006; Molina et al., 2007)

Unknown


(Beausoleil et al., 2006)

(Beausoleil et al., 2006; Dai et al., 2007) (Hoffert et al., 2006)

(Gruhler et al., 2005; Dai et al., 2007) (Dai et al., 2007; Smolka et al., 2007) (Dai et al., 2007; Smolka et al., 2007) (Gevaert et al., 2003; Trinidad et al., 2006) (Trinidad et al., 2006)

(Johnson and Blobel, 1999)

(Johnson and Blobel, 1999)

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Interaction with chaperone/disaggregase Hsp104? (genetic suppression) Unknown


Not conserved

Not conserved

Not conserved

HsSEPT9 R88, S93F

HsSEPT9 T142

RnSEPT9 T150

Phosphorylation (unknown) Phosphorylation (unknown)

N/A


Unknown

Septin–septin interaction (cellular co-localization phenotype)j Unknown


(Hoffert et al., 2006)

(Molina et al., 2007)

(Sudo et al., 2007)

Reference(s)

a Amino acid residues are identified using the standard one-letter abbreviation. The identity of the septin protein in question is given with a two-letter prefix identifying the host species (e.g. ‘Sc’, Saccharomyces cerevisiae). b Indicates the (predicted) location of this residue is shown in Plate Figure 3.3. c N/A, not applicable. d The sequence of HsSEPT2 presented in (Sirajuddin et al., 2007) includes an isoleucine at position 67 rather than the valine predicted by all available sequences in public databases. e Mutation of ScCdc10 at G179 (in the cdc10-11 allele) causes a temperature-sensitive phenotype, and the inability of the mutant to assemble at the bud neck in the presence of wild-type ScCdc10, even at the permissive temperature (Cid et al., 1998). Both phenotypes also apply to the cdc10-1 allele, isolated by Hartwell (1971) and determined by sequencing to harbour the substitution D182N (M. A. McMurray and J. Thorner, unpublished data). f The temperature-sensitive cdc12-1 allele identified by Hartwell (1971) encodes a mutant ScCdc12 with the substitution G227E (Casamayor and Snyder, 2003). When shifted to the non-permissive temperature, yeast cells carrying this mutation cannot assemble new septin structures, but existing structures are not affected (Dobbelaere et al., 2003). Mutation at the equivalent position in ScCdc11 (G230) also renders cells temperature-sensitive (Casamayor and Snyder, 2003). g Residue is conserved in HsSEPT10. h ScCdc12 mutations K351N or E368Q eliminate the adverse effects on septin organization and function of overexpression of Hsp104G17S T499I , an allele of the chaperone/disaggregase harbouring mutations in both the coiled coil–forming middle region, and in the nucleotide-binding domain (Schirmer et al., 2004). i Equivalent to HsSEPT9 T49. j Mutations R88W and S93F in the uniquely extended N terminus of SEPT9 are associated with hereditary neuralgic amyotrophy (HN) (Kuhlenbaumer et al., 2005). The mutant forms appear to interact with certain other septins, whereas the wild type does not (Sudo et al., 2007).

Unknown

Unknown

Extreme N terminus


(continued)

Residue(s)a

Table 3.1



87

Thorner, 2004). Moreover, the effect of this modification is thought to function in conjunction with the effect that nucleotide binding has on Cdc10 and Cdc12 for three reasons (Versele and Thorner, 2004). The phenotype of cdc10(S46N) cdc12(T48N) double mutant cells is ameliorated by overexpression of CLA4 specifically. Conversely, the phenotype of cdc10(S46N) cdc12(T48N) double mutant cells is markedly exacerbated by a cla4 mutation. Finally, a cdc10(S46N S256A) double mutant has a more severe phenotype that either a cdc10(S46N) or a cdc10(S256A) single mutant. Interestingly, Ser256 (equivalent to G261 in human SEPT2; see Table 3.1), while not conserved as a phosphorylatable residue in any of the three human subunits whose structures have been solved, is predicted to be located near the G-domain interface and at a position where phosphorylation could conceivably influence the strength of the G-dimer interaction (Plate Figure 3.3a). Accordingly, the severe consequences on septin function of combining the nucleotide binding mutations with the non-phosphorylatable substitution at S256A probably stem from additive effects that weaken the G-dimer interaction between Cdc10 and Cdc3. Another phosphorylation event that potentially affects a G-dimer interface is Ser221 in Shs1 (equivalent to T186 in human SEPT2; see Table 3.1) and its counterparts (T183 and/or T185) in Cdc12. Phosphorylation of Shs1 on S221 occurs in response to DNA replication stress and appears to play a role in this Rad53 (human ortholog is CHK2)-dependent checkpoint (Smolka et al., 2006). Hence, Shs1 is presumably the target of one of the kinases activated in response to the DNA damage that occurs when DNA replication is compromised (Smolka et al., 2007). Based on its location, it is tempting to speculate that phosphorylation of Shs1 at this position diminishes its ability to interact with Cdc12 and that phosphorylation of Cdc12 at the equivalent position reinforces this effect, perhaps by charge repulsion between the two negatively charged phosphate groups. By extension of this idea, phosphorylation of SEPT2 at T186 may affect its interaction with SEPT7, and might be a consequence of the DNA damage response in human cells, in which septins have recently been shown to function (Kremer, Adang and Macara, 2007). Many similar, intriguing and readily testable predictions can be generated by comparing the locations and functions of specific septin residues between different species, now made possible by our new level of understanding of septin structure and about the organization of septin rods.

Lipophilia and the structural basis for septin–membrane interaction The 10-nm filaments at the yeast bud neck are closely apposed to the plasma membrane (Figure 3.4a, right) (Byers and Goetsch, 1976a). Moreover, it has been noted that, upon fractionation of cell extracts, septins often co-purify with the membrane-containing material under conditions in which it would be expected that interaction with other factor(s) that might mediate septin-membrane interaction would be disrupted (Xie et al., 1999). Indeed, during a typical preparation

88


from animal cells, the soluble septin complexes are retained, whereas those in the membrane fraction are removed. For example, in the preparation of Pnutcontaining septin complexes from D. melanogaster (Field et al., 1996), ∼50 % of the total Pnut was in the particulate fraction and was discarded. It is at least conceivable that the membrane-associated septin complexes could be different in a significant way from those in the soluble fraction. Nonetheless, it is generally accepted that septin complexes and filaments interact directly with membranes. This interaction could influence, and certainly places constraints on, the ultrastructure of septin filaments and may facilitate or preclude interactions between any given septin in a filament and other proteins. Conversely, as has been pointed out by others, such membrane-associated septin ‘fences’ can act as barriers to the diffusion of integral membrane proteins (Takizawa et al., 2000; Dobbelaere and Barral, 2004). At first glance, the curvature of the plasma membrane itself seems an ideal candidate to act as a template figure to direct assembly of otherwise linear septin rods and filaments into ring-shaped structures. However, some purified septin complexes display ring-forming ability even in the test tube (Kinoshita et al., 2002; Farkasovsky et al., 2005). This property of self-closure may be related to those septin rods that possess a ‘kink’ observable by EM, such as seen in purified preparations of the recombinant human SEPT2-SEPT6-SEPT7 hexamer (Sirajuddin et al., 2007). If the kink were always on the same side of each rod in a filament, it would bend in a curved arc even in the absence of membrane structures. Furthermore, septins can form similarly shaped rings in cells that have drastically different membrane curvatures. For example, the septin rings formed at the convex plasma membranes of ovoid S. cerevisiae cells are indistinguishable from those formed at a concave membrane when budding occurs at the isthmus of a dumbbell-shaped zygote (Cid et al., 2001). Indeed, at least in part by virtue of their recruitment of membrane- and wall-remodelling enzymes, septins sculpt the cell envelope, and not vice versa (Gladfelter, Pringle and Lew, 2001; Schmidt et al., 2003; Gladfelter, 2006). How do septins contact the plasma membrane? Eukaryotic membranes contain various phosphoinositides that often display discrete localization patterns and facilitate the assembly of higher-order protein complexes at the membrane via the specific binding of membrane-associated proteins. Direct phospholipid binding has been reported for human SEPT4 (Zhang et al., 1999) and budding yeast Cdc3, Cdc10, Cdc11 and Cdc12 (Casamayor and Snyder, 2003). In the case of SEPT4, binding is specific for phosphatidylinositol 4,5-bisphosphate (PtdIns[4,5]P2 ) (Zhang et al., 1999), which is enriched at sites of septin function (e.g. the cleavage furrow of dividing animal cells) (Saul et al., 2004; Field et al., 2005) and required for normal SEPT4 localization (Zhang et al., 1999). PtdIns[4,5] P2 is also enriched at the yeast bud neck (Garrenton, 2007); C. Stefan and S.D. Emr, personal communication; L.E. Stolz and J.D. York, personal communication) and is required for normal septin localization there (Rodriguez-Escudero et al., 2005). It has been reported that GST-yeast septin fusions expressed in and


89

purified from yeast cells bind with highest affinity to filter-immobilized PtdIns[4]P and PtdIns[5]P, and not at all to PtdIns[4,5]P2 (Casamayor and Snyder, 2003). However, in studies assessing binding to phosphoinositide-containing liposomes, yeast septins individually expressed in and purified from bacteria bind better to vesicles containing PtdIns[4,5]P2 than to those containing PtdIns[4]P (T. Allyn, V. Votin and J. Thorner, unpublished results). Additional studies are clearly needed to clarify the lipid-binding specificities of septin subunits, septin rods and septin filaments. Protein-membrane interaction is often mediated by motifs in the protein that contain multiple basic residues that can interact electrostatically with the negatively charged head groups of phosphoinositides and/or other acidic glycerophospholipids. Most septins have a polybasic motif located just N-terminal to the P-loop of the G domain (Plate Figure 3.3c), and mutations in this motif reportedly disrupt lipid binding in vitro (Zhang et al., 1999; Casamayor and Snyder, 2003) and interfere with normal septin function in vivo (T. Allyn, V. Votin and J. Thorner, unpublished results). However, the crystal structure of the human septin hexamer reveals that the polybasic motif falls squarely within the α0 helix and is substantially buried at each NC dimer interface. Nonetheless, one or two of the basic side chains might be sufficiently exposed so as to contribute to membrane interaction. It should be appreciated that, in the context of the complex, even just one or two such contacts contributed by each subunit would represent 8-16 basic residues all projecting off essentially the same face of a rod (Plate Figure 3.3b). Indeed, this arrangement could provide a zipper-like effect that would lock a rod or filament all along its length to the surface of the membrane, consistent with the EM images showing that the 10-nm neck filaments are closely and continuously apposed to the plasma membrane (Byers and Goetsch, 1976a) (Figure 3.4a, right). Also, it is noteworthy that of the seven septins encoded in the S. cerevisiae genome, the three subunits known (Cdc11) or deduced (Shs1 and Spr28), as described earlier, to be at the ends of a rod have the most highly basic α0 sequences: Cdc11, -RKRKHLKR-; Shs1, -RRKKEHKR-; and, Spr28, -RRRKGYKK- (as compared, for example, with the corresponding segment of Cdc12, -RYKIVNEE-). At the ends of the hetero-octamer (a potential, but unfilled, NC interface), the α0 helix is completely exposed (Plate Figure 3.3b). Therefore, these basic ‘fingers’ at each end of a rod could provide a robust mechanism by which rods are recruited to the membrane as soon as they are assembled or, alternatively, to ensure that assembly of rods (or filaments) is nucleated only in association with membranes.

Byers and Goetsch revisited We feel that, as a fitting conclusion to this chapter, we must return to where we began, with the 10-nm filaments at the bud neck of S. cerevisiae cells (Byers and Goetsch, 1976a; Frazier et al., 1998). How has what we’ve learned more

90


recently about septin ultrastructure and biochemistry improved our ability to interpret those EM images? Remarkably, despite the advances described in this article, a long-standing question in the field has remained how the filaments at the bud neck are aligned. Byers and Goetsch (1976a) interpreted their images as reflecting continuous filaments (∼10 nm in cross-section) (Figure 3.4a) wrapping circumferentially around the bud neck, either as separate bands or as a helix with a wide gyre, spaced ∼30 nm apart (Figure 3.4b). However, others have suggested (Field et al., 1996) that yeast septin filaments actually run in the orthogonal direction, extending from the mother cell into the bud, and that the pattern observed by Byers and Goetsch (1976a) arises artifactually. Specifically, it was proposed that (i) the filaments run parallel to the mother-bud axis, (ii) the filaments contain some regularly repeated structure that occurs at intervals of ∼30 nm, (iii) for some reason, this periodic structure stains more intensely than any other part of the filament and (iv) the filaments are in register, so that the heavily staining blobs on one filament are aligned with the blobs of the filaments to its right and to its left. In this way, it was posited, the more densely stained striations made up of all the blobs would appear like filaments running circumferentially. Some, albeit weak, evidence for this highly speculative alternative model of filament organization at the bud neck comes from the fact that certain mutants (e.g. gin4) cause aberrant septin structures to be formed and these include aggregated filament bundles with their long dimension parallel to the mother-bud axis (Longtine, Fares and Pringle, 1998). More recently, using a strategy designed to tag septins with GFP in a ‘fixed’ orientation so as to take advantage of the fluorescence anisotropy of the chromophore in this fluorescent probe, valiant attempts were made to examine filament orientation in vivo (Vrabioiu and Mitchison, 2006). The GFP molecules in such septin-GFP fusions appeared to be oriented all in the same direction, confirming that in the filaments at the bud neck the septins themselves are uniformly and linearly arrayed (Vrabioiu and Mitchison, 2006). Intriguingly, whatever direction this represents (parallel to the bud neck or orthogonal to it), the orientation of the GFP shifts by 90◦ when the septin collar at the bud neck is split into two distinct rings during cytokinesis (Vrabioiu and Mitchison, 2006). One interpretation of this observation (if one assumes that each septin-GFP chimera is immutably rigid) is that every septin (or rod or filament) rotates by 90◦ during this structural transition, a feat of molecular gymnastics truly astounding to envision. Of course, some other change in the septins (or rods or filaments), like association with another protein or some post-translational modification, could cause the GFP to rotate its position by 90◦ relative to the septin to which it is attached. In any event, to answer the original question – the orientation of filaments in the collar in relation to the mother-bud axis – the fluorescence polarization of the septin-GFP probe at the bud neck must be compared with a septin-GFP-containing structure of known orientation. When bundles of filaments formed in vitro by purified yeast complexes are used (visualizing the polarized fluorescence of an individual filament requires a level of sensitivity that cannot yet be achieved for


91

obvious technical reasons), the results obtained were interpreted to indicate that the septin filaments in the filamentous collar run parallel to the mother-bud axis (Vrabioiu and Mitchison, 2006). The subsequent 90◦ shift observed would then mean that, at cytokinesis, the filaments rotate to become perpendicular to the mother-bud axis. However, there are many important caveats to the use of this approach. One is that the preparation used as the known standard for orientation was a bundle of septin aggregates. Typically, the formation of such aggregates reflects some degree of denaturation or unfolding of the constituent septin(s). Hence, the analysis may not reflect the true situation when septins are in their native state. Furthermore, another defining feature of such septin bundles is a greatly reduced spacing between the filaments within it (Bertin et al., 2008). If the normal spacing between unbundled filaments pairs (15–25 nm) reflects cross-filament bridges composed of the CTE–CTE elements projecting orthogonally off each filament, as we have argued, then it is conceivable that the collapse of the spacing observed when filaments bundle is accompanied by a ∼90◦ rotation, so that the CTE–CTE elements that were formerly perpendicular to the filament axis are now approximately parallel to it. Thus, the standard used as the ‘known’ may be the exact opposite of what was assumed. In any event, the rotation observed in vivo during cytokinesis might reflect a difference in the orientation of the CTE–CTE elements rather than any change in the filaments themselves. Another potential concern is that GFP was fused to the C terminus of either Cdc3 or Cdc12, with the assumption that the orientation of the GFP relative to the septin would be fixed by virtue of the putative α-helical nature of the presumptive Cdc3-Cdc12 coiled coil and the known α-helical structure of the N-terminal segment of GFP (Vrabioiu and Mitchison, 2006). Accordingly, the crux of the interpretation of the polarized fluorescence experiments is that the orientation of the coiled coils in the collar at the bud neck remains rigidly fixed and the same as in the standard (the bundles of filaments formed in vitro) . However, as described earlier, both EM and crystal structure analyses clearly demonstrate that, at least in vitro, the coiled coils can be ‘floppy’, occupying a variety of positions relative to the filament axis of nearly 180◦ (John et al., 2007). Consequently, the jury is still out on the issue of the orientation of the septin filaments in the collar relative to the mother-bud axis. There are many reasons to be reassured that the images of the neck filaments observed by Byers and Goetsch (1976a) actually do represent continuous circumferential hoops (or the gyres of a continuous long helix filament) (Byers and Goetsch, 1976a). Based on what we now know about the ultrastructure and atomic level structure of the rods, which constitute the fundamental building block of filaments, the molecular significance of the dimensions observed and described by Byers and Goetsch (1976a) fall neatly into place, but only if the septin filaments in the collar are continuous and circumferential around the bud neck. First, the 10-nm diameter of neck filaments is approximately double the width of the yeast octameric rod (∼4–5 nm) as measured by high-resolution EM (Sirajuddin et al., 2007; Bertin et al., 2008) or

92


of the human hexameric rod (4–5 nm) from either high-resolution EM or X-ray crystallography (Sirajuddin et al., 2007). This suggests that the neck filaments could be pairs of filaments, adjoined via contacts between two filaments mediated by the face of the G domains opposite to that from which the α6 helix and CTEs project (Figure 3.4b and Plate Figure 3.3b). Indeed, in bud neck sections in which the filaments can been viewed longitudinally, the 10-nm thickness can occasionally be resolved as two fine lines ∼5 nm apart (Byers and Goetsch, 1976a) (Figure 3.4b, right). These fine lines are likely the ‘spines’ of each of the individual ∼5-nm-wide filaments, visible only near the edges of the cylindrical bud neck because elsewhere the sections slice through the thicker filament cores (Figure 3.4b, left). G-domain-mediated pairing of filaments might even explain the otherwise incongruous homotypic interactions observed when differentially tagged versions of the same yeast septin were tested for the capacity to self-associate in vitro (Versele et al., 2004) or by the two-hybrid method (Farkasovsky et al., 2005). The 28-nm interval between filaments (presumably measured from centre to centre) translates into 18 nm of physical separation (given a 5-nm thickness of each filament within the pair). This distance is quite similar to the spacing between the paired filaments formed in the test tube (e.g. 15–25 nm (Frazier et al., 1998; Bertin et al., 2008). The most parsimonious interpretation of this spacing is that it represents the separation imposed by extension of the CTEs from one member of each of two opposed filament pairs (Figure 3.4a (right) and 3.4b (right)), assuming that the CTE–CTE elements (calculated to be ∼10 nm if α-helical and fully extended) only overlap/conjoin via contacts within their very distal coiled coil–forming segments (Figure 3.2). The coiled coils (and/or proteins associated with them in vivo) would, according to this model, represent the 3-nm-thick lateral extensions interconnecting adjacent filaments seen by Byers and Goetsch (1976a) (Figure 3.4a, right). A final consideration is the relative position of the filaments with respect to the plasma membrane, from which they appear slightly separated by density that Byers and Goetsch dubbed the ‘connecting zone’ (Byers and Goetsch, 1976a). It does not seem likely that residues from the polybasic motif could protrude away from the G domain far enough to explain this gap (Figure 3.4a, right). It should be recalled, however, that the N terminus of Cdc3 is 80-100 residues larger than those of the other three core yeast septins. Thus, one possibility is that the N terminus of Cdc3 is specialized to ensure membrane contact and occupies the connecting zone. Alternatively, the connecting zone could be occupied by any of the numerous non-septin proteins at the bud neck that interact both with septins and the plasma membrane.

FUTURE DIRECTIONS Although only a small fraction of the last 30 years of research on septins has focused on their ultrastructure and biochemistry, the recent major accomplishments summarized and analysed in this chapter reveal that we now know more

REFERENCES

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about the organization and architecture of these polymer-forming proteins than we do about their physiological functions and the molecular basis for how they contribute to the cellular processes in which they participate. Hence, what remains is to bridge this gap and connect the features of septin structure to the demands of the biological processes in which septin-containing structures participate. In this regard, the application of modern EM techniques, such as 3-D tomography, in combination with mutant or tagged septin alleles, designed on the basis of our improved understanding of rod and filament architecture, should permit analysis of septin ultrastructure that will ultimately solve unequivocally the riddle of the orientation and organization of the septin filaments within the collar at the yeast bud neck. Furthermore, the actual mechanisms that control septin rod assembly and regulate the dynamic cell cycle-dependent transitions that septin-containing structures undergo also remain largely ill-defined. However, the newly acquired structural information now provides a set of basic tools with which to attack all of these remaining questions and to move forward, with the promise of achieving the kind of profound understanding accomplished by others who have studied historically more cytologically prominent cytoskeletal polymers.

ACKNOWLEDGEMENTS We want to acknowledge in particular the stimulating and fruitful collaboration we have had with our colleague, Prof. Eva Nogales, and various members of her group, specifically Dr Aurelie Bertin, Dr Patricia Grob and Galo Garcia. We also thank another colleague, Prof. Thomas C. Alber and his associate, Dr Ho-leung Ng, for very helpful discussions and technical assistance, Dr Mark S. Longtine, Dr Douglas R. Kellogg, Dr John D. York and Dr Scott D. Emr for the communication of unpublished information, and Raymond E. Chen for useful conversations and comments on a draft of the manuscript. We also gratefully acknowledge the support of a Postdoctoral Research Fellowship (#61-1295) from the Jane Coffin Childs Memorial Fund for Medical Research (to M.A.M.) and of NIH Research Grant GM218421 (to J.T.).

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Bertin, A., McMurray, M.A., Grob, P. et al. (2008) Assembly and supramolecular organization of the septin hetero-oligomers and filaments of the yeast, Saccharomyces cerevisiae. Proceedings of the National Academy of Sciences of the United States of America, 105, 8274–9. Byers, B. and Goetsch, L. (1976a) A highly ordered ring of membrane-associated filaments in budding yeast. Journal of Cell Biology, 69, 717–21. Byers, B. and Goetsch, L. (1976b) Loss of the filamentous ring in cytokinesis-defective mutants of budding yeast. Journal of Cell Biology, 70, 35. Carroll, C.W., Altman, R., Schieltz, D. et al. (1998) The septins are required for the mitosis-specific activation of the Gin4 kinase. Journal of Cell Biology, 143, 709–17. Casamayor, A. and Snyder, M. (2003) Molecular dissection of a yeast septin: distinct domains are required for septin interaction, localization, and function. Molecular and Cellular Biology, 23, 2762–77. Caviston, J.P., Longtine, M., Pringle, J.R. and Bi, E. (2003) The role of Cdc42p GTPase-activating proteins in assembly of the septin ring in yeast. Molecular Biology of the Cell , 14, 4051–66. Chi, A., Huttenhower, C., Geer, L.Y. et al. (2007) Analysis of phosphorylation sites on proteins from Saccharomyces cerevisiae by electron transfer dissociation (ETD) mass spectrometry. Proceedings of the National Academy of Sciences of the United States of America, 104, 2193–98. Chu, S., DeRisi, J., Eisen, M. et al. (1998) The transcriptional program of sporulation in budding yeast. Science, 282, 699–705. Cid, V.J., Adamikova, L., Cenamor, R. et al. (1998) Cell integrity and morphogenesis in a budding yeast septin mutant. Microbiology, 144, 3463–74. Cid, V.J., Adamikova, L., Sanchez, M. et al. (2001) Cell cycle control of septin ring dynamics in the budding yeast. Microbiology, 147, 1437–50. Dai, J., Jin, W.H., Sheng, Q.H. et al. (2007) Protein phosphorylation and expression profiling by Yin-yang multidimensional liquid chromatography (Yin-yang MDLC) mass spectrometry. Journal of Proteome Research, 6, 250–62. DeMarini, D.J., Adams, A.E., Fares, H. et al. (1997) A septin-based hierarchy of proteins required for localized deposition of chitin in the Saccharomyces cerevisiae cell wall. Journal of Cell Biology, 139, 75–93. Deng, Y., Liu, J., Zheng, Q. et al. (2006) Antiparallel four-stranded coiled coil specified by a 3-3-1 hydrophobic heptad repeat. Structure, 14, 247–55. De Virgilio, C., DeMarini, D.J. and Pringle, J.R. (1996) SPR28, a sixth member of the septin gene family in Saccharomyces cerevisiae that is expressed specifically in sporulating cells. Microbiology, 142 (Pt 10), 2897–905. Dobbelaere, J. and Barral, Y. (2004) Spatial coordination of cytokinetic events by compartmentalization of the cell cortex. Science, 305, 393–96. Dobbelaere, J., Gentry, M.S., Hallberg, R.L. and Barral, Y. (2003) Phosphorylationdependent regulation of septin dynamics during the cell cycle. Developmental Cell , 4, 345–57. Douglas, L.M., Alvarez, F.J., McCreary, C. and Konopka, J.B. (2005) Septin function in yeast model systems and pathogenic fungi. Eukaryotic Cell , 4, 1503–12. Drees, B.L., Sundin, B., Brazeau, E. et al. (2001) A protein interaction map for cell polarity development. Journal of Cell Biology, 154, 549–71.

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4 Yeast septins: a cortical organizer Yves Barral Institute of Biochemistry, ETH Zurich, Schaffmatstrasse 18, 8093 Zurich, Switzerland

INTRODUCTION Hartwell and colleagues identified the septins in the early 1970s as four yeast cell division cycle (cdc) genes (CDC3 , CDC10 , CDC11 , CDC12 ) that caused cytokinesis defects when inactivated (Pringle Chapter 1, this book; Hartwell, Culotti and Reid, 1970; Hartwell, 1971; Hartwell et al., 1974; Longtine et al., 1996). Genetic analysis showed that septin mutants were also defective in cell morphogenesis, leading to the formation of elongated buds. These four septin genes encode related proteins, characterized by the presence of a Ras-related GTPase domain followed by a coiled-coil domain (for review see Faty, Fink and Barral (2002)). The C-terminal coiled-coil indicates that unlike Ras small GTPases septin proteins stably associate with binding partners. All four proteins localize together and in an interdependent manner to the yeast bud neck (Haarer and Pringle, 1987; Kim, Haarer and Pringle, 1991). Moreover, septin mutants lacked the so-called bud-neck filaments, an ordered structure apparent at the bud neck in Electron Microscpy (EM) images (Byers and Goetsch, 1976). Accordingly, septins assemble together into a novel cytoskeletal structure made of filaments, and recombinant complexes composed of Cdc3, Cdc10, Cdc11 and Cdc12 form filaments in vitro (Frazier et al., 1998). Over the past twenty years, the yeast septin collar at the bud neck have been implicated in a number of cellular processes, ranging from cell division and cell cycle control, to bud site selection, cell polarity and the compartmentalization of organelles. Here, I will review these different functions, and their known molecular mechanisms. I will subsequently try to build on this knowledge to evaluate different models about what the actual molecular function of septins might be. However, before discussing the details of septin function in yeast, let’s consider first the structures they form. The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

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SEPTIN STRUCTURES IN YEAST Budding yeast septins are involved in two major structures. During vegetative divisions, the five septins Cdc3, Cdc10, Cdc11, Cdc12 and Shs1/Sept7 (Mino et al., 1998; Kim, Haarer and Pringle, 1991; Haarer and Pringle, 1987; Ford and Pringle, 1991) form the bud-neck filaments. Upon completion of meiosis, another complex comprising Cdc3, Cdc10, Cdc11, Spr28 (De Virgilio, DeMarini and Pringle, 1996) and Spr3 (Fares, Goetsch and Pringle, 1996) contribute to spore wall formation. Our knowledge of this second process remains fragmentary. This review focuses on vegetative septin structures.

Three septin structures at the bud neck Septins form at the bud neck three successive structures during the budding cycle (Faty, Fink and Barral, 2002). At bud emergence, septins form a cap that covers the forming budding site (Versele and Thorner, 2004). This cap structure is transient and has not been observed identically by all authors. Some have described a mesh of randomly oriented filaments (Cid et al., 2001), while others reported diffuse staining throughout the presumptive bud site (Versele and Thorner, 2004). Yet other investigators documented the formation a ring encircling the future bud site (Haarer and Pringle, 1987; Kim, Haarer and Pringle, 1991). The reason for these differences are unclear, but probably reflects the method of visualization used. Green fluorescent protein (GFP)-tagged septins revealed rather the diffuse cap, while septin visualization by indirect immunofluorescence on fixed cells showed either a filament mesh or a ring. Fixation probably stabilizes the structural intermediates that are the least dynamic, trapping septins in their filamentous form. GFP-tagged septins might also shift the population towards the most dynamic intermediates. Thus, diffuse and filamentous structures, might correspond to two intermediates that coexist in the reality. Very soon after bud emergence, septins form a sharp ring identical by all visualization techniques. This ring, also called septin collar, is formed of septin filaments tightly apposed to the plasma membrane and corresponds to the bud neck filaments observed by electron microscopy (Byers and Goetsch, 1976). Following the curvature of the plasma membrane at the bud neck, this ‘ring’ takes an hourglass shape (Cid et al., 2001; Lippincott et al., 2001); it is a cylinder constricted in the middle (review Gladfelter, Pringle and Lew, 2001; Longtine and Bi, 2003). How the filaments are organized in this cylinder was solved only recently. The bud-neck filaments observed by EM run parallel to the plane of the bud neck (Byers and Goetsch, 1976). However, these filamentous densities might correspond to cross-links between filaments. Indeed, in vitro and under low salt conditions recombinant septin filaments assemble into sheets (Frazier et al., 1998), where

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stripes become visible at regular intervals perpendicular to the actual filaments. These stripes very much resemble the neck filaments. Furthermore, mutants lacking the septin-dependent kinase (SDK) Gin4 form irregular rings where filaments seem to become visible as bundles that align with the mother-bud axis (Longtine and Bi, 2003). Therefore, in the septin collar the septin filaments might not assemble in a spring shape, as originally expected, but seem to orient parallel to the mother-bud axis like the boards of a barrel (Frazier et al., 1998). The current, and probably definitive solution to this discussion was provided by the Mitchison lab in a set of very innovative experiments, using polarized light microscopy (Vrabioiu and Mitchison, 2006; Vrabioiu and Mitchison, 2007). These data show that septin filaments run parallel to the mother-bud axis (Vrabioiu and Mitchison, 2006). Thus, septins filaments form a sort of gaze through lateral interactions at the plasma membrane. The septin collar stays at the bud neck throughout bud growth and mitosis, i.e. for half to three quarters of the cell cycle. Upon mitotic exit it then splits into two rings, one on each side of the bud neck (Lippincott et al., 2001). Polarized light microscopy indicates that during this transition the filaments turn 90◦ and align with the plane of the bud neck (Vrabioiu and Mitchison, 2006). Simultaneously, the actomyosin ring starts to contract, and the curvature of the plasma membrane inverses to close the bud neck (Figure 4.1, IV). Therefore, the septin rings must shift from a nearly cylindrical organization (septin collar) to two flat rings (Figure 4.1, IV). This shape change could explain the reorientation of the filaments (see below). During these changes, the diameter of the septin collar and the septin rings remains constant (Lippincott and Li, 1998a). The septin ring(s) does not contract at cytokinesis.

Septin dynamics Cytoskeletal networks generally are highly dynamic, such as microtubules and actin microfilaments. Continuous alternation between growth and shrinkage promotes rapid reorganization and reshaping of the cell in response to intra and extracellular signals. In contrast, septin filaments show very little dynamics during most of the cell cycle. In fluorescence recovery after photobleaching (FRAP) experiments, septin turnover is observed only during short intervals prior to bud emergence (phase I, Figure 4.1), at ring splitting (phase III), and as the ring disassembles in the next G1 phase (Dobbelaere et al., 2003; Caviston et al., 2003). These intervals correspond to rapid reorganization steps. During phase II (septin collar) and phase IV (split rings), little to no recovery is observed, indicating that the septin filaments do not turnover. During the dynamic intervals, fluorescence recovery appears to take place at the cost of the unbleached part of the ring, suggesting that the ring is in a ‘fluid’


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I

II

III

IV

Figure 4.1 The different forms adopted by the septin cytoskeleton at the yeast bud neck throughout the budding process. During bud emergence (stage I) the filaments assemble into a cap or flat ring morphology structure around the site of bud emergence. As the bud emerges (stage II) the filaments adopt a ring like structure that follows the outward curvature of the plasma membrane at the bud neck. In stage III, which corresponds to the period of bud growth, the ring now forms an hourglass shaped collar at the transition between mother and bud. This collar is tightly apposed to the inner surface of the plasma membrane. Upon cytokinesis onset, the collar splits into two rings (Stage IV) that are remodeled again to follow the changing curvature of the plasma membrane as the neck closes. Thereby, the rings probably adopt back the flat ring morphology. In stage II and III, the filaments are aligned with the mother-bud axis. We suggest that they open in an iris-like manner at each extremities of the collar, where it increases in diameter. In the stage I and IV, the filaments slide along each other to accommodate the flat topology of the ring.

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state, where subunits move mainly inside the ring. Accordingly, Dobbelaere et al. observed that completely photobleached rings did not recover, indicating that there is little or no exchange of subunits between the ring and the cytoplasm (Dobbelaere et al., 2003). This observation was, however, not reproduced by Caviston and coworkers, who observed some recovery of fully photobleached rings (Caviston et al., 2003). This discrepancy is possibly due to these experiments focusing on different stages in phase I. Indeed, Dobbelaere et al. focused on cells undergoing bud emergence, while Caviston et al. focused on cells prior to bud emergence. Therefore, we suggest that during the first half of phase I the newly recruited septins do rapidly exchange with the cytoplasm. During the second half, the filaments start to be more stable, and movement is restricted within the ring. Thus, we distinguish three different dynamic states for septin higher-order structures: (i) During assembly and disassembly rings are dynamic and exchange subunits with the cytoplasm. (ii) Concomitant with changes in the curvature of the plasma membrane at bud emergence and cytokinesis, the septin ring is in the fluid state. It adapts to the new topology of the cell, and exchanges are due to filament movements within the ring. (iii) During the rest of the time, which is most, the septin ring is in a frozen state. The subunit do neither move within the ring nor exchange with the cytoplasm.

A model for filament organization in the septin ring Thus, septin structures adapt remarkably well to the complex topological changes of the plasma membrane at bud emergence and cytokinesis, without loosing membrane interaction. How this behaviour is supported by the structure of the collar is not known. It is unclear how the filaments, which are parallel to the bud neck axis, support the hourglass shape of the collar. It is unlikely that the filaments are either compressed in the middle of the cylinder, or separate from each other at its extremities. Instead, we propose that the filaments might slide along each other as the barrel becomes widens, like in an iris (Figure 4.1, X). At ring splitting, this movement might reach the extreme, such as to fit the ring in a disc, on the cleavage plane (Figure 4.1, X). This model is attractive because it sheds light onto what the fluid phase might correspond to. Indeed, in late phase I the septin ring rearranges from a disc-crown to a cylinder, requiring extensive sliding of the filaments along each other. This would explain the fluorescence recovery observed in FRAP experiments, despite the lack of exchange with the cytoplasm. Likewise, the recovery observed during phase III might be due to similar sliding events as the membrane curvature changes again. Thus, our current picture of septin organization reveals a sophisticated system of filaments that covers the topologically complex surface of the plasma membrane at the bud neck and precisely follows its changes.

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THE CELLULAR FUNCTIONS OF SEPTINS: VARIETY AND RECURRENCE Yeast septins have been implicated in a broad panel of functions in cytokinesis, cell cycle control, and cellular compartmentalization. These processes involve septins in both structural and regulatory functions. This structural/regulatory duality has long represented an important difficulty in septin research.

Septins in cell division In yeast, division plane determination is subordinated to the budding process and cytokinesis always takes place at the bud neck. Thus, the division plane is set at bud emergence, early in the cycle, even before DNA replication and mitosis (for review on cytokinesis in budding yeast (Tolliday, Bouquin and Li, 2001; Bi, 2001)). Therefore, yeast cells need (i) to memorize the position of the cleavage plane up until cytokinesis, and (ii) to position the spindle perpendicular to this plane. This second condition ensures that spindle elongation correctly segregates one daughter nucleus into the mother and one into the bud. The yeast septin collar is involved in the solution to these problems. Its position serves as a memory for the position of the bud neck until cytokinesis, and as a spatial cue to direct spindle positioning.

Septins in spindle positioning The septin ring at the bud neck plays at least three distinct roles in spindle positioning to the bud neck and alignment with the mother-bud axis. Its first role is to organize capture sites for microtubule plus-ends at the cortex of the bud neck. Upon capture, force is generated on attached microtubules to pull the spindle towards the bud neck (Kusch et al., 2002). Septin contribution to microtubule attachment is mainly indirect. Indeed, the septin ring on the inner side of the plasma membrane at the bud neck is in fact not in direct contact with the cytoplasm, but shielded by a sheet of endoplasmic reticulum (ER, see below) (Byers and Goetsch, 1976; Luedeke et al., 2005). Thus, microtubules are unlikely to directly interact with septins at the bud neck. Furthermore, microtubule attachment at the bud neck also depends on Bud6. Bud6 is a peripheral-membrane protein that localizes to the bud neck, probably to the endoplasmic membrane, in a septin-dependent manner (Amberg et al., 1997; Luedeke et al., 2005; Huisman et al., 2004). Thus, microtubule attachment sites might be located on the surface of the ER at the bud neck. There, Bud6 might directly interact with microtubule ends during microtubule capture (Huisman et al., 2004). The second role of the septin collar is in the regulation of microtubule dynamics at the bud neck, downstream of attachment. There, septin function depends on the SDKs Hsl1 and Gin4 (Barral et al., 1999; Altman and Kellogg, 1997; Kusch et al.,

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2002; Tanaka and Nojima, 1996; Ma, Lu and Grunstein, 1996). In the absence of SDKs, microtubules attached to the bud neck show a reduced catastrophe frequency and grow longer. They also fail to stay attached once they finally start to shrink. As a consequence the spindle pole body (SPB) (functionally equivalent to the centrosome of animal cells) at the other end of the attached microtubule is pushed back into the mother cell instead of being pulled towards the cleavage plane (Kusch et al., 2002). The nature of the substrates regulated by the SDKs remains unknown. Finally, a third process depending on the septin collar is the regulation of dynein distribution. Indeed, in wild-type cells dynein accumulates at the tip of the microtubules emanating from only one SPB, the SPB directed towards the bud (and hence towards the bud neck), here called SPBbud (Shaw et al., 1997; Segal et al., 2000; Grava et al., 2006). Cells lacking SDKs localize dynein to microtubules emanating from both SPBs, and dynein level on microtubules remains weak (Grava et al., 2006). How SDKs control the distribution of dynein is still unknown, but time-lapse analysis indicate that dynein recruitment to the SPBbud and emanating microtubules depends on contact events between the microtubule plus end and the bud neck (Grava et al., 2006). In turn, dynein asymmetry ensures that only one SPB is finally pulled into the bud upon anaphase onset (Grava et al., 2006). Thus, septins contribute to spindle positioning mainly by providing a spatial cue for spindle organization and the control of spindle movements. In these functions, the septins provide a scaffold for the assembly and regulation of other structures, such as the ER and SDKs.

Septins and the assembly of the cytokinetic machinery Cytokinesis is a highly complex process, requiring the coordination of multiple events. Like in animal cells, yeast cytokinesis proceeds in two steps: furrow ingression, which reduces the cytoplasmic connection between the daughter cells to a narrow bridge, and abscission, which resolves the plasma membrane and definitely separates the two daughters (for review on cytokinesis see Guertin, Trautmann and McCollum, 2002; Glotzer, 2005). Furrow ingression depends on myosin II, Myo1, which is recruited in a septin-dependent manner to the bud neck shortly after bud emergence, but does not function before late anaphase, when it contributes to actin recruitment and assembly of the actomyosin contractile ring (Lippincott and Li, 1998b; Bi et al., 1998). Upon mitotic exit, the kinase Mob1/Dbf2, a downstream effector of the mitotic exit network (MEN) triggers ring contraction (Lippincott and Li, 1998b; Lippincott et al., 2001; Luca et al., 2001; Menssen, Neutzner and Seufert, 2001). MEN is equivalent to the septation initiation network (SIN) in fission yeast (for review see Krapp, Gulli and Simanis (2004). Although septins are involved in the recruitment of myosin II to the bud neck, they are not part of the actomyosin ring itself, which contracts between the two split septin rings (Lippincott et al., 2001; Dobbelaere and Barral, 2004).

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Remarkably, myosin II is not essential for cytokinesis in yeast. Thus, septins must play additional roles besides myosin II recruitment. One such role is the recruitment of other cytokinetic factors, such as the proteins Hof1 (Lippincott and Li, 1998a), Cyk3 (Korinek et al., 2000; Vallen, Caviston and Bi, 2000; Iwase et al., 2007). Hof1, related to fission yeast cdc14, localizes like Myo1 to the bud neck throughout most of bud growth and focuses between the split septin rings at cytokinesis (Vallen, Caviston and Bi, 2000; Lippincott and Li, 1998a; Dobbelaere and Barral, 2004). Its exact function is not yet clear, but is independent of and overlapping with that of Myo1. While both myo1 and hof1 single mutant cells are alive, the myo1 hof1 double mutant is dead (Vallen, Caviston and Bi, 2000). Thus, in the absence of the Myo1 Hof1 is able to support furrow ingression and vice versa, while co-inactivation of both prevents ingression. The molecular function of Cyk3 is also unclear. It is recruited to the bud neck only in late mitosis (Korinek et al., 2000) and functions independently of both Myo1 and Hof1. Indeed, while each of the cyk3, hof1 and myo1 single mutants are alive in most genetic backgrounds, each double mutant combining two of these mutations is dead (Korinek et al., 2000). Thus, Cyk3 can support cytokinesis in the absence of any one of Hof1 or Myo1. Together, these data indicate that at least three parallel pathways contribute to yeast cytokinesis. Each of these pathways depends on septin function for their proper recruitment to the cleavage plane. Thus, one aspect of septin function in yeast cytokinesis is to serve as a scaffold for the assembly of the cytokinetic machinery prior to cell cleavage.

Septins during cytokinesis The role of septins in cytokinesis is however not restricted to pre-cytokinetic events. Disruption of the split septin rings during cytokinesis does not affect the localization of Hof1 and Myo1 anymore but still leads to cytokinetic failure (Dobbelaere and Barral, 2004). During cytokinesis, these rings located on each side of the abscission site function as diffusion barrier to confine numerous membrane-remodeling factors to the abscission site (Dobbelaere and Barral, 2004). Such factors comprise the exocyst, a complex involved in the targeting of exocytic vesicles to the plasma membrane (Finger, Hughes and Novick, 1998) and the polarisome (Sheu et al., 1998), involved in actin-cable nucleation at the cell cortex (Dobbelaere and Barral, 2004). The exocyst is absolutely required for proper furrowing and abscission. The implication of several independent molecular pathways in cytokinesis clearly provides robustness to the process. In turn, their involvement in all of these pathways explains why yeast septins are so crucial for cytokinesis. These data also establish that septins provide at least two independent functions: a scaffolding function prior to cytokinesis and a confinement role during cell cleavage. As we will see, this second duality, scaffold/barrier, is also recurring in septin function.


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Conservation of septin functions It is not clear whether the cell division functions of septin are conserved. Septins do not appear to mediate spindle positioning in another cell type than yeast. However, septins certainly interact with and contribute to the regulation of interphase and mitotic microtubules in mammals (Surka, Tsang and Trimble, 2002; Spiliotis, Kinoshita and Nelson, 2005; Kremer, Haystead and Macara, 2005). Thus, a conserved connection between septin and microtubules might exist, yet be implicated in different processes in different cell types. Similarly, while septins are not necessary for cytokinesis in many cells. For example, C. elegans embryos and fission yeast assemble and contract their actomyosin ring and complete cytokinesis fine in the absence of septins (Nguyen et al., 2000; An et al., 2004; Tasto, Morrell and Gould, 2003; Berlin, Paoletti and Chang, 2003; Wu et al., 2003). Still, septins localize to both sides of the cleavage plane in all tested cell types (for review see Kinoshita and Noda, 2001; Joo, Tsang and Trimble (2005)). Furthermore, recent data established that insect, nematode and mammalian septins interact with anillins (Field and Alberts, 1995) and contribute to the proper regulation and functional robustness of myosin II (Oegema et al., 2000; Kinoshita et al., 2002; Hime, Brill and Fuller, 1996; Maddox et al., 2007). Anillins form a family of peripheral-membrane proteins that interact with both actin- and septin-based structures. In tissue culture, septins are required for cytokinesis events posterior to furrow ingression, suggesting that they contribute to abscission (Kinoshita et al., 1997; Joo, Tsang and Trimble, 2005). Thus, the molecular functions of septin in both actomyosin-related and abscission processes are likely to be conserved, although the impact of these functions on the cleavage process is variable. The nature and strength of this impact probably depends on the biology of the cell. For example, the role of septins in ‘memorizing’ the location of the bud neck during bud growth might play a determining role in making budding yeast septins so central. By contrast, the size of the nematode oocyte is probably the reason why emphasis is rather put on actomyosin ring contraction in these cells.

Septins and cell morphogenesis Septins contribute in two ways to budding. First, they contribute to bud-site selection in late G1. Second, they mediate proper bud morphogenesis.

Septins and bud site selection Yeast buds are not positioned randomly on their mothers, but according to selection programs that depend on the plo¨ıdy of the cell. Haploid cells bud axially, i.e. adjacent to the previous budding site. Diploids cells display a bipolar budding pattern. They bud alternatively adjacent and distal to the original cite of cytokinesis (see Casamayor and Snyder (2002) for a review on bud-site selection). In both cases, the septin rings set the spatial cues that the cell use to position new buds. In

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haploid cells, the septin rings of telophase cells, which persist during G1, activate a signalling cascade comprising the anillin-related protein Bud4 (Sanders and Herskowitz, 1996; Park and Bi, 2007; Gladfelter et al., 2005), the Bud4-interacting protein Bud3 (Chant et al., 1995), the membrane proteins Axl1 and Axl2 (Fujita et al., 1994; Roemer et al., 1996), the ras-related GTPase Rsr1/Bud1 (Bender and Pringle, 1991), and its guanine nucleotide exchange factor (GEF), Bud5 (Bender, 1993) (reviewed in Park and Bi (2007)). Although we do not understand this cascade in detail, its activation is initiated by the septin-dependent recruitment of the Bud4/Bud3 complex to the cortex (Sanders and Herskowitz, 1996; Chant et al., 1995). This initial step depends on septin function and appears to involve a direct interaction between Bud4 and septins. Subsequent activation of Bud1 by this cascade at the previous site of cytokinesis triggers the recruitment of the Rho-like GTPase Cdc42, and the assembly of the new bud site (Kozminski et al., 2003). In turn, a novel septin collar is assembled at the new bud neck. In diploid cells, septins similarly contribute to the recruitment of the cortical proteins Bud8 and Bud9, involved in the spatial regulation of Bud5 and Bud1 (Park and Bi, 2007; Schenkman et al., 2002). Thus, in haploids and diploids the role of septins in bud-site selection involves their scaffolding function.

Septins and bud morphogenesis The involvement of septins in bud morphogenesis was first evidenced by the elongated buds formed by septin-defective cells (Hartwell, 1971). This morphology first suggested a role for septins, which are located at the bud neck, in the spatial control of growth deposition within the bud. Normal bud morphogenesis takes place in two steps. During bud emergence, buds grow apically; the localization of growth, i.e. plasma membrane expansion and cell wall remodelling, is restricted to the bud tip. Subsequently, buds grow isotropically, i.e. growth becomes deposited uniformly over the bud surface (Lew and Reed, 1993; Adams and Pringle, 1984). Because of its rigidity, the extension of the cell wall is determined locally by the targeted delivery of new material and cell wall remodelling enzymes by exocytosis (reviewed in Pruyne and Bretscher, 2000a; Pruyne and Bretscher 2000b). Bud morphology is hence determined by the pattern of vesicle delivery, first to the bud tip during bud emergence, apical bud growth and to the entire bud periphery during isotropic bud growth. Vesicles delivery to the plasma membrane is ensured by myosin V-dependent transport along actin cables (Johnston, Prendergast and Singer, 1991; Liu and Bretscher, 1992; Govindan, Bowser and Novick, 1995) reviewed in Pruyne et al., (2004)). Because Myosin V moves its cargos unidirectionally towards the barbed end of microfilaments, vesicles are delivered to the sites of actin-cable nucleation by formins (Pruyne et al., 2002; Evangelista et al., 2002; Sagot et al., 2002), enclosed in the polarizome (Sheu, Barral and Snyder, 2000). Further spatial regulation is provided by the fact that exocytosis of vesicle content also requires the activity of a multiprotein complex, the exocyst (TerBush


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et al., 1996) and of its regulators the Rho-like GTPases Rho1 and Cdc42 (Zhang et al., 2001; Roumanie et al., 2005). Thus, the localization of the exocyst and the polarizome determines the distribution of cell growth. Accordingly, at bud emergence exocyst and polarizome co-localize to the bud site, accumulate at the bud tip during apical growth, and redistribute to the entire bud cortex at the shift to isotropic bud growth (reviewed in Pruyne et al. 2004). In addition, bud morphogenesis is coordinated with cell cycle progression (Lew and Reed, 1993). Bud emergence and apical bud growth is initiated in late G1 by Cdk1 and the G1 cyclins Cln1 and Cln2. Subsequently, the B-type cyclins Clb1 and Clb2 trigger the shift to isotropic bud growth during the S phase. Consequently, cells that fail to activate Clb1 and Clb2 keep growing apically and form elongated buds. These observations suggested two models to explain how septins contribute to bud morphogenesis. They might help control the spatial pattern of vesicle delivery inside the bud, or participate to the control of cell cycle progression and the activation of Clb1,2/Cdk1 complexes. The second model is the most accurate.

The morphogenetic checkpoint Beside septin-defective cells, a number of other mutants form elongated buds (Blacketer et al., 1993; Blacketer, Madaule and Myers, 1995; Kellogg and Murray, 1995). Among them, the nap1, gin4, hsl1, hsl7, and elm1 cells also delay anaphase onset, indicating that cell cycle progression is affected (Barral et al., 1999; Bouquin et al., 2000; Altman and Kellogg, 1997). Both the morphological and cell cycle phenotypes of these mutants are suppressed by disrupting the SWE1 gene, the ortholog of the wee1 kinase in Schizzosccharomyces pombe (Booher, Deshaies and Kirschner, 1993), and by mutation of the tyrosine 19 of Cdk1/Cdc28 to phenylalanine (Barral et al., 1999; Shulewitz, Inouye and Thorner, 1999; Longtine et al., 2000). Tyrosine 19 of Cdk1 is the residue that is phosphorylated by Swe1 in vivo and the Y19F mutation prevents Cdk1 inhibition by Swe1. Thus, the anaphase delay and the elongated bud phenotype of these mutants is caused by Swe1-dependent repression of Clb1,2/Cdk1. Together, these and others data have established a model in which Hsl1, Gin4, Hsl7 and Elm1 act in a cascade that control activation of the kinase Cdk1/Clb1,2 through inhibition of its negative regulator Swe1 (see Figure 4.2). Remarkably, this pathway links Cdk1 activity to the status of septin assembly. Indeed, Hsl1, Hsl7, Gin4, Elm1 and Swe1 all localize to the bud neck during bud growth, in a septin-dependent manner (Carroll et al., 1998; Barral et al., 1999; Shulewitz, Inouye and Thorner, 1999; Moriya and Isono, 1999). Among them, both Hsl1 and Gin4 are SDKs; they directly interact with and depend on septins for activity (Carroll et al., 1998; Barral et al., 1999; Bouquin et al., 2000). Accordingly, anaphase onset is delayed in septin-defective cells and both this delay and the elongated bud phenotypes of septin mutants are suppressed upon disruption of SWE1 (Barral et al., 1999). Thus, the abnormal bud morphology of


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septin collar

Elm1 ?

Gin4

Hsl1 Hsl7

? Swe1

Cdc28/Clb1,2 Y19

Cdc28/Clb1,2 Y19

anaphase isotropic bud growth

P

Mih1

Slt2/Mpk1

Mkk1, 2

Bck1

Rho1

actin defects

Figure 4.2 Schematic organization of the two branches of the morphogenetic pathway. On the top, the Hsl1/Hsl7 cassette responds to proper septin organization by targeting the Swe1 kinase to degradation. Thereby, it links activation of the Cdk complex Cdc28/Clb1,2 by the Mih1 phosphatase to proper assembly of the septin collar. Cdc28/Clb1,2 can in turn trigger the shift of bud growth from apical to isotropic, as well as the activation of the anaphase promoting complex. From the bottom, the Slt2/Mpk1 MAP kinase pathway prevents Mih1 activation in response to actin defects. Thereby it ensures that no adaptation from Swe1-dependent cell cycle arrest can take place before the bud has reached the proper size.

septin mutants is due to a cell cycle response that maintains the cells in apical bud when the septin collar is disrupted. What is the physiological relevance of linking Swe1-control, cell cycle progression and septins in budding yeast? There are two possible interpretations. On one hand, the septin collar might provide a scaffold to organize and control the


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Swe1 pathway. On the other hand, the Swe1 pathway might monitor septins to delay anaphase in response to septin organization defects. Current data are consistent with a Swe1/Mih1 cassette monitoring both actin and septin organization, independently of each other (Figure 4.2). Indeed, cells where actin organization is disrupted early in bud growth arrest prior to anaphase in a Swe1-dependent manner for hours, much longer than septin-defective cells do (McMillan, Sia and Lew, 1998). This arrest depends on the Rho1-Mpk1/Slt2 pathway (Harrison et al., 2001), which acts to inhibit the phosphatase Mih1. Mih1, the homolog of fission yeast cdc25 counteracts Swe1 by removing the inhibitory phosphate that Swe1 puts on Cdk1 (Russell, Moreno and Reed, 1989). Thereby, it stimulates the resumption of cell cycle progression after Swe1-dependent arrest. Thus, actin defects occurring early in bud growth, when the septin collar is not yet fully stabilized, prevent adaptation from Swe1-dependent arrest and impair anaphase onset. Accordingly, actin defects no longer cause a cycle arrest if the cells have progressed beyond the time at which the Hsl1/Hsl7 regulatory cassette becomes active and represses Swe1, thanks to proper septin assembly. In contrast, septin-defective cells are fully proficient for the induction of a Swe1-dependent arrest, yet adapt and escape the arrest after 40 to 50 minutes. Thus, on one hand, the SDK/Swe1 pathway monitors septin organization and prevents activation of late Cdk1 complexes when the septin ring is not properly assembled. On the other hand, the Rho1-Mpk1/Slt2-Mih1 pathway monitors actin function and bud growth and allow adaptation to Swe1-dependent arrest only if the bud has already reached a decent size. In the first pathway, SDKs function as sensors for proper septin organization (Barral et al., 1999; Bouquin et al., 2000; Hanrahan and Snyder, 2003; Carroll et al., 1998). An interesting twist to this story is that the Hsl1/Hsl7 module can therefore be viewed as a sensor for bud emergence. Indeed, septin reorganization following the changes of plasma membrane curvature during bud emergence appears to be necessary for proper activation of the Hsl1 kinase (Theesfeld et al., 2003). However, it must be pointed out that the two lines of thinking (i) Hsl1 monitors proper septin organization or (ii) the Hsl1/septin module monitors the topology of the plasma membrane at the bud neck, are only semantically different. Molecularly speaking, they probably correspond to one same mechanism, in which bud emergence is required for the proper formation of a septin collar that is the only septin structure supporting SDK activation. In any case, the morphogenetic checkpoint derived from these mechanisms ensures that anaphase onset depends on the presence of a bud, of a properly assembled cleavage apparatus and on the cell cortex and the cell wall being correctly organized.

Septin and cell polarity Although preventing anaphase completion in response to defects of the cytokinetic machinery makes biologically sense, it is not immediately clear what the advantages are of preventing the shift to isotropic growth in response to septin

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assembly defects. The phenotypic characterization of cells lacking both Swe1 and septin function, i.e. of cells undergoing the shift to isotropic bud growth in the absence of the septin collar, helped answer this question. Indeed, although at restrictive temperature the swe1 cdc12-6 double mutant cells fail to form elongated buds, they are still morphologically abnormal. The buds remain small, and the mother cells swell (Barral et al., 2000), indicating that septin defective these cells fail to restrict growth to the bud. Similarly, the swe1 cdc12-6 cells fail to confine the polarizome and the exocyst to the bud only (Barral et al., 2000). Thus, an important function of the septin ring is to help restrict the localization of polarity factors and cell growth to the bud. Thus, the septin collar appears to lay at the bud neck a boundary for the organization of cortical polarity. A major mechanism by which the septin collar bounds the distribution of cortical components is through the formation of a diffusion barrier in the plasma membrane. This has been clearly established for the case of Ist2, an integral membrane protein localized to the bud plasma membrane. FRAP experiments indicated that Ist2 diffuses rapidly within the plasma membrane, yet fails to equilibrate between mother and bud. Upon disruption of the septin ring at the bud neck, however, the distribution of Ist2 equilibrates rapidly between mother and bud, indicating that septins contribute to preventing Ist2 from diffusing through the bud neck (Takizawa et al., 2000). The phenotype of the swe1 cdc12-6 cells indicate that this diffusion barrier helps maintaining the asymmetric distribution of polarity markers. There are at least two possible models for how septins function in the assembly of this diffusion barrier. First, septins might form this barrier themselves (Faty, Fink and Barral, 2002). Septins might directly interact with phospholipids such as phosphatidylinositol 4, 5-bisphosphate (PtdIns(4,5)P(2)) and phosphatidylinositol 3,4,5-trisphosphate (PtdIns(3,4,5)P(3)) (Zhang et al., 1999), which in yeast are required for their localization to the plasma membrane at the bud neck (Casamayor and Snyder, 2003; Rodriguez-Escudero et al., 2005). The many interactions linking septins subunits and phospholipids underneath a single septin filament might therefore lead to two simultaneous outcomes. The filaments would be tightly apposed to the cytoplasmic face of the plasma membrane, impairing the passage of integral plasma membrane proteins with any sizable cytoplasmic domain. These septin-lipid interactions would also recruit and immobilize phospholipids assembling a specialized lipid domain underneath the filament. Such a domain would in turn limit the diffusion of other lipids and the passage of membrane proteins poorly soluble in this lipid phase. Second, septins might not compose the barrier itself but rather be involved in its assembly and maintenance. In this model, other proteins must form the barrier itself. However, we failed so far to identify other bud neck proteins besides septins for being required for the restriction of polarized cortical proteins to the bud. Extensive studies will be required to test both types of models and determine their relative contribution to the control of lateral diffusion of plasma membrane proteins at the bud neck.

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Cellular compartmentalization Cell polarity consists in the asymmetric distribution of cellular factors between two poles of the cell. We recently described a related process, which consists in the compartmentalization of cellular organelles into mother and bud domains. A well-characterized example concerns the ER. While the yeast ER is a single continuous organelle extending throughout the cell cortex and the nuclear envelope, a lateral diffusion barrier restricts the exchange of ER-membrane proteins between mother and bud. This barrier does not seem to play a direct role in cell polarity. First, the proteins subjected to compartmentalization are not polarized. Second, disruption of the barrier in sur2 (Buvelot Frei and Barral, unpublished data), shs1 and bud6 (Luedeke et al., 2005) cells causes no obvious defect in cell polarity. However, ER compartmentalization might help maintaining or reinforcing cellular asymmetry downstream of polarity establishment and bud growth. For example, this barrier might help confining polarized mRNAs (Shepard et al., 2003; Takizawa et al., 2000) to the bud cortex by preventing their diffusion back into the mother cell. Thereby, it might contribute to the bud-specific expression of cell fate determinants such as the transcription factor Ash1 (Bobola et al., 1996). How do septins influence ER organization? Most of the surface of the yeast ER is covered with ribosomes, indicating that yeast cells contain mostly rough ER. At the bud neck, however, a continuous ring of smooth ER covers the septin collar. This sheet disappears in septin defective cells, where it is replaced with rough ER tubules as in the remainder of the cortex. Furthermore, the smooth ER at the bud neck lacks many ER-membrane proteins, such as the translocon and Hmg1 (Luedeke et al., 2005). Thus, smooth ER formation involves the exclusion of these proteins from that ER-domain. The peripheral membrane protein bud6 though appears to be enriched on the bud-neck ER. This protein, which localizes to the bud neck in a septin-dependent manner, is required for the restriction of lateral diffusion in the ER-membrane at the bud neck. Yet, it is not involved in the exclusion of ER-membrane proteins from the bud-neck ER (Luedeke et al., 2005). Thus, barrier formation might be a two steps process, where the septins first direct the formation of a ring of smooth ER at the bud neck, to which Bud6 binds to restrict diffusion. How septins direct smooth ER formation is not yet known. However, cells lacking the SDKs Hsl1 and Gin4 fail to assemble a barrier (Luedeke et al., 2005) and to exclude rough ER markers from neck region. Thus, formation of the barrier throughout the entire ER-membrane appears to be independent of direct septin interactions with the membrane. Rather, it seems to involve the phosphorylation of yet unidentified ER factors by SDKs.

MOLECULAR FUNCTIONS FOR YEAST SEPTINS While we start to know much about septin functions, we still know very little about septin biochemistry. Genetic and cytological studies indicate that septins

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fulfil their cellular functions in either of two ways: through the recruitment of specialized proteins to the bud neck, or through formation of lateral diffusion barriers in membranes passing the bud neck. In all these processes the main function of septins boils down to establishing, maintaining and signalling the position of the bud neck, defined as both a boundary and a functionally specialized area of the cortex. This septin collar forms a stable anchor point around which the cell organizes. As we will discuss below, these observations already make predictions about what septin’s function(s) must be at the molecular level. Septin function mostly depends on the formation of higher-order structures. First, this appears to be the condition for them to stay at the bud neck and not diffuse away. Second, the individual septins show no functional specialization, and so far no function could be genetically separated from collar assembly. Thus, collar assembly is a pre-requisite for all cellular aspects of septin function. Consequently, most septin-interacting proteins might bind septin filaments or sheets of filaments rather than individual subunits, explaining the difficulty of purifying septin-interacting proteins using traditional biochemistry techniques. Therefore, one fundamental challenge for future studies in septin research is to understand the mechanisms governing septin dynamics, the assembly and disassembly of higher order structures, and the interaction of septin-binding proteins with individual septins and with the septin collar. As mentioned earlier, a key feature of septin sheets resides in their ability to precisely cover the plasma membrane at the bud neck, despite its complex topology and the complex changes it undergoes during bud emergence and cytokinesis. We suspect that the plasticity of septin assemblies lies at the heart of their functions. Particularly, it is very possible that the reorganization that the septin gaze undergoes during the cell cycle serves not only to maintain contact with a changing plasma membrane, but also to guide and control these changes and to coordinate with them the ability of septins to interact with and modulate the activity of other proteins, such as SDKs.

CONCLUSION AND PERSPECTIVES In light of what we start to understand about septin function in yeast, septins form a cytoskeletal system that is fundamentally different from microtubules and microfilaments. Septin structures are poorly dynamic, do not appear to serve as tracks for motor-driven and directional transport of cargos, and are non-polar (John et al., 2007; Sirajuddin et al., 2007). Instead, they are able to mark a specific location for an extended period of time, to tightly interact with membranes, which they help to organize and to serve as a complex scaffold that can recruit and activate proteins in a manner that seem to be coordinated with the topological changes of the underlying membrane. Thus, the yeast septin cytoskeleton forms structures that are more reminiscent of an organizer like centriols and centrosomes, rather than of a force generator as microtubules are. To keep with the skeletal metaphore,

REFERENCES

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the functional characterization of the microfilaments and the microtubules showed that they are the muscle of the cell. Septins now emerge as the missing bones on which these muscles anchor to articulate the forces that they generate. This pivotal position of septins in the biology of the yeast cell makes the septin collar an outstanding candidate to contribute to many long-term processes in the cell. In that sense, further research on septin biology is likely to provide us many insights about the long process through which the yeast bud emerges and differentiates itself from its mother. Thus, the septin collar has the perfect properties and is exactly at the right place to not only help us understand how cells divide but also how old cells generate successive new-born daughters while they continue to age. In years to come, we predict that septins will turn out to take a central role in cell biology well beyond yeast.

ACKNOWLEDGEMENTS I would like to express my deep gratitude to Stanie Buvelot Frei for her critical reading of the manuscript, her numerous and helpful comments, and her help with establishing and formating the final version of the manuscript. I would also like to acknowledge the members of the septin field for their comments and criticisms throughout our work, and particularly Peter Hall, Hilary Russel and John Pringle for taking on the challenge to edit this book.

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5 Septins in four model fungal systems: diversity in form and function Amy S. Gladfelter Department of Biology, Dartmouth College, Hanover, NH, USA

Peter Sudbery Department of Molecular Biology and Biotechnology, Sheffield University, Western Bank, Sheffield S10 2TN, UK

INTRODUCTION After their initial discovery by the analysis of genes found in the screen for cell-division-cycle mutants in the yeast Saccharomyces cerevisiae (Hartwell, 1971; Longtine et al., 1996), septin proteins have been identified based on sequence similarities in a variety of fungal genomes and indeed appear to be ubiquitous in the fungal kingdom (Lindsey and Momany, 2006; Pan, Malmberg and Momany, 2007). Septin orthologues clearly emerge in the amino acid sequences based on characteristic regions such as guanosine triphosphate (GTP)-binding and coiled– coiled domains. However, a remarkable feature of the fungal septins is the apparent diversity of functions, localizations and higher-order structures found in different cell types. In this chapter, we focus on septin function in four fungal species in which septins have been analysed in some depth. We begin with the fission yeast Schizosaccharomyces pombe, which grows exclusively as a unicellular, uninucleate yeast; then we consider the pathogen Candida albicans which switches between yeast, pseudohyphal, and true hyphal


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CH05 SEPTINS IN FOUR MODEL FUNGAL SYSTEMS: DIVERSITY IN FORM AND FUNCTION

Table 5.1

Summary of septin proteins in four fungi

Organism

Septins

Localization

Function

S. pombe

Spn1p-4p Spn5p-7p (sporulation specific)

Spn1-4 : Double ring (apparently derived from initial single ring) assembles at division plane in anaphase. Spn5-7: pro-spore membrane. Yeast cells and pseudohyphae: assemble at prebud site and form collar at mother-bud neck Hyphae: Band of septin ‘bars’ transiently present at base of germ tube (lacks Cdc3p), cap at hyphal tips, rings within germ tubes Always assembled as discrete, filamentous bars of variable length depending on maturity of septin ring. Appear at hyphal tips, along hyphae, and at the bases of lateral branches. AspB: localizes post-mitotically as a single ring at septation sites; matures to a double ring; then the ring on apical side of septa persists while ring on basal side disassembles at septation.

Efficient septation (non-essential). Orderly pro-spore membrane extension.

C. albicans

Cdc3p, 10p, 11p, 12p, Sep7p, Spr3p, Spr28p

A. gossypii

Cdc3p, 10p, 11ap, 11bp, 12p, Sep7p, Spr3p, Spr28p

A. nidulans

AspA, B, C, D, E

Cell separation, morphogenesis, exocytosis (Cdc3p and Cdc12p essential).

Mitosis control, sporulation, hyphal morphogenesis, septum formation (non-essential).

Chitin deposition in septa, branching pattern, conidiophore development (AspB non-essential).

morphologies; and we conclude with two fungi, Ashbya gossypii and Aspergillus nidulans, that grow exclusively as multinucleate filaments (summarized in Table 5.1). The connections and comparisons between these different species will illustrate the plasticity of form and function displayed by septins within the fungal kingdom.

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SCHIZOSACCHAROMYCES POMBE: A UNINUCLEATE FISSION YEAST S. pombe has rod-shaped cells formed through polarized growth at the cell ends followed by division in the middle of the cell. The molecular mechanisms driving this reproducible morphogenetic program have been studied intensively, and much detail is known about the basis for polarized growth and cytokinesis in S. pombe (Brunner and Nurse, 2000). Notably, fission yeast cells polarize growth and execute cytokinesis using mechanisms that appear to have some fundamental differences from those in the similarly well-studied S. cerevisiae, as might have been expected given their significant evolutionary divergence (on the order of 900 million years) (Heckman et al., 2001). Likewise, the higher-order structure, assembly, and function of septins in S. pombe share some striking similarities but also display many differences from those of S. cerevisiae. The S. pombe genome contains seven genes encoding septins. Spn1p, 2p, 3p and 4p are expressed in vegetatively growing cells, whereas Spn5p, 6p and 7p are expressed exclusively during sporulation where they (along with Spn2p but not Spn1p, Spn3p, or Spn4p) are important for the normal formation of the forespore membrane and spore wall (Longtine et al., 1996; Mata et al., 2002; Rustici et al., 2004; M. Onishi, Y. Fukui and J. Pringle, personal communication). Notably, there is no Shs1p/Sep7p orthologue apparent in the genome or present in septin complexes when purified from vegetatively grown cells (An et al., 2004). Intriguingly, Spn7p, which in overall sequence is most similar to septins in the Cdc11 group, appears to have lost all GTP-binding motifs and lacks any coiled-coil domain but nonetheless still seems to localize and function together with the other septins during sporulation (J. Pringle, personal communication), calling into question the absolute importance of these domains for septin function. Spn1p-4p share relatively low amino-acid-sequence identity with their orthologues in ¡i>S. cerevisiae¡/i>, which are ScCdc3p, 10p, 11p and 12p respectively (Longtine et al., 1996); strikingly, Spn2p, like Cdc10p, also lacks a predicted coiled-coil domain. Despite amino acid identities of only 42–52 %, an antibody raised against ScCdc3p recognized Spn1p, and some S. pombe septins can complement the function of S. cerevisiae septins inactivated by temperature-sensitive mutations (H.B. Kim. T. Pugh, and J. Pringle, personal communication). Purified Spn1p-4p assemble into a predicted linear array composed of subunits of homodimers of each septin protein associating in the order Spn3-4-1-2 (An et al., 2004), similar to the predicted arrangement of S. cerevisiae septins (Versele et al., 2004; Bertin et al., 2008) and mammalian septins (Sirajuddin et al., 2007). Although these data indicate some fundamental structural conservation in the septin assemblies, there are also some striking differences in the localization and function of septins in fission yeast compared to S. cerevisiae. In the latter, septins form a ring at the prebud site (the future site of cytokinesis) during late G1 of

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Nucleus Septin ring

Figure 5.1 Septin-ring assembly during the S. pombe cell cycle. The septin ring in S. pombe assembles in mitosis in an apparent single ring that then matures into a double ring upon the onset of anaphase. The double ring persists until cell separation

the cell cycle and arrive at this site before other components of the cytokinetic machinery such as actin and type II myosin (Kim, Haarer and Pringle, 1991; Bi et al., 1998; Iwase et al., 2006). In contrast, in S. pombe, the septins coalesce into a ring around the middle of the cell only in mitosis after many other cytokinesis proteins have already gathered at that site (Berlin, Paoletti and Chang, 2003; Tasto, Morrell and Gould, 2003; An et al., 2004); Figure 5.1). Notably, Spn1p-4p do not appear to be functionally equivalent, presumably reflecting their differential positions and roles in septin filament assembly (see above). Deletion of spn1 or spn4 leads to the loss of all organized septins at the division site; whereas in the absence of Spn2p or 3p, the remaining septins still mostly find the cell middle, although additional ectopic, septin assemblies appear in these mutants in other locations in the cell cortex, and the medial ring does not mature normally (An et al., 2004); (John Pringle, personal communication). Thus, Spn1p, 4p and at least one other vegetative septin protein are required for normal assembly and maturation of the ring. Although initial assembly may be as a single ring, it is soon apparent that the septins form a double ring that brackets the medial actomyosin ring and persists until septin-ring disassembly at cell separation (Berlin, Paoletti

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and Chang, 2003; An et al., 2004). The septin rings do not contract but instead maintain a constant diameter at all times while other cytokinetic factors invaginate within the domain bounded by the septin ring (Berlin, Paoletti and Chang, 2003); (J.Q. Wu and J. Pringle, personal communication). Moreover, fluorescence recovery after photobleaching (FRAP) of Spn4-GFP, suggests that the septin subunits are stably incorporated within the ring rather than rapidly exchanging with free subunits in the cytosol (Berlin, Paoletti and Chang, 2003). The septin ring retains its original diameter through cytokinesis in both S. cerevisiae and S. pombe, yet is essential in the former case but not in the latter. Thus, the ability of the septin structure to invaginate with the cleavage furrow (as seen in many animal cells) is not a reliable predictor of the essentiality of septins in cytokinesis. Interestingly, the initial accumulation of septins in the cell middle appears to be distinctly regulated from the later steps of ring organization and maturation to a double ring. The septins can congregate at the cortex in the middle of the cell in the absence of the protein Mid2p, an anaphase-specific protein that is weakly similar in sequence to the S. cerevisiae Bud4p (involved in bud-site selection and perhaps in septin organization at the bud site); however, Mid2p is required to form the compact septin rings (Berlin, Paoletti and Chang, 2003; Tasto, Morrell and Gould, 2003). In cells lacking Mid2p, the dynamics of the septin assembly also change dramatically such that there is a 30-fold increase in the exchangability of septins in the ring compared to the stable rings of wild-type cells (Berlin, Paoletti and Chang, 2003). Additionally, instead of the clear double ring, the septins assemble into a ‘disc’ or ‘washer’-like higher-order structure that seems to track the cleavage furrow rather than maintaining its dimensions as acquired at assembly (Berlin, Paoletti and Chang, 2003; Tasto, Morrell and Gould, 2003). Similarly, overexpression of Mid2p leads to persistence of the septin rings (Tasto, Morrell and Gould, 2003; An et al., 2004). Thus, fission yeast septin-ring organization and function late in mitosis are tightly regulated either directly or indirectly by Mid2p. Despite this carefully coordinated assembly and maturation of the septin rings at the site of cytokinesis, the septins are not essential for cytokinesis in fission yeast. Loss of Spn1p or Spn4p produces cells with a delay in cell separation that is more severe than that in cells lacking Spn2p or Spn3p, which only show defects under certain stress conditions (Longtine et al., 1996); (O. Al-Awar, M. Valencik and J. Pringle, personal communication). This difference in the severity of phenotype parallels the differential effects on septin localization as seen in these mutants and further supports the idea that not all septin subunits are functionally equivalent in fission yeast (An et al., 2004). The mild phenotypes are not due to functional redundancy among the septins, because even a sextuple mutant lacking Spn1p-5p and Spn7p has a phenotype indistinguishable from an spn1 or spn4 mutant (J.-Q. Wu, J. Bähler, and J. Pringle, personal communication). Septin mutants form normal primary and secondary septa based on transmission electron micrographs however hydrolytic enzymes (Eng1p and Agn1p) are mislocalized to a broad disk instead of a tight ring in spn4 mutants (Martin-Cuadrado et al., 2005). These data suggest that septin mutants may have

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defects in targeting the hydrolytic enzymes that help execute cell separation. The delay in cell separation leads to an appearance of chains of cells linked together with prominent but not separated septa. The extent of cell-separation delay varies from cell to cell and septum to septum in septin mutant populations. Based on genetic interactions between the S. pombe septin mutations and mutations effecting factors in the cell-wall-integrity pathway (which appears to be similar to the well-studied pathway in S. cerevisiae (Levin, 2005)), it has been hypothesized that the variable kinetics of cell separation may be due to variable rates of repair of partially defective septa formed in the absence of a normal septin assembly (J.Q. Wu and J. Pringle, personal communication).

CANDIDA ALBICANS: A POLYMORPHIC HUMAN PATHOGEN C. albicans is a fungus that is usually found growing harmlessly in the human gastrointestinal and genitourinary tract. However, it can also be an important pathogen, causing conditions that range from painful mucosal infections of the vagina in otherwise healthy women to severe, life-threatening, blood-stream infections among vulnerable groups such as newborns and certain intensive-care patients, especially those undergoing cancer chemotherapy, immunosuppressant therapy or catheterization (Kibbler et al., 2003). A striking feature of C. albicans pathogenesis and biology is its ability to switch among several morphological forms (Figure 5.2). These range from unicellular budding yeasts to true hyphae with parallel-sided walls. Between these two extremes, C. albicans can exhibit a variety of growth forms that are filamentous but retain a constriction between adjacent cellular compartments. These are

(a)

(b)

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Figure 5.2 Morphological forms of C. albicans. Pseudohyphae (a), yeast (b), hyphae (c). Like hyphae, pseudohyphal cells are polarized, but they have constrictions at the sites of septation (arrows), Hyphae are thinner than pseudohyphae, and there are no constrictions at sites of septation (arrowed and inset)

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collectively referred to as pseudohyphae (Sudbery, 2001; Berman and Sudbery, 2002; Sudbery, Gow and Berman, 2004). In yeast form cells, daughter buds separate from the mother cell after cytokinesis. In contrast, the daughters remain associated after cytokinesis in pseudohyphae, but they are readily separated by brief sonication or by the mild mechanical shearing forces present in shaken liquid cultures. In hyphae, the cellular compartments remain firmly attached to each other and are not separated even by vigorous shaking or prolonged sonication. The differences between yeast, pseudohyphae and hyphae have recently been reviewed in detail Sudbery, Gow and Berman, (2004). Notably, septins assemble into different higher-order structures depending on the cell type. Therefore, the same set of septin proteins can be differentially regulated and have different appearances and functions, depending upon cell fate.

Septin organization The C. albicans genome contains seven septin genes, CDC3, CDC10, CDC11, CDC12 , SEP7, SPR3 and SPR28 (Warenda and Konopka, 2002). The gene designations follow the S. cerevisiae nomenclature, where SEP7 is the homologue of S. cerevisiae SHS1/SEP7. The similarities with the S. cerevisiae homologues are 54 % for Cdc3p, 69 % for Cdc10p, 48 % for Cdc11p, 61 % for Cdc12p and 33 % for Sep7p. CDC3 and CDC12 are essential for viability, but cdc10/ and cdc11/ mutants are viable although they show conditional morphological and cell-wall defects (Warenda and Konopka, 2002). (C. albicans is an obligate diploid, hence both copies of a gene must be deleted to generate a null mutant.) The homology is lower in the case of SPR3 (34 %) and SPR28 (33 %). In S. cerevisiae, expression of SPR3 and SPR28 is meiosis-specific, and their function in C. albicans is unclear since meiosis has not yet been demonstrated in this organism. The organization of septin structures is different in yeast and pseudohyphae compared to hyphae, and this reflects a fundamental difference in the regulation of the cell cycle in these morphologies (Figure 5.3; Plate 5.1 [see p. 246 for Plates]) (Sudbery, 2001; Warenda and Konopka, 2002; Sudbery, Gow and Berman, 2004). In yeast and pseudohyphae, a septin patch forms at the site of bud evagination composed of Cdc3p, Cdc10p, Cdc11p and Cdc12p. As the bud forms, the septin complex remains at the neck, forming first a ring and then a collar covering the bud neck, similar to what occurs in S. cerevisiae. Mitosis takes place across the plane of the bud neck (Colour plate 5.1a and b). At cytokinesis the septin collar splits into two as it organizes the formation of the primary and secondary septa. A hyphal germ tube evaginates from a mother yeast cell before the Start of the cell cycle (Figure 5.3). Evagination is accompanied by the formation of a septin patch which remains at the germ-tube neck (Sudbery, 2001; Warenda and Konopka, 2002). However, this septin complex does not mature into a ring; rather, it forms a band of longitudinal septin bars (Figure 5.4; Colour plate 5.1c). This structure is known as the basal septin band (Berman and Sudbery, 2002) and is

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Yeast

Pseudohyphae

Hyphae

Septin collar

Basal septin band

Septin ring

Septin Cap

Nucleus

Primary septum

Figure 5.3 Septin organization in C. albicans yeast, pseudohyphae and hyphae. In yeast and pseudohyphae, a septin ring forms just prior to bud emergence. As the bud forms, the septins spread to form a collar or hourglass structure at the bud neck. Mitosis takes place across the plane of the septin collar, which splits into two during cytokinesis. In hyphae, a septin patch forms at the same time as, or just before, the emergence of the hyphal germ tube. As the germ tube elongates, the septin complex becomes a disorganized band of longitudinal bars its base and a cap of septins is present at the hyphal tip. As the cell cycle commences, a septin ring forms from the septin cap and remains in place as the germ tube continues to elongate while the basal septin band fades and disappears. At the end of the first cell cycle, the nucleus migrates out of the mother cell into the germ tube, and mitosis takes place across the septin ring. After anaphase, one nucleus migrates back into the mother cell. The septin ring then splits into two as the primary and secondary septa form

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(c)

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Figure 5.4 Formation of the septin ring during germ-tube emergence in C. albicans. Frames taken from a time-lapse movie. (a) A young germling displays a basal septin band and septin cap. (b) A faint ring forms from the cap. (c) The ring becomes brighter and the cap and basal band fade. (d). The septin ring remains stationary as the germ tube elongates, and the cap and basal band completely disappear (Images courtesy of Laura Jones)

similar to the structures seen at the base of S. cerevisiae mating projections (shmoos) (Ford and Pringle, 1991) and vegetative cells lacking Gin4p, Nap1p, or Cla4p (Longtine, Fares and Pringle, 1998; Longtine et al., 2000). In addition to the basal band, a septin cap is visible at the germ-tube tip (Figure 5.4; Colour plate 5.1c). As the germ tube elongates, a septin ring forms from this cap so that septin ring is present within the germ tube rather than at its neck (Figure 5.4, Colour plate 5.1d). This septin ring is not sharply defined and is probably equivalent to the septin collar at the base of a yeast or pseudohyphal bud (Colour plate 5.1d, inset). The basal band and septin cap disappear after the septin ring forms (Figure 5.4), possibly due to competition between the different structures for septin subunits. The band and cap persist longer in gin4/ mutants, which can form the basal septin band and cap but not the septin ring within the hypha (Wightman et al., 2004) (see below). The nucleus migrates out of the mother cell, mitosis takes place within the germ tube, and one daughter nucleus then migrates back into the mother cell (Colour plate 5.1d). After anaphase, the septin ring splits into two sharply defined rings and the primary septum forms between the two rings (Colour plate 5.1e).

Control of septin organization by the protein kinases Gin4p and Ccn1p-Cdc28p The role of the basal septin band in hyphal cells is unclear, but it is possible that it is involved in determining the shape of the germ-tube base, which lacks a constriction in contrast to that present between the mother cell and a bud in yeast or pseudohyphae. As noted above, the basal septin band resembles the septin array in S. cerevisiae mutants lacking Gin4p suggesting that the formation of the band does not require Gin4p. Indeed, in Gin4p-depleted hyphae, the basal band formed normally but the septin ring failed to assemble (Wightman et al., 2004). Thus, the basal band and septin ring have different genetic requirements for their formation and maintenance. Interestingly, in Gin4p-depleted cells, nuclear migration and mitosis occurred normally within the germ tube. Thus, the septin ring is not the

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spatial marker that determines where mitosis will occur. Furthermore, in wild-type hyphae the septin rings form before nuclear migration is initiated; therefore nuclear position cannot be the spatial cue for septin ring formation. Thus, there must be at least one independent spatial marker determining the location of the septin ring and position where mitosis will occur in C. albicans hyphae. What is the substrate of Gin4p that is crucial for septin-ring formation? Recent work has demonstrated that Gin4p can directly phosphorylate the septin Cdc11p at Ser395 (Sinha et al., 2007). This phosphorylated Cdc11p is now ‘primed’ to be a substrate for Cdc28p (a cyclin-dependent kinase) in complex with the Ccn1p cyclin, which phosphorylates the adjacent Ser394 residue. If phosphorylation of Ser394 is prevented, either by deletion of CCN1 or by a cdc11 S394A mutation, germ tubes evaginate normally and septins localize to a ring. However, after the septin ring forms, growth becomes isotropic, resulting in a swollen hyphal tip. Tip swelling in a ccn1/ strain is rescued by cdc11 S394D S395D phosphomimetic mutations, showing that phosphorylation of Cdc11p is the only Ccn1p function required for normal hyphal development. Cdc11p phosphorylation by Gin4p is a cell cycle regulated event occurring during the mitosis and cytokinesis phases of the previous cell cycle. Thus, Ser395 is already phosphorylated when unbudded yeast cells are induced to form hyphae. After hyphal induction, Ccn1p-Cdc28p associates with the septin complex within 5–10 minutes. This rapid response appears to be independent of the signal-transduction pathways that promote the hyphal-specific pattern of transcription, because it occurs in an efg1/ cph1/ mutant. This suggests that at least one early event of hyphal morphogenesis is independent of the hyphal-specific transcriptional response. Under the alias CLN1 , CCN1 had previously been shown to be essential for the maintenance but not the establishment of hyphal growth (Loeb et al., 1999), consistent with the conclusions of Sinha et al. (2007). However, it should be noted that phylogenetic analysis has shown that Ccn1p is not an orthologue of S. cerevisiae Cln1p and indeed is not a member of the S. cerevisiae G1 (Cln1-3p) or G2 (Clb1-6p) cyclin families (unpublished observations). The phosphorylation of Cdc11p does not appear to be important for septin organization but clearly impacts septin-ring function. How does the phosphorylation at early times after hyphal induction promote normal hyphal morphogenesis much later after the septin ring has formed? Polarized growth in fungi involves the directed flow of secretory vesicles to the site of growth to generate new cell wall and membrane upon exocytosis. Septins have been implicated in directing exocytosis in several systems (Beites et al., 1999; Beites, Peng and Trimble, 2001; Beites, Campbell and Trimble, 2005; Spiliotis and Nelson, 2006). It has been suggested that septins play a role in generating the characteristic hourglass shape of the bud neck of S. cerevisiae yeast cells by targeting secretion to the bud base, the resulting lateral growth producing the characteristic swelling on the daughter side of the neck (Gladfelter et al., 2005). Although C. albicans yeast cells have a similar hourglass shape at the bud neck, there is no constriction at the site of the septin ring in hyphae. Thus, the propensity of the septin ring to target secretion

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must be suppressed in hyphae. Perhaps phosphorylation of Ser395 of Cdc11p by Ccn1p-Cdc28p is a critical modification that limits septin–exocyst interactions. The exocycst is a multi-protein structure to which secretory vesicles dock before fusion with the plasma membrane. In S. cerevisiae, Sec3p acts as the spatial cue for the formation of the exocyst: localization of other exocyst components depends on Sec3p, but Sec3p localization is autonomous of the other components (Finger and Novick, 1997; Finger, Hughes and Novick, 1998). Support for a mechanism involving regulation of septin–exocyst associations in hyphae is provided by the observation that the exocyst component Sec3p co-immunoprecipitates with the septin Cdc11p in C. albicans (Li et al., 2007). Furthermore, sec3/ mutants could initiate apparently normal germ tubes, but when the septin ring formed, growth became isotropic and the tip became swollen – a phenotype reminiscent of that of the ccn1/ and cdc11 S394A mutants. However, this tip swelling did not occur in a cdc10/ sec3/ double mutant (Li et al., 2007). A model that accomodates all of these data is as follows. At early stages in hyphal development, growth is directed towards the tip. When the septin ring forms, continued polarized growth depends on tip localization of Sec3p. However, Sec3p affinty for Cdc11p in the ring complex leads to competition for Sec3p between the tip and the septin ring. Phosphorylation of Cdc11p Ser394, which depends on the formation of a septin complex that includes Cdc10p, weakens the affinity of Sec3p for Cdc11p. Thus, the tip localization of Sec3p dominates, thereby promoting polarized growth at the tip at the expense of lateral growth directed from the septin ring.

Control of septin organization by Cdc42p GDP/GTP cycling In S. cerevisiae, the Cdc42p Rho-type guanosine triphosphatase (GTPase) controls many aspects of morphogenesis including the formation of the septin ring at the bud neck (Johnson, 1999; Park and Bi, 2007). Like other GTPases, Cdc42p cycles between GTP- and guanosine diphosphate (GDP)-bound forms. Hydrolysis of GTP is enhanced by guanosine triphosphatase-activating proteins (GAPs) that bring Cdc42p to the GDP-bound form. The C. albicans genome encodes two Cdc42 GAPs, Rga2p and Bem3p (named by homology to their S. cerevisiae orthologues). Because the GTPases are normally active in the GTP-bound form, Cdc42p is expected to be hyperactive in cells lacking Rga2p or Bem3p. In rga2/ bem3/ cells grown under pseudohyphal-promoting conditions, the septin ring was found to form within the germ tube instead of at the bud neck, suggesting that the position of the septin ring is determined by the level of Cdc42p activation (Court and Sudbery, 2007). Furthermore, such cells also assumed a more hyphal-like shape, and a Spitzenkörper (a cluster of vesicles characteristic of the tips of hyphae) was present, indicating that Cdc42p also promoted the extreme polarized growth characteristic of hyphae. However, when Cdc42p was locked into the GTP-bound form by the conditional expression of a cdc42 G12V allele, cells grown in pseudohyphal-promoting conditions were swollen, and the septins formed prominent bars at the bud neck. The differences in phenotype between

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cells with Cdc42p activated either through loss of the GAPs or through being locked in the GTP-bound state suggest that Cdc42p cycling between GDP- and GTP-bound forms is required for the maturation of the septin collar. This conclusion is consistent with studies in S. cerevisiae that suggest that formation of the septin ring is a two-stage process: Cdc42p-GTP recruits septin subunits to form a patch, but maturation into a collar requires Cdc42p cycling between the GDP- and GTP-bound forms together with the action of Gin4p, Nap1p, Cla4p, Gic1p and Gic2p (Gladfelter et al., 2002; Caviston et al., 2003; Gladfelter, Zyla and Lew, 2004; Iwase et al., 2006). In C. albicans, germ-tube evagination from the mother cell occurs before the cell cycle is initiated (Hazan, Sepulveda-Becerra and Liu, 2002). Presumably, Cdc42p-GTP at the site of evagination will recruit septin subunits, but in the absence of Rga2p and Bem3p these subunits do not mature into a collar. When the cell cycle is initiated, the expression of RGA2 and BEM3 facilitates Cdc42 GDP/GTP cycling to promote maturation of the septin subunits into the collar that forms within the growing germ tube. In summary, septins clearly play important roles in generating the different morphological forms of C. albicans. As in S. cerevisiae, they organize the formation of primary and secondary septa during cytokinesis and cell cycle regulated phosphorylation may modify the septin ring in hyphae to prevent it from programming the lateral growth that produces the constriction in yeast and pseudohyphal buds.

Sep7 modifies the septin ring in hyphae to abrogate the cell-separation program Cell separation occurs after cytokinesis in C. albicans yeast and pseudohyphae, but not in hyphae. In the yeast form, Cdc14p localizes to the site of septation and is required for the localization and activation of the Ace2p-dependent transcriptional program that results in the synthesis of hydrolytic enzymes that will destroy the primary septum (Clemente-Blanco et al., 2006). In contrast, Cdc14p does not localize to the septation site in hyphae and the resulting absence of the Ace2p-dependent transcription program explains the failure of hyphal compartments to separate (Clemente-Blanco et al., 2006). FRAP experiments show that the septin subunits Cdc3p, Cdc12p and Sep7 are stable in both hyphae and yeast, that is, once the septin ring forms; it does not exchange subunits with the free cytoplasmic pool. However, in hyphae, but not yeast, the septin Cdc10p does exchange subunits with the cytoplasmic pool (Gonzalez-Novo et al., 2008). This dynamic property of Cdc10p in hyphae depends on the septin Sep7p. In sep7/ mutants, the septin ring assumes the properties of the yeast ring – Cdc10p in the septin ring is stabilized, Cdc14p is recruited and adjacent compartments separate after cytokinesis. The hyphal properties of the septin ring also depend on Hgc1p, a G1 cyclin that is only expressed in hyphae. The phosphorylation pattern of Sep7p is different, and total Sep7p levels are reduced in an hgc1/ mutant. This suggests that Hgc1p-Cdc28p phosphorylates and stabilizes Sep7p, allowing it to modify the

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properties of the septin ring to abrogate the cell-separation program. This is one of the first examples in fungi where the post-translational modification of a single septin subunit has been shown to change both the dynamics and function of the septin ring in a normal developmental program.

ASHBYA GOSSYPII: A MULTINUCLEATE, FILAMENTOUS HEMIASCOMYCETE A. gossypii is an ascomycete that unlike C. albicans does not switch between different morphologies but rather is constitutively filamentous. Based on their genome sequences, A. gossypii is predicted to have diverged from a common ancestor with S. cerevisiae approximately 100 million years ago. About 90 % of the A. gossypii genome is homologous and syntenic to S. cerevisiae (Dietrich et al., 2004) so that A. gossypii genes are named according to their syntenic homologues in S. cerevisiae. While the overall sets of proteins are similar, the sequence identity between orthologous proteins is often less than 50 %, consistent with the observed divergence in morphology and lifestyles between the two organisms. A. gossypii grows exclusively as filaments in which nuclear division occurs in the absence of cytokinesis, thereby producing large syncytia in which many nuclei share a common cytoplasm (Gladfelter, Hungerbuehler and Philippsen, 2006). It has never been observed to form uninucleate yeast in which a single nuclear division is coupled to a single cell division. Thus, A. gossypii is an excellent model for understanding septin functions unrelated to cytokinesis. Eight septin genes are found in the A. gossypii genome, including homologues to all of the S. cerevisiae vegetative and sporulation-specific septins. An additional CDC11 (called b) gene lies adjacent to the ancestoral CDC11 (called a) and is probably a product of a tandem duplication occurring after the divergence from S. cerevisiae (Dietrich et al., 2004). Preliminary expression data suggest that CDC11b may only be expressed under specific growth conditions (Peter Philippsen, personal communication). Identity between the S. cerevisiae and A. gossypii septins ranges from 58 % (for Cdc3p) to 78 % (for Cdc12p), and typical septin domains including the GTP-binding domain, predicted phosphoinositidebinding elements and coiled-coil domains in the C-termini are clearly identifiable in the A. gossypii sequences.

Morphologically diverse septin organizations assemble within one cell Septins assemble and mature through morphologically distinct cortical organizations in A. gossypii hyphal cells (Figure 5.5, Colour plate 5.2). Septins initially assemble into a loose, elongated band of filaments at the growing tips of hyphae (Colour plate 5.2, panels a and b). Upon an unidentified signal, this band coalesces into a more compact and apparently organized ring that becomes fixed to

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1

2

5

3

Spore

4 Nucleus Septin rings

Figure 5.5 Morphologically diverse septin organizations assemble in A. gossypii. In the cortex of a single, multinucleate A. gossypii cell septins appear in a variety of different organizations. Diffuse filamentous septins appear at the growing tips of hyphae (1) and then the filaments compress into more compact rings made of bars of septin protein (2). The consolidation of the septin filament structure is accompanied by an ‘anchoring’ of the ring at the cortex and detachment from the growing tip. The bars of septin protein persist for variable times before the bars elongate and the single ring transforms into two adjacent rings that persist (3). Septins also congregate in a cap at emerging branches (4); this pool of protein remains at the cortex at the base of the branch, forming a ring made of bars. This ring then appears to split into two rings while the growing branch tip assembles new septin filaments (5). These two rings at the branch base are asymmetric, with very faint septin bars visible at the side of the main hyphae and more compact and bright bars of septins on the branch side (5). (Schematic based on observations of Sep7p-GFP in living cells and Cdc11p by immunofluorescence; see Colour plate 5.2)

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one location on the cortex while tip growth continues (Colour plate 5.2, panels c and d). Unlike the smooth septin collars observed to date in other fungi, this newly ‘anchored’ ring remains in a filamentous organization of discrete short bars that dot the cortex, reminiscent of the transient basal septin band seen in C. albicans germlings (Figures 5.4 and 5.5, Colour plate 5.2, panels c, d, f and g). Meanwhile, the growing tip assembles a new band of filaments. In some cases, the bars split when a septum is built and this doublet ring then persist. Diffuse clouds of short septin filaments begin to coalesce prior to branch emergence on the sides of hyphae, and these organize into a ring as branch hyphae emerge (Colour plate 5.2, panels e and f). These rings persist at the base of the branch and develop an asymmetric organization, while new filaments assemble at the growing branch tips (Colour plate 5.2, panels g and h). In any single septin mutant (cdc3, cdc10, cdc11a or cdc12), Sep7p does not assemble into any of these organized structures (Helfer and Gladfelter, 2006); (B. DeMay and A. Gladfelter, unpublished data). Septin-ring assembly and maturation occur simultaneously in multiple spots on the cortex of A. gossypii hyphae. Thus, a common cytoplasm underlies these apparently independent septin-organization events. Current questions about septin localization and organization that are under investigation in this system include: is there a maturation program built into the organization of septin rings that essentially acts as a timer to execute the changes in appearance without additional input? Do the rings respond to internal and/or external cues to modulate their organization? What are the differences in septin-subunit composition and dynamics of the different organizations? What regulatory proteins act on the septin cortex to direct changes in organization? Do these regulatory factors respond to signals from the nuclear division cycle? Answers to these questions will greatly expand our understanding of how different molecular signals are translated into diverse higher-order septin structures.

Septin function in spatial control of mitosis Although deletion of any septin gene except CDC11b perturbs assembly of an organized septin ring, A. gossypii cells lacking any one septin are viable and show only a 30 % reduction in radial growth rate (Helfer and Gladfelter, 2006). Hyphal morphology is somewhat aberrant in cells lacking septins, and the phenotypes of septin null mutants are especially pronounced in mature mycelia, in which hyphae become kinked and ‘wavy’ and the incomplete septa that normally form between multinucleate compartments are absent. In addition, deletion of CDC3, CDC10, CDC11a or CDC12 led to sporulation defects suggesting these septins contribute to normal spore morphogenesis (H. Helfer, P. Philippsen and A. Gladfelter, unpublished data). The viable phenotypes of septin mutants, combined with a cell cycle lacking cytokinesis, make A. gossypii a powerful organism for uncovering novel cellular roles for septins.

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One novel function of the septins in A. gossypii has emerged from studies of the spatial control of mitosis in these hyphal cells. In multinucleate hyphae, mitosis is asynchronous such that neighbouring nuclei divide independently despite residing in a common cytoplasm (Gladfelter, 2006). Notably, however, the locations where nuclei most frequently divide do not appear to be random. In particular, it was shown recently that the septin cortex can direct the position of mitosis such that nuclear division preferentially occurs near assembled septin rings, particularly at branch points (Helfer and Gladfelter, 2006). In wild-type cells, 45 % of mitoses occur near branch points, whereas in septin mutants only 12 % of mitoses occur near branches and instead nuclei appear to divide randomly throughout hyphae independent of morphological features. How does the septin cortex create mitosis-promoting zones within the continuous cytoplasm of hyphae? Genetic evidence and localization data support a model in which the septins locally reduce the activity of the cyclin-dependent kinase antagonist, Swe1p (Wee1 homologue) by concentrating its negative regulators (Helfer and Gladfelter, 2006). This localized inhibition of Swe1p then favours division of nuclei in the vicinity of the septin cortex. This signalling network of septins-Swe1p-regulators and Swe1p is also responsive to external nutrient status, and we hypothesize that the septins therefore integrate both internal and external signals to promote localized mitoses. The septins enable A. gossypii cells to control precisely where new nuclei are made, which may be key for responding to localized nutrient gradients in the natural environment. Thus, in this system, the septin cortex links the cell interior and exterior by transmitting and responding to a variety of signals. In summary, A. gossypii , despite its close relationship on the genome level to S. cerevisiae, shows distinct septin organizations and seemingly unique septin functions. These features of the septin cortex probably reflect the multinucleate, hyphal architecture of these cells. Not only do these studies show how different septins assemblies can appear between species but they also indicate how diverse higher-order septin structures can assemble and mature within a single cell. Future analyses of the bases for the varied septin rings in A. gossypii hold much promise for understanding how specific post-translational modifications to septins lead to changes in septin organization.

ASPERGILLUS NIDULANS: A TRUE FILAMENTOUS ASCOMYCETE A. nidulans is a filamentous ascomycete that shows diverse morphologies during vegetative growth and the formation of asexual spores by conidiation (Harris and Momany, 2004). After germination of a spore, several rounds of mitosis occur while a hyphal germ tube emerges by polarized growth. The first septation site is formed after the third mitosis, and as the hypha grows, compartments containing 2–6 nuclei will be generated that are demarcated by additional septa. Only the

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compartments at the tips or from which branches emerge contain mitotically active nuclei. Powerful genetics and well-developed cell biological tools have allowed A. nidulans to make major contributions to understanding of the eukaryotic cell cycle, polarized growth, septation and metabolism. Analysis of the septins in A. nidulans has begun to impact nearly all of these diverse areas of cell biology, and the unique morphology of A. nidulans has placed it in a central position for understanding septin function outside of the uninucleate yeasts. A. nidulans has five genes encoding septins, which correspond to the core vegetative septins in S. cerevisae plus an additional septin that is unique to other filamentous fungi although is not present in A. gossypii. The septins (and their orthologues) are named AspA (Cdc11p), AspB (Cdc3p), AspC (Cdc12p), AspD (Cdc10p) and AspE (filamentous-fungus specific) (Lindsey and Momany, 2006; Pan, Malmberg and Momany, 2007).

Localization and function AspB has been most intensively investigated for localization and function (Momany and Hamer, 1996; Westfall and Momany, 2002). AspB was shown to localize to newly forming septa as a single ring that colocalizes with the actin ring (Figure 5.6). As in S. pombe, the AspB ring does not contract but maintains a constant diameter over time. As the septum matures, two rings of AspB become visible, and then only a single ring on the apical (growth tip) side of the septum persists while the ring on the basal side disassembles. Thus, intriguingly, the septin rings react to the overall spatial organization of these cells and display different patterns of organization relative to the axis of polarity. Based on analysis of nuclear-cycle stages in the apical compartment, AspB ring assembly appears to occur post-mitotically. In contrast, AspB assembles into rings that ‘anticipate’ the sites of branching in subapical compartments, before the onset of mitosis. Thus, depending on the morphogenetic context, septin-ring assembly can be either preor post-mitotic in A. nidulans (Figure 5.6). Although AspB was initially thought to be essential, cells lacking this protein are viable, although they grow slowly (R. Lindsey and M. Momany, personal communication). aspB deletion mutants and mutants harbouring a conditional allele of aspB are hyperbranching, form septa deficient in chitin, and have aberrant conidiophores (Momany and Hamer, 1996; Westfall and Momany, 2002). Thus, even in these multinucleate cells in which a single cell division is not coordinated with a single nuclear division, septins are integral to normal growth and morphogenesis.

CONCLUSIONS Studies to date in these four morphologically diverse fungi have demonstrated a tantalizing spectrum of septin organizations and functions. One unifying principle is that septins assemble where septa form. However, notable differences

142


4

3

2

1 Nucleus arrested in interphase (basal) Post-mitotic nucleus (apical) 5 Septin rings

Chitin ring, mature septum

Figure 5.6 Septin rings assemble post-mitotically in the apical compartment and pre-mitotically at branching sites in A. nidulans. The thick disk (1) represents a mature septum that separates the growing apical compartment from the basal compartments and this lacks any septin proteins. The ring adjacent to this (2) is the youngest septum in the growing apical compartment and the septins are shown here as they initially assemble as a single ring. This single septin ring splits into a double ring as the septum matures (3). This double ring then is dismantled asymmetrically, so that only the half of the ring closest to the tip persists, while the more basal ring dissipates (shown as the discontinuous ring in 4). Septin rings also assemble at sites of lateral branch emergence(s) before the nuclei in this compartment re-enter the cell cycle (5). (From data in Westfall and Momany, 2002)

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are seen in how the assembly of septin rings coordinates with the timing of the cell cycle, and in some cases, in how septins actually influence the timing of the nuclear cycle. The exact higher-order structures vary both between species and also between single cell types or even single cells of the same species. This suggests a tremendous regulatory flexibility in how the septin subunits may interact. Furthermore, whether or not the septins are essential varies from organism to organism, and this does not simply reflect whether the cells are uninucleate or filamentous or if the septin assemblage contracts or remains a fixed diameter at cell division. Finally, the septins contribute to diverse processes in these different fungi, including regulation of the timing of mitosis, the polarized growth of hyphae, hyphal morphology and the formation of spores. Thus, work in these four diverse fungi demonstrates that septin function varies as much as their forms of assembly and each system holds great promise for addressing the mechanisms of septin function and organization.

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Court, H. and Sudbery, P. (2007) Regulation of Cdc42 GTPase activity in the formation of hyphae in Candida albicans. Molecular Biology of the Cell , 18, 265–81. Dietrich, F.S., Voegeli, S., Brachat, S. et al. (2004) The Ashbya gossypii genome as a tool for mapping the ancient Saccharomyces cerevisiae genome. Science, 304, 304–7. Finger, F.P., Hughes, T.E. and Novick, P. (1998) Sec3p is a spatial landmark for polarized secretion in budding yeast. Cell , 92, 559–71. Finger, F.P. and Novick, P. (1997) Sec3p is involved in secretion and morphogenesis in Saccharomyces cerevisiae. Molecular Biology of the Cell, 8, 647–62. Ford, S.K. and Pringle, J.R. (1991) Cellular morphogenesis in the Saccharomyces cerevisiae cell cycle: localization of the CDC11 gene product and the timing of events at the budding site. Developmental Genetics, 12, 281–92. Gladfelter, A.S. (2006) Nuclear anarchy: mitosis in multinucleated cells. Current Opinion in Microbiology, 9 (6), 547–52. Gladfelter, A.S., Bose, I., Zyla, T.R. et al. (2002) Septin ring assembly involves cycles of GTP loading and hydrolysis by Cdc42p. Journal of Cell Biology, 156, 315–26. Gladfelter, A.S., Hungerbuehler, A.K. and Philippsen, P. (2006) Asynchronous nuclear division cycles in multinucleated cells. Journal of Cell Biology, 172, 347–62. Gladfelter, A.S., Kozubowski, L., Zyla, T.R. and Lew, D.J. (2005) Interplay between septin organization, cell cycle and cell shape in yeast. Journal of Cell Science, 118, 1617–28. Gladfelter, A.S., Zyla, T.R. and Lew, D.J. (2004) Genetic interactions among regulators of septin organization. Eukaryotic Cell , 3, 847–54. Gonzalez-Novo, A., Correa-Bordes, J., Labrador, L., Sánchez, M., Vázquez de Aldana, C.R., Jiménez, J. (2008) Sep7 is essential to modify septin ring dynamics and inhibit cell separation during Candida albicans hyphal growth. Molecular Biology of the Cell , 19 (4), 1509–18. Harris, S.D. and Momany, M. (2004) Polarity in filamentous fungi: moving beyond the yeast paradigm. Fungal Genetics and Biology, 41, 391–400. Hartwell, L.H. (1971) Genetic control of the cell division cycle in yeast. IV. Genes controlling bud emergence and cytokinesis. Experimental Cell Research, 69, 265–76. Hazan, I., Sepulveda-Becerra, M. and Liu, H. (2002) Hyphal elongation is regulated independently of cell cycle in Candida albicans. Molecular Biology of the Cell , 13, 134–45. Heckman, D.S., Geiser, D.M., Eidell, B.R. et al. (2001) Molecular evidence for the early colonization of land by fungi and plants. Science, 293, 1129–33. Helfer, H. and Gladfelter, A.S. (2006) AgSwe1p regulates mitosis in response to morphogenesis and nutrients in multinucleated Ashbya gossypii cells. Molecular Biology of the Cell , 17, 4494–512. Iwase, M., Luo, J., Nagaraj, S. et al. (2006) Role of a Cdc42p effector pathway in recruitment of the yeast septins to the presumptive bud site. Molecular Biology of the Cell , 17, 1110–25. Johnson, D.I. (1999) Cdc42: an essential Rho-type GTPase controlling eukaryotic cell polarity. Microbiology and Molecular Biology Reviews, 63, 54–105. Kibbler, C.C., Seaton, S., Barnes, R.A. et al. (2003) Management and outcome of bloodstream infections due to Candida species in England and Wales. Journal of Hospital Infection, 54, 18–24.

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6 Septins in the metazoan model systems Drosophila melanogaster and Caenorhabditis elegans Christine M. Field Department of Systems Biology, Harvard Medical School, Boston, MA 02115, USA

Amy Shaub Maddox Institute for Research in Immunology and Cancer, Department of Pathology and Cell Biology, University of Montréal, Montréal H3C 3J7, Canada

John R. Pringle Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305, USA

Karen Oegema Ludwig Institute for Cancer Research, Department of Cellular and Molecular Medicine, University of California, San Diego, CA 92093, USA

INTRODUCTION The fruit fly Drosophila melanogaster and the nematode worm Caenorhabditis elegans have made enormous contributions to genetics, cell biology, and developmental biology. The key role of these organisms has resulted from their numerous experimental advantages, including particularly their tractability for genetic analysis. It also reflects the intensive attention that has been focused on these organisms over the past 100 (Drosophila) or 30 (C. elegans) years; this attention has led to the development both of broad and deep knowledge of the biology of the organisms and of powerful methodologies for further study. In the hope (and The Septins Edited by Peter A. Hall, S.E. Hilary Russell and John R. Pringle © 2008 John Wiley & Sons, Ltd. ISBN: 978-0-470-51969-1

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CH06 METAZOAN MODEL SYSTEMS

expectation) that these metazoan model systems will prove equally important in the further elucidation of the properties and biological roles of the septins, we have attempted to summarize here what is currently known of the septins in these organisms.

THE GENE AND PROTEIN FAMILIES Drosophila has five known septins, Pnut, Sep1, Sep2, Sep4, and Sep5, as summarized in Table 6.1 (The ‘Sep3’ position was left free for Pnut, but that name now seems unlikely to change.) pnut was found serendipitously in the course of a genetic analysis of photoreceptor development (Neufeld and Rubin, 1994). (Because the gene was originally identified by a recessive mutation, Drosophila convention dictates that its symbol begins with a lower-case letter.) Sep1 and Sep2 were found in deliberate searches by PCR using degenerate primers based on the sequences of previously known septins (Fares, Peifer and Pringle, 1995; Al-Awar, 1996; Longtine et al., 1996; Field et al., 1996; see also Chapter 1 for more historical context). For Sep2 , a PCR product amplified from genomic DNA was used to recover cDNA clones, which were sequenced (GenBank entry U28966, June 1995); the sequences matched the coding sequence predicted from the subsequently released genome sequence. Sep4 was identified independently by A. Maeland and N. Brown and by J. Kramer and S. Hawley in 1999 (personal communications); each group noticed a septin-encoding gene very near the site of a mutationally defined gene that it was trying to clone (see Kramer and Hawley, 2003; Estrada et al., 2007), as noted first in FlyBase in 1999. At least one cDNA was sequenced and matched the coding sequence predicted from the genome sequence, but contemporary annotation of the databases appears to have been absent or incomplete, so that Sep4 had to be ‘rediscovered’ by Muñoz-Soriano and Paricio (2007). Sep5 was found by searching the expressed sequence tag (EST) sequences released by the Berkeley Drosophila Genome Project. Full sequencing of one of these clones (LD12056) revealed what appeared to be a full-length septin gene (K. Hales, C. Brown, J. Pringle, and M. Peifer, unpublished results; GenBank entry AF167578, July 1999); this sequence matched the coding sequence predicted from the genome sequence. Because some significant regions of the Drosophila genome remain unsequenced (notably in the heterochromatic regions), it remains possible that an additional septin gene(s) will be found there. Remarkably, given the larger number of septins in every other animal and fungal species that has been examined carefully, C. elegans appears to have just two septins, UNC-59 and UNC-61 (Nguyen et al., 2000; Table 6.1). The genes encoding these proteins were identified by a deliberate search of C. elegans EST and genomic sequences for septin-like genes, but they proved to correspond to unc-59 and unc-61 , which had been defined (genetically) much earlier by the isolation and analysis of uncoordinated mutants (White, Horvitz and Sulston, 1982).

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149

BIOCHEMISTRY OF THE DROSOPHILA AND C. ELEGANS SEPTINS Studies in both Drosophila and C. elegans have been central to our current understanding of septin biochemistry.

Isolation of a three-septin complex from Drosophila embryos Co-immunoprecipitation and density-gradient centrifugation experiments had suggested that Sep1 and Pnut formed a complex and that this complex probably contained additional polypeptides (Fares, Peifer and Pringle, 1995). These conclusions were confirmed and greatly extended when Field et al. (1996) achieved the first purification of a septin complex. This complex was isolated in a single step from Drosophila embryo extracts by binding to a polyclonal antibody, directed against the C-terminal 14 amino acids of Pnut, that had been immobilized on a resin, and then eluting with a buffer containing the peptide against which the antibody had been made. The isolated complex contained three-septin polypeptides, Pnut, Sep1 and Sep2, in a stoichiometry of ∼1 : 1 : 1. Analysis by gel filtration and density-gradient centrifugation indicated that the complex had a Stokes radius of 9.9 nm and sedimented at ∼8 S, suggesting a native molecular mass of ∼340 kDa. The large Stokes radius relative to the molecular weight indicated that the complex was highly asymmetric. Given the predicted molecular weights of the three septins and the stoichiometry of the complex, it was proposed to contain two copies of each polypeptide (Field et al., 1996). It is interesting that neither Sep4 nor Sep5 appeared to be present in the isolated septin complexes, particularly given that Sep4, at least, appears to co-localize with other septins in the cleavage furrows of at least some cells (Table 6.1, footnote i). Similarly, although Shih et al. (2002) observed multiple interactions of Pnut, Sep1, and Sep2 with each other during two-hybrid screening, as well as interactions of these three proteins with various other proteins, neither Sep4 nor Sep5 was recovered. It is possible that the failure to detect Sep4 and Sep5 reflects low levels of expression, or expression only in certain cell types (as has been shown for mammalian septins: Hall et al., 2005; see also Chapters 7, 8 and 14), and thus poor representation in the extracts and cDNA libraries used. Further exploration of this issue should be very interesting.

Binding, slow exchange, and hydrolysis of GTP by the isolated septin complex Nearly all known septins contain a predicted nucleotide-binding sequence, or P-loop (Saraste, Sibbald and Wittinghofer, 1990), and additional consensus sequences that define the GTPase superfamily (Bourne, Sanders and

FBgn0026351 CG2916

Sep5

1B

2B

CC

CC

Cellularization, cytokinesis (L).g

At least some Cytokinesis (L).i cleavage furrows, various cell-cortex regions.i No information. No information.

FBgn0026362 CG9699

Sep4

1B

PB, CC

NT,h PB, CC

FBgn0014029 CG4173

Sep2

2B

FBgn0011710 CG1403

Sep1

Mutants

None reported.

None reported.

sep2

pnut Cellularization, cytokinesis (L,M)e . Nervous system (L,M). Cellularization, None reported. cytokinesis (L).

Functionc

Cleavage furrows, some intercellular bridges, neurons, various cell-cortex regions. Cleavage furrows, some intercellular bridges, neurons, various cell-cortex regions. Cleavage furrows, some intercellular bridges, neurons, various cell-cortex regions.g

2B

FBgn0013726 CG8705

Pnut

Localization

NT, PB, CC

Classa Featuresb

Properties of fly and worm septin genes and proteins

Protein FlyBase or WormBase ID

Table 6.1 Referencesd

73 % identical to Sep2.

73 % identical to Sep5. Sep2-GFP-expressing flies available. Can localize in some (but not all) contexts even when Pnut is absent.

Fails to localize when Pnut is absent

9

14, 15

3–10, 22

2–8, 10, 13f

Zygotic lethal when 1–8, 10–12, 22 homozygous; germ-line clones have been analyzed.

Comments

150 CH06 METAZOAN MODEL SYSTEMS

WBGene00006795

UNC-61

1B

2B

Cortical contractile patches during polarity establishment. Contractile rings during polar body extrusion and cytokinesis. Cortex of cellularizing gonad. Same as UNC-59

PB, CC

NT,h CC

Asymmetric furrow ingression during early embryonic cytokinesis (R). Increases robustness of embryonic cytokinesis (R). Post-embryonic cytokinesis (M). Neuronal migration, distal-tip-cell migration (M). Same as UNC-59 e228 and n3169 (point mutations)

e261 and e1005 (premature stops)

17–21

16–21

b

As defined by Pan, Malmberg and Momany (2007); see also Chapter 2. All seven proteins have the signature motifs of a GTP-binding site, and Pnut, Sep1, and Sep2 have been shown biochemically to bind GTP. NT, an extended N-terminus relative to other septins; PB, a cluster of three (UNC-59), four (Pnut), or five (Sep1 and Sep4) basic residues lying just N-terminal to the GTP-binding domain that may be involved in phosphoinositide binding (see Chapter 3); CC, predicted coiled-coil domain. c Evidence for function can be derived from localization (L), mutant phenotypes (M), and/or RNAi (R). d (1) Neufeld and Rubin, 1994; (2) Fares, Peifer and Pringle, 1995; (3) Longtine et al., 1996; (4) Field et al., 1996; (5) Hime, Brill and Fuller, 1996; (6) Al-Awar, 1996; (7) Adam, Pringle and Peifer, 2000; (8) Sisson et al., 2000; (9) Hales, Peifer and Pringle, 2000; (10) Shih et al., 2002; (11) Somma et al., 2002; (12) Field et al., 2005; (13) Maleszka, de Couet and Miklos, 1998; (14) Kramer and Hawley, 2003; (15) Muñoz-Soriano and Paricio, 2007; (16) White, Horvitz and Sulston, 1982; (17) Nguyen et al., 2000; (18) Finger, Kopish and White, 2003; (19) Maddox et al., 2005; (20) Maddox et al., 2007; (21) John et al., 2007; (22) Echard et al., 2004.

a

WBGene00006793

UNC-59

BIOCHEMISTRY OF THE DROSOPHILA AND C. ELEGANS SEPTINS 151

(continued )

f Considerable

text for discussion of the apparent nonuniformity of the Pnut requirement for cytokinesis. information about the cytogenetics (the endpoints of relevant deletions, etc.) of the Sep1 region is available in reference 13 and earlier publications from the Miklos group that are cited therein. However, it does not appear possible to deduce the phenotype (if any) resulting from loss of Sep1 (referred to as innocent bystander in reference 13) from any of the published information. See Chapter 1 for a summary of the historical context. g Reflecting the similarity in sequence between Sep2 and Sep5, the anti-Sep2 antibodies used in both biochemical (Field et al., 1996; Al-Awar, 1996) and protein-localization (Al-Awar, 1996; Adam, Pringle and Peifer, 2000; Figure 1c) studies were found later to interact weakly with Sep5. However, multiple lines of evidence, including peptide sequencing of immunoprecipitated protein and corroboration of many central immunofluorescence results with observations on flies expressing Sep2-GFP (Figure 1a and b), suggest that the conclusions reached with respect to Sep2 are valid (Field et al., 1996; J. Adam, K. Hales, G. Dillard, H.-P. Shih, O. Al-Awar, M. Peifer and J. Pringle, unpublished results). h The N-terminal extensions on Sep4 and UNC-61 are considerably shorter than those on some septins such as Pnut and S. cerevisiae Cdc3p. i In immunoblots, the antibody generated against Sep4 also recognized a second protein that was expressed throughout embryogenesis and in some adult tissues (C. Field, J. Adam and K. Oegema, unpublished results), precluding the successful use of this antibody in immunofluorescence experiments on embryos. However, this cross-reacting protein did not appear to be expressed in adult ovaries, where staining of both cleavage furrows (in dividing germline stem cells) and the cortical regions of germline cells was observed (J. Adam, C. Field, J. Pringle and M. Peifer, unpublished results). Interestingly, the cortical regions of follicle cells, which are rich in Pnut, Sep1, and Sep2 (Fares, Peifer and Pringle, 1995; J. Adam, C. Field, K. Oegema, J. Pringle and M. Peifer, unpublished results), did not stain with the Sep4 antibodies.

e See

Table 6.1

152 CH06 METAZOAN MODEL SYSTEMS


153

McCormick, 1991). Thus, septins were predicted to bind and perhaps hydrolyse GTP. GTP binding was confirmed for the isolated Drosophila complex by using HPLC ion-exchange chromatography to analyse the nucleotide released following denaturation of the complex with urea (Field et al., 1996). On average, 1.1 mol of guanine nucleotide was released per mole of septin polypeptide (corresponding to ∼6 per native complex), with an average GDP/GTP ratio of 2.6. These data suggested that each septin polypeptide in the complex binds one molecule of guanine nucleotide and also indicated tight binding, given that the isolation procedure involved extensive washes in nucleotide-free buffer. Subsequently, 1 : 1 binding of guanine nucleotide to septin polypeptide has been observed for complexes isolated from all other species examined, with the possible exception of the bacterially expressed C. elegans complex (see below). The ability of the purified Drosophila septins to exchange and hydrolyse GTP was also measured. In experiments in which [α-32 P]GTP was added to septin complexes attached to antibody-coated beads, approximately 10 % of the total nucleotide exchanged after 18 h at room temperature and ∼75 % of the exchanged nucleotide was hydrolysed within 2 h (Field et al., 1996). These data indicate that the Drosophila septin complex is relatively inert in exchange assays, either because it does not exchange GTP in cells or because the purified septins lack appropriate exchange factors. It remains unclear whether nucleotide exchange or hydrolysis regulates septin function in vivo. Many investigators favour a role similar to that in small GTPases, although there is no direct support for such a model. Mitchison and Field (2002) have argued that GTP binding may play a structural role in protein folding and perhaps complex formation, akin to the role of GTP bound to the non-exchanging site on α-tubulin, but that GTP exchange/hydrolysis does not regulate septin function once the complex is assembled (see also Vrabioiu et al., 2004).

Filament formation by septin complexes in vitro Early studies of wild-type and mutant yeast by electron microscopy (EM) and immunofluorescence led to the hypothesis that septin polypeptides are the major structural components of the 10-nm ‘neck filaments’ observed in the neck region of budding cells (Byers and Goetsch, 1976; Longtine et al., 1996; and see Chapter 3). It was thus satisfying to find that when dialyzed into a buffer of physiological ionic strength, the immunopurified Drosophila septin complex assembled into filaments in vitro. The assembled filaments were approximately 7 nm in diameter and of variable length. Length histograms revealed that filaments were multiples of an ∼26-nm subunit, presumably corresponding to the 2 : 2 : 2 septin complex (Field et al., 1996). The distribution of lengths, with short filaments co-existing with longer ones, suggested that pure septin complexes polymerize in vitro by an isodesmic mechanism, without a nucleation step (see, for example, Romberg, Simon and Erickson, 2001). In this mechanism, assembly and disassembly are not

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CH06 METAZOAN MODEL SYSTEMS

limited to polymer ends. Adjacent polymer subunits in the middle of a filament can dissociate from each other, breaking the filament in two, and, in a converse reaction, two filaments can also anneal to form a longer one. Analysis of septin complexes isolated from other organisms demonstrated that the polymerization behaviour exhibited by the Drosophila complex is typical (Frazier et al., 1998; Hsu et al., 1998; Kinoshita et al., 2002). Characterization of yeast and human septin complexes also revealed additional features suggesting that in addition to linear polymers, septin filaments can associate laterally to form higher-order structures in cells. For example, yeast septin filaments tend to align side by side (Frazier et al., 1998), whereas human septin filaments tend to roll up into torus-shaped assemblies (Kinoshita et al., 2002). The 26-nm periodicity of the Drosophila septin filaments led to the speculation that the yeast neck filaments might not run circumferentially around the mother-bud axis, as suggested by the early EM studies, but instead be oriented axially, parallel to the mother-bud axis (Field et al., 1996), with the alignment between monomers in adjacent filaments generating the circumferential striations seen in the EM images. Although still controversial (see Chapters 3 and 4), this model has recently been supported by a new technique utilizing polarized fluorescence microscopy of GFP-septin fusions (Vrabioiu and Mitchison, 2006, 2007).

Assembly of a two-septin C. elegans complex Septins are found in fungi and animals, but not in plants. Based on phylogenetic comparisons, all septins can be partitioned into five groups, with the animal septins falling exclusively into groups 1 and 2 (Pan, Malmberg and Momany, 2007; Chapter 2). The genomes of most sequenced animals encode between four and thirteen septins, with all animals containing at least one, and typically multiple, septins from each group. In contrast, the C. elegans genome encodes just two septins, one in group 1 (UNC-61) and one in group 2 (UNC-59) (Nguyen et al., 2000; Pan, Malmberg and Momany, 2007). Depletion of either one or both septins gives essentially identical phenotypes (Nguyen et al., 2000; Finger, Kopish and White, 2003; Maddox et al., 2007), and each protein is required for the other’s localization (Nguyen et al., 2000), suggesting that both proteins are required to form a functional complex in vivo. This relative simplicity has made the C. elegans septins an attractive system for functional characterization and biochemical reconstitution. The C. elegans septin complex has been assembled after expression either in E. coli or in insect cells (John et al., 2007). In both cases, the two proteins were insoluble when expressed individually but formed a soluble complex when co-expressed. The predominant soluble complex was a tetramer with a 2 : 2 UNC-59:UNC-61 stoichiometry, but dimers with a 1 : 1 stoichiometry of the two subunits were also observed. EM of negatively stained tetramers revealed an


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elongated complex ∼7-nm wide and 17–20-nm long. After classification and averaging, these elongated particles resolved into four linearly arranged globular densities. By performing EM on preparations in which either UNC-59 or UNC-61 was tagged with GFP, the heterotetramer was shown to exist in an UNC-59/UNC-61/UNC-61/UNC-59 configuration, suggesting that the UNC-59/ UNC-61 dimers form a non-polar tetramer via the association of their UNC-61 subunits. It should be noted that in contrast to previously purified septin complexes, these heterologously expressed C. elegans complexes exhibited low nucleotide content (

The Septins

The

The

The Cutie (The Mercenaries)

The Defector [The Delivery]

The Dreamseller: The Calling

The Memnoch the Devil

The Sky's the Limit

The The Awakening

The The Battle Master

The Beyond the Grave

The The Grey King

The The Witching Hour

The Lady, the Lion

The Demon, the City

The The Struggle

The

The

The Defector [The Delivery]

The More the Terrier

The Carpenter, The Fairy

The Temple and the

The Sky's the Limit

The Deader the Better

The The Kingless Land

The The Wizard Heir

The The Shattered Land

The The Omen Machine

The Captive, The Fugitive

The The Vacant Throne

The Septins

The

The

The Cutie (The Mercenaries)

The Defector [The Delivery]

The Dreamseller: The Calling

The Memnoch the Devil

The Sky's the Limit

The The Awakening

The The Battle Master

The Beyond the Grave

The The Grey King

The The Witching Hour

The Lady, the Lion

The Demon, the City

The The Struggle

The

The

The Defector [The Delivery]

The More the Terrier

The Carpenter, The Fairy

The Temple and the

The Sky's the Limit

The Deader the Better

The The Kingless Land

The The Wizard Heir

The The Shattered Land

The The Omen Machine

The Captive, The Fugitive

The The Vacant Throne

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