The Nucleus - PDF Free Download

The Nucleus A Subject Collection from C o ld Spring H arbor Perspectives in Biology Articles online at www.cshperspecti...

Author: David L. Spector | Tom Misteli (editors)

98 downloads 2005 Views 25MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

The Nucleus A Subject Collection from C o ld Spring H arbor Perspectives in Biology Articles online at www.cshperspectives.org A ll rights reserved < 2 0 1 1 by C old Spring Harbor Laboratory Press, Cold Spring Harbor, N ew York Printed in the United States o f America Publisher

|ohn Inglis

Acquisition Editor

Richard Sever

Project M anager

M aryliz Dickerson

Permissions C oordinator

Carol Brown

Production Editor

Kaaren Kockenmeister

Cover Designer

Denise Weiss

Front cover artwork: Diagram illustrating the organization of the eukaryotic nucleus. Depicted are chromatin (orange), enclosed by and anchored to the nuclear envelope (gray), and transcrib ing RNPs (purple), w hich after processing are exported through nuclear pore complexes (green and blue). (Illustration courtesy o f M ichael P. Rout. The Rockefeller University.)

Library o f Congress Cataloging-in-Publication Data

The nucleus/edited by Tom Mistelli, David L. Spector, p. cm. Includes bibliographical references and index. ISBN 978-0-87969-894-2 (hardcover: alk. paper) 1. C ell nuclei. I. Mistelli, Tom. II. Spector, David L. III. Title. Q H 5 9 5 .N 8 5 2011 571,6'6--dc22 2010039936 10

9

8

7

6

5

4

3

2

1

A ll W orld W id e W eb addresses are accurate to the best of our knowledge at the time of printing. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Cold Spring Harbor Laboratory Press, provided that the appropriate fee is paid directly to the Copyright Clearance Center (CCC). Write or call CC C at 222 Rosewood Drive, Danvers, M A 01923 (978-750-8400) for information about fees and regulations. Prior to photocopying items for educational classroom use, contact C C C at the above address. Additional information on C C C can be obtained at C C C O nline at http://www.copyright.com/. A ll Cold Spring Harbor Laboratory Press publications may be ordered directly from Cold Spring Harbor Laboratory Press, 500 Sunnyside Blvd., Woodbury', New York 11797-2924. Phone: 1-800843-4388 in Continental U.S. and Canada. A ll other locations: (516) 422-4100. FAX: (516) 4224097. E-mail: cshpress
Contents

Preface, ix

OVERVIEW The Nucleus Introduced, 1 Thoru Pederson

THE NU CLEA R PERIPHERY The Nuclear Envelope, 17 M a r lin W. Helzer

Nuclear Lamins, 33 Thomas Dechat, Stephen A. Adam , Pekka Taimen, Takeshi Shimi, and Robert D. Goldman

Lamin-binding Proteins, 55 Katherine I.. Wilson a n d Roland Foisner

The Nuclear Pore Complex and Nuclear Transport, 73 Susan R. Wente a n d M ich a e l P. Rout

C H R O M O S O M ES & CHROM ATIN Chromosome Territories, 93 Thomas Cremer and M a rio n Cremer

Gene Pbsitioning, 115 Carm elo Ferrai, Ines Jesus de Castro, Liron Lavitas, M ita Chotalia, a n d Ana Pombo

Chromatin Higher-order Structure and Dynamics, 133 Christopher L Woodcock and Rajarshi P. Ghosh

Nuclear Organization and Dosage Compensation, 159 Jennifer C. Chow and F.dith Heard

The Budding Yeast Nucleus, 175 Angela Taddei, Heiko Schober, and Susan M . Gasser

Nuclear Functions of Actin, 195 Neus Visa and Piergiorgio Percipalle

NU CLEA R BODIES The Nucleolus, 209 Thoru Pederson

Nuclear Speckles, 225 D av id I.. Speclor and Angus I. Lam ond

The Cajal Body and Histone Locus Body, 237 Zehra Nizam i, Svetlana Deryiisheva, and Joseph G. G a ll

PM L Nuclear Bodies, 249 Valerie Lallemand-Breitenbach and Hugues de The

The Perinucleolar Compartment, 267 C allie Pollock and Sui H uang

Rjraspeckles, 277 Archa H . Fox and Angus I. Lam ond

Nuclear Stress Bodies, 291 Giuseppe Biam onti a n d Claire Vourc’h

Orphan Nuclear Bodies, 303 M a r ia Carmo-Fonseca, M a ria T. Berciano, a n d M iguel Lafarga

Biogenesis of Nuclear Bodies, 315 M iroslav D u n d r and Tom M is le li

F U N C T IO N A L ORGANIZATION Organization of Transcription, 331 Lyubomira Chakalova a n d Peter Fraser

Organization of D N A Replication, 347 Vadim O. Chagin, Jeffrey II. Stear, a n d M . Cristina Cardoso

D N A Damage Response, 361 Giuseppina G ig lia -M a ri, Angelika Zotter, and Wint Vermeulen

RN A Processing and Export, 381 Sam i Hocine, Robert H . Singer, a n d D avid G riinw ald

N UCLEAR ARCHITECTURE IN DIFFERENTIATION A N D DISEASE Diseases of the Nuclear Envelope, 401 H ow ard J. Worman, Cecilia Ostlund, a n d Yuexia Wang

Higher-order Genome Organization in Human Disease, 419 Tom M isteli

Nuclear Ataxias, 437 I Iarry T. O rr

Developments in RN A Splicing and Disease, 449 M ich ael G. Poulos, Ranjan Batra, Konstantinos Charizanis, and M a urice S. Swanson

APPENDIX Tables of Data on the Nucleus, 463 (Adapted by kind permission of Federation o f American Societies for Experimental Biology [FASEB] from B iological Handbooks. I. Cell Biology (Altman P.L. and Katz DD; 1976], Bethesda, Maryland.)

Index, 507

The Nucleus Introduced Thoru Pederson Program in Cell and Developmental Dynamics, Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts 01605 Correspondence: [email protected]

Now is an opportune moment to address the confluence of cell biological form and function that is the nucleus. Its arrival is especially timely because the recognition that the nucleus is extremely dynamic has now been solidly established as a paradigm shift over the past two decades, and also because we now see on the horizon numerous ways in which organization itself, including gene location and possibly self-organizing bodies, underlies nuclear functions.

“We have entered the cell, the Mansion of our birth, and started the inventory of our acquired wealth.” —Albert Claude

hen I first read that morsel from Albert Claude’s 1974 Nobel Prize lecture it seemed Solomonic wisdom, as it indeed was. Though he was referring to cell biology en toto, the study of the nucleus was then at a tipping point and new advances were just at hand. Since then, the nucleus field has literally nucleated and we are now at a position to both admire the recent past and register excitement about the present and where the nucleus field may be headed.

W

THE NUCLEUS DISCOVERED

We cannot know who first saw the nucleus but we do know that the father of optical

microscopy, Antony van Leeuwenhoeck, did so with amphibian and avian erythrocytes in 1710 and that in 1781 Felice Fontana did so as well in eel skin cells. More definitive accounts followed by Franz Bauer, who in 1802 sketched orchid cells and pointed out the nucleus (Bauer 1830 –1838), as well as by Jan Purkyneˇ, who described it as the vesicula germanitiva in chicken oocytes (Purkyneˇ 1825), and Robert Brown who observed it in a variety of plant cells (Brown 1829 – 1832), earning additional fame for coining the term “nucleus” (for excellent accounts of these early descriptions of the nucleus see Gall 1996; Harris 1999). Of course, these early observations did not ascribe particular significance to this structure, the given name simply conveying its central location. Later, the nucleus was increasingly observed and became, with some prescience, a key tenet of the cell theory. The nucleus remained a rather lonely item in the eukaryotic cell’s parts list for many

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000521

T. Pederson

decades, until the discoveries of discrete cytoplasmic entities, e.g., mitochondria, the Golgi apparatus etc. (reviewed in Wilson 1925). THE NUCLEUS INHERITED

We do not know how and when the genome of an ancestral cell first became encased in a primitive nucleus. We have no evidence that cells living in the RNA world ever had a membrane (or any other structure) around the genome, i.e., that they ever became nucleate. Once a ribozyme RNA replicase arose, anything would have been possible including the emergence of ribozymes with lipid biosynthetic activities. Enthalpy-favored or free energy-driven events could then have led to stabilizing selection of RNA-lipid affinities and on from there. A cottage industry of experiments on the interactions of lipids with RNA has emerged in the chemical biology field in the past decade but the significance of these studies to prebiotic systems and the earliest cells remains speculative. As for the advent of the DNA world, and of eukaryotes, a major concept is that a prokaryote organism was invaded by another, nonnucleated cell, setting up an endosymbiotic relationship, with the entering organism’s outer membrane seeding what would become the nuclear envelope. The major proponent of this plausible idea has also suggested that this hypothetical invader also brought in a centriole, the forerunner of what we know as the centriole/ basal body in extant eukaryotes (Margulis et al. 2000). We can not play the videotape of life on Earth backwards and although we can reconstruct some things with a degree of empirical confidence, albeit amidst debate (reviewed by Misteli 2001a; Poole and Penny 2001; Rotte and Martin 2001), or speculation (e.g., Lake 2009), when it comes to how the nucleus arrived, we just do not know. THE NUCLEUS ENVELOPED

Notwithstanding the uncertainty of its evolutionary origin, the nucleus is bounded by a double membrane, the nuclear envelope, which in many cells is contiguous with the endoplasmic

reticulum. The frequently observed intimacy of the nuclear envelope with the endoplasmic reticulum has been often under-appreciated, particularly as it bears on the isolation of nuclei and issues of resulting purity. There is also growing interest in how nuclear membrane proteins may be integrators of nuclear and cytoplasmic organization and dynamics (e.g., King et al. 2008; Roux et al. 2009; Starr 2009). A seminal finding was that the nuclear envelope contains pores (reviewed by Gall 1964; 1967), which have now been defined in considerable compositional and structural detail (e.g., Alber et al. 2007; Fernandez-Martinez and Rout 2009). A somewhat less familiar but equally important area of investigation has revealed that the nuclear envelope harbors a signal transduction system of its own, featuring players in common with the plasma membrane, for example the lipid-linked inositol trisphosphate system (Martelli et al. 1991; for reviews see Divecha et al. 1993; Cocco et al. 2009; Barton et al. 2010). THE NUCLEUS DIVERSIFIED

However the nucleus arose, it went on to display a variety of organizations. These range from the highly condensed nuclei of mature erythrocytes in nonmammalian vertebrates to the bimorphic nuclei in almost all ciliates. In the latter organisms a micronucleus perpetuates the genome whereas a macronucleus contains DNA fragments that represent only a fraction of the organism’s genome complexity and which encode the RNAs and proteins needed for vegetative life. It was a particular feature of this genomic strategy, namely the macronuclear amplification of the ribosomal RNA genes (Gall 1974), that led to the discovery of the telomere DNA sequence (Blackburn and Gall 1978). It was also in these ciliates that selfsplicing RNA was discovered and in which the era of chromatin epigenetic marks was launched (reviewed in Pederson 2010). THE NUCLEUS VIEWED

Beyond early observations made by bright field microscopy, the staining method developed by

The Nucleus, Introduced

Robert Feulgen, which attaches a dye to aciddepurinated DNA, was a major tool in advancing the DNA ¼ gene theory, based on studies of the DNA content of haploid versus diploid cells by a graduate student, Hewson Swift, in the laboratory of Arthur Pollister at Columbia University, and concurrent ones by Hans Ris in the laboratory of Alfred Mirsky at the Rockefeller Institute (reviewed by Pederson 2005). Although biochemical measurements of the DNA contents of germ versus somatic cells had revealed a twofold difference, these findings mostly remained in the biochemical community and did not have as much impact as might have been expected. It was the Feulgen cytophotometry results that helped catalyze the idea that DNA is genes. This was, to borrow part of Winston Churchill’s famous phrase—“the end of the beginning” (Avery et al. 1944). Meanwhile, phase contrast microscopy had beautifully revealed the interphase nucleus and mitotic chromosomes, the latter not “nuclear” in the strictest sense. Electron microscopy led to the visualization of the double nuclear membrane and nuclear pores (Gall 1964; 1967), the tripartite structure of the nucleolus (Bernhard et al. 1952), the nuclear lamina (Fawcett 1966), and subsequently the observation of chromatin n-bodies (Olins and Olins 1974), later re-named nucleosomes (Oudet et al. 1975). An intriguing nuclear structure first observed by the Spanish neuroanatomist Santiago Ramon y Cajal (Cajal 1903, 1910) underwent a renaissance of interest when one of its protein components was identified by Eng Tan and colleagues (Andrade et al. 1991; Raska et al. 1991). Initially termed the coiled body, a campaign in 1999 by Joseph Gall resulted in a consensus to rename this structure the Cajal body. Gall has made some of the most seminal advances in our understanding of this nuclear body (for reviews see Gall 2000; Handwerger et al. 2006; Gall 2009; Nizami et al. 2010). Similarly, in the 1990s the nucleolus, interchromatin granule clusters (a.k.a. nuclear speckles) and other nuclear bodies also underwent advances in molecular definition (reviewed by Spector 1993; Pederson 2002a; Spector 2001; 2006).

THE NUCLEUS ISOLATED

Using pus-soaked bandages from a local hospital, Friedrich Miescher discovered DNA in the late 1860s. As described in a richly detailed historical account (Portugal and Cohen 1977), Miescher wrote a manuscript and submitted it to a top journal but had to sit by and wait while the editor (and his former mentor), Felix Hoppe-Seyler, checked the findings in his own lab, resulting in the publication of two concurrent papers (Miescher 1871; Hoppe-Seyler 1871). Over the next 90 years, efforts to isolate nuclei did not go very far. In the 1960s Alfred Dounce at the University of Rochester, Van Potter and Conrad Elvehjem at the University of Wisconsin, and Philip Siekevitz and George Palade at the Rockefeller Institute pioneered the isolation of nuclei from animal tissue. (The Rochester and Wisconsin investigators had the homogenizers they developed named for them.) Important advances in isolating nuclei from plant tissue were made at approximately the same time by James Bonner’s group at Caltech. Later, groups at Baylor College of Medicine (Harris Busch), Rockefeller University (Alfred Mirsky) and Albert Einstein College of Medicine (Sheldon Penman) published refined methods. As is the case for most cell fractionation methods, none of these proved to be perfect but they were major advances nonetheless. The fact that the endoplasmic reticulum and the nuclear envelope are contiguous in many cells is but one example of this challenge, and the tendency of the cytoplasmic intermediate filament system to often collapse upon the nucleus during cell fractionation is yet another. The pioneer methods of nuclear isolation employed sucrose concentrations in which the tissue homogenate was exposed to hydrodynamic forces and/or sucrose density differences that caused cytoplasmic elements and adherent endoplasmic reticulum to separate from the nuclei. The most popular of the early methods (Chauveau et al. 1956) was subsequently refined in important ways to further minimize cytoplasmic contamination (Maggio et al. 1963; Blobel and Potter 1966), with these two latter advances constituting the gold standard of

T. Pederson

nuclear isolation for many years, and still today for many tissues. As regards methods aimed at single-cell studies, an important method (Penman 1966) involved swelling cells in a hypotonic buffer so that the expanded cell volume would collide with the hydrostatic shear forces delivered by a 0.0015-inch clearance stainless steel device, patterned after ones introduced by Dounce. Detergent-based methods were also introduced (Traub et al. 1964). These typically employed Triton X-100 or Royal Dutch Shell’s “nonionic P-40” (a.k.a. NP-40). These breach the plasma membrane and, depending on cell type, also strip away the endoplasmic reticulum. In some cases, NP-40 collapses some of the plasma membrane onto the nucleus, so that nuclei prepared by this method may not always be as pure as some investigators have assumed. For some cells, the main advantage of the NP-40 method is less the nuclear fraction, but the relatively high purity of the ribosomes that are released (Borun et al. 1967). All methods of nuclear isolation must be monitored both for what remains attached from the cytoplasm (or is taken up from it), as well as what is lost from within. Various markers have been used to assess the depletion of cytoplasmic material in isolated nuclei, based on the presumption that the true intracellular locations of such markers are known to some degree of certainty. In one study, HeLa cell nuclei were isolated in a buffer consisting of a previous cytoplasmic fraction from 3H-leucine-labeled cells (Bhorjee and Pederson 1972), thus allowing the level of cytoplasmic protein contamination of the nuclei to be readily estimated. Issues of nuclear purity loomed large in early studies claiming that protein synthesis occurs in the isolated nuclei (and thus presumably within nuclei in vivo) but this work was challenged on several grounds (reviewed by Goldstein 1970; Pederson 1976). More recently this issue has resurfaced (e.g., Iborra et al. 2001, although this study also included intact cell experiments). Various controls for nuclear purity have been better but still not sufficient to quiet all doubts as to the source of apparent protein synthesis in isolated nuclei (for reviews

see Pederson 2001a; Dahlberg et al. 2003; Nathanson et al. 2003). Meanwhile, less attention has been devoted to analyzing not what is adsorbed to nuclei as opposed to what is lost from nuclei during various isolation methods. There have been other instructive guideposts over the years for judging the purity of nuclear fractions. An instructive vignette is the saga of the biosynthesis of the U-rich spliceosomal small nuclear RNAs. We now know that they are exported to the cytoplasm as 30 extended precursors, there to be trimmed, 50 -cap hypermethylated and assembled with various proteins before re-entry into the nucleus as functional snRNPs. Although many studies had pointed to this pathway at varying degrees of cogency, it was the use of two particular nuclear isolation methods, each with exceptionally strong credentials, that led to acceptance of this pathway of snRNP biogenesis (Gurney and Eliceiri 1980; Zieve et al. 1988). This was further solidified by the demonstration that exogenous U2 snRNA precursor molecules introduced into the cytoplasm of mammalian cells enter this pathway and become 30 trimmed, cap hypermethylated, and assembled into snRNPs, followed by nuclear uptake (Kleinschmidt and Pederson 1990). Another example of how the critical issue of nuclear purity was addressed arose in a study in which the total sequence complexity of nuclear versus cytoplasmic RNA was measured (Holland et al. 1980). Because the nuclear RNA was anticipated to have a higher sequence complexity than cytoplasmic RNA, contamination of the cytoplasmic fraction by leaked nuclear RNA loomed large. Accordingly, an experiment was undertaken in which the cells were labeled with an RNA tracer for a very short time (2 min), far too brief for any labeled transcripts to be exported to the cytoplasm in vivo, before cell fractionation. The label was followed throughout the isolation of the cytoplasmic fraction and polyribosome-associated mRNA, which revealed that no more than 0.03% of the nuclear RNA synthesized in the previous 2 min was present in the final mRNA preparation (Holland et al. 1980). Given that these

The Nucleus, Introduced

kinds of controls are conceptually obvious and experimentally quite facile, it is surprising how rarely they have been employed in studies in which knowing the purity of a nuclear or cytoplasmic fraction is essential to weighing the findings. Beyond the isolation of nuclei in bulk, there is the gold standard of manual isolation of nuclei from material that affords this opportunity (e.g., Duryee 1954; later refined and reviewed by Lund and Paine 1990; Paine et al. 1992). In a recent application of this approach, frog oocyte nuclei were manually isolated under oil and the relative densities of the nucleolus versus surrounding nucleoplasmic bodies were determined by differential interference light microscopy, with results that defied most expectations (Handwerger et al. 2005). For most of the cells and tissues from which one wants to isolate nuclei, manual isolation is of course out of the question. But it is worth emphasizing that what has been learned from occasional studies employing manually isolated nuclei is likely to be a more reliable guide to understanding nuclear organization and function than has sometimes been realized by those members of the nucleus research community whose work is based on bulk nuclear isolation. A general point to be made is that although isolated nuclei served many important experimental uses during the modern era of research on the nucleus, their role in advancing the biochemistry of gene expression was surprisingly short-lived and quite limited. Cell-free systems based on isolated nuclei played only transitory and rather minor roles in the areas of DNA replication, transcription, and mRNA processing—usually giving way to more efficient soluble systems in short order. An exception was the use of isolated nuclei to allow RNA polymerase II to continue transcription and thus, by hybridization analysis of the extended chains, determine the boundaries of a transcription unit (Weber et al. 1977). In contrast to these limited roles of isolated nuclei in the investigation of gene expression, a very different cell-free system derived from frog eggs (Newport and Forbes 1987) was of monumental importance in advancing our understanding of chromatin

and nuclear envelope assembly and the control of cell-cycle progression. THE NUCLEUS COMPOSED

In 1976, the Federation of American Societies of Experimental Biology (FASEB) heroically collated all available information on the “properties of cells” and I was chosen to organize and edit the chapter on the nucleus. At the suggestion of the editors, and with FASEB’s permission, we are republishing here the tables of data on the nucleus from the original volume (Altman and Katz 1976) (see supplemental data online). These tables speak to painstaking analytical work carried out in a time gone by and rarely pursued today. But let there be no mistake, the concentrations of protein or RNA in various preparations of isolated nuclei, or extracts derived from them, can be a critical factor in the interpretation of certain kinds of studies. What is needed now to complement these important compositional data is additional biophysical information (reviewed in Pederson 2002a). One important step in this direction is that the fluid viscosity of the interchromatin space has now been estimated, based on diffusion coefficients of reporter molecules, to be approximately one-fifth that in water (Wachsmuth et al. 2000). The considerations of fluid viscosity and free versus anomalous diffusion are critical to the understanding of intranuclear transport and nuclear body dynamics (Wachsmuth et al. 2000). In terms of chemical kinetics one would also want to know the water concentration in the nucleus. Only then could one know, or at least estimate, the ionic strength of the solvent phase and the true concentrations of solutes. The importance of these biophysical parameters is conveyed by the fact that only a relatively small change in the intracellular ion concentration elicits a dramatic mitosis-like state, which is completely reversible (Robbins et al. 1970). Polyamines are abundant constituents of the nucleus in most cells and yet their possible functions are rarely considered. Iron is a surprisingly abundant component of the interphase nucleus and mitotic chromosomes and is essential for DNA

T. Pederson

replication (Robbins and Pederson, 1970). Many compositional issues such as these are rarely considered in current research on the nucleus and yet they can be extremely determinative. But, happily, there are steps in this direction, as biophysicists increasingly join the nucleus research community (for an “interdisciplinary” review see O’Brien et al. 2003). THE NUCLEUS DECONSTRUCTED

Major advances in biology were made by teasing a cell or its parts out, as from the squid giant axon, leading to the discovery of kinesin, or by glycerinating rabbit muscle to discover the biochemistry of its contraction. The major conceptual discovery about the nucleus, that it contains the genome, did not involve its dissection. But a desire to know its parts naturally arose in due course. Early attempts to isolate chromatin were made by the laboratories of Alfred Mirsky (Rockefeller) and James Bonner (Caltech) but these fractions were of dubious enrichment and the major advances in the field of chromatin and chromosome structure came from in situ studies (reviewed by DuPraw 1970). One of the first, convincing isolations of a subnuclear component was that of the nucleolus from starfish oocytes (Vincent 1955). Later, three groups went further. Palade and Siekevitz at Rockefeller purified guinea pig liver nuclei, subjected them to sonication and isolated nucleolar and nucleoplasmic fractions (Maggio et al. 1963). The demonstrated degree of fractionation was impressive and this work constituted one of the most important advances in the nucleus field at the time. Meanwhile, the laboratory of Harris Busch at Baylor College of Medicine developed a similar, sonication-based method for isolating nucleoli from rat hepatoma cells (Muramatsu et al. 1963) and subsequently this group, notably Ramachandra Reddy, exploited this method to identify a number of small nucleolar and nucleoplasmic RNAs long before their functions became known (for a review see Reddy and Busch 1988). Shortly thereafter, Sheldon Penman developed a method to resolve a HeLa cell nuclear fraction into nucleoli and nucleoplasm (Penman 1966).

His method involved the use of DNAse and high ionic strength and produced nucleolar and nucleoplasmic fractions that were less native than those obtained by the sonication method, although it led to spectacular advances in our understanding of RNA biosynthesis. More recently, mammalian and plant cell nucleoli have been purified and subjected to extensive proteomic analyses (Andersen et al. 2002; Scherl et al. 2002; Pendle et al. 2005; Hinsby et al. 2006; for reviews see Dundr and Misteli 2002; Pederson 2002b; Coute´ et al. 2006 ; Leung et al. 2006). Nucleoplasmic bodies known as interchromatin granule clusters (a.k.a. “speckles”) were isolated by David Spector and colleagues (Mintz et al. 1999) and later subjected by this group to proteomics analysis (Saitoh et al. 2004; reviewed in Lamond and Spector 2003). Meanwhile, Cajal bodies (reviewed by Gall 2000, 2009; Nizami et al. 2010) were isolated by Angus Lamond’s laboratory (Lam et al. 2002). An exciting aspect of this recent period was that these nuclear bodies were being analyzed in such molecular detail at the very time the dynamics of their components were being observed in live cell studies, as will be discussed below. Another nuclear structure appears in some cells, the perinucleolar compartment (Huang et al. 1997; 1998), and its presence has recently been found to have promising prognostic value in the staging of breast carcinoma (Kamath et al. 2005). It is a discoid, caplike structure intimately attached to the nucleolus and has thus resisted isolation so far. There presently are no clues to its function, beyond its accretion of certain small RNAs transcribed by RNA polymerase III (Matera et al. 1995; Weng et al. 2003). THE NUCLEUS TERRITORIALIZED

A major development was the discovery that interphase chromosomes occupy specific locations (Zorn et al. 1979; Cremer et al. 1982), which to some extent had been anticipated from classical studies of the arrangement of chromosomes in the metaphase plate (Rabl 1885) and by the actual coining of the term “chromosome territory” (Boveri 1909). The

The Nucleus, Introduced

modern concept of chromosome territories was promptly adopted in certain sectors of the human genetics and radiation biology communities but took more than two decades to gain broader traction (reviewed by Cremer and Cremer 2001; 2006), now with appreciated relevance to chromosomal translocations (see Pederson 2003 for a review), genome evolution (Tanabe et al. 2002; Foster and Bridger 2005) and the entire issue of how chromosome location relates to gene density and/or expression. This latter area has been hyperactive in the past few years (reviewed by Spector 2003; Pederson 2004; Gilbert et al. 2005; Sprout et al. 2005; Cremer et al. 2006; Akhtar and Gasser 2007; Misteli et al. 2007; Sexton et al. 2007; Takizawa et al. 2008; Towbin et al. 2009; Deniaud and Bickmore 2009) but the results have been surprisingly variable and confounding (for a particularly lucid summary see the Introduction in Meaburn et al. 2008). A distinct possibility that has arisen recently is that gene positioning is self-organizing, based not on expression per se but on the potential for sets of genes to be coregulated (Rajapakse et al. 2009; reviewed by Misteli 2009). A clinical feature of chromosome territoriality is the fact that the extreme cytological manifestation of gene repression, i.e., heterochromatin, has long served as a key landmark for pathologists, both in its extent and location. But an exciting new dimension of diagnostic potential lies in studies, mentioned earlier, of intranuclear gene locations in relation to not only reciprocal translocations (Roix et al. 2003; reviewed in Pederson 2003) but in more recent work on gene repositioning in solid tumors (e.g., Meaburn et al. 2009). The nucleus is territorialized not only with respect to the locations of the chromosomes themselves but also to a considerable extent with respect to the layout of expressed versus silenced genomic regions. This aspect of the nuclear structure field has recently taken one particularly surprising turn, reminding us that we are still in the early days. In most cells, heterochromatin is located at the nuclear periphery or in close approximation to the nucleolus. Remarkably, it has recently been reported that

in the rod photoreceptor cell nuclei of nocturnal animals, the heterochromatin is coalesced into a large central domain in the nucleus with the euchromatin displaced to more peripheral locations, with plausible speculations as to the efficiency of light transmission and retinal harvesting (Solovei et al. 2009; reviewed by Ragoczy and Groudine 2009). Another important recent development has been the introduction of methods to capture the interchromosome regions that lie in closest juxtaposition (Dekker et al. 2002; reviewed by Dostie and Dekker 2007). With this new methodology the field of genome organization is moving to a 4-D spatialtemporal registration, which is itself likely to be determinative of phenotype. The extraordinary packing density of interphase chromatin has been recently investigated with “Hi-C”, a powerful variation of the chromosome conformation capture methodology (LiebermanAiden et al. 2009; for a review see Langowski 2010).

THE NUCLEUS NOT LIKELY MATRIXED

Breaking up nuclei with sonication is one thing, extracting them with salt is another. The latter was a major effort of many labs in the 1950s and thereafter, most notably for isolating and characterizing histones. But others (e.g., Zbarsky and Georgiev 1959) employed graded salt extractions to fractionate total nuclear components. This yielded successive fractions of differing composition, not unexpected because the zwitterion concept of protein net charge had long been discovered by Arne Tiselius and advanced by John Edsall. There was therefore no reason to believe that cell components such as nuclei, mitochondria, or ribosomes would not release different sets of proteins as the ionic strength is elevated. Few cell biologists were comfortable with assigning a residue of the nucleus any relationship to the in vivo situation. But in 1974 this nuclear residue got renamed a “nuclear matrix” (Berezney and Coffey 1974) and later work gave rise to the idea that chromatin and nascent RNA are attached to it.

T. Pederson

The notion took hold in some quarters and gained a degree of traction as all sorts of entities were observed to be present in this fraction. But there was never any proof of the in vivo existence of such a structure nor is there proof today (for detailed reviews see Pederson 1998; 2000) and many leaders in the field of nuclear structure and function question the nuclear matrix concept. The consensual skepticism as to a nucleuswide, arborized network of filaments extending throughout the nucleoplasm certainly does not rule out the existence of short-range structural motifs. THE NUCLEUS IN DIVISION

The current focus of the cell division field on chromosome capture by kinetochore– microtubule interactions, how anaphase works mechanistically and the operation of mitotic checkpoints is of course well justified. But it is also possible that this emphasis may be missing one of the most intriguing of all events—the formation of daughter nuclei. Precursors of the nucleolus, nuclear envelope, and nuclear lamina congress, mature, and assemble in “daughter to be” cells as early as anaphase and definitively in telophase. This presently subsidiary effort in the cell division field needs more momentum as it is no less intriguing than anaphase, and likely involves very different mechanisms such as the free energy of DNA–lipid affinities and other chemical phenomena that are unique to telophase and the assembly of daughter nuclei, seemingly via pathways whose memory had not been erased from the previous mitosis. THE NUCLEUS IMAGED

In his detailed recipes, the histochemistry pioneer A.G. Everson Pearse described various ways to stain cells, including the nucleus (Pearse 1961). I tried many of these methods as a student including the aforementioned Feulgen reaction. The combination of methyl green and pyronin ended up as my favorite, with which I first saw nucleoli, to become a longstanding interest. At the end of the classical era, immunostaining and susbsequently the

advent of GFP advanced the localization of nuclear proteins and bodies, and the discovery of in situ nucleic acid hybridization (Gall and Pardue 1969) made possible the localization of both genes and RNAs. These methods, especially when powerfully used in combination, have made it possible to even demarcate subregions of nuclear domains formerly thought to be relatively homogenous, such as the granular component of the nucleolus, which now appears to be a landscape of distinct molecular zones rather than a uniform lawn of nascent ribosomes (Politz et al. 2002; 2005). Of course, these light microscopy advances have left ample room for the continuing application of electron microscopy. Particular progress has been made in the past two decades with respect to the ultrastructural analysis of both nuclear pore complexes, as mentioned earlier, as well as a defined messenger RNP, sometimes the two caught together (Mehlin et al. 1992). THE NUCLEUS IN MOTION, WITHIN

There was never much doubt that, around the less mobile chromosomes and nucleoli, molecules roam the nucleus. This emerged from numerous studies in which tagged molecules were microinjected into the nucleus and observed to display high mobility (e.g., Wang et al. 1990). Subsequently it became possible to express fluorescently tagged nuclear proteins and observe their dynamics. The first such study examined the dynamics of interchromatin granule clusters (Misteli et al. 1997) which revealed these nucleoplasmic bodies to be more dynamic than had been anticipated. Then, a new wave of studies appeared resulting from the application of the method of fluorescence recovery after photobleaching (FRAP). In the first of these, FRAP was employed to study the mobility of the DNA repair machinery tagged with GFP (Houtsmuller et al. 1999). This paper represented both a technical and conceptual milestone and soon thereafter FRAP was applied to several other nuclear proteins in a boomlet of important studies (Kruhlak et al. 2000; Lever et al. 2000; Misteli et al. 2000; Phair and Misteli 2000; Boisvert et al. 2001; Chen and

The Nucleus, Introduced

Huang 2001; reviewed in Pederson 2000b; Misteli 2001b; Pederson 2001b; Phair and Misteli 2001). These revealed much more rapid and/ or more extensive dynamics than would have been anticipated from either earlier in vitro work, or from the apparent stasis of certain nuclear bodies, constituting a true paradigm shift in the nucleus field (reviewed by Misteli 2001b). Even the nuclear lamina, which had long been viewed as one of the most stable structures in the nucleus, was found to undergo dynamic exchange of subunits, leading to an appreciable nucleoplasmic concentration (Moir et al. 2000; reviewed by Dechat et al. 2008) and there is increasing evidence that the dynamic lamins have functions while in the nucleoplasm (Malhas et al. 2007; Lee et al. 2009; reviewed by Shimi et al. 2008; Shumaker et al. 2008), and it now appears that nucleoporins do as well (Capelson et al. 2010; Kalverda et al. 2010). This wave of FRAP studies addressed the dynamics of protein constituents of nuclear bodies and was, of course, based on GFP. But GFP also enabled investigation of the movements of nuclear bodies and their constituents. Studies of the movements of chromosomes (Marshall et al. 1997; reviewed by Gasser 2002), promyelocytic leukemia (PML) bodies (Muratani et al. 2002) and Cajal bodies (Platani et al. 2002) were the first of these. The Marshall et al. study was ahead of its time and it was amusing to recall the incredulity expressed by some that interphase chromosomes, relatively giant structures, are moving, and with no dependence on metabolic energy. But others properly anticipated that there is of course no reason the chromosomes would not display this microscopic biophysical property, viz. the manifestation of the kinetic energy of any particle. Although GFP enabled these studies (for reviews see Misteli et al. 1997; Eils et al. 2000; Pederson 2000b; Misteli 2001b; Pederson 2001b; Chubb and Bickmore 2003; Dundr et al. 2002; Gasser 2002), investigating the intranuclear movements of the other major nuclear species, RNA, required different innovation. The fact that microinjection of a pre-mRNAs

or small nucleolar RNAs into the nucleus of mammalian cells resulted in localization with sites known to contain splicing machinery or the nucleoli, respectively (Wang et al. 1990; Jacobson et al. 1995; Jacobson and Pederson 1998; reviewed in Pederson 2001c) showed that introduced RNA is mobile. Further innovations led to tagging of endogenous RNA and these studies confirmed that poly(A) RNA is highly diffusive in the nucleus (Politz et al. 1998; 1999; reviewed in Politz and Pederson 2000; Pederson 2001c), and subsequent studies revealed, importantly, that this was true of specific mRNAs (Singh et al. 1999; Shav-Tal et al. 2004) and also 28S ribosomal RNA (Politz et al. 2003). These approaches also made it possible to investigate the live cell dynamics of RNA in relation to specific intranuclear structures, e.g., interchromatin granules (Politz et al. 2006). An increasing number of single-molecule level studies of nuclear molecular dynamics have appeared recently (Dange et al. 2008; Gru¨nwald et al. 2008; Siebrasse et al. 2009) and, together with revolutionary advances in the spatial resolution of diffraction-limited optical microscopy (for a brief review see Pederson 2006), one senses the dawning of a new era as systems biology enters the nucleus, or vice-versa (reviewed by Gorski and Misteli 2005). Another major advance was the introduction of a method to tag specific chromatin regions via the binding of GFP-tagged lac repressor to an integrated tandem array of the lac operator (Robinett et al. 1996), which in turn allowed the visualization in living cells of gene activation at discrete loci (Tsukamoto et al. 2000; Janicki et al. 2004) and the dynamics of transcription factors (Becker et al. 2002; Karpova et al. 2008; Sprouse et al. 2008). As mentioned earlier, the movements of interphase chromosomes have implications for the statistics of reciprocal exchanges between interphase chromosomes, with disease relevance (Roix et al. 2003; reviewed by Pederson 2003). Nucleoli also move to the extent that the chromosomes which contain the repeated rRNA genes are mobile. Diffusion, by definition, arises from the thermal energy inherent in the particle itself. However, the possibility that gene repositioning may be mediated by a process

T. Pederson

that uses metabolic energy has recently been the topic of initial studies (and debate). There is a growing body of evidence for actin and myosin in the nucleus, a field that has moved past its initial uncertainties and is now addressing functions (reviewed by Pederson and Aebi 2002; 2005; Pederson 2008). In particular, three recent studies have implicated actin-based processes in gene repositioning (Chuang et al. 2006; Dundr et al. 2007; Hu et al. 2008). This emerging concept is among the most deserving of vigilance in the nucleus field at this time. THE NUCLEUS THAT AWAITS US

Speculation is always risky. One thing is clear— the question of how nuclear bodies arise in the first place (reviewed by Misteli 2001c) has taken on a new dimension because of the discovery that a Cajal body can assemble when key components are experimentally addressed to a specific nuclear site (Kaiser et al. 2008). Other recent studies raise the possibility that noncoding, nucleus-retained RNAs may play roles in nuclear architecture (Clemson et al. 2009; Sasaki et al. 2009; Sunwoo et al. 2009; reviewed by Wilusz et al. 2009). Also on the present horizon are stem cell issues that call for increasing input from experts on the nucleus. Stem cells have been hyped but it is early days and the nucleus cell biology community has much to offer, both by science and skepticism (for the latter perspective see Lander 2009). The essence of “stem-cellness” is an asymmetric descent of phenotypic potential vs. maintaining a continual seed of replenishment. So the two daughter cells have to be profoundly different but it is a difference we presently grasp very dimly. This is as deep a problem in cell biology as there is today, and there are already encouraging signs of progress (e.g., Parnell and Stillman 2008). Another field that is ripe for the intervention of nucleus experts is somatic nuclear transfer, in which it is the reaction of the introduced nucleus to the maternal environment, known to be dominant from classical studies, that sets in motion the reprogrammed developmental events. What are the molecules that underlie this powerful influence of the egg cytoplasm?

Finally, there can be no doubt that the increasing ability to study nuclear dynamics and function in living cells, now reaching single molecule level detection, constitutes one of the most powerful advances. The recent breaking of the classical diffraction limit of optical microscopy brings the nucleus into nanoscale range. The possible coaptation of these breakthrough approaches with instructive cell types and model systems stirs excitement for what lies ahead. The nucleus research community is living in very interesting times.

CODA

The assumption underlying this collection on nuclear structure and function is that the nucleus has reached a stage of enabling coherence as part of the epistemological structure of modern biological science. However, there are likely to be many things about the nucleus that we don’t yet know and may not know anytime soon. We can only hope that what the geneticist J.B.S. Haldane posited on the cosmos will prove not to be true for the nucleus: “Now, my suspicion is that the universe is not only queerer than we suppose, but queerer than we can suppose.” If we appropriately bear in mind that the nucleus may be more complicated than we may have once thought, and yet just may be knowable, then this very belief may empower us and our students and successors to penetrate the subject’s awaiting depths, the next of which now beckon. There is every reason to believe in this program. So let us be of good cheer.

ACKNOWLEDGMENTS

I am indebted to Joseph Gall for characteristically insightful suggestions on the manuscript. Cited work from the author’s laboratory was funded by grants from the National Institutes of Health (GM-21595, GM-23914 and GM-60551), the National Science Foundation (MCB-0445841), the American Cancer Society (CD-126), and the G. Harold and Leila Y. Mathers Foundation. Some important ideas

The Nucleus, Introduced

that stirred in the modern era of the nucleus field were catalyzed by a series of symposia convened by the author at the Marine Biological Laboratory, Woods Hole, supported by a grant from the Grass Foundation and the late Ellen Grass. REFERENCES Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprato A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Altman PL, Katz DD. 1976. Biological Handbooks. I. Cell Biology. Federation of American Societies for Experimental Biology, Bethesda, Maryland. Andersen JS, Lyon CE, Fox AH, Leung AKL, Lam YW, Steen H, Mann M, Lamond AI. 2002. Directed proteomic analysis of the human nucleolus. Curr Biol 12: 1 –11. Andrade LEC, Chan EKL, Rasˇka I, Peebles CL, Roos G, Tan EM. 1991. Human autoantibody to a novel protein of the nuclear coiled body: immunological characterization and cDNA cloning of p80-coilin. J Exp Med 173: 1407– 1419. Avery OT, MacLeod CM, McCarty M. 1944. Studies on the chemical nature of the substance inducing transformation of pneumococcal types. I. Induction of transformation by a deoxyribonucleic acid fraction isolated from pneumococcus type III. J Exp Med 79: 137–158. Barton CA, Laishram RS, Anderson RA. 2010. Nuclear phosphoinositides: a signaling enigma wrapped in a compartmental conundrum. Trends Cell Biol 20: 25–35. Bauer F. 1830–1838. Illustrations of orchidaceous plants Ridegway, London. Becker M, Baumann C, John S, Walker DA, Vigneron M, McNally JG, Hager GL. 2002. Dynamic behavior of transcription factors on a natural promoter in living cells. EMBO reports 3: 1188– 1194. Berezney R, Coffey DS. 1974. Identification of a nuclear protein matrix. Biochem Biophys Res Commun 60: 1410– 1417. Bernhard W, Haguenau F, Oberling C. 1952. L’ultrastructure du nucleole de quelques cellules animals, re´ve´le´e par le microscope e´lectronique. Experientia 8: 58– 63. Bhorjee JS, Pederson T. 1972. Nonhistone chromosomal proteins in synchronized HeLa cells. Proc Natl Acad Sci 69: 3345–3349. Blobel G, Potter VR. 1966. Nuclei from rat liver: isolation method that combines purity with high yield. Science 154: 1662–1665. Blackburn EH, Gall JG. 1978. A tandemly repeated sequence at the termini of the extrachromosomal ribosomal RNA genes in Tetrahymena. J Mol Biol 120: 33–53. Boisvert FM, Kruhlak MJ, Box AK, Hendzel MJ, BazettJones DP. 2001. The transcription coactivator CBP is a dynamic component of the promyelocytic leukemia nuclear body. J Cell Biol 152: 1089–1106.

Borun TW, Scharff MD, Robbins E. 1967. Preparation of mammalian polyribosomes with the detergent Nonidet-P40. Biochim Biophys Acta 149: 302– 304. Boveri T. 1909. Die blastomerenkern von Ascaris megalocephala und die theorie der chromosomenindividualita¨t. Arch Zellforsch 3: 181– 268. Brown R. 1829– 1832. Observations on the organs and mode of fecundation in Orchideae and Asclepiadeae. Trans Linn Soc Lond 16: 685– 746. Cajal SRy. 1903. Un sencillo metodo de coloracion selectiva del reticulo protoplasmatico y sus efectos en los diversos organos nerviosos de vertebrados e invertebrados. Trab Lab Invest Biol Univ Madrid 2: 129– 221. Cajal SRy. 1910. El nu´cleo de las ce´lulas piramidales del cerebro humano y del algunos mamiferos. Trab Lab Invest Biol Univ Madrid 8: 27–62. Capelson M, Liang Y, Schulte R, Mair W, Wagner U, Hetzer MW. 2010. Chromatin-bound nuclear pore components regulate gene expression in higher eukaryotes. Cell 140: 372 –383. Chauveau J, Moule Y, Rouiller C. 1956. Isolation of pure and unaltered liver nuclei: morphology and biochemical composition. Exp Cell Res 11: 317– 321. Chen D, Huang S. 2001. Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells. J Cell Biol 153: 169– 176. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 831. Chubb JR, Bickmore WA. 2003. Considering nuclear compartmentalization in the light of nuclear dynamics. Cell 112: 403–406. Clemson CM, Hutchinson JN, Sara SA, Ensminger AW, Fox AH, Chess A, Lawrence JB. 2009. An architectural role for a nuclear noncoding RNA: NETA1 RNA is essential for the structure of paraspeckles. Mol Cell 33: 717– 726. Cocco L, Faenza I, Follo MY, Billi AM, Ramazzotti G, Papa V, Martelli AM, Manzoli L. 2009. Nuclear inositides: Pl-PLC signaling in cell growth, differentiation and pathology. Adv Enzyme Reg 49: 2 –10. Coute´ Y, Burgess JA, Diaz JJ, Chichester C, Lisacek F, Greco A, Sanchez JC. 2006. Deciphering the human nucleolar proteome. Mass Spect Rev 25: 215– 234. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292–301. Cremer T, Cremer C. 2006. Rise, fall and resurrection of chromosome territories: a historical perspective. Part II: Fall and resurrection of chromosome territories during the 1950s and 1960s. Part III: Chromosome territories and the functional nuclear architecture: experiments and models from the 1990s to the present. Eur J Histochem 50: 223 –272. Cremer T, Cremer M, Dietzel S, Muller S, Solovei I, Fakan S. 2006. Chromosome territories—a functional nuclear landscape. Curr Opin Cell Biol 18: 307 –316. Cremer T, Cremer C, Schenider T, Baumann H, Hens L, Kirsch-Volders M. 1982. Analysis of chromosome positions in the interphase nucleus of Chinese hamster cells

T. Pederson by laser-UV-microirradiation experiments. Hum Genet 62: 201 –209. Dahlberg JE, Lund E, Goodwin EB. 2003. Nuclear translation: what is the evidence? RNA 9: 1 –8. Dange T, Gru¨nwald D, Gru¨nwald A, Peters R, Kubitscheck U. 2008. Autonomy and robustness of translocation through the nuclear pore complex: a single-molecule study. J Cell Biol 183: 77– 86. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832 –853. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306–1311. Deniaud E, Bickmore WA. 2009. Transcription and the nuclear periphery: edge of darkness? Curr Opin Genet Dev 19: 187 –191. Divecha N, Banfic´ F, Irvine RF. 1993. Inositides and the nucleus and inositides in the nucleus. Cell 74: 405– 407. Dostie J, Dekker J. 2007. Mapping networks of physical interactions between genomic elements using 5C technology. Nat Protocols 2: 988 –1002. Dundr M, McNally JG, Cohen J, Misteli T. 2002. Quantitation of GFP-fusion proteins in single living cells. J Struct Biol 140: 92–99. Dundr M, Misteli T. 2002. Nucleolomics: an inventory of the nucleolus. Mol Cell 9: 5 –7. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095– 1103. DuPraw EJ. 1970. DNA and Chromosomes Holt, Rinehart & Winston, New York. Duryee WR. 1954. Microdissection studies on human ovarian eggs. Trans NY Acad Sci 17: 103– 108. Eils R, Gerlich D, Tvarusko´ W, Spector DL, Misteli T. 2000. Quantitative imaging of pre-mRNA splicing factors in living cells. Mol Biol Cell 11: 413– 418. Fawcett DW. 1966. On the occurrence of a fibrous lamina on the inner aspect of the nuclear envelope in certain cells of vertebrates. Am J Anat 119: 129 –145. Fernandez-Martinez J, Rout MP. 2009. Nuclear pore complex biogenesis. Curr Opin Cell Biol 21: 603– 612. Foster HA, Bridger JM. 2005. The genome and the nucleus: a marriage made by evolution. Chromosoma 114: 212 –229. Gall JG. 1964. Electron microscopy of the nuclear envelope. Protoplasmatologia 5: 4 –25. Gall JG. 1967. Octagonal nuclear pores. J Cell Biol 32: 391 – 399. Gall JG. 1974. Free ribosomal RNA genes in the macronucleus of Tetrahymena. Proc Natl Acad Sci USA 71: 3078– 3081. Gall JG. 1996. A pictorial history. Views of the cell. American Society for Cell Biology, Bethesda, MD. Gall JG. 2000. Cajal bodies: the first 100 years. Ann Rev Cell Dev Biol 16: 273– 300. Gall JG. 2009. Chromosome odds and ends. Ann Rev Cell Dev Biol, in press.

Gall JG, Pardue ML. 1969. Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci 63: 378– 383. Gasser SM. 2002. Visualizing chromatin dynamics in interphase nuclei. Science 296: 1412–1416. Gilbert N, Gilchrist S, Bickmore WA. 2005. Chromatin organization in the mammalian nucleus. Int Rev Cytol 242: 283–336. Goldstein L. 1970. On the question of protein synthesis by cell nuclei. Adv Cell Biol 1: 187–210. Gorski S, Misteli T. 2005. Systems biology in the cell nucleus. J Cell Sci 118: 4083– 4092. Gru¨nwald D, Martin RM, Buschmann V, Bazett-Jones DP, Leonhardt H, Kubitscheck U, Cardoso MC. 2008. Probing intranuclear environments at the single-molecule level of detection. Biophys J 94: 2847–2858. Gurney T, Eliceiri GL. 1980. Intracellular distribution of low molecular weight RNA species in HeLa cells. J Cell Biol 87: 398– 403. Handwerger KE, Cordero JA, Gall JG. 2005. Cajal bodies, nucleoli, and speckles in the Xenopus oocyte nucleus have a low-density, sponge-like structure. Mol Biol Cell 16: 202– 211. Handwerger KE, Gall JG. 2006. Subnuclear organelles: new insights into form and function. Trends Cell Biol 16: 19– 26. Harris H. 1999. The birth of the cell. Yale University Press, New Haven. Hinsby AM, Kiemer L, Karlberg EO, Lage K, Fausbøll A, Juncker AS, Andersen JS, Mann M, Brunak S. 2006. A wiring of the human nucleolus. Mol Cell 22: 285– 295. Holland CA, Mayrand SH, Pederson T. 1980. Sequence complexity of nuclear and messenger RNA in HeLa cells. J Mol Biol 138: 755 –778. ¨ ber die chemische zusammensetHoppe-Seyler F. 1871. U zung des eiters. Hoppe-Seyler’s Medizinisch-Chemischen Untersuchungen 4: 486–501. Houtsmuller AB, Rademakers S, Nigg AL, Hoogstraten D, Hoeijmakers JHJ, Vermuelen W. 1999. Action of DNA repair endonuclease ERCC1/XPF in living cells. Science 284: 958–961. Hu Q, Kwon YS, Nunez E, Cardamone MD, Hutt KR, Ohgi KA, Garcia-Bassets I, Rose DW, Glass CK, Rosenfeld MG, et al. 2008. Enhancing nuclear receptor-induced transcription requires nuclear motor and LSD1-dependent gene networking in interchromatin granules. Proc Natl Acad Sci USA 105: 19199– 19204. Huang S, Derrinck TJ, Ellisman MH, Spector DL. 1997. The dynamic organization of the perinucleolar compartment in the cell nucleus. J Cell Biol 137: 965– 974. Huang S, Derrinck TJ, Ellisman MH, Spector DL. 1998. The perinucleolar compartment and transcription. J Cell Biol 143: 35–47. Iborra FJ, Jackson DA, Cook PR. 2001. Coupled transcription and translation within nuclei of mammalian cells. Science 293: 1139– 1142. Jacobson MR, Cao LC, Wang YL, Pederson T. 1995. Dynamic localization of RNase MRP RNA in the nucleolus observed by fluorescent RNA cytochemistry in living cells. J Cell Biol 131: 1649–1658.

The Nucleus, Introduced Jacobson MR, Pederson T. 1998. A 7-methylguanosine cap commits U3 and U8 small nuclear RNAs to the nucleolar localization pathway. Nucleic Acids Res 26: 756– 760. Janicki SM, Tsukamoto T, Salghetti SE, Tansey WP, Sachidanandam R, Prasanth KV, Ried T, Shav-Tal Y, Bertrand E, Singer RH, et al. 2004. From silencing to gene expression: real-time analysis in single cells. Cell 116: 683– 698. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713– 1717. Kalverda B, Pickersgill H, Shloma VV, Fornerod M. 2010. Nucleoporins directly stimulate expression of developmental and cell-cycle genes inside the nucleoplasm. Cell 140: 360– 371. Kamath RV, Thor AD, Wang C, Edgerton SM, Slusarczyk A, Leary DJ, Wang J, Wiley EL, Jovanovic B, Wu Q, et al. 2005. Perinucleolar compartment prevalence has an independent prognostic value for breast cancer. Cancer Res 65: 246– 253. Karpova TS, Kim MJ, Spriel C, Nalley K, Stasevich TJ, Kherrouche Z, Heliot L, McNally JG. 2008. Concurrent fast and slow cycling of a transcriptional activator at an endogenous promoter. Science 319: 466– 469. King MC, Drivas TG, Blobel G. 2008. A network of nuclear envelope membrane proteins linking centromeres to microtubules. Cell 134: 427–438. Kleinschmidt AM, Pederson T. 1990. RNA processing and ribonucleoprotein assembly studied in vivo by RNA transfection. Proc Natl Acad Sci 87: 1283–1287. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazaett-Jones DP, Hendzel MJ. 2000. Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150: 41–51. Lake JA. 2009. Evidence of an early prokaryotic endosymbiosis. Nature 460: 967– 971. Lam YW, Lyon CE, Lamond AI. 2002. Large-scale isolation of Cajal bodies from HeLa cells. Mol Biol Cell 13: 2461– 2473. Lamond AI, Spector DL. 2003. Nuclear speckles: a model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605–612. Lander AD. 2009. The ‘stem cell’ concept: is it holding us back? J Biol 8: 70. Langowski J. 2010. Chromosome conformation by crosslinking: polymer physics matters. Nucleus 1: 13–16. Lee DC, Welton KL, Smith ED, Kennedy BK. 2009. A-type nuclear lamins act as transcriptional repressors when targeted to promoters. Exp Cell Res 315: 996 –1007. Lever MA, Th’ng JP, Sun X, Hendzel MJ. 2000. Rapid exchange of histone H1.1 on chromatin in living cells. Nature 408: 873–876. Leung AK, Trinkle-Mullcahy L, Lam YW, Andersen JS, Mann M, Lamond AI. 2006. NOPdb: nucleolar proteome database. Nucleic Acid Res 34: D218–220. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –293. Lund E, Paine P. 1990. Nonaqueous isolation of transcriptionally active nuclei from Xenopus oocytes. Methods Enzymol 181: 36–43.

Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. I. Isolation and chemical characterization of a nuclear fraction from guinea pig liver. J Cell Biol 18: 267 –291. Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. II. Isolation and chemical characterization of nucleolar and nucleoplasmic subfractions. J Cell Biol 18: 292– 312. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593 –603. Margulis L, Dolan MF, Guerrero R. 2000. The chimeric eukaryote: origin of the nucleus from the karyomastigont in amitochondriate protists. Proc Natl Acad Sci 97: 6954–6959. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Martelli AM, Gilmour RS, Bertagnolo V, Neri LM, Manzoli L, Cocco L. 1991. Nuclear localization and signalling activity of phosphoinositidase Cb in Swiss 3T3 cells. Nature 358: 242 –244. Matera AG, Frey MR, Margelot K, Wolin SL. 1995. A perinucleolar compartment contains several RNA polymerase III transcripts as well as the polypyrimidine tract binding protein, hnRNP I. J Cell Biol 129: 1181–1193. Meaburn KJ, Newbold RF, Bridger JM. 2008. Positioning of human chromosomes in murine cell hybrids according to synteny. Chromosoma 117: 579– 591. Meaburn KJ, Gugla PR, Khan S, Lockett SJ, Misteli T. 2009. Disease-specific gene repositioning in breast cancer. J Cell Biol 187: 801 –812. Mehlin H, Daneholt B, Skoglund U. 1992. Translocation of a specific premessenger ribonucleoprotein particle through the nuclear pore studied with electron microscope tomography. Cell 15: 605– 613. ¨ ber die chemische zusammensetzung Miescher F. 1871. U der eiterzellen. Hoppe-Seyler’s Medizinisch-Chemischen Untersuchungen 4: 441–460. Mintz PJ, Patterson SD, Neuwald AF, Spahr CS, Spector DL. 1999. Purification and biochemical characterization of interchromatin granule clusters. EMBO J 18: 4308–4320. Misteli T, Gunjan A, Hock R, Bustin M, Brown DT. 2000. Dynamic binding of histone H1 to chromatin in living cells. Nature 408: 877–881. Misteli T. 2001a. Where the nucleus comes from. Trends Cell Biol 11: 149. Misteli T. 2001b. Protein dynamics: implications for nuclear architecture and gene expression. Science 291: 843– 847. Misteli T. 2001c. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Misteli T. 2007. Beyond the sequence: cellular organization of genome function. Cell 128: 787– 800. Misteli T. 2009. Self-organization in the genome. Proc Natl Acad Sci USA 106: 6885– 6886. Misteli T, Ca´ceres JF, Spector DL. 1997. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523 –527.

T. Pederson Moir RD, Yoon M, Khuon S, Goldman RD. 2000. Nuclear lamins A and B1: different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Muramatsu M, Smetana K, Busch H. 1963. Quantitative aspects of isolation of nucleoli of the Walker carcinosarcoma and liver of the rat. Cancer Res 23: 510– 518. Muratani M, Gerlich D, Janicki SM, Gebhard M, Eils R, Spector DL. 2002. Metabolic-energy-dependent movement of PML bodies within the mammalian cell nucleus. Nature Cell Biol 4: 106 –110. Nathanson L, Xia T, Deutscher MP. 2003. Nuclear protein synthesis: a re-evaluation. RNA 9: 9– 13. Newport JW, Forbes DJ. 1987. The nucleus: structure, function, and dynamics. Ann Rev Biochem 56: 535–565. Nizami Z, Deryusheva S, Gall J. 2010. The Cajal body and histone locus body. Cold Spring Harb Perspect Biol 2: a000653. O’Brien TP, Bult CJ, Cremer C, Grunze M, Knowles BB, Langowski J, McNally J, Pederson T, Politz JC, Pombo A, et al. 2003. Genome function and nuclear architecture: from gene expression to nanoscience. Genome Res 13: 1029– 1041. Olins AL, Olins DE. 1974. Spheroid chromatin units (n bodies). Science 183: 330 –332. Paine PL, Johnson ME, Lau YT, Tluczek LJM, Miller DS. 1992. The oocyte nucleus isolated in oil retains in vivo structure and functions. Biotechniques 13: 238 –245. Parnell EJ, Stillman DJ. 2008. Getting a transcription factor to only one nucleus following mitosis. PLoS Biol 6: 1829– 1832. Pearse AGE. 1961. Histochemistry. 2nd edition, Little, Brown and Co., Boston. Pederson T. 1976. Cellular aspects of histone synthesis. In Protein synthesis (ed. E.H. McConkey), Vol. 2: pp. 69– 123, Marcel Dekker, New York. Pederson T. 1998. Thinking about a nuclear matrix. J Mol Biol 277: 147–159. Pederson T. 2000a. Half a century of “the nuclear matrix”. Mol Biol Cell 11: 799– 805. Pederson T. 2000b. Diffusional protein transport within the nucleus: a message in the medium. Nature Cell Biol 2: E73– 74. Pederson T. 2001a. Is the nucleus in need of translation? Trends Cell Biol 11: 395–397. Pederson T. 2001b. Protein mobility within the nucleus— what are the right moves? Cell 104: 635–638. Pederson T. 2001c. Fluorescent RNA cytochemistry: tracking gene transcripts in living cells. Nucleic Acids Res 29: 1013– 1016. Pederson T. 2002a. Dynamics and genome-centricity of inter-chromatin domains in the nucleus. Nat Cell Biol 4: E287– E291. Pederson T. 2002b. Proteomics of the nucleolus: more proteins, more functions? Trends Biochem. Sci 27: 111 –112. Pederson T. 2003. Gene territories and cancer. Nature Genetics 34: 242–243. Pederson T. 2004. The spatial organization of the genome in mammalian cells. Curr Opin Genet Dev 14: 203–209.

Pederson T. 2005. In memoriam- Hans Ris. Chromosoma 114: 307– 309. Pederson T. 2006. Diffraction at the Brandeburg g(r)ate. EMBO reports 7: 1202–1205. Pederson T. 2008. As functional nuclear actin comes into view, is it globular, filamentous, or both? J Cell Biol 180: 1061– 1064. Pederson T. 2010. An olympian protozoan. Nucleus 1: 1– 3. Pederson T, Aebi U. 2002. Actin in the nucleus: what form and what for? J Struct Biol 140: 3 –9. Pederson T, Aebi U. 2005. Nuclear actin extends, with no contraction in sight. Mol Biol Cell 16: 5055– 5060. Pendle AF, Clark GP, Boon R, Lewandowska D, Lam YW, Andersen J, Mann M, Lamond AI, Brown JW, Shaw PJ. 2005. Proteomic analysis of the Arabidopsis nucleolus suggest novel nucleolar functions. Mol Biol Cell 16: 260–269. Penman S. 1966. RNA metabolism in the Hela cell nucleus. J Mol Biol 17: 117– 130. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Phair RD, Misteli T. 2001. Kinetic modeling approaches to in vivo imaging. Nat Rev Mol Cell Biol 2: 898 –907. Platani M, Goldberg I, Lamond AI, Swedlow JR. 2002. Cajal body dynamics and association with chromatin are ATP-dependent. Nat Cell. Biol 4: 502– 508. Politz JCR, Pederson T. 2000. Review: movement of mRNA from transcription site to nuclear pores. J Struc Biol 129: 252 –257. Politz JC, Lewandowski LB, Pederson T. 2002. Signal recognition particle RNA localization within the nucleolus differs from the classical sites of ribosome synthesis. J Cell Biol 159: 411– 418. Politz JCR, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805– 4812. Politz JC, Brown ES, Wolf DE, Pederson T. 1998. Intranuclear diffusion and hybridization state of oligonucleotides measured by fluorescence correlation spectroscopy in living cells. Proc Natl Acad Sci 95: 6043–6048. Politz JCR, Polena I, Trask I, Bazett-Jones DP, Pederson T. 2005. A nonribosomal landscape in the nucleolus revealed by the stem cell protein nucleostemin. Mol Biol Cell 16: 3401– 4310. Politz JC, Tuft RA, Pederson T, Singer RH. 1999. Movement of nuclear poly(A) RNA throughout the interchromatin space in living cells. Curr Biol 9: 285– 291. Politz JCR, Tuft RA, Prasanth KV, Baudendistel N, Fograt KE, Lifshitz LM, Langowski J, Spector DL, Pederson T. 2006. Rapid, diffusional shuttling of poly(A) RNA between nuclear speckles and the nucleoplasm. Mol Biol Cell 17: 1239–1249. Poole A, Penny D. 2001. Does endo-symbiosis explain the origin of the nucleus? Nat Cell Biol 3: E173– E174. Portugal FH, Cohen JS. 1977. ACentury of DNA. MIT Press, Cambridge, MA. Purkyneˇ JE. 1825. Symbolae ad ovi avium historiam ante incubationem. 1st ed. Vratislaviae. Reprinted 1948: Opera selecta. Tomus 1, Opera facultatis medicae universitatis carolinae pragensis. ¨ ber zelltheilung. Morph Jb 10: 214–330. Rabl C. 1885. U

The Nucleus, Introduced Ragoczy T, Groudine M. 2009. The nucleus inside outthrough a rod darkly. Cell 137: 205–207. Rajapakse I, Perlman MD, Scalzo D, Kooperberg C, Groudine M, Kosak ST. 2009. The emergence of lineage-specific chromosomal topologies from coordinate gene regulation. Proc Natl Acad Sci USA 106: 6679– 6684. Rasˇka I, Andrade LEC, Ochs RL, Chan EKL, Chang CM, Roos G, Tan EM. 1991. Immunological and ultrastructural studies of the nuclear coiled body with autoimmune antibodies. Exp Cell Res 195: 27–37. Reddy R, Busch H. 1988. Small nuclear RNAs: sequences, structure and modifications. In Structure and function of major and minor small nuclear ribonucleoprotein particles M.L. Birnstiel, ed. Springer-Verlag, Berlin, pp. 1–37. Robinett CC, Straight A, Li G, Willhelm C, Sudlow G, Murray A, Belmont AS. 1996. In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J Cell Biol 135: 1685–1700. Robbins E, Pederson T. 1970. Iron: its intracellular localization and possible role in cell division. Proc Natl Acad Sci USA 66: 1244– 1251. Robbins E, Pederson T, Klein P. 1970. Comparison of mitotic phenomena and effects induced by hypertonic solutions in HeLa cells. J Cell Biol 44: 400 –416. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in lymphomas. Nat Genetics 34: 287– 291. Rotte C, Martin W. 2001. Does endo-symbiosis explain the origin of the nucleus? Nat Cell Biol 3: E173–E174. Roux KJ, Crisp ML, Liu Q, Kim D, Kozlov S, Stewart CL, Burke B. 2009. Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc Natl Acad USA 106: 2194– 2199. Saitoh N, Spahr CS, Patterson SD, Bubulya P, Neuwald AF, Spector DL. 2004. Proteomic analysis of interchromatin granule clusters. Mol Biol Cell 15: 3876–3890. Sasaki YTF, Ideue T, Sano M, Mituyama T, Hirose T. 2009. MEN e/b noncoding RNAs are essential for structural integrity of nuclear paraspeckles. Proc Natl Acad Sci USA 106: 2525– 2530. Scherl A, Coute´ Y, Deon C, Calle K, Kindbeiter K, Sanchez JD, Greco A, Hochstrasser D, Diaz JJ. 2002. Functional proteomic analysis of human nucleolus. Mol Biol Cell 13: 4100–4109. Sexton T, Schober H, Fraser P, Gasser SM. 2007. Gene regulation through nuclear organization. Nat Struc. Mol Biol 14: 1049–1055. Shav-Tal Y, Darzacq X, Shenoy SM, Fusco D, Janicki SM, Spector DL, Singer RH. 2004. Dynamics of single mRNPs in nuclei of living cells. Science 304: 1797– 1800. Shimi T, Pfleghaar K, Kojima S, Pack CG, Solovei I, Goldman AE, Adam SA, Shumaker DK, Kinjo M, Cremer T, et al. 2008. The A- and B-type nuclear lamin networks: microdomains involved in chromatin organization and transcription. Genes Dev 22: 3409– 3421. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin

Ig-fold binds to PCNA: its role in DNA replication. J Cell Biol 181: 269– 280. Siebrasse JP, Veith R, Dobay A, Leonhardt H, Daneholt B, Kubitscheck U. 2009. Discontinuous movement of mRNP particles in nucleoplasmic regions devoid of chromatin. Proc Natl Acad Sci 105: 20291–20296. Singh OP, Bjo¨rkroth B, Masich S, Wieslander L, Daneholt B. 1999. The intranuclear movement of Balbiani ring premessenger ribonucleoprotein particles. Exp Cell Res 251: 135–146. Solovei I, Kreysing M, Lanctoˆt C, Ko¨sem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356 –368. Spector DL. 1993. Macromolecular domains within the cell nucleus. Ann Rev Cell Biol 9: 265–315. Spector DL. 2001. Nuclear domains. J Cell Sci 114: 2891– 2893. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Ann Rev Biochem 72: 573 –608. Spector DL. 2006. SnapShot: cellular bodies. Cell 127: 1070. Sprouse RO, Karpova TS, Mueller F, Dasgupta A, McNally JG, Auble DT. 2008. Regulation of TATA-binding protein dynamics in living yeast cells. Proc Natl Acad Sci USA 105: 13304– 13308. Sprout D, Gilbert N, Bickmore WA. 2005. The role of chromatin structure in regulating the expression of clustered genes. Nat Rev Genet 6: 775 –781. Starr DA. 2009. A nuclear-envelope bridge positions nuclei and moves chromosomes. J Cell Sci 122: 577–586. Sunwoo H, Dinger ME, Wilusz JE, Amaral PP, Mattick JS, Spector DL. 2009. MEN e/b nuclear-retained noncoding RNAs are up-regulated upon muscle differentiation and are essential components of paraspeckles. Genome Res 19: 347 –359. Takizawa T, Meaburn KJ, Misteli T. 2008. The meaning of gene positioning. Cell 135: 9– 13. Tanabe H, Muller S, Neusser M, von Hase J, Calcagno E, Cremer M, Solovei I, Cremer C, Cremer T. 2002. Evolutionary conservation of chromosome territory arrangements in cell nuclei from higher primates. Proc Natl Acad Sci USA 99: 4424– 4429. Towbin BD, Meister P, Gasser SM. 2009. The nuclear envelope- a scaffold for silencing? Curr Opin Genet Dev 19: 180– 186. Traub A, Kaufmann E, Ginzburg-Tietz Y. 1964. Studies on nuclear ribosomes. I. Association of DPN-pyrophosphorylase with nuclear ribosomes in normal and neoplastic tissues. Exp Cell Res 34: 371–383. Tsukamoto T, Hashiguchi N, Janicki SM, Tumbar T, Belmont AS, Spector DL. 2000. Visualization of gene activity in living cells. Nat Cell Biol 2: 871 –878. Vincent WS. 1955. Structure and chemistry of nucleoli. Int Rev Cytol 4: 269–298. Wachsmuth M, Waldeck W, Langowski J. 2000. Anomalous diffusion of fluorescent probes inside living cell nuclei investigated by spatially-resolved fluorescence correlation spectroscopy. J Mol Biol 298: 677– 689.

T. Pederson Wang J, Cao LG, Wang YL, Pederson T. 1990. Localization of pre-messenger RNA at discrete nuclear sites. Proc Natl Acad Sci 88: 7391–7395. Weber J, Jelinek W, Darnell JE. 1977. The definition of large viral transcription unit late in Ad2 infection of HeLa cells: mapping of nascent RNA molecules labeled in isolated nuclei. Cell 10: 611– 616. Weng C, Politz JC, Pederson T, Huang S. 2003. RNA polymerase III transcripts and the PTB protein are essential for the integrity of the perinucleolar compartment. Mol Biol Cell 14: 2425– 2435. Wilson EB. 1925. The cell in development and heredity. 3rd edition. Macmillan, New York, NY, pp. 28–57.

Wilusz JE, Sunwoo H, Spector DL. 2009. Long noncoding RNAs: functional surprises from the RNA world. Genes Dev 23: 1494–1504. Zbarsky IB, Georgiev GP. 1959. Cytological characteristics of protein and nucleoprotein fractions of cell nuclei. Biochim Biophys Acta 32: 301 –302. Zieve GW, Sauterer RA, Feeney RJ. 1988. Newly synthesized small nuclear RNAs appear transiently in the cytoplasm. J Mol Biol 199: 259– 267. Zorn C, Cremer C, Cremer T, Zimmer J. 1979. Unscheduled DNA synthesis after partial UV irradiation of the cell nucleus: distribution in interphase and metaphase. Exp Cell Res 124: 111 –119.

The Nuclear Envelope Martin W. Hetzer Salk Institute for Biological Studies, Molecular and Cell Biology Laboratory, La Jolla, California 92037 Correspondence: [email protected]

The nuclear envelope (NE) is a highly regulated membrane barrier that separates the nucleus from the cytoplasm in eukaryotic cells. It contains a large number of different proteins that have been implicated in chromatin organization and gene regulation. Although the nuclear membrane enables complex levels of gene expression, it also poses a challenge when it comes to cell division. To allow access of the mitotic spindle to chromatin, the nucleus of metazoans must completely disassemble during mitosis, generating the need to re-establish the nuclear compartment at the end of each cell division. Here, I summarize our current understanding of the dynamic remodeling of the NE during the cell cycle.

he NE, a hallmark of eukaryotic cells, is a highly organized double membrane that encloses the nuclear genome (Kite 1913). Early electron microscopy (EM) images revealed that the inner (INM) and outer nuclear membranes (ONM) are continuous with the endoplasmic reticulum (ER) (Watson 1955). Despite the lipid continuity between the NE and the ER, both ONM and INM are comprised of diverse groups of proteins that are typically not enriched in the ER (Hetzer et al. 2005) (Table 1). The first group consists of 30 different polypeptides, called nucleoporins or Nups, which form the 40 – 70 MD nuclear pore complexes (NPCs) (Tran and Wente 2006; D’Angelo and Hetzer 2008). NPCs are aqueous channels that show eightfold rotational symmetry with an outer diameter of 100 nm and a central transport channel measuring 40 nm in diameter, through which bidirectional exchange of proteins, RNA, and

T

ribonucleoprotein complexes between the nucleoplasm and cytoplasm occurs (Beck et al. 2004; Beck et al. 2007; Terry et al. 2007). A subset of Nups is stably embedded in the NE, forming a scaffold structure or NPC core (Rabut et al. 2004; D’Angelo et al. 2009), which is thought to stabilize the highly curved and energetically unfavorable pore membrane (Alber et al. 2007; Boehmer et al. 2008). This core includes the Nup107/160 complex (Nup84 complex in yeast) and the Nup205 complex (yeast Nup170), which together constitute 50% of the entire NPC (Fig. 1) (Brohawn et al. 2009). Attached to this scaffold are peripheral Nups, many of which contain phenylalanine-glycine (FG) rich repeats that establish a permeability barrier and also mediate active, receptor-dependent transport across the NE (Peters 2009). A second group of NE proteins, specifically localizes to the INM (Fig. 1) (Schirmer and Gerace 2005).

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000539 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000539

M.W. Hetzer Table 1. List of NE proteins. NPC

INM

ONM

Lamina

Nup35

LBR

Nup37

Lap1

Nup43

Lap2b

Nup50

Emerin

Lamin A Lamin B1 Lamin B2 Lamin C

Nup54 Nup58/45 Nup62 Nup75 Nup88 Nup96

MAN1 Nurim NET8 NET38 NET56 LEM2/ NET25 NET9 NET32 NET37 Sun1 Sun2 LUMA 60 NETs

Nesprin-3/ Net35 Syne/Myne/ Nesprin 1 Nesprin-2a and b Syne/ Nesprin-2G Samp1

Nup98 Nup107 Nup133 Nup153 Nup155 Nup160 Nup188 Nup205 Nup214 Nup358/ RanBP2 Sec13R Seh1 Pom121 Ndc1 Gp210 Tpr Rae1 Aladin Nlp1/hCG1

The nuclear pore complex (NPC) contains 30 nucleoporins (Nups). Nup98 and Nup96 are synthesized as a single polypeptide that becomes autoproteolytically cleaved to give rise to Nup98 and Nup96. Inner nuclear membrane (INM) proteins: lamin B receptor (LBR), lamin-associated protein (LAP), Nuclear Envelope Transmembrane protein (NET). More than 60 NETs have no assigned function. Outer nuclear membrane (ONM): spectrin repeat containing nuclear envelope protein (syne), spindle-associated membrane protein 1 (Samp1). Lamina: lamin A and C are products of alternative splicing.

Although most of these .60 integral membrane proteins (also referred to as NE transmembrane proteins or NETs [Schirmer et al. 2003]) remain largely uncharacterized, interaction with lamins (see later) and chromatin have been shown for some of them, such as lamin B receptor (LBR), lamina-associated polypeptide (LAP) 1, LAP2, emerin, and MAN1 (Akhtar and Gasser 2007; Dorner et al. 2007; Schirmer and Foisner 2007). It is becoming increasingly clear that INM proteins play vital and diverse roles in nuclear function such as chromatin organization, gene expression, and DNA metabolism (Mattout et al. 2006; Heessen and Fornerod 2007; Reddy et al. 2008). Importantly, improper localization and function of INM proteins have been linked to numerous human diseases, which has sparked considerable interest in NE biology over the last decade (Vlcek and Foisner 2007; Worman and Bonne 2007; Neilan 2009). A third class of NE proteins specifically resides in the ONM (Fig. 1). This diverse group of integral membrane proteins shares a small KASH (Klarsicht, ANC-1, Syne Homolgy) domain, which has been shown to interact with Sad1p/UNC-84 (SUN)-domain proteins of the inner nuclear membrane within the periplasmic space of the NE (Starr and Han 2003; Wilhelmsen et al. 2006). Two other related ONM proteins, nuclear envelope spectrin repeat (nesprin)-1 and -2, have been shown to directly interact with the actin cytoskeleton through their amino-terminal actin-binding domain (ABD) (Wilhelmsen et al. 2005). These ONM proteins are implicated in nuclear positioning that is essential for processes such as cell polarization, pronuclear migration, and the organization of syncitia (Fridkin et al. 2009). In addition, ONM and INM proteins form “bridges” across the perinuclear space that might be involved in separating the two NE membrane leaflets at an even distance of 50 nm (Voeltz and Prinz 2007). These lumenal proteinaceous bridges could establish physical connections between the cytoskeleton and chromatin,

Organization of the Nuclear Membrane

Cytoplasm

Actin, intermediate filaments

BAF

BAF

nesprin

nesprin

ONM

SUN

nurim ?

emerin

?

LEM2

HP1

LAP2

LBR

BAF

NETs

MAN1

ER lumen (PNS)

SUN

NPC

INM

Lamina (lamin A/C and B)

Chromatin Nucleoplasm

Figure 1. Topology of the NE. Inner and outer nuclear membranes (INM and ONM, respectively) are separated

by the ER lumen or perinuclear space (PNS). The nuclear lamina interacts with NE proteins and chromatin. INM proteins link the NE to chromatin and the lamina. ONM proteins provide a connection from the nucleus to the cytoskeleton. The lamin B receptor (LBR) interacts both with B-type lamins and chromatin-associated heterochromatin protein 1 (HP1) in conjunction with core histones. Members of the LEM (lamina-associated protein 2 [LAP2], emerin, MAN1)-domain family ( pink) bind to lamins and interact with chromatin through barrier-to-autointegration factor (BAF). SUN proteins (SUN 1 and 2) interact with nesprins in the ONM, thereby forming so-called LINC complexes that establish connections to actin and intermediate filaments in the cytoplasm. Nurim is a multi-pass membrane protein with unknown function. Proteomic approaches have identified 60 putative transmembrane proteins (NETs), most of which remain uncharacterized.

which might be relevant for transcription, replication, and DNA repair mechanisms (Tzur et al. 2006; Stewart et al. 2007). The final group of NE proteins constitutes the lamina, a meshwork of intermediate filaments that is composed of A- and B-type lamins (Gruenbaum et al. 2000). Although the lamina has been shown to be critical for nuclear stability, particularly in tissues that are exposed to mechanical forces such as muscle fibers (Cohen et al. 2008), it has become clear that lamins also play major roles in chromatin function and gene expression (Gruenbaum et al. 2005; Dechat et al. 2008; Reddy et al. 2008). Similar to INM proteins, mutations in lamins are linked to a large number of diverse human diseases (Mounkes et al. 2003; Muchir and Worman 2004; Mattout et al. 2006) and to aging (Gruenbaum et al.

2005; Scaffidi and Misteli 2006), highlighting the crucial role of the NE protein network for normal cell function. In summary, the NE fulfills a critical role in shielding the genome from cytoplasmic components, but also represents a highly specialized membrane that provides anchoring sites for chromatin and the cytoskeleton (D’Angelo and Hetzer 2006). NE REMODELING IN DIVIDING CELLS

Although a membrane-enclosed nuclear genome can be found in all eukaryotes, there is a critical difference in the cell-cycle dependent dynamics of the NE between “lower” eukaryotes (e.g., yeast and filamentous fungi) and metazoa (i.e., “higher” eukaryotes). The former undergo

M.W. Hetzer

closed mitosis, where spindle microtubules can either form inside the nucleus or are able to penetrate an intact nuclear membrane (Heywood 1978; Byers 1981; Ribeiro et al. 2002). In contrast, the NE of metazoan cells completely disintegrates during cell division to allow the mitotic spindle to access chromosomes (Kutay and Hetzer 2008). As a consequence, every dividing cell has to reform the NE and re-establish the identity of the nuclear compartment (Hetzer et al. 2005; Anderson and Hetzer 2008b; Guttinger et al. 2009). NE remodeling in proliferating cells is a highly dynamic process that involves a vast number of molecular players (Figs. 2,3). By the end of interphase in G2, the nuclei have duplicated their genome, doubled the number of NPCs, and increased the surface area of the NE. The surrounding ER network is continuous with the NE, but not enriched in NE proteins. The entry of mitosis, i.e., prophase, is marked by NE breakdown (NEBD) and the loss of the nucleo-cytoplasmic compartmentalization (Burke and Ellenberg 2002). Between NEBD

G2 ER sheets

and early anaphase, when chromosomes align in the metaphase plate and subsequently segregate, chromatin is essentially free of membranes (Puhka et al. 2007; Anderson and Hetzer 2008a). During these cell-cycle stages, the majority of soluble NE proteins are distributed throughout the cytoplasm and transmembrane NE proteins reside in the mitotic ER (Ellenberg et al. 1997; Anderson and Hetzer 2007; Puhka et al. 2007) (Fig 2). In anaphase, ER membranes begin to reassociate with and rapidly enclose the chromatin mass (Anderson and Hetzer 2008a). Chromatin association of a subset of Nups, decondensation of chromatin, and the assembly of new NPCs occurs concomitantly with NE formation (Anderson and Hetzer 2008c). At the end of cell division, the NE has reformed as a closed membrane barrier and re-establishes the nuclear compartment by enabling selective nucleo-cytoplasmic transport (Dultz et al. 2008). After its formation, the NE expands and undergoes additional structural changes necessary for cell-cycle progression and transcription, including the assembly of new

NEBD in prophase

ER tubules

Metaphase

Dispersal of NE components into ER

Mitotic spindle NE/ER membranes

Chromatin

Chromatin

NPC

Chromatin

Intact NE with NPCs

Disassembled NPCs

Nups at kinetochores

Figure 2. Schematic illustration of NE breakdown. In G2, the cell nucleus has completed DNA replication and

NPC duplication. The NE (dark green), which is continuous with the ER network (green), encloses the chromosomes (blue). NPCs (red/blue) mediate nuclear transport. When cells enter mitosis, NPCs disassemble and the NE gets reabsorbed into the ER, which at this stage is composed of tubules. NPC components are dispersed into the cytoplasm and NE proteins are partitioned into the ER (dashed green lines). Centrosomes (orange dots) move to the NE and microtubules ( purple) participate in the rupturing of the NE. In metaphase, a subset of NPC components has associated with kinetochores and the spindle is established. At this stage, chromosomes at the metaphase plate are devoid of membranes.

Organization of the Nuclear Membrane Anaphase Re-attaching NE/ER membranes

Telophase NE flattening and sealing

G1 NE expansion

Chromatin Chromatin

Chromatin

Chromatin-bound Nups

Assembling NPCs

Nuclear transport

Figure 3. Schematic illustration of NE reformation around segregated chromosomes in one of the two daughter

cells. In anaphase, ER membranes associate with chromatin (red arrows) and a subset of nucleoporins associates with chromatin. Additional membrane tubules and NE proteins are recruited to the chromatin surface and mediate NE flattening. At this stage, most NE proteins are cleared from the ER network. In late anaphase/ early telophase, a closed NE has formed with pores being assembled in a step-wise manner. Once pores become transport competent, the NE expands and cells move into G1.

NPCs (Winey et al. 1997; D’Angelo et al. 2006) (Fig. 3).

BREAKING DOWN THE NUCLEUS

Live cell imaging of mammalian cells has revealed that NEBD, which occurs rapidly within a few minutes in prophase (Lenart et al. 2003), occurs in a step-wise fashion. One of the initial events is the selective loss of Nups from the NPCs (Terasaki et al. 2001; Dultz et al. 2008; Katsani et al. 2008). Nup98, a dynamic NPC component (Griffis et al. 2002), is released before the rest of the NPC components, which disassemble synchronously (Dultz et al. 2008). Collective dispersal of Nups can also be observed in Drosophila, suggesting that pore disassembly is evolutionarily conserved (Katsani et al. 2008). Biochemical evidence suggests that the NPC disassembles into stable subcomplexes and not necessarily into individual polypeptides (Miller and Forbes 2000; Harel et al. 2003b; Walther et al. 2003a), which might stabilize the nucleoporins and allow more rapid reformation of NPCs at the end of mitosis.

Unfortunately, we do not have a clear picture of the biochemical nature of the disassembled NPC components. Recent evidence suggests that some nuclear pore components are degraded in a proteasome-dependent manner during mitosis, a process that might regulate total pore numbers. Furthermore, in the same study, Nup43 has been shown to be absent from the mitotic Nup107/160 complex, which therefore seems to differ from its counterpart in interphase (Chakraborty et al. 2008).

THE ROLE OF KINASES IN NE BREAKDOWN

NPC disassembly is critical for early stages of NEBD, such as disrupting the permeability barrier and allowing the influx of molecules, which are thought to be critical for further disassembly steps. For example, protein kinases like cyclindependent kinase 1(Cdk1/CycB1) may require access to proteins of the INM and lamins for proper mitotic regulation (Wu et al. 1998). That is consistent with the idea that the bulk of lamin disassembly occurs after the NPCs are permeabilized (Lee et al. 2000; Beaudouin

M.W. Hetzer

et al. 2002). As the lamina is disrupted, many INM proteins lose their anchor and are released into the ER (Ellenberg et al. 1997; Yang et al. 1997; Puhka et al. 2007; Anderson et al. 2009a; Lu et al. 2009). One of the critical events during NEBD is the hyperphosphorylation of NE proteins that is thought to disrupt protein complexes and/ or play a role in the activation of factors involved in this process. Several kinases, such as Cdk1/ CycB1, protein kinase C (PKC), Nima, and Aurora A, have been implicated in NEBD (Collas 1999; Burke and Ellenberg 2002; Gong et al. 2007; Portier et al. 2007). The key kinase seems to be Cdk1, which phosphorylates lamins and Nups, including the Nup107/160 complex, Nup93 subcomplex, Nup53, Nup98, Ndc1, and gp210 (Guttinger et al. 2009). The disassembly of INM proteins, such as Lap2a and b and LBR, seem to depend on Cdk1 (Courvalin et al. 1992; Dechat et al. 1998; Dreger et al. 1999). Several other kinases have been shown to participate in NEBD including PKC (Goss et al. 1994), Aurora A (Portier et al. 2007), and polo-like kinase (PLK1) (Chase et al. 2000); however, their targets remain to be determined. Lamin B is the only known target of PKC during NEBD (Goss et al. 1994). In this context, it is worth mentioning that NPCs in organisms with closed mitosis are also partially disassembled. For instance, in Aspergillus nidulans, 14 Nups are released, leaving a partial NPC that allows the entry of the Cdk1/ cyclin B complex (Osmani et al. 2006). This partial disassembly requires the kinases Nima and Cdk1 (De Souza et al. 2004). Thus, structural reorganization of the NPC during mitosis might be an evolutionarily conserved feature shared by all eukaryotes. MEMBRANE DYNAMICS DURING NEBD

An interesting aspect of NEBD was recently revealed by the finding that the nuclear membranes are actively participating in the process. Work in the nematode Caenorhabditis elegans has shown that depletion of reticulons and the GTPase Rab5 inhibits NEBD (Audhya et al. 2007). The reticulons are part of a family

of membrane-associated proteins that together with a distinct class of membrane-bending proteins like DP1 have been shown to tubulate the ER (Voeltz et al. 2006). These results suggest that active reshaping of the ER might be a critical step in NEBD. Two nucleoporins have also been implicated in this process; however, their roles remain unclear. The transmembrane nucleoporin gp210 is enriched in its phosphorylated form at the NE just before NEBD occurs (Galy et al. 2008). The mechanism by which gp210 contributes to the disassembly of NPCs and the lamina remains to be characterized. Depletion of Nup153, a dynamic component of the nuclear basket, inhibits NEBD and was shown to recruit the COPI complex, which mediates retrograde transport from the Golgi to the ER, to the NE (Liu et al. 2003). What roles Nup153 and the COPI complex play in this process has yet to be elucidated. New data suggests that lipid synthesis itself might play a role in NE dynamics. The knock down of lipin, a conserved phosphatidic acid (PA) phosphatase that catalyzesthe dephosphorylation of PA to yield diacylglycerol (DAG) (Siniossoglou 2009), causes defects in NEBD, abnormal chromosome segregation, and irregular nuclear morphology in C. elegans. Interestingly, down-regulation of nematode Lpin-1 was required for disassembly of the nuclear lamina during late NEBD, suggesting that the Lpin-1 requirement appears to be separable from the effect of Lpin-1 on the peripheral ER (Golden et al. 2009; Gorjanacz and Mattaj 2009). Although phosphorylation is clearly a key event in NEBD, interactions of the NE with microtubules are thought to generate mechanical forces that assist in rupturing the nuclear membrane and presumably the lamina in a dynein-mediated process (Beaudouin et al. 2002; Salina et al. 2002; Muhlhausser and Kutay 2007). This process might also involve the small GTPase Ran, which has been shown to regulate microtubule dynamics during NEBD, potentially uncovering yet another potential mitotic function of Ran (Muhlhausser and Kutay 2007). However, it should be kept in mind that nuclear disassembly can also occur in the absence of microtubules in vitro (Lenart

Organization of the Nuclear Membrane

et al. 2003), indicating that dynein-dependent rupturing of the NE may not be essential. STRUCTURE AND FUNCTION OF NE COMPONENTS DURING MITOSIS

As mentioned above, chromosomes are essentially membrane-free between metaphase and anaphase. EM and fluorescence microscopy has provided compelling evidence that the NE is reabsorbed into the ER (Ellenberg et al. 1997; Anderson and Hetzer 2007; Anderson and Hetzer 2008a; Anderson et al. 2009a; Lu et al. 2009). For instance, fluorescence timelapse microscopy in living cells has shown that NE proteins such as the nucleoporins gp210, Ndc1, and Pom121 are partitioned into the mitotic ER (Ellenberg et al. 1997; Puhka et al. 2007). Likewise, INM proteins can be found in the ER during mitosis (Puhka et al. 2007). The topology of the mitotic ER, however, remains controversial. Three-dimensional modeling of the ER by electron tomography suggested that the mitotic ER remains an intact network of membrane tubules that is essentially free of sheets (Puhka et al. 2007). This is supported by recent data suggesting that the intrinsic propensity of the ER to oscillate between tubules and sheets is used during mitosis and affects the fate of the NE during the mitosis of C. elegans (Audhya et al. 2007). However, compelling data from nematodes and mammalian cells challenges the view of an entirely tubular ER and instead suggest that the ER is largely composed of membrane sheets. For instance, it was shown that the rough ER of metaphase HeLa cells appears to be exclusively cisternal and concentrates at the cell cortex, often following the contours of the plasma membrane (McCullough and Lucocq 2005). In a study using three-dimensional (3D) reconstructions of living cells, the ER appeared entirely formed of cisternae and very few tubules were observed (Lu et al. 2009). Similar results were obtained in C. elegans (Poteryaev et al. 2005). Determining the exact nature of the ER and the disassembled NE components remain critical future goals, because this has important implications for NE formation.

NE PROTEINS HAVE MITOTIC FUNCTIONS

Although the dynamic organization of the ER membranes remains elusive, it has become clear that disassembled NE components fulfill critical functions in mitosis. For instance, the multimeric Nup107-160 complex, which in interphase is essential for pore assembly and function (D’Angelo et al. 2006), and Nup358, which shows SUMO E3 ligase activity that modifies RanGAP, Ran’s GTPase activating protein (Joseph et al. 2002; Pichler et al. 2002), can both be detected in association with kinetochores in mitosis. In addition, ELYS/MEL-28 and Nup107-160 are also found with spindle poles and proximal microtubules (Loiodice et al. 2004; Galy et al. 2006; Rasala et al. 2006). This association is not critical in Drosophila, suggesting that the role of Nups in kinetochore function is a relatively late evolutionary event (Katsani et al. 2008). However, in mammalian cells, the absence of the Nup107-160 complex perturbs bipolar spindle formation (Orjalo et al. 2006), and inhibits Nup358 recruitment to microtubule-bound kinetochores (Salina et al. 2003; Joseph et al. 2004). Recently it was shown that the kinesin-binding domain (KBD) of Nup358 associates with kinesin-1, KIF5B/ KIF5C. Interestingly, the KBD stimulates the ATPase activity of KIF5B in the presence of microtubules, thereby increasing its activity (Cho et al. 2009). The Nup107-160 complex might have an independent role at kinetochores because in a Xenopus in vitro spindle assembly assay, the Nup358/SUMO-RanGAP complex does not associate with kinetochores (Arnaoutov and Dasso 2005). Because mNup133 (a mNup107160 member) and CENP-F (a kinetochore protein) interact with each other, this function might be linked to the dynein partners Nde1 and Nde1l, which also bind CENP-F (Vergnolle and Taylor 2007; Zuccolo et al. 2007). Additional roles for other NE proteins in cell-cycle and mitotic progression have also been reported. Depletion of lamin B, a type V intermediate-filament protein and a component of the nuclear lamina, also results in mitotic spindle defects (Zheng and Tsai 2006).

M.W. Hetzer

Interestingly, the spindle-associated lamin B appears to be present in a membranous, matrixlike network and seems to facilitate spindle microtubule organization in a dynein-dependent manner (Tsai et al. 2006; Ma et al. 2009). Although the mechanistic details of spindlematrix function with respect to lamin B remain to be determined, these results contribute to the emerging paradigm for structural components of the NE having roles in mitosis. This idea is further supported by the recent finding of spindle-associated membrane protein 1 (Samp1), which during interphase is localized to the inner nuclear membrane, specifically localized to the polar regions of the mitotic spindle (Buch et al. 2009). Depletion of Samp1 expression resulted in separation of centrosomes from the NE, indicating that it is functionally connected to the cytoskeleton. This provides evidence for the existence of interactions between mitotic ER membranes with the spindle. RE-ESTABLISHING ORDER: REFORMATION OF THE NE

NE formation has been studied in various cellfree systems, cell types, and organisms (Burke and Ellenberg 2002; Hetzer et al. 2005; Anderson and Hetzer 2008c). Because membranes are typically delicate structures that are easily disrupted during cell fractionation and fixation, it is easy to see how data obtained from different systems lead to the postulation of sometimes opposing models. There are essentially two different ideas about how the NE is formed: (1) by vesicle fusion and (2) by reshaping of ER into NE sheets. Based on biochemical data and EM observations, it was initially proposed that the NE fragments into NE vesicles (Collas and Courvalin 2000). This idea is derived from cell-free systems mainly of Xenopus, starfish, and Drosophila (Lohka and Masui 1983; Newport 1987). Frog egg extracts have the advantage of containing large stockpiles of disassembled NE components (Newport and Spann 1987) and nuclei can be assembled within 60 minutes in a test tube by mixing a chromatin source, cytosol, and membranes. These artificial nuclei are

capable of nucleocytoplasmic transport, DNA replication, as well as NEBD (Sheehan et al. 1988; Gant and Wilson 1997; Hetzer et al. 2002). The idea of vesicle fusion as the principal mechanism of NE formation was further supported by findings that GTPgS, a nonhydrolyzable GTP analog, blocks NE formation and results in assembly intermediates covered with chromatin-bound vesicles (Boman et al. 1992; Hetzer et al. 2001). Interestingly, depletion of the small GTPase Ran mimicked the effect of GTPgS addition (Hetzer et al. 2000; Zhang and Clarke 2000). Because the Ran exchange factor RCC1 (Ohtsubo et al. 1989) is stably associated with chromatin, it was proposed that high levels of RanGTP are generated around chromatin that provide a spatial signal for chromatin-associated processes such as NE formation and NPC assembly (Bilbao-Cortes et al. 2002; Hetzer et al. 2002; Hutchins et al. 2004). It was subsequently shown that Ran releases the nuclear transport receptor Importin b (Nachury et al. 2001; Harel et al. 2003a; Harel and Forbes 2004) from nucleoporins and thereby triggers NPC assembly (Harel et al. 2003a; Walther et al. 2003b). A similar mechanism might be required for the formation of a closed NE, but the targets of Importin b remain unknown. Corroborating the in vitro data, Ran has been shown to be required for NE formation in C. elegans (Askjaer et al. 2002) and yeast (Ryan et al. 2003). Although egg extracts provide a unique experimental system to study ER and NE reconstitution, it is important to realize that the assembly reactions are initiated with isolated membranes, which are in a highly fragmented state that does not represent the intact in vivo organization of the mitotic ER. Recent evidence shows that an intact NE can also form from preformed ER in vitro (Anderson and Hetzer 2007). Strikingly, nuclear assembly from a preformed ER is insensitive to fusion inhibitors such as GTPgS, ATPgS, or antibodies that inhibit the function of the AAA-ATPase p97. In addition, factors that are essential for ER tubule formation, such as ATP (Dreier and Rapoport 2000) and reticulons (Voeltz et al. 2006), only block nuclear assembly if added

Organization of the Nuclear Membrane

before the ER network is organized (Anderson and Hetzer 2007). The results strongly support the notion that the ER is the source of NE membranes. According to this idea, the observed inhibition of nuclear assembly by fusion inhibitors and the role of SNARE proteins in NE formation are likely to be an indirect effect of blocked ER reconstitution (Baur et al. 2007). Although these experiments show that the NE can form from an ER network in vitro, they do not address the question of how NE formation occurs in intact cells. High resolution imaging in intact cells has revealed ER tubules contacting chromatin at early stages of NE formation. Shortly after contact, the rapid coating of chromatin by ER membranes can be observed (Anderson and Hetzer 2007). Recent data suggest that endogenous concentrations of NE-promoting transmembrane proteins are ratelimiting for nuclear assembly. Because NE formation is also affected by endogenous levels of the ER-shaping reticulon proteins that slow NE formation, these findings suggest a tug of war between reticulons and their membrane-curving activity and NE proteins, which promote membrane attachment and spreading around chromatin (Anderson and Hetzer 2008a). The massive membrane-restructuring event that results in the formation of the sheet-like NE involves functionally diverse groups of NE proteins that collaborate during mitosis to tether membranes to the chromatin surface and thereby drive NE formation (Anderson et al. 2009b). Recent findings that DNA-binding activity of some INM proteins is required for NE formation in vitro (Ulbert et al. 2006) and that membrane sheets formed efficiently on protein-free immobilized DNA in vitro support this idea. In addition, NE formation also seems to involve the ability of several INM proteins to bind to chromatin factors. For example, it has been shown that the integral INM protein LBR, which binds to heterochromatin-binding protein 1 (HP1), is required for targeting and anchoring NE membranes to chromatin in vitro (Collas et al. 1996; Pyrpasopoulou et al. 1996). In a similar fashion, the barrier-to-autointegration factor (BAF), a chromatin-binding protein (Segura-Totten et al. 2002) and its kinase Vrk have recently been shown to play a direct role in

NE formation by recruiting LEM domain proteins to chromatin (Gorjanacz et al. 2007). Vrk seems to be a key regulator in this process because BAF phosphorylation reduces chromatin binding and interactions with LEM domain proteins such as emerin (Bengtsson and Wilson 2004; Hirano et al. 2005). Thus, NE formation is likely to involve a complex interplay of transmembrane NE proteins distributed into mitotic ER with the reorganizing chromatin. ROLE OF CHROMATIN IN NE FORMATION

Recent studies of cell-free nuclear assembly systems suggest that initial ER/chromatin contacts are mediated via tubule ends. In a second step, these tethered ER tubules are reorganized into flat nuclear membrane sheets by DNA-binding-NE-specific membrane proteins (Ulbert et al. 2006; Anderson and Hetzer 2007). In cells, the ER appears to be largely cisternal (Lu et al. 2009) and thus it is likely that pre-existing membrane sheets are directly tethered to the chromatin surface in vivo. This is consistent with the finding that overexpression of reticulons delays NE formation, whereas their knock down accelerates NE formation (Anderson and Hetzer 2008a). During NE formation, chromatin undergoes a series of conformational changes, from a state of maximal chromatin compaction in late anaphase, right before NE formation (MoraBermudez et al. 2007), to transcription- and replication-competent decondensation in interphase. Anaphase compaction requires the kinesin-like DNA binding protein (Kid), which ensures the formation of a compact chromosome cluster during anaphase and the properenclosure of the segregated chromatin mass into a single nucleus (Ohsugi et al. 2008). Interestingly, Kid loading onto anaphase chromosomes is dependent on Importin b (Tahara et al. 2008), adding to the growing number of mitotic processes regulated by this transport receptor. Recently, a compelling case has been made that mechanistically links NE formation and chromatin decondensation. The hexameric ATPase Cdc48/p97, which was implicated in membrane fusion (Kondo et al. 1997) and

M.W. Hetzer

ubiquitin-dependent processes (Ye et al. 2003), extracts Aurora B from chromatin, which results in its inactivation and subsequent chromatin decondensation. Interestingly, the inhibition of Cdc48/p97 blocked NE formation, suggesting that chromatin decondensation is required for NE formation (Ramadan et al. 2007), possibly by opening chromatin structure. It is tempting to speculate that this open chromatin structure provides binding sites for INM proteins and therefore drives NE formation. Concomitantly, with the coating of chromatin by the NE, NPCs assemble in the reforming nuclei. This fascinating example of protein selfassembly is coordinated by the stepwise recruitment of a subset of NPC proteins to chromatin (Dultz et al. 2008) and some progress has been made in determining how this process occurs. A protein called Mel-28/ELYS, which was identified in a screen in C. elegans for factors involved in pronuclear formation, is critical for the association of the Nup107/160 complex to chromatin (Rasala et al. 2006; Franz et al. 2007). Because Mel-28/ELYS contains an AT-hook domain (Kimura et al. 2002), a likely scenario is that this step occurs by direct binding to DNA. Interestingly, RanGTP stimulates Mel-28/ELYS recruitment (Franz et al. 2007), presumably by releasing Importin b from one of the Nup107/ 160/ELys components (Hetzer et al. 2005). Although other nucleoporins have been implicated in pore assembly, their exact role remains to be determined. For instance, a complex of Nup53 and Nup155 has recently been shown to be essential for NE formation in nematodes and vertebrates (Franz et al. 2005; HawrylukGara et al. 2008); however, how the membraneassociated Nup53 coordinates interactions between chromatin, membranes, and soluble Nup155 remains unclear. A group of nucleoporins have been shown to be essential for pore assembly (Hetzer et al. 2005), the largest subcomplex being the Nup107/160 complex, whose depletion results in NPC-free nuclear membranes (Harel et al. 2003b; Walther et al. 2003a). In vertebrates, two transmembrane nucleoporins have also been shown to participate in NE formation, Ndc1 and POM121 (Mansfeld et al. 2006; Stavru

et al. 2006; Anderson et al. 2009b). The third known transmembrane Nup, gp210, is not expressed in all cell types and thus is unlikely to be essential for pore assembly (Eriksson et al. 2004). Whereas the molecular role of these scaffold Nups is unclear, an interesting link between POM121 and the Nup107/160 complex has been made and nuclear membrane formation might actually be linked to pore assembly by a poorly understood checkpoint by which the Nup107/160 complex “senses” nuclear membranes (Antonin et al. 2005). REMODELING THE INTERPHASE NE

In order for cells to properly progress through multiple cell divisions, the number of NPCs doubles in interphase (Maul et al. 1972). It is unclear if the increase simply reflects the necessity to double pores for daughter cells of the next division cycle, or whether an increase in NPC number is important for interphase cellcycle progression, e.g., efficient replication or transcription. Interphase nuclear pore assembly is particularly interesting because NPC formation occurs from both sides of the NE (D’Angelo et al. 2006). Thus, the question arises, what is the mechanism of communication between the double membranes? Interestingly, Nup133, a member of the Nup107/160 complex, contains an ALPS-like motif including an amphipathic a-helical domain that has been shown to act as a membrane curvature sensor in vitro (Drin et al. 2007). It is possible that this domain is involved in targeting the Nup107/160 complex to membranes during NE formation. A prediction from this idea is that the membrane hole is formed before Nup107/160 is recruited. Whereas the fusion of INM and ONM remains elusive in mammalian cells, significant progress has been made in deciphering this process in yeast. Genetic studies in S. cerevisiae revealed a network of protein– protein interactions that appears to initiate NPC formation. The nucleoporins Nup59/53 and the integral pore membrane nucleoporins Pom152 and Pom34 tether Nup170 and a third integral membrane nucleoporin Ndc1 to the NE at new assembly sites (Onischenko et al. 2009). Furthermore, depletion

Organization of the Nuclear Membrane

of Nup170 and its homolog Nup157 causes the accumulation of NPC-like structures in the INM and at cytoplasmic foci rather than properly localized to nuclear pores spanning the NE. It is interesting to note that the yeast reticulons and Yop1 not only display genetic interactions with the Poms, but also seem to play an essential role in the formation of new NPCs (Dawson et al. 2009). Similar to the depletion of Nup170, in the absence of reticulons, NPClike intermediates also accumulate in an aberrant manner in the INM and ONM of the NE. The current idea is that a complex consisting of the three membrane-spanning nucleoporins Pom152, Pom34, and Ndc1 marks a new assembly site by joining or juxtaposing ONM and INM, which is consistent with the observation that NPC assembly proceeds from both sides of the NE (D’Angelo et al. 2006). According to this idea, Nup53/59 and the reticulons Yop1/ Dbp1 are recruited to the nuclear membrane and inserted into the outer leaflet of the membrane using monotopic membrane insertion domains, inducing or stabilizing local curvature to the membrane. However, this idea was recently challenged on theoretical grounds, arguing that reticulons induce positive membrane curvature (e.g., giving rise to the convex surface of a tubule), but not negative curvature, which is likely to occur during NPC assembly. Instead, it was proposed that reticulons might participate in the local enrichment of negative curvatureinducing proteins involved in NE formation (Antonin 2009). Whether transmembrane nucleoporins, reticulons, oran unidentified machinery induces the fusion of INM and ONM during NPC assembly remains an open question. During interphase, the NE expands considerably and thus requires the supply of additional nuclear membrane and proteins. In vitro nuclear expansion is blocked by disrupting the connection of nuclei with the peripheral ER (D’Angelo et al. 2006), suggesting that membranes feed into the ONM via connections with ER tubules. Growth of the INM requires passage of membrane components through the fusion sites with the ONM at the NPCs. The current view is that INM proteins are retained once they reach the INM by interactions

with either the lamina or chromatin. How INM proteins are targeted in metazoa is less clear. One model suggested that ATP-driven changes in nucleoporin interactions might allow membrane proteins to travel across the NPC (Ohba et al. 2004). In yeast, integral INM proteins have been shown to directly interact with specific nucleoporins and transport receptors to promote their movement past the NPC (King et al. 2006). By the end of interphase, the NE has undergone major changes in protein composition and is ready to break down at the onset of mitosis to start a new life cycle in the two daughter cells. During interphase, NE connections with the cytoskeleton have to withstand external forces. Recent work has suggested that SUN and KASH domain proteins play key roles in this process (Adam 2001). KASH domain proteins interact with centrosomes/SPBs or cytoskeletal elements in the cytoplasm and bind to SUN domain proteins in the INM to connect the NE to structures on the outside of the nucleus. The nuclear lamina could serve in metazoa to distribute the forces across a broader area (Stewart et al. 2007) (Roux and Burke 2007). CONCLUDING REMARKS

The NE is well known for its role in shielding the nuclear genome from cytoplasmic components and mediating nucleocytoplasmic transport. Less well understood is its dynamic behavior in dividing cells and its role in the distribution of the genetic material. Furthermore, it has become clear that the NE might actively control the organization of the genome and directly regulate gene expression. This opens some interesting opportunities for future research activities. For instance, the nature of most NE proteinchromatin interactions remain uncharacterized and many of the INM proteins are expressed in a tissue-specific manner. This raises the important question of whether NE composition is a critical component of cell fate determination. If NE proteins control chromatin organization, then is there a link between NE formation and nuclear organization? In other words, are the NE-chromatin contact points that are established

M.W. Hetzer

in anaphase/telophase maintained in interphase? Because many NE proteins are linked to a large number of diseases, answers to these questions are likely to have a direct impact on human health.

ACKNOWLEDGMENTS

I would like to thank members in my laboratory, in particular, Maximiliano D’Angelo, Roberta Schulte, and Jesse Vargas for critically reading the manuscript. This work was supported by the National Institute of General Medical Sciences (award number R01GM073994).

REFERENCES Adam SA. 2001. The nuclear pore complex. Genome Biol 2: REVIEWS0007. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695 –701. Anderson DJ, Hetzer MW. 2007. Nuclear envelope formation by chromatin-mediated reorganization of the endoplasmic reticulum. Nat Cell Biol 9: 1160–1166. Anderson DJ, Hetzer MW. 2008a. Reshaping of the endoplasmic reticulum limits the rate for nuclear envelope formation. J Cell Biol 182: 911 –924. Anderson DJ, Hetzer MW. 2008b. Shaping the endoplasmic reticulum into the nuclear envelope. J Cell Sci 121: 137–142. Anderson DJ, Hetzer MW. 2008c. The life cycle of the metazoan nuclear envelope. Curr Opin Cell Biol 20: 386 –392. Anderson DJ, Vargas J, Hisao J, Hetzer MW. 2009a. Recruitment of functionally distinct membrane proteins to chromatin mediates nuclear envelope formation in vivo. J Cell Biol 186: 183 –191. Anderson DJ, Vargas JD, Hsiao JP, Hetzer MW. 2009b. Recruitment of functionally distinct membrane proteins to chromatin mediates nuclear envelope formation in vivo. J Cell Biol 186: 183 –191. Antonin W. 2009. Nuclear envelope: Membrane bending for pore formation? Curr Biol 19: R410–412. Antonin W, Franz C, Haselmann U, Antony C, Mattaj IW. 2005. The integral membrane nucleoporin pom121 functionally links nuclear pore complex assembly and nuclear envelope formation. Mol Cell 17: 83–92. Arnaoutov A, Dasso M. 2005. Ran-GTP regulates kinetochore attachment in somatic cells. Cell Cycle 4: 1161–1165. Askjaer P, Galy V, Hannak E, Mattaj IW. 2002. Ran GTPase cycle and importins a and b are essential for spindle formation and nuclear envelope assembly in living Caenorhabditis elegans embryos. Mol Biol Cell 13: 4355–4370.

Audhya A, Desai A, Oegema K. 2007. A role for Rab5 in structuring the endoplasmic reticulum. J Cell Biol 178: 43– 56. Baur T, Ramadan K, Schlundt A, Kartenbeck J, Meyer HH. 2007. NSF- and SNARE-mediated membrane fusion is required for nuclear envelope formation and completion of nuclear pore complex assembly in Xenopus laevis egg extracts. J Cell Sci 120: 2895– 2903. Beaudouin J, Gerlich D, Daigle N, Eils R, Ellenberg J. 2002. Nuclear envelope breakdown proceeds by microtubuleinduced tearing of the lamina. Cell 108: 83– 96. Beck M, Forster F, Ecke M, Plitzko JM, Melchior F, Gerisch G, Baumeister W, Medalia O. 2004. Nuclear pore complex structure and dynamics revealed by cryoelectron tomography. Science 306: 1387–1390. Beck M, Lucic V, Forster F, Baumeister W, Medalia O. 2007. Snapshots of nuclear pore complexes in action captured by cryo-electron tomography. Nature 449: 611– 615. Bengtsson L, Wilson KL. 2004. Multiple and surprising new functions for emerin, a nuclear membrane protein. Curr Opin Cell Biol 16: 73–79. Bilbao-Cortes D, Hetzer M, Langst G, Becker PB, Mattaj IW. 2002. Ran binds to chromatin by two distinct mechanisms. Curr Biol 12: 1151– 1156. Boehmer T, Jeudy S, Berke IC, Schwartz TU. 2008. Structural and functional studies of Nup107/Nup133 interaction and its implications for the architecture of the nuclear pore complex. Mol Cell 30: 721 –731. Boman AL, Delannoy MR, Wilson KL. 1992. GTP hydrolysis is required for vesicle fusion during nuclear envelope assembly in vitro. J Cell Biol 116: 281 –294. Brohawn SG, Partridge JR, Whittle JR, Schwartz TU. 2009. The nuclear pore complex has entered the atomic age. Structure 17: 1156–1168. Buch C, Lindberg R, Figueroa R, Gudise S, Onischenko E, Hallberg E. 2009. An integral protein of the inner nuclear membrane localizes to the mitotic spindle in mammalian cells. J Cell Sci 122: 2100– 2107. Burke B, Ellenberg J. 2002. Remodelling the walls of the nucleus. Nat Rev Mol Cell Biol 3: 487– 497. Byers B. 1981. Cytology of the yeast life cycle. In Molecular Biology of the Yeast Saccharomyces. In Life Cycle and Inheritance (ed. Strathern E.W.J.a.J.R.B. J. N. Strathern), pp. 59– 96. Cold Spring Harbor Laboratory Press, NY. Chakraborty P, Wang Y, Wei JH, van Deursen J, Yu H, Malureanu L, Dasso M, Forbes DJ, Levy DE, Seemann J, et al. 2008. Nucleoporin levels regulate cell cycle progression and phase-specific gene expression. Dev Cell 15: 657– 667. Chase D, Serafinas C, Ashcroft N, Kosinski M, Longo D, Ferris DK, Golden A. 2000. The polo-like kinase PLK-1 is required for nuclear envelope breakdown and the completion of meiosis in Caenorhabditis elegans. Genesis 26: 26– 41. Cho KI, Yi H, Desai R, Hand AR, Haas AL, Ferreira PA. 2009. RANBP2 is an allosteric activator of the conventional kinesin-1 motor protein, KIF5B, in a minimal cell-free system. EMBO Rep 10: 480 –486. Cohen TV, Hernandez L, Stewart CL. 2008. Functions of the nuclear envelope and lamina in development and disease. Biochem Soc Trans 36: 1329–1334.

Organization of the Nuclear Membrane Collas P. 1999. Sequential PKC- and Cdc2-mediated phosphorylation events elicit zebrafish nuclear envelope disassembly. J Cell Sci 112: 977– 987. Collas I, Courvalin JC. 2000. Sorting nuclear membrane proteins at mitosis. Trends Cell Biol 10: 5 –8. Collas P, Courvalin JC, Poccia D. 1996. Targeting of membranes to sea urchin sperm chromatin is mediated by a lamin B receptor-like integral membrane protein. J Cell Biol 135: 1715– 1725. Courvalin JC, Segil N, Blobel G, Worman HJ. 1992. The lamin B receptor of the inner nuclear membrane undergoes mitosis-specific phosphorylation and is a substrate for p34cdc2-type protein kinase. J Biol Chem 267: 19035–19038. D’Angelo MA, Hetzer MW. 2006. The role of the nuclear envelope in cellular organization. Cell Mol Life Sci 63: 316–332. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456–466. D’Angelo MA, Anderson DJ, Richard E, Hetzer MW. 2006. Nuclear pores form de novo from both sides of the nuclear envelope. Science 312: 440–443. D’Angelo MA, Raices M, Panowski SH, Hetzer MW. 2009. Age-dependent deterioration of nuclear pore complexes causes a loss of nuclear integrity in postmitotic cells. Cell 136: 284– 295. Dawson TR, Lazarus MD, Hetzer MW, Wente SR. 2009. ER membrane-bending proteins are necessary for de novo nuclear pore formation. J Cell Biol 184: 659–675. De Souza CP, Osmani AH, Hashmi SB, Osmani SA. 2004. Partial nuclear pore complex disassembly during closed mitosis in Aspergillus nidulans. Curr Biol 14: 1973– 1984. Dechat T, Gotzmann J, Stockinger A, Harris CA, Talle MA, Siekierka JJ, Foisner R. 1998. Detergent-salt resistance of LAP2a in interphase nuclei and phosphorylationdependent association with chromosomes early in nuclear assembly implies functions in nuclear structure dynamics. EMBO J 17: 4887– 4902. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832 –853. Dorner D, Gotzmann J, Foisner R. 2007. Nucleoplasmic lamins and their interaction partners, LAP2a, Rb, and BAF, in transcriptional regulation. FEBS J 274: 1362– 1373. Dreier L, Rapoport TA. 2000. In vitro formation of the endoplasmic reticulum occurs independently of microtubules by a controlled fusion reaction. J Cell Biol 148: 883–898. Dreger M, Otto H, Neubauer G, Mann M, Hucho F. 1999. Identification of phosphorylation sites in native lamina-associated polypeptide 2 b. Biochemistry 38: 9426– 9434. Drin G, Casella JF, Gautier R, Boehmer T, Schwartz TU, Antonny B. 2007. A general amphipathic a-helical motif for sensing membrane curvature. Nat Struct Mol Biol 14: 138–146. Dultz E, Zanin E, Wurzenberger C, Braun M, Rabut G, Sironi L, Ellenberg J. 2008. Systematic kinetic analysis of

mitotic dis- and reassembly of the nuclear pore in living cells. J Cell Biol 180: 857 –865. Ellenberg J, Siggia ED, Moreira JE, Smith CL, Presley JF, Worman HJ, Lippincott-Schwartz J. 1997. Nuclear membrane dynamics and reassembly in living cells: Targeting of an inner nuclear membrane protein in interphase and mitosis. J Cell Biology 138: 1193–1206. Eriksson C, Rustum C, Hallberg E. 2004. Dynamic properties of nuclear pore complex proteins in gp210 deficient cells. FEBS Lett 572: 261 –265. Franz C, Askjaer P, Antonin W, Iglesias CL, Haselmann U, Schelder M, de Marco A, Wilm M, Antony C, Mattaj IW. 2005. Nup155 regulates nuclear envelope and nuclear pore complex formation in nematodes and vertebrates. EMBO J 24: 3519–3531. Franz C, Walczak R, Yavuz S, Santarella R, Gentzel M, Askjaer P, Galy V, Hetzer M, Mattaj IW, Antonin W. 2007. MEL-28/ELYS is required for the recruitment of nucleoporins to chromatin and postmitotic nuclear pore complex assembly. EMBO Rep 8: 165– 172. Fridkin A, Penkner A, Jantsch V, Gruenbaum Y. 2009. SUN-domain and KASH-domain proteins during development, meiosis and disease. Cell Mol Life Sci 66: 1518–1533. Galy V, Antonin W, Jaedicke A, Sachse M, Santarella R, Haselmann U, Mattaj I. 2008. A role for gp210 in mitotic nuclear-envelope breakdown. J Cell Sci 121: 317– 328. Galy V, Askjaer P, Franz C, Lopez-Iglesias C, Mattaj IW. 2006. MEL-28, a novel nuclear-envelope and kinetochore protein essential for zygotic nuclear-envelope assembly in C. elegans. Curr Biol 16: 1748–1756. Gant TM, Wilson KL. 1997. Nuclear assembly. Annu Rev Cell Dev Biol 13: 669 –695. Golden A, Liu J, Cohen-Fix O. 2009. Inactivation of the C. elegans lipin homolog leads to ER disorganization and to defects in the breakdown and reassembly of the nuclear envelope. J Cell Sci 122: 1970–1978. Gong D, Pomerening JR, Myers JW, Gustavsson C, Jones JT, Hahn AT, Meyer T, Ferrell JE Jr. 2007. Cyclin A2 regulates nuclear-envelope breakdown and the nuclear accumulation of cyclin B1. Curr Biol 17: 85– 91. Gorjanacz M, Mattaj IW. 2009. Lipin is required for efficient breakdown of the nuclear envelope in Caenorhabditis elegans. J Cell Sci 122: 1963–1969. Gorjanacz M, Klerkx EP, Galy V, Santarella R, Lopez-Iglesias C, Askjaer P, Mattaj IW. 2007. Caenorhabditis elegans BAF-1 and its kinase VRK-1 participate directly in postmitotic nuclear envelope assembly. EMBO J 26: 132– 143. Goss VL, Hocevar BA, Thompson LJ, Stratton CA, Burns DJ, Fields AP. 1994. Identification of nuclear bII protein kinase C as a mitotic lamin kinase. J Biol Chem 269: 19074– 19080. Griffis ER, Altan N, Lippincott-Schwartz J, Powers MA. 2002. Nup98 is a mobile nucleoporin with transcriptiondependent dynamics. Mol Biol Cell 13: 1282– 1297. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol 6: 21–31. Gruenbaum Y, Wilson KL, Harel A, Goldberg M, Cohen M. 2000. Review: nuclear lamins– structural proteins with fundamental functions. J Struct Biol 129: 313–323.

M.W. Hetzer Guttinger S, Laurell E, Kutay U. 2009. Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat Rev Mol Cell Biol 10: 178– 191. Harel A, Forbes DJ. 2004. Importin b: Conducting a much larger cellular symphony. Mol Cell 16: 319 –330. Harel A, Chan RC, Lachish-Zalait A, Zimmerman E, Elbaum M, Forbes DJ. 2003a. Importin b negatively regulates nuclear membrane fusion and nuclear pore complex assembly. Mol Biol Cell 14: 4387–4396. Harel A, Orjalo AV, Vincent T, Lachish-Zalait A, Vasu S, Shah S, Zimmerman E, Elbaum M, Forbes DJ. 2003b. Removal of a single pore subcomplex results in vertebrate nuclei devoid of nuclear pores. Mol Cell 11: 853–864. Hawryluk-Gara LA, Platani M, Santarella R, Wozniak RW, Mattaj IW. 2008. Nup53 is required for nuclear envelope and nuclear pore complex assembly. Mol Biol Cell 10.1091/mbc.E07-08-0820. Heessen S, Fornerod M. 2007. The inner nuclear envelope as a transcription factor resting place. EMBO Rep 8: 914–919. Hetzer M, Gruss OJ, Mattaj IW. 2002. The Ran GTPase as a markerof chromosome position in spindle formation and nuclear envelope assembly. Nat Cell Biol 4: E177– 184. Hetzer M, Walther TC, Mattaj IW. 2005. Pushing the envelope: Structure, function, and dynamics of the nuclear periphery. Annu Rev Cell Dev Biol 21: 367– 380. Hetzer M, Bilbao-Cortes D, Walther TC, Gruss OJ, Mattaj IW. 2000. GTP hydrolysis by Ran is required for nuclear envelope assembly. Mol Cell 5: 1013–1024. Hetzer M, Meyer HH, Walther TC, Bilbao-Cortes D, Warren G, Mattaj IW. 2001. Distinct AAA-ATPase p97 complexes function in discrete steps of nuclear assembly. Nat Cell Biol 3: 1086–1091. Heywood P. 1978. Ultrastructure of mitosis in the chloromonadophycean alga Vacuolaria virescens. J Cell Sci 31: 37–51. Hirano Y, Segawa M, Ouchi FS, Yamakawa Y, Furukawa K, Takeyasu K, Horigome T. 2005. Dissociation of emerin from barrier-to-autointegration factor is regulated through mitotic phosphorylation of emerin in a Xenopus egg cell-free system. J Biol Chem 280: 39925–39933. Hutchins JR, Moore WJ, Hood FE, Wilson JS, Andrews PD, Swedlow JR, Clarke PR. 2004. Phosphorylation regulates the dynamic interaction of RCC1 with chromosomes during mitosis. Curr Biol 14: 1099– 1104. Joseph J, Liu ST, Jablonski SA, Yen TJ, Dasso M. 2004. The RanGAP1-RanBP2 complex is essential for microtubulekinetochore interactions in vivo. Curr Biol 14: 611 –617. Joseph J, Tan SH, Karpova TS, McNally JG, Dasso M. 2002. SUMO-1 targets RanGAP1 to kinetochores and mitotic spindles. J Cell Biol 156: 595– 602. Katsani KR, Karess RE, Dostatni N, Doye V. 2008. In vivo dynamics of Drosophila nuclear envelope components. Mol Biol Cell 19: 3652–3666. Kimura N, Takizawa M, Okita K, Natori O, Igarashi K, Ueno M, Nakashima K, Nobuhisa I, Taga T. 2002. Identification of a novel transcription factor, ELYS, expressed predominantly in mouse foetal haematopoietic tissues. Genes Cells 7: 435 –446.

King MC, Lusk CP, Blobel G. 2006. Karyopherin-mediated import of integral inner nuclear membrane proteins. Nature 442: 1003– 1007. Kite GL. 1913. The relative permeability of the surface and interior portions of the cytoplasm of animal and plant cells. Biol Bull 25: 1–7. Kondo H, Rabouille C, Newman R, Levine TP, Pappin D, Freemont P, Warren G. 1997. p47 is a cofactor for p97-mediated membrane fusion. Nature 388: 75– 78. Kutay U, Hetzer MW. 2008. Reorganization of the nuclear envelope during open mitosis. Curr Opin Cell Biol 20: 669 –677. Lee KK, Gruenbaum Y, Spann P, Liu J, Wilson KL. 2000. C. elegans nuclear envelope proteins emerin, MAN1, lamin, and nucleoporins reveal unique timing of nuclear envelope breakdown during mitosis. Mol Biol Cell 11: 3089–3099. Lenart P, Rabut G, Daigle N, Hand AR, Terasaki M, Ellenberg J. 2003. Nuclear envelope breakdown in starfish oocytes proceeds by partial NPC disassembly followed by a rapidly spreading fenestration of nuclear membranes. J Cell Biol 160: 1055– 1068. Liu J, Prunuske AJ, Fager AM, Ullman KS. 2003. The COPI complex functions in nuclear envelope breakdown and is recruited by the nucleoporin Nup153. Dev Cell 5: 487 –498. Lohka MJ, Masui Y. 1983. Formation in vitro of sperm pronuclei and mitotic chromosomes induced by amphibian ooplasmic components. Science 220: 719–721. Loiodice I, Alves A, Rabut G, Van Overbeek M, Ellenberg J, Sibarita JB, Doye V. 2004. The entire Nup107–160 complex, including three new members, is targeted as one entity to kinetochores in mitosis. Mol Biol Cell 15: 3333–3344. Lu L, Ladinsky MS, Kirchhausen T. 2009. Cisternal organization of the endoplasmic reticulum during mitosis. Mol Biol Cell 15: 3471– 3480. Ma L, Tsai MY, Wang S, Lu B, Chen R, Iii JR, Zhu X, Zheng Y. 2009. Requirement for Nudel and dynein for assembly of the lamin B spindle matrix. Nat Cell Biol 11: 247–256. Mansfeld J, Guttinger S, Hawryluk-Gara LA, Pante N, Mall M, Galy V, Haselmann U, Muhlhausser P, Wozniak RW, Mattaj IW, et al. 2006. The conserved transmembrane nucleoporin NDC1 is required for nuclear pore complex assembly in vertebrate cells. Mol Cell 22: 93–103. Mattout A, Dechat T, Adam SA, Goldman RD, Gruenbaum Y. 2006. Nuclear lamins, diseases and aging. Curr Opin Cell Biol 18: 335– 341. Maul GG, Maul HM, Scogna JE, Lieberman MW, Stein GS, Hsu BY, Borun TW. 1972. Time sequence of nuclear pore formation in phytohemagglutinin-stimulated lymphocytes and in HeLa cells during the cell cycle. J Cell Biol 55: 433– 447. McCullough S, Lucocq J. 2005. Endoplasmic reticulum positioning and partitioning in mitotic HeLa cells. J Anat 206: 415– 425. Miller BR, Forbes DJ. 2000. Purification of the vertebrate nuclear pore complex by biochemical criteria. Traffic 1: 941 –951. Mora-Bermudez F, Gerlich D, Ellenberg J. 2007. Maximal chromosome compaction occurs by axial shortening in

Organization of the Nuclear Membrane anaphase and depends on Aurora kinase. Nat Cell Biol 9: 822–831. Mounkes L, Kozlov S, Burke B, Stewart CL. 2003. The laminopathies: Nuclear structure meets disease. Curr Opin Genet Dev 13: 223 –230. Muchir A, Worman HJ. 2004. The nuclear envelope and human disease. Physiology 19: 309–314. Muhlhausser P, Kutay U. 2007. An in vitro nuclear disassembly system reveals a role for the RanGTPase system and microtubule-dependent steps in nuclear envelope breakdown. J Cell Biol 178: 595– 610. Nachury MV, Maresca TJ, Salmon WC, Waterman-Storer CM, Heald R, Weis K. 2001. Importin b is a mitotic target of the small GTPase Ran in spindle assembly. Cell 104: 95–106. Neilan EG. 2009. Laminopathies, other progeroid disorders, and aging: Common pathogenic themes and possible treatments. Am J Med Genet A 149A: 563– 566. Newport J. 1987. Nuclear reconstitution in vitro: Stages of assembly around protein-free DNA. Cell 48: 205 –217. Newport J, Spann T. 1987. Disassembly of the nucleus in mitotic extracts: Membrane vesicularization, lamin disassembly, and chromosome condensation are independent processes. Cell 48: 219– 230. Ohba T, Schirmer EC, Nishimoto T, Gerace L. 2004. Energyand temperature-dependent transport of integral proteins to the inner nuclear membrane via the nuclear pore. J Cell Biol 167: 1051–1062. Ohsugi M, Adachi K, Horai R, Kakuta S, Sudo K, Kotaki H, Tokai-Nishizumi N, Sagara H, Iwakura Y, Yamamoto T. 2008. Kid-mediated chromosome compaction ensures proper nuclear envelope formation. Cell 132: 771–782. Ohtsubo M, Okazaki H, Nishimoto T. 1989. The RCC1 protein, a regulator for the onset of chromosome condensation locates in the nucleus and binds to DNA. J Cell Biol 109: 1389– 1397. Onischenko E, Stanton LH, Madrid AS, Kieselbach T, Weis K. 2009. Role of the Ndc1 interaction network in yeast nuclear pore complex assembly and maintenance. J Cell Biol 185: 475–491. Orjalo AV, Arnaoutov A, Shen Z, Boyarchuk Y, Zeitlin SG, Fontoura B, Briggs S, Dasso M, Forbes DJ. 2006. The Nup107–160 nucleoporin complex is required for correct bipolar spindle assembly. Mol Biol Cell 17: 3806– 3818. Osmani AH, Davies J, Liu HL, Nile A, Osmani SA. 2006. Systematic deletion and mitotic localization of the nuclear pore complex proteins of Aspergillus nidulans. Mol Biol Cell 17: 4946–4961. Peters R. 2009. Translocation through the nuclear pore: Kaps pave the way. Bioessays 31: 466– 477. Pichler A, Gast A, Seeler JS, Dejean A, Melchior F. 2002. The nucleoporin RanBP2 has SUMO1 E3 ligase activity. Cell 108: 109– 120. Portier N, Audhya A, Maddox PS, Green RA, Dammermann A, Desai A, Oegema K. 2007. A microtubule-independent role for centrosomes and aurora a in nuclear envelope breakdown. Dev Cell 12: 515– 529. Poteryaev D, Squirrell JM, Campbell JM, White JG, Spang A. 2005. Involvement of the actin cytoskeleton and

homotypic membrane fusion in ER dynamics in Caenorhabditis elegans. Mol Biol Cell 16: 2139–2153. Puhka M, Vihinen H, Joensuu M, Jokitalo E. 2007. Endoplasmic reticulum remains continuous and undergoes sheet-to-tubule transformation during cell division in mammalian cells. J Cell Biol 179: 895 –909. Pyrpasopoulou A, Meier J, Maison C, Simos G, Georgatos SD. 1996. The lamin B receptor (LBR) provides essential chromatin docking sites at the nuclear envelope. EMBO J 15: 7108–7119. Rabut G, Doye V, Ellenberg J. 2004. Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nat Cell Biol 6: 1114– 1121. Ramadan K, Bruderer R, Spiga FM, Popp O, Baur T, Gotta M, Meyer HH. 2007. Cdc48/p97 promotes reformation of the nucleus by extracting the kinase Aurora B from chromatin. Nature 450: 1258–1262. Rasala BA, Orjalo AV, Shen Z, Briggs S, Forbes DJ. 2006. ELYS is a dual nucleoporin/kinetochore protein required for nuclear pore assembly and proper cell division. Proc Natl Acad Sci 103: 17801– 17806. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243 –247. Ribeiro KC, Pereira-Neves A, Benchimol M. 2002. The mitotic spindle and associated membranes in the closed mitosis of trichomonads. Biol Cell 94: 157– 172. Roux KJ, Burke B. 2007. Nuclear envelope defects in muscular dystrophy. Biochim Biophys Acta 1772: 118– 127. Ryan KJ, McCaffery JM, Wente SR. 2003. The Ran GTPase cycle is required for yeast nuclear pore complex assembly. J Cell Biol 160: 1041–1053. Salina D, Bodoor K, Eckley DM, Schroer TA, Rattner JB, Burke B. 2002. Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell 108: 97–107. Salina D, Enarson P, Rattner JB, Burke B. 2003. Nup358 integrates nuclear envelope breakdown with kinetochore assembly. J Cell Biol 162: 991–1001. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Schirmer EC, Foisner R. 2007. Proteins that associate with lamins: Many faces, many functions. Exp Cell Res 313: 2167–2179. Schirmer EC, Gerace L. 2005. The nuclear membrane proteome: Extending the envelope. Trends Biochem Sci 30: 551– 558. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2003. Nuclear membrane proteins with potential disease links found by subtractive proteomics. Science 301: 1380–1382. Segura-Totten M, Kowalski AK, Craigie R, Wilson KL. 2002. Barrier-to-autointegration factor: Major roles in chromatin decondensation and nuclear assembly. J Cell Biol 158: 475–485. Sheehan MA, Mills AD, Sleeman AM, Laskey RA, Blow JJ. 1988. Steps in the assembly of replication-competent nuclei in a cell-free system from Xenopus eggs. J Cell Biol 106: 1 –12. Siniossoglou S. 2009. Lipins, Lipids and Nuclear Envelope Structure. Traffic 10: 1181– 1187.

M.W. Hetzer Starr DA, Han M. 2003. ANChors away: an actin based mechanism of nuclear positioning. J Cell Sci 116: 211–216. Stavru F, Hulsmann BB, Spang A, Hartmann E, Cordes VC, Gorlich D. 2006. NDC1: A crucial membrane-integral nucleoporin of metazoan nuclear pore complexes. J Cell Biol 173: 509–519. Stewart CL, Roux KJ, Burke B. 2007. Blurring the boundary: The nuclear envelope extends its reach. Science 318: 1408– 1412. Tahara K, Takagi M, Ohsugi M, Sone T, Nishiumi F, Maeshima K, Horiuchi Y, Tokai-Nishizumi N, Imamoto F, Yamamoto T, et al. 2008. Importin-b and the small guanosine triphosphatase Ran mediate chromosome loading of the human chromokinesin Kid. J Cell Biol 180: 493–506. Terasaki M, Campagnola P, Rolls MM, Stein PA, Ellenberg J, Hinkle B, Slepchenko B. 2001. A new model for nuclear envelope breakdown. Mol Biol Cell 12: 503–510. Terry LJ, Shows EB, Wente SR. 2007. Crossing the nuclear envelope: Hierarchical regulation of nucleocytoplasmic transport. Science 318: 1412–1416. Tran EJ, Wente SR. 2006. Dynamic nuclear pore complexes: Life on the edge. Cell 125: 1041–1053. Tsai MY, Wang S, Heidinger JM, Shumaker DK, Adam SA, Goldman RD, Zheng Y. 2006. A mitotic lamin B matrix induced by RanGTP required for spindle assembly. Science 311: 1887–1893. Tzur YB, Wilson KL, Gruenbaum Y. 2006. SUN-domain proteins: ’Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782– 788. Ulbert S, Platani M, Boue S, Mattaj IW. 2006. Direct membrane protein-DNA interactions required early in nuclear envelope assembly. J Cell Biol 173: 469– 476. Vergnolle MA, Taylor SS. 2007. Cenp-F links kinetochores to Ndel1/Nde1/Lis1/dynein microtubule motor complexes. Curr Biol 17: 1173–1179. Vlcek S, Foisner R. 2007. Lamins and lamin-associated proteins in aging and disease. Curr Opin Cell Biol 19: 298–304. Voeltz GK, Prinz WA. 2007. Sheets, ribbons and tubules— how organelles get their shape. Nat Rev Mol Cell Biol 8: 258–264. Voeltz GK, Prinz WA, Shibata Y, Rist JM, Rapoport TA. 2006. A class of membrane proteins shaping the tubular endoplasmic reticulum. Cell 124: 573– 586. Walther TC, Alves A, Pickersgill H, Loiodice I, Hetzer M, Galy V, Hulsmann BB, Kocher T, Wilm M, Allen T, et al.

2003a. The conserved Nup107– 160 complex is critical for nuclear pore complex assembly. Cell 113: 195– 206. Walther TC, Askjaer P, Gentzel M, Habermann A, Griffiths G, Wilm M, Mattaj IW, Hetzer M. 2003b. RanGTP mediates nuclear pore complex assembly. Nature 424: 689 –694. Watson ML. 1955. The nuclear envelope; its structure and relation to cytoplasmic membranes. J Biophys Biochem Cytol 1: 257– 270. Wilhelmsen K, Ketema M, Truong H, Sonnenberg A. 2006. KASH-domain proteins in nuclear migration, anchorage and other processes. J Cell Sci 119: 5021–5029. Wilhelmsen K, Litjens SH, Kuikman I, Tshimbalanga N, Janssen H, van den Bout I, Raymond K, Sonnenberg A. 2005. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J Cell Biol 171: 799– 810. Winey M, Yarar D, Giddings TH Jr, Mastronarde DN. 1997. Nuclear pore complex number and distribution throughout the Saccharomyces cerevisiae cell cycle by three-dimensional reconstruction from electron micrographs of nuclear envelopes. Mol Biol Cell 8: 2119–2132. Worman HJ, Bonne G. 2007. “Laminopathies”: Awide spectrum of human diseases. Exp Cell Res 313: 2121– 2133. Wu L, Osmani SA, Mirabito PM. 1998. A role for NIMA in the nuclear localization of cyclin B in Aspergillus nidulans. J Cell Biol 141: 1575– 1587. Yang L, Guan T, Gerace L. 1997. Integral membrane proteins of the nuclear envelope are dispersed throughout the endoplasmic reticulum during mitosis. J Cell Biol 137: 1199–1210. Ye Y, Meyer HH, Rapoport TA. 2003. Function of the p97Ufd1-Npl4 complex in retrotranslocation from the ER to the cytosol: Dual recognition of nonubiquitinated polypeptide segments and polyubiquitin chains. J Cell Biol 162: 71– 84. Zhang C, Clarke PR. 2000. Chromatin-independent nuclear envelope assembly induced by Ran GTPase in Xenopus egg extracts. Science 288: 1429–1432. Zheng Y, Tsai MY. 2006. The mitotic spindle matrix: A fibromembranous lamin connection. Cell Cycle 5: 2345–2347. Zuccolo M, Alves A, Galy V, Bolhy S, Formstecher E, Racine V, Sibarita JB, Fukagawa T, Shiekhattar R, Yen T, et al. 2007. The human Nup107–160 nuclear pore subcomplex contributes to proper kinetochore functions. EMBO J 26: 1853–1864.

Nuclear Lamins Thomas Dechat1, Stephen A. Adam1, Pekka Taimen2, Takeshi Shimi1, and Robert D. Goldman1 1

Department of Cell and Molecular Biology, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611

2

Department of Pathology, University of Turku, Kiinamyllynkatu 10 20520 Turku, Finland

Correspondence: [email protected]

The nuclear lamins are type V intermediate filament proteins that are critically important for the structural properties of the nucleus. In addition, they are involved in the regulation of numerous nuclear processes, including DNA replication, transcription and chromatin organization. The developmentally regulated expression of lamins suggests that they are involved in cellular differentiation. Their assembly dynamic properties throughout the cell cycle, particularly in mitosis, are influenced by posttranslational modifications. Lamins may regulate nuclear functions by direct interactions with chromatin and determining the spatial organization of chromosomes within the nuclear space. They may also regulate chromatin functions by interacting with factors that epigenetically modify the chromatin or directly regulate replication or transcription.

filamentous layer located between the inner nuclear membrane (INM) and peripheral heterochromatin was evident even in early electron microscopic studies of vertebrate cell nuclei (Fawcett 1966). This layer, later termed the nuclear lamina, is also found to be closely associated with nuclear pore complexes (NPCs) and contains three major structurally related polypeptides (Aaronson and Blobel 1975). These proteins are named nuclear lamins A, B, and C according to their molecular weights (Gerace and Blobel 1980). Further biochemical characterization and cDNA cloning of the nuclear lamins classifies them as type V intermediate filament proteins (Goldman et al. 1986; McKeon et al. 1986).

A

STRUCTURE AND BIOCHEMICAL PROPERTIES OF NUCLEAR LAMINS Lamin Isoforms and Expression Patterns

Lamins are present in all metazoans examined to date ranging from hydra to human, but are not found in unicellular organisms and plants (Cohen et al. 2001; Melcer et al. 2007). Extensive characterization in several model organisms including humans, mice, frogs, fruit flies and nematodes shows that their properties are shared across species (Melcer et al. 2007; Dechat et al. 2008b). Based on their sequence homologies, expression patterns, structural features, and biochemical and dynamic properties, lamins are subdivided into A- and B-types. All

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000547 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000547

T. Dechat et al.

metazoans express at least one B-type lamin. Typically, invertebrates have only a single lamin gene of the B-type, with some exceptions such as Drosophila, which expresses one B-type (lamin Dm0) and one A-type lamin (lamin C) encoded by two distinct genes. Most vertebrates have one A-type lamin and two B-type lamin genes except for Xenopus, which has three B-type genes (Melcer et al. 2007). The two major isoforms of vertebrate A-type lamins, lamins A and C, are derived from a single gene (LMNA) by alternative splicing (see Fig. 1) (Lin and Worman 1993). In some vertebrates, alternative splicing can also produce two less abundant isoforms, lamins AD10 and C2, from LMNA (Nakajima and Abe 1995; Machiels et al. 1996). Lamins B1 and B2 are the two major B-type lamins in most vertebrates. They are encoded by the LMNB1 and LMNB2 genes, respectively (Peter et al. 1989; Vorburger et al. 1989). The latter also encodes the minor isoform lamin B3 (Furukawa and Hotta 1993). Although at least one B-type lamin is expressed in all cells throughout development, the expression of A-type lamins is developmentally regulated (Benavente et al. 1985; Schatten et al. 1985; Lehner et al. 1987). During mouse development, lamins A and C are not expressed until days 10 –12 of mouse embryogenesis and then primarily in primordial muscle cells (Stewart and Burke 1987; Rober et al. 1989). Lamin A/C expression in other organs does not occur until after birth (Rober et al. 1989). Cells of hematopoietic lineage express only B-type lamins (Guilly et al. 1990; Rober et al. 1990). Similar patterns of expression of A and B-type lamins take place during the developmental α-helical rod L12

L1 Lamin A

Lamin C

1A

1A

1B

1B

progression of other vertebrates (Benavente et al. 1985; Lehner et al. 1987; Prather et al. 1989) and Drosophila (Frasch et al. 1988; Riemer et al. 1995). The regulated expression of A- and B-type lamins is also evident during differentiation of stem cells in culture. For example, undifferentiated human and mouse embryonic stem (ES) cells lack lamins A and C, but express lamins B1 and B2 (Constantinescu et al. 2006). The minor mammalian isoforms, lamins C2 and B3, are expressed exclusively in germ cells (Furukawa and Hotta 1993; Machiels et al. 1995; Nakajima and Abe 1995), whereas small amounts of lamin AD10 appear to be present in a variety of cell types (Machiels et al. 1995). The importance of the developmental regulation of lamin expression is evident from studies in LMNA knockout (LMNA 2/2 ) and LMNB1 mutant (LMNB D/D) mice. In mice null for the A-type lamins, no obvious embryonic defects can be detected, but these animals show severe postnatal growth retardation and muscular dystrophy (Sullivan et al. 1999). On the other hand, mice with an insertional mutation in LMNB1 develop defects in their lungs and bones during embryogenesis and die at birth, even though they continue to express lamin B2 (Vergnes et al. 2004). Structure and Assembly of the Nuclear Lamins

The nuclear lamins have the typical tripartite structure of intermediate filament (IF) proteins, consisting of a highly a-helical central rod domain flanked by a short globular amino-terminal “head” domain and a longer carboxy-terminal NLS

lg-fold

L2 2A

2A

2B 389

417 436

544

389

417 436

544 572

646

2B

Figure 1. Structure of nuclear lamins. Schematic drawing of mature lamin A and lamin C polypeptide chains.

The lamin structure consists of a short amino terminal head domain, a central a-helical rod domain (red), and the carboxy-terminal domain containing the NLS and the Ig-fold (blue; the nine b-strands of the Ig-fold motif are depicted). Modified, with permission, from (Dechat et al. 2008b).

Nuclear Lamins

“tail” domain (Parry et al. 1986). The central rod domain is composed of four subhelical regions comprised of heptad repeats and designated as coil 1A, 1B, 2A, and 2B. These individual “coils” are separated from each other by three short linker segments, L1, L12, and L2, of which L12 is the most flexible (Parry et al. 1986). Although the head domain appears to be unstructured, the tail domain contains a highly conserved structural motif similar to a type s immunoglobulin fold (Ig-fold) (Dhe-Paganon et al. 2002; Krimm et al. 2002). A nuclear localization signal (NLS), required for transport into the nucleus, is present in all lamins between the carboxy-terminal end of the central rod domain and the Ig-fold (Loewinger and McKeon 1988) (Fig. 1). In vitro, lamins self-assemble into higher order structures through a series of steps. The first step involves the formation of a coiled-coil dimer by the in register and in parallel association of two a-helical rod domains into a left handed superhelix (Heitlinger et al. 1991) (Fig. 2A). Next, the lamin dimers associate in a head-to-tail fashion (Heitlinger et al. 1991; Stuurman et al. 1996) and these polarized arrays interact in an antiparallel fashion to form apolar tetrameric protofilaments (Heitlinger et al. 1991) (Fig. 2B). The interaction of four lamin protofilaments leads to the formation of 10-nm filaments (Heitlinger et al. 1991; Stuurman et al. 1996; Ben-Harush et al. 2009) (Fig. 2C,D). Although the assembly of some invertebrate lamins appears to terminate at this stage of the assembly process, vertebrate lamins continue to assemble into well ordered paracrystalline arrays in vitro (Zackroff and Goldman 1979; Heitlinger et al. 1991; Stuurman et al. 1996; Ben-Harush et al. 2009) (Fig. 2E). The assembly of lamin proteins differs from their cytoplasmic counterparts in several ways. For example, lamin dimers associate by polar “head-to-tail” interactions to form protofilaments, which is very different from the halfstaggered, antiparallel side-by-side association of cytoplasmic IF dimers, which assemble into 10 nm filaments (Strelkov et al. 2004). This difference is at least in part explained by the crystal structure of the coiled-coil dimer of the lamin A coil 2B region (Strelkov et al. 2004). Although

Figure 2. Assembly of the nuclear lamins in vitro. Lamins self assemble to form dimers (A) which then join to form linear head-to-tail polymers ( protofilaments) (B). Bar ¼ 100 nm; electron micrograph of rotary shadowed chicken lamin B2. These protofilaments further assemble into “beaded” filaments or fibers (C ) which in turn associate laterally into thicker fibers (D), and eventually into paracrystalline arrays (E); C,D are negatively stained electron microscope preparations. Reprinted from Stuurman et al. (1998) with permission from Elsevier.

this structure appears to be similar to the overall structure of the homologous segment of the cytoplasmic IF protein vimentin, the distribution of charged residues varies causing significant changes in the patterns of intra- and interhelical salt bridges. Additionally, vertebrate lamins have six extra heptads in the central rod domain (McKeon et al. 1986), which may also help to explain assembly differences between cytoplasmic IF and lamins. The central rod domains of some invertebrates lamins contain fewer extra heptads and these lamins stop assembling into higher order structures at the 10-nm (IF) stage of assembly, similar to cytoplasmic IF proteins (Ben-Harush et al. 2009). Posttranslational Processing and Modifications of the Nuclear Lamins

Lamins A, B1, and B2 are expressed as prelamins that require extensive posttranslational modifications of their carboxy-terminal – CAAX box

T. Dechat et al.

to become mature lamins (Rusinol and Sinensky 2006; Davies et al. 2009). Modification of the – CAAX box takes place in a highly regulated temporal sequence starting with the farnesylation of the cysteine residue (Farnsworth et al. 1990; Lutz et al. 1992) by a farnesyltransferase (Zhang and Casey 1996). This modification initiates a sequence of processing steps (Beck et al. 1990) beginning with the removal of the – AAX by a CAAX prenyl protease. Two members of this zinc metalloproteinase family have been identified in humans: Rce1 (Ras-converting enzyme 1) and Zmpste24 (Zinc metalloprotease related to the STE24 homolog in yeast), also known as FACE1 (farnesylated-proteins converting enzyme)(Boyartchuk et al. 1997; Leung et al. 2001; Corrigan et al. 2005); followed by carboxymethylation of the carboxy-terminal

cysteine by isoprenylcysteine carboxyl methyltransferase (Icmt) (Winter-Vann and Casey 2005). In contrast to B-type lamins, which remain permanently farnesylated and carboxymethylated, an additional 15 amino acids are removed from the carboxyl terminus of farnesylated/carboxymethylated prelamin A by Zmpste24/FACE1 (Corrigan et al. 2005). This final processing step results in the production of mature lamin A lacking the carboxy-terminal farnesyl- and carboxymethyl-modifications (Fig. 3). This sequence of lamin processing steps is highly regulated and each step depends on the preceding modification (Kilic et al. 1997). Lamin C, which is 74 residues shorter than mature lamin A, does not possess a – CAAX box and therefore is not farnesylated or otherwise modified (see Fig. 1). The precise location

Pre-lamin CAAX-COOH Farnesyltransferase S

CAAX-COOH AAX endopeptidase S

Carboxylmethyltransferase

C-COOH

S

C-COCH3 Zmpste24/FACE1 S

C-COCH3 417 436

544 Mature lamin A

646

545 415 438 Mature lamin B1

583

S

C-COCH3 577 435 470 Mature lamin B2

617

Figure 3. Posttranslational processing of the carboxyl terminus of prelamins A, B1, and B2. Processing takes place in a series of steps: (1) addition of a farnesyl group to the cysteine residue of the –CAAX box of pre-lamin A, prelamin B1 and prelamin B2 by a farnesyltransferase; (2) removal of the last three residues (2AAX) by an AAX endopeptidase; (3) methylation of the terminal carboxylic acid group (2COOH) by a carboxyl methyltransferase; (4) removal of the carboxyl terminal 15 amino acids of lamin A with the farnesyl attached by the metalloprotease Zmpste24/FACE1. This last proteolysis step does not occur on B-type lamins and therefore they remain farnesylated. Modified, with permission, from Dechat et al. (2008b).

Nuclear Lamins

of the intracellular post-translational processing sites for the lamins remains largely unknown. Although the enzymes required for processing the carboxyl terminus of the lamins are present in both the inner nuclear membrane and endoplasmic reticulum membranes, the complete processing of prelamin A can occur when the protein is confined either to the cytoplasm or to the nucleus (Barrowman et al. 2008). Even though the prelamins can be processed in the cytoplasm, it is more likely that all lamin processing occurs in the nucleus (Lutz et al. 1992), because lamins are rapidly transported into the nucleus after translation in the cytoplasm (Lehner et al. 1986). In addition to farnesylation and carboxymethylation, lamins are also posttranslationally modified by phosphorylation (Ottaviano and Gerace 1985), sumoylation (Zhang and Sarge 2008), ADP-ribosylation (Adolph 1987), and possibly by glycosylation (Ferraro et al. 1989). Phosphorylation of serine and threonine residues proximal to the NLS by protein kinase C is known to play a role in the regulated import of lamins into the nucleus (Hennekes et al. 1993; Leukel and Jost 1995). At the onset of mitosis, the phosphorylation of lamins at specific sites by cyclin-dependent kinase (Cdk) 1 and protein kinase C (PKC) is required to drive disassembly of the lamina (Gerace and Blobel 1980; Dessev and Goldman 1988; Dessev et al. 1988; Dessev et al. 1989; Heald and McKeon 1990; Peter et al. 1990; Ward and Kirschner 1990; Dessev et al. 1991; Molloy and Little 1992; Goss et al. 1994; Collas 1999). Subsequently, dephosphorylation of the mitotic sites by protein phosphatase 1a is required for lamin/lamina assembly during the telophase/early G1 transition (Thompson et al. 1997). Lamin Structure and Dynamic Properties within the Lamina and Nucleoplasm

Very little is known about the formation, composition, and structure of lamin polymers in vivo. The structures of the lamins within the lamina that have been described to date range from a regular meshwork of 10 – 15 nm filaments observed in Xenopus oocyte germinal

vesicles and in sperm pronuclei assembled in vitro (Aebi et al. 1986; Zhang and Casey 1996; Goldberg et al. 2008a) (Fig. 4A), to a more irregular filamentous meshwork seen in mammalian cells (Belmont et al. 1993; Schermelleh et al. 2008; Shimi et al. 2008) (Fig. 4B,C). These meshworks have long been assumed to be stable structures based on the resistance of the nuclear lamins within the lamina to extraction in detergent containing high salt solutions, and the propensity of purified lamins to polymerize into higher order insoluble structures in vitro at relatively low critical concentrations (Aebi et al. 1986; Lourim and Lin 1989; Glass and Gerace 1990). In further support of this assumption, measurements of lamin mobility by fluorescence recovery after photobleaching (FRAP) reveal that GFP-tagged A- and B-type lamins do not appreciably exchange subunits within the lamina for up to several hours throughout most of interphase (Broers et al. 1999; Moir et al. 2000; Dahl et al. 2006). Recently it has become apparent that the Aand B-type lamins form separate filamentous networks in the lamina. These structures can be identified by both high resolution light microscopy (Schermelleh et al. 2008; Shimi et al. 2008) and whole mount electron microscopy in lamin B containing Xenopus oocyte nuclei ectopically expressing A-type lamins (Goldberg et al. 2008b). Even though the A-type and B-type lamin networks appear to be mainly separate structures (Shimi et al. 2008), there is evidence that the individual networks overlap and interact to varying degrees. Evidence for interaction between these two lamin meshworks is based on studies using fluorescence resonance energy transfer (FRET) in combination with time domain fluorescence lifetime imaging and high resolution confocal immunofluorescence (Moir et al. 2000; Delbarre et al. 2006; Shimi et al. 2008). Other evidence supporting the interactions between the A and B type lamins is derived from studies of nuclei in which either A or B type lamins are perturbed. Fibroblasts derived from LMNA 2/2 mouse embryos (MEFs) or a limb girdle muscular dystrophy patient with a homozygous nonsense mutation in LMNA

T. Dechat et al.

Figure 4. The nuclear lamins form a meshwork of fil-

aments within the lamina. (A) Spread nuclear envelope from Xenopus oocytes after detergent extraction and preparation for transmission electron microscopy by freeze-drying/unidirectional metal shadowing. The micrograph shows the nuclear lamina meshwork partially studded with nuclear pore complexes. (Inset) Higher-magnification view of a particularly wellpreserved area clearly shows the near-tetragonal lamina meshwork. Bars, 1 mm. Reprinted from Stuurman et al. (1998), with permission from Elsevier. (B) Structured illumination microsopy (SIM) reveals that there is an irregular meshwork of nuclear lamin B as revealed by immunofluorescence (green). This preparation is also stained with antibodies directed against nuclear pores and is stained with DAPI for DNA/ chromatin. Pores (red), DAPI (blue). From Schermelleh et al. (2008). Reprinted with permission from AAAS. (C) Confocal immunofluorescence localization of lamin A/C (green) and lamin B1 (red) in HeLa cells. Lamins A/C and B1 in a single nucleus are seen in an equatorial section (left panels) and the nuclear surface (right panels). The areas indicated by white squares in the top panels are enlarged fivefold in the lower panels. These images demonstrate that lamins form mainly separate networks with some overlapping regions. Bar, 5 mm. Adapted, with permission, from Shimi et al. (2008).

contain abnormally shaped nuclei containing blebs or lobules (Sullivan et al. 1999; Muchir et al. 2003). A significant number of these blebbed regions lack B-type lamins and NPCs. In addition, nuclei in LMNB D/D MEFs are also highly lobulated, showing a dramatic increase in the mesh size of the lamin A/C network (Vergnes et al. 2004). HeLa cells depleted of lamin B1 by shRNA silencing frequently develop extensively enlarged lamin A/C meshworks and nuclear envelope blebs which lack lamin B2 and NPCs (Shimi et al. 2008). Many of these nuclear phenotypes are also found in cells endogenously or ectopically expressing either point mutations or truncations of lamins A/C (Ostlund et al. 2001; Vaughan et al. 2001; Vigouroux et al. 2001; Bechert et al. 2003; Favreau et al. 2003; Muchir et al. 2003; Goldman et al. 2004). It is also noteworthy that lamin A is more mobile in the nuclear lamina in nuclei with decreased amounts of lamin B1 (Tang et al. 2008). The A- and B-type lamins have different disassembly and assembly properties during mitosis as revealed by biochemical fractionation, immunofluorescence, and live cell imaging analysis, further supporting the idea that the two types of lamins form separate networks in the lamina. As described earlier, depolymerization of the lamins is regulated by mitotic kinases and reassembly requires dephosphorylation by protein phosphatase 1a (Thompson et al. 1997; Ito et al. 2007). When the nuclear envelope is disassembled during late prophase, A-type lamins become dispersed throughout the cytoplasm in an apparently freely diffusible state whereas the majority of B-type lamins remain associated with the nuclear membranes, which appear to be mainly dispersed into the endoplasmic reticulum (Gerace and Blobel 1980; Stick et al. 1988). This association of B-type lamins with membranes during mitosis appears to be mainly attributable to their permanently farnesylated state (Rusinol and Sinensky 2006). Further support for differences in the properties of the A- and B-type lamins is derived from studies revealing their spatial and temporal order of assembly into the nuclear lamina at anaphase/telophase and in the early stages of

Nuclear Lamins

G1. For example, in mouse keratinocytes and hamster cells, lamin B1 accumulates along the entire periphery of the decondensing daughter chromosomes and assembles into a relatively stable polymer by mid- to late- telophase (Broers et al. 1999; Moir et al. 2000). In these cells, lamin A begins to accumulate within the nucleus mainly after the major components of the nuclear envelope including NPCs are assembled in daughter cells in late telophase. In HeLa cells, however, a small fraction of A-type lamins associates with chromosomes much earlier than in mouse cells, first assembling in the central or “core” region of chromosomes in close association with kinetochores (Dechat et al. 2004; Dechat et al. 2007; Haraguchi et al. 2008) (Fig. 5). The remainder of the A-type lamins is then transported into the nucleus after formation of an intact nuclear envelope. Later, other elements of the nuclear envelope begin to assemble in daughter cells (Dechat et al. 2004; Dechat et al. 2007; Haraguchi et al. 2008). Although the major fraction of the various lamin isoforms is associated with the nuclear lamina, these proteins are also present

throughout the nucleoplasm, especially in interphase (Goldman et al. 1992; Lutz et al. 1992). These nucleoplasmic lamins are likely to have different functions, for example in DNA replication and transcription (Dechat et al. 2008b), and a small fraction may represent assembly intermediates that are subsequently incorporated into the nuclear lamina (Goldman et al. 1992). Interestingly, B-type lamins in the nucleoplasm appear to be relatively static, similar to those in the lamina, whereas the nucleoplasmic A-type lamins are much more dynamic. These differences in the properties of nucleoplasmic A- and B-type lamins are supported experimentally by their biochemical extractability and their dynamic properties as determined in vivo by FRAP analyses and fluorescence correlation spectroscopy (FCS) (Broers et al. 1999; Moir et al. 2000; Shimi et al. 2008). For example, FCS measurements of GFP-tagged lamins A/C and lamins B1/B2 showed that nucleoplasmic lamins A/C are highly mobile, whereas lamin B1 and lamin B2 are mainly immobile (Shimi et al. 2008). This latter observation suggests that nucleoplasmic B-type lamins are either assembled into some type of

Figure 5. Association of lamin A with chromosomes during mitosis. HeLa cells expressing GFP-lamin A were

followed by time-lapse microscopy from the metaphase/anaphase transition (far left panels) into early G1 (far right panels). GFP-lamin A first associates with the core regions of chromosomes during telophase and spreads to cover the entire chromatin surface by early G1. DIC images of the same series are shown in the bottom row (Dechat et al. 2007).

T. Dechat et al.

structure or are tightly associated with other unknown immobile structural components. Interestingly, decreasing the amount of nucleoplasmic lamin B1 by shRNA silencing increases the rate of mobility for a large fraction of nucleoplasmic lamin A (Shimi et al. 2008). Taken together, these FCS results suggest that, as in the case of the lamina, the A- and B-type lamins form separate yet interacting structures within the nucleoplasm. Obviously, it will be of great interest in the future to determine the specific functions of these nucleoplasmic lamins.

FUNCTIONS OF THE LAMINS Regulation of Nuclear Shape and Mechanical Stability

It is becoming increasingly obvious that some of the functions of nuclear lamins are analogous to those of cytoskeletal intermediate filament proteins, which are known to be involved in the determination and maintenance of cell shape and mechanical properties (Goldman et al. 2008). In support of this, the nuclei of LMNA 2/2 MEFs display increased deformability and impaired viability under mechanical strain compared to the nuclei in control MEFs (Houben et al. 2007). Increased nuclear deformability is also observed in human ES cells lacking A-type lamins as compared to nuclei in differentiated cells expressing the A-type lamins (Pajerowski et al. 2007). In addition, cells either deficient in lamins or expressing mutant lamin proteins often contain misshapen nuclei (Ostlund et al. 2001; Vaughan et al. 2001; Vigouroux et al. 2001; Bechert et al. 2003; Favreau et al. 2003; Muchir et al. 2003; Goldman et al. 2004). Quantitative rheological measurements, particle tracking methods and differential interference contrast microscopy reveal that in vitro reconstituted lamin B1 networks are extremely porous, with elastic stiffness providing resistance to shear deformations (Panorchan et al. 2004a; Panorchan et al. 2004b). Interestingly, measurements made by atomic force microscopy show that germinal vesicles isolated from Xenopus oocytes, which normally contain only B-type lamins, display a significant

increase in stiffness on the ectopic expression of lamin A (Schape et al. 2009). Nuclear Lamins and the Regulation of Chromatin Positioning and Gene Expression

Lamins may regulate transcription by organizing chromatin into active and inactive domains. For example, electron microscopic and light microscopic observations show that there is a close association between peripherally localized heterochromatin and the nuclear lamina (Fawcett 1966; Paddy et al. 1990) (Fig. 6). This suggests that lamins may be involved in anchoring or organizing interphase chromosomes (Sullivan et al. 1999; Goldman et al. 2004; Nikolova et al. 2004; Galiova et al. 2008). Additional support for this idea comes from biochemical experiments showing that lamins A/C bind to mitotic chromosomes and to polynucleosomal particles in vitro (Burke 1990; Glass and Gerace 1990; Yuan et al. 1991). The interaction of lamins with chromatin could be mediated either by their direct binding to histones (Taniura et al. 1995; Goldberg et al. 1999; Mattout et al. 2007), or to specific DNA sequences in the matrix attachment/scaffold-associated regions (MARs/ SARs) (Luderus et al. 1992; Luderus et al. 1994; Baricheva et al. 1996; Zhao et al. 1996). The Role of Lamins in Mitosis

In addition to nuclear disassembly, several lines of evidence suggest a direct role for lamins in nuclear assembly following mitosis. The

Figure 6. The nuclear lamina is a fibrous network (fibrous lamina) located between peripheral heterochromatin and the inner nuclear membrane. A transmission electron micrograph of a thin section showing a portion of a smooth muscle nucleus in guinea pig epedidymis. Adapted from Fawcett (1966), copyright 1995, Wiley-Liss, Inc. Reprinted with permission of John Wiley & Sons, Inc. American Journal of Anatomy, Vol. 119, No. 1, pg. 140.

Nuclear Lamins

microinjection of lamin antibodies into mitotic cells causes arrest in a telophase-like state with the chromosomes remaining condensed (Benavente and Krohne 1986). In addition, decreased expression of lamin Dm0 in Drosophila inhibits nuclear membrane assembly and causes an enrichment of NPCs in cytoplasmic annulate lamellae (Lenz-Bohme et al. 1997). In Caenorhabditis elegans, the down-regulation of the single lamin leads to a loss of chromosomes, defects in chromosome separation into daughter cell nuclei, and abnormal condensation of chromatin (Liu et al. 2000). In vitro nuclear assembly in frog egg extracts can be inhibited by addition of the C-terminal tail fragment of lamin B3, in part because of the Ig-fold motif in the lamin tail, which can inhibit lamin polymerization in vitro (Shumaker et al. 2005). Lastly, the expression of the mutant lamin A which causes Hutchinson-Gilford Progeria Syndrome (HGPS) leads to defects in cell division including a delay in cytokinesis, delayed completion of nuclear reassembly at the end of mitosis and an increase in mitotic cells (Cao et al. 2007; Dechat et al. 2007). B-type lamins also play a role in the formation of the mitotic spindle. In cultured cells, a diffuse localization of B-type lamins can be observed associated with the mitotic spindle (Georgatos et al. 1997; Maison et al. 1997; Moir et al. 2000; Beaudouin et al. 2002; Tsai et al. 2006). In frog egg extracts, a mitotic spindle can be induced to form around added DNA or on beads coated with a variety of factors important for mitotic spindle formation (Tsai and Zheng 2005). In these lysates, lamin B3, the major lamin in frog eggs, is required to organize a “spindle matrix” structure. This matrix contains membrane vesicles and proteins known to regulate the assembly and dynamic properties of the microtubules comprising the mitotic spindle (Tsai et al. 2006). Depletion of lamin B3 from the egg extracts with antibodies or the addition of dominant negative fragments of lamin B3 known to disrupt lamin assembly, also disrupt the formation of the mitotic spindle. Furthermore, lamin B3 is associated with several spindle assembly factors within the spindle matrix such as

NuMA and Nudel and it appears to play an important role in regulating the activity of these proteins during mitosis (Tsai et al. 2006; Ma et al. 2009).

The Role of Lamins in DNA Replication and Repair

Several lines of evidence suggest that lamins play a role in DNA replication. In cultured cells, sites of DNA replication can be visualized as discrete early or late replication foci in the nucleoplasm. These foci can be detected by incorporation of bromodeoxyuridine into sites of replication (Moir et al. 1994) or by immunolocalization of replication-associated proteins such as proliferating cell nuclear antigen (PCNA) (Shumaker et al. 2008). It has been shown that lamin B1 colocalizes with these replication foci during late S phase in mouse 3T3 cells (Moir et al. 1994) and that lamins A/C are present at sites of early replication in normal human fibroblasts (Kennedy et al. 2000). Furthermore, DNA replication in Xenopus nuclei assembled in vitro is inhibited by depletion of lamin B3 or by the addition of a dominant-negative fragment of lamin B3, which drives the disassembly of the endogenous lamin network (Lopez-Soler et al. 2001; Shumaker et al. 2005; Shumaker et al. 2008). There is evidence that the lamins play a direct role in regulating replication by their binding to DNA replication factors such as PCNA. The lamin binding site for PCNA resides within the highly conserved Ig-fold motif located in the carboxyl terminus of both the A- and B-type lamins (Shumaker et al. 2008). In addition, there is a close association between lamin B3 and PCNA at the surface of sperm head chromatin during the earliest stages of nuclear assembly in Xenopus interphase egg extracts (Shumaker et al. 2008). Nuclear lamins have also been implicated in DNA repair, although the mechanistic basis of this remains unclear. For example, the expression of disease-causing mutant lamins impairs the formation of DNA repair foci (Liu et al. 2005; Manju et al. 2006). Furthermore, genetic instability because of defects in telomere

T. Dechat et al.

function and DNA repair has been implicated in progeria, the premature aging syndrome (Gonzalez-Suarez et al. 2009a; Gonzalez-Suarez et al. 2009b).

The Role of Lamins in Transcription

Several lines of experimental evidence support the possibility that lamins play a role in transcription. An amino-terminally deleted dominant negative lamin A, which disassembles lamin networks, specifically inhibits RNA pol II activity in hamster cells or in nuclei isolated from Xenopus embryos (Spann et al. 2002). In addition, over-expression of lamins A/C or silencing of lamin B1 leads to a significant inhibition of pol II transcription in HeLa cells (Kumaran and Spector 2008; Shimi et al. 2008; Tang et al. 2008). Lamins also associate with several transcription factors leading to the suggestion that they are involved in specific regulatory pathways (Heessen and Fornerod 2007; Andres and Gonzalez 2009). For example, the association of Oct-1 with lamin B1 at the nuclear envelope appears to be important for the oxidative stress response in MEFs, as lamin B1 deficiency leads to a dysregulation of Oct-1-dependent genes and to an increase in reactive oxygen species (Malhas et al. 2009). Lamins A/C also interact with the transcription factors c-Fos, MOK2, and sterol response element binding protein 1 (SREBP1) (Dreuillet et al. 2002; Lloyd et al. 2002; Ivorra et al. 2006; Dreuillet et al. 2008; Harper et al. 2009). In the case of c-Fos, its interaction with lamins A/C at the nuclear envelope appear to suppress AP-1 (activating protein 1) binding to DNA and transcriptional activity in an extracellular signal-regulated kinase (ERK) 1/2 activity-dependent fashion (Ivorra et al. 2006; Gonzalez et al. 2008). In addition to regulating transcription factor function by direct interaction, lamins are also associated with transcription factors indirectly via several lamin-binding proteins including emerin, LAP2b and pRb. Although the evidence supporting a role for the lamins in regulating transcription is convincing, it remains unclear

whether this regulation involves direct or indirect binding to transcription factors. Lamins as Regulators of the Positioning and Organization of Chromatin

The positioning of genes near or in contact with the nuclear lamina may be a mechanism for modulating gene expression. Inactive genes and gene-poor chromatin are frequently found in association with the lamina region, which suggests that the nuclear lamina may be a transcriptionally silent microdomain (Boyle et al. 2001; Cremer et al. 2001). Furthermore, lamins may play a role in establishing this microdomain by directly interacting with chromatin. The interactions of lamins with chromatin/ DNA at the nuclear lamina can be identified in more detail by DamID labeling (Pickersgill et al. 2006). In this technique, an expression vector encoding a potential chromatin/DNA binding protein fused to the Escherichia coli enzyme DNA adenine methyl transferase (Dam) is expressed in cells. DNA methylated by the lamin Dam fusion protein is then analyzed by sequencing. In Drosophila Kc cells, lamin Dm0 seems to mainly associate with transcriptionally inactive, mid-to-late replicating genome regions, lacking active histone marks and enriched in large intergenic regions (Pickersgill et al. 2006). In human lung fibroblasts lamin B1-associated domains (LADs) are present along chromosomes in a distinct pattern interspersed with “lamin B1 poor” regions (Guelen et al. 2008). These LADs are 1 Mb in size, are mostly heterochromatic. Genes present within LADs are 5–10-fold less active than genes outside LADs. The lamina appears to have gene or chromatin-specific effects on activation or repression of transcription. Evidence for the lamina constituting a repressive environment for transcription comes from studies localizing inactive gene loci to the nuclear periphery (Kosak et al. 2002; Zink et al. 2004; Williams et al. 2006). For example, the inactive Ig-heavy and Ig-k loci are preferentially positioned at the nuclear periphery in non-B-cell lineages, but are centrally located in B-cell nuclei where

Nuclear Lamins

they are actively engaged in transcription (Kosak et al. 2002). In addition, transcriptionally inactive testis-specific gene clusters are frequently associated with lamin Dm0 in the lamina of Drosophila S2 cells. Upon downregulation of lamin Dm0, these gene clusters relocate from the lamina to the nuclear interior and become transcriptionally active (Shevelyov et al. 2009). Artificial tethering of a reporter gene to the nuclear envelope using the membrane spanning domain of the inner nuclear membrane protein emerin results in the transcriptional inactivation of the gene in mouse cells (Reddy et al. 2008). However, the relocalization of chromosomes from the nuclear interior to the nuclear periphery using the inner membrane protein Lap2b inactivates some genes, but not others (Finlan et al. 2008). The idea that the nuclear lamina is not repressive to all transcription is further supported by the finding that targeting a genetic locus to the nuclear lamina by artificial tethering to lamin B1 does not interfere with its transcriptional activation (Kumaran and Spector 2008). In addition, lamin A can act as a transcriptional repressor in mammalian cells and in yeast when artificially targeted to specific promoters (Lee et al. 2009). This latter finding is intriguing because yeast do not contain lamin genes. These somewhat conflicting results demonstrate that more research is required to determine the role of lamins in transcription both in the region of the lamina and throughout the nucleoplasm. Lamins are Involved in the Epigenetic Regulation of Chromatin

The dramatic loss of peripheral heterochromatin in the nuclei of cells expressing one of the mutant forms of lamin A, progerin, that causes the premature aging disorder HGPS emphasizes the important role of lamins in the modification and/or the organization of chromatin (Shumaker et al. 2006) (Fig. 7). A partial loss of peripheral heterochromatin is also seen in LMNA 2/2 MEFs (Sullivan et al. 1999; Nikolova et al. 2004). Furthermore, down-regulation of lamin B1 in HeLa cells results in lamin A/C-rich and lamin B2 deficient nuclear blebs

or microdomains characterized by the absence of heterochromatin (Shimi et al. 2008). These changes in chromatin organization related to lamin expression are reflected in alterations in histone modifications including reductions in trimethylated H3K9, trimethylated H3K27 and an increase in trimethylation of H4K20 in HGPS cells (Scaffidi and Misteli 2006; Shumaker et al. 2006; Shimi et al. 2008) (Fig. 7). These studies demonstrate that the interaction of chromatin with lamins, whether direct or indirect, has strong effects on the epigenetic modification of histones, and that laminassociated microdomains in the nucleus might regulate modification of chromatin (Pegoraro et al. 2009). One potential link between lamins and chromatin modification may involve the tumor suppressor ING1. This protein, which interacts with histone acetyltransferases and histone deacetylases, appears to be stabilized and targeted to the nucleus by its interaction with A-type lamins (Han et al. 2008). HGPS cells have reduced ING1 levels and the expression of ING1 protein lacking its lamin interacting domain causes a loss of peripheral heterochromatin in normal cells. Lamins and Interphase Chromosome Organization

The organization of chromosome territories and domains is influenced by the expression of lamins. For example, in LMNB D/D MEFs and in cells derived from patients suffering from some types of laminopathies, gene-poor chromosome 18 is positioned away from its normal location at the nuclear periphery toward the nuclear interior (Malhas et al. 2007; Meaburn et al. 2007). Furthermore, in HeLa cells silenced for the expression of lamin B1, the lamin A/C rich and lamin B2-deficient nuclear envelope blebs that form are associated predominantly with gene-rich chromosome regions (Shimi et al. 2008) (Fig. 8). Together, these findings suggest that A-type lamins are preferentially associated with gene-rich chromatin regions and B-type lamins are preferentially associated with gene-poor regions of chromosomes. It must be emphasized that these studies are of a

T. Dechat et al. Figure 7. The association of the nuclear lamina with

chromatin in normal and progeria patients’ cells. (A) Thin section electron micrograph of a late passage blebbed nucleus in an HGPS patient’s skin fibroblast (left and center panels) and normal human foreskin fibroblasts (right panel). A high-magnification view of the nuclear envelope in a normal human foreskin fibroblast shows a normal array of heterochromatin adjacent to the nuclear envelope, making any lamina structure difficult to detect (right panel). A higher-magnification view of a HGPS cell showing a loss of peripheral heterochromatin and a prominent electron-dense lamina region associated with the inner nuclear envelope membrane (the nucleus is to the left in center and right panels) (Goldman et al. 2004). (B) Alterations of histone methylation patterns in HGPS fibroblasts. Normal and HGPS fibroblasts from female donors were double-labeled with antibodies against lamins A/C (red) and trimethylated Lys 9 in histone H3 (H3K9me3), Lys 27 in histone H3 (H3K27me3), or Lys 20 in histone H4 (H4K20me3) (all green). Note the decrease of H3K9me3 and H3K27me3 and the increase of H4K20me3 in the lobulated HGPS nuclei compared with normal nuclei. The decrease in H3K27me3 is best observed at the inactive X chromosome, which is normally enriched in this histone modification (see arrowhead in center left panel). Bars, 10 mm. Reprinted, with permission, from Dechat et al. (2008b).

preliminary nature and that much more work is required to establish the precise relationships between the different types of lamins and gene rich/gene poor chromosome domains within the interphase nucleus. Lamins may also play a role in the localization and function of centromeres and telomeres in the nucleus. Centromeres are typically arranged near the nuclear periphery in an interphase nucleus (Solovei et al. 2004). However, the lamin A/C-rich, lamin B2-deficient nuclear envelope blebs that form in lamin B1-silenced cells are devoid of centromeres, suggesting that B-type lamins are involved in anchoring these

heterochromatic structures at the nuclear lamina (Shimi et al. 2008). Telomeres are also influenced by the composition and structure of the lamina, because in LMNA 2/2 MEFs, the distribution, length and structure of telomeres is altered leading to increased genomic instability compared to control MEFs (Gonzalez-Suarez et al. 2009a). Human fibroblasts ectopically expressing HGPS lamin A mutant proteins show rapid telomere shortening and an accelerated replicative senescence phenotype (McClintock et al. 2006; Huang et al. 2008). Furthermore, the proliferative defects seen in normal dermal fibroblasts expressing progerin can be overcome

Nuclear Lamins

Figure 8. Localization of specific gene rich chromosomal regions in nuclear blebs in LB1-silenced HeLa cells. Chromosomes are detected by fluorescence in situ hybridization. DNA is counterstained with Hoechst 33258 (blue). Chromosome 6p (or 19), and chromosome 6q (or 18) are shown in green and red, respectively. Bars, 5 mm. Adapted, with permission, from Shimi et al. (2008).

by expression of the catalytic subunit of telomerase or the inactivation of p53 (Kudlow et al. 2008). Other support for the involvement of lamins in telomere function comes from the finding that the D4Z4 human subtelomeric repeat can localize an adjacent telomere to the nuclear periphery probably by interacting with A-type lamins and CTCF (CCCTC-binding factor) (Ottaviani et al. 2009a; Ottaviani et al. 2009b). The D4Z4 repeat acts as a CTCF and A-type lamin-dependent transcriptional insulator, suggesting that nuclear lamin architecture may organize specific regions of chromatin and influence gene expression (Ottaviani et al. 2009a). Strong support for the role of the lamins in interphase chromosome organization comes from a study of cells obtained from patients with a form of atypical HGPS (Taimen et al. 2009b). Unlike the most common form of HGPS, which results in only defective lamin A, these patients have a mutation resulting in the expression of a point mutation, E145K, in the 1B segment of the central rod domain of both lamins A and C (see Fig. 1). The cells from these patients have a flower-shaped nucleus with centrally clustered centromeres and abnormal peripherally displaced telomeres (Fig. 9). This abnormal configuration of chromosomes requires a round of cell division and persists throughout interphase. The multilobulated

Figure 9. Mislocalization of heterochromatin and centromeres in HGPS E145K cells. Control and E145K patient cells were stained with anti-lamin A (red), CREST antiserum (green), and Hoechst (white). Maximum projections of series of z-sections spanning the entire nucleus and side projections are shown. Centromeres are clustered in the central region of the nucleus in E145K cells, while they are either closely associated with the peripheral lamina region or elsewhere in the nucleoplasm in control cells. Inset shows an example of the close association of one centromere with the lamina in a single confocal section. In the right hand panels, side projections from merged images are shown. Centromeres are distributed throughout the nuclei in normal cells. In E145K cells centromeres are clustered in the middle of one region of the nucleus (Taimen et al. 2009a).

nuclei are therefore unlikely to correctly establish normal interphase chromosome territory arrangements. Both the abnormal nuclear shapes and chromosome configurations in E145K cells appear to be due to the aberrant assembly of the nuclear lamina by the mutant lamin A. In vitro analysis of E145K lamin A polymerization shows that this mutant lamin forms disorganized higher-order structures, a defect not seen in the more common form of progeria resulting from a truncation near the carboxyl terminus (Taimen et al. 2009b). Lamins in Cell Proliferation and Differentiation

Deficiencies in A-type lamins or the expression of mutant A-type lamins both lead to an array of proliferative defects because of cell cycle

T. Dechat et al.

acceleration, cell cycle arrest and/or premature senescence (Verstraeten et al. 2007; Dechat et al. 2008a). There is evidence that lamins A/ C are involved in the regulation of the pRb/ E2F pathway responsible for the G1/S cell cycle transition. One current model suggests that pRb in its suppressive hypophosphorylated state is in a complex with lamins A/C and LAP2a, and that the loss of lamins A/C leads to the degradation of pRb (Ozaki et al. 1994; Markiewicz et al. 2002; Johnson et al. 2004). In addition to pRb, cyclin D3, another critical regulator of G1 progression, interacts directly with lamins A/C (Mariappan et al. 2007). The findings that lamins are involved in the regulation of cell cycle progression and that they are differentially expressed during development, suggest that they also function in the regulation of cell differentiation. In support of this, during myoblast differentiation there is a decrease in the soluble nucleoplasmic lamin A/C pool, most likely in a pRb and cyclin D3dependent manner (Muralikrishna et al. 2001; Mariappan and Parnaik 2005). Furthermore, the down-regulation of lamins A/C or the expression of lamin A/C mutants associated with Autosomal Dominant Emery Dreifuss Muscular Dystrophy (AD-EDMD) lead to impaired myoblast differentiation possibly by disrupting the Rb-MyoD pathway (Bakay et al. 2006; Frock et al. 2006). As further evidence for the importance of the lamin A/pRb interaction, the expression of either an AD-EDMD lamin A mutant protein or the HGPS lamin A mutant protein, progerin, results in the inhibition of the phosphorylation of pRb (Markiewicz et al. 2005; Dechat et al. 2007). In the case of the AD-EDMD mutation, a further decrease in pRb levels along with the failure to hyperphosphorylate the available pRB inhibits myoblast differentiation (Favreau et al. 2004; Kandert et al. 2009). In adipocyte differentiation, the overexpression of either wild-type lamin A or a lamin A mutant associated with familial partial lipodystrophy impairs the potential of mouse fibroblasts to differentiate into mature adipocytes; and LMNA 2/2 mouse embryonic fibroblasts differentiate more readily into fat-containing cells compared to control cells

(Boguslavsky et al. 2006). There is also evidence that lamins A/C regulate adipocyte differentiation in a complex with the inner nuclear membrane protein emerin, by influencing the nucleocytoplasmic distribution of b-catenin (Tilgner et al. 2009). In addition, lamins A/C appear to be involved in osteoblast differentiation, as silencing their expression causes impaired osteoblastogenesis and accelerated osteoclastogenesis in human bone marrow stromal cells (Akter et al. 2009; Rauner et al. 2009). Lamin A may also be involved in adult stem cell differentiation as the expression of progerin causes defects mouse stem cell populations (Espada et al. 2008) in the differentiation potential of human mesenchymal stem cells, probably by affecting the Notch-signaling pathway (Espada et al. 2008; Scaffidi and Misteli 2008). Another aspect of the involvement of the lamins in differentiation is related to aging. In this regard, progerin can be detected in cells from healthy individuals and the protein appears to accumulate with increasing age (McClintock et al. 2006; Scaffidi and Misteli 2006; Cao et al. 2007; Rodriguez et al. 2009). Furthermore, dermal fibroblasts from older individuals (Scaffidi and Misteli 2006), and cells in aged C. elegans (Haithcock et al. 2005) display changes in nuclear shape similar to those found in premature aging. The Lamins are Connected to the Cytoskeleton

There is evidence that the nuclear lamins are connected to the cytoskeleton through a complex of proteins called the LINC complex (linker of nucleoskeleton and cytoskeleton) (Crisp et al. 2006; Ketema et al. 2007; StewartHutchinson et al. 2008; Burke and Roux 2009). In this complex the integral proteins of the inner nuclear membrane, Sun1 and Sun2, interact with the outer nuclear membrane proteins, nesprin-1, nesprin-2, and nesprin-3a,in the luminal space between the inner and outer nuclear membranes. Although the Sun proteins bind to the lamins (Crisp et al. 2006), the nesprins are associated with both the microfilament and intermediate filament cytoskeletons

Nuclear Lamins

via direct actin binding and plectin (Tzur et al. 2006). Another lamin binding protein of the inner nuclear membrane, emerin, binds directly to nesprin isoforms, providing additional linkages between the lamins and the cytoskeleton (Wheeler et al. 2007; Zhang et al. 2007). These interactions between the nuclear lamina and the cytoskeleton may have important functional significance. For example, nuclei in fibroblasts deficient in lamin B1 rotate at a higher frequency than nuclei in control cells (Ji et al. 2007). LMNA 2/2 MEFs, on the other hand, are reported to show a decrease in mechanical stiffness, defects in mechanotransduction resulting in impaired strain-induced signaling, defects in cell polarization and cell migration, and a disturbed organization of microfilaments, vimentin IF networks and microtubules (Houben et al. 2007; Lee et al. 2007; Dahl et al. 2008; Houben et al. 2009). CONCLUSIONS AND FUTURE PERSPECTIVES

Over the past decade it has become clear that the nuclear lamins are one of the key players in determining nuclear architecture and function. There is growing evidence that these Type V IF proteins provide a filamentous scaffold built from interconnecting A- and B-type lamin networks that pervade the entire nucleus. This scaffold not only determines the shape and mechanical properties of the nucleus, but also serves as a docking site for chromatin and for numerous proteins involved in chromatin organization and various nuclear functions. Besides the extensive lamin structures located within the lamina, smaller and more dynamic lamin polymers appear to be components of large protein complexes known to be involved in a wide range of nuclear housekeeping functions such as DNA replication, DNA repair and RNA PolII transcription. We have learned much about the nuclear lamins from the analysis of the changes in nuclear form and function that take place in cells from patients suffering from the remarkably large number of diseases attributable to hundreds of mutations in the LMNA gene.

However, the precise roles that normal lamins play in nuclear functions remain largely unknown. To gain more insights into their specific functions we need a better understanding of their normal structure at the highest levels of resolution. To date only a few subdomains of the lamins have been shown to be amenable to analysis by X-ray diffraction, because of the well known difficulties inherent in crystallizing IF proteins. Other challenges for future studies include determining the steps in lamin polymerization and depolymerization in vivo and determining how complex networks of the Aand B-type lamins are assembled and interact with each other both at the nuclear periphery and within the nucleoplasm throughout the cell cycle. Finally, it will be of great interest to determine the different roles of the A- and B-type lamins in determining the overall architecture of the nucleus and the mechanisms involved in their linkages to other nuclear structures, such as the nuclear membrane and pore complexes.

ACKNOWLEDGMENTS

Work in the Goldman lab on nuclear lamins is funded by the National Institute on Aging and the National Cancer Institute. PT received a fellowship from Sigrid Juselius Foundation, Orion-Farmos Research Foundation, Cancer Society of Southwestern Finland, and Finnish Cultural Foundation.

REFERENCES Aaronson RP, Blobel G. 1975. Isolation of nuclear pore complexes in association with a lamina. Proc Natl Acad Sci 72: 1007–1011. Adolph KW. 1987. ADPribosylation of nuclear proteins labeled with [3H]adenosine: changes during the HeLa cycle. Biochim Biophys Acta 909: 222– 230. Aebi U, Cohn J, Buhle L, Gerace L. 1986. The nuclear lamina is a meshwork of intermediate-type filaments. Nature 323: 560–564. Akter R, Rivas D, Geneau G, Drissi H, Duque G. 2009. Effect of lamin A/C knockdown on osteoblast differentiation and function. J Bone Miner Res 24: 283 –293. Andres V, Gonzalez JM. 2009. Role of A-type lamins in signaling, transcription, and chromatin organization. J Cell Biol 187: 945 –957.

T. Dechat et al. Bakay M, Wang Z, Melcon G, Schiltz L, Xuan J, Zhao P, Sartorelli V, Seo J, Pegoraro E, Angelini C, et al. 2006. Nuclear envelope dystrophies show a transcriptional fingerprint suggesting disruption of Rb-MyoD pathways in muscle regeneration. Brain 129: 996– 1013. Baricheva EA, Berrios M, Bogachev SS, Borisevich IV, Lapik ER, Sharakhov IV, Stuurman N, Fisher PA. 1996. DNA from Drosophila melanogaster b-heterochromatin binds specifically to nuclear lamins in vitro and the nuclear envelope in situ. Gene 171: 171– 176. Barrowman J, Hamblet C, George CM, Michaelis S. 2008. Analysis of prelamin A biogenesis reveals the nucleus to be a CaaX processing compartment. Mol Biol Cell 19: 5398– 5408. Beaudouin J, Gerlich D, Daigle N, Eils R, Ellenberg J. 2002. Nuclear envelope breakdown proceeds by microtubule-induced tearing of the lamina. Cell 108: 83–96. Bechert K, Lagos-Quintana M, Harborth J, Weber K, Osborn M. 2003. Effects of expressing lamin A mutant protein causing Emery-Dreifuss muscular dystrophy and familial partial lipodystrophy in HeLa cells. Exp Cell Res 286: 75–86. Beck LA, Hosick TJ, Sinensky M. 1990. Isoprenylation is required for the processing of the lamin A precursor. J Cell Biol 110: 1489–1499. Belmont AS, Zhai Y, Thilenius A. 1993. Lamin B distribution and association with peripheral chromatin revealed by optical sectioning and electron microscopy tomography. J Cell Biol 123: 1671– 1685. Ben-Harush K, Wiesel N, Frenkiel-Krispin D, Moeller D, Soreq E, Aebi U, Herrmann H, Gruenbaum Y, Medalia O. 2009. The supramolecular organization of the C elegans nuclear lamin filament. J Mol Biol 386: 1392– 1402. Benavente R, Krohne G. 1986. Involvement of nuclear lamins in postmitotic reorganization of chromatin as demonstrated by microinjection of lamin antibodies. J Cell Biol 103: 1847– 1854. Benavente R, Krohne G, Franke WW. 1985. Cell typespecific expression of nuclear lamina proteins during development of Xenopus laevis. Cell 41: 177– 190. Boguslavsky RL, Stewart CL, Worman HJ. 2006. Nuclear lamin A inhibits adipocyte differentiation: implications for Dunnigan-type familial partial lipodystrophy. Hum Mol Genet 15: 653– 663. Boyartchuk VL, Ashby MN, Rine J. 1997. Modulation of Ras and a-factor function by carboxyl-terminal proteolysis. Science 275: 1796–1800. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211– 219. Broers JL, Machiels BM, van Eys GJ, Kuijpers HJ, Manders EM, van Driel R, Ramaekers FC. 1999. Dynamics of the nuclear lamina as monitored by GFP-tagged A-type lamins. J Cell Sci 112: 3463–3475. Burke B. 1990. On the cell-free association of lamins A and C with metaphase chromosomes. Exp Cell Res 186: 169–176. Burke B, Roux KJ. 2009. Nuclei take a position: Managing nuclear location. Dev Cell 17: 587–597.

Cao K, Capell BC, Erdos MR, Djabali K, Collins FS. 2007. A lamin A protein isoform overexpressed in HutchinsonGilford progeria syndrome interferes with mitosis in progeria and normal cells. Proc Natl Acad Sci 104: 4949–4954. Cohen M, Lee KK, Wilson KL, Gruenbaum Y. 2001. Transcriptional repression, apoptosis, human disease and the functional evolution of the nuclear lamina. Trends Biochem Sci 26: 41–47. Collas P. 1999. Sequential PKC- and Cdc2-mediated phosphorylation events elicit zebrafish nuclear envelope disassembly. J Cell Sci 112: 977–987. Constantinescu D, Gray HL, Sammak PJ, Schatten GP, Csoka AB. 2006. Lamin A/C expression is a marker of mouse and human embryonic stem cell differentiation. Stem Cells 24: 177–185. Corrigan DP, Kuszczak D, Rusinol AE, Thewke DP, Hrycyna CA, Michaelis S, Sinensky MS. 2005. Prelamin A endoproteolytic processing in vitro by recombinant Zmpste24. Biochem J 387: 129– 138. Cremer M, von Hase J, Volm T, Brero A, Kreth G, Walter J, Fischer C, Solovei I, Cremer C, Cremer T. 2001. Nonrandom radial higher-order chromatin arrangements in nuclei of diploid human cells. Chromosome Res 9: 541 –567. Crisp M, Liu Q, Roux K, Rattner JB, Shanahan C, Burke B, Stahl PD, Hodzic D. 2006. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J Cell Biol 172: 41– 53. Dahl KN, Ribeiro AJ, Lammerding J. 2008. Nuclear shape, mechanics, and mechanotransduction. Circ Res 102: 1307–1318. Dahl KN, Scaffidi P, Islam MF, Yodh AG, Wilson KL, Misteli T. 2006. Distinct structural and mechanical properties of the nuclear lamina in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 103: 10271– 10276. Davies BS, Fong LG, Yang SH, Coffinier C, Young SG. 2009. The posttranslational processing of prelamin A and disease. Annu Rev Genomics Hum Genet 10: 153– 174. Dechat T, Adam SA, Goldman RD. 2008a. Nuclear lamins and chromatin: When structure meets function. Adv Enzyme Regul 49: 157 –166. Dechat T, Gajewski A, Korbei B, Gerlich D, Daigle N, Haraguchi T, Furukawa K, Ellenberg J, Foisner R. 2004. LAP2a and BAF transiently localize to telomeres and specific regions on chromatin during nuclear assembly. J Cell Sci 117: 6117– 6128. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008b. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832– 853. Dechat T, Shimi T, Adam SA, Rusinol AE, Andres DA, Spielmann HP, Sinensky MS, Goldman RD. 2007. Alterations in mitosis and cell cycle progression caused by a mutant lamin A known to accelerate human aging. Proc Natl Acad Sci 104: 4955– 4960. Delbarre E, Tramier M, Coppey-Moisan M, Gaillard C, Courvalin JC, Buendia B. 2006. The truncated prelamin A in Hutchinson-Gilford progeria syndrome alters segregation of A-type and B-type lamin homopolymers. Hum Mol Genet 15: 1113– 1122.

Nuclear Lamins Dessev G, Goldman R. 1988. Meiotic breakdown of nuclear envelope in oocytes of Spisula solidissima involves phosphorylation and release of nuclear lamin. Dev Biol 130: 543–550. Dessev G, Iovcheva-Dessev C, Bischoff JR, Beach D, Goldman R. 1991. A complex containing p34cdc2 and cyclin B phosphorylates the nuclear lamin and disassembles nuclei of clam oocytes in vitro. J Cell Biol 112: 523 –533. Dessev G, Iovcheva C, Tasheva B, Goldman R. 1988. Protein kinase activity associated with the nuclear lamina. Proc Natl Acad Sci 85: 2994– 2998. Dessev G, Palazzo R, Rebhun L, Goldman R. 1989. Disassembly of the nuclear envelope of spisula oocytes in a cellfree system. Dev Biol 131: 496– 504. Dhe-Paganon S, Werner ED, Chi YI, Shoelson SE. 2002. Structure of the globular tail of nuclear lamin. J Biol Chem 277: 17381– 17384. Dreuillet C, Harper M, Tillit J, Kress M, Ernoult-Lange M. 2008. Mislocalization of human transcription factor MOK2 in the presence of pathogenic mutations of lamin A/C. Biol Cell 100: 51– 61. Dreuillet C, Tillit J, Kress M, Ernoult-Lange M. 2002. In vivo and in vitro interaction between human transcription factor MOK2 and nuclear lamin A/C. Nucleic Acids Res 30: 4634–4642. Espada J, Varela I, Flores I, Ugalde AP, Cadinanos J, Pendas AM, Stewart CL, Tryggvason K, Blasco MA, Freije JM, et al. 2008. Nuclear envelope defects cause stem cell dysfunction in premature-aging mice. J Cell Biol 181: 27– 35. Farnsworth CC, Gelb MH, Glomset JA. 1990. Identification of geranylgeranyl-modified proteins in HeLa cells. Science 247: 320– 322. Favreau C, Dubosclard E, Ostlund C, Vigouroux C, Capeau J, Wehnert M, Higuet D, Worman HJ, Courvalin JC, Buendia B. 2003. Expression of lamin A mutated in the carboxyl-terminal tail generates an aberrant nuclear phenotype similar to that observed in cells from patients with Dunnigan-type partial lipodystrophy and EmeryDreifuss muscular dystrophy. Exp Cell Res 282: 14– 23. Favreau C, Higuet D, Courvalin JC, Buendia B. 2004. Expression of a mutant lamin A that causes EmeryDreifuss muscular dystrophy inhibits in vitro differentiation of C2C12 myoblasts. Mol Cell Biol 24: 1481–1492. Fawcett DW. 1966. On the occurrence of a fibrous lamina on the inner aspect of the nuclear envelope in certain cells of vertebrates. Am J Anat 119: 129 –145. Ferraro A, Eufemi M, Cervoni L, Marinetti R, Turano C. 1989. Glycosylated forms of nuclear lamins. FEBS Lett 257: 241– 246. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Frasch M, Paddy M, Saumweber H. 1988. Developmental and mitotic behaviour of two novel groups of nuclear envelope antigens of Drosophila melanogaster. J Cell Sci 90: 247 –263. Frock RL, Kudlow BA, Evans AM, Jameson SA, Hauschka SD, Kennedy BK. 2006. Lamin A/C and emerin are critical for skeletal muscle satellite cell differentiation. Genes Dev 20: 486 –500.

Furukawa K, Hotta Y. 1993. cDNA cloning of a germ cell specific lamin B3 from mouse spermatocytes and analysis of its function by ectopic expression in somatic cells. Embo J 12: 97–106. Galiova G, Bartova E, Raska I, Krejci J, Kozubek S. 2008. Chromatin changes induced by lamin A/C deficiency and the histone deacetylase inhibitor trichostatin A. Eur J Cell Biol 87: 291– 303. Georgatos SD, Pyrpasopoulou A, Theodoropoulos PA. 1997. Nuclear envelope breakdown in mammalian cells involves stepwise lamina disassembly and microtubuledrive deformation of the nuclear membrane. J Cell Sci 110: 2129– 2140. Gerace L, Blobel G. 1980. The nuclear envelope lamina is reversibly depolymerized during mitosis. Cell 19: 277 –287. Glass JR, Gerace L. 1990. Lamins A and C bind and assemble at the surface of mitotic chromosomes. J Cell Biol 111: 1047–1057. Goldberg M, Harel A, Brandeis M, Rechsteiner T, Richmond TJ, Weiss AM, Gruenbaum Y. 1999. The tail domain of lamin Dm0 binds histones H2A and H2B. Proc Natl Acad Sci 96: 2852–2857. Goldberg MW, Fiserova J, Huttenlauch I, Stick R. 2008a. A new model for nuclear lamina organization. Biochem Soc Trans 36: 1339– 1343. Goldberg MW, Huttenlauch I, Hutchison CJ, Stick R. 2008b. Filaments made from A- and B-type lamins differ in structure and organization. J Cell Sci 121: 215 –225. Goldman RD, Grin B, Mendez MG, Kuczmarski ER. 2008. Intermediate filaments: versatile building blocks of cell structure. Curr Opin Cell Biol 20: 28–34. Goldman AE, Maul G, Steinert PM, Yang HY, Goldman RD. 1986. Keratin-like proteins that coisolate with intermediate filaments of BHK-21 cells are nuclear lamins. Proc Natl Acad Sci 83: 3839– 3843. Goldman AE, Moir RD, Montag-Lowy M, Stewart M, Goldman RD. 1992. Pathway of incorporation of microinjected lamin A into the nuclear envelope. J Cell Biol 119: 725–735. Goldman RD, Shumaker DK, Erdos MR, Eriksson M, Goldman AE, Gordon LB, Gruenbaum Y, Khuon S, Mendez M, Varga R, et al. 2004. Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci 101: 8963– 8968. Gonzalez JM, Navarro-Puche A, Casar B, Crespo P, Andres V. 2008. Fast regulation of AP-1 activity through interaction of lamin A/C, ERK1/2, and c-Fos at the nuclear envelope. J Cell Biol 183: 653–666. Gonzalez-Suarez I, Redwood AB, Gonzalo S. 2009a. Loss of A-type lamins and genomic instability. Cell Cycle 8: 3860–3865. Gonzalez-Suarez I, Redwood AB, Perkins SM, Vermolen B, Lichtensztejin D, Grotsky DA, Morgado-Palacin L, Gapud EJ, Sleckman BP, Sullivan T, et al. 2009b. Novel roles for A-type lamins in telomere biology and the DNA damage response pathway. Embo J 28: 2414–2427. Goss VL, Hocevar BA, Thompson LJ, Stratton CA, Burns DJ, Fields AP. 1994. Identification of nuclear b II protein

T. Dechat et al. kinase C as a mitotic lamin kinase. J Biol Chem 269: 19074–19080. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948–951. Guilly MN, Kolb JP, Gosti F, Godeau F, Courvalin JC. 1990. Lamins A and C are not expressed at early stages of human lymphocyte differentiation. Exp Cell Res 189: 145–147. Haithcock E, Dayani Y, Neufeld E, Zahand AJ, Feinstein N, Mattout A, Gruenbaum Y, Liu J. 2005. Age-related changes of nuclear architecture in Caenorhabditis elegans. Proc Natl Acad Sci USA 102: 16690–16695. Han X, Feng X, Rattner JB, Smith H, Bose P, Suzuki K, Soliman MA, Scott MS, Burke BE, Riabowol K. 2008. Tethering by lamin A stabilizes and targets the ING1 tumour suppressor. Nat Cell Biol 10: 1333–1340. Haraguchi T, Kojidani T, Koujin T, Shimi T, Osakada H, Mori C, Yamamoto A, Hiraoka Y. 2008. Live cell imaging and electron microscopy reveal dynamic processes of BAF-directed nuclear envelope assembly. J Cell Sci 121: 2540– 2554. Harper M, Tillit J, Kress M, Ernoult-Lange M. 2009. Phosphorylation-dependent binding of human transcription factor MOK2 to lamin A/C. Febs J 276: 3137– 3147. Heald R, McKeon F. 1990. Mutations of phosphorylation sites in lamin A that prevent nuclear lamina disassembly in mitosis. Cell 61: 579– 589. Heessen S, Fornerod M. 2007. The inner nuclear envelope as a transcription factor resting place. EMBO Rep 8: 914–919. Heitlinger E, Peter M, Haner M, Lustig A, Aebi U, Nigg EA. 1991. Expression of chicken lamin B2 in Escherichia coli: Characterization of its structure, assembly, and molecular interactions. J Cell Biol 113: 485 –495. Hennekes H, Peter M, Weber K, Nigg EA. 1993. Phosphorylation on protein kinase C sites inhibits nuclear import of lamin B2. J Cell Biol 120: 1293– 1304. Houben F, Ramaekers FC, Snoeckx LH, Broers JL. 2007. Role of nuclear lamina-cytoskeleton interactions in the maintenance of cellular strength. Biochim Biophys Acta 1773: 675–686. Houben F, Willems CH, Declercq IL, Hochstenbach K, Kamps MA, Snoeckx LH, Ramaekers FC, Broers JL. 2009. Disturbed nuclear orientation and cellular migration in A-type lamin deficient cells. Biochim Biophys Acta 1793: 312 –324. Huang S, Risques RA, Martin GM, Rabinovitch PS, Oshima J. 2008. Accelerated telomere shortening and replicative senescence in human fibroblasts overexpressing mutant and wild-type lamin A. Exp Cell Res 314: 82–91. Ito H, Koyama Y, Takano M, Ishii K, Maeno M, Furukawa K, Horigome T. 2007. Nuclear envelope precursor vesicle targeting to chromatin is stimulated by protein phosphatase 1 in Xenopus egg extracts. Exp Cell Res 313: 1897– 1910. Ivorra C, Kubicek M, Gonzalez JM, Sanz-Gonzalez SM, Alvarez-Barrientos A, O’Connor JE, Burke B, Andres V.

2006. A mechanism of AP-1 suppression through interaction of c-Fos with lamin A/C. Genes Dev 20: 307–320. Ji JY, Lee RT, Vergnes L, Fong LG, Stewart CL, Reue K, Young SG, Zhang Q, Shanahan CM, Lammerding J. 2007. Cell nuclei spin in the absence of lamin b1. J Biol Chem 282: 20015– 20026. Johnson BR, Nitta RT, Frock RL, Mounkes L, Barbie DA, Stewart CL, Harlow E, Kennedy BK. 2004. A-type lamins regulate retinoblastoma protein function by promoting subnuclear localization and preventing proteasomal degradation. Proc Natl Acad Sci 101: 9677–9682. Kandert S, Wehnert M, Muller CR, Buendia B, Dabauvalle MC. 2009. Impaired nuclear functions lead to increased senescence and inefficient differentiation in human myoblasts with a dominant p.R545C mutation in the LMNA gene. Eur J Cell Biol 88: 593– 608. Kennedy BK, Barbie DA, Classon M, Dyson N, Harlow E. 2000. Nuclear organization of DNA replication in primary mammalian cells. Genes Dev 14: 2855–2868. Ketema M, Wilhelmsen K, Kuikman I, Janssen H, Hodzic D, Sonnenberg A. 2007. Requirements for the localization of nesprin-3 at the nuclear envelope and its interaction with plectin. J Cell Sci 120: 3384–3394. Kilic F, Dalton MB, Burrell SK, Mayer JP, Patterson SD, Sinensky M. 1997. In vitro assay and characterization of the farnesylation-dependent prelamin A endoprotease. J Biol Chem 272: 5298–5304. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158 –162. Krimm I, Ostlund C, Gilquin B, Couprie J, Hossenlopp P, Mornon JP, Bonne G, Courvalin JC, Worman HJ, ZinnJustin S. 2002. The Ig-like structure of the C-terminal domain of lamin A/C, mutated in muscular dystrophies, cardiomyopathy, and partial lipodystrophy. Structure 10: 811 –823. Kudlow BA, Stanfel MN, Burtner CR, Johnston ED, Kennedy BK. 2008. Suppression of proliferative defects associated with processing-defective lamin A mutants by hTERT or inactivation of p53. Mol Biol Cell 19: 5238–5248. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Lee JS, Hale CM, Panorchan P, Khatau SB, George JP, Tseng Y, Stewart CL, Hodzic D, Wirtz D. 2007. Nuclear lamin A/C deficiency induces defects in cell mechanics, polarization, and migration. Biophys J 93: 2542–2552. Lee DC, Welton KL, Smith ED, Kennedy BK. 2009. A-type nuclear lamins act as transcriptional repressors when targeted to promoters. Exp Cell Res 315: 996– 1007. Lehner CF, Furstenberger G, Eppenberger HM, Nigg EA. 1986. Biogenesis of the nuclear lamina: in vivo synthesis and processing of nuclear protein precursors. Proc Natl Acad Sci 83: 2096–2099. Lehner CF, Stick R, Eppenberger HM, Nigg EA. 1987. Differential expression of nuclear lamin proteins during chicken development. J Cell Biol 105: 577– 587. Lenz-Bohme B, Wismar J, Fuchs S, Reifegerste R, Buchner E, Betz H, Schmitt B. 1997. Insertional mutation of the

Nuclear Lamins Drosophila nuclear lamin Dm0 gene results in defective nuclear envelopes, clustering of nuclear pore complexes, and accumulation of annulate lamellae. J Cell Biol 137: 1001– 1016. Leukel M, Jost E. 1995. Two conserved serines in the nuclear localization signal flanking region are involved in the nuclear targeting of human lamin A. Eur J Cell Biol 68: 133–142. Leung GK, Schmidt WK, Bergo MO, Gavino B, Wong DH, Tam A, Ashby MN, Michaelis S, Young SG. 2001. Biochemical studies of Zmpste24-deficient mice. J Biol Chem 276: 29051– 29058. Lin F, Worman HJ. 1993. Structural organization of the human gene encoding nuclear lamin A and nuclear lamin C. J Biol Chem 268: 16321–16326. Liu J, Rolef Ben-Shahar T, Riemer D, Treinin M, Spann P, Weber K, Fire A, Gruenbaum Y. 2000. Essential roles for Caenorhabditis elegans lamin gene in nuclear organization, cell cycle progression, and spatial organization of nuclear pore complexes. Mol Biol Cell 11: 3937–3947. Liu B, Wang J, Chan KM, Tjia WM, Deng W, Guan X, Huang JD, Li KM, Chau PY, Chen DJ, et al. 2005. Genomic instability in laminopathy-based premature aging. Nat Med 11: 780 –785. Lloyd DJ, Trembath RC, Shackleton S. 2002. A novel interaction between lamin A and SREBP1: Implications for partial lipodystrophy and other laminopathies. Hum Mol Genet 11: 769 –777. Loewinger L, McKeon F. 1988. Mutations in the nuclear lamin proteins resulting in their aberrant assembly in the cytoplasm. Embo J 7: 2301– 2309. Lopez-Soler RI, Moir RD, Spann TP, Stick R, Goldman RD. 2001. A role for nuclear lamins in nuclear envelope assembly. J Cell Biol 154: 61–70. Lourim D, Lin JJ. 1989. Expression of nuclear lamin A and muscle-specific proteins in differentiating muscle cells in ovo and in vitro. J Cell Biol 109: 495– 504. Luderus ME, de Graaf A, Mattia E, den Blaauwen JL, Grande MA, de Jong L, van Driel R. 1992. Binding of matrix attachment regions to lamin B1. Cell 70: 949– 959. Luderus ME, den Blaauwen JL, de Smit OJ, Compton DA, van Driel R. 1994. Binding of matrix attachment regions to lamin polymers involves single-stranded regions and the minor groove. Mol Cell Biol 14: 6297– 6305. Lutz RJ, Trujillo MA, Denham KS, Wenger L, Sinensky M. 1992. Nucleoplasmic localization of prelamin A: Implications for prenylation-dependent lamin A assembly into the nuclear lamina. Proc Natl Acad Sci 89: 3000–3004. Ma L, Tsai MY, Wang S, Lu B, Chen R, Iii JR, Zhu X, Zheng Y. 2009. Requirement for Nudel and dynein for assembly of the lamin B spindle matrix. Nat Cell Biol 11: 247– 256. Machiels BM, Broers JL, Raymond Y, de Ley L, Kuijpers HJ, Caberg NE, Ramaekers FC. 1995. Abnormal A-type lamin organization in a human lung carcinoma cell line. Eur J Cell Biol 67: 328 –335. Machiels BM, Henfling ME, Schutte B, van Engeland M, Broers JL, Ramaekers FC. 1996. Subcellular localization of proteasomes in apoptotic lung tumor cells and persistence as compared to intermediate filaments. Eur J Cell Biol 70: 250– 259.

Maison C, Pyrpasopoulou A, Theodoropoulos PA, Georgatos SD. 1997. The inner nuclear membrane protein LAP1 forms a native complex with B-type lamins and partitions with spindle-associated mitotic vesicles. Embo J 16: 4839–4850. Malhas AN, Lee CF, Vaux DJ. 2009. Lamin B1 controls oxidative stress responses via Oct-1. J Cell Biol 184: 45– 55. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593 –603. Manju K, Muralikrishna B, Parnaik VK. 2006. Expression of disease-causing lamin A mutants impairs the formation of DNA repair foci. J Cell Sci 119: 2704–2714. Mariappan I, Parnaik VK. 2005. Sequestration of pRb by cyclin D3 causes intranuclear reorganization of lamin A/C during muscle cell differentiation. Mol Biol Cell 16: 1948–1960. Mariappan I, Gurung R, Thanumalayan S, Parnaik VK. 2007. Identification of cyclin D3 as a new interaction partner of lamin A/C. Biochem Biophys Res Commun 355: 981–985. Markiewicz E, Ledran M, Hutchison CJ. 2005. Remodelling of the nuclear lamina and nucleoskeleton is required for skeletal muscle differentiation in vitro. J Cell Sci 118: 409 –420. Markiewicz E, Dechat T, Foisner R, Quinlan RA, Hutchison CJ. 2002. Lamin A/C binding protein LAP2alpha is required for nuclear anchorage of retinoblastoma protein. Mol Biol Cell 13: 4401– 4413. Mattout A, Goldberg M, Tzur Y, Margalit A, Gruenbaum Y. 2007. Specific and conserved sequences in D melanogaster and C elegans lamins and histone H2A mediate the attachment of lamins to chromosomes. J Cell Sci 120: 77–85. McClintock D, Gordon LB, Djabali K. 2006. HutchinsonGilford progeria mutant lamin A primarily targets human vascular cells as detected by an anti-Lamin A G608G antibody. Proc Natl Acad Sci 103: 2154– 2159. McKeon FD, Kirschner MW, Caput D. 1986. Homologies in both primary and secondary structure between nuclear envelope and intermediate filament proteins. Nature 319: 463–468. Meaburn KJ, Cabuy E, Bonne G, Levy N, Morris GE, Novelli G, Kill IR, Bridger JM. 2007. Primary laminopathy fibroblasts display altered genome organization and apoptosis. Aging Cell 6: 139– 153. Melcer S, Gruenbaum Y, Krohne G. 2007. Invertebrate lamins. Exp Cell Res 313: 2157– 2166. Moir RD, Montag-Lowy M, Goldman RD. 1994. Dynamic properties of nuclear lamins: lamin B is associated with sites of DNA replication. J Cell Biol 125: 1201–1212. Moir RD, Yoon M, Khuon S, Goldman RD. 2000. Nuclear lamins A and B1: Different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Molloy S, Little M. 1992. p34cdc2 kinase-mediated release of lamins from nuclear ghosts is inhibited by cAMPdependent protein kinase. Exp Cell Res 201: 494– 499. Muchir A, van Engelen BG, Lammens M, Mislow JM, McNally E, Schwartz K, Bonne G. 2003. Nuclear envelope

T. Dechat et al. alterations in fibroblasts from LGMD1B patients carrying nonsense Y259X heterozygous or homozygous mutation in lamin A/C gene. Exp Cell Res 291: 352 –362. Muralikrishna B, Dhawan J, Rangaraj N, Parnaik VK. 2001. Distinct changes in intranuclear lamin A/C organization during myoblast differentiation. J Cell Sci 114: 4001– 4011. Nakajima N, Abe K. 1995. Genomic structure of the mouse A-type lamin gene locus encoding somatic and germ cellspecific lamins. FEBS Lett 365: 108– 114. Nikolova V, Leimena C, McMahon AC, Tan JC, Chandar S, Jogia D, Kesteven SH, Michalicek J, Otway R, Verheyen F, et al. 2004. Defects in nuclear structure and function promote dilated cardiomyopathy in lamin A/C-deficient mice. J Clin Invest 113: 357– 369. Ostlund C, Bonne G, Schwartz K, Worman HJ. 2001. Properties of lamin A mutants found in Emery-Dreifuss muscular dystrophy, cardiomyopathy and Dunnigan-type partial lipodystrophy. J Cell Sci 114: 4435–4445. Ottaviani A, Rival-Gervier S, Boussouar A, Foerster AM, Rondier D, Sacconi S, Desnuelle C, Gilson E, Magdinier F. 2009a. The D4Z4 macrosatellite repeat acts as a CTCF and A-type lamins-dependent insulator in facio-scapulohumeral dystrophy. PLoS Genet 5: e1000394. Ottaviani A, Schluth-Bolard C, Rival-Gervier S, Boussouar A, Rondier D, Foerster AM, Morere J, Bauwens S, Gazzo S, Callet-Bauchu E, et al. 2009b. Identification of a perinuclear positioning element in human subtelomeres that requires A-type lamins and CTCF. Embo J 28: 2428– 2436. Ottaviano Y, Gerace L. 1985. Phosphorylation of the nuclear lamins during interphase and mitosis. J Biol Chem 260: 624–632. Ozaki T, Saijo M, Murakami K, Enomoto H, Taya Y, Sakiyama S. 1994. Complex formation between lamin A and the retinoblastoma gene product: Identification of the domain on lamin A required for its interaction. Oncogene 9: 2649–2653. Paddy MR, Belmont AS, Saumweber H, Agard DA, Sedat JW. 1990. Interphase nuclear envelope lamins form a discontinuous network that interacts with only a fraction of the chromatin in the nuclear periphery. Cell 62: 89–106. Pajerowski JD, Dahl KN, Zhong FL, Sammak PJ, Discher DE. 2007. Physical plasticity of the nucleus in stem cell differentiation. Proc Natl Acad Sci 104: 15619–15624. Panorchan P, Wirtz D, Tseng Y. 2004b. Structure-function relationship of biological gels revealed by multipleparticle tracking and differential interference contrast microscopy: The case of human lamin networks. Phys Rev E Stat Nonlin Soft Matter Phys 70: 041906. Panorchan P, Schafer BW, Wirtz D, Tseng Y. 2004a. Nuclear envelope breakdown requires overcoming the mechanical integrity of the nuclear lamina. J Biol Chem 279: 43462– 43467. Parry DA, Conway JF, Steinert PM. 1986. Structural studies on lamin. Similarities and differences between lamin and intermediate-filament proteins. Biochem J 238: 305 –308. Pegoraro G, Kubben N, Wickert U, Gohler H, Hoffmann K, Misteli T. 2009. Ageing-related chromatin defects through loss of the NURD complex. Nat Cell Biol 11: 1261– 1267.

Peter M, Kitten GT, Lehner CF, Vorburger K, Bailer SM, Maridor G, Nigg EA. 1989. Cloning and sequencing of cDNA clones encoding chicken lamins A and B1 and comparison of the primary structures of vertebrate Aand B-type lamins. J Mol Biol 208: 393– 404. Peter M, Nakagawa J, Doree M, Labbe JC, Nigg EA. 1990. In vitro disassembly of the nuclear lamina and M phase-specific phosphorylation of lamins by cdc2 kinase. Cell 61: 591–602. Pickersgill H, Kalverda B, de Wit E, Talhout W, Fornerod M, van Steensel B. 2006. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat Genet 38: 1005–1014. Prather RS, Sims MM, Maul GG, First NL, Schatten G. 1989. Nuclear lamin antigens are developmentally regulated during porcine and bovine embryogenesis. Biol Reprod 41: 123– 132. Rauner M, Sipos W, Goettsch C, Wutzl A, Foisner R, Pietschmann P, Hofbauer LC. 2009. Inhibition of lamin A/C attenuates osteoblast differentiation and enhances RANKL-dependent osteoclastogenesis. J Bone Miner Res 24: 78–86. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243 –247. Riemer D, Stuurman N, Berrios M, Hunter C, Fisher PA, Weber K. 1995. Expression of Drosophila lamin C is developmentally regulated: Analogies with vertebrate A-type lamins. J Cell Sci 108: 3189– 3198. Rober RA, Weber K, Osborn M. 1989. Differential timing of nuclear lamin A/C expression in the various organs of the mouse embryo and the young animal: A developmental study. Development 105: 365–378. Rober RA, Sauter H, Weber K, Osborn M. 1990. Cells of the cellular immune and hemopoietic system of the mouse lack lamins A/C: distinction versus other somatic cells. J Cell Sci 95: 587–598. Rodriguez S, Coppede F, Sagelius H, Eriksson M. 2009. Increased expression of the Hutchinson-Gilford progeria syndrome truncated lamin A transcript during cell aging. Eur J Hum Genet 17: 928– 937. Rusinol AE, Sinensky MS. 2006. Farnesylated lamins, progeroid syndromes and farnesyl transferase inhibitors. J Cell Sci 119: 3265– 3272. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059–1063. Scaffidi P, Misteli T. 2008. Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10: 452 –459. Schape J, Prausse S, Radmacher M, Stick R. 2009. Influence of lamin A on the mechanical properties of amphibian oocyte nuclei measured by atomic force microscopy. Biophys J 96: 4319– 4325. Schatten G, Maul GG, Schatten H, Chaly N, Simerly C, Balczon R, Brown DL. 1985. Nuclear lamins and peripheral nuclear antigens during fertilization and embryogenesis in mice and sea urchins. Proc Natl Acad Sci 82: 4727– 4731. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of

Nuclear Lamins the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332– 1336. Shevelyov YY, Lavrov SA, Mikhaylova LM, Nurminsky ID, Kulathinal RJ, Egorova KS, Rozovsky YM, Nurminsky DI. 2009. The B-type lamin is required for somatic repression of testis-specific gene clusters. Proc Natl Acad Sci 106: 3282–3287. Shimi T, Pfleghaar K, Kojima S, Pack CG, Solovei I, Goldman AE, Adam SA, Shumaker DK, Kinjo M, Cremer T, et al. 2008. The A- and B-type nuclear lamin networks: microdomains involved in chromatin organization and transcription. Genes Dev 22: 3409– 3421. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Shumaker DK, Lopez-Soler RI, Adam SA, Herrmann H, Moir RD, Spann TP, Goldman RD. 2005. Functions and dysfunctions of the nuclear lamin Ig-fold domain in nuclear assembly, growth, and Emery-Dreifuss muscular dystrophy. Proc Natl Acad Sci USA 102: 15494–15499. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin Ig-fold binds to PCNA: Its role in DNA replication. J Cell Biol 181: 269 –280. Solovei I, Schermelleh L, During K, Engelhardt A, Stein S, Cremer C, Cremer T. 2004. Differences in centromere positioning of cycling and postmitotic human cell types. Chromosoma 112: 410 –423. Spann TP, Goldman AE, Wang C, Huang S, Goldman RD. 2002. Alteration of nuclear lamin organization inhibits RNA polymerase II-dependent transcription. J Cell Biol 156: 603– 608. Stewart-Hutchinson PJ, Hale CM, Wirtz D, Hodzic D. 2008. Structural requirements for the assembly of LINC complexes and their function in cellular mechanical stiffness. Exp Cell Res 314: 1892– 1905. Stewart C, Burke B. 1987. Teratocarcinoma stem cells and early mouse embryos contain only a single major lamin polypeptide closely resembling lamin B. Cell 51: 383–392. Stick R, Angres B, Lehner CF, Nigg EA. 1988. The fates of chicken nuclear lamin proteins during mitosis: Evidence for a reversible redistribution of lamin B2 between inner nuclear membrane and elements of the endoplasmic reticulum. J Cell Biol 107: 397 –406. Strelkov SV, Schumacher J, Burkhard P, Aebi U, Herrmann H. 2004. Crystal structure of the human lamin A coil 2B dimer: Implications for the head-to-tail association of nuclear lamins. J Mol Biol 343: 1067– 1080. Stuurman N, Heins S, Aebi U. 1998. Nuclear lamins: Their structure, assembly, and interactions. J Struct Biol 122: 42–66. Stuurman N, Sasse B, Fisher PA. 1996. Intermediate filament protein polymerization: Molecular analysis of Drosophila nuclear lamin head-to-tail binding. J Struct Biol 117: 1 –15. Sullivan T, Escalante-Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B. 1999. Loss of A-type

lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147: 913 –920. Taimen P, Pfleghaar K, Shimi T, Moller D, Ben-Harush K, Erdos MR, Adam SA, Herrmann H, Medalia O, Collins FS, et al. 2009a. A progeria mutation reveals functions for lamin A in nuclear assembly, architecture, and chromosome organization. Proc Natl Acad Sci 106: 20788– 20793. Taimen P, Pfleghaar K, Shimi T, Moller D, Ben-Harush K, Erdos MR, Adam SA, Herrmann H, Medalia O, Collins FS, et al. 2009b. A progeria mutation reveals functions for lamin A in nuclear assembly, architecture, and chromosome organization. Proc Natl Acad Sci 106: 20788– 20793. Tang CW, Maya-Mendoza A, Martin C, Zeng K, Chen S, Feret D, Wilson SA, Jackson DA. 2008. The integrity of a lamin-B1-dependent nucleoskeleton is a fundamental determinant of RNA synthesis in human cells. J Cell Sci 121: 1014– 1024. Taniura H, Glass C, Gerace L. 1995. A chromatin binding site in the tail domain of nuclear lamins that interacts with core histones. J Cell Biol 131: 33– 44. Thompson LJ, Bollen M, Fields AP. 1997. Identification of protein phosphatase 1 as a mitotic lamin phosphatase. J Biol Chem 272: 29693– 29697. Tilgner K, Wojciechowicz K, Jahoda C, Hutchison C, Markiewicz E. 2009. Dynamic complexes of A-type lamins and emerin influence adipogenic capacity of the cell via nucleocytoplasmic distribution of beta-catenin. J Cell Sci 122: 401–413. Tsai MY, Zheng Y. 2005. Aurora A kinase-coated beads function as microtubule-organizing centers and enhance RanGTP-induced spindle assembly. Curr Biol 15: 2156–2163. Tsai MY, Wang S, Heidinger JM, Shumaker DK, Adam SA, Goldman RD, Zheng Y. 2006. A mitotic lamin B matrix induced by RanGTP required for spindle assembly. Science 311: 1887– 1893. Tzur YB, Wilson KL, Gruenbaum Y. 2006. SUN-domain proteins: ‘Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782–788. Vaughan A, Alvarez-Reyes M, Bridger JM, Broers JL, Ramaekers FC, Wehnert M, Morris GE, Whitfield WGF, Hutchison CJ. 2001. Both emerin and lamin C depend on lamin A for localization at the nuclear envelope. J Cell Sci 114: 2577– 2590. Vergnes L, Peterfy M, Bergo MO, Young SG, Reue K. 2004. Lamin B1 is required for mouse development and nuclear integrity. Proc Natl Acad Sci 101: 10428– 10433. Verstraeten VL, Broers JL, Ramaekers FC, van Steensel MA. 2007. The nuclear envelope, a key structure in cellular integrity and gene expression. Curr Med Chem 14: 1231–1248. Vigouroux C, Auclair M, Dubosclard E, Pouchelet M, Capeau J, Courvalin JC, Buendia B. 2001. Nuclear envelope disorganization in fibroblasts from lipodystrophic patients with heterozygous R482Q/W mutations in the lamin A/C gene. J Cell Sci 114: 4459– 4468. Vorburger K, Kitten GT, Nigg EA. 1989. Modification of nuclear lamin proteins by a mevalonic acid derivative

T. Dechat et al. occurs in reticulocyte lysates and requires the cysteine residue of the C-terminal CXXM motif. Embo J 8: 4007– 4013. Ward GE, Kirschner MW. 1990. Identification of cell cycle-regulated phosphorylation sites on nuclear lamin C Cell 61: 561– 577. Wheeler MA, Davies JD, Zhang Q, Emerson LJ, Hunt J, Shanahan CM, Ellis JA. 2007. Distinct functional domains in nesprin-1a and nesprin-2b bind directly to emerin and both interactions are disrupted in X-linked Emery-Dreifuss muscular dystrophy. Exp Cell Res 313: 2845– 2857. Williams RR, Azuara V, Perry P, Sauer S, Dvorkina M, Jorgensen H, Roix J, McQueen P, Misteli T, Merkenschlager M, et al. 2006. Neural induction promotes largescale chromatin reorganisation of the Mash1 locus. J Cell Sci 119: 132–140. Winter-Vann AM, Casey PJ. 2005. Post-prenylationprocessing enzymes as new targets in oncogenesis. Nat Rev Cancer 5: 405– 412. Yuan J, Simos G, Blobel G, Georgatos SD. 1991. Binding of lamin A to polynucleosomes. J Biol Chem 266: 9211– 9215.

Zackroff RV, Goldman RD. 1979. In vitro assembly of intermediate filaments from baby hamster kidney (BHK-21) cells. Proc Natl Acad Sci 76: 6226–6230. Zhang FL, Casey PJ. 1996. Protein prenylation: Molecular mechanisms and functional consequences. Annu Rev Biochem 65: 241– 269. Zhang YQ, Sarge KD. 2008. Sumoylation regulates lamin A function and is lost in lamin A mutants associated with familial cardiomyopathies. J Cell Biol 182: 35– 39. Zhang Q, Bethmann C, Worth NF, Davies JD, Wasner C, Feuer A, Ragnauth CD, Yi Q, Mellad JA, Warren DT, et al. 2007. Nesprin-1 and -2 are involved in the pathogenesis of Emery Dreifuss muscular dystrophy and are critical for nuclear envelope integrity. Hum Mol Genet 16: 2816–2833. Zhao K, Harel A, Stuurman N, Guedalia D, Gruenbaum Y. 1996. Binding of matrix attachment regions to nuclear lamin is mediated by the rod domain and depends on the lamin polymerization state. FEBS Lett 380: 161– 164. Zink D, Amaral MD, Englmann A, Lang S, Clarke LA, Rudolph C, Alt F, Luther K, Braz C, Sadoni N, et al. 2004. Transcription-dependent spatial arrangements of CFTR and adjacent genes in human cell nuclei. J Cell Biol 166: 815 –825.

Lamin-binding Proteins Katherine L. Wilson1 and Roland Foisner2 1

Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, Maryland 21205

2

Max F. Perutz Laboratories, Department of Medical Biochemistry, Medical University of Vienna, Vienna 1030

Correspondence: [email protected]

A- and B-type lamins are the major intermediate filaments of the nucleus. Lamins engage in a plethora of stable and transient interactions, near the inner nuclear membrane and throughout the nucleus. Lamin-binding proteins serve an amazingly diverse range of functions. Numerous inner-membrane proteins help anchor lamin filaments to the nuclear envelope, serving as part of the nuclear “lamina” network that is essential for nuclear architecture and integrity. Certain lamin-binding proteins of the inner membrane bind partners in the outer membrane and mechanically link lamins to the cytoskeleton. Inside the nucleus, lamin-binding proteins appear to serve as the “adaptors” by which the lamina organizes chromatin, influences gene expression and epigenetic regulation, and modulates signaling pathways. Transient interactions of lamins with key components of the transcription and replication machinery may provide an additional level of regulation or support to these essential events.

he eukaryotic cell nucleus is a complex membrane-bounded organelle that houses, organizes, and regulates the genome. The nucleus is structurally organized into functional domains, one of which is the nuclear envelope (NE). The NE has two concentric membranes, named the “inner” and “outer” nuclear membranes (INM and ONM, respectively). These membranes are separated by a 30– 50 nm lumen, and fuse to form holes ( pores) occupied by nuclear pore complexes (NPCs), which mediate active and passive movement of molecules between the cytoplasm and nucleoplasm (Gruenbaum et al. 2005; Stewart et al. 2007). The NE and its lumen are continuous with the endoplasmic reticulum (ER) and share many

T

ER functions. However, the INM and ONM are also each structurally and functionally unique, because of specific enrichments for distinct integral membrane proteins (Schirmer and Gerace 2005; Schirmer and Foisner 2007). In mammals, the INM in particular appears to be populated by over 50 different membrane proteins, most of which are uncharacterized. Among characterized INM proteins, most can bind directly to A- or B-type lamins, or both. A- and B-type lamins polymerize to form separate networks of nuclear intermediate filaments that concentrate near the INM in metazoans (Dechat et al. 2008). Many INM proteins are localized by binding directly or indirectly to lamin filaments. This network of filaments and

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000554 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000554

K.L. Wilson and R. Foisner

lamin-binding proteins at the INM is known as the “peripheral lamina.” B-type lamins are essential for cell viability and development, whereas A-type lamins arose later in evolution and are nonessential. Lamins are important structurally and as “scaffolds” for many other proteins and complexes in the nucleus. Lamin filaments are important for the assembly, structure, shape, and mechanical stability of metazoan nuclei. There is also growing evidence that lamins regulate chromatin organization and gene expression, and influence signaling (Gruenbaum et al. 2005; Dechat et al. 2008). These functions involve a full spectrum of biochemical interactions between lamins, chromatin, and a variety of partners, including regulatory proteins responsive to external and intrinsic signals (Fig. 1). The pace of discovering new lamin-binding proteins has nearly

overwhelmed our capacity to characterize them. Each new partner has the potential to provide fresh insight into the structure and regulation of the nucleoskeleton and its relationships with the genome. Further motivation for research in this area comes with the discovery that an increasing number of human diseases are linked to defects in lamins or lamin-binding proteins. THE CONCEPT OF LAMINA-ASSOCIATED POLYPEPTIDES

The term lamina-associated-polypeptide (LAP) was coined to define a novel group of nuclear proteins that cofractionated with lamins and required high concentrations of monovalent salts and nonionic detergents to extract them from nuclei (Senior and Gerace 1988; Foisner

Cytoskeleton NPC ONM INM

Reinforce nucleoskeleton

Link nucleoskeleton to cytoskeleton

Regulate signaling

NPC anchoring assembly

Chromatin tethering

Retention at INM Lamin filaments

Gene regulation

Lamin assembly

Regulate signaling/ transcription activity

Lamin-binding proteins

Figure 1. Overview of known roles for lamin-binding proteins. Many lamin-binding proteins located in the

INM, on chromatin, and in the nucleoplasm are thought to have mechanical and structural roles, such as reinforcing the nucleoskeleton, interlinking the nucleoskeleton and cytoskeleton, anchoring NPCs, and tethering chromatin to the nuclear envelope. Others regulate signaling or transcription. Many lamin-binding proteins require lamins for their correct localization, whereas others regulate or facilitate lamin assembly.

Lamin-binding Proteins

and Gerace 1993). The first characterized LAPs were all integral INM proteins: LAP1 (Martin et al. 1995) and LAP2 (Harris et al. 1994; Furukawa et al. 1995; Berger et al. 1996) each has one transmembrane domain and is expressed as multiple isoforms, whereas the so-called lamin B receptor (LBR) has eight transmembrane domains (Worman et al. 1990). These proteins all bind lamins directly in vitro (Worman et al. 1988; Foisner and Gerace 1993; Furukawa et al. 1998). Since then, around 80 additional putative NE membrane proteins have been discovered (Schirmer et al. 2005; Schirmer and Foisner 2007). Most fit the original LAP criterion (resistance to biochemical extraction) and localize at the NE, but are untested for binding to lamins. The LAP concept expanded as lamins (Moir et al. 2000b; Naetar et al. 2008) and lamin-binding proteins including Narf (Barton and Worman 1999), LAP2a (Naetar et al. 2008), and Lco1 (Vlcek et al. 2004; Naetar et al. 2008) were discovered in the nuclear interior. Laminbinding proteins can have widely differing, and differentially regulated, affinities for lamins that might influence their biochemical extraction. Some proteins are known to bind lamins directly and relatively stably at the NE or in the nuclear interior. By contrast, other proteins bind lamins more transiently, probably as a means of regulating their activities; these partners include PCNA (Shumaker et al. 2008), c-Fos (Gonzalez et al. 2008), retinoblastoma (Rb) (Johnson et al. 2004; Pekovic et al. 2007), and Oct-1 (Malhas et al. 2009). LEM-DOMAIN PROTEINS AND BAF: TETHERING CHROMATIN TO THE NUCLEOSKELETON

One prominent family of lamin-binding proteins involved in nuclear architecture and chromatin organization share the LEM (LAP2, Emerin, MAN) domain, an 45-residue motif that folds as two a-helices (Laguri et al. 2001). The LEM domain binds a conserved metazoan chromatin protein named Barrier to Autointegration Factor, BAF (Furukawa 1999; Cai et al. 2001; Lee et al. 2001; Shumaker et al. 2001; Shimi et al. 2004; Cai et al. 2007). BAF, a mobile

lamin-binding protein, can “bridge” DNA and interacts with histones (Margalit et al. 2007). Most, but not all, LEM proteins are integral INM proteins with one or two transmembrane domains (Fig. 2). Mammals have four genes encoding characterized LEM proteins, LAP2 (a, b, and other isoforms), MAN1, emerin (Lin et al. 2000), and LEM2/NET25 (Schirmer et al. 2003; Brachner et al. 2005; Chen et al. 2006; Ulbert et al. 2006), and three uncharacterized genes encoding predicted nonmembrane proteins, LEM3, LEM4, and LEM5 (Lee and Wilson 2004), known respectively as ANKLE1, ANKLE2, and LEMD1. Three LEM proteins (emerin, LEM2, and LEM3) are conserved in Caenorhabditis elegans (Lee et al. 2000; Gruenbaum et al. 2002; Liu et al. 2003). Four are expressed in Drosophila: MAN1 (Wagner et al. 2006), otefin (Goldberg et al. 1998; Jiang et al. 2008), and Bocksbeutel a and b(Wagner et al. 2004). The functions and interactions of LEM proteins, lamins, and BAF are strongly conserved between mammals, nematodes, and flies, suggesting they have fundamental roles in the nucleus. Some LEM-domain proteins have additional domains that bind DNA, or other chromatin proteins. For example, LAP2b can bind HA95, a chromatin protein involved in DNA replication (Martins et al. 2003). All isoforms of LAP2 have a second “LEM-like” domain that binds DNA, rather than BAF (Cai et al. 2001; Laguri et al. 2001). Similarly, MAN1 and probably LEM2 binds DNA directly via a carboxyterminal “winged helix” domain (Caputo et al. 2006). Furthermore, all tested LEM proteins bind either A- or B-type lamins, or both, directly (Clements et al. 2000; Lee et al. 2001; Sakaki et al. 2001; Brachner et al. 2005; Mansharamani and Wilson 2005), and some require this interaction to localize at the NE (Sullivan et al. 1999; Vaughan et al. 2001; Brachner et al. 2005). Lamins, in turn, also directly bind DNA (Shoeman and Traub 1990; Luderus et al. 1992; Stierle et al. 2003) and histones (Taniura et al. 1995; Mattout et al. 2007). Thus, a complex network of interactions between lamins, LEM proteins, BAF, and, most likely, other INM proteins is involved in anchoring chromatin to the NE and lamins.

K.L. Wilson and R. Foisner

Ce-Emerin Dm-Bocksbeutel Dm-Otefin h-Emerin

Ce-LEM2 Dm-MAN1 h-MAN1 h-LEM2

h-LAP2-beta

ONM INM Lamina

HA95 BAF

HDAC3

BAF

BAF

BAF

Rb

Gcl

Gcl Lmo7 Btf β-catenin

Gcl Smad

h-LAP2-alpha Lamin BAF

LEM

LEM-like

Winged Helix

Figure 2. LEM-domain proteins organize chromatin and regulate signaling and transcription. Domain

organizations of characterized LEM proteins; the LEM motif confers direct binding to barrier to autointegration factor (BAF). All LEM proteins can bind A- and/or B-type lamins. Some, including human (h-)emerin, C. elegans (Ce-) emerin, D. melanogaster (Dm-) bocksbeutel a, and Dm-Otefin, have one transmembrane (TM) domain. LAP2 proteins, expressed only in vertebrates, also have a DNA-binding “LEM-like” domain. Others (e.g., hLem2, Ce-Lem2, hMAN1, and Dm-MAN1) have two TM domains and a carboxy-terminal Winged Helix domain that binds DNA. Some, including hLAP2a (shown) and three uncharacterized LEM proteins (not shown), have no TM domain. LEM proteins have additional functional domains; e.g., LAP2b partners include replication protein HA95 and HDAC3; LAP2b recruits HDAC3 to the NE and contributes to epigenetic regulation. Emerin binds many transcription factors including GCL, Lmo7, Btf, and b-catenin. GCL also binds hLAP2b and hMAN1, suggesting functional overlap. MAN1 is unique in binding regulatory Smads, and inhibits BMP and TGF-b signaling.

LEM proteins might appear to function redundantly in chromatin organization because there are so many of them, and functional inactivation of any single tested LEM gene does not grossly disrupt chromatin organization. Functional overlap is supported by C. elegans studies, which showed that double-knockdown of two LEM proteins (Ce-emerin and Ce-lem2) caused embryonic lethality at the 100-cell stage, when embryonic gene expression normally begins, with phenotypes that included aneuploidy, grossly defective chromatin organization and nuclear structure, and failure to assemble nuclei after mitosis (Liu et al. 2003). Nearly identical phenotypes are caused by down-regulating

either Ce-lamin or BAF alone (Gruenbaum et al. 2005; Margalit et al. 2005a), strongly suggesting that BAF and LEM-domain proteins are key components of nuclear architecture. Nevertheless, despite potential redundancies, eliminating just one LEM protein, for example emerin in mice, can affect nuclear structure: Emerin-null skeletal muscle nuclei are fragile (Ozawa et al. 2006), and show significantly increased autophagic degradation of structurally aberrant regions of the nucleus (Park et al. 2009a). Lamin mutations also cause obvious defects in nuclear shape and chromatin organization (Sabatelli et al. 2001; Scaffidi and Misteli 2006; Shumaker et al. 2006; Wang

Lamin-binding Proteins

et al. 2006; Hakelien et al. 2008; Park et al. 2009b), as well as enhanced autophagic degradation of nuclei (Park et al. 2009a). Further evidence for the physiological relevance of both lamins and LEM proteins in chromatin organization comes from recent genomewide studies to identify lamina-bound DNA. The analysis of methylated DNA in cultured Drosophila cells expressing lamin B1 fused to DNA methyltransferase (Dam), revealed 500 genes in close contact with the lamina (Pickersgill et al. 2006). In human fibroblasts, this strategy revealed both lamin B1 and emerin in close contact with .1300 sharply defined domains in the genome, most of which were flanked either by binding sites for the insulator protein CTCF, or by E2F/DP-regulated promoters (Guelen et al. 2008). Interestingly, at least two INM LEM proteins, LAP2b and emerin, are implicated in the regulation of E2F/DP promoters (Nili et al. 2001; Holaska and Wilson 2006), and both CTCF and A-type lamins are required for the insulator function of a subtelomeric array of D4Z4 repeats, shortening of which cause Facio-Scapulo-Humeral Dystrophy (FSHD) (Ottaviani et al. 2009). Future application of genomewide “close contact” studies to other LEM proteins, and other types of lamin-binding proteins, may provide much-needed insight into their relationships with chromosomes. Interestingly, in proliferating cells, these “relationships” might be established actively while nuclei reassemble during mitotic anaphase and telophase. Nuclear assembly involves a complex series of interactions between chromatin, membranes, INM and ONM proteins, lamins, and other mitotically solubilized components, the mechanisms and regulation of which remain poorly understood (Margalit et al. 2005b; Anderson and Hetzer 2008; Guttinger et al. 2009). However, protein – protein interactions, for example between BAF, lamins, and LEM proteins, are required because nuclear assembly fails when one component is missing, as shown in C. elegans studies (Margalit et al. 2005b), and also fails in HeLa cells that express emerin and BAF mutants with impaired interactions (Haraguchi et al. 2001).

LINC COMPLEXES LINK THE NUCLEAR INTERIOR TO THE CYTOPLASM

Many independent studies in yeast, worms, flies, and mammals converged on a major discovery: Membrane proteins of the NE mechanically interlink the nucleoskeleton and cytoskeleton. These “LINC” complexes (Crisp et al. 2006) enable the regulated, cytoplasmic-motor-driven movement of the entire nucleus to new positions within the cell, and the movement of entire NE-tethered chromosomes to new positions along the NE (for reviews, see Tzur et al. 2006b; Wilhelmsen et al. 2006; Starr 2009). In addition, gene-regulatory responses to external force must now be assumed to potentially involve not only indirect signaling from the cell surface, but also direct signaling via mechanotransducing complexes at the NE. Considered at the simplest level, LINC complexes consist of lamin-binding SUN-domain proteins in the INM that interact in the NE lumen with KASH-domain proteins (Fig. 3) anchored in the ONM (Hodzic et al. 2004; Padmakumar et al. 2005; Crisp et al. 2006; Haque et al. 2006); KASH proteins also bind directly or indirectly to cytoplasmic actin, microtubules, centrosomes, or intermediate filaments (Zhang et al. 2001; Zhen et al. 2002; Padmakumar et al. 2004; Wilhelmsen et al. 2005; McGee et al. 2006; Roux et al. 2009). It is important to note that at least two of the many KASH-domain protein isoforms (nesprin-1a and nesprin-2b) are INM proteins that bind directly to lamins and emerin (Mislow et al. 2002a; Mislow et al. 2002b; Zhang et al. 2005). SUN-domain proteins, conserved from yeast to humans, are defined by a 120-residue motif located in the lumenal space of the NE; the name SUN was based on S. pombe Sad1 and D. melanogaster UNC-84 (Malone et al. 1999). C. elegans and D. melanogaster each have two SUN genes: UNC84 and germ-cell-specific matefin/SUN1 in C. elegans, and Klaroid and CG6589 in Drosophila. Mammals have four: SUN1 and SUN2, and testis-specific genes SUN3 and Spag4 (reviewed in Starr 2009). Human SUN1 (hSUN1) and hSUN2 each have a coiled-coil domain near the SUN domain that

K.L. Wilson and R. Foisner

KASH-domain proteins Ce-ANC-1 Nesprin-1/2

h-Nesprin 3

Ce-ZYG-12 Dm-Klarsicht

Ce-ZYG-12

Intermediate Filament Actin

Centrosomes

Microtubule

Plectin Dynein Dynein

ONM INM Him-8 ?

Lamina h-Nesprin 1/2 Ce-UNC84 h-SUN1/2

Ce-UNC84 Dm-Klaroid

Meiotic chromosome h-SUN1/2

Ce-Matefin h-SUN 1

SUN-domain proteins

SUN domain

KASH domain

Actin binding domain

Figure 3. SUN- and KASH-domain proteins interact in the NE lumen to form LINC (Links Nucleoskeleton and

Cytoskeleton) complexes. SUN-domain proteins span the INM and dimerize; each nucleoplasmic domain binds lamins and each SUN-domain in the NE lumen binds the KASH domain of an ONM-embedded KASH protein, which binds actin, centrosomes/microtubules, or plectin/intermediate filaments in the cytoplasm. Only one KASH domain partner of a SUN dimer is depicted. SUN proteins also bind, potentially via Him-8, the pairing centers of meiotic chromosomes. Many additional nesprin isoforms reside elsewhere, including the INM; e.g., human nesprins-1a and -2b in the INM bind lamins and emerin (not shown).

mediates dimerization and is also involved in binding the KASH domain; their exposed nucleoplasmic amino-terminal domain binds lamins (reviewed by Tzur et al. 2006b; Starr 2009). KASH- (Klarsicht, ANC-1, Syne homology) domain nomenclature is confusing because these proteins were identified in diverse species before their conservation was recognized. Mammalian KASH proteins are now known as “nesprins.” The KASH domain comprises a single carboxy-terminal transmembrane domain followed by a 35-residue lumenal domain. ONM-localized KASH proteins require the interaction with SUN-domain proteins for retention at the ONM. C. elegans encodes

three known KASH-domain proteins (ANC-1, UNC-83, and ZYG-12) (Starr and Han 2002; Malone et al. 2003; McGee et al. 2006) and D. melanogaster encodes the two proteins Klarsicht and MSP 300 (Patterson et al. 2004; Xie and Fischer 2008). Nesprin isoform complexity is much higher in mammals, which have four known genes: Nesprin-1 (SYNE1), nesprin-2 (SYNE2), nesprin-3, and nesprin-4. The Nesprin-1 and -2 genes are alternatively transcribed and alternatively spliced to produce more than 12 protein isoforms each, from small to enormous (.1 MDa). Some isoforms lack the KASH domain and are not membrane-localized, whereas others localize at the INM or even the Golgi complex (Starr and Fischer 2005; Wilhelmsen et al.

Lamin-binding Proteins

2006). All nesprins have multiple “spectrin repeat” domains that confer an extended configuration, but can also mediate specific protein – protein interactions. For example, specific spectrin repeat domains in the small, INM-localized isoform nesprin-1a mediate direct binding to lamins, emerin, and other nesprin-1a molecules (Mislow et al. 2002a). The largest isoforms of nesprin-1 and -2 bind actin (Zhen et al. 2002; Padmakumar et al. 2004). Nesprin-3 binds plectin, a cytoskeletal protein that binds cytoplasmic intermediate filaments (Wilhelmsen et al. 2005), whereas nesprin-4 binds kinesin 1, a plus-enddirected microtubule-dependent motor, and may be involved in dislocalizing the centrosome and Golgi membranes away from the nucleus in epithelial cells (Roux et al. 2009). LINC complexes formed by SUN- and KASHdomain proteins are known to determine the spacing between the INM and ONM (Crisp et al. 2006), position NPCs (Liu et al. 2007), and control nuclear size and NE architecture (Luke et al. 2008). Different combinations of SUN- and KASH-domain proteins mediate different functions (Fig. 3), including forcetransfer across the NE, and the stiffness of the cytoskeleton (Stewart-Hutchinson et al. 2008). In C. elegans, the SUN protein UNC84 anchors nuclei to cytoplasmic actin filaments viathe KASHdomain protein ANC-1 (Malone et al. 1999; Starr and Han 2002), mediates movement of nuclei via a different KASH partner (UNC-83) (Starr et al. 2001), and attaches centrosomes to the NE via a third KASH partner (ZYG-12) and the dynein subunit DLI-1 (Malone et al. 2003) (Fig. 3). The other C. elegans SUN-domain protein, matefin/SUN1, is also required to recruit the proapoptotic protein CED-4 to the NE (Tzur et al. 2006a), implying additional roles for LINC complexes in death signaling. In Drosophila, LINC complexes mediate centrosome attachment to the NE, and nuclear migration in developing photoreceptor cells (Patterson et al. 2004; Kracklauer et al. 2007). In mice, deletions in the nesprin-1 or nesprin-2 genes, or overexpression of the KASH-domain, which destroys endogenous LINC complexes, disrupts nuclear organization, results in failure of nuclei to cluster at neuromuscular junctions (Grady et al. 2005;

Zhang et al. 2007), and can cause muscular dystrophy (Puckelwartz et al. 2009). Amazingly, LINC complexes can also transmit mechanical force from the cytoskeleton through the NE to move chromosomes, forexample during meiotic prophase, when telomeres attach to the NE and cluster to form bouquet-like structures that facilitate chromosome pairing (Chikashige et al. 2007). In a C. elegans matefin/ SUN1 mutant strain, chromosome reorganization is disrupted in early meiosis, and ZYG-12 “patches” do not form on the NE (Penkner et al. 2007). This suggests meiotic chromosome clustering and homolog recognition requires both matefin/SUN1 and ZYG-12 (a KASH-domain protein), potentially via HIM-8-mediated attachment of pairing centers to matefin/SUN1 (Penkner et al. 2007). Similarly, mammalian SUN1 and SUN2 (Ding et al. 2007; Schmitt et al. 2007) are telomere-associated between the leptotene and diplotene stages of meiosis, and SUN1-knockout mice have both impaired NE-telomere association and impaired homolog pairing (Ding et al. 2007). The meiotic functions of mammalian SUN proteins do not require A-type lamins (Schmitt et al. 2007). Potential roles for B-type lamin(s) remain to be tested in mammals, and seem likely because in C. elegans the nuclear migration and nuclear anchoring functions of LINC complexes require the one (B-type) lamin expressed in this organism (Lee et al. 2002). However, similar SUN- and KASH-dependent pathways exist in S. pombe and S. cerevisiae, which do not have lamins (Starr 2009). It is unknown how the functions and interactions of SUN and KASH proteins are regulated (Starr 2009). One interesting possibility is that LINC complexes might be regulated by proteins located in the NE lumen. This possibility is suggested by Torsin A, a so-called “AAAþ ATPase” enzyme located in the NE/ER lumen that is required to localize nesprin (Nery et al. 2008). REGULATION OF SIGNALING AND DIFFERENTIATION BY LAMIN-BINDING PROTEINS

The plasma membrane contains many structural complexes (e.g., focal adhesions and dystroglycan

K.L. Wilson and R. Foisner

complex) that can be dynamically regulated and generate signals to which cells respond in a tissue-specific manner, in many cases at the level of gene regulation. There is increasing evidence that this paradigm might also apply to laminassociated protein complexes at the NE, and in the nucleoplasm. The concept of “lamin-linked dynamic signaling” is supported by studies on several LEM-domain proteins. For example, LAP2b, the largest INM-localized LAP2 isoform, binds lamin B specifically, and also directly interacts with transcription regulators (e.g., GCL) and epigenetic modifiers including HDAC3, and contributes to transcriptional repression (Nili et al. 2001; Somech et al. 2005). Interestingly, another INM-localized LEM protein, MAN1, directly binds and inhibits R-Smads and functions as a major regulator of BMP- and TGF-b-signaling in early vertebrate development (Osada et al. 2003; Raju et al. 2003; Lin et al. 2005; Mansharamani and Wilson 2005; Pan et al. 2005; Cohen et al. 2007). Roles for lamin-binding proteins in TGF-b-signaling are also reported in Drosophila: The INM-localized LEM protein Otefin binds directly to Medea (Smad-4) and they physically tether their repressed target locus, bam (“bag of marbles”), at the NE (Jiang et al. 2008). The INM protein emerin also supports the paradigm of complex regulation of protein – protein interactions and signaling at the NE. Emerin directly binds more than ten known partners, suggesting a degree of biochemical complexity at the NE that rivals many cell surface structures. Among these partners are structural proteins (nesprin-1a, nesprin-2b, actin, nuclear myosin 1c, lamins, and tubulin), other INM proteins (MAN1 and LUMA), proteins involved in signaling, transcription, mRNA splicing (b-catenin, GCL, Lmo7, Btf, and YT521-B), and of course, BAF (Lee et al. 2001; Mislow et al. 2002a; Holaska et al. 2003; Haraguchi et al. 2004; Holaska et al. 2004; Mansharamani and Wilson 2005; Holaska et al. 2006; Markiewicz et al. 2006; Wheeler et al. 2007; Bengtsson and Otto 2008) (for reviews, see Bengtsson and Wilson 2004; Wagner and Krohne 2007). The biochemical purification of emerin-associated

protein complexes from HeLa cell nuclei suggests emerin can form at least six distinct multiprotein complexes (Holaska and Wilson 2007). Some putative complexes included primarily architectural components (e.g., lamins, actin, nuclear myosin 1c, and nuclear aII-spectrin), whereas others included chromatin and gene regulators (e.g., histones, Lmo7, Rb, and components of the Nuclear Co-Repressor [NCoR] complex). Emerin may also be involved in sensing and responding to mechanical tension at the NE (Lammerding et al. 2005). The interaction of Lmo7 (Holaska et al. 2006) and b-catenin (Markiewicz et al. 2006) with emerin regulates their nuclear export and represses their activity. These regulatory functions may be important for muscle and adipocyte differentiation, respectively (Holaska et al. 2006; Tilgner et al. 2009). It is still unclear how emerin regulates the nuclear export of these factors. Emerin was also reported to localize in the ONM and mediate centrosome-NE attachment (Salpingidou et al. 2007). How emerin “knows” when to assemble or disassemble particular complexes, and how its localization in the INM versus ONM is regulated, are important open questions. Growing evidence that emerin is phosphorylated suggests mechanisms by which different kinases and signaling pathways might directly control its binding to specific partners, including BAF (Hirano et al. 2005; Roberts et al. 2006; Schlosser et al. 2006; Luo et al. 2009). REGULATION OF SIGNALING AND PROLIFERATION CONTROL BY LAMIN-BINDING PROTEINS IN THE NUCLEAR INTERIOR

The largest LAP2 isoform, LAP2a, lacks a transmembrane domain, localizes to the nuclear interior, and specifically binds A-type lamins (Dechat et al. 2000; Vlcek et al. 2002; Dechat et al. 2004). In proliferating cells, LAP2a is essential and sufficient to target and retain a subset of lamins within the nuclear interior (Naetar et al. 2007; Naetar et al. 2008). Nucleoplasmic lamins are stable, but more mobile than peripheral lamins, suggesting interior lamins assemble

Lamin-binding Proteins

differently or bind to chromatin (Moir et al. 2000b). LAP2a and lamin A also bind directly to tumor suppressor Rb (Ozaki et al. 1994; Markiewicz et al. 2002; Pekovic et al. 2007), and LAP2a is involved in Rb-mediated repression of E2F/Rb target genes (Dorner et al. 2006) and Rb-dependent proliferation control in tissue progenitor stem cells (Naetar et al. 2008). The physiological relevance of intranuclear LAP2a-lamin complexes in Rb regulation is supported by studies showing destabilization and proteolytic degradation (Johnson et al. 2004; Nitta et al. 2006; Nitta et al. 2007) and impaired phosphorylation-dependent repressor activity control (Van Berlo et al. 2005) of Rb in cells lacking A-type lamins. “FAMILIAR” REGULATORY PROTEINS AND ENZYMES DISCOVERED TO BIND LAMINS

In addition to Rb, other known transcription factors and regulators have been discovered to bind A- and/or B-type lamins directly. This binding, whether transient or stable in vivo, suggests these proteins—and pathways they represent—either require or are influenced by lamins. These lamin-binding proteins include cyclin D3 (Mariappan et al. 2007), transcription factors cFos, Oct-1, SREBP1, MOK2, and tumor suppressor ING1 (Han et al. 2008), an enzyme named 12(S)-lipoxygenase (12[S] LOX) (Tang et al. 2000), and at least two kinases: protein kinase C a (PKCa) and JIL-1 kinase (Bao et al. 2005) (reviewed in Zastrow et al. 2004; Vlcek and Foisner 2007). Binding to lamin B1 sequesters transcription factor Oct-1 at the NE, away from promoters; in lamin B1-deficient cells, Oct-1-dependent genes that respond to oxidative stress and reactive oxygen species are misregulated (Malhas et al. 2009). Similarly, c-Fos, an early response transcription factor, is sequestered at the NE by A-type lamins (Ivorra et al. 2006) and during mitogen activated protein (MAP) kinase signaling, activated ERK1/2 interacts with lamin A and c-Fos at the NE to rapidly release c-Fos and facilitate cell proliferation (Gonzalez et al. 2008). Deregulation of this pathway in emerin and lamin A-deficient cells might up-regulate ERK-mediated signaling, a

known phenotype of certain lamin- or emerindeficient cells (Muchir et al. 2007; Muchir et al. 2009). Lamin A is also reported to affect Notch signaling, required to differentiate mesenchymal stem cells, by tethering SKIP, a coactivator of Notch-dependent target genes (Scaffidi and Misteli 2008). A better understanding of these interactions is sorely needed because it might begin to illuminate how and why nuclear architecture defects, for example in accelerated aging or other diseases linked to mutations in lamins and lamin-binding proteins, might disrupt signaling and gene regulation pathways. The nuclear import receptor, importin a, is also a lamin-binding protein (Adam et al. 2008). Importin a inhibits lamin assembly in vitro, suggesting it prevents newly synthesized lamins from assembling in the cytoplasm; this interaction is disrupted by Ran-GTP, found at high levels only within nuclei, releasing the assembly block once lamins are in the nucleus. PCNA, the Sliding Clamp “Ring” for DNA Replication, Binds Lamins

Lamins are also involved in DNA replication. Replication is a structural challenge, because replication complexes must traverse each huge chromosome, including regions tethered as heterochromatin to the NE. DNA replication complexes are anchored to nucleoskeletal structures (Hozak et al. 1993). During replication, the NE-anchored heterochromatin becomes disconnected, moves to replication complexes, and is then retethered at the NE (Li et al. 1998). DNA replication arrests at the elongation phase in nuclei subjected to lamina network disruption by “dominant –negative” lamin fragments (Spann et al. 1997; Moir et al. 2000a). In a breakthrough for this understudied area, PCNA ( proliferating cell nuclear antigen), which forms a sliding clamp “ring” ahead of the replication fork, was found to bind lamins directly (Shumaker et al. 2008). Further study of PCNA, the first but perhaps not the only lamin-binding component of DNA replication machinery, might shed light on how and why this fundamental process came to depend on the nucleoskeleton and lamins in metazoans.

K.L. Wilson and R. Foisner

LAMIN-BINDING PROTEINS AND HUMAN DISEASE

The diversity and physiological relevance of lamin-binding proteins is reflected in the growing number of human diseases linked to genes encoding lamins or lamin-binding proteins (Broers et al. 2006; Capell and Collins 2006; Worman and Bonne 2007). These diseases are known collectively as “laminopathies” or “envelopathies.” More than 12 overlapping or distinct syndromes including autosomal dominant Emery-Dreifuss muscular dystrophy (ADEDMD) are caused by mutations in the gene encoding A-type lamins (LMNA). In several cases, the same or clinically indistinguishable diseases can also be caused by mutations in one or more lamin-binding proteins. For example, EDMD is also caused by X-linked recessive loss of emerin (Bione et al. 1994), or dominant missense mutations in SYNE1 or SYNE2, encoding nesprin-1 and nesprin-2, respectively (Zhang et al. 2007; Puckelwartz et al. 2009). These similar pathologies suggest the disruption of a functional complex(es) that requires products of all four genes (Gotzmann and Foisner 2006). On the other hand, mutations in emerin can also cause other syndromes including Limb-Girdle muscular dystrophy, cardiomyopathy with conduction defects, or familial atrial fibrillation (Ben Yaou et al. 2007; Ura et al. 2007; Karst et al. 2008). “Classical” EDMD is characterized by progressive muscle wasting, contractures of tendons, and cardiomyopathy with ventricular conduction system defects (Bione et al. 1994; Manilal et al. 1996). Most X-linked EDMD patients are functionally null for emerin because of instability of the mutated protein or, more rarely, failed INM retention of mutant emerin proteins. In many AD-EDMD patients with LMNA mutations, emerin drifts into the ER and is functionally lost from the NE (Broers et al. 2006; Worman and Bonne 2007); however, with LMNA mutations, one must assume many additional INM or ONM proteins similarly mislocalize, if their retention at the NE requires A-type lamins. Gene-expression profiling of muscle biopsies from EDMD patients (Bakay et al. 2006) and emerin-down-regulated mouse cells (Melcon

et al. 2006) suggested shared defects in the expression of genes regulated by Rb1 and MyoD, which are required for muscle stem cell regulation and muscle differentiation (Gotzmann and Foisner 2006). A mutation in LAP2a that disrupts binding to lamin A is known to cause dilated cardiomyopathy (Taylor et al. 2005). Whether this mutation might also impair Rb activity, or reflect a novel role of LAP2a, is unknown. Additional disease mechanisms are suggested by the impaired organization and function of the neuromuscular junction in lamin-null mice and AD-EDMD patients (Mejat et al. 2009), and evidence that emerin is regulated by Her2 signaling (Tifft et al. 2009). “Signaling” Laminopathies Linked to MAN1 and LBR

Heterozygous loss-of-function mutations in MAN1 (LEMD3) cause osteopoikilosis, Buschke-Ollendorff syndrome, and melorheostosis, characterized by increased bone density (Hellemans et al. 2004). This pathology may result from impaired MAN1 regulation of TGF-b (Smad) signaling (discussed above), which is important for bone development. LBR, the INM-localized sterol reductase that also binds lamin B, is required for nuclei to change shape and reorganize chromatin in differentiating neutrophils (Hoffmann et al. 2002). LBR is proposed to associate with heterochromatic under-acetylated chromatin (Polioudaki et al. 2001; Makatsori et al. 2004) through binding to heterochromatin protein 1 (HP1) (Ye and Worman 1996). Heterozygous LBR mutations cause Pelger-Huet anomaly, a benign autosomal dominant syndrome characterized by abnormal nuclear shape and chromatin organization in blood granulocytes (Hoffmann et al. 2002). Greater loss of LBR protein correlates with more severe phenotypes, including developmental delay, epilepsy, and skeletal abnormalities (Hoffmann et al. 2007). Mutations that significantly reduce the sterol reductase activity of LBR are linked to autosomal recessive Greenberg’s skeletal dysplasia, characterized by lethal skeletal and visceral anomalies (Waterham et al. 2003).

Lamin-binding Proteins

Central Nervous System (CNS) Laminopathies

Brain pathologies with genetic links to lamins or lamin-binding proteins were noticeably missing, until recently. Surprisingly, the duplication of LMNB1, encoding lamin B1, causes autosomal dominant leukodystrophy, a slowly progressive disorder characterized by widespread demyelination of the CNS (Padiath et al. 2006; Brussino et al. 2009). Proper regulation of lamin B1 expression is crucial for oligodendrocyte development and myelination (Lin and Fu 2009). The lamin B1-binding proteins and pathways that underlie these functions in the brain are open questions. Primary dystonia is a CNS-based autosomal-dominant movement disorder caused by mutations in the AAAþATPase torsin A. This brain pathology was traced to the aberrantly tight binding of the mutated torsin A protein to the lumenal domain of LAP1 (Naismith et al. 2004; Goodchild and Dauer 2005), an INM protein that binds A- and B-type lamins. Wild-type torsin A is recruited to the NE by LAP1 when ATP is bound, whereas torsin A distributes throughout the ER lumen in the absence of ATP; the immobilization of mutant torsin A in the NE lumen correlates with cytoskeletal protein accumulation near the NE and disrupted cytoskeletal dynamics, neurite extension, and cell adhesion (Hewett et al. 2006). In addition to causing EDMD, other mutations in nesprin-1 (SYNE1) can cause autosomal-recessive cerebellar ataxia (Gros-Louis et al. 2007), characterized by impaired walking. Although this pathology might reflect impaired function of a brain-specific postsynaptic nesprin-1 isoform, parallel disruption of nuclear isoforms cannot be ruled out.

CONCLUDING REMARKS

Lamin-binding proteins are an emerging class of proteins that may explain how chromosomes are organized, how nuclei are assembled after mitosis or moved to new positions in the cell, and how transcription, replication, signaling, and many other activities are supported by lamin filaments and the nucleoskeleton. Further

exploration of this understudied area of cell biology may lead to improved and potentially therapeutic understanding of many human diseases. ACKNOWLEDGMENTS

We gratefully acknowledge grant support from the Austrian Science Research Fund (FWF P17871) and the EURO-Laminopathies research project of the European Commission (Contract LSHM-CT-2005-018690) to R.F. and National Institutes of Health Directors Bridge Award (2R56GM08646-13A1) to K.L.W. REFERENCES Adam SA, Sengupta K, Goldman RD. 2008. Regulation of nuclear lamin polymerization by importin a. J Biol Chem 283: 8462– 8468. Anderson DJ, Hetzer MW. 2008. Shaping the endoplasmic reticulum into the nuclear envelope. J Cell Sci 121: 137 –142. Bakay M, Wang Z, Melcon G, Schiltz L, Xuan J, Zhao P, Sartorelli V, Seo J, Pegoraro E, Angelini C, et al. 2006. Nuclear envelope dystrophies show a transcriptional fingerprint suggesting disruption of Rb-MyoD pathways in muscle regeneration. Brain 129: 996 –1013. Bao X, Zhang W, Krencik R, Deng H, Wang Y, Girton J, Johansen J, Johansen KM. 2005. The JIL-1 kinase interacts with lamin Dm0 and regulates nuclear lamina morphology of Drosophila nurse cells. J Cell Sci 118: 5079–5087. Barton RM, Worman HJ. 1999. Prenylated prelamin A interacts with Narf, a novel nuclear protein. J Biol Chem 274: 30008– 30018. Ben Yaou R, Toutain A, Arimura T, Demay L, Massart C, Peccate C, Muchir A, Llense S, Deburgrave N, Leturcq F, et al. 2007. Multitissular involvement in a family with LMNA and EMD mutations: Role of digenic mechanism? Neurology 68: 1883–1894. Bengtsson L, Otto H. 2008. LUMA interacts with emerin and influences its distribution at the inner nuclear membrane. J Cell Sci 121: 536 –548. Bengtsson L, Wilson KL. 2004. Multiple and surprising new functions for emerin, a nuclear membrane protein. Curr Opin Cell Biol 16: 73–79. Berger R, Theodor L, Shoham J, Gokkel E, Brok-Simoni F, Avraham KB, Copeland NG, Jenkins NA, Rechavi G, Simon AJ. 1996. The characterization and localization of the mouse thymopoietin/lamina- associated polypeptide 2 gene and its alternatively spliced products. Genome Res 6: 361– 370. Bione S, Maestrini E, Rivella S, Mancini M, Regis S, Romeo G, Toniolo D. 1994. Identification of a novel X-linked gene responsible for Emery-Dreifuss muscular dystrophy. Nature Genet 8: 323–327.

K.L. Wilson and R. Foisner

Brachner A, Reipert S, Foisner R, Gotzmann J. 2005. LEM2 is a novel MAN1-related inner nuclear membrane protein associated with A-type lamins. J Cell Sci 118: 5797– 5810. Broers JL, Ramaekers FC, Bonne G, Yaou RB, Hutchison CJ. 2006. Nuclear lamins: Laminopathies and their role in premature ageing. Physiol Rev 86: 967 –1008. Brussino A, Vaula G, Cagnoli C, Mauro A, Pradotto L, Daniele D, Di Gregorio E, Barberis M, Arduino C, Squadrone S, et al. 2009. A novel family with Lamin B1 duplication associated with adult-onset leucoencephalopathy. J Neurol Neurosurg Psychiatry 80: 237–240. Cai M, Huang Y, Ghirlando R, Wilson KL, Craigie R, Clore GM. 2001. Solution structure of the constant region of nuclear envelope protein LAP2 reveals two LEM-domain structures: One binds BAF and the other binds DNA. EMBO J 20: 4399– 4407. Cai M, Huang Y, Suh JY, Louis JM, Ghirlando R, Craigie R, Clore GM. 2007. Solution NMR structure of the barrierto-autointegration factor-Emerin complex. J Biol Chem 282: 14525– 14535. Capell BC, Collins FS. 2006. Human laminopathies: Nuclei gone genetically awry. Nat Rev Genet 7: 940– 952. Caputo S, Couprie J, Duband-Goulet I, Konde E, Lin F, Braud S, Gondry M, Gilquin B, Worman HJ, Zinn-Justin S. 2006. The carboxyl-terminal nucleoplasmic region of MAN1 exhibits a DNA binding winged helix domain. J Biol Chem 281: 18208–18215. Chen IH, Huber M, Guan T, Bubeck A, Gerace L. 2006. Nuclear envelope transmembrane proteins (NETs) that are up-regulated during myogenesis. BMC Cell Biol 7: 38. Chikashige Y, Haraguchi T, Hiraoka Y. 2007. Another way to move chromosomes. Chromosoma 116: 497–505. Clements L, Manilal S, Love DR, Morris GE. 2000. Direct interaction between emerin and lamin A. Biochem Biophys Res Commun 267: 709 –714. Cohen TV, Kosti O, Stewart CL. 2007. The nuclear envelope protein MAN1 regulates TGFb signaling and vasculogenesis in the embryonic yolk sac. Development 134: 1385– 1395. Crisp M, Liu Q, Roux K, Rattner JB, Shanahan C, Burke B, Stahl PD, Hodzic D. 2006. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J Cell Biol 172: 41–53. Dechat T, Gajewski A, Korbei B, Gerlich D, Daigle N, Haraguchi T, Furukawa K, Ellenberg J, Foisner R. 2004. LAP2a and BAF transiently localize to telomeres and specific regions on chromatin during nuclear assembly. J Cell Sci 117: 6117–6128. Dechat T, Korbei B, Vaughan OA, Vlcek S, Hutchison CJ, Foisner R. 2000. Lamina-associated polypeptide 2a binds intranuclear A-type lamins. J Cell Sci 113: 3473–3484. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832–853. Ding X, Xu R, Yu J, Xu T, Zhuang Y, Han M. 2007. SUN1 is required for telomere attachment to nuclear envelope and gametogenesis in mice. Dev Cell 12: 863– 872.

Dorner D, Vlcek S, Foeger N, Gajewski A, Makolm C, Gotzmann J, Hutchison CJ, Foisner R. 2006. Lamina-associated polypeptide 2a regulates cell cycle progression and differentiation via the retinoblastoma-E2F pathway. J Cell Biol 173: 83–93. Foisner R, Gerace L. 1993. Integral membrane proteins of the nuclear envelope interact with lamins and chromosomes, and binding is modulated by mitotic phosphorylation. Cell 73: 1267–1279. Furukawa K. 1999. LAP2 binding protein 1 (L2BP1/BAF) is a candidate mediator of LAP2- chromatin interaction. J Cell Sci 112: 2485–2492. Furukawa K, Fritze CE, Gerace L. 1998. The major nuclear envelope targeting domain of LAP2 coincides with its lamin binding region but is distinct from its chromatin interaction domain. J Biol Chem 273: 4213–4219. Furukawa K, Pante N, Aebi U, Gerace L. 1995. Cloning of a cDNA for lamina-associated polypeptide 2 (LAP2) and identification of regions that specify targeting to the nuclear envelope. EMBO J 14: 1626– 1636. Goldberg M, Lu H, Stuurman N, Ashery-Padan R, Weiss AM, Yu J, Bhattacharyya D, Fisher PA, Gruenbaum Y, Wolfner MF. 1998. Interactions among Drosophila nuclear envelope proteins lamin, otefin, and YA. Mol Cell Biol 18: 4315– 4323. Gonzalez JM, Navarro-Puche A, Casar B, Crespo P, Andres V. 2008. Fast regulation of AP-1 activity through interaction of lamin A/C, ERK1/2, and c-Fos at the nuclear envelope. J Cell Biol 183: 653– 666. Goodchild RE, Dauer WT. 2005. The AAAþ protein torsinA interacts with a conserved domain present in LAP1 and a novel ER protein. J Cell Biol 168: 855– 862. Gotzmann J, Foisner R. 2006. A-type lamin complexes and regenerative potential: A step towards understanding laminopathic diseases? Histochem Cell Biol 125: 33–41. Grady RM, Starr DA, Ackerman GL, Sanes JR, Han M. 2005. Syne proteins anchor muscle nuclei at the neuromuscular junction. Proc Natl Acad Sci 102: 4359–4364. Gros-Louis F, Dupre N, Dion P, Fox MA, Laurent S, Verreault S, Sanes JR, Bouchard JP, Rouleau GA. 2007. Mutations in SYNE1 lead to a newly discovered form of autosomal recessive cerebellar ataxia. Nat Genet 39: 80– 85. Gruenbaum Y, Lee KK, Liu J, Cohen M, Wilson KL. 2002. The expression, lamin-dependent localization and RNAi depletion phenotype for emerin in C. elegans. J Cell Sci 115: 923 –929. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol 6: 21–31. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948–951. Guttinger S, Laurell E, Kutay U. 2009. Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat Rev Mol Cell Biol 10: 178 –191. Hakelien AM, Delbarre E, Gaustad KG, Buendia B, Collas P. 2008. Expression of the myodystrophic R453W mutation

Lamin-binding Proteins

of lamin A in C2C12 myoblasts causes promoter-specific and global epigenetic defects. Exp Cell Res 314: 1869– 1880. Han X, Feng X, Rattner JB, Smith H, Bose P, Suzuki K, Soliman MA, Scott MS, Burke BE, Riabowol K. 2008. Tethering by lamin A stabilizes and targets the ING1 tumour suppressor. Nat Cell Biol 10: 1333–1340. Haque F, Lloyd DJ, Smallwood DT, Dent CL, Shanahan CM, Fry AM, Trembath RC, Shackleton S. 2006. SUN1 interacts with nuclear lamin A and cytoplasmic nesprins to provide a physical connection between the nuclear lamina and the cytoskeleton. Mol Cell Biol 26: 3738–3751. Haraguchi T, Holaska JM, Yamane M, Koujin T, Hashiguchi N, Mori C, Wilson KL, Hiraoka Y. 2004. Emerin binding to Btf, a death-promoting transcriptional repressor, is disrupted by a missense mutation that causes EmeryDreifuss muscular dystrophy. Eur J Biochem 271: 1035– 1045. Haraguchi T, Koujin T, Segura-Totten M, Lee KK, Matsuoka Y, Yoneda Y, Wilson KL, Hiraoka Y. 2001. BAF is required for emerin assembly into the reforming nuclear envelope. J Cell Sci 114: 4575–4585. Harris CA, Andryuk PJ, Cline S, Chan HK, Natarajan A, Siekierka JJ, Goldstein G. 1994. Three distinct human thymopoietins are derived from alternatively spliced mRNAs. Proc Natl Acad Sci 91: 6283–6287. Hellemans J, Preobrazhenska O, Willaert A, Debeer P, Verdonk PC, Costa T, Janssens K, Menten B, Van Roy N, Vermeulen SJ, et al. 2004. Loss-of-function mutations in LEMD3 result in osteopoikilosis, Buschke-Ollendorff syndrome and melorheostosis. Nat Genet 36: 1213– 1218. Hewett JW, Zeng J, Niland BP, Bragg DC, Breakefield XO. 2006. Dystonia-causing mutant torsinA inhibits cell adhesion and neurite extension through interference with cytoskeletal dynamics. Neurobiol Dis 22: 98–111. Hirano Y, Segawa M, Ouchi FS, Yamakawa Y, Furukawa K, Takeyasu K, Horigome T. 2005. Dissociation of emerin from barrier-to-autointegration factor is regulated through mitotic phosphorylation of emerin in a Xenopus egg cell-free system. J Biol Chem 280: 39925–39933. Hodzic DM, Yeater DB, Bengtsson L, Otto H, Stahl PD. 2004. Sun2 is a novel mammalian inner nuclear membrane protein. J Biol Chem 279: 25805– 25812. Hoffmann K, Dreger CK, Olins AL, Olins DE, Shultz LD, Lucke B, Karl H, Kaps R, Muller D, Vaya A, et al. 2002. Mutations in the gene encoding the lamin B receptor produce an altered nuclear morphology in granulocytes (Pelger-Huet anomaly). Nat Genet 31: 410– 414. Hoffmann K, Sperling K, Olins AL, Olins DE. 2007. The granulocyte nucleus and lamin B receptor: Avoiding the ovoid. Chromosoma 116: 227– 235. Holaska JM, Wilson KL. 2006. Multiple roles for emerin: Implications for Emery-Dreifuss muscular dystrophy. Anat Rec A Discov Mol Cell Evol Biol 288: 676– 680. Holaska JM, Wilson KL. 2007. An emerin “proteome”: Purification of distinct emerin-containing complexes from HeLa cells suggests molecular basis for diverse roles including gene regulation, mRNA splicing, signaling, mechanosensing, and nuclear architecture. Biochemistry 46: 8897–8908. Holaska JM, Kowalski AK, Wilson KL. 2004. Emerin caps the pointed end of actin filaments: Evidence for an actin

cortical network at the nuclear inner membrane. PLoS Biol 2: E231. Holaska JM, Rais-Bahrami S, Wilson KL. 2006. Lmo7 is an emerin-binding protein that regulates the transcription of emerin and many other muscle-relevant genes. Hum Mol Genet 15: 3459– 3472. Holaska JM, Lee KK, Kowalski AK, Wilson KL. 2003. Transcriptional repressor germ cell-less (GCL) and barrier to autointegration factor (BAF) compete for binding to emerin in vitro. J Biol Chem 278: 6969–6975. Hozak P, Hassan AB, Jackson DA, Cook PR. 1993. Visualization of replication factories attached to nucleoskeleton. Cell 73: 361–373. Ivorra C, Kubicek M, Gonzalez JM, Sanz-Gonzalez SM, Alvarez-Barrientos A, O’Connor JE, Burke B, Andres V. 2006. A mechanism of AP-1 suppression through interaction of c-Fos with lamin A/C. Genes Dev 20: 307–320. Jiang X, Xia L, Chen D, Yang Y, Huang H, Yang L, Zhao Q, Shen L, Wang J. 2008. Otefin, a nuclear membrane protein, determines the fate of germline stem cells in Drosophila via interaction with Smad complexes. Dev Cell 14: 494 –506. Johnson BR, Nitta RT, Frock RL, Mounkes L, Barbie DA, Stewart CL, Harlow E, Kennedy BK. 2004. A-type lamins regulate retinoblastoma protein function by promoting subnuclear localization and preventing proteasomal degradation. Proc Natl Acad Sci 101: 9677– 9682. Karst ML, Herron KJ, Olson TM. 2008. X-linked nonsyndromic sinus node dysfunction and atrial fibrillation caused by emerin mutation. J Cardiovasc Electrophysiol 19: 510– 515. Kracklauer MP, Banks SM, Xie X, Wu Y, Fischer JA. 2007. Drosophila klaroid encodes a SUN domain protein required for Klarsicht localization to the nuclear envelope and nuclear migration in the eye. Fly (Austin) 1: 75–85. Laguri C, Gilquin B, Wolff N, Romi-Lebrun R, Courchay K, Callebaut I, Worman HJ, Zinn-Justin S. 2001. Structural characterization of the LEM motif common to three human inner nuclear membrane proteins. Structure 9: 503 –511. Lammerding J, Hsiao J, Schulze PC, Kozlov S, Stewart CL, Lee RT. 2005. Abnormal nuclear shape and impaired mechanotransduction in emerin-deficient cells. J Cell Biol 170: 781–791. Lee KK, Wilson KL. 2004. All in the family: Evidence for four new LEM-domain proteins Lem2 (NET-25), Lem3, Lem4 and Lem5 in the human genome. Symp Soc Exp Biol: 329–339. Lee KK, Gruenbaum Y, Spann P, Liu J, Wilson KL. 2000. C. elegans nuclear envelope proteins emerin, MAN1, lamin, and nucleoporins reveal unique timing of nuclear envelope breakdown during mitosis. Mol Biol Cell 11: 3089–3099. Lee KK, Haraguchi T, Lee RS, Koujin T, Hiraoka Y, Wilson KL. 2001. Distinct functional domains in emerin bind lamin A and DNA-bridging protein BAF. J Cell Sci 114: 4567–4573. Lee KK, Starr D, Cohen M, Liu J, Han M, Wilson KL, Gruenbaum Y. 2002. Lamin-dependent localization of UNC-84, a protein required for nuclear migration in Caenorhabditis elegans. Mol Biol Cell 13: 892– 901.

K.L. Wilson and R. Foisner

Li G, Sudlow G, Belmont AS. 1998. Interphase cell cycle dynamics of a late-replicating, heterochromatic homogeneously staining region: Precise choreography of condensation/decondensation and nuclear positioning. J Cell Biol 140: 975–989. Lin ST, Fu YH. 2009. miR-23 regulation of lamin B1 is crucial for oligodendrocyte development and myelination. Dis Model Mech 2: 178 –188. Lin F, Blake DL, Callebaut I, Skerjanc IS, Holmer L, McBurney MW, Paulin-Levasseur M, Worman HJ. 2000. MAN1, an inner nuclear membrane protein that shares the LEM domain with lamina-associated polypeptide 2 and emerin. J Biol Chem 275: 4840– 4847. Lin F, Morrison JM, Wu W, Worman HJ. 2005. MAN1, an integral protein of the inner nuclear membrane, binds Smad2 and Smad3 and antagonizes transforming growth factor-b signaling. Hum Mol Genet 14: 437 –445. Liu J, Lee KK, Segura-Totten M, Neufeld E, Wilson KL, Gruenbaum Y. 2003. MAN1 and emerin have overlapping function(s) essential for chromosome segregation and cell division in Caenorhabditis elegans. Proc Natl Acad Sci 100: 4598–4603. Liu Q, Pante N, Misteli T, Elsagga M, Crisp M, Hodzic D, Burke B, Roux KJ. 2007. Functional association of Sun1 with nuclear pore complexes. J Cell Biol 178: 785–798. Luderus ME, de Graaf A, Mattia E, den Blaauwen JL, Grande MA, de Jong L, van Driel R. 1992. Binding of matrix attachment regions to lamin B1. Cell 70: 949 –959. Luke Y, Zaim H, Karakesisoglou I, Jaeger VM, Sellin L, Lu W, Schneider M, Neumann S, Beijer A, Munck M, et al. 2008. Nesprin-2 Giant (NUANCE) maintains nuclear envelope architecture and composition in skin. J Cell Sci 121: 1887–1898. Luo Y, Yang C, Jin C, Xie R, Wang F, McKeehan WL. 2009. Novel phosphotyrosine targets of FGFR2IIIb signaling. Cell Signal 21: 1370– 1378. Makatsori D, Kourmouli N, Polioudaki H, Shultz LD, McLean K, Theodoropoulos PA, Singh PB, Georgatos SD. 2004. The inner nuclear membrane protein lamin B receptor forms distinct microdomains and links epigenetically marked chromatin to the nuclear envelope. J Biol Chem 279: 25567–25573. Malhas AN, Lee CF, Vaux DJ. 2009. Lamin B1 controls oxidative stress responses via Oct-1. J Cell Biol 184: 45–55. Malone CJ, Fixsen WD, Horvitz HR, Han M. 1999. UNC-84 localizes to the nuclear envelope and is required for nuclear migration and anchoring during C. elegans development. Development 126: 3171– 3181. Malone CJ, Misner L, Le Bot N, Tsai MC, Campbell JM, Ahringer J, White JG. 2003. The C. elegans hook protein, ZYG-12, mediates the essential attachment between the centrosome and nucleus. Cell 115: 825 –836. Manilal S, Nguyen TM, Sewry CA, Morris GE. 1996. The Emery-Dreifuss muscular dystrophy protein, emerin, is a nuclear membrane protein. Hum Mol Genet 5: 801– 808. Mansharamani M, Wilson KL. 2005. Direct binding of nuclear membrane protein MAN1 to emerin in vitro and two modes of binding to barrier-to-autointegration factor. J Biol Chem 280: 13863–13870.

Margalit A, Brachner A, Gotzmann J, Foisner R, Gruenbaum Y. 2007. Barrier-to-autointegration factor—a BAFfling little protein. Trends Cell Biol 17: 202–208. Margalit A, Segura-Totten M, Gruenbaum Y, Wilson KL. 2005a. Barrier-to-autointegration factor is required to segregate and enclose chromosomes within the nuclear envelope and assemble the nuclear lamina. Proc Natl Acad Sci 102: 3290– 3295. Margalit A, Vlcek S, Gruenbaum Y, Foisner R. 2005b. Breaking and making of the nuclear envelope. J Cell Biochem 95: 454– 465. Mariappan I, Gurung R, Thanumalayan S, Parnaik VK. 2007. Identification of cyclin D3 as a new interaction partner of lamin A/C. Biochem Biophys Res Commun 355: 981–985. Markiewicz E, Dechat T, Foisner R, Quinlan RA, Hutchison CJ. 2002. Lamin A/C binding protein LAP2a is required for nuclear anchorage of retinoblastoma protein. Mol Biol Cell 13: 4401–4413. Markiewicz E, Tilgner K, Barker N, van de Wetering M, Clevers H, Dorobek M, Hausmanowa-Petrusewicz I, Ramaekers FC, Broers JL, Blankesteijn WM, et al. 2006. The inner nuclear membrane protein emerin regulates b-catenin activity by restricting its accumulation in the nucleus. EMBO J 25: 3275–3285. Martin L, Crimaudo C, Gerace L. 1995. cDNA cloning and characterization of lamina-associated polypeptide 1C (LAP1C), an integral protein of the inner nuclear membrane. J Biol Chem 270: 8822–8828. Martins S, Eikvar S, Furukawa K, Collas P. 2003. HA95 and LAP2 b mediate a novel chromatin-nuclear envelope interaction implicated in initiation of DNA replication. J Cell Biol 160: 177– 188. Mattout A, Goldberg M, Tzur Y, Margalit A, Gruenbaum Y. 2007. Specific and conserved sequences in D. melanogaster and C. elegans lamins and histone H2A mediate the attachment of lamins to chromosomes. J Cell Sci 120: 77– 85. McGee MD, Rillo R, Anderson AS, Starr DA. 2006. UNC-83 is a KASH protein required for nuclear migration and is recruited to the outer nuclear membrane by a physical interaction with the SUN protein UNC-84. Mol Biol Cell 17: 1790– 1801. Mejat A, Decostre V, Li J, Renou L, Kesari A, Hantai D, Stewart CL, Xiao X, Hoffman E, Bonne G, et al. 2009. Lamin A/ C-mediated neuromuscular junction defects in EmeryDreifuss muscular dystrophy. J Cell Biol 184: 31– 44. Melcon G, Kozlov S, Cutler DA, Sullivan T, Hernandez L, Zhao P, Mitchell S, Nader G, Bakay M, Rottman JN, et al. 2006. Loss of emerin at the nuclear envelope disrupts the Rb1/E2F and MyoD pathways during muscle regeneration. Hum Mol Genet 15: 637– 651. Mislow JM, Holaska JM, Kim MS, Lee KK, Segura-Totten M, Wilson KL, McNally EM. 2002a. Nesprin-1a self-associates and binds directly to emerin and lamin A in vitro. FEBS Lett 525: 135– 140. Mislow JM, Kim MS, Davis DB, McNally EM. 2002b. Myne-1, a spectrin repeat transmembrane protein of the myocyte inner nuclear membrane, interacts with lamin A/C. J Cell Sci 115: 61–70. Moir RD, Spann TP, Herrmann H, Goldman RD. 2000a. Disruption of nuclear lamin organization blocks the

Lamin-binding Proteins

elongation phase of DNA replication. J Cell Biol 149: 1179– 1192. Moir RD, Yoon M, Khuon S, Goldman RD. 2000b. Nuclear lamins A and B1: different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151: 1155–1168. Muchir A, Pavlidis P, Bonne G, Hayashi YK, Worman HJ. 2007. Activation of MAPK in hearts of EMD null mice: Similarities between mouse models of X-linked and autosomal dominant Emery Dreifuss muscular dystrophy. Hum Mol Genet 16: 1884–1895. Muchir A, Shan J, Bonne G, Lehnart SE, Worman HJ. 2009. Inhibition of extracellular signal-regulated kinase signaling to prevent cardiomyopathy caused by mutation in the gene encoding A-type lamins. Hum Mol Genet 18: 241–247. Naetar N, Hutter S, Dorner D, Dechat T, Korbei B, Gotzmann J, Beug H, Foisner R. 2007. LAP2a-binding protein LINT-25 is a novel chromatin-associated protein involved in cell cycle exit. J Cell Sci 120: 737– 747. Naetar N, Korbei B, Kozlov S, Kerenyi MA, Dorner D, Kral R, Gotic I, Fuchs P, Cohen TV, Bittner R, et al. 2008. Loss of nucleoplasmic LAP2a-lamin A complexes causes erythroid and epidermal progenitor hyperproliferation. Nat Cell Biol 10: 1341– 1348. Naismith TV, Heuser JE, Breakefield XO, Hanson PI. 2004. TorsinA in the nuclear envelope. Proc Natl Acad Sci 101: 7612–7617. Nery FC, Zeng J, Niland BP, Hewett J, Farley J, Irimia D, Li Y, Wiche G, Sonnenberg A, Breakefield XO. 2008. TorsinA binds the KASH domain of nesprins and participates in linkage between nuclear envelope and cytoskeleton. J Cell Sci 121: 3476– 3486. Nili E, Cojocaru GS, Kalma Y, Ginsberg D, Copeland NG, Gilbert DJ, Jenkins NA, Berger R, Shaklai S, Amariglio N, et al. 2001. Nuclear membrane protein LAP2b mediates transcriptional repression alone and together with its binding partner GCL (germ-cell-less). J Cell Sci 114: 3297– 3307. Nitta RT, Smith CL, Kennedy BK. 2007. Evidence that proteasome-dependent degradation of the retinoblastoma protein in cells lacking A-type lamins occurs independently of gankyrin and MDM2. PLoS ONE 2: e963. Nitta RT, Jameson SA, Kudlow BA, Conlan LA, Kennedy BK. 2006. Stabilization of the retinoblastoma protein by A-type nuclear lamins is required for INK4A-mediated cell cycle arrest. Mol Cell Biol 26: 5360– 5372. Osada S, Ohmori SY, Taira M. 2003. XMAN1, an inner nuclear membrane protein, antagonizes BMP signaling by interacting with Smad1 in Xenopus embryos. Development 130: 1783– 1794. Ottaviani A, Rival-Gervier S, Boussouar A, Foerster AM, Rondier D, Sacconi S, Desnuelle C, Gilson E, Magdinier F. 2009. The D4Z4 macrosatellite repeat acts as a CTCF and A-type lamins-dependent insulator in facioscapulo-humeral dystrophy. PLoS Genet 5: e1000394. Ozaki T, Saijo M, Murakami K, Enomoto H, Taya Y, Sakiyama S. 1994. Complex formation between lamin A and the retinoblastoma gene product: Identification of the domain on lamin A required for its interaction. Oncogene 9: 2649– 2653.

Ozawa R, Hayashi YK, Ogawa M, Kurokawa R, Matsumoto H, Noguchi S, Nonaka I, Nishino I. 2006. Emerinlacking mice show minimal motor and cardiac dysfunctions with nuclear-associated vacuoles. Am J Pathol 168: 907–917. Padiath QS, Saigoh K, Schiffmann R, Asahara H, Yamada T, Koeppen A, Hogan K, Ptacek LJ, Fu YH. 2006. Lamin B1 duplications cause autosomal dominant leukodystrophy. Nat Genet 38: 1114–1123. Padmakumar VC, Abraham S, Braune S, Noegel AA, Tunggal B, Karakesisoglou I, Korenbaum E. 2004. Enaptin, a giant actin-binding protein, is an element of the nuclear membrane and the actin cytoskeleton. Exp Cell Res 295: 330 –339. Padmakumar VC, Libotte T, Lu W, Zaim H, Abraham S, Noegel AA, Gotzmann J, Foisner R, Karakesisoglou I. 2005. The inner nuclear membrane protein Sun1 mediates the anchorage of Nesprin-2 to the nuclear envelope. J Cell Sci 118: 3419– 3430. Pan D, Estevez-Salmeron LD, Stroschein SL, Zhu X, He J, Zhou S, Luo K. 2005. The integral inner nuclear membrane protein MAN1 physically interacts with the R-Smad proteins to repress signaling by the transforming growth factor-b superfamily of cytokines. J Biol Chem 280: 15992–16001. Park YE, Hayashi YK, Bonne G, Arimura T, Noguchi S, Nonaka I, Nishino I. 2009a. Autophagic degradation of nuclear components in mammalian cells. Autophagy 5: 795 –804. Park YE, Hayashi YK, Goto K, Komaki H, Hayashi Y, Inuzuka T, Noguchi S, Nonaka I, Nishino I. 2009b. Nuclear changes in skeletal muscle extend to satellite cells in autosomal dominant Emery-Dreifuss muscular dystrophy/ limb-girdle muscular dystrophy 1B. Neuromuscul Disord 19: 29–36. Patterson K, Molofsky AB, Robinson C, Acosta S, Cater C, Fischer JA. 2004. The functions of Klarsicht and nuclear lamin in developmentally regulated nuclear migrations of photoreceptor cells in the Drosophila eye. Mol Biol Cell 15: 600–610. Pekovic V, Harborth J, Broers JL, Ramaekers FC, van Engelen B, Lammens M, von Zglinicki T, Foisner R, Hutchison C, Markiewicz E. 2007. Nucleoplasmic LAP2a-lamin A complexes are required to maintain a proliferative state in human fibroblasts. J Cell Biol 176: 163 –172. Penkner A, Tang L, Novatchkova M, Ladurner M, Fridkin A, Gruenbaum Y, Schweizer D, Loidl J, Jantsch V. 2007. The nuclear envelope protein Matefin/SUN-1 is required for homologous pairing in C. elegans meiosis. Dev Cell 12: 873 –885. Pickersgill H, Kalverda B, de Wit E, Talhout W, Fornerod M, van Steensel B. 2006. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat Genet 38: 1005–1014. Polioudaki H, Kourmouli N, Drosou V, Bakou A, Theodoropoulos PA, Singh PB, Giannakouros T, Georgatos SD. 2001. Histones H3/H4 form a tight complex with the inner nuclear membrane protein LBR and heterochromatin protein 1. EMBO Rep 2: 920 –925. Puckelwartz MJ, Kessler E, Zhang Y, Hodzic D, Randles KN, Morris G, Earley JU, Hadhazy M, Holaska JM, Mewborn

K.L. Wilson and R. Foisner

SK, et al. 2009. Disruption of nesprin-1 produces an Emery Dreifuss muscular dystrophy-like phenotype in mice. Hum Mol Genet 18: 607 –620. Raju GP, Dimova N, Klein PS, Huang HC. 2003. SANE, a novel LEM domain protein, regulates bone morphogenetic protein signaling through interaction with Smad1. J Biol Chem 278: 428–437. Roberts RC, Sutherland-Smith AJ, Wheeler MA, Jensen ON, Emerson LJ, Spiliotis II, Tate CG, Kendrick-Jones J, Ellis JA. 2006. The Emery-Dreifuss muscular dystrophy associated-protein emerin is phosphorylated on serine 49 by protein kinase A. Febs J 273: 4562–4575. Roux KJ, Crisp ML, Liu Q, Kim D, Kozlov S, Stewart CL, Burke B. 2009. Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc Natl Acad Sci 106: 2194–2199. Sabatelli P, Lattanzi G, Ognibene A, Columbaro M, Capanni C, Merlini L, Maraldi NM, Squarzoni S. 2001. Nuclear alterations in autosomal-dominant Emery-Dreifuss muscular dystrophy. Muscle Nerve 24: 826–829. Sakaki M, Koike H, Takahashi N, Sasagawa N, Tomioka S, Arahata K, Ishiura S. 2001. Interaction between emerin and nuclear lamins. J. Biochem 129: 321 –327. Salpingidou G, Smertenko A, Hausmanowa-Petrucewicz I, Hussey PJ, Hutchison CJ. 2007. A novel role for the nuclear membrane protein emerin in association of the centrosome to the outer nuclear membrane. J Cell Biol 178: 897– 904. Scaffidi P, Misteli T. 2006. Lamin A-dependent nuclear defects in human aging. Science 312: 1059– 1063. Scaffidi P, Misteli T. 2008. Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10: 452–459. Schirmer EC, Foisner R. 2007. Proteins that associate with lamins: Many faces, many functions. Exp Cell Res 313: 2167– 2179. Schirmer EC, Gerace L. 2005. The nuclear membrane proteome: Extending the envelope. Trends Biochem Sci 30: 551–558. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2003. Nuclear membrane proteins with potential disease links found by subtractive proteomics. Science 301: 1380– 1382. Schirmer EC, Florens L, Guan T, Yates JR 3rd, Gerace L. 2005. Identification of novel integral membrane proteins of the nuclear envelope with potential disease links using subtractive proteomics. Novartis Found. Symp 264: 63–76; discussion 76– 80, 227– 230. Schlosser A, Amanchy R, Otto H. 2006. Identification of tyrosine-phosphorylation sites in the nuclear membrane protein emerin. FEBS J 273: 3204– 3215. Schmitt J, Benavente R, Hodzic D, Hoog C, Stewart CL, Alsheimer M. 2007. Transmembrane protein Sun2 is involved in tethering mammalian meiotic telomeres to the nuclear envelope. Proc Natl Acad Sci 104: 7426– 7431. Senior A, Gerace L. 1988. Integral membrane proteins specific to the inner nuclear membrane and associated with the nuclear lamina. J Cell Biol 107: 2029– 2036. Shimi T, Koujin T, Segura-Totten M, Wilson KL, Haraguchi T, Hiraoka Y. 2004. Dynamic interaction between BAF

and emerin revealed by FRAP, FLIP, and FRET analyses in living HeLa cells. J Struct Biol 147: 31–41. Shoeman RL, Traub P. 1990. The in vitro DNA-binding properties of purified nuclear lamin proteins and vimentin. J Biol Chem 265: 9055–9061. Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, et al. 2006. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci 103: 8703– 8708. Shumaker DK, Lee KK, Tanhehco YC, Craigie R, Wilson KL. 2001. LAP2 binds to BAF.DNA complexes: requirement for the LEM domain and modulation by variable regions. EMBO J 20: 1754–1764. Shumaker DK, Solimando L, Sengupta K, Shimi T, Adam SA, Grunwald A, Strelkov SV, Aebi U, Cardoso MC, Goldman RD. 2008. The highly conserved nuclear lamin Ig-fold binds to PCNA: Its role in DNA replication. J Cell Biol 181: 269– 280. Somech R, Shaklai S, Geller O, Amariglio N, Simon AJ, Rechavi G, Gal-Yam EN. 2005. The nuclear-envelope protein and transcriptional repressor LAP2b interacts with HDAC3 at the nuclear periphery, and induces histone H4 deacetylation. J Cell Sci 118: 4017–4025. Spann TP, Moir RD, Goldman AE, Stick R, Goldman RD. 1997. Disruption of nuclear lamin organization alters the distribution of replication factors and inhibits DNA synthesis. J Cell Biol 136: 1201– 1212. Starr DA. 2009. A nuclear-envelope bridge positions nuclei and moves chromosomes. J Cell Sci 122: 577–586. Starr DA, Fischer JA. 2005. KASH ’n Karry: The KASH domain family of cargo-specific cytoskeletal adaptor proteins. Bioessays 27: 1136– 1146. Starr DA, Han M. 2002. Role of ANC-1 in tethering nuclei to the actin cytoskeleton. Science 298: 406–409. Starr DA, Hermann GJ, Malone CJ, Fixsen W, Priess JR, Horvitz HR, Han M. 2001. unc-83 encodes a novel component of the nuclear envelope and is essential for proper nuclear migration. Development 128: 5039–5050. Stewart CL, Roux KJ, Burke B. 2007. Blurring the boundary: The nuclear envelope extends its reach. Science 318: 1408–1412. Stewart-Hutchinson PJ, Hale CM, Wirtz D, Hodzic D. 2008. Structural requirements for the assembly of LINC complexes and their function in cellular mechanical stiffness. Exp Cell Res 314: 1892–1905. Stierle V, Couprie J, Ostlund C, Krimm I, Zinn-Justin S, Hossenlopp P, Worman HJ, Courvalin JC, DubandGoulet I. 2003. The carboxyl-terminal region common to lamins A and C contains a DNA binding domain. Biochemistry 42: 4819– 4828. Sullivan T, Escalante-Alcalde D, Bhatt H, Anver M, Bhat N, Nagashima K, Stewart CL, Burke B. 1999. Loss of A-type lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J Cell Biol 147: 913 –920. Tang K, Finley RL Jr, Nie D, Honn KV. 2000. Identification of 12-lipoxygenase interaction with cellular proteins by yeast two-hybrid screening. Biochemistry 39: 3185–3191.

Lamin-binding Proteins

Taniura H, Glass C, Gerace L. 1995. A chromatin binding site in the tail domain of nuclear lamins that interacts with core histones. J Cell Biol 131: 33–44. Taylor MR, Slavov D, Gajewski A, Vlcek S, Ku L, Fain PR, Carniel E, Di Lenarda A, Sinagra G, Boucek MM, et al. 2005. Thymopoietin (lamina-associated polypeptide 2) gene mutation associated with dilated cardiomyopathy. Hum Mutat 26: 566 –574. Tifft KE, Bradbury KA, Wilson KL. 2009. Tyrosine phosphorylation of nuclear-membrane protein emerin by Src, Abl and other kinases. J Cell Sci 122: 3780–3790. Tilgner K, Wojciechowicz K, Jahoda C, Hutchison C, Markiewicz E. 2009. Dynamic complexes of A-type lamins and emerin influence adipogenic capacity of the cell via nucleocytoplasmic distribution of b-catenin. J Cell Sci 122: 401– 413. Tzur YB, Wilson KL, Gruenbaum Y. 2006b. SUN-domain proteins: ’Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat Rev Mol Cell Biol 7: 782– 788. Tzur YB, Margalit A, Melamed-Book N, Gruenbaum Y. 2006a. Matefin/SUN-1 is a nuclear envelope receptor for CED-4 during Caenorhabditis elegans apoptosis. Proc Natl Acad Sci 103: 13397– 13402. Ulbert S, Antonin W, Platani M, Mattaj IW. 2006. The inner nuclear membrane protein Lem2 is critical for normal nuclear envelope morphology. FEBS Lett 580: 6435– 6441. Ura S, Hayashi YK, Goto K, Astejada MN, Murakami T, Nagato M, Ohta S, Daimon Y, Takekawa H, Hirata K, et al. 2007. Limb-girdle muscular dystrophy due to emerin gene mutations. Arch Neurol 64: 1038–1041. Van Berlo JH, Voncken JW, Kubben N, Broers JL, Duisters R, van Leeuwen RE, Crijns HJ, Ramaekers FC, Hutchison CJ, Pinto YM. 2005. A-type lamins are essential for TGF-b1 induced PP2A to dephosphorylate transcription factors. Hum Mol Genet 14: 2839–2849. Vaughan A, Alvarez-Reyes M, Bridger JM, Broers JL, Ramaekers FC, Wehnert M, Morris GE, Whitfield WGF, Hutchison CJ. 2001. Both emerin and lamin C depend on lamin A for localization at the nuclear envelope. J Cell Sci 114: 2577–2590. Vlcek S, Foisner R. 2007. A-type lamin networks in light of laminopathic diseases. Biochim Biophys Acta 1773: 661–674. Vlcek S, Foisner R, Wilson KL. 2004. Lco1 is a novel widely expressed lamin-binding protein in the nuclear interior. Exp Cell Res 298: 499 –511. Vlcek S, Korbei B, Foisner R. 2002. Distinct functions of the unique C terminus of LAP2a in cell proliferation and nuclear assembly. J Biol Chem 277: 18898–18907. Wagner N, Krohne G. 2007. LEM-Domain proteins: New insights into lamin-interacting proteins. Int Rev Cytol 261: 1 –46. Wagner N, Schmitt J, Krohne G. 2004. Two novel LEMdomain proteins are splice products of the annotated Drosophila melanogaster gene CG9424 (Bocksbeutel). Eur J Cell Biol 82: 605– 616. Wagner N, Kagermeier B, Loserth S, Krohne G. 2006. The Drosophila melanogaster LEM-domain protein MAN1. Eur J Cell Biol 85: 91–105.

Wang Y, Herron AJ, Worman HJ. 2006. Pathology and nuclear abnormalities in hearts of transgenic mice expressing M371K lamin A encoded by an LMNA mutation causing Emery-Dreifuss muscular dystrophy. Hum Mol Genet 15: 2479–2489. Waterham HR, Koster J, Mooyer P, Noort Gv G, Kelley RI, Wilcox WR, Wanders RJ, Hennekam RC, Oosterwijk JC. 2003. Autosomal recessive HEM/Greenberg skeletal dysplasia is caused by 3 b-hydroxysterol delta 14-reductase deficiency due to mutations in the lamin B receptor gene. Am J Hum Genet 72: 1013–1017. Wheeler MA, Davies JD, Zhang Q, Emerson LJ, Hunt J, Shanahan CM, Ellis JA. 2007. Distinct functional domains in nesprin-1a and nesprin-2b bind directly to emerin and both interactions are disrupted in X-linked Emery-Dreifuss muscular dystrophy. Exp Cell Res 313: 2845–2857. Wilhelmsen K, Ketema M, Truong H, Sonnenberg A. 2006. KASH-domain proteins in nuclear migration, anchorage and other processes. J Cell Sci 119: 5021–5029. Wilhelmsen K, Litjens SH, Kuikman I, Tshimbalanga N, Janssen H, van den Bout I, Raymond K, Sonnenberg A. 2005. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J Cell Biol 171: 799– 810. Worman HJ, Bonne G. 2007. “Laminopathies”: Awide spectrum of human diseases. Exp Cell Res 313: 2121– 2133. Worman HJ, Evans CD, Blobel G. 1990. The lamin B receptor of the nuclear envelope inner membrane: A polytopic protein with eight potential transmembrane domains. J Cell Biol 111: 1535–1542. Worman HJ, Yuan J, Blobel G, Georgatos SD. 1988. A lamin B receptor in the nuclear envelope. Proc Natl Acad Sci 85: 8531–8534. Xie X, Fischer JA. 2008. On the roles of the Drosophila KASH domain proteins Msp-300 and Klarsicht. Fly 2: 74– 81. Ye Q, Worman HJ. 1996. Interaction between an integral protein of the nuclear envelope inner membrane and human chromodomain proteins homologous to Drosophila HP1. J Biol Chem 271: 14653–14656. Zastrow MS, Vlcek S, Wilson KL. 2004. Proteins that bind A-type lamins: Integrating isolated clues. J Cell Sci 117: 979 –987. Zhang Q, Ragnauth CD, Skepper JN, Worth NF, Warren DT, Roberts RG, Weissberg PL, Ellis JA, Shanahan CM. 2005. Nesprin-2 is a multi-isomeric protein that binds lamin and emerin at the nuclear envelope and forms a subcellular network in skeletal muscle. J Cell Sci 118: 673 –687. Zhang Q, Skepper JN, Yang F, Davies JD, Hegyi L, Roberts RG, Weissberg PL, Ellis JA, Shanahan CM. 2001. Nesprins: A novel family of spectrin-repeat-containing proteins that localize to the nuclear membrane in multiple tissues. J Cell Sci 114: 4485– 4498. Zhang X, Xu R, Zhu B, Yang X, Ding X, Duan S, Xu T, Zhuang Y, Han M. 2007. Syne-1 and Syne-2 play crucial roles in myonuclear anchorage and motor neuron innervation. Development 134: 901 –908. Zhen YY, Libotte T, Munck M, Noegel AA, Korenbaum E. 2002. NUANCE, a giant protein connecting the nucleus and actin cytoskeleton. J Cell Sci 115: 3207– 3222.

The Nuclear Pore Complex and Nuclear Transport Susan R. Wente1 and Michael P. Rout2 1

Department of Cell and Developmental Biology, Vanderbilt University Medical Center, Nashville, Tennessee 37232

2

Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, New York 10065

Correspondence: [email protected] and [email protected]

Internal membrane bound structures sequester all genetic material in eukaryotic cells. The most prominent of these structures is the nucleus, which is bounded by a double membrane termed the nuclear envelope (NE). Though this NE separates the nucleoplasm and genetic material within the nucleus from the surrounding cytoplasm, it is studded throughout with portals called nuclear pore complexes (NPCs). The NPC is a highly selective, bidirectional transporter for a tremendous range of protein and ribonucleoprotein cargoes. All the while the NPC must prevent the passage of nonspecific macromolecules, yet allow the free diffusion of water, sugars, and ions. These many types of nuclear transport are regulated at multiple stages, and the NPC carries binding sites for many of the proteins that modulate and modify the cargoes as they pass across the NE. Assembly, maintenance, and repair of the NPC must somehow occur while maintaining the integrity of the NE. Finally, the NPC appears to be an anchor for localization of many nuclear processes, including gene activation and cell cycle regulation. All these requirements demonstrate the complex design of the NPC and the integral role it plays in key cellular processes.

axonomically speaking, all life on earth falls into one of two fundamental groups, the prokaryotes and the eukaryotes. The prokaryotes, the first group to evolve, are single cell organisms bounded by a single membrane. About 1.5 billion years later, a series of evolutionary innovations led to the emergence of eukaryotes. Eukaryotes have multiple inner membrane structures that allow for compartmentalization within the cell, and therefore differentiation of the cell and regulation within it. Ultimately, the greater cellular complexity of

T

eukaryotes allowed them to adopt a multicellular lifestyle, as seen in the plants, fungi and animals of today (reviewed in Field and Dacks 2009). Internal membrane bound structures sequester all genetic material in eukaryotic cells. The most prominent of these structures, which gives the eukaryotes their Greek-rooted name, is the nucleus—the central “kernel” (gr. “karyo-”) of the cell. The nucleus is bounded by a double membrane termed the nuclear envelope (NE), which separates the nucleoplasm and genetic

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000562 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000562

S.R. Wente and M.P. Rout

material from the surrounding cytoplasm. However the genetic material in the nucleus is not totally isolated from the rest of the cell. Studded throughout the NE are portals called nuclear pore complexes (NPCs). The NPC is a highly selective, bidirectional transporter for a tremendous range of cargoes. Going into the nucleus, these cargoes include inner nuclear membrane proteins and all the proteins in the nucleoplasm. Going out are RNA-associated proteins that are assembled into ribosomal subunits or messenger ribonucleoproteins (mRNPs). Once transported, the NPC must ensure these cargos are retained in their respective nuclear and cytoplasmic compartments. All the while the NPC must prevent the passage of nonspecific macromolecules, yet allow the free diffusion of water, sugars, and ions. These many types of nuclear transport are regulated at multiple stages, providing a powerful extra level of cellular control that is not necessary in prokaryotes. Assembly, maintenance, and repair of the NPC must somehow occur while maintaining the integrity of the NE. Finally, the NPC appears to be an anchor for localization of many nuclear processes, including gene activation and cell cycle regulation (reviewed in Ahmed and Brickner 2007; Hetzer and Wente 2009). All these requirements demonstrate the complex design of the NPC and the integral role it plays in key cellular processes. STRUCTURE OF THE NPC: SET UP OF THE MACHINE

The specifications of the NPC’s transport machinery represent a huge engineering challenge for evolution. No transitional forms of this elaborate transport system have yet been found in modern day organisms to reveal how it evolved. However, recent clues show that the NPC itself retains in its core a fossil of its ancient origins, indicating that the same mechanism that generated the internal membranes of eukaryotes might also have been responsible for the NPCs and the transport machinery. In the electron microscope, the NPC appears as a complex cylindrical structure with strong octagonal symmetry, measuring some

100 – 150 nm in diameter and 50 – 70 nm in thickness depending on the organism (reviewed in Wente 2000; Lim et al. 2008). This overall appearance seems broadly conserved throughout all eukaryotes. The two membranes of the NE, the outer and inner membranes, join only in a specialized, sharply curved piece of “pore membrane” that forms a grommet in the NE within which the NPC sits. Within each NPC is a core structure containing eight spokes surrounding a central tube. This central hole (30 nm diameter and 50 nm long) is where the nucleoplasm connects to the cytoplasm and where macromolecular exchange occurs. Peripheral filaments are attached to the core, filling the central hole as well as emanating into the nucleoplasm and cytoplasm. These filaments form a basket-like structure on the nuclear side of the NPC (Fig. 1). One can envision the NPC as being comprised of layers of interacting proteins, starting with the core structure, moving outwards through its peripheral filaments, and then to associating clouds of soluble transport factors and peripherally associating protein complexes in the nucleus and cytoplasm (Rout and Aitchison 2001). These protein interactions can occur on radically different time scales. Some proteins form relatively permanent associations with the core structure, and so are termed nuclear pore complex components or “nucleoporins” (“Nups”). Other proteins associate transiently with the NPC, either constantly cycling on and off or attaching only at particular times in the cell’s life cycle. The NPC is covered in binding sites for these transiently associating proteins. Because the NPC is neither a motor nor an enzyme, the interactions provided by its binding sites wholly define the function of the NPC. Recent work, mainly in the yeast Saccharomyces cerevisiae and in vertebrates, has begun to elucidate the molecular architecture of the NPC (Rout et al. 2000; Cronshaw et al. 2002; Alber et al. 2007b). Given its large size, the main body of the NPC comprises a surprisingly small number of 30 unique proteins (Table 1). However, because of the NPC’s eightfold symmetry, these Nups are each present in

Nuclear Pore Complexes and Nuclear Transport Central tube

Core scaffold

Outer ring Inner ring

Lumenal ring

Cytoplasmic filaments Cytoplasmic FG nups Symmetric FG nups Nuclear envelope

Pore membrane

Linker nups

Nucleoplasmic FG nups Nuclear basket

Figure 1. Major structural features of the NPC (based on the architectural map of Alber et al. (2007b); see Table 1

and main text for details).

multiple copies (usually 16 per NPC) resulting in around 400 polypeptides for each NPC in every eukaryote (Rout et al. 2000; Cronshaw et al. 2002; DeGrasse et al. 2009). Further redundancy is evident from the recent mapping of the yeast NPC. Indeed, the NPC’s structure is modular, consisting of a few highly repetitive protein fold types (Devos et al. 2006; Alber et al. 2007b; DeGrasse et al. 2009). This suggests that the bulk of the NPC’s structure has evolved through multiple duplications of a small precursor set of genes encoding just a handful of progenitor Nups.

To understand its evolutionary origins, the NPC of the highly divergent Trypanosoma was recently characterized (DeGrasse et al. 2009). Despite significant divergence in primary structure, the Trypanosome NPC consists mainly of the motifs and domains found in vertebrate and yeast NPCs, indicating on a molecular level that the basic structural components of the NPC are conserved across all eukaryotes. Importantly, this also strongly implies that the last common eukaryotic ancestor had many features in common with contemporary NPCs, and perhaps provided a key adaptive advantage

Table 1. Nucleoporin homologs of yeast and vertebrates NPC substructure

Outer Ring

Inner Ring

Cytoplasmic FG Nups and Filaments Lumenal Ring Symmetric FG Nups Linker Nups Nucleoplasmic FG Nups and Filaments

Yeast components

Vertebrate components

Nup84 subcomplex (Nup84, Nup85, Nup120, Nup133, Nup145C, Sec13, Seh1) Nup170 subcomplex (Nup170, Nup157, Nup188, Nup192, Nup59, Nup53) Nup159, Nup42

Nup107-160 complex (Nup160, Nup133, Nup107, Nup96, Nup75, Seh1, Sec13, Aladin, Nup43, Nup37) Nup155 subcomplex (Nup155, Nup205, Nup188, Nup35)

Ndc1, Pom152, Pom34 Nsp1, Nup57, Nup49, Nup145N, Nup116, Nup100 Nup82, Nic96 Nup1, Nup60, Mlp1, Mlp2

Gp210, Ndc1, Pom121 Nup62, Nup58/45, Nup54, Nup98

Nup358, Nup214, Nlp1

Nup88, Nup93 Nup153, Tpr

S.R. Wente and M.P. Rout

for this organism that has been retained, little changed, ever since. The structural proteins making up the bulk of the spokes and rings give the NPC its shape and strength (Fig. 1). These core proteins of the NPC also maintain the stability of the nuclear envelope and facilitate the bending of the pore membrane into the inner and outer NE membranes. The most equatorial rings, termed the inner rings, are comprised of the Nup170 complex (yeast) or Nup155 complex (vertebrates) (Aitchison et al. 1995; Grandi et al. 1997; Miller et al. 2000) (Fig. 1). The inner rings are sandwiched between the outer rings, which are comprised of the Nup84 complex (yeast) or Nup107 complex (vertebrates) (Table 1) (Siniossoglou et al. 1996; Fontoura et al. 1999; Siniossoglou et al. 2000; Belgareh et al. 2001; Vasu et al. 2001). Together, these Nup complexes form a scaffold that hugs the curved surface of the pore membrane and helps form the central tube through which macromolecular exchange occurs (Alber et al. 2007a; Alber et al. 2007b). Nups in the core scaffold represent roughly half the mass of the whole NPC and are composed almost entirely of either b-propeller folds, a-solenoid folds, or a distinct arrangement of both in an amino-terminal b-propeller followed by a carboxy-terminal a-solenoid fold. The core scaffold of all eukaryotes appears to retain this basic fold composition (Devos et al. 2004; Devos et al. 2006; DeGrasse et al. 2009). Strikingly, there are similarities between the structures of the core NPC scaffold curving around the pore membrane and other membrane-associated complexes such as clathrin/ adaptin, COPI, and COPII (Fig. 1) (Devos et al. 2004; Devos et al. 2006). Clathrin/adaptin is involved in coat-mediated endocytosis at the plasma membrane, and COPI and COPII are responsible for coat-mediated vesicular transport between the plasma membrane and endomembrane systems such as the Golgi and ER. Indeed, the similarities between core scaffold Nups and coating complexes have been borne out in numerous crystallographic studies (Berke et al. 2004; Hsia et al. 2007; Brohawn et al. 2008; Debler et al. 2008; Brohawn et al. 2009; Leksa et al. 2009; Seo et al. 2009; Whittle and Schwartz

2009), although nearly 2 billion years of evolution have made it difficult at first glance to recognize the common origin of these two groups. However, their common b-propeller and helixturn-helix repeat structure is still unmistakable (Brohawn et al. 2008; Field and Dacks 2009). In NPCs the “coat” comprises the core scaffold of the NPC, where—analogous to the curved membrane of a vesicle being stabilized by a COP or clathrin coat—it stabilizes the curved pore membrane. These similarities also give a tantalizing glimpse into the deep evolutionary origins of eukaryotes. It seems early protoeukaryotes distinguished themselves from their prokaryotes by acquiring a membrane-curving protein module, the “proto-coatomer” (likely composed of a simple b-propeller/a-solenoid protein), that allowed them to mold their plasma membranes into internal compartments. Modern eukaryotes diversified this module into many specialized membrane coating complexes, accounting for the evolution of their internal membrane systems (Devos et al. 2004; Devos et al. 2006). The framework of the NPC serves two key transport purposes: to form a barrier of defined permeability within the pore, and to facilitate transport of selected macromolecules across it. Both processes are dependent on the correct positioning of critical Nups in the NPC architecture (Radu et al. 1995; Strawn et al. 2004; Liu and Stewart 2005). Attached to the inside face of the NPC core scaffold, facing the central tube’s cavity, are groups of nucleoporins termed “linker nucleoporins” (Fig. 1). Together with the inner ring, these seem to form most of the attachment sites for a last set of nucleoporins, termed “FG Nups” (Alber et al. 2007b). These FG Nups, named for their phenylalanine-glycine repeats, are the direct mediators of nucleocytoplasmic transport (Radu et al. 1995; Strawn et al. 2004; Liu and Stewart 2005) (see the following section). The core NPC scaffold is connected to a set of integral membrane proteins, which form an outer luminal ring in the NE lumen and anchor the NPC into the NE (Nehrbass et al. 1996; Alber et al. 2007a; Alber et al. 2007b) (Fig. 1). Oddly, the membrane nucleoporins seem poorly

Nuclear Pore Complexes and Nuclear Transport

conserved—if at all—across the eukaryotes. The fact that all the currently known pore membrane proteins in Aspergillus nidulans can seemingly be dispensed with for NPC function and assembly might indicate that there are not strong pressures for their conservation, and that there are other membrane proteins that can serve the role (Liu et al. 2009). This fact also sets up a quandary—if most or all of the NPC’s presumed membrane anchors are dispensible, how then is the NPC reliably anchored to the membrane? Several groups are seeking the answer to this question (Hetzer and Wente 2009). OPERATION OF THE MACHINE: THE SOLUBLE PHASE

Understanding the transport machine requires resolving both its barrier and binding activities. How the NPC machine balances both of these selective functions has been a challenging mystery. Studies of fluorescently labeled sized dextrans or gold particles microinjected into cells (Feldherr and Akin 1997; Keminer and Peters 1999) have defined the practical permeability limits of the NPC, showing that under physiological time scales, macromolecules greater than 40 kDa in size do not show any measureable redistribution between the nucleus and cytoplasm and thus, no movement through the NPC. Conversely, metal ions, small metabolites, and molecules less than 40 kDa in mass or 5 nm in diameter can pass relatively freely. NPC permeability is altered in several yeast nup mutants, pinpointing NPC structural elements, including core scaffold components, that are critical to the assembly or maintenance of this barrier (Shulga et al. 2000; Denning et al. 2001; Shulga and Goldfarb 2003; Strawn et al. 2004; Patel et al. 2007). Larger macromolecules overcome this permeability barrier by interacting either directly with the NPC themselves or through soluble transport factors. These macromolecules account for a tremendous variety of cargo including proteins, tRNAs, ribosomal subunits, and viral particles (reviewed in Macara 2001). Overall, the NPC is capable of transporting cargo up to 39 nm in diameter. This is on par with the size of the ribosomal subunits

and viral capsids that are known to move as intact complexes (Pante and Kann 2002). Macromolecules larger than this can still be translocated across the NPC, including mRNPs (mRNAs coated with RNA-binding proteins) with masses reaching several hundred thousand daltons. EM images of Balbiani ring mRNP particles associated with the NPC show the posttranscriptional 50 nm mRNA–protein particles to rearrange into rodlike structures, decreasing their maximum diameter to 25 nm (Mehlin et al. 1992). Thus, cargoes above a limiting diameter must rearrange to pass through the selective barrier of the NPC (Daneholt 2001). A transport signal and a shuttling receptor for that transport signal are the minimal requirements for any facilitated translocation (reviewed in Mattaj and Englmeier 1998; Pemberton and Paschal 2005). The targeting of proteins into or out of the nucleus requires specific amino acid sequence spans, termed nuclear localization sequences (NLSs) or nuclear export sequences (NESs). All the information required to target a protein to the nucleus is within these short sequences. In fact, fusion of an NLS to a nonnuclear protein is sufficient to mediate its transport and import to the nucleus (Goldfarb et al. 1986). For proteins, there are many distinct types of NLSs and NESs. For example, the classical NLS (cNLS) is the simple five amino acid peptide KKKRK, necessary and sufficient for targeting its attached protein to the nucleus (Goldfarb et al. 1986), whereas many proteins carry a more complex “bipartite” NLS consisting of two clusters of basic amino acids, separated by a spacer of roughly 10 amino acids (Dingwall et al. 1988). However, the full spectrum of sequences recognized by each transport receptor has not yet been carefully and fully defined. The most in depth analysis of NLS structural recognition by a transport receptor and extrapolation to predicting cargoes on a broader genome level has only been reported for one transport receptor (Lee et al. 2006). The key parameters defining an NLS or NES include critical tests for necessity and sufficiency in the endogenous protein. Importantly, some proteins undergo dynamic cycles of nuclear import and export and harbor both NLSs and

S.R. Wente and M.P. Rout

NESs. This can increase the complexity of identifying the signals. Moreover, the recognition and accessibility of the signals can be controlled by signaling, cell cycle, and developmental events (reviewed in Weis 2003; Terry et al. 2007). During NPC translocation, soluble transport factors are required to either bring cargo to the NPC or modulate cargo translocation across the NPC. Most of these soluble transport factors come from the family of proteins known

as the karyopherins (Table 2). The karyopherins (also called importins, exportins, and tranportins) were the first family of shuttling transport factors discovered. Fourteen karyopherin family members are found in Saccharomyces cerevisiae whereas at least 20 have been found in metazoans (reviewed in Fried and Kutay 2003; Pemberton and Paschal 2005). Most karyopherins bind their cargoes directly. However, in some cases an adaptor protein is needed in addition to the

Table 2. Karyopherin transport factors of yeast and vertebrates S. cerevisiae Karyopherins

Vertebrate Karyopherins

Kap95

Importin-b1

Kap104

Transportin or Transportin 2

Kap108/Sxm1

Importin 8

Kap109/Cse1 Kap111/Mtr10

CAS Transportin SR1 or SR2

Kap114

Importin 9

Kap119/Nmd5

Importin 7

Kap120 Kap121/Pse1

HsRanBP11 Importin 5/Importin b3/RanBP5

Kap122/Pdr6 Kap123

Importin 4

Kap127/Los1 Kap142/Msn5

Exportin-t Exportin 5

Crm1/Xpo1

Importin 13 CRM1/Exportin 1

— — —

Exportin 4 Exportin 6 Exportin 7/RanBP16

Examples of Cargo(s): (v) – vertebrate, (sc) – S. cerevisiae

Imports via sc-Kap60/v-importin-a adaptor proteins with cNLS; Imports via v-Snurportin the UsnRNPs; with no adaptor, imports v-cargo SREBP-2, HIV Rev, HIV TAT, cyclin B Imports sc-cargo – Nab2, Hrp1; v-cargo – PY-NLS proteins, mRNA-binding proteins, histones, ribosomal proteins Imports sc-cargo – Lhp1, ribosomal proteins; v-cargo – SRP19, Smad Imports sc-cargo – Kap60/Srp1; v-cargo – importin as Imports sc-cargo – Npl3, tRNAs; v-cargo – SR proteins, HuR Imports sc-cargo – TBP, histones, Nap1, Sua7; v-cargo – histones, ribosomal proteins Imports sc-cargo – Hog1, Crz1, Dst1, ribosomal proteins, histones; v-cargo – Smad, ERK, GR, ribosomal proteins Imports sc-cargo – Rpf1 Imports sc-cargo – Yra1, Spo12, Ste12, Yap1, Pho4, histones, ribosomal proteins; v-cargo – histones, ribosomal proteins Imports sc-cargo – Toa1 and Toa2, TFIIA Imports sc-cargo – SRP proteins, histones, ribosomal proteins; v-cargo - Transition Protein 2, histones, ribosomal protein S3a Exports tRNAs sc-cargo – imports replication protein A; exports Pho4, Crz1, Cdh1; v-cargo - exports pre-miRNA v-cargo – imports UBC9, Y14; exports eIF1A Exports proteins with leucine-rich NES, 60S ribosomal subunits (via NMD3 adaptor), 40S ribosomal subunits v-cargo – imports SOX2, SRY; exports Smad3, eIF5A Exports profilin, actin Exports p50-RhoGAP

Based on references cited within and adapted from Tran et al. 2007a and DeGrasse et al. 2009.

Nuclear Pore Complexes and Nuclear Transport

karyopherin to recognize signals. Not only do karyopherins have a cargo-binding domain, they also have an NPC-binding domain(s) as well as a binding domain at the amino-terminus for the small Ras-like GTPase Ran (see the following paragraph) (reviewed in Macara 2001; Harel and Forbes 2004). Overall, karyopherin family members share only modest sequence homology, with the greatest similarity being within their Ran-binding domains (Gorlichet al. 1997). However, a hallmark architecture within the karyopherins, as determined by recent highresolution structural studies, is the tandem HEAT-repeat fold formed by antiparallel helices connected by a short turn (reviewed in Conti and Izaurralde 2001). The HEAT-repeats arrange to form a superhelical structure, similar to a snail’s shell. This folding is reminiscent of the helix-turn-helix repeats found in the NPC’s core scaffold proteins. This similarity raises the intriguing possibility that karyopherins diverged from a common structure involved in both stationary and soluble phases of transport.

Import cycle

The association and dissociation of a karyopherin-cargo complex is regulated by direct binding of the small GTPase Ran (Fig. 2) (reviewed in Fried and Kutay 2003; Madrid and Weis 2006; Cook et al. 2007). In vitro binding studies show that import complexes are dissociated by RanGTP binding. Conversely, export complexes are formed via RanGTP association (Rexach and Blobel 1995; Floer and Blobel 1996; Chi and Adam 1997; Floer et al. 1997; Kutay et al. 1997a; Kutay et al. 1997b; Nakielny et al. 1999). Based on the localizations of the Ran GTPase activating protein (RanGAP) in the cytoplasm and the Ran guanine nucleotide exchange factor (RanGEF) in the nucleoplasm, cytoplasmic Ran is primarily in the GDP-bound state whereas nucleoplasmic Ran is kept primarily in the GTP-bound state (Fig. 2). The gradient formed from these localizations has been elegantly demonstrated by imaging fluorescence resonance energy transfer-based biosensors (Kalab et al. 2002). The RanGTP gradient across the two faces of the NPC is essential

Export cycle

NLS cargo

T

Import karyopherin

T

D

D

RanGDP

RanGDP

RanGTP

RanGTP T T

T T

T

T

Export karyopherin NES cargo

Figure 2. The nuclear transport cycle for karyopherins and their cargos. See main text for details.

S.R. Wente and M.P. Rout

for establishing the directionality of karyopherin-mediated transport. The pathway for karyopherin-mediated translocation is well described (reviewed in Weis 2003; Terry et al. 2007) (Fig. 2). For import, a specific karyopherin recognizes its cognate cargo in the cytoplasm where RanGTP levels are low. The karyopherin mediates the binding of the import complex to the NPC and facilitates translocation through the NPC. Once the complex moves through the NPC, release and dissociation of the karyopherin-cargo complex are stimulated by RanGTP in the nucleus. The karyopherin bound to RanGTP is then recycled back to the cytoplasm. Finally, GTP hydrolysis of Ran on the cytoplasmic side frees the karyopherin to interact with a second cargo molecule for further cycles of transport. Overall, Ran decreases the affinity of the karyopherin for its cargo (reviewed in Macara 2001; Cook et al. 2007). For export complexes, an analogous process occurs, but in this case, RanGTP binding increases the affinity of the karyopherin b for the export cargo. For example, for the exporting karyopherin Crm1 and an export cargo SPN1 (snurportin 1 adaptor for UsnRNPs), the Crm1 affinities for RanGTP and SPN1 in the ternary RanGTP-Crm1-SPN1 complex are increased 1000-fold (Paraskeva et al. 1999; Monecke et al. 2009). Actual movement through the NPC does not require energy input. The Ran affinity switches provide the energy for efficient cargo delivery and release. The only possible exception to this rule involves the import of large cargoes, where the presence of Ran and hydrolyzable GTP may be required for the import of cargoes .500 kDa in vitro (Lyman et al. 2002). It was also originally thought that an individual karyopherin was adapted for either import or export, but not both. However, there are now documented examples of karyopherins functioning in both import and export, although with different cargoes in each direction (Yoshida and Blobel 2001). In addition to protein import and export, karyopherins can also transport RNAs. For example, the karyopherins Crm1 and exportin-t mediate the export of uridine-rich small nuclear RNAs (U snRNAs) and tRNAs, respectively

(Simos et al. 2002; Rodriguez 2004). Transport is accomplished via direct binding of karyopherins to RNA or to signal sequences within the protein components of the RNP complexes. For example, Crm1 does not bind UsnRNAs directly and requires the adaptor PHAX that binds the cap-complex on the RNA (Ohno et al. 2000). However, exportin-t directly interacts with tRNAs (Arts et al. 1998; Hellmuth et al. 1998; Kutay et al. 1998; Lipowsky et al. 1999). Karyopherins are also involved in the export of some viral RNAs, including their mRNAs (Carmody and Wente 2009). However, the primary mRNP export transport receptor is a nonkaryopherin designated Mex67 in yeast and NXF1 in metazoans, which heterodimerizes with a protein termed respectively Mtr2 or p15/Nxt1 (Erkmann and Kutay 2004). Even though Mex67 is unrelated in sequence and structure to the karyopherin family, it has all the requirements of a transport receptor: cargo binding, nucleocytoplasmic shuttling, and NPC-binding. The stoichiometry of the Mex67-Mtr2 heterodimer per transported mRNP is unknown; Mex67-Mtr2 either is recruited directly to the mRNA or interacts cotranscriptionally with proteins of the mRNP assembly (Erkmann and Kutay 2004; Carmody and Wente 2009). Like karyopherins, Mex67Mtr2 heterodimers bind directly to FG Nups, although it seems they prefer different subsets of FG Nups to their karyopherin counterparts, which might reflect how the karyopherin mediated transport pathways and mRNPexport pathways are kept apart at the NPC (Strawn et al. 2001; Terry et al. 2007; Terry and Wente 2007). As mRNP is a major source of traffic across the NPC, it is interesting that most of this transit is facilitated by non-karyopherin carriers. Because Ran is not utilized to establish a gradient, directionality in the mRNA export pathway is conferred by proteins that modify the mRNPs as they cross the NPC. Chief among these is the protein Dbp5. Dbp5 is a member of the SF2 helicase superfamily of RNAdependent ATPases (Snay-Hodge et al. 1998; Tseng et al. 1998) and carries a DEAD/H-box sequence motif. Such DEAD-box proteins are

Nuclear Pore Complexes and Nuclear Transport

involved in the alteration of RNA structure, such as the facilitation of RNA duplex formation and unwinding or aiding the association and dissociation of RNA-binding proteins (Jankowsky et al. 2001; Fairman et al. 2004; Yang and Jankowsky 2006). Much like the GDP/GTP triggered switches for Ran, DEADbox proteins potentially use ADP/ATP for nucleotide-dependent conformational switches (Henn et al. 2002; Tran et al. 2007b; Henn et al. 2008; Fan et al. 2009). Dbp5 ATPase activity is activated by Gle1, a protein that exchanges between the nucleus and cytoplasm with a docking site on the cytoplasmic filaments of the NPC (Cole and Scarcelli 2006). Gle1 activation of Dbp5 in turn requires the soluble molecule inositol hexakisphosphate (IP6) to increase the binding affinity of Dbp5 to RNA (Alcazar-Roman et al. 2006; Weirich et al. 2006). This raises the possibility that IP6 is acting as a signaling molecule to somehow regulate the mRNA export machinery (York et al. 1999). The Dbp5 on the cytoplasmic side of the NPC triggers the release of mRNP proteins such as Mex67 and Nab2, effectively remodeling the mRNP protein composition as they exit the NPC as well as imparting directionality to mRNP transport (Tran et al. 2007b). There also seems to be a checkpoint at the NPC basket, to ensure that mRNAs have been correctly spliced and packaged into mRNPs immediately before export. SUMO is a ubiquitin-like molecule that can be attached to proteins as a signal or modulator of their function. The SUMO protease, Ulp1, is retained at the NPC via its interaction with the nuclear basket component Mlp/Tpr, and it has been suggested that sumoylation of mRNPs acts to signal that they are correctly assembled. Conversely, the desumoylation of incorrectly spliced mRNPs at the nuclear basket prevents their export and instead targets them for degradation (Lewis et al. 2007; Xu et al. 2008). Other nonkaryopherin carriers for NPC translocation exist with the same capacity to dock at various NPC sites via the FG Nups. Ntf2/p10 is a carrier for RanGDP importing Ran to the nucleus. Ntf2/p10 also uses the localization of RanGEF in the nucleus to create

a differential affinity for RanGDP versus RanGTP (reviewed in Pemberton and Paschal 2005; Cook et al. 2007). There are also some proteins with karyopherin-like HEAT-repeats that require no receptor and translocate independently of a carrier. The driving force for their accumulation in the nucleus is their affinity for binding targets in the nucleoplasm (Fagotto et al. 1998). Perhaps the most abundant nuclear transport cargos in many cells are the 60S and 40S ribosomal subunits, on the way from synthesis in the nucleolus to form mature ribosomes in the cytoplasm. Export of ribosomal subunits is still poorly understood, and it seems that unlike the case of a single karyopherin bound to each protein cargo molecule as described earlier, several transport receptors must chaperone each ribosomal subunit across the NPC. The earliest identified ribosomal exporter was Crm1 (Fornerod et al. 1997; Fukuda et al. 1997; Stade et al. 1997). For export of the 60S subunit, the NES is provided by the export adaptor Nmd3 (Johnson et al. 2002). Nmd3 binds reversibly to the 60S subunit while the latter is being assembled in the nucleus. After export into the cytoplasm, a GTPase called Lsg1 appears to mediate the release of Nmd3 from the 60S ribosomal subunit (Zemp and Kutay 2007). In yeast the Mex67-Mtr2 heterodimer has also been suggested to serve as an export receptor for the 60S subunit (Yao et al. 2007). Although more karyopherin-like proteins have recently been identified that play a role in ribosomal export (Oeffinger et al. 2004), other nonkaryopherin receptors like Arx1 have been found, though Arx1 still interacts with the same NPC docking sites as karyopherins (Bradatsch et al. 2007). Why might so many transport factors be required to ferry the ribosomal subunits out of the nucleus? One reason might be explained by the analogy of maneuvering ships out of the harbor. One tugboat is all that is required to move a small barge quickly and efficiently, but on its own does not have the power to move an ocean liner, which instead needs many tugboats together to take it safely out of harbor. Similarly, as the efficiency of passage through the NPC decreases with cargo size

S.R. Wente and M.P. Rout

(see the preceding section) but increases with binding strength to FG Nups (see the following section), many transport factors are needed to carry large cargos across the NPC. Regardless, the fundamental mechanism for transport in all these cases of different carriers is still dependent on the same features of the NPC machine. OPERATION OF THE NPC: THE “FG” TRANSLOCATION MACHINERY

Although the cargo binding mechanisms of the transport factors discussed earlier vary, almost all binding sites with the NPC are found in the same class of Nups. These Nups are collectively termed FG Nups because they contain “FG repeat regions.” The NPC core framework provides the correct positioning of these FG Nups so that they flank and fill the central tube. Approximately a third of all Nups contain FG repeat regions, which consist of multiple small hydrophobic clusters containing an FG (Phe-Gly) dipeptide (usually FG, FXFG, or GLFG) separated by 20 – 70 residue hydrophilic linkers (reviewed in Rout and Wente 1994; Allen et al. 2001; Tran and Wente 2006). Binding of these repeats is cooperative, with two to four repeats associating with each transport factor molecule (Bayliss et al. 2002; Grant et al. 2003; Isgro and Schulten 2005; Liu and Stewart 2005). With 5– 50 repeats per FG Nup and 200 FG Nups per NPC, there is a potential for .1000 transport factor binding sites per NPC. Imaging of the trajectories of a single translocating molecule through the NPC is consistent with movement between multiple binding sites ( presumably the many FG repeats) within the NPC (Yang et al. 2004; Kubitscheck et al. 2005; Yang and Musser 2006). FG repeat regions are natively disordered and having no secondary structure they instead form filaments that can diffusively writhe around their attachment sites at the NPC. Although some will be diffuse random coils, others may have enough internal cohesion to form more compact “molten globules” (Denning et al. 2003; Lim et al. 2006; Patel et al. 2007; Krishnan et al. 2008). By forming a dense meshwork in and around the central tube, the FG repeats appear to set

up an entropic potential that excludes macromolecules from their vicinity, while permitting the approach of small molecules such as water and nucleotides (Lim et al. 2006). However, karyopherins overcome this barrier as they bind FG Nups and, passing from FG Nup to FG Nup, diffuse readily through the NPC. Indeed, this model is supported by the fact that a simple device with similar properties—a nanotube coated with FG Nups—replicated many of the features of nuclear transport (Jovanovic-Talisman et al. 2009). In terms of the Gibb’s free energy required to cross the NPC, the enthalpy for binding the FG Nups cancels out the entropic barrier of the central tube region, flattening the energy landscape and thus lowering the activation energy of translocation across the NE. This is analogous to how an enzyme or catalyst lowers the activation energy of the transition state permitting a substrate to convert to product. This concept has been termed “virtual gating” (Rout et al. 2000; Rout et al. 2003). Nevertheless, despite much work over the last few years, how exactly this gating works at the molecular level remains unclear and several models have attempted to explain it. One possibility is that the natively unfolded FG Nups could act as a “polymer brush” where rapid diffusion-driven movements of their unfolded domains would sweep away macromolecules from their vicinity. Although a central tube filled with such brushes would exclude nonbinding macromolecules, transport factors would overcome this exclusion by binding to the FG repeats and so pass through the NPC (Rout et al. 2000; Rout et al. 2003; Lim et al. 2006). In another model, regions of FG repeats may collapse upon binding transport factors, as indicated by atomic force microscopy data (Lim et al. 2007a; Lim et al. 2007b). As they pass through the central tube, transport factors would open up their own passageway through the meshwork of FG repeats. A different view is provided by the “saturated” model (Frey et al. 2006; Frey and Gorlich 2007), originally the “hydrophobic gel” model (Ribbeck and Gorlich 2002). Here it is suggested that the phenylalanines in the FG repeat regions are crosslinked with each other through their Phe residues to

Nuclear Pore Complexes and Nuclear Transport

form a dense gel of FG repeat filaments. Transport factors pass through this gel by binding the FG repeats, and in doing so they dissolve the crosslinks. Other macromolecules are therefore excluded because they cannot do this. A more recent “reduction in dimensionality”-model posits that the FG repeat regions form a layer coating the inner walls of the central tube (Peters 2005). Although transport factors enter this layer through binding giving them full access to the tube’s volume, nonbinding molecules can only pass through the narrow FG Nupfree middle. Recently, work has emerged that suggests while crossing the NPC, transport factors not only carry cargoes but also enhance the selectivity of the NPC by competing away nonspecific macromolecules, indicating that the transport factors themselves are also important components of the selectivity barrier (Timney et al. 2006; Zilman et al. 2007; Jovanovic-Talisman et al. 2009). Not all of these ideas are mutually exclusive, and “hybrid” models have also been proposed (Strawn et al. 2004; Patel et al. 2007; Krishnan et al. 2008). FG Nup repeat regions seem to fall into several classes on the basis of their charge density and FG repeat motif type. Moreover, these classes seem to have somewhat different physical behaviors (Krishnan et al. 2008). Each class of FG Nup appears to be preferred by only certain kinds of transport factor. Thus, these different FG Nup classes might serve to mediate multiple, simultaneous, functionally independent transport pathways through the NPC (reviewed in Tran and Wente 2006). Supporting evidence for this has come from in vivo studies using target combinations of different FG region deletions. Certain deletion combinations can affect passage of one transport factor through the NPC without affecting another (Strawn et al. 2004; Terry and Wente 2007). FG Nups organization around the NPC can be categorized as either symmetric or asymmetric. “Symmetric” FG Nups are found on both sides of the NPC closely packed around and within the central tube. “Asymmetric” FG Nups are found primarily on either the nuclear or cytoplasmic side only, and significantly further from the tube (Rout et al. 2000; Alber et al.

2007b). Deletion mutagenesis experiments in yeast have shown that the FG regions of asymmetric Nups are not essential for transport (Strawn et al. 2004; Zeitler and Weis 2004). Instead, they may guide the directionality of transport-factor/cargo complexes by providing a high-affinity binding platform at the far end of a transport factor’s route through the NPC. A transport factor would be drawn to this site and prevented by it from returning through the NPC until Ran terminates the transport reaction, in this way increasing the efficiency of transport (Rout et al. 2003; Strawn et al. 2004; Zilman et al. 2007). REGULATORS OF TRANSLOCATION: THE NON-FG DETERMINANTS

Unidirectional transport through the NPC can be viewed as being driven by the formation of gradients, in which cargos move from a “source” to a “sink.” Karyopherin-cargo complexes form on one side of the NPC and diffuse down their concentration gradient to the other side, where their concentration is low because they are disassembled there. This “source and sink” process also drives other forms of nucleocytoplasmic transport, such as the attachment of RNA binding proteins on one side of the NPC and their removal on the other side during RNP export (reviewed in Rout et al. 2003; Stewart 2007). The NPC on its own does not enforce directionality (Nachury and Weis 1999). Single molecule studies show the transport receptor-cargo complexes seem to move randomly within the NPC channel until they exit either face of the NPC (Yang et al. 2004). Thus, in a physiological context, other NPC events and non-FG Nup binding sites are required to provide transport efficiency, regulation, and directionality (Fig. 3). As detailed earlier, there are specific factors for mediating transport directionality and cargo release: The RanGTPase for karyopherins and the Gle1-IP6 activated Dbp5 for mRNA export. These use non-FG NPC docking sites that increase their local concentration at their respective sites of action. Dbp5 and Gle1 bind to distinct but structurally juxtaposed Nups on the cytoplasmic filaments. In budding yeast,

S.R. Wente and M.P. Rout Transport factors & cargos

Transport modulators

Nuclear organization

NLS cargo Import karyopherin

SUMOylation/ RanGAP complex

Membrane cargo

Gle1/1P6/Dbp5 complex Ran binding sites

Active gene tethered at NPC

SUMO peptidase

Exporting RNP

Ran Export karyopherin

RNP proofreading complex

NES cargo Mex67 mRNP export complex

Ran binding sites

Microtubule attachment NPC assembly factors Lamina and Mlp/Tpr tethering of NPC Mad proteins

Mobile nup

Figure 3. The main NPC interactors: left, transport factors and their cargos, center, modulators of RNP export,

karyopherins or Ran, right, cytoskeletal, nucleoskeletal, and chromosomal elements.

Dbp5 interacts with Nup159 (Hodge et al. 1999; Schmitt et al. 1999) and Gle1 with Nup42 (Murphy and Wente 1996; Strahm et al. 1999), with both sets of binding partners fully conserved in metazoans. The NPC can also help to regulate the GTP-bound state of Ran in its vicinity. Thus, mammalian cells have binding sites for sumoylated-RanGAP on the cytoplasmic filament Nup358, likely to help enhance the dissociation of exporting karyopherin complexes as they exit the NPC (Mahajan et al. 1997; Matunis et al. 1998). In organisms like plants that lack Nup358, unique NE associated proteins facilitate the localized anchoring of RanGAP (Xu et al. 2007; Zhao et al. 2008), so it seems that regulating the GTP-bound state in the immediate vicinity of the NPC is important for transport efficiency in many eukaryotes. Although the FG domains do not form a gradient of docking sites in the translocation channel, there are non-FG high affinity docking sites located at terminal transport steps that influence efficiency. In particular, two Nups on the opposing cytoplasmic and nuclear faces provide a virtual smorgasbord of binding sites for different factors that regulate cell physiology: Nup358 and Nup153, respectively (Fig. 3). In metazoan cells, the cytoplasmic facing Nup358 provides a high affinity binding site for the mRNA export factor NXF1 on the cytoplasmic face of the NPC (Forler et al. 2004). Similarly, Nup153 provides a high affinity binding site for an importing karyopherin on the nuclear face of the NPC (Moroianu et al. 1997).

Although both have FG domains it is the nonFG docking sites that are functionally distinguishing. Nup358 was originally identified as a RanGTP binding protein (Wu et al. 1995; Yokoyama et al. 1995) and binds sumoylated RanGAP. However, it also has a domain with E3 SUMO1 ligase activity and docks the sumolylation cofactor enzyme Ubc9 to facilitate the addition of SUMO to targets directly at the cytoplasmic filaments (Pichler et al. 2002; Reverter and Lima 2005). Nup358 also harbors a cyclophilin A homologous domain that associates with subunits of the 26S proteasome and might modulate ubiquitin-proteasome function (Wu et al. 1995; Yi et al. 2007). The amino-terminal domain of Nup358 allows interaction with microtubules during interphase as well as with kinetochores in mitosis (Salina et al. 2003; Joseph et al. 2004; Joseph and Dasso 2008). On the nuclear side of the NPC, Nup153 has a Zn finger Ran binding domain (Nakielny et al. 1999; Higa et al. 2007) attracting Ran to the vicinity of the NPC to regulate karyopherinbased transport events at its nuclear face. In sharp contrast to the Nup358-SUMO1 ligase activity, Nup153 actually binds a SUMO1/ sentrin/SMT3 specific peptidase 2, perhaps to help desumolyate misassembled mRNPs (Hang and Dasso 2002; Zhang et al. 2002). Nup153 also possesses a novel single-stranded RNA binding domain (Ball et al. 2004) that is perhaps concomitant with a function in RNP export. Nup153 also plays a role in recruiting the

Nuclear Pore Complexes and Nuclear Transport

COPI coatomer complex, possibly to mediate NPC mitotic disassembly (Liu et al. 2003). Taken together, Nup358 and Nup153 are but two examples of an emerging paradigm for multifunctional nucleoporins. They demonstrate that the NPC serves as a scaffold for a diversity of processes that are distinct from those strictly associated with simply moving a cargo from one side of the NPC to the other. Transport can also be modulated at the level of the transport factors and their cargos. A common method is to regulate the binding strength or accessibility of an NLS or NES in a cargo for its cognate karyopherin. This often involves phosphorylation in or near the signal sequence of the cargo. Another method involves modulating the level of transport machinery components, usually particular subtypes of karyopherin. Using this method, transport of particular proteins recognized by that karyopherin is prevented until the karyopherin itself is expressed (reviewed in Poon and Jans 2005). THE NPC AS A NUCLEAR COMPONENT

Although the most dynamic process associated with the NPC is nucleocytoplasmic transport, the NPC should not be regarded as a static structure. New NPCs are assembled de novo during interphase (D’Angelo et al. 2006) and, in higher eukaryotes at least, the NPC is reversibly disassembled during mitosis (reviewed in Hetzer et al. 2005; Fernandez-Martinez and Rout 2009). There is emerging evidence for multiple nonnucleoporin accessory factors that play roles in NPC biogenesis and maintenance (Fig. 3). This includes proteins that regulate membrane composition and membrane dynamics such as Acc1 and Apq12, respectively (Schneiter et al. 1996; Scarcelli et al. 2007). Changes in the lipid composition or membrane fluidity can impact NPC structure and localization. Chromatin associated proteins, such as ELYS, are required specifically for postmitotic assembly of NPCs (Franz et al. 2007). Recent studies have found several discrete roles for the reticulon (Rtn) family of membrane-associated proteins in NE disassembly and reassembly during open mitosis. The Rtns

are critical for regulating transitions between the endoplasmic reticulum tubular network and the NE sheet (Anderson and Hetzer 2008). However, during interphase, an Rtn1 pool at the NE facilitates de novo NPC assembly (Dawson et al. 2009). This most likely is to provide a transient stabilizer of the highly convex region of the pore membrane during early steps in its formation. It is likely that additional non-NPC accessory factors will continue to be revealed, and this is an active area of research that has been well covered in several recent reviews (Antonin et al. 2008; D’Angelo and Hetzer 2008; Kutay and Hetzer 2008; Lim et al. 2008). Even after its assembly, many of the interactions holding the bulk of the NPC together are actually surprisingly transient. Upon completion of NPC construction, some Nups remain stably associated, others turn over promptly, and some shuttle rapidly between the nucleoplasm and cytoplasm, spending little time on the NPC itself (Rabut et al. 2004; Dilworth et al. 2001; Dilworth et al. 2005). For example, there are multiple roles for Nups in kinetochore function and mitosis, as well as in the maintenance of the boundaries between silent and active chromatin (Roux and Burke 2006; Kalverda et al. 2008; Lim et al. 2008). Despite these processes requiring Nups, none appear to occur at the NPC itself. Indeed, recent work underscores how a great number of Nup-chromatin interactions occur within the nuclear interior and away from the NPC itself; these interactions have also been shown to be crucial in controlling gene expression during cell differentiation in higher eukaryotes (Capelson et al. 2010; Kalverda et al. 2010). Conversely, the NPC can act as a reservoir for cell cycle proteins when they are inactive. One example, the Mad proteins, are docked to the NPC in interphase but are released when they function in mitosis (Iouket al. 2002; Scott et al. 2005; Lee et al. 2008). The NPC also plays an important role in the organization of internal nuclear architecture (Fig. 3) and is used as an attachment site for such nuclear peripheral proteins as lamins and yMlp/vTpr. An exciting recent discovery is that certain inducible genes carry specific sequences

S.R. Wente and M.P. Rout

that can cause them to associate with the NPC’s vicinity upon transcriptional activation; once there, they retain a “memory” of their active state for some time after they cease transcription, leaving the genes poised for rapid reactivation (Ahmed et al. 2010). This tethering of induced genes to the NPC might also be also aided by factors required for posttranscriptional mRNP processing and export (Dieppois et al. 2006; Chekanova et al. 2008; Rougemaille et al. 2008). CONCLUSIONS

Our structural and functional understanding of the nuclear transport machinery has blossomed in the past decade, revealing new and unexpected aspects about the NPC and its interacting transport factors. Perhaps most surprising has been the discovery of the evolutionary connections between the NPC and the vesicular trafficking apparatus, the unusual nature of the FG Nup-mediated selective barrier, and the increasing range of connectivity between the NPC and global aspects of cellular physiology and gene expression. Yet as is so often the case, the solving of one question has raised many others. Many of the unanswered questions regarding the mechanisms and regulation of the nuclear transport machinery present exciting challenges for the years ahead. ACKNOWLEDGMENTS

We acknowledge that many of the primary references for critical studies are included in reviews cited, and appreciate our colleagues’ consideration in this limitation. We thank Renee Dawson and Kelli Mullin for assistance with manuscript preparation. This work was supported by funding from the National Institutes of Health (R01 GM51219 to S. R. W. and R01 GM062427 and R01 GM071329 to M. P. R.). REFERENCES Ahmed S, Brickner JH. 2007. Regulation and epigenetic control of transcription at the nuclear periphery. Trends Genet 23: 396 –402. Ahmed S, Brickner DG, Light WH, Cajigas I, McDonough M, Froyshteter AB, Volpe T, Brickner JH. 2010. DNA zip codes control an ancient mechanism for gene

targeting to the nuclear periphery. Nat Cell Biol 12: 111 –118. Aitchison JD, Rout MP, Marelli M, Blobel G, Wozniak RW. 1995. Two novel related yeast nucleoporins Nup170p and Nup157p: Complementation with the vertebrate homologue Nup155p and functional interactions with the yeast nuclear pore-membrane protein Pom152p. J Cell Biol 131: 1133–1148. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007a. Determining the architectures of macromolecular assemblies. Nature 450: 683 –694. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007b. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Alcazar-Roman AR, Tran EJ, Guo S, Wente SR. 2006. Inositol hexakisphosphate and Gle1 activate the DEAD-box protein Dbp5 for nuclear mRNA export. Nat Cell Biol 8: 711– 716. Allen NP, Huang L, Burlingame A, Rexach M. 2001. Proteomic analysis of nucleoporin interacting proteins. J Biol Chem 276: 29268– 29274. Anderson DJ, Hetzer MW. 2008. Reshaping of the endoplasmic reticulum limits the rate for nuclear envelope formation. J Cell Biol 182: 911– 924. Antonin W, Mattaj IW. 2005. Nuclear pore complexes: Round the bend? Nat Cell Biol 7: 10– 12. Antonin W, Ellenberg J, Dultz E. 2008. Nuclear pore complex assembly through the cell cycle: Regulation and membrane organization. FEBS Lett 582: 2004– 2016. Arts GJ, Kuersten S, Romby P, Ehresmann B, Mattaj IW. 1998. The role of exportin-t in selective nuclear export of mature tRNAs. EMBO J 17: 7430– 7441. Ball JR, Dimaano C, Ullman KS. 2004. The RNA binding domain within the nucleoporin Nup153 associates preferentially with single-stranded RNA. RNA 10: 19– 27. Bayliss R, Leung SW, Baker RP, Quimby BB, Corbett AH, Stewart M. 2002. Structural basis for the interaction between NTF2 and nucleoporin FxFG repeats. EMBO J 21: 2843–2853. Belgareh N, Rabut G, Bai SW, van Overbeek M, Beaudouin J, Daigle N, Zatsepina OV, Pasteau F, Labas V, FromontRacine M, et al. 2001. An evolutionarily conserved NPC subcomplex, which redistributes in part to kinetochores in mammalian cells. J Cell Biol 154: 1147–1160. Berke IC, Boehmer T, Blobel G, Schwartz TU. 2004. Structural and functional analysis of Nup133 domains reveals modular building blocks of the nuclear pore complex. J Cell Biol 167: 591– 597. Bradatsch B, Katahira J, Kowalinski E, Bange G, Yao W, Sekimoto T, Baumgartel V, Boese G, Bassler J, Wild K, et al. 2007. Arx1 Functions as an Unorthodox Nuclear Export Receptor for the 60S Preribosomal Subunit. Mol Cell 27: 767–779. Brohawn SG, Leksa NC, Spear ED, Rajashankar KR, Schwartz TU. 2008. Structural evidence for common ancestry of the nuclear pore complex and vesicle coats. Science 322: 1369– 1373.

Nuclear Pore Complexes and Nuclear Transport Brohawn SG, Partridge JR, Whittle JR, Schwartz TU. 2009. The nuclear pore complex has entered the atomic age. Structure 17: 1156– 1168. Capelson M, Liang Y, Schulte R, Mair W, Wagner U, Hetzer MW. 2010. Chromatin-bound nuclear pore components regulate gene expression in higher eukaryotes. Cell 140: 372–383. Carmody SR, Wente SR. 2009. mRNA nuclear export at a glance. J Cell Sci 122: 1933– 1937. Chekanova JA, Abruzzi KC, Rosbash M, Belostotsky DA. 2008. Sus1, Sac3, and Thp1 mediate post-transcriptional tethering of active genes to the nuclear rim as well as to non-nascent mRNP. RNA 14: 66–77. Chi NC, Adam SA. 1997. Functional domains in nuclear import factor p97 for binding the nuclear localization sequence receptor and the nuclear pore. Mol Biol Cell 8: 945–956. Cole CN, Scarcelli JJ. 2006. Transport of messenger RNA from the nucleus to the cytoplasm. Curr Opin Cell Biol 18: 299 –306. Conti E, Izaurralde E. 2001. Nucleocytoplasmic transport enters the atomic age. Curr Opin Cell Biol 13: 310–319. Cook A, Bono F, Jinek M, Conti E. 2007. Structural biology of nucleocytoplasmic transport. Annu Rev Biochem 76: 647–671. Cronshaw JM, Krutchinsky AN, Zhang W, Chait BT, Matunis MJ. 2002. Proteomic analysis of the mammalian nuclear pore complex. J Cell Biol 158: 915– 927. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456–466. D’Angelo MA, Anderson DJ, Richard E, Hetzer MW. 2006. Nuclear pores form de novo from both sides of the nuclear envelope. Science 312: 440–443. Daneholt B. 2001. Assembly and transport of a premessenger RNP particle. Proc Natl Acad Sci 98: 7012– 7017. Dawson TR, Lazarus MD, Hetzer MW, Wente SR. 2009. ER membrane-bending proteins are necessary for de novo nuclear pore formation. J Cell Biol 184: 659–675. Debler EW, Ma Y, Seo HS, Hsia KC, Noriega TR, Blobel G, Hoelz A. 2008. A fence-like coat for the nuclear pore membrane. Mol Cell 32: 815 –826. DeGrasse JA, DuBois KN, Devos D, Siegel TN, Sali A, Field MC, Rout MP, Chait BT. 2009. Evidence for a shared nuclear pore complex architecture that is conserved from the last common eukaryotic ancestor. Mol Cell Proteomics 8: 2119–2130. Denning D, Mykytka B, Allen NP, Huang L, Al B, Rexach M. 2001. The nucleoporin Nup60p functions as a Gsp1p-GTP-sensitive tether for Nup2p at the nuclear pore complex. J Cell Biol 154: 937–950. Denning DP, Patel SS, Uversky V, Fink AL, Rexach M. 2003. Disorder in the nuclear pore complex: The FG repeat regions of nucleoporins are natively unfolded. Proc Natl Acad Sci 100: 2450–2455. Devos D, Dokudovskaya S, Alber F, Williams R, Chait BT, Sali A, Rout MP. 2004. Components of coated vesicles and nuclear pore complexes share a common molecular architecture. PLoS Biol 2: e380. Devos D, Dokudovskaya S, Williams R, Alber F, Eswar N, Chait BT, Rout MP, Sali A. 2006. Simple fold composition

and modular architecture of the nuclear pore complex. Proc Natl Acad Sci 103: 2172– 2177. Dieppois G, Iglesias N, Stutz F. 2006. Cotranscriptional recruitment to the mRNA export receptor Mex67p contributes to nuclear pore anchoring of activated genes. Mol Cell Biol 26: 7858– 7870. Dilworth DJ, Suprapto A, Padovan JC, Chait BT, Wozniak RW, Rout MP, Aitchison JD. 2001. Nup2p dynamically associates with the distal regions of the yeast nuclear pore complex. J Cell Biol 153: 1465–1478. Dilworth DJ, Tackett AJ, Rogers RS, Yi EC, Christmas RH, Smith JJ, Siegel AF, Chait BT, Wozniak RW, Aitchison JD. 2005. The mobile nucleoporin Nup2p and chromatin-bound Prp20p function in endogenous NPC-mediated transcriptional control. J Cell Biol 171: 955 –965. Dingwall C, Robbins J, Dilworth SM, Roberts B, Richardson WD. 1988. The nucleoplasmin nuclear location sequence is larger and more complex than that of SV-40 large T antigen. J Cell Biol 107: 841 –849. Erkmann JA, Kutay U. 2004. Nuclear export of mRNA: From the site of transcription to the cytoplasm. Exp Cell Res 296: 12–20. Fagotto F, Gluck U, Gumbiner BM. 1998. Nuclear localization signal-independent and importin/karyopherinindependent nuclear import of b-catenin. Curr Biol 8: 181 –190. Fairman ME, Maroney PA, Wang W, Bowers HA, Gollnick P, Nilsen TW, Jankowsky E. 2004. Protein displacement by DExH/D "RNA helicases" without duplex unwinding. Science 304: 730–734. Fan JS, Cheng Z, Zhang J, Noble C, Zhou Z, Song H, Yang D. 2009. Solution and crystal structures of mRNA exporter Dbp5p and its interaction with nucleotides. J Mol Biol 388: 1– 10. Feldherr CM, Akin D. 1997. The location of the transport gate in the nuclear pore complex. J Cell Sci 110: 3065–3070. Fernandez-Martinez J, Rout MP. 2009. Nuclear pore complex biogenesis. Curr Opin Cell Biol 21: 603 –612. Field MC, Dacks JB. 2009. First and last ancestors: Reconstructing evolution of the endomembrane system with ESCRTs, vesicle coat proteins, and nuclear pore complexes. Curr Opin Cell Biol 21: 4 –13. Floer M, Blobel G. 1996. The nuclear transport factor karyopherin b binds stoichiometrically to Ran-GTP and inhibits the Ran GTPase activating protein. J Biol Chem 271: 5313–5316. Floer M, Blobel G, Rexach M. 1997. Disassembly of RanGTP-karyopherin b complex, an intermediate in nuclear protein import. J Biol Chem 272: 19538– 19546. Fontoura BM, Blobel G, Matunis MJ. 1999. A conserved biogenesis pathway for nucleoporins: Proteolytic processing of a 186-kilodalton precursor generates Nup98 and the novel nucleoporin, Nup96. J Cell Biol 144: 1097–1112. Forler D, Rabut G, Ciccarelli FD, Herold A, Kocher T, Niggeweg R, Bork P, Ellenberg J, Izaurralde E. 2004. RanBP2/Nup358 provides a major binding site for NXF1-p15 dimers at the nuclear pore complex and functions in nuclear mRNA export. Mol Cell Biol 24: 1155–1167.

S.R. Wente and M.P. Rout Fornerod M, Ohno M, Yoshida M, Mattaj IW. 1997. CRM1 is an export receptor for leucine-rich nuclear export signals. Cell 90: 1051–1060. Franz C, Walczak R, Yavuz S, Santarella R, Gentzel M, Askjaer P, Galy V, Hetzer M, Mattaj IW, Antonin W. 2007. MEL-28/ELYS is required for the recruitment of nucleoporins to chromatin and postmitotic nuclear pore complex assembly. EMBO Rep 8: 165 –172. Frey S, Gorlich D. 2007. A saturated FG-repeat hydrogel can reproduce the permeability properties of nuclear pore complexes. Cell 130: 512 –523. Frey S, Richter RP, Gorlich D. 2006. FG-rich repeats of nuclear pore proteins form a three-dimensional meshwork with hydrogel-like properties. Science 314: 815– 817. Fried H, Kutay U. 2003. Nucleocytoplasmic transport: Taking an inventory. Cell Mol Life Sci 60: 1659– 1688. Fukuda M, Asano S, Nakamura T, Adachi M, Yoshida M, Yanagida M, Nishida E. 1997. CRM1 is responsible for intracellular transport mediated by the nuclear export signal. Nature 390: 308 –311. Goldfarb DS, Gariepy J, Schoolnik G, Kornberg RD. 1986. Synthetic peptides as nuclear localization signals. Nature 322: 641– 644. Gorlich D, Dabrowski M, Bischoff FR, Kutay U, Bork P, Hartmann E, Prehn S, Izaurralde E. 1997. A novel class of RanGTP binding proteins. J Cell Biol 138: 65–80. Grandi P, Dang T, Pane N, Shevchenko A, Mann M, Forbes D, Hurt E. 1997. Nup93, a vertebrate homologue of yeast Nic96p, forms a complex with a novel 205-kDa protein and is required for correct nuclear pore assembly. Mol Biol Cell 8: 2017– 2038. Grant RP, Neuhaus D, Stewart M. 2003. Structural basis for the interaction between the Tap/NXF1 UBA domain and FG nucleoporins at 1A resolution. J Mol Biol 326: 849–858. Hang J, Dasso M. 2002. Association of the human SUMO-1 protease SENP2 with the nuclear pore. J Biol Chem 277: 19961–19966. Harel A, Forbes DJ. 2004. Importin b: Conducting a much larger cellular symphony. Mol Cell 16: 319 –330. Hellmuth K, Lau DM, Bischoff FR, Kunzler M, Hurt E, Simos G. 1998. Yeast Los1p has properties of an exportinlike nucleocytoplasmic transport factor for tRNA. Mol Cell Biol 18: 6374– 6386. Henn A, Cao W, Hackney DD, De La Cruz EM. 2008. The ATPase cycle mechanism of the DEAD-box rRNA helicase, DbpA. J Mol Biol 377: 193 –205. Henn A, Shi SP, Zarivach R, Ben-Zeev E, Sagi I. 2002. The RNA helicase DbpA exhibits a markedly different conformation in the ADP-bound state when compared with the ATP- or RNA-bound states. J Biol Chem 277: 46559–46565. Hetzer MW, Wente SR. 2009. Border control at the nucleus: Biogenesis and organization of the nuclear membrane and pore complexes. Dev Cell 17: 606– 616. Hetzer MW, Walther TC, Mattaj IW. 2005. Pushing the envelope: Structure, function, and dynamics of the nuclear periphery. Annu Rev Cell Dev Biol 21: 347– 380.

Higa MM, Alam SL, Sundquist WI, Ullman KS. 2007. Molecular characterization of the Ran-binding zinc finger domain of Nup153. J Biol Chem 282: 17090–17100. Hodge CA, Colot HV, Stafford P, Cole CN. 1999. Rat8p/ Dbp5p is a shuttling transport factor that interacts with Rat7p/Nup159p and Gle1p and suppresses the mRNA export defect of xpo1–1 cells. EMBO J 18: 5778– 5788. Hsia KC, Stavropoulos P, Blobel G, Hoelz A. 2007. Architecture of a coat for the nuclear pore membrane. Cell 131: 1313–1326. Iouk T, Kerscher O, Scott RJ, Basrai MA, Wozniak RW. 2002. The yeast nuclear pore complex functionally interacts with components of the spindle assembly checkpoint. J Cell Biol 159: 807– 819. Isgro TA, Schulten K. 2005. Binding dynamics of isolated nucleoporin repeat regions to importin-b. Structure 13: 1869–1879. Jankowsky E, Gross CH, Shuman S, Pyle AM. 2001. Active disruption of an RNA-protein interactionby a DExH/D RNA helicase. Science 291: 121–125. Johnson AW, Lund E, Dahlberg J. 2002. Nuclear export of ribosomal subunits. Trends Biochem Sci 27: 580– 585. Joseph J, Dasso M. 2008. The nucleoporin Nup358 associates with and regulates interphase microtubules. FEBS Lett 582: 190– 196. Joseph J, Liu ST, Jablonski SA, Yen TJ, Dasso M. 2004. The RanGAP1-RanBP2 complex is essential for microtubulekinetochore interactions in vivo. Curr Biol 14: 611– 617. Jovanovic-Talisman T, Tetenbaum-Novatt J, McKenney AS, Zilman A, Peters R, Rout MP, Chait BT. 2009. Artificial nanopores that mimic the transport selectivity of the nuclear pore complex. Nature 457: 1023–1027. Kalab P, Weis K, Heald R. 2002. Visualization of a Ran-GTP gradient in interphase and mitotic Xenopus egg extracts. Science 295: 2452– 2456. Kalverda B, Roling MD, Fornerod M. 2008. Chromatin organization in relation to the nuclear periphery. FEBS Lett 582: 2017–2022. Kalverda B, Pickersgill H, Shloma VV, Fornerod M. 2010. Nucleoporins Directly Stimulate Expression of Developmental and Cell-Cycle Genes Inside the Nucleoplasm. Cell 140: 360 –371. Keminer O, Peters R. 1999. Permeability of single nuclear pores. Biophys J 77: 217– 228. Krishnan VV, Lau EY, Yamada J, Denning DP, Patel SS, Colvin ME, Rexach MF. 2008. Intramolecular cohesion of coils mediated by phenylalanine–glycine motifs in the natively unfolded domain of a nucleoporin. PLoS Comput Biol 4: e1000145. Kubitscheck U, Grunwald D, Hoekstra A, Rohleder D, Kues T, Siebrasse JP, Peters R. 2005. Nuclear transport of single molecules: Dwell times at the nuclear pore complex. J Cell Biol 168: 233 –243. Kutay U, Hetzer MW. 2008. Reorganization of the nuclear envelope during open mitosis. Curr Opin Cell Biol 20: 669 –677. Kutay U, Bischoff FR, Kostka S, Kraft R, Gorlich D. 1997a. Export of importin a from the nucleus is mediated by a specific nuclear transport factor. Cell 90: 1061–1071. Kutay U, Izaurralde E, Bischoff FR, Mattaj IW, Gorlich D. 1997b. Dominant-negative mutants of importin-b block

Nuclear Pore Complexes and Nuclear Transport multiple pathways of import and export through the nuclear pore complex. EMBO J 16: 1153–1163. Kutay U, Lipowsky G, Izaurralde E, Bischoff FR, Schwarzmaier P, Hartmann E, Gorlich D. 1998. Identification of a tRNA-specific nuclear export receptor. Mol Cell 1: 359–369. Lee BJ, Cansizoglu AE, Suel KE, Louis TH, Zhang Z, Chook YM. 2006. Rules for nuclear localization sequence recognition by karyopherin b 2. Cell 126: 543– 558. Lee SH, Sterling H, Burlingame A, McCormick F. 2008. Tpr directly binds to Mad1 and Mad2 and is important for the Mad1-Mad2-mediated mitotic spindle checkpoint. Genes Dev 22: 2926–2931. Leksa NC, Brohawn SG, Schwartz TU. 2009. The structure of the scaffold nucleoporin Nup120 reveals a new and unexpected domain architecture. Structure 17: 1082– 1091. Lewis A, Felberbaum R, Hochstrasser M. 2007. A nuclear envelope protein linking nuclear pore basket assembly, SUMO protease regulation, and mRNA surveillance. J Cell Biol 178: 813 –827. Lim RY, Fahrenkrog B, Koser J, Schwarz-Herion K, Deng J, Aebi U. 2007a. Nanomechanical basis of selective gating by the nuclear pore complex. Science 318: 640– 643. Lim RY, Huang NP, Koser J, Deng J, Lau KH, SchwarzHerion K, Fahrenkrog B, Aebi U. 2006. Flexible phenylalanine-glycine nucleoporins as entropic barriers to nucleocytoplasmic transport. Proc Natl Acad Sci 103: 9512– 9517. Lim RY, Koser J, Huang NP, Schwarz-Herion K, Aebi U. 2007b. Nanomechanical interactions of phenylalanineglycine nucleoporins studied by single molecule forcevolume spectroscopy. J Struct Biol 159: 277 –289. Lim RY, Ullman KS, Fahrenkrog B. 2008. Biology and biophysics of the nuclear pore complex and its components. Int Rev Cell Mol Biol 267: 299–342. Lipowsky G, Bischoff FR, Izaurralde E, Kutay U, Schafer S, Gross HJ, Beier H, Gorlich D. 1999. Coordination of tRNA nuclear export with processing of tRNA. Rna 5: 539–549. Liu SM, Stewart M. 2005. Structural basis for the highaffinity binding of nucleoporin Nup1p to the Saccharomyces cerevisiae importin-b homologue, Kap95p. J Mol Biol 349: 515–525. Liu HL, De Souza CP, Osmani AH, Osmani SA. 2009. The three fungal transmembrane nuclear pore complex proteins of Aspergillus nidulans are dispensable in the presence of an intact An-Nup84–120 complex. Mol Biol Cell 20: 616 –630. Liu J, Prunuske AJ, Fager AM, Ullman KS. 2003. The COPI complex functions in nuclear envelope breakdown and is recruited by the nucleoporin Nup153. Dev Cell 5: 487–498. Lyman SK, Guan T, Bednenko J, Wodrich H, Gerace L. 2002. Influence of cargo size on Ran and energy requirements for nuclear protein import. J Cell Biol 159: 55– 67. Macara IG. 2001. Transport into and out of the nucleus. Microbiol Mol Biol Rev 65: 570– 594. Madrid AS, Weis K. 2006. Nuclear transport is becoming crystal clear. Chromosoma 115: 98–109. Mahajan R, Delphin C, Guan T, Gerace L, Melchior F. 1997. A small ubiquitin-related polypeptide involved in

targeting RanGAP1 to nuclear pore complex protein RanBP2. Cell 88: 97–107. Mattaj IW, Englmeier L. 1998. Nucleocytoplasmic transport: The soluble phase. Annu Rev Biochem 67: 265– 306. Matunis MJ, Wu J, Blobel G. 1998. SUMO-1 modification and its role in targeting the Ran GTPase-activating protein, RanGAP1, to the nuclear pore complex. J Cell Biol 140: 499–509. Mehlin H, Daneholt B, Skoglund U. 1992. Translocation of a specific premessenger ribonucleoprotein particle through the nuclear pore studied with electron microscope tomography. Cell 69: 605– 613. Miller BR, Powers M, Park M, Fischer W, Forbes DJ. 2000. Identification of a new vertebrate nucleoporin, Nup188, with the use of a novel organelle trap assay. Mol Biol Cell 11: 3381– 3396. Monecke T, Guttler T, Neumann P, Dickmanns A, Gorlich D, Ficner R. 2009. Crystal structure of the nuclear export receptor CRM1 in complex with Snurportin1 and RanGTP. Science 324: 1087–1091. Moroianu J, Blobel G, Radu A. 1997. RanGTP-mediated nuclear export of karyopherin a involves its interaction with the nucleoporin Nup153. Proc Natl Acad Sci 94: 9699–9704. Murphy R, Wente SR. 1996. An RNA-export mediator with an essential nuclear export signal. Nature 383: 357– 360. Nachury MV, Weis K. 1999. The direction of transport through the nuclear pore can be inverted. Proc Natl Acad Sci 96: 9622–9627. Nakielny S, Shaikh S, Burke B, Dreyfuss G. 1999. Nup153 is an M9-containing mobile nucleoporin with a novel Ran-binding domain. EMBO J 18: 1982–1995. Nehrbass U, Rout MP, Maguire S, Blobel G, Wozniak RW. 1996. The yeast nucleoporin Nup188p interacts genetically and physically with the core structures of the nuclear pore complex. J Cell Biol 133: 1153–1162. Oeffinger M, Dlakic M, Tollervey D. 2004. A pre-ribosomeassociated HEAT-repeat protein is required for export of both ribosomal subunits. Genes Dev 18: 196 –209. Ohno M, Segref A, Bachi A, Wilm M, Mattaj IW. 2000. PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101: 187 –198. Pante N, Kann M. 2002. Nuclear pore complex is able to transport macromolecules with diameters of about 39 nm. Mol Biol Cell 13: 425– 434. Paraskeva E, Izaurralde E, Bischoff FR, Huber J, Kutay U, Hartmann E, Luhrmann R, Gorlich D. 1999. CRM1mediated recycling of snurportin 1 to the cytoplasm. J Cell Biol 145: 255– 264. Patel SS, Belmont BJ, Sante JM, Rexach MF. 2007. Natively unfolded nucleoporins gate protein diffusion across the nuclear pore complex. Cell 129: 83–96. Pemberton LF, Paschal BM. 2005. Mechanisms of receptormediated nuclear import and nuclear export. Traffic 6: 187 –198. Peters R. 2005. Translocation through the nuclear pore complex: Selectivity and speed by reduction-of-dimensionality. Traffic 6: 421– 427.

S.R. Wente and M.P. Rout Pichler A, Gast A, Seeler JS, Dejean A, Melchior F. 2002. The nucleoporin RanBP2 has SUMO1 E3 ligase activity. Cell 108: 109– 120. Poon IK, Jans DA. 2005. Regulation of nuclear transport: Central role in development and transformation? Traffic 6: 173 –186. Rabut G, Doye V, Ellenberg J. 2004. Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nat Cell Biol 6: 1114– 1121. Radu A, Moore MS, Blobel G. 1995. The peptide repeat domain of nucleoporin Nup98 functions as a docking site in transport across the nuclear pore complex. Cell 81: 215 –222. Reverter D, Lima CD. 2005. Insights into E3 ligase activity revealed by a SUMO-RanGAP1-Ubc9-Nup358 complex. Nature 435: 687–692. Rexach M, Blobel G. 1995. Protein import into nuclei: Association and dissociation reactions involving transport substrate, transport factors, and nucleoporins. Cell 83: 683–692. Ribbeck K, Gorlich D. 2002. The permeability barrier of nuclear pore complexes appears to operate via hydrophobic exclusion. EMBO J 21: 2664– 2671. Rodriguez MS, Dargemont C, Stutz F. 2004. Nuclear export of RNA. Biol Cell 96: 639–655. Rougemaille M, Dieppois G, Kisseleva-Romanova E, Gudipati RK, Lemoine S, Blugeon C, Boulay J, Jensen TH, Stutz F, Devaux F, et al. 2008. THO/Sub2p functions to coordinate 30 -end processing with gene-nuclear pore association. Cell 135: 308 –321. Rout MP, Wente SR. 1994. Pores for thought: Nuclear pore complex proteins. Trends Cell Biol 4: 357– 365. Rout MP, Aitchison JD. 2001. The nuclear pore complex as a transport machine. J Biol Chem 276: 16593– 16596. Rout MP, Aitchison JD, Magnasco MO, Chait BT. 2003. Virtual gating and nuclear transport: The hole picture. Trends Cell Biol 13: 622–628. Rout MP, Aitchison JD, Suprapto A, Hjertaas K, Zhao Y, Chait BT. 2000. The yeast nuclear pore complex: Composition, architecture, and transport mechanism. J Cell Biol 148: 635– 651. Roux KJ, Burke B. 2006. From pore to kinetochore and back: Regulating envelope assembly. Dev Cell 11: 276– 278. Salina D, Enarson P, Rattner JB, Burke B. 2003. Nup358 integrates nuclear envelope breakdown with kinetochore assembly. J Cell Biol 162: 991– 1001. Scarcelli JJ, Hodge CA, Cole CN. 2007. The yeast integral membrane protein Apq12 potentially links membrane dynamics to assembly of nuclear pore complexes. J Cell Biol 178: 799–812. Schmitt C, von Kobbe C, Bachi A, Pante N, Rodrigues JP, Boscheron C, Rigaut G, Wilm M, Seraphin B, CarmoFonseca M, et al. 1999. Dbp5, a DEAD-box protein required for mRNA export, is recruited to the cytoplasmic fibrils of nuclear pore complex via a conserved interaction with CAN/Nup159p. EMBO J 18: 4332–4347. Schneiter R, Hitomi M, Ivessa AS, Fasch EV, Kohlwein SD, Tartakoff AM. 1996. A yeast acetyl coenzyme A carboxylase mutant links very-long-chain fatty acid synthesis to the structure and function of the nuclear membrane-pore complex. Mol Cell Biol 16: 7161–7172.

Scott RJ, Lusk CP, Dilworth DJ, Aitchison JD, Wozniak RW. 2005. Interactions between Mad1p and the nuclear transport machinery in the yeast Saccharomyces cerevisiae. Mol Biol Cell 16: 4362– 4374. Seo HS, Ma Y, Debler EW, Wacker D, Kutik S, Blobel G, Hoelz A. 2009. Structural and functional analysis of Nup120 suggests ring formation of the Nup84 complex. Proc Natl Acad Sci 106: 14281–14286. Shulga N, Goldfarb DS. 2003. Binding dynamics of structural nucleoporins govern nuclear pore complex permeability and may mediate channel gating. Mol Cell Biol 23: 534 –542. Shulga N, Mosammaparast N, Wozniak R, Goldfarb DS. 2000. Yeast nucleoporins involved in passive nuclear envelope permeability. J Cell Biol 149: 1027–1038. Simos G, Grosshans H, Hurt E. 2002. Nuclear export of tRNA. Results Probl Cell Differ 35: 115– 131. Siniossoglou S, Lutzmann M, Santos-Rosa H, Leonard K, Mueller S, Aebi U, Hurt E. 2000. Structure and assembly of the Nup84p complex. J Cell Biol 149: 41– 54. Siniossoglou S, Wimmer C, Rieger M, Doye V, Tekotte H, Weise C, Emig S, Segref A, Hurt EC. 1996. A novel complex of nucleoporins, which includes Sec13p and a Sec13p homolog, is essential for normal nuclear pores. Cell 84: 265–275. Snay-Hodge CA, Colot HV, Goldstein AL, Cole CN. 1998. Dbp5p/Rat8p is a yeast nuclear pore-associated DEADbox protein essential for RNA export. EMBO J 17: 2663–2676. Stade K, Ford CS, Guthrie C, Weis K. 1997. Exportin 1 (Crm1p) is an essential nuclear export factor. Cell 90: 1041–1050. Stewart M. 2007. Ratcheting mRNA out of the nucleus. Mol Cell 25: 327–330. Strahm Y, Fahrenkrog B, Zenklusen D, Rychner E, Kantor J, Rosbach M, Stutz F. 1999. The RNA export factor Gle1p is located on the cytoplasmic fibrils of the NPC and physically interacts with the FG-nucleoporin Rip1p, the DEAD-box protein Rat8p/Dbp5p and a new protein Ymr 255p. EMBO J 18: 5761– 5777. Strawn LA, Shen T, Wente SR. 2001. The GLFG regions of Nup116p and Nup100p serve as binding sites for both Kap95p and Mex67p at the nuclear pore complex. J Biol Chem 276: 6445– 6452. Strawn LA, Shen T, Shulga N, Goldfarb DS, Wente SR. 2004. Minimal nuclear pore complexes define FG repeat domains essential for transport. Nat Cell Biol 6: 197– 206. Terry LJ, Wente SR. 2007. Nuclear mRNA export requires specific FG nucleoporins for translocation through the nuclear pore complex. J Cell Biol 178: 1121– 1132. Terry LJ, Shows EB, Wente SR. 2007. Crossing the nuclear envelope: Hierarchical regulation of nucleocytoplasmic transport. Science 318: 1412– 1416. Timney BL, Tetenbaum-Novatt J, Agate DS, Williams R, Zhang W, Chait BT, Rout MP. 2006. Simple kinetic relationships and nonspecific competition govern nuclear import rates in vivo. J Cell Biol 175: 579– 593. Tran EJ, Wente SR. 2006. Dynamic nuclear pore complexes: Life on the edge. Cell 125: 1041– 1053. Tran EJ, Bolger TA, Wente SR. 2007a. SnapShot: Nuclear transport. Cell 131: 420.

Nuclear Pore Complexes and Nuclear Transport Tran EJ, Zhou Y, Corbett AH, Wente SR. 2007b. The DEADbox protein Dbp5 controls mRNA export by triggering specific RNA:Protein remodeling events. Mol Cell 28: 850–859. Tseng SS, Weaver PL, Liu Y, Hitomi M, Tartakoff AM, Chang TH. 1998. Dbp5p, a cytosolic RNA helicase, is required for poly(A)þ RNA export. EMBO J 17: 2651–2662. Vasu S, Shah S, Orjalo A, Park M, Fischer WH, Forbes DJ. 2001. Novel vertebrate nucleoporins Nup133 and Nup160 play a role in mRNA export. J Cell Biol 155: 339–354. Weirich CS, Erzberger JP, Flick JS, Berger JM, Thorner J, Weis K. 2006. Activation of the DExD/H-box protein Dbp5 by the nuclear-pore protein Gle1 and its coactivator InsP6 is required for mRNA export. Nat Cell Biol 8: 668–676. Weis K. 2003. Regulating Access to the Genome. Nucleocytoplasmic Transport throughout the Cell Cycle. Cell 112: 441–451. Wente SR. 2000. Gatekeepers of the nucleus. Science 288: 1374– 1377. Whittle JR, Schwartz TU. 2009. Architectural nucleoporins Nup157/170 and Nup133 are structurally related and descend from a second ancestral element. J Biol Chem Wu J, Matunis MJ, Kraemer D, Blobel G, Coutavas E. 1995. Nup358, a cytoplasmically exposed nucleoporin with peptide repeats, Ran-GTP binding sites, zinc fingers, a cyclophilin A homologous domain, and a leucine-rich region. J Biol Chem 270: 14209–14213. Xu XM, Meulia T, Meier I. 2007. Anchorage of plant RanGAP to the nuclear envelope involves novel nuclearpore-associated proteins. Curr Biol 17: 1157–1163. Xu Z, Lam LS, Lam LH, Chau SF, Ng TB, Au SW. 2008. Molecular basis of the redox regulation of SUMO proteases: A protective mechanism of intermolecular disulfide linkage against irreversible sulfhydryl oxidation. FASEB J 22: 127–137. Yang W, Musser SM. 2006. Nuclear import time and transport efficiency depend on importin b concentration. J Cell Biol 174: 951 –961.

Yang Q, Jankowsky E. 2006. The DEAD-box protein Ded1 unwinds RNA duplexes by a mode distinct from translocating helicases. Nat Struct Mol Biol 13: 981–986. Yang W, Gelles J, Musser SM. 2004. Imaging of singlemolecule translocation through nuclear pore complexes. Proc Natl Acad Sci 101: 12887–12892. Yao W, Roser D, Kohler A, Bradatsch B, Bassler J, Hurt E. 2007. Nuclear export of ribosomal 60S subunits by the general mRNA export receptor Mex67-Mtr2. Mol Cell 26: 51–62. Yi H, Friedman JL, Ferreira PA. 2007. The cyclophilin-like domain of Ran-binding protein-2 modulates selectively the activity of the ubiquitin-proteasome system and protein biogenesis. J Biol Chem 282: 34770– 34778. Yokoyama N, Hayashi N, Seki T, Pante N, Ohba T, Nishii K, Kuma K, Hayashida T, Miyata T, Aebi U, et al. 1995. A giant nucleopore protein that binds Ran/TC4. Nature 376: 184–188. York JD, Odom AR, Murphy R, Ives EB, Wente SR. 1999. A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science 285: 96– 100. Yoshida K, Blobel G. 2001. The karyopherin Kap142p/ Msn5p mediates nuclear import and nuclear export of different cargo proteins. J Cell Biol 152: 729–740. Zeitler B, Weis K. 2004. The FG-repeat asymmetry of the nuclear pore complex is dispensible for bulk nucleocytoplasmic transport in vivo. J Cell Biol 167: 583–590. Zemp I, Kutay U. 2007. Nuclear export and cytoplasmic maturation of ribosomal subunits. FEBS Lett 581: 2783–2793. Zhang H, Saitoh H, Matunis MJ. 2002. Enzymes of the SUMO modification pathway localize to filaments of the nuclear pore complex. Mol Cell Biol 22: 6498–6508. Zhao Q, Brkljacic J, Meier I. 2008. Two distinct interacting classes of nuclear envelope-associated coiled-coil proteins are required for the tissue-specific nuclear envelope targeting of Arabidopsis RanGAP. Plant Cell 20: 1639– 1651. Zilman A, Di Talia S, Chait BT, Rout MP, Magnasco MO. 2007. Efficiency, selectivity, and robustness of nucleocytoplasmic transport. PLoS Comput Biol 3: e125.

Chromosome Territories Thomas Cremer1,2 and Marion Cremer1 1

Biozentrum, Department of Biology II (Chair of Anthropology and Human Genetics), Ludwig-MaximiliansUniversity, Grosshadernerstrasse 2, 82152 Martinsried, Germany

2

Munich Center for Integrated Protein Sciences (CIPSM), 81377 Munich, Germany

Correspondence: [email protected]

Chromosome territories (CTs) constitute a major feature of nuclear architecture. In a brief statement, the possible contribution of nuclear architecture studies to the field of epigenomics is considered, followed by a historical account of the CT concept and the final compelling experimental evidence of a territorial organization of chromosomes in all eukaryotes studied to date. Present knowledge of nonrandom CT arrangements, of the internal CT architecture, and of structural interactions with other CTs is provided as well as the dynamics of CT arrangements during cell cycle and postmitotic terminal differentiation. The article concludes with a discussion of open questions and new experimental strategies to answer them.

mpressive progress has been achieved during the last decade with regard to the functional implications of DNA methylation, histone modifications, and chromatin remodeling events for gene regulation (Fuks 2005; Kouzarides 2007; Maier et al. 2008; Jiang and Pugh 2009). It has, however, also become obvious that decoding the chromatin language does not suffice to fully understand the ways in which the diploid genome contributes to the formation of the different epigenomes present in the various cell types of a multicellular organism. Different epigenomes and their functional implications also depend on differences in higher-order chromatin organization and nuclear architecture at large. Epigenomic research aims

I

for an integrated understanding of the structural and functional aspects of epigenetics with nuclear architecture during the differentiation of toti- or pluripotent cells to functionally distinct cell types. The territorial organization of chromosomes in interphase (chromosome territories, CTs) constitutes a basic feature of nuclear architecture. This article starts with a brief historical account of the CT concept and the compelling experimental evidence in favor of a territorial organization of chromosomes in all eukaryotes studied to date. A survey of what is presently known about nonrandom arrangements of CTs, about changes of such arrangements in cycling cells as a result of internal or external

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a003889 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a003889

T. Cremer and M. Cremer

influences and about the internal architecture of CTs and their structural interactions with each other is provided. The article concludes with a discussion of open questions on CT organization and new experimental strategies to answer them.

ORIGIN OF THE CHROMOSOME TERRITORY CONCEPT

Since the late 19th century, an uncounted number of microscopic studies has appeared on numerous aspects of nuclear structure and on the observation of mitotic chromosomes. A territorial organization of interphase chromosomes was first suggested for animal cell nuclei by Carl Rabl (Rabl 1885) (Fig. 1A,B), but it was Theodor Boveri who introduced the term chromosome territory (CT) in his seminal studies of blastomere stages of the horse roundworm Parascaris equorum or Ascaris megalocephala, as the species was called at his time (Boveri 1909). The worm exists in two varieties, one with two pairs (Ascaris bivalens), the other with one pair of germ line chromosomes (Ascaris univalens). Boveri argued that each chromosome visible in mitosis retains its individuality during interphase and occupies a distinct part of the nuclear space. As can be seen in his drawings, Boveri was able to distinguish chromosome ends sticking out in protrusions of prophase nuclei (Fig. 1C) and used these protrusions as markers for the nuclear position of the asserted CTs in interphase nuclei (Fig. 1D). In fixed two-cell embryos, Boveri noted that their nuclear topography was strikingly similar during interphase and prophase, when chromosomes became visible as distinct entities. In four cell embryos, however, he typically observed two pairs of nuclei each carrying distinctly different protrusion patterns (Fig. 1D). His ingenious speculative talent led him to the following predictions on chromosome arrangements in Ascaris nuclei during the first steps of postzygotic development (for a comprehensive review, see Cremer and Cremer

2006a): (I) CTorder is stably maintained during interphase; (II) Chromosome neighborhood patterns change from prophase to metaphase; and (III) New chromosome neighborhood arrangements established in the metaphase plate are conserved to a considerable extent throughout anaphase and telophase, resulting in rather symmetrical arrangements of CTs in the two daughter nuclei. Different opinions on the structural organization of CTs were put forward in the early days of the 20th century. Eduard Strasburger (1905) published a colored cartoon claiming that CTs are built up from little chromatin clumps (Fig. 1E), whereas Theodor Boveri discussed a sponge-like CT structure built up from networks of anastomizing chromatin bundles (Fig. 1F). Despite light microscopic evidence in favor of chromosome territories at least in some species and cell types (for review see Stack et al. 1977) the weight of electron microscopic evidence established since the 1950s apparently argued for an unraveling of chromosomes in interphase nuclei into intermingling chromatin fibers of 10 – 30 nm in diameter with no sign of individual chromosomes. As a consequence, the concept of chromosome territories fell into oblivion or was even considered to be experimentally disproved (Wischnitzer 1973). During the 1970s and 1980s, most researchers seemed content with the assumption that the nucleus is filled with intermingling chromatin fibers and loops like a dish of spaghetti, an assumption widely reflected by textbooks of cell biology.

EXPERIMENTAL EVIDENCE FOR CHROMOSOME TERRITORIES

Stephen M. Stack, David B. Brown, and William C. Dewey were among the few researchers in the 1970s that still adhered to Boveri’s seemingly outdated concept (Stack et al. 1977; for review, see Cremer and Cremer 2006b). They squashed fixed cells from Allium cepa root tips, as well as Chinese hamster cells in acetic acid, and

Chromosome Territories

subjected the air-dried cells to salt solutions containing NaOH, and finally performed a Giemsa staining. This treatment yielded clumps of condensed chromatin arguably reflecting interphase chromosomes (Fig. 2A). The authors concluded that “chromosomes . . . remain in distinct domains throughout interphase.” Our group obtained early evidence for CTs in nuclei of diploid Chinese hamster cells with the help of laser-UV-microirradiation experiments (Zorn et al. 1976; Zorn et al. 1979; Cremer et al. 1982a; Cremer et al. 1982b; for review, see Cremer and Cremer 2006b; Meaburn and Misteli 2007). A laser microbeam (l ¼ 257 nm) was used to induce UV-damaged DNA within a small part of the nucleus. It was predicted that DNA damage inflicted within a small volume of the nucleus would yield different results depending on how chromosomes were arranged. Figure 2B exemplifies the experimental rationale with the example of woolen threads assembled within a “nuclear” space (Cremer et al. 1982a). Each thread reflects a chromatin fiber constituting an individual chromosome. In case of threads distributed throughout the whole nuclear space (upper panel), the “damage” label would become scattered over many threads. If individual threads occupy distinct territories (lower panel), localized label would mark only a small subset of threads. This experimental rationale was realized in the following way (Fig. 2C – E): Nuclei of living cells were microirradiated in G1. Locally damaged DNA was pulse-labeled with 3H-thymidine reflecting unscheduled DNA synthesis during excision repair of DNA photolesions. 3H-thymidine incorporation was detected by autoradiography in nuclei fixed immediately after the pulse (Fig. 2C) or in metaphase spreads prepared from cells that were allowed to proceed to the next mitosis (Fig. 2D). Alternatively, microirradiated DNA was visualized by immunostaining with antibodies raised against UV-damaged DNA (Cremer et al. 1984b) (Fig. 2E). As predicted by the CT concept, both approaches clearly showed that microirradiation of a small part of the nucleus damaged only a small

subset of the mitotic chromosome complement (Fig. 2D). The labeled parts of mitotic chromosomes revealed the UV damaged segments of neighboring chromosome territories hit by the microbeam during the preceding interphase. Consistent with this result, microirradiation of a small part of the metaphase plate of a living cell yielded a mirror-like pattern of distinctly labeled domains in the resulting daughter nuclei (Fig. 2F). These experiments provided the first compelling, although still indirect, evidence for the existence of chromosome territories. The direct visualization of individual CTs was made possible by in situ hybridization techniques developed during the mid 1980s. Initial experiments were performed with total human genomic DNA on cell hybrids that contained only one or few human chromosomes in a mouse or hamster genome complement (Manuelidis 1985; Schardin et al. 1985) (Fig. 3A). The achievement of chromosome sorting by flow cytometry of fluorescently labeled mitotic chromosomes (Cremer et al. 1984a; Gray et al. 1987; Fawcett et al. 1994) enabled the generation of chromosome specific painting probes for a large number of species. Subsequent amplification of DNA by cloning in bacterial vectors or by universal PCR (Telenius et al. 1992) and novel techniques for the suppression of ubiquitous repetitive sequences by COT-1 DNA (Cremer et al. 1988; Pinkel et al. 1988; Lichter et al. 1988a; Lichter et al. 1988b) or by depletion of these sequences from the respective probes (Bolzer et al. 1999) made it possible to delineate individual chromosomes in metaphase plates (Fig. 3B) and their territories in the interphase nucleus (Fig. 3C – F,K). To study the spatial arrangement of CTs, elaborate three-dimensional (3D) FISH protocols were developed (Cremer et al. 2008). 3D FISH in combination with light optical serial sectioning of nuclei by laser confocal microscopy and 3D image reconstruction has become the method of choice for studies of higher-order arrangements of CTs. The increasing availability of DNA probes for specific subchromosomal regions from chromosome arms down to bands

T. Cremer and M. Cremer

Figure 1. Early concepts in favor of a territorial organization of chromosomes in interphase nuclei. (A–B) Carl

Rabl´s hypothetical view (Rabl 1885) of a territorial chromosome arrangement in the interphase nucleus was based on studies of Proteus and Salamandra, in particular on epithelial cells from Salamandra maculata larvae. (A) Side view; supposed CTs are built up from primary chromatin threads (left side), from which (Continued )

Chromosome Territories

and single gene loci enabled studies of CT substructures (Fig. 3G – J). NONRANDOM ARRANGEMENTS OF CHROMOSOME TERRITORIES AND CHROMOSOMAL SUBREGIONS

Higher-order chromatin arrangements may primarily reflect geometrical constraints, which obviously affect the 3D distribution of larger and smaller CTs crowded together in the nuclear space (Neusser et al. 2007). Occasionally, however, proximity patterns assembled by chance, may have provided functional advantageous and consequently were favored by natural selection. The search for nonrandom chromatin assemblies, the mechanisms responsible for their formation and their functional implications is one of the major goals of nuclear architecture research. This search is still in its beginning. Methodological Problems in Distinguishing Random and Nonrandom Higher-Order Chromatin Arrangements

An overview on methods and problems for the spatial quantitative analysis of fluorescently labeled CTs and other nuclear structures is provided in Ronneberger et al. (2008). To decide about the random or nonrandom distribution

of a given target, such as a CT, chromosomal subregion, or gene, it is important to define a single or sometimes multiple 3D reference points, which represent the target in question in the nuclear space. For a painted CT, its intensity gravity center can be chosen as a single reference point or the CT surface may be used to define multiple reference points. Next, proper reference structures must be defined to decide whether the chosen target is distributed randomly or nonrandomly with respect to them. For reference structures, one can choose other chromatin targets or distinct nuclear structures, e.g., the nuclear lamina, nucleoli, or splicing speckles. Two cases of nonrandom arrangements of specific chromatin targets are considered here: nonrandom radial arrangements and nonrandom neighborhood arrangements, also referred to as proximity patterns. For studies of the radial arrangement, the nuclear space can be divided into a number of concentric shells with equal volume. 3D distances measured in a random sample of nuclei between the fluorescence intensity gravity center of a given labeled target and either the nuclear center or the closest point of the nuclear lamina provide information about its preferred radial nuclear position. Alternatively, all voxels contributing to a given target and reference structure, respectively, may be used to measure 3D

Figure 1. (Continued ). secondary and tertiary threads branch out and form a chromatin network (right side). Spindle attachment sites, now known as centromeres, congress at one site of the nucleus (top, Rabl’s Polfeld), whereas the telomeres cluster at the opposite site (bottom, Rabl’s Gegenpolseite). (B) View from above on the Polfeld. (C) Drawings made by Theodor Boveri (1909) from two Ascaris megalocephala univalens embryos fixed during prophase of the two-cell stage. Arrows point to nuclear protrusions containing the distal parts of the two germ line chromosomes. (D) Boveri’s drawing of a fixed four-cell embryo shows two pairs of cells with a distinctly different arrangement of these protrusions in interphase nuclei. Boveri argued that the two upper and two lower cells, respectively, represent daughter cells and explained the strikingly different nuclear protrusion patterns observed in the two cell pairs as a result of chromosome movements during prometaphase. (E) Eduard Strasburger’s colored model view of a tissue cell nucleus from the plant Galtonia candicans (Strasburger 1905). Chromosome territories supposedly are built up from higher-order chromatin foci delineated in red and blue. (F ) The white and gray shaded bundles in Boveri’s sketch from 1909 reflect two neighboring CTs with sponge-like structures built up from networks of anastomizing chromatin bundles. The continuous lines reflect Rabl’s primary threads, and the dotted line depicts the possibility of a rare pathological situation, in which secondary chromatin threads from one CT encompass the primary thread of the other, a situation possibly leading to an exchange of chromatin material between the chromosomes or a segregation failure during the next mitosis.

T. Cremer and M. Cremer

B

A

C

D 2

E

2 1

Figure 2. Experimental evidence for a territorial organization of interphase chromosomes. (A) Giemsa stained interphase nucleus of a fixed Chinese hamster cell (CHO line) reveals chromatin clumps likely representing individual CTs (reprinted with permission from Stack et al. 1977). (B) Experimental rationale of laser-UVmicrobeam experiments to distinguish between a nonterritorial (upper row) and a territorial (bottom row) chromosome arrangement in cell nuclei (compare Cremer et al. 1982a). (C) Autoradiograph of a diploid Chinese Hamster cell. The nucleus of the living cell was microirradiated in G1, pulse-labeled with 3H thymidine and fixed immediately thereafter. The arrow points to a cluster of silver grains detected over the site of microirradiation. (D) Metaphase spread from the same experiment obtained about 40 hours after microirradiation. One chromosome 1 and one chromosome 2 are intensely marked with silver grains, indicating that the microbeam hit the respective territories during interphase, whereas their homologs are unlabeled, arguing against the spatial association of the homolog territories. (E) Immunocytochemical identification of microirradiated DNA (arrows) in a pair of Chinese hamster daughter nuclei fixed around 4 hours after microirradiation of a small part of a metaphase plate. The mirror-like distribution of microirradiated chromatin in the two daughter nuclei argues for a similar arrangement of chromosome territories (reprinted from Cremer et al. [1984b] with permission).

distances. An equal frequency of voxels for a given chromatin target in all nuclear shells strongly argues for its random radial nuclear interior or peripheral arrangement of the target.

Nonrandom proximity patterns between sets of targets are indicated by 3D distance measurements, which are significantly smaller than expected in case of their random distribution

Chromosome Territories

in the nuclear space. It is important to note that nonrandom proximity patterns between genes located at distant sites of the same chromosome or on different chromosomes provide hints for but do not prove functional implications. For example, an apparently nonrandom proximity pattern of certain targets may simply reflect their nonrandom radial distribution in an interior nuclear shell, resulting in significantly smaller mean distances and a significantly higher frequency of contacts compared with targets randomly arranged in a peripheral nuclear shell. The importance of clearly defining targets and reference structures for which possible random or nonrandom arrangements are analyzed, can hardly be overemphasized. Certain targets may be arranged highly nonrandomly in the nuclear space, but still entirely at random in an internal nuclear shell or with respect to other more limited reference volumes or other reference structures. Before a detected nonrandom proximity pattern between certain targets gives rise to speculations about functional interactions, it should always be shown that this pattern cannot be simply explained as a consequence of a nonrandom radial distribution. As another caveat, it should be noted that the size of samples of nuclei subjected to a detailed 3D analysis is typically small (in the order of 20 – 50). It is of utmost importance to avoid a biased selection of such nuclei.

Radial Nuclear Arrangements of Chromosome Territories and Chromosome Subregions

Early evidence for a nonrandom radial distribution of entire CTs was based on (3D) FISH experiments in human lymphocyte nuclei using chromosome painting probes for human (HSA) chromosome 19, the chromosome with the highest gene density and for HSA 18, a gene poor chromosome. HSA 19 CTs were consistently found in the interior of human lymphocyte nuclei and of numerous other cell

types, whereas the territories of the gene poor HSA 18 were located at the nuclear periphery (Fig. 3C) (Croft et al. 1999; Cremer et al. 2001; Cremer et al. 2003). These observations of a gene density correlated radial arrangement in the nucleus were completed and confirmed by analyses comprising all human chromosomes (Boyle et al. 2001). An evolutionary comparison of lymphoblastoid cells from various primate species showed that this nonrandom radial nuclear distribution has been evolutionary conserved despite major evolutionary chromosome rearrangements. Orthologous segments of HSA 19 are positioned in the nuclear interior, whereas segments corresponding to human chromosome 18 locate at the nuclear periphery (Fig. 3D) (Tanabe et al. 2002). Nonrandom radial nuclear arrangements of CTs, depending on their gene density, were also observed in rodents (Mayer et al. 2005; Neusser et al. 2007), cattle (Koehler et al. 2009), and birds (Habermann et al. 2001). Recently, it was shown in bovine preimplantion embryos that this difference was not yet present in nuclei of early blastomere stages. Its first appearance correlated with major genome activation and was fully established in blastocysts (Koehler et al. 2009) (Fig. 3E). A multicolor 3D FISH approach was established for diploid human fibroblasts (46,XY) and allowed the colorful discrimination of the 22 pairs of autosmal CTs and the CTs of the two sex chromosomes (Bolzer et al. 2005) (Fig. 3F). For the flat-ellipsoid fibroblast nuclei, a radial arrangement of CTs, mainly according to chromosome size or DNA content, was shown in contrast to the clearly gene density correlated radial pattern observed in spherically shaped nuclei, such as nuclei in lymphocytes. Still, gene density correlated patterns on the subchromosomal level were also present in fibroblast nuclei, as shown by the preference of gene dense, Alu sequence-rich chromatin in the nuclear interior. 3D FISH experiments using sets of BAC clones with inserts from gene dense and gene poor chromosome segments confirmed such a distinct gene density correlated radial

T. Cremer and M. Cremer

A

D

B

#18 (5 genes/Mbp) #19 (20 genes/Mbp)

C

Human

E

Orangutan

6

F

White handed gibbon 3

6

1

6 Y

3

Y

3 6

2

1 1

1 2

G

H

2

I

Mb 0.5 1.0 1.5 2.0

11p15.5

J

K

Xa Xa Xi

Xa

Xi

Xi

Figure 3. Direct evidence for chromosome territories (CTs) by in situ hybridization experiments. (A) In situ

hybridization with biotinylated human genomic DNA of a Chinese hamster human hybrid cell line carrying a single human X chromosome reveals the specifically labeled human X chromosome (arrow) in metaphase spreads (top; other chromosomes are Giemsa stained) and its respective human X territory in interphase nuclei (bottom); for details, see Schardin et al. (1985). (B) Visualization of individual chromosomes in a human (HSA) metaphase plate (chr. HSA18 in red, chr. HSA 19 in green) after fluorescence in situ hybridization (FISH) using labeled chromosome painting probes. (C) A single light optical mid-section through the nucleus of a human lymphoblastoid cell after 3D FISH with the (Continued )

Chromosome Territories

nuclear arrangement (Fig. 3G,H). Present data provide strong evidence that the local gene density within windows of 2 – 10 Mb is apparently a strong and likely pivotal key player for the radial position of chromatin in the nucleus (Kozubek et al. 2002; Cremer et al. 2003; Murmann et al. 2005; Kupper et al. 2007). Other parameters, in particular transcriptional activity, replication timing, and GC content have also been

correlated with nonrandom radial nuclear arrangements of CTs and chromosomal subregions (Mayer et al. 2005; Federico et al. 2006; Goetze et al. 2007; Grasser et al. 2008; Hepperger et al. 2008). However, the interdependence of these parameters within a given DNA segment makes it difficult to dissect their impact on radial chromatin distribution in the nucleus (Kupper et al. 2007). The observation

Figure 3. (Continued ). same painting probes shows a HSA 19 CT (green) in the nuclear center and a HSA 18 CT

(red) at the periphery; for details, see Tanabe et al. (2002). (D) Top: Idiogramatic illustration of primate chromosomes or subchromosomal regions from the Orangutan ( pongus pygmaeus, PPY, middle) and white handed gibbon (Hylobates lar, HLA, right) orthologous to human chromosomes 18 (red) and 19 (green). Note the pronounced chromosomal rearrangements between the human and the HLA karyotype: The arm of a large HLA chromosome is orthologous to HSA 18, whereas four HSA 19 orthologous chromosome segments are distributed on three other large HLA chromosomes. Bottom: 3D reconstructions of representative HSA (left), PPY (middle), and HLA (right) lymphoblastoid cell nuclei reveal the same nonrandom radial nuclear positions of orthologous gene dense HSA 19 (green) and gene poor chromatin HSA 18 (red); for details, including quantitative evaluation, see Tanabe et al. (2002). (E) Painted CTs of the gene dense chromosome BTA 19 (green) and gene poor BTA 20 chromosome (red) in a domestic cattle (Bos taurus, BTA) embryo during blastocyst stage. Left: Maximum intensity projections of painted CTs in DAPI stained nuclei (blue). Right: 3D reconstructions of nuclei of the same embryo from a different perspective. Gene dense BTA 19 chromatin is preferentially distributed in the nuclear interior and gene poor BTA 20 chromatin at the nuclear periphery: for details, including quantitative evaluation, see Koehler et al. (2009). (F ) Simultaneous delineation of all chromosomes in a human fibroblast nucleus (left) and a prometaphase rosette (right) by multi-color FISH. Light optical mid-sections with false color representation of all CTs and prometaphase chromosomes, respectively, are shown. Examples of individual CTs and mitotic chromosomes are denoted with their respective karyotypic number; for details, see Bolzer et al. (2005). (G) Partial 3D reconstruction of a human lymphocyte nucleus after 3D-FISH of two differently labeled sets of BAC clones from HSA 12 carrying sequences from several gene-dense (green) and several gene-poor chromosome segments (red), respectively. This nucleus illustrates two neighboring HSA 12 CTs with distinct gene density correlated radial nuclear arrangement. (H ) Left: 3D reconstruction of a single painted HSA 12 territory (blue) showing the distinctly different, polarized arrangements of these two sets of BAC-clones. Right: 3D reconstruction of a HSA 12 CT recorded after 3D FISH with two differentially labeled sets of BAC clones containing sequences from highly expressed (green) and repressed genes (red), respectively, shows that active and silent genes are distributed throughout the CT; for details, see Kupper et al. (2007). (I ) Multicolor 3D FISH to a human fibroblast reveals the two HSA 11 CTs (blue) together with the particular gene dense region 11p15.5 (green, yellow, and red). For the delineation of this region, 15 BACs were used and differentially labeled as shown in the inset. Z-projections of light optical serial sections illustrate different shapes of this region: a finger-like chromatin protrusion in the lower CT and a much more compact shape in the upper CT; for details, see Albiez et al. (2006). (J ) Top: Two human X chromosomes in a human fibroblast metaphase plate are shown after multicolor FISH with four differentially labeled BAC pools representing four segments from qter to pter (q-arm: green, blue; p-arm: yellow, red). Bottom: Projections of light optical sections through the Xa- and Xi-territory of a human fibroblast nucleus following 3D FISH with the same BAC pools show four separate domains of these segments within the Xa- and Xi-territory (images courtesy of Kathrin Teller, Univ of Munich). (K) Left: Painting of the Xa- and Xi-territory in a female human fibroblast nucleus (46,XX) exemplifies the different shape and painting intensity of the two X-territories. The Xi-territory was independently identified by Barr body staining (not shown). Right: intensity profiles of enlarged images of the Xi- and Xa-territory. The color code white, yellow, and red reflects high, medium, and low intensities (images courtesy of Irina Solovei, Univ of Munich).

T. Cremer and M. Cremer

that radial positions of specific gene regions can differ significantly between cell types argues for additional cell-type-specific factors (Hepperger et al. 2008). The picture gets even more complex when we consider movements toward the nuclear interior shown for individual genes on transcriptional activation (see later). These movements emphasize that radial nuclear positions of genes do not solely depend on the local DNA sequence environment, where a given gene is embedded. Proximity Patterns of Chromosome Territories

Whereas little evidence for nonrandom neighborhood/proximity patterns was found between specific homologous and heterologous chromosomes in nuclei of human fibroblasts (Bolzer et al. 2005), nonrandom proximity patterns for smaller subsets of nonhomologous CTs were described by Misteli and coworkers in cell nuclei from various mouse tissues (Parada et al. 2002; Roix et al. 2003; Parada et al. 2004) and in human lymphocytes (Brianna Caddle et al. 2007; Khalil et al. 2007). These proximity patterns, however, were of a rather probabilistic nature, i.e., their presence was shown by an excess of certain heterologous (and occasionally also homologous) CTs in a population of nuclei, not as an event consistently observed in each nucleus. Differences of statistically preferred proximity patterns were recently also described for CT subsets in different fibroblast-derived cell lines (Zeitz et al. 2009). It should be emphasized that a pronounced cell-to-cell variation of CT neighborhood arrangements was found in all cell types studied so far. Neither functional implications of cell type or cell-line-specific probabilistic neighborhood arrangements nor mechanisms responsible for their establishment are presently understood. Such studies resemble snap-shots of individuals present in a room. A single snapshot does not allow conclusions about the dynamic behavior of these individuals. Accordingly, a decision whether certain CTs are permanently close to each other, have met before or will meet after the snap-shot was taken, would

require many snap-shots of well defined cell types in fixed samples at the same stage of interphase or postmitotic terminal differentiation, which are presently not available.

CT STRUCTURE, SHAPES, AND PLASTICITY

Apparently, 1 Mb-chromatin domains, first detected in S-phase nuclei as replication foci (Ma et al. 1998) and later shown to be persistent higher-order chromatin structures, are basic structural units, which build up CTs (Jackson and Pombo 1998; Visser and Aten 1999; Berezney et al. 2005; Albiez et al. 2006). To date, neither their ultrastructural organization nor the packaging of chromatin connecting these domains has been fully clarified. In addition, chromonema fibers may play an important role in the higher-order organization of CTs (Belmont and Bruce 1994) and their structural interconnection to global chromatin networks (Albiez et al. 2006). Single 1-Mb domains are likely built up from smaller loop domains, whereas larger chromatin clumps may be composed of clusters of 1-Mb domains. CTs visualized by 3D FISH appear as structures with manifold shapes composed of higher-order chromatin domains (Dietzel et al. 1998; Khalil et al. 2007; Kupper et al. 2007) (Fig. 3H–K). Separate arm domains were disclosed by painting of chromosome arms or parts of them in human cell nuclei (Dietzel et al. 1998) (Fig. 3K). The outer surface of an individual CTapparently does not generally provide a particular compartment for gene dense and/or transcriptionally active chromatin as originally suggested (Zirbel et al. 1993). A few chromatin regions with particular high gene density and/or transcriptional activity, such as the 11p15.5 segment (Fig. 3I), the MHC and EDC loci, or the HOX gene cluster have been consistently found looping out as protrusions from the core territory (Volpi et al. 2000; Williams et al. 2002; Chambeyron et al. 2005; Kupper et al. 2007). On a more global level, however, gene-dense and/or highly expressed sequences were found equally distributed throughout their respective territories (Mahy et al. 2002; Kupper et al. 2007)

Chromosome Territories

(Fig. 3H). The dissociation of gene localization relative to their CTs and gene regulation was shown for the murine Hoxd locus during differentiation and development: Although both decondensation and movement outside of the CT occur during gene activation in ES cells and the tail bud of the embryo, in the limb bud, gene activation and chromatin condensation occur without any looping out from the CT (Morey et al. 2007). This topic is described in detail in Heard and Bickmore (2007) and Morey et al. (2009). A particular staining intensity of the inactive X chromosome (Xi) with DNA-specific dyes was observed decades ago (so called Barr body [Lyon 1962]), indicating a higher compactness in Xi compared with the active X chromosome (Xa) or other autosomes (Fig. 3K, L). Previous studies showed distinct differences between Xi and Xa in shape and surface structure but surprisingly little differences in volume (Eils et al. 1996). In Xi, Clemson et al. (2006) observed a preferential positioning of genes —irrespective of their transcriptional activity— at the nuclear periphery, whereas repetitive sequences were found rather in the nuclear interior. Such a distinct radial arrangement was not reported for Xa, possibly because of larger invaginations in Xa. At large, however, structural differences between Xi and Xa on the subchromosomal level are poorly defined to date, although the CTs X provide an excellent model system for structure/function relationships in homologous chromosomes with different functional allocation. Detailed knowledge of the folding structure (conformation) of small contiguous chromatin segments within a given CT is limited so far to a few regions in autosomes and further studies are definitely required before a comprehensive picture can be drawn on the internal structure and local conformation of CTs. Shopland et al. (2006) analyzed the 3D structure of a highly conserved 4.3-Mb region on mouse chromosome 14 containing four clusters of genes separated by gene “deserts.” In Drosophila melanogaster, gene dense and gene poor segments within a 7-Mb region of chromosome 2 were described to form spatially segregated

clusters of different stability and folding structure (Boutanaev et al. 2005), and Goetze et al. (2007) showed for several cell lines that ridges (contiguous genomic regions harboring adjacent genes with high ubiquitous transcriptional activity) are in general less condensed and more irregularly shaped than “antiridges”, arguing that the structure of these two types of genomic domains is largely independent of tissuespecific variations in gene expression. MateosLangerak et al. (2009) provide data on the folding structure of subregions of CT 1 and 11 that are in accordance with a suggested random loop model with 10 – 30 loops/Mb. The relevance of 3D FISH studies for conclusions regarding the fine structure of CTs in vivo has to consider some technical issues. Differences in the fixation procedure of cells, amplification, and labeling efficiency of chromosome painting probes may have an impact on CT delineation. The necessary suppression or depletion of interspersed repetitive sequences prevents a complete visualization of CTs even under optimal hybridization conditions. The most crucial step for CT fine structure occurs during the denaturation of the nuclear DNA, a step necessary for successful probe hybridization (Solovei et al. 2002). These limitations emphasize the need for the visualization of CTs in living cells under conditions which do not interfere with chromatin or other nuclear functions (Zink et al. 1998; Walter et al. 2003). Although urgently required, in vivo approaches have their own technical problems, such as an interference of in vivo labeling strategies with the function of labeled chromatin and phototoxicity effects during prolonged observation periods. Different approaches have been established for the quantitative assessment of CT conformation. These include density, volume, roundness, and smoothness factors, as well as principal component analyses, which are also potentially useful to assess CT orientation within the nucleus (Eils et al. 1996; Roix et al. 2003; Ronneberger et al. 2008). The subjective choice of threshold setting for confocal image stacks influences the determination of CT borders and accordingly of measured CT volumes. Low threshold settings can result in the inclusion of

T. Cremer and M. Cremer

background, leading to overestimation of CT volumes and apparent chromatin intermingling between neighboring CTs, whereas high thresholds can lead to the visual loss of fine DNA structures looping out from the chromatin bulk of a given territory. STABILITY AND CHANGES OF CT ARRANGEMENTS IN CYCLING CELLS

The question of to what extent a given proximity pattern established between CTs during a given interphase may be transmitted through mitosis to the next interphase has remained controversial (Gerlich et al. 2003; Walter et al. 2003; Thomson et al. 2004; Kalmarova et al. 2008). These studies were based on live-cell approaches using transgenic cell lines with either FP-tagged or photoconvertible tagged histones. This allowed following up fluorescently labeled chromatin through mitosis into the next interphase. Present evidence argues for the stability of a given CT neighborhood arrangement once established at the onset of interphase until the next prophase, whereas chromosome movements during prometaphase necessary to establish the metaphase plate can lead to major changes of side-by-side chromosome arrangements in the metaphase plate compared with the side-by-side arrangements of the respective CTs during the preceding interphase (Walter et al. 2003; Cvackova et al. 2009). DYNAMICS OF CT ARRANGEMENTS DURING POSTMITOTIC CELL DIFFERENTIATION AND IN TERMINALLY DIFFERENTIATED CELLS

In a seminal investigation, Barr and Bertram described that on electric stimulation of cat motor neurons, a “nucleolar satellite” (now known as the Barr body) moved from its usual position adjacent to the nucleolus toward the nuclear membrane within a time course of several days (Barr and Bertram 1949). Borden and Manuelidis (1988) showed a pronounced repositioning of the human X-territories in neurons of both males and females in electrophysiologically

defined seizure foci. Repositioning of CTs during cellular differentiation of murine cerebellar Purkinje neurons was described by Martou and De Boni (2000) in terms of changes of centromere positions that adopted their final position around day 5 post partum. A striking example of CT reorganization during terminal differentiation was recently shown by Solovei et al. (2009) in a study of the mammalian retina. In mammals adapted to nocturnal life, all heterochromatin becomes located in the nuclear interior during postmitotic terminal differentiation of rod cells, whereas all euchromatin is shifted toward the nuclear periphery. This transformation starts around day 6 post partum and takes several weeks for completion. Rod cells of mammals with diurnal life styles do not show such a chromatin reorganization. Their nuclei reveal the conventional pattern with heterochromatin enriched at the nuclear periphery and around the nucleoli, whereas euchromatin is mostly distributed in the nuclear interior. This global nuclear reorganization in rod cells of nocturnal species necessitates a profound reorganization of radial chromatin arrangements rather than a change of CT proximity patterns. Unexpectedly, the inverted pattern of rod cell nuclei in nocturnal mammals reflects an adaptation to vision in low light conditions. Because of the somewhat higher refractive index of heterochromatin compared with less condensed euchromatin, inverted nuclei act as microlenses, which help to channel photons to the photoreceptors. This may be the first example in which a specific adaptive advantage of a celltype-specific nuclear architecture could be shown. Interestingly, this functional specialty shows to what extent the nuclear architecture can be modified under an overriding selective pressure. DYNAMICS AND INTERACTIONS OF SPECIFIC GENE LOCI LOCATED ON THE SAME OR ON DIFFERENT CTs

This question is particularly important with respect to long-range chromatin movements

Chromosome Territories

involved in the congression of coregulated genes. Live-cell experiments performed with cultured mammalian and Drosophila cells showed locally constrained movements of subchromosomal domains (Abney et al. 1997; Marshall et al. 1997; Bornfleth et al. 1999; Edelmann et al. 2001). The typical prevalence of heterochromatin localized at the lamina and the observation of silenced genes in this peripheral nuclear subcompartment has supported the concept that the nuclear periphery is a largely repressive environment for transcription, and vice versa, the nuclear interior a compartment for transcriptional activity (Schneider and Grosschedl 2007). This would require corresponding movements of gene loci in correlation to their (cell type) correlated transcriptional activity. Recent studies, however, support a more complex picture (Taddei et al. 2006; Akhtar and Gasser 2007; Deniaud and Bickmore 2009). Several groups succeeded to tether specific chromatin segments to the nuclear envelope in living cells. They found that some genes were suppressed when closely associated with the envelope, but that others were not (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Of particular interest are hints that a longrange spatial nuclear convergence of genes, which are located many megabases apart in cis, i.e., on the same chromosome, or trans, i.e., on different chromosomes, might be involved in mechanisms of gene activation or silencing (Zuckerkandl and Cavalli 2007; Bartkuhn and Renkawitz 2008). This phenomenon has been referred to as “gene kissing” (Kioussis 2005) or “chromosome kissing” (Cavalli 2007). As an early example, LaSalle and Lalande (1996) presented 3D FISH evidence for the transient spatial association of the AS/PWS loci during late S phase. These loci comprise the genes involved in two imprinting disorders, the Angelman syndrome and the Prader–Willi syndrome. The authors argued that transient “kissing” between the two loci is required for maintaining opposite imprints in cycling cells. This specific case of “kissing,” however, could not be confirmed in a later study (Teller et al. 2007).

A “nonmicroscopic” biochemical approach with a great potential for the disclosure of spatial interactions of specific genomic loci is the technique of “chromosome conformation capture” (3C) introduced by Dekker et al. (2002) and its recent extension to 4C (chromosome conformation capture-on-chip [Simonis et al. 2006] and “circular chromosome conformation capture” [Zhao et al. 2006]), respectively. With this “high throughput” approach, it has become possible to determine genome wide nonrandom spatial interactions between specific DNA segments in cis and trans to a reference locus on a given CT (for review, see Dekker 2008). This method is based on in situ formaldehyde cross-linking of proximal DNA– protein interactions with a distance of several A˚ngstro¨m, subsequent fragmentation of cross linked DNA, recircularization (ligation) and amplification, and finally identification of these products by microarray techniques. To date, numerous spatial interactions of CTs were reported in cis and trans (Ling et al. 2006; Lomvardas et al. 2006; Babu et al. 2008). These assays demand strict controls to escape the danger of false positive or negative findings (Simonis et al. 2007). By combining this proximity based ligation assay with massive parallel sequencing (Hi-C), the construction of spatial proximity maps of an entire genome down to a resolution level of 1 Mb was recently achieved as shown for a human lymphoblastoid cell line (LiebermanAiden et al. 2009). These maps confirmed the presence of chromosome territories, the spatial proximity of small, gene dense chromosomes, and the spatial segregation of open and closed chromatin, a parameter that strongly correlates with gene density (Gilbert et al. 2004). At the megabase scale, a chromatin conformation consistent with a fractal globule was suggested. High throughput assays with a superior resolution and microscopic approaches are complementary to further elucidate CT conformation and spatial interactions in trans (Simonis and de Laat 2008). Ligation techniques to date require millions of cells and their principle excludes an application at the single cell level. More important, microscopic approaches are

T. Cremer and M. Cremer

the only way to reveal the entire structure of nuclear components and to determine their topography with respect to each other. MODELS OF NUCLEAR ARCHITECTURE: OPEN QUESTIONS AND EXPERIMENTAL STRATEGIES TO ANSWER THEM

The territorial organization of interphase chromosomes is now generally accepted as a basic principle of nuclear organization in both animals (Cremer and Cremer 2001) and plants (Shaw et al. 2002; Pecinka et al. 2004; Berr et al. 2006), and may even hold for single-cell eukaryotes, such as budding and fission yeast (Bystricky et al. 2005; Molnar and Kleckner 2008). However, severe limitations in our present knowledge of the functional architecture of CTs and the nucleus at large become obvious when we consider currently proposed models (Fig. 4). The chromosome territory-interchromatin compartment (CT-IC) model (Fig. 4A,B) argues that nuclei are built up from two principal components, chromosome territories (CTs) and the interchromatin compartment (IC). Individual CTs form an interconnected higherorder chromatin network (Visser et al. 2000; Albiez et al. 2006). According to the CT-IC model, this chromatin network is spatially associated with a second contiguous 3D spatial network, the interchromatin compartment (IC), which was observed both at the light and electron microscopic level (Albiez et al. 2006; Rouquette et al. 2009). The IC concept asserts a DNA free or at least largely free, contiguous space of channels, which start at the nuclear pores and expand as larger channels and lacunes between the higher-order chromatin network described earlier. The IC harbors splicing speckles and a variety of nonchromatin nuclear bodies (Verschure et al. 1999; Visser et al. 2000; Albiez et al. 2006). The IC concept evolved from the interchromosomal domain (ICD) concept originally proposed by Peter Lichter and co-workers (Zirbel et al. 1993), who defined the ICD as a network-like space expanding mainly around CTs with little penetration into the CT interior (Cremer et al. 1993).

Supposedly, genes were preferentially transcribed in a region of decondensed chromatin at the CT periphery and RNA transcripts would be directly released into the ICD compartment. This concept was supported by a series of studies from this group (Bridger et al. 1998; Reichenzeller et al. 2000; Bridger et al. 2005; Richter et al. 2005). Accumulating evidence for genes transcribed both outside and in the interior of CTs (Cmarko et al. 1999; Verschure et al. 1999; Mahy et al. 2002; Kupper et al. 2007) is consistent with electron microscopic evidence for a (nearly) network-like DNA free space both outside and inside CTs (Visser et al. 2000; Rouquette et al. 2009). Based on this evidence, the hypothetical CT structure suggested by the CT-IC model can be compared with a sponge of chromatin permeated by intraterritorial IC channels (Fig. 4B). The entire IC is separated from the more condensed interior of chromatin domains and/or higher-order chromatin fibers by a thin (,200 nm) layer of rather decondensed chromatin, termed the perichromatin region (PR) (Fakan and van Driel 2007). EM evidence has supported the view that the PR topographically represents the utmost periphery of a given chromatin domain bordering the IC and functionally represents the major nuclear subcompartment for transcription, cotranscriptional RNA splicing (Fakan and Bernhard 1971; Cmarko et al. 1999; Trentani et al. 2003), as well as DNA replication (Jaunin and Fakan 2002) and possibly also DNA repair (Solimando et al. 2009). Transcription yields perichromatin fibrils, which are generated in the PR as nascent pre-mRNA transcripts of single genes complexed with hnRNPs and are served by nearby speckles with factors for cotranscriptional splicing. The PR concept is a decisive part of the CT-IC model but has been disregarded by proponents of other models (Dehghani et al. 2005; Branco and Pombo 2006; Fraser and Bickmore 2007; Alberts et al. 2008; Fedorova and Zink 2008; Fedorova and Zink 2009). The “lattice” model of interphase chromatin proposed by Dehghani et al. (2005) suggests a lattice-like network of 10- and 30-nm fibers. This structure yields a porous organization of chromatin with

Chromosome Territories

A

C

1 Mb chromatin domains

Perichromatin region

IC

5 P

CTs

B Chromosome territory

D Speckle

IC Genes

Chromatin domain

IC

PF Genes

Interchromatin compartment

Figure 4. Different models of nuclear architecture. (A) Chromosome territory-interchromatin compartment

(CT-IC) model (for description, see text). (B) Hypothetical view of the functional nuclear architecture according to the CT-IC model. Chromatin domains are considered the major constituents of a CT. The IC expands between these domains as a rather DNA free nuclear compartment carrying splicing speckles and nuclear bodies. The width of the IC space is highly variable depending on Brownian movements of chromatin domains and allowing transient contacts of domain surfaces in cis and trans. During ongoing transcription, genes are at least partially decondensed at any given time into the perichromatin region (PR) located at the domain periphery. Perichromatin fibrils (PF) are generated there. Each PF carries a nascent transcript (green) from a different gene. White dots with a line symbolize RNA Pol II molecules with their CTD domain, which may play a role in the structural organization of splicing events. Splicing speckles located in the IC provide the splicing factors to PFs, which also represent the structures in which cotranscriptional splicing occurs. (C) According to the interchromatin network (ICN) model (Branco and Pombo 2006), intermingling chromatin fibers/loops from the same CT, as well as from neighboring CTs, can make contact in cis and trans. Blue dots represent sites of intrachromosomal and interchromosomal contacts with unknown composition. Although there is extensive space between chromatin fibers/loops, this space should not be confused with the functional relationship of the IC and PR predicted by the CT-IC model. (D) Model suggested by Fraser and Bickmore (Fraser and Bickmore 2007, figure reprinted with permission from Macmillan Publishers Ltd). These authors review evidence arguing for the colocalization of genes in the nucleus for expression or coregulation. Transcription factories (dark pink) can recruit genes in cis and trans located on decondensed chromatin loops that extend outside chromosome territories. The pale pink area on the left represents a splicing-factor enriched speckle. The blue circle exemplifies an interaction for coregulation in trans, which can occur between regulatory elements and/or gene loci.

fibers intermingling at the borders of neighboring CTs. The interchromatin network (ICN) model (Fig. 4C) (Branco and Pombo 2006) predicts that chromatin fibers and loops intermingle in a rather uniform way both in the interior of individual CTs and between differentially labeled neighboring CTs, making any

distinction between the interior or periphery of distinct chromatin domains functionally meaningless. In this interchromatin network, loops may expand from one CT to meet loops from another CT. Arguably, active genes on decondensed chromatin loops that extend outside chromosome territories can colocalize both

T. Cremer and M. Cremer

in cis in so called expression hubs (Kosak and Groudine 2004) or be transcribed at preassembled transcription factories (Fig. 4D) (Fraser and Bickmore 2007). Some models emphasize the functional importance of giant chromatin loops, which emanate from chromosome territories (Chubb and Bickmore 2003; Fraser and Bickmore 2007). Supposedly, giant loops may even expand across the nuclear space (Alberts et al. 2008). According to these models, giant loops may carry genes to even very remote sites in the nuclear space for coregulation in an expression hub (Kosak and Groudine 2004). Alternatively, giant loops may transport genes to a remote repressive nuclear compartment (Alberts et al. 2008). The lack of quantitative rigor with respect to the predicted numbers, length, and compaction levels of such fibers and their 3D distribution has limited the usefulness of the models described earlier. To overcome this limitation, chromatin polymer models have been developed, which make experimentally testable, quantitative predictions about functionally important features of the nuclear architecture, such as the expected size distribution of chromatin loops and chromatin compaction levels (van den Engh et al. 1992; Munkel and Langowski 1998; Munkel et al. 1999). A recent chromatin polymer model has assumed a broad range of loop sizes (Mateos-Langerak et al. 2009). None of the present models for functional nuclear architecture is fully supported by compelling experimental evidence. In particular, two problems need urgent clarification: 1) the possible speed and extent of chromatin movements driven by Brownian motion in case of random walks and by unknown mechanisms in case of directed chromatin movements; 2) the topography of the major nuclear functions and the validation or experimental falsifications of the functional marriage between the IC and the PR as predicted by the CT-IC model (Fig. 4B) but not by other models (Fig. 4C, D). Although estimates of the number of transcription factories vary widely, the number of genes transcribed at any given time seems to be much larger. Accordingly, it has been suggested that a single transcription factory is able to

transcribe several genes simultaneously in cis and trans and that an extraordinarily large part of the genome passes through the limited number of transcription factories in a cell nucleus (Chakalova et al. 2005; Sutherland and Bickmore 2009). As a consequence, one would also expect an extraordinary mobility of chromatin in the nucleus. In contrast, the concept of perichromatin fibrils argues that most transcription takes place in the PR and that simultaneous transcription in a single factory is rather the exception than the rule, if this concept can finally be proven (Sutherland and Bickmore 2009). On the basis of FISH experiments with differentially labeled chromosome painting probes on cryosections (140 – 180 nm) from cell nuclei, Branco and Pombo (2006) detected zones of color overlap between pained CTs (Fig. 5A– C), which they interpreted as the result of intermingling chromatin fibers. Evidence for intermingling was substantiated by transmission electron microscopy with colloidal gold particles of different size for “intermingling” CTs. In control experiments, Branco and Pombo showed that the positions of gold grains reflecting the location of histone H2B molecules did not significantly change when cryosections were studied before and after mock FISH, excluding the possibility that chromatin intermingling was an artifact of chromatin denaturation. Examination of published electron micrographs does not, however, allow a morphological orientation in the section with regard to chromatin domains and other nuclear structures. Controversial issues both with respect to CT structure and the interchromatin space are in part because of the limited resolution of present light microscopy techniques and the difficulties to obtain 3D data sets by electron microscopy. Recent developments of laser-based light microscopic techniques with ultra-high resolution has now opened possibilities for light optical nanoscopy, which will help to overcome these limitations (Hell 2007; Schermelleh et al 2008; Gunkel et al. 2009). The combination of a focused ion beam with high resolution scanning electron microscopy is an additional, very promising tool for 3D analysis of chromosome architecture (Schroeder-Reiter et al. 2009).

Chromosome Territories

A

B

C

D

E

H 1

1

3

3

2

2

4

4

5

5

F

G

I 1

1

3

4 2

4 2 5

Figure 5. Experimental evidence against and in favor of an interchromatin space. Fluorescence microscopic

images of a painted HSA 5 CT (A) and HSA 7 CT (B) in a cryosection of approximately 150 nm thickness from a human lymphocyte nucleus (Branco and Pombo 2006). (C ) Overlap of images A and B shows the two CTs in different colors. The area of assumed intermingling between the two CTs is delineated by the yellow line. (D –G) Topographical relationships between double minute (DM) chromosomes carrying active MYCN genes (red) and the painted 3q-arm domain (green) observed in a nucleus of the human neuroblastoma cell HDN-16 (for details, see Solovei et al. 2000). (D) Laser confocal section. DMs denoted by 1 –5 are located both at the periphery of the 3q-arm domain and in channel-like invaginations of the interchromatin compartment (IC) penetrating into the interior of the arm domain. (E) IC invaginations are emphasized in this gray image of the same 3q-arm domain section. Numbers indicate the location of DMs 1 – 5 in D. (F–G) 3D reconstruction of the entire 3q-domain (green) with all associated DMs (red). (F) top-view and (G) section through the reconstructed domain. (H,I) 3D reconstruction of a rat liver cell nucleus specifically stained for DNA and imaged by Serial Block-Face—Scanning Electron Microscopy (Rouquette et al. 2009). (H ) A reconstructed mid part of the nucleus with 250-nm thickness shows a strongly inhomogeneous distribution of DNA in higher-order chromatin clusters (gray) and the wide mostly DNA free interchromatin compartment (white) expanding between these clusters. (I ) Reconstruction of the major part of the nucleus. The dotted line indicates the removal of the nuclear periphery exposed to the viewer. Here, the nuclear volume seems to be entirely filled with chromatin, most likely—as we suggest—as a consequence of the sponge-like organization of CTs built up from interconnected chromatin domains/bundles permeated by the IC.

T. Cremer and M. Cremer

ACKNOWLEDGMENTS

This work was supported by DFG (Cr 59/28-1) and CIPSM. The authors are grateful to the many colleagues of the group of the Cremer laboratory who contributed with their experiments, ideas, and enthusiasm to unravel the nature of CTs.

REFERENCES Abney JR, Cutler B, Fillbach ML, Axelrod D, Scalettar BA. 1997. Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J Cell Biol 137: 1459– 1468. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. 2008. Molecular biology of the cell. Garland Science, New York. Albiez H, Cremer M, Tiberi C, Vecchio L, Schermelleh L, Dittrich S, Kupper K, Joffe B, Thormeyer T, von Hase J, et al. 2006. Chromatin domains and the interchromatin compartment form structurally defined and functionally interacting nuclear networks. Chromosome Res 14: 707 – 733. Babu DA, Chakrabarti SK, Garmey JC, Mirmira RG. 2008. Pdx1 and b2/NeuroD1 participate in a transcriptional complex that mediates short-range DNA looping at the insulin gene. J Biol Chem 283: 8164– 8172. Barr ML, Bertram EG. 1949. A morphological distinction between neurons of the male and female, and the behavior of the nucleolar satellite during accelerated nucleoprotein synthesis. Nature 163: 676–677. Bartkuhn M, Renkawitz R. 2008. Long range chromatin interactions involved in gene regulation. Biochim Biophys Acta 1783: 2161–2166. Belmont AS, Bruce K. 1994. Visualization of G1 chromosomes: A folded, twisted, supercoiled chromonema model of interphase chromatid structure. J Cell Biol 127: 287– 302. Berezney R, Malyavantham KS, Pliss A, Bhattacharya S, Acharya R. 2005. Spatio-temporal dynamics of genomic organization and function in the mammalian cell nucleus. Adv Enzyme Regul 45: 17– 26. Berr A, Pecinka A, Meister A, Kreth G, Fuchs J, Blattner FR, Lysak MA, Schubert I. 2006. Chromosome arrangement and nuclear architecture but not centromeric sequences are conserved between Arabidopsis thaliana and Arabidopsis lyrata. Plant J 48: 771 –783. Bolzer A, Craig JM, Cremer T, Speicher MR. 1999. A complete set of repeat-depleted, PCR-amplifiable, human chromosome-specific painting probes. Cytogenet Cell Genet 84: 233 –240. Bolzer A, Kreth G, Solovei I, Koehler D, Saracoglu K, Fauth C, Muller S, Eils R, Cremer C, Speicher MR, et al. 2005. Three-dimensional maps of all chromosomes in human male fibroblast nuclei and prometaphase rosettes. PLoS Biol 3: e157.

Borden J, Manuelidis L. 1988. Movement of the X chromosome in epilepsy. Science 242: 1687– 1691. Bornfleth H, Edelmann P, Zink D, Cremer T, Cremer C. 1999. Quantitative motion analysis of subchromosomal foci in living cells using four-dimensional microscopy. Biophys J 77: 2871–2886. Boutanaev AM, Mikhaylova LM, Nurminsky DI. 2005. The pattern of chromosome folding in interphase is outlined by the linear gene density profile. Mol Cell Biol 25: 8379–8386. Boveri T. 1909. Die Blastomerenkerne von Ascaris megalocephala und die Theorie der Chromosomenindividualita¨t. Arch Zellforsch 3: 181– 268. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211 –219. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol 4: e138. Brianna Caddle L, Grant JL, Szatkiewicz J, van Hase J, Shirley BJ, Bewersdorf J, Cremer C, Arneodo A, Khalil A, Mills KD. 2007. Chromosome neighborhood composition determines translocation outcomes after exposure to high-dose radiation in primary cells. Chromosome Res 15: 1061–1073. Bridger JM, Herrmann H, Munkel C, Lichter P. 1998. Identification of an interchromosomal compartment by polymerization of nuclear-targeted vimentin. J Cell Sci 111: 1241– 1253. Bridger JM, Kalla C, Wodrich H, Weitz S, King JA, Khazaie K, Krausslich HG, Lichter P. 2005. Nuclear RNAs confined to a reticular compartment between chromosome territories. Exp Cell Res 302: 180–193. Bystricky K, Laroche T, van Houwe G, Blaszczyk M, Gasser SM. 2005. Chromosome looping in yeast: telomere pairing and coordinated movement reflect anchoring efficiency and territorial organization. J Cell Biol 168: 375 –387. Cavalli G. 2007. Chromosome kissing. Curr Opin Genet Dev 17: 443– 450. Chakalova L, Debrand E, Mitchell JA, Osborne CS, Fraser P. 2005. Replication and transcription: shaping the landscape of the genome. Nat Rev Genet 6: 669–677. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215–2223. Chubb JR, Bickmore WA. 2003. Considering nuclear compartmentalization in the light of nuclear dynamics. Cell 112: 403–406. Clemson CM, Hall LL, Byron M, McNeil J, Lawrence JB. 2006. The X chromosome is organized into a gene-rich outer rim and an internal core containing silenced nongenic sequences. Proc Natl Acad Sci 103: 7688– 7693. Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R, Fakan S. 1999. Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211 –223.

Chromosome Territories Cremer C, Rappold G, Gray JW, Muller CR, Ropers HH. 1984a. Preparative dual-beam sorting of the human Y chromosome and in situ hybridization of cloned DNA probes. Cytometry 5: 572 –579. Cremer M, Grasser F, Lanctot C, Muller S, Neusser M, Zinner R, Solovei I, Cremer T. 2008. Multicolor 3D fluorescence in situ hybridization for imaging interphase chromosomes. Methods Mol Biol 463: 205 –239. Cremer M, Kupper K, Wagler B, Wizelman L, von Hase J, Weiland Y, Kreja L, Diebold J, Speicher MR, Cremer T. 2003. Inheritance of gene density-related higher order chromatin arrangements in normal and tumor cell nuclei. J Cell Biol 162: 809–820. Cremer M, von Hase J, Volm T, Brero A, Kreth G, Walter J, Fischer C, Solovei I, Cremer C, Cremer T. 2001. Nonrandom radial higher-order chromatin arrangements in nuclei of diploid human cells. Chromosome Res 9: 541 – 567. Cremer T, Baumann H, Nakanishi K, Cremer C. 1984b. Correlation between interphase and metaphase chromosome arrangements as studied by laser-uv-microbeam experiments. Chromosomes Today 8: 203– 212. Cremer T, Cremer C. 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2: 292– 301. Cremer T, Cremer C. 2006a. Rise, fall and resurrection of chromosome territories: a historical perspective. Part I. The rise of chromosome territories. Eur J Histochem 50: 161 –176. Cremer T, Cremer C. 2006b. Rise, fall and resurrection of chromosome territories: a historical perspective. Part II. Fall and resurrection of chromosome territories during the 1950s to 1980s. Part III. Chromosome territories and the functional nuclear architecture: experiments and models from the 1990s to the present. Eur J Histochem 50: 223–272. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982a. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Cremer T, Cremer C, Schneider T, Baumann H, Hens L, Kirsch-Volders M. 1982b. Analysis of chromosome positions in the interphase nucleus of Chinese hamster cells by laser-UV-microirradiation experiments. Hum Genet 62: 201 –209. Cremer T, Kurz A, Zirbel R, Dietzel S, Rinke B, Schrock E, Speicher MR, Mathieu U, Jauch A, Emmerich P, et al. 1993. Role of chromosome territories in the functional compartmentalization of the cell nucleus. Cold Spring Harb Symp Quant Biol 58: 777– 792. Cremer T, Lichter P, Borden J, Ward DC, Manuelidis L. 1988. Detection of chromosome aberrations in metaphase and interphase tumor cells by in situ hybridization using chromosome-specific library probes. Hum Genet 80: 235–246. Croft JA, Bridger JM, Boyle S, Perry P, Teague P, Bickmore WA. 1999. Differences in the localization and morphology of chromosomes in the human nucleus. J Cell Biol 145: 1119–1131.

Cvackova Z, Masata M, Stanek D, Fidlerova H, Raska I. 2009. Chromatin position in human HepG2 cells: although being non-random, significantly changed in daughter cells. J Struct Biol 165: 107– 117. Dehghani H, Dellaire G, Bazett-Jones DP. 2005. Organization of chromatin in the interphase mammalian cell. Micron 36: 95–108. Dekker J. 2008. Gene regulation in the third dimension. Science 319: 1793– 1794. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Deniaud E, Bickmore WA. 2009. Transcription and the nuclear periphery: Edge of darkness? Curr Opin Genet Dev 19: 187– 191. Dietzel S, Jauch A, Kienle D, Qu G, Holtgreve-Grez H, Eils R, Munkel C, Bittner M, Meltzer PS, Trent JM, et al. 1998. Separate and variably shaped chromosome arm domains are disclosed by chromosome arm painting in human cell nuclei. Chromosome Res 6: 25– 33. Edelmann P, Bornfleth H, Zink D, Cremer T, Cremer C. 2001. Morphology and dynamics of chromosome territories in living cells. Biochim Biophys Acta 1551: M29–39. Eils R, Dietzel S, Bertin E, Schrock E, Speicher MR, Ried T, Robert-Nicoud M, Cremer C, Cremer T. 1996. Threedimensional reconstruction of painted human interphase chromosomes: Active and inactive X chromosome territories have similar volumes but differ in shape and surface structure. J Cell Biol 135: 1427– 1440. Fakan S, Bernhard W. 1971. Localisation of rapidly and slowly labelled nuclear RNA as visualized by high resolution autoradiography. Exp Cell Res 67: 129 –141. Fakan S, van Driel R. 2007. The perichromatin region: A functional compartment in the nucleus that determines large-scale chromatin folding. Semin Cell Dev Biol 18: 676 –681. Fawcett JJ, Longmire JL, Martin JC, Deaven LL, Cram LS. 1994. Large-scale chromosome sorting. Methods Cell Biol 42: 319 –330. Federico C, Scavo C, Cantarella CD, Motta S, Saccone S, Bernardi G. 2006. Gene-rich and gene-poor chromosomal regions have different locations in the interphase nuclei of cold-blooded vertebrates. Chromosoma 115: 123– 128. Fedorova E, Zink D. 2008. Nuclear architecture and gene regulation. Biochim Biophys Acta 1783: 2174–2184. Fedorova E, Zink D. 2009. Nuclear genome organization: common themes and individual patterns. Curr Opin Genet Dev 19: 166– 171. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Fraser P, Bickmore W. 2007. Nuclear organization of the genome and the potential for gene regulation. Nature 447: 413–417. Fuks F. 2005. DNA methylation and histone modifications: teaming up to silence genes. Curr Opin Genet Dev 15: 490 –495. Gerlich D, Beaudouin J, Kalbfuss B, Daigle N, Eils R, Ellenberg J. 2003. Global chromosome positions are transmitted through mitosis in mammalian cells. Cell 112: 751 –764.

T. Cremer and M. Cremer Gilbert N, Boyle S, Fiegler H, Woodfine K, Carter NP, Bickmore WA. 2004. Chromatin architecture of the human genome: gene-rich domains are enriched in open chromatin fibers. Cell 118: 555–566. Goetze S, Mateos-Langerak J, Gierman HJ, de Leeuw W, Giromus O, Indemans MH, Koster J, Ondrej V, Versteeg R, van Driel R. 2007. The three-dimensional structure of human interphase chromosomes is related to the transcriptome map. Mol Cell Biol 27: 4475–4487. Grasser F, Neusser M, Fiegler H, Thormeyer T, Cremer M, Carter NP, Cremer T, Muller S. 2008. Replicationtiming-correlated spatial chromatin arrangements in cancer and in primate interphase nuclei. J Cell Sci 121: 1876– 1886. Gray JW, Dean PN, Fuscoe JC, Peters DC, Trask BJ, van den Engh GJ, Van Dilla MA. 1987. High-speed chromosome sorting. Science 238: 323 –329. Gunkel M, Erdel F, Rippe K, Lemmer P, Kaufmann R, Hormann C, Amberger R, Cremer C. 2009. Dual color localization microscopy of cellular nanostructures. Biotechnol J 4: 927 –938. Habermann FA, Cremer M, Walter J, Kreth G, von Hase J, Bauer K, Wienberg J, Cremer C, Cremer T, Solovei I. 2001. Arrangements of macro- and microchromosomes in chicken cells. Chromosome Res 9: 569–584. Heard E, Bickmore W. 2007. The ins and outs of gene regulation and chromosome territory organisation. Curr Opin Cell Biol 19: 311 –316. Hell SW. 2007. Far-field optical nanoscopy. Science 316: 1153– 1158. Hepperger C, Mannes A, Merz J, Peters J, Dietzel S. 2008. Three-dimensional positioning of genes in mouse cell nuclei. Chromosoma 117: 535–551. Jackson DA, Pombo A. 1998. Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J Cell Biol 140: 1285– 1295. Jaunin F, Fakan S. 2002. DNA replication and nuclear architecture. J Cell Biochem 85: 1 –9. Jiang C, Pugh BF. 2009. Nucleosome positioning and gene regulation: advances through genomics. Nat Rev Genet 10: 161 –172. Kalmarova M, Smirnov E, Kovacik L, Popov A, Raska I. 2008. Positioning of the NOR-bearing chromosomes in relation to nucleoli in daughter cells after mitosis. Physiol Res 57: 421– 425. Khalil A, Grant JL, Caddle LB, Atzema E, Mills KD, Arneodo A. 2007. Chromosome territories have a highly nonspherical morphology and nonrandom positioning. Chromosome Res 15: 899 –916. Kioussis D. 2005. Gene regulation: Kissing chromosomes. Nature 435: 579–580. Koehler D, Zakhartchenko V, Froenicke L, Stone G, Stanyon R, Wolf E, Cremer T, Brero A. 2009. Changes of higher order chromatin arrangements during major genome activation in bovine preimplantation embryos. Exp Cell Res 315: 2053–2063. Kosak ST, Groudine M. 2004. Form follows function: The genomic organization of cellular differentiation. Genes Dev 18: 1371–1384.

Kouzarides T. 2007. Chromatin modifications and their function. Cell 128: 693 –705. Kozubek S, Lukasova E, Jirsova P, Koutna I, Kozubek M, Ganova A, Bartova E, Falk M, Pasekova R. 2002. 3D Structure of the human genome: order in randomness. Chromosoma 111: 321– 331. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Kupper K, Kolbl A, Biener D, Dittrich S, von Hase J, Thormeyer T, Fiegler H, Carter NP, Speicher MR, Cremer T, et al. 2007. Radial chromatin positioning is shaped by local gene density, not by gene expression. Chromosoma 116: 285–306. LaSalle JM, Lalande M. 1996. Homologous association of oppositely imprinted chromosomal domains. Science 272: 725–728. Lichter P, Cremer T, Borden J, Manuelidis L, Ward DC. 1988a. Delineation of individual human chromosomes in metaphase and interphase cells by in situ suppression hybridization using recombinant DNA libraries. Hum Genet 80: 224– 234. Lichter P, Cremer T, Tang CJ, Watkins PC, Manuelidis L, Ward DC. 1988b. Rapid detection of human chromosome 21 aberrations by in situ hybridization. Proc Natl Acad Sci 85: 9664–9668. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289– 293. Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269–272. Lomvardas S, Barnea G, Pisapia DJ, Mendelsohn M, Kirkland J, Axel R. 2006. Interchromosomal interactions and olfactory receptor choice. Cell 126: 403 –413. Lyon MF. 1962. Sex chromatin and gene action in the mammalian X-chromosome. Am J Hum Genet 14: 135 –148. Ma H, Samarabandu J, Devdhar RS, Acharya R, Cheng PC, Meng C, Berezney R. 1998. Spatial and temporal dynamics of DNA replication sites in mammalian cells. J Cell Biol 143: 1415–1425. Mahy NL, Perry PE, Gilchrist S, Baldock RA, Bickmore WA. 2002. Spatial organization of active and inactive genes and noncoding DNA within chromosome territories. J Cell Biol 157: 579– 589. Maier VK, Chioda M, Becker PB. 2008. ATP-dependent chromatosome remodeling. Biol Chem 389: 345– 352. Manuelidis L. 1985. Individual interphase chromosome domains revealed by in situ hybridization. Hum Genet 71: 288– 293. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Martou G, De Boni U. 2000. Nuclear topology of murine, cerebellar Purkinje neurons: changes as a function of development. Exp Cell Res 256: 131– 139.

Chromosome Territories Mateos-Langerak J, Bohn M, de Leeuw W, Giromus O, Manders EM, Verschure PJ, Indemans MH, Gierman HJ, Heermann DW, van Driel R, et al. 2009. Spatially confined folding of chromatin in the interphase nucleus. Proc Natl Acad Sci 106: 3812– 3817. Mayer R, Brero A, von Hase J, Schroeder T, Cremer T, Dietzel S. 2005. Common themes and cell type specific variations of higher order chromatin arrangements in the mouse. BMC Cell Biol 6: 44. Meaburn KJ, Misteli T. 2007. Cell biology: chromosome territories. Nature 445: 379 –781. Molnar M, Kleckner N. 2008. Examination of interchromosomal interactions in vegetatively growing diploid Schizosaccharomyces pombe cells by Cre/loxP site-specific recombination. Genetics 178: 99– 112. Morey C, Da Silva NR, Perry P, Bickmore WA. 2007. Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134: 909–919. Morey C, Kress C, Bickmore WA. 2009. Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to up-regulate gene expression. Genome Res 19: 1184–1194. Munkel C, Eils R, Dietzel S, Zink D, Mehring C, Wedemann G, Cremer T, Langowski J. 1999. Compartmentalization of interphase chromosomes observed in simulation and experiment. J Mol Biol 285: 1053– 1065. Munkel C, Langowski J. 1998. Chromosome structure predicted by a polymer model. Phys Rev E57: 5888– 5896. Murmann AE, Gao J, Encinosa M, Gautier M, Peter ME, Eils R, Lichter P, Rowley JD. 2005. Local gene density predicts the spatial position of genetic loci in the interphase nucleus. Exp Cell Res 311: 14–26. Neusser M, Schubel V, Koch A, Cremer T, Muller S. 2007. Evolutionarily conserved, cell type and species-specific higher order chromatin arrangements in interphase nuclei of primates. Chromosoma 116: 307 –320. Parada LA, McQueen PG, Misteli T. 2004. Tissue-specific spatial organization of genomes. Genome Biol 5: R44. Parada LA, McQueen PG, Munson PJ, Misteli T. 2002. Conservation of relative chromosome positioning in normal and cancer cells. Curr Biol 12: 1692–1697. Pecinka A, Schubert V, Meister A, Kreth G, Klatte M, Lysak MA, Fuchs J, Schubert I. 2004. Chromosome territory arrangement and homologous pairing in nuclei of Arabidopsis thaliana are predominantly random except for NOR-bearing chromosomes. Chromosoma 113: 258–269. Pinkel D, Landegent J, Collins C, Fuscoe J, Segraves R, Lucas J, Gray J. 1988. Fluorescence in situ hybridization with human chromosome-specific libraries: detection of trisomy 21 and translocations of chromosome 4. Proc Natl Acad Sci 85: 9138–9142. ¨ ber Zelltheilung. Morph Jb 10: 214– 330. Rabl C. 1885. U Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243–247. Reichenzeller M, Burzlaff A, Lichter P, Herrmann H. 2000. In vivo observation of a nuclear channel-like system: evidence for a distinct interchromosomal domain compartment in interphase cells. J Struct Biol 129: 175 –185.

Richter K, Reichenzeller M, Gorisch SM, Schmidt U, Scheuermann MO, Herrmann H, Lichter P. 2005. Characterization of a nuclear compartment shared by nuclear bodies applying ectopic protein expression and correlative light and electron microscopy. Exp Cell Res 303: 128 –137. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Ronneberger O, Baddeley D, Scheipl F, Verveer PJ, Burkhardt H, Cremer C, Fahrmeir L, Cremer T, Joffe B. 2008. Spatial quantitative analysis of fluorescently labeled nuclear structures: Problems, methods, pitfalls. Chromosome Res 16: 523– 562. Rouquette J, Genoud C, Vazquez-Nin GH, Kraus B, Cremer T, Fakan S. 2009. Revealing the high-resolution threedimensional network of chromatin and interchromatin space: A novel electron-microscopic approach to reconstructing nuclear architecture. Chromosome Res 17: 801 –810. Schardin M, Cremer T, Hager HD, Lang M. 1985. Specific staining of human chromosomes in Chinese hamster x man hybrid cell lines demonstrates interphase chromosome territories. Hum Genet 71: 281– 287. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332–1336. Schneider R, Grosschedl R. 2007. Dynamics and interplay of nuclear architecture, genome organization, and gene expression. Genes Dev 21: 3027–3043. Schroeder-Reiter E, Perez-Willard F, Zeile U, Wanner G. 2009. Focused ion beam (FIB) combined with high resolution scanning electron microscopy: A promising tool for 3D analysis of chromosome architecture. J Struct Biol 165: 97– 106. Shaw PJ, Abranches R, Paula Santos A, Beven AF, Stoger E, Wegel E, Gonza´lez-Melendi P. 2002. The architecture of interphase chromosomes and nucleolar transcription sites in plants. J Struct Biol 140: 31–38. Shopland LS, Lynch CR, Peterson KA, Thornton K, Kepper N, Hase J, Stein S, Vincent S, Molloy KR, Kreth G, et al. 2006. Folding and organization of a contiguous chromosome region according to the gene distribution pattern in primary genomic sequence. J Cell Biol 174: 27– 38. Simonis M, de Laat W. 2008. FISH-eyed and genome-wide views on the spatial organisation of gene expression. Biochim Biophys Acta 1783: 2052– 2060. Simonis M, Klous P, Splinter E, Moshkin Y, Willemsen R, de Wit E, van Steensel B, de Laat W. 2006. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat Genet 38: 1348–1354. Simonis M, Kooren J, de Laat W. 2007. An evaluation of 3C-based methods to capture DNA interactions. Nat Methods 4: 895– 901. Solimando L, Luijsterburg MS, Vecchio L, Vermeulen W, van Driel R, Fakan S. 2009. Spatial organization of nucleotide excision repair proteins after UV-induced DNA damage in the human cell nucleus. J Cell Sci 122: 83– 91.

T. Cremer and M. Cremer Solovei I, Cavallo A, Schermelleh L, Jaunin F, Scasselati C, Cmarko D, Cremer C, Fakan S, Cremer T. 2002. Spatial preservation of nuclear chromatin architecture during three-dimensional fluorescence in situ hybridization (3D-FISH). Exp Cell Res 276: 10–23. Stack SM, Brown DB, Dewey WC. 1977. Visualization of interphase chromosomes. J Cell Sci 26: 281 –299. Solovei I, Kreysing M, Lanctot C, Kosem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356– 368. Strasburger E. 1905. Die stofflichen Grundlagen der Vererbung im organischen Reich. Gustav Fischer, Jena, Germany. Sutherland H, Bickmore WA. 2009. Transcription factories: gene expression in unions? Nat Rev Genet 10: 457–466. Taddei A, Van Houwe G, Hediger F, Kalck V, Cubizolles F, Schober H, Gasser SM. 2006. Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature 441: 774– 778. Tanabe H, Muller S, Neusser M, von Hase J, Calcagno E, Cremer M, Solovei I, Cremer C, Cremer T. 2002. Evolutionary conservation of chromosome territory arrangements in cell nuclei from higher primates. Proc Natl Acad Sci 99: 4424–4429. Telenius H, Pelmear AH, Tunnacliffe A, Carter NP, Behmel A, Ferguson-Smith MA, Nordenskjold M, Pfragner R, Ponder BA. 1992. Cytogenetic analysis by chromosome painting using DOP-PCR amplified flow-sorted chromosomes. Genes Chromosomes Cancer 4: 257 –263. Teller K, Solovei I, Buiting K, Horsthemke B, Cremer T. 2007. Maintenance of imprinting and nuclear architecture in cycling cells. Proc Natl Acad Sci 104: 14970–14975. Thomson I, Gilchrist S, Bickmore WA, Chubb JR. 2004. The radial positioning of chromatin is not inherited through mitosis but is established de novo in early G1. Curr Biol 14: 166 –172. Trentani A, Testillano PS, Risueno MC, Biggiogera M. 2003. Visualization of transcription sites at the electron microscope. Eur J Histochem 47: 195– 200. Van den Engh G, Sachs R, Trask BJ. 1992. Estimating genomic distance from DNA sequence location in cell nuclei by a random walk model. Science 257: 1410– 1412. Verschure PJ, van Der Kraan I, Manders EM, van Driel R. 1999. Spatial relationship between transcription sites and chromosome territories. J Cell Biol 147: 13–24. Visser AE, Aten JA. 1999. Chromosomes as well as chromosomal subdomains constitute distinct units in interphase nuclei. J Cell Sci 112: 3353–3360.

Visser AE, Jaunin F, Fakan S, Aten JA. 2000. High resolution analysis of interphase chromosome domains. J Cell Sci 113: 2585– 2593. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J, et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–1576. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. 2003. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol 160: 685 –697. Williams RR, Broad S, Sheer D, Ragoussis J. 2002. Subchromosomal positioning of the epidermal differentiation complex (EDC) in keratinocyte and lymphoblast interphase nuclei. Exp Cell Res 272: 163–175. Wischnitzer S. 1973. The submicroscopic morphology of the interphase nucleus. Int Rev Cytol 34: 1– 48. Zeitz MJ, Mukherjee L, Bhattacharya S, Xu J, Berezney R. 2009. A probabilistic model for the arrangement of a subset of human chromosome territories in WI38 Human fibroblasts. J Cell Physiol 221: 120–129. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, Singh U, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347. Zink D, Cremer T, Saffrich R, Fischer R, Trendelenburg MF, Ansorge W, Stelzer EH. 1998. Structure and dynamics of human interphase chromosome territories in vivo. Hum Genet 102: 241– 251. Zirbel RM, Mathieu UR, Kurz A, Cremer T, Lichter P. 1993. Evidence for a nuclear compartment of transcription and splicing located at chromosome domain boundaries. Chromosome Res 1: 93–106. Zorn C, Cremer C, Cremer T, Zimmer J. 1979. Unscheduled DNA synthesis after partial UV irradiation of the cell nucleus. Distribution in interphase and metaphase. Exp Cell Res 124: 111 –119. Zorn C, Cremer T, Cremer C, Zimmer J. 1976. Laser UV microirradiation of interphase nuclei and post-treatment with caffeine. A new approach to establish the arrangement of interphase chromosomes. Hum Genet 35: 83–89. Zuckerkandl E, Cavalli G. 2007. Combinatorial epigenetics, “junk DNA”, and the evolution of complex organisms. Gene 390: 232 –242.

Gene Positioning Carmelo Ferrai, Ineˆs Jesus de Castro, Liron Lavitas, Mita Chotalia, and Ana Pombo Genome Function Group, MRC Clinical Sciences Centre, Imperial College School of Medicine, Hammersmith Hospital Campus, Du Cane Road, London W12 0NN, United Kingdom Correspondence: [email protected]

Eukaryotic gene expression is an intricate multistep process, regulated within the cell nucleus through the activation or repression of RNA synthesis, processing, cytoplasmic export, and translation into protein. The major regulators of gene expression are chromatin remodeling and transcription machineries that are locally recruited to genes. However, enzymatic activities that act on genes are not ubiquitously distributed throughout the nucleoplasm, but limited to specific and spatially defined foci that promote preferred higher-order chromatin arrangements. The positioning of genes within the nuclear landscape relative to specific functional landmarks plays an important role in gene regulation and disease.

egulation of gene expression is vital to establishing the metabolic activities in every cell in all organisms, and allows their adaptation to both stress and developmental processes. In humans, more than 20,000 genes and at least an equal number of noncoding transcription units are distributed among the 22 pairs of autosomes and two sex chromosomes. The physical organization of chromosomes within the nucleus occurs in a hierarchical manner. At the most basic level of organization, DNAwinds around octamers of histone molecules leading to the formation of nucleosomes and adjacent nucleosomes are linked via DNA. Nucleosome arrays have an inherent propensity to form compacted, filamentous structures with the extent of condensation proposed to have regulatory potential. Condensed nucleosome arrays form fibers with a diameter of approximately 30 nm (Dorigo et al. 2004), a conformation in which the accessibility of DNA may be

R

restricted. Chromatin condensation can result in masking of regulatory sequences thereby affecting transcription (Cairns 2005). Decondensed, open chromatin is thought to be more accessible to factors necessary for the activation and transcription of genes (Cosma 2002; Sproul et al. 2005), and is also segregated toward the nuclear interior in most cell types (Solovei et al. 2009). Important processes that affect the condensation status of chromatin include the covalent modification of histone tails within nucleosomes and the repositioning of nucleosomes along the DNA fiber (Cosma 2002). At higher levels of chromatin organization, the three-dimensional arrangement of genes in the nuclear space also influences gene expression. Radial positioning of genes mostly correlates with states of expression, with active genes often located in the interior and silent genes at the periphery (Takizawa et al. 2008a; Takizawa et al. 2008b). However, radial

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000588 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000588

C. Ferrai et al.

positioning of genes is unlikely to be a major determinant of gene expression per se, as chromatin regulator complexes do not display radial gradients of abundance, but are instead homogenously distributed or preferentially localized in nuclear subcompartments. The association of inactive genes with the nuclear periphery, which is enriched for transcriptionally silent heterochromatin in most cell types, suggests a molecular mechanism for gene repression. For example, the human CFTR gene associates with peripheral heterochromatin in cell types where the locus is silent, but occupies more internal positions in cell types where CFTR is expressed (Zink et al. 2004). In contrast, the highly conserved murine CFTR homologue and adjacent genes are more centrally positioned irrespective of their expression (Sadoni et al. 2008), possibly because of chromosomal arrangement constraints outside of the Cftr locus. A correlation between gene silencing and positioning at the nuclear periphery has also been shown for several differentiationspecific genes in various cell systems (Brown et al. 1999; Skok et al. 2001; Kosak et al. 2002; Kim et al. 2004; Williams et al. 2006). More recently, the silencing effect of direct associations to the nuclear lamina have been investigated by artificial tethering of reporter genes (Andrulis et al. 1998; Finlan et al. 2008; Reddy et al. 2008). Many genes are inactivated in response to an induced association to the lamina, despite other genes remaining active; these conflicting responses suggest that individual genes have differential sensitivity to the silencing mechanisms at the periphery. Gene neighborhoods constitute a major determinant of gene positioning as each gene is not free to establish its preferred position, but responds to the properties of neighboring genes in the same chromosome. Chromosomes occupy discrete territories within the interphase nucleus in preferred, yet nondeterministic positions that correlate with cell type-specific patterns of gene expression (Cremer et al. 1982; Parada and Misteli 2002). Painting of whole chromosomes in human activated lymphocytes, at high-resolution using a fluorescence in situ hybridization (FISH) approach on

ultrathin cryosections, shows that 20% of the nuclear volume contains intermingled chromatin from neighboring chromosomes, and that a specific inducible locus (MHCII) can be positioned within other CTs (Branco and Pombo 2006). The close spatial proximity between chromosomes has cell type specificity and correlates with increased translocation probability (Roix et al. 2003; Kuroda et al. 2004; Osborne et al. 2007). Chromosome positioning depends on the physical constraints of packaging large chromatin polymers within a small nuclear volume, and is influenced by cell-type specific gene expression programs. In spherical nuclei, such as lymphocytes, the radial positioning of chromosomes correlates with their gene density, with gene-dense and gene-poor chromosomes positioned centrally or at the periphery, respectively (Boyle et al. 2001; Branco et al. 2008). In cells with flat nonspherical nuclei, such as fibroblasts, the size of the chromosome correlates with the radial position, with smaller chromosomes occupying central positions of the nucleus and larger chromosomes positioned toward the periphery independently of gene density (Bolzer et al. 2005). Gene expression is generally controlled by the combinatorial binding of a multitude of transcription factors to specific DNA sequences, which can be located either immediately upstream of protein coding sequences (at promoters) or at considerable distances, upstream or downstream, from the gene. For example, the locus-control region (LCR) at the murine b-globin cluster is located 60 kb upstream of the adult bmajor globin gene (Hbb1) (Dean 2006). Activating transcription factors act either by promoting interactions between transcription start sites and RNA polymerase (RNAP) enzymes or associated transcription machineries, or by recruiting chromatin modifiers that establish local open chromatin states, which are permissive for transcription (Szutorisz et al. 2005). The complex three-dimensional network of chromatin interactions likely reflects the transcriptional program of the cell, in which genes assume “on” or “off ” states according to their local chromatin associations and to their

Gene Positioning

positioning within specific nuclear neighborhoods with activating or silencing features, respectively (Fig. 1) (Dillon 2006; Misteli 2007). GENE POSITIONING AT THE LOCAL LEVEL: CHROMATIN INTERACTIONS BETWEEN REGULATORY ELEMENTS

The transcriptional activation of genes is often described by the simple binding of transcription factors to specific regulatory sequences at variable distances from transcription start sites, followed by the recruitment of RNAP and associated regulatory factors. Novel methodological approaches have enabled the identification of long-range chromatin interactions between regulatory elements and the genes they regulate across the genome, thereby permitting the study of gene regulation in three dimensions. A major breakthrough in this area was the development of the chromosome conformation capture (3C) method, a molecular approach first used to study the three-dimensional folding of Saccharomyces cerevisiae chromosome III (Dekker et al. 2002). In the 3C technique, chromatin is extracted from whole nuclei after chemical cross-linking of DNA-protein complexes and digestion with restriction enzymes or sonication. Intermolecular DNA ligation at low DNA concentration promotes ligation between protein-crosslinked DNA fragments because of their local proximity (Dekker et al. 2002). Interaction partners can then be identified using primers. 3C-based technologies have been extensively used to examine the mechanism of gene regulation at the b-globin gene cluster where the LCR and additional hypersensitive sites form an active chromatin hub with the Hbb1 promoter in a tissue-specific manner (Tolhuis et al. 2002). The conformation of the active hub changes throughout erythrocyte development, in agreement with gene activity at the locus (Palstra et al. 2003), and is dependent on specific regulatory factors (Drissen et al. 2004; Vakoc et al. 2005; Splinter et al. 2006). These findings illustrate that intrachromosomal associations, driven by specific interactions between chromatin binding factors, are an

additional feature of gene regulation. 3C-based approaches have also been widely used to show long-range chromatin interactions at other loci, specifically between distal enhancers and gene promoters (Eivazova and Aune 2004; Spilianakis and Flavell 2004; Liu and Garrard 2005; Amano et al. 2009), but also between promoter and terminator sequences of the same gene, dependent on RNAPII (O’Sullivan et al. 2004; Tan-Wong et al. 2009). Chromatin looping has also been observed at silencers, insulators and boundary elements, which are thought to limit the interaction of enhancers to their target gene promoters. For example, differentially methylated regions (DMRs), involved in the control of parental specific gene expression, establish specific loop structures that may partition differentially expressed genes into distinct loops of active versus silent domains (Murrell et al. 2004). Cell-type specific chromatin interactions mediated by chromatin-associated proteins such as Polycomb repressor complexes, SATB1 and 2, CTCF, and cohesins, have also been identified. For example, cohesin and CTCF binding sites at the developmentally regulated cytokine locus IFNg, in human TH1 lymphocytes, correlate with chromatin interactions in cis, although CTCF is not sufficient to maintain these interactions in the absence of cohesin (Hadjur et al. 2009). GENE POSITIONING AT HIGHER LEVEL: CHROMATIN INTERACTIONS BETWEEN DISTAL GENES

Chromatin interactions between functionally related genes on the same or different chromosomes have also been described by both 3Cbased approaches and by single-cell imaging using FISH. The most extensively studied example is that of the murine b-globin gene, Hbb1, which has been shown to interact with other erythroid-specific genes, Eraf and Uros that are located approximately 40 Mb downstream of Hbb1 on chromosome 7 (Osborne et al. 2004; Simonis et al. 2006). Although it remains unclear whether such interactions are necessary for the correct temporal onset of erythroid

C. Ferrai et al. PcG bodies

Splicing speckles

Centromeric clusters

Cajal bodies

RNAPII factories

Chromosome intermingling Nucleolus

Chromosome territories

Nuclear lamina

PML bodies

Gene

Cajal bodies

PML bodies

RNAPII factories

PcG bodies

Nuclear lamina

Splicing speckles

Centromeric clusters

Nuclear pore

Figure 1. Nuclear subcompartments. An illustration of the mammalian cell nucleus showing nuclear domains and features of gene positioning (orange bar) that have been identified thus far. The nuclear positioning of genes and their associations with different nuclear landmarks are implicated in gene activation and gene repression as discussed in the text. DNA counterstain in blue where applicable. Centromeric clusters. Human CD2 transgenes (green) colocalize with a centromeric cluster (red) in nonexpressing T cells from hCD2 transgenic mice (line 1.3A14). Reprinted by permission from (Hiragami-Hamada et al. 2009).

Gene Positioning

transcription, they appear to involve transcription factories (Osborne et al. 2004; Schoenfelder et al. 2010) or the larger splicing speckles (Brown et al. 2006; Brown et al. 2008; Hu et al. 2009). Both compartments are associated with phosphorylated forms of RNAPII (Xie et al. 2006), but transcription factories are active sites of transcription containing many active RNAPs and associated nascent transcripts (Iborra et al. 1996; Jackson et al. 1998; Pombo et al. 1999), whereas splicing speckles are enriched for a pool of splicing factors and RNAPII complexes that remain stably associated upon transcriptional inhibition (Hall et al. 2006; Xie et al. 2006; Lawrence and Clemson 2008). Association of speckles with highly active genes or with genes that have complex splicing reactions (Xing et al. 1995; Jolly et al. 1999; Smith et al. 1999; Johnson et al. 2000; Shopland et al. 2002; Shopland et al. 2003; Hall et al. 2006; Smith et al. 2007) suggests that gene proximity to splicing speckles may increase the efficiency of RNA splicing and thereby influence the levels of gene expression. Other interchromosomal associations between genes, which correlate with either gene activation or repression, have been described in many different systems (Harmon and Sedat 2005; Spilianakis et al. 2005; Bacher et al. 2006; Ling et al. 2006; Lomvardas et al. 2006; Xu et al. 2006; Apostolou and Thanos 2008; Noordermeer et al. 2008; Schoenfelder et al. 2010). The initial 3C technique revealed many

interesting long-range interactions but was limited to the analyses of a small number of sequences requiring prior knowledge of both interacting baits. Developments of the 3C technique, including circular chromosomal conformation capture (Zhao et al. 2006) and chromosome conformation capture-on-chip (Simonis et al. 2006) (both called 4C), use inverse PCR primers to amplify circular DNA obtained by the ligation of two proximal segments of chromatin. These extended applications of 3C technology enable the simultaneous identification of many interactions using single known baits. Unbiased genome-wide strategies have also emerged. In yeast high-throughput sequencing of 3C-ligation products and bioinformatic identification of ligated fragments has revealed novel chromatin arrangements (Rodley et al. 2009). Two independent approaches in mammalian systems, Hi-C (Lieberman-Aiden et al. 2009) and chromatin interaction analysis by paired-end tag sequencing (ChIA-PET) (Fullwood et al. 2009) have allowed unbiased identification of ligation products by tagging the ligation junction and then enriching the region using affinity purification. In contrast to Hi-C, the ChIA-PET approach enriches 3C products associated with a specific protein of interest thereby identifying protein-based interchromatin networks. One inherent limitation of the 3C-derived approaches is its population-based analysis

Figure 1. (Continued ) Splicing speckles. The C3/C4 genomic region of murine chromosome 8 (red) associates

with splicing speckles (green) in fetal liver cells. Reprinted by permission from (Noordermeer et al. 2008). PcG bodies. Bxd gene (red) positioned at PcG bodies (green) in the posterior part of wild-type stage 5-8 hour Drosophila embryos examined by 2D-FISH. Reprinted by permission from Macmillan Publishers Ltd: Nature Cell Biology (Lanzuolo et al. 2007), copyright (2007). Chromosome intermingling. Intermingling between chromosomes 5 (green) and 7 (red) in human lymphocytes. Reproduced with permission from (Branco and Pombo 2006). Chromosome territories. uPA locus (green) is positioned inside of its chromosome territory (CT; chromosome 10, red) in HepG2 cells. Reproduced with permission from (Ferrai et al. 2010). Nuclear lamina. Igh locus (green) interaction with the nuclear lamina (LMNB1, red) in NIH3T3. Reprinted by permission from Macmillan Publishers Ltd: Nature (Reddy et al. 2008), copyright (2008). PML bodies. MHCII region (red) is proximal to PML bodies (green) in control fibroblasts. Reproduced with permission from (Shiels et al. 2001). Cajal bodies. Simultaneous hybridization of RNU1, RNU2, RNU3, RNU7, RNU12, HIST1, and HIST2 loci show several simultaneous gene interactions with Cajal bodies (red). Reprinted from (Frey et al. 1999). Copyright (1999), with permission from Elsevier. RNAPII factories. Association of hCD2 transgenes with RNAPII-S2p in T cells from hCD2 transgenic mouse line 1.3B (SQ Xie, R Festenstein, A Pombo, unpubl.).

C. Ferrai et al.

that gives rise to average maps of chromatin interactions unlikely to exist in an individual cell. High resolution analyses of physical proximity between 3C partners using cryo-FISH (Simonis et al. 2006) or confocal microscopy (Lieberman-Aiden et al. 2009; Schoenfelder et al. 2010) show remarkably low frequencies of association at distances below 500 nm, which are not consistent with close molecular interactions. Chromatin associations detected by 3C may not always be directly related to regulatory interactions, but may be inherent to chromosome organization and/or promoted by independent coassociation to large nuclear structures. Cytological approaches that measure the frequency of locus-specific interactions at the single cell level remain an important complement to molecular 3C approaches, albeit limited to the analyses of few sequences. GENE POSITIONING RELATIVE TO SITES OF TRANSCRIPTION: TRANSCRIPTION FACTORIES

Different classes of genes are transcribed by the three nuclear RNAPs type I, II, or III: RNAPI transcribes the 45S ribosomal RNA genes tandemly repeated in the genome, RNAPII transcribes messenger RNAs and small structural RNAs, and RNAPIII transcribes the 5S rRNA, transfer RNA and other small structural RNA genes. Visualization of transcriptional activity after pulse-labeling of tagged nucleotides showed that genes are active in distinct subnuclear domains termed transcription factories (Jackson et al. 1993; Wansink et al. 1993). The limited number of transcription sites per nucleus as compared with the larger number of active transcription units suggests that many active RNAPs and associated transcription units are present in each transcription factory (Iborra et al. 1996; Jackson et al. 1998; Kimura et al. 1999; Pombo et al. 1999). Evidence that most active genes are associated with only one or a few active RNAPs at any given time led to the idea that each transcription factory simultaneously transcribes more than one gene (Cook 1999). Support for this model

initially came from observations that Hbb1, Eraf, and Uros genes, separated by several Mb, coassociate in a shared factory when transcribed (Osborne et al. 2004). A recent analysis of a larger number of genes in the same cell type shows proximity between Hbb1 and additional genes separated by large genomic regions within the same chromosome and between genes from different chromosomes, albeit specific associations are not required for transcriptional activity of the genes analyzed (Schoenfelder et al. 2010). Other studies also support the functional specialization of transcription factories, depending on the type of RNAP (Pombo et al. 1999), gene structure (Xu and Cook 2008) (Schoenfelder et al. 2010), or activation state of RNAPII (Ferrai et al. 2010).

GENE POSITIONING RELATIVE TO NUCLEOPLASMIC SUBCOMPARTMENTS Splicing Speckles

Splicing speckles are major subnuclear compartments characterized by an accumulation of components of the splicing machinery, polyAþ RNA and numerous mRNA metabolic factors, as well as RNAPII (Hall et al. 2006; Xie et al. 2006; Lawrence and Clemson 2008). Several splicing speckle domains (10 – 30) are typically present in each nucleus, measuring 0.5 – 3 mm in diameter and occupying 5% –10% of the nucleoplasmic volume (Spector 1993; Fay et al. 1997; Xie et al. 2006). Subchromosomal R bands, containing 210 Mb blocks of gene-rich genomic sequence, are frequently found associated with speckles (Shopland et al. 2003). A subset of genes such as heat-shock genes, erythroid specific genes, collagen genes, and muscle specific genes preferentially associate with SC35 domains when active (Xing et al. 1995; Jolly et al. 1999; Smith et al. 1999; Johnson et al. 2000; Shopland et al. 2002; Shopland et al. 2003; Hall et al. 2006; Smith et al. 2007). In vivo studies using multicopy BAC transgenes, containing the inducible Hsp70 gene tagged with Lac operator sequences and visualized on GFP-LacI expression, show increased association of the transgene with

Gene Positioning

splicing speckles within 30 min of heat shock activation (Hu et al. 2009). Coassociation of genes that share the same pathway has been observed at speckles in skeletal muscle and in erythroid cells. The musclespecific genes, myf-4, MyoD, and cMyHC, important for skeletal muscle differentiation, localize to SC35 domains in differentiated muscle cells, but not in the undifferentiated satellite myoblasts or fibroblasts (Moen et al. 2004). Erythroid genes, Hbb, Hba, Eraf, Slc4a1, and Gata1, located on four different murine chromosomes, have also been found nonrandomly positioned relative to common splicing speckles (Brown et al. 2008). Importantly, the association with speckles of genes that have complex splicing reactions and the accumulation of unspliced RNAs at these sites (Johnson et al. 2000; Smith et al. 2007) suggests that gene associations with speckles may influence the yield of mature mRNAs. Cajal Bodies and PML Bodies

Cajal bodies are nucleoplasmic structures, typically one to six per nucleus, with diameters of 0.1 – 1 mm (Gall 2003). They are involved in the biogenesis of several classes of spliceosomal small nuclear ribonucleoprotein particles (snRNPs) and are enriched in RNAP subunits, specific basal transcription factors, and RNA processing factors. U2 snRNA genes associate within Cajal bodies in a mechanism that depends on the U2 gene promoter and is thought to be important for the maturation of newly synthesized U2 snRNAs (Frey et al. 1999; Smith and Lawrence 2000; Frey and Matera 2001). In vivo imaging of U2 snRNA arrays showed that they only associate with Cajal bodies upon induction. The mechanism involves long-range movements toward relatively stably positioned Cajal bodies and is thought to be mediated by actin polymerization (Dundr et al. 2007). Cajal bodies contain the U7 snRNP, a key component of the histone mRNA 30 -end formation machinery (Frey and Matera 1995). Although replication-dependent histone genes that are cell-cycle regulated, including HIST1

and HIST2 multigene clusters, associate with Cajal bodies, their transcriptional activity is not dependent on this association, which appears to be mediated by the nascent transcripts (Jacobs et al. 1999; Schul et al. 1999; Liu et al. 2006; Stanek and Neugebauer 2006). In contrast, cell cycle-independent histone genes, such as H3.3A, which are constitutively active and produce polyadenylated transcripts, do not associate with Cajal bodies (Shopland et al. 2001). Promyelocytic leukemia nuclear bodies (PML) are nuclear multiprotein domains, (Five to 30 per nucleus) with diameters between 0.1 – 1.0 mm that can vary in relation to the cell cycle stage and cell type (Bernardi and Pandolfi 2007). PML bodies contain many proteins and have been implicated in multiple regulatory pathways, from the control of cell proliferation to DNA repair or apoptosis, and in viral infection. PML bodies colocalize with the TP53 locus (tumor protein 53 gene) in Jurkat cells, raising the possibility that PML bodies play a role in regulating TP53 transcription (Sun et al. 2003). Interaction with PML bodies has also been detected in living cells (Tsukamoto et al. 2000), but ongoing transcription appeared independent of PML body association (Tsukamoto et al. 2000). Furthermore, the analysis of multiple genomic regions showed that PML bodies associate with genomic regions characterized by high transcriptional activity (Wang et al. 2004). However, detection of active alleles by RNA-FISH revealed that transcriptional activity is independent of gene association with PML bodies, and PML knock-down does not alter transcription levels (Wang et al. 2004), suggesting that expression does not require association with PML bodies. Polycomb Bodies

Polycomb group (PcG) complexes are involved in heritable gene repression by epigenetic chromatin modifications and by regulating the nuclear organization of their target genes (Schuettengruber et al. 2007). In mammalian and Drosophila cells, several PcG proteins have a nonhomogeneous distribution in the

C. Ferrai et al.

nucleoplasm being enriched in discrete foci called PcG bodies (Saurin et al. 1998; Orlando 2003), which can be preferentially localized near centromeric heterochromatin (Saurin et al. 1998). Whole-mount FISH in Drosophila melanogaster embryos showed that Polycomb response elements (PREs) such as Fab-7 or bxd of the inactive homeotic bithorax complex (BX-C) colocalize with PcG bodies in more than 80% of the anterior nuclei (Lanzuolo et al. 2007). In the posterior part of the embryo, inactive PREs (bxd) remain predominately bound to PcG bodies, whereas active elements (Fab-7) clearly lose contact with these domains, supporting the idea that gene association with PcG bodies is important for gene silencing. The pairing of two Fab-7 PREs from different genomic regions, as well as the coassociation of three PREs (Fab-7, Mcp, and bxd) within the BX-C Hox gene locus, occur within shared PcG bodies (Grimaud et al. 2006; Lanzuolo et al. 2007). A coassociation of several PcG regulated genes in PcG bodies is also suggested by the existence of fewer PcG bodies than PcG target genes identified by genome-wide chromatin immunoprecipitation (ChIP) (Sexton et al. 2009), further supporting the idea that repositioning of genomic regions with respect to nuclear compartments may be important for the regulation of gene expression. ChIP analysis of a major Polycomb protein, Enhancer of Zeste (Ezh2), combined with 3C showed that Ezh2 is present at intra- and interchromosomal hubs that are associated with gene silencing (Tiwari et al. 2008a; Tiwari et al. 2008b). Centromeric Heterochromatin Regions

Gene associations with centromeric heterochromatin regions also correlate with gene silencing. Brown and colleagues analyzed the nuclear position of several genes (CD2, l5, CD8a, Rag, Sox1 and Hoxb4-b9) relative to centromeric regions in mature B lymphocytes, before and following mitogenic CD40mediated activation (Brown et al. 1999). Genes that are not expressed in mature B cells (l5, CD8a, Rag, Sox1, and Hoxb4-b9) were not associated with centromeric chromatin in resting

mature B cells, but became juxtaposed to these domains in cycling mature B cells after mitogenic stimulation. The CD2 gene, which is widely expressed in mature B lymphocytes, remained distant from centromeric domains irrespectively of stimulation. These findings indicate that some inactive genes become associated with centromeric heterochromatic regions during cell division. Analyses of Rag and TdT genes, expressed in CD4þCD8þ thymocytes but down-regulated on T-cell receptor engagement, showed repositioning of these genes to centromeric regions during induced differentiation, resulting in heritable silencing (Brown et al. 1999). Surprisingly, transient silencing of Rag and TdT in a transformed thymocyte cell line was reversible and was not accompanied by repositioning of these genes to centromeric regions, implicating that silencing is initiated through a more complex mechanism. Genes also reposition to centromeric regions on allelic exclusion during B-cell maturation (Skok et al. 2001). In immature B lymphocytes, IgH and IgL alleles are not associated with centromeric regions, but become associated with these regions upon allelic exclusion which leads to B cells expressing functional receptors encoded by a single IgH allele and one of four IgL alleles. The repositioning of genes to centromeric regions is proposed to be mediated through a DNA binding protein, Ikaros/Lyf-1, which is important for lymphocyte development (Brown et al. 1997; Brown et al. 1999). Nevertheless, transgenes integrated in centromeric regions, are not always silenced (Sabbattini et al. 2001). The mouse l5 and VpreB1 genes are expressed in pro- and pre-B cells but silenced in immature and mature B cells, where they are relocated to centromeric regions. A l5-VpreB1-transgene with a mutated Ikaros binding site was released from Ikaros-mediated silencing (Sabbattini et al. 2001). Interestingly, the integration of the mutated l5-VpreB1transgene into centromeric heterochromatin region does not restore silencing at the mature B-cell stage (Sabbattini et al. 2001), showing that transcription can take place in centromeric regions and Ikaros’ pivotal role in mediating l5-VpreB1-silencing on B-cell differentiation.

Gene Positioning

Nuclear Lamina

The nuclear lamina underlays the inner nuclear membrane and consists of type V intermediate filament proteins called nuclear lamins (Dechat et al. 2008). Markers of gene repression, such as enrichment for hypoacetylated chromatin (Sadoni et al. 1999), depletion of both RNAPII and the open chromatin histone modification H3K4me2 (dimethylation of lysine 4 of histone 3) are associated with the nuclear lamina (Luco et al. 2008). Genome-lamina interactions in human occur through more than 1,300 sharply defined large chromatin domains, referred to as lamina associated domains (LADs) (Guelen et al. 2008). These domains are 0.1 – 10 Mb in size in human fibroblasts, are enriched five- 10-fold for genes with lower expression levels than genes outside LADs and are enriched in repressive histone marks, H3K27me3 (trimethylation of lysine 27 on histone 3) and H3K9me2 (dimethylation of lysine 9 on histone 3), but are depleted of the active histone mark H3K4me2. Dynamic repositioning of some genes to the lamina correlates with changes in gene expression. Activation of genes including Hbb during differentiation of mouse erythroid cells (Ragoczy et al. 2006), IgH and IgK in murine B-cell differentiation (Kosak et al. 2002) or Mash1 during murine neuronal differentiation (Williams et al. 2006), correlates with their separation from the repressive lamina environment. Interestingly, some genes initiate transcription while at the nuclear lamina before internal repositioning (Hbb) (Ragoczy et al. 2006) or remain in close proximity to the nuclear periphery upon activation (IFNg) (Hewitt et al. 2004) indicating that the peripheral gene positioning per se is not restrictive for transcription. Induced tethering of reporter genes to the nuclear lamina has shown in yeast and cultured mammalian cells the relevance of physical gene associations with the nuclear periphery for transcriptional activity (Andrulis et al. 1998; Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Although some genes were repressed by the interaction with the nuclear periphery (Andrulis et al. 1998; Finlan et al.

2008; Reddy et al. 2008), others remained fully transcriptionally competent (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). Finlan and colleagues showed that the mechanisms of repression at the nuclear periphery could be alleviated by inhibition of histone deacetylase (HDAC) activity without affecting gene association with the lamina (Finlan et al. 2008), indicating the importance of HDACs in promoting gene repression at the lamina. In all three mammalian studies, transgene repositioning required the passage through mitosis (Finlan et al. 2008; Kumaran and Spector 2008; Reddy et al. 2008). In summary, gene repression at the nuclear lamina is therefore not a general phenomenon but gene specific, depending on multiple parameters such as transcription factor accessibility, promoter strength, existence of insulator elements and pre-existing chromatin marks, which may counteract the mechanisms of transcription repression at the nuclear lamina. Gene silencing mechanisms that correlate with chromatin association to the nuclear periphery have also been observed in lower eukaryotes (Akhtar and Gasser 2007; Towbin et al. 2009). In yeast nuclei, which do not have a nuclear lamina, repressive proteins such as Sir proteins (Sir2, Sir3, and Sir4) are known to interact with subtelomeric heterochromatic regions and such interactions are mostly confined to foci localized at the nuclear envelope. Transcription-dependent associations of specific groups of genes with nuclear pores have also been described from yeast to mammals, suggesting that the nuclear periphery promotes distinct functional environments compatible with both gene repression and activation. CHROMATIN DYNAMICS

Live cell imaging in interphase nuclei shows that chromatin movements occur within a radius of 0.5 – 1.0 mm, which given the large size of the mammalian nucleus (10 mm diameter) suggests that a given locus is constrained to a very small fraction of the total nuclear volume (Abney et al. 1997; Chubb et al. 2002; Walter et al. 2003). Larger chromatin movements

C. Ferrai et al.

(.2 mm) were observed in early G1 phase of the cell cycle, suggesting that chromatin repositioning occurs during the re-establishment of interphase nuclear architecture (Walter et al. 2003). The importance and complexity of chromatin mobility in gene expression is highlighted in examples of gene looping out of their own chromosome territories (CTs) upon gene induction, such as observed for MHCII, EDC and Hox clusters and for the uPA gene (Volpi et al. 2000; Chambeyron and Bickmore 2004; Williams et al. 2006; Ferrai et al. 2010). The repositioning of Hox clusters B and D relative to their CTs and the condensation level of clusters have been studied during retinoic acid-induced differentiation of murine embryonic stem (ES) cells and in embryos (Chambeyron and Bickmore 2004; Chambeyron et al. 2005; Morey et al. 2007). Hox clusters extrude outside their territories and decondense concomitantly with expression. However, locus decondensation occurs independently of locus position relative to its CT and the HoxD locus is preferentially positioned at the CT interior in limb bud embryo cells (embryonic day 9.5) where it is active (Morey et al. 2007), suggesting that locus positioning relative to the CTmay not always be important for gene expression upon induction. RNA-FISH experiments have shown that in the induced state active alleles locate inside, at the edge, and outside of the CT, with a preferential positioning of inactive alleles within the CT (Morey et al. 2009). A more recent analysis of the inducible uPA gene in human HepG2 cells shows that it relocates out of the CT on activation with phorbol esters within 3 hours of treatment (Ferrai et al. 2010). In the active state, uPA gene activity is independent of its position in relation to its CT. Unexpectedly, before tetradecanoyl phorbol acetate (TPA) induction of the gene, the majority of uninduced uPA genes positioned inside the CT are silenced, whereas a small proportion of uPA loci located outside of the CT are more likely to be transcribed. Collectively, these studies suggest that locus positioning inside the CT correlates in some situations with gene silencing, despite the presence of active RNAPII and transcription occurring throughout the volume

of CTs (Abranches et al. 1998; Verschure et al. 1999; Verschure et al. 2003; Branco and Pombo 2006; Ferrai et al. 2010). ALTERED GENE POSITIONING IN DISEASE

Nuclear architecture and chromatin structure are important features for the regulation of gene expression, and can be altered in diseases. Unbalanced levels or mutations of lamin proteins or emerin, an integral protein of the nuclear envelope, are linked to a group of rare genetic disorders called nuclear envelopathies (Webster et al. 2009). Well known nuclear envelopathies such as Hutchinson-Gilford progeria syndrome (HGPS; premature aging) and Emery-Dreifuss muscular dystrophy (EDMD) are associated with mutations of the lamin A/C or emerin genes and are often characterized by alterations in nuclear morphology that affect chromatin organization and may compromise gene expression (Fig. 2A) (Webster et al. 2009). Studies in primary fibroblasts of several nuclear envelopathies (including HGPS and EDMD) show the repositioning of chromosomes 13 and 18 toward the nuclear interior (Fig. 2B) (Meaburn et al. 2007). Repositioning of chromosome 18 to the nuclear interior and up-regulation of genes within this chromosome are also observed after lamin B1 knockout in mouse embryonic fibroblasts (Malhas et al. 2007). Aberrant chromosome distribution in envelopathies only affects specific tissues and cell lineages (Meaburn et al. 2007), as it is not observed in cell types such as cultured lymphoblastoid cells derived from patients with emerin and lamin A mutations (Boyle et al. 2001; Meaburn et al. 2005). Repositioning of genes, small chromatin regions, or CTs is also seen in other diseases, such as facioscapulohumeral muscular dystrophy (FSHD) (Masny et al. 2004), immunodeficiency centromeric instability facial anomalities (ICF) (Matarazzo et al. 2007), epilepsy (Borden and Manuelidis 1988), or in male factor infertility (Finch et al. 2008). ICF syndrome is caused by mutations in DNA methyltransferase 3B (DNMT3B) and is associated with DNA

Gene Positioning

A

Abnormal nuclei in progeria

B

Chromosome repositioning in progeria

C

Gene repositioning in ICF

D

Gene repositioning in cancer

Figure 2. Nuclear organization defects in disease. (A) Abnormal nuclear shape in Hutchinson-Gilford Progeria

Syndrome. Nuclei in patients with progeria show an abnormal morphology (right) contrasting with the regular nuclei shape found in healthy individuals (left). Reproduced with permission from (Scaffidi et al. 2005). (B) Chromosome repositioning in Hutchinson-Gilford Progeria Syndrome. Interior repositioning of human chromosome 13 (green, arrows) in a HGPS fibroblast cell line (right), in comparison to a control cell line (left). Proliferation marker pKi-67 (red). Reproduced with permission from (Meaburn et al. 2007). (C ) Gene repositioning in immunodeficiency centromeric instability facial anomalies (ICF). In control male cells (left), the SYBL1 (red, arrows) inactive allele (Y chromosome) is localized inside or at the edge of the Y CT, whereas in ICF cells (right) it loops out of its CT and escapes silencing. Reproduced with permission from (Matarazzo et al. 2007). Copyright (2007) National Academy of Sciences. (D) Gene repositioning in early mammary tumorigenesis. Position of the cancer AKT1 locus (red, arrows) before (left) and after (right) oncogenic activation of an epithelial cell line. After activation there is a radial shift in the locus position toward the periphery. Reproduced with permission from (Meaburn and Misteli 2008).

hypomethylation at specific genomic sites, such as the SYBL1 gene promoter which becomes derepressed. Interestingly, in ICF male and female patients, the SYBL1 inactive allele, located on the Y and inactive X chromosome, respectively, relocates outside of its respective CT, recapitulating the position of the active alleles (Fig. 2C) (Matarazzo et al. 2007). In females, but not in males, both inactive SYBL1 and its silent neighbor SPRY3 loop out from the CT, suggesting that the reorganization extends beyond the derepressed, hypomethylated gene (Matarazzo et al. 2007). Altered chromosome positioning has also been associated with numerous types of cancer

in which altered gene expression patterns lead to malignant transformation of the cell. Chronic leukemias are associated with enlarged nuclei in lymphoid cells (Rozycka et al. 1988). Abnormal relocation of chromosome 18 from the nuclear periphery to the interior has been documented in several types of tumor cell lines, including melanoma-derived line, cervix carcinoma, colon carcinoma, Hodgkin-derived cell line and colon carcinoma metastasis cells (Cremer et al. 2003). A radial shift of chromosome 8 and a significant change in its shape was observed in pancreatic neoplastic tissue as well as a radial shift of centromeric regions of chromosome 17 and HER2 domains in tumor

C. Ferrai et al.

breast tissues (Wiech et al. 2005). Individual gene loci are also repositioned during early tumorigenesis. In vitro induction of tumorigenesis in early breast cancer showed altered positioning of a set of cancer-associated genes such as AKT1, BCL2, ERB2, and VEGF loci although no correlation was found between radial redistribution and gene activity level (Fig. 2D; (Meaburn and Misteli 2008). The altered positioning of genes has recently been analyzed in invasive breast cancers toward devising prognosis markers of cancer. Eight genes (HES5, ERBB2, MYC, FOSL2, HSP90AA1, AKT1, TGFB3, and CSF1R) were robustly repositioned in breast cancer affected patients in comparison to normal tissues (Meaburn et al. 2009). The origin of chromosomal rearrangements has long been assumed to be a random process, in which a DNA break in one chromosome results in stochastic joining with a DNA segment of another chromosome by defective DNA repair. However, recurrent translocation partners in specific types of cancer have hinted for a contribution of proximal nuclear positioning of the loci involved. Chromosomes that are known to preferentially translocate in specific cell types are more often associated in the nucleus (Roix et al. 2003; Brianna Caddle et al. 2007). Furthermore, the extent of intermingling between specific pairs of chromosomes correlates with the frequency of translocations between the same chromosomes observed after ionizing radiation (Branco and Pombo 2006). Analyses of single genes close to or at translocation breakpoints have also shown that their nuclear positioning is more proximal in cell types where the translocation is observed (Neves et al. 1999; Nikiforova et al. 2000; Roix et al. 2003; Kuroda et al. 2004; Osborne et al. 2007). More recently, studies on anaplastic large cell lymphoma, often associated with t(2;5) translocations, and prostate cancer, with frequent translocations between TMPRSS2 and ERG or ETV1, have shown a direct correlation between increased gene proximity upon transcription induction and translocation (Lin et al. 2009; Mani et al. 2009; Mathas et al. 2009). Considered together, these observations strongly support the notion that physical

proximity of genomic regions, influenced by states of gene activity and cell-type specific genome architecture, can predispose translocations that are characteristic of specific cell types and cancers. CONCLUDING REMARKS

The absence of “membrane-enclosed” subcompartments in the eukaryotic cell nucleus led to early notions that molecules and processes were randomly distributed within this cellular compartment. Work in past decades has shown many features of the structural and functional organization of the eukaryotic cell nucleus. Most nuclear events do not occur throughout the nucleoplasm, but are limited to specific and spatially defined sites (Cook 1999; Misteli 2007). The nuclear volume contains morphological distinct higher-order chromatin domains, such as CTs and condensed heterochromatin, and a number of proteinaceous subcompartments including transcription factories, nuclear lamina, the nucleolus, speckles, and Cajal bodies (Handwerger and Gall 2006). The physically distinct nature of each compartment not only contributes to spatial partitioning of the nuclear space, but also creates distinct functional subdomains within the nucleus. The mechanisms by which spatial positioning of the genome contributes to functional regulation are far from understood, and represent one of the biggest challenges in modern molecular and cell biology. One possibility is that subcompartmentalization of the nucleus influences multiple parameters including transcription factor accessibility, promoter strength, existence of insulator elements and pre-existing chromatin marks, which cumulatively lead to the correct expression signature unique for each cell type. ACKNOWLEDGMENTS

We thank Sheila Q. Xie, Andre´ Mo¨ller and Miguel R. Branco for help researching literature for this article, and the Medical Research Council (UK) for support. C.F. was a recipient of a Wellcome Trust Value in People (VIP) Award.

Gene Positioning

I.J.C. is supported by Fundac¸a˜o para a Cieˆncia e a Tecnologia (Portugal). REFERENCES Abney JR, Cutler B, Fillbach ML, Axelrod D, Scalettar BA. 1997. Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J Cell Biol 137: 1459– 1468. Abranches R, Beven AF, Aragon-Alcaide L, Shaw PJ. 1998. Transcription sites are not correlated with chromosome territories in wheat nuclei. J Cell Biol 143: 5–12. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507 –517. Amano T, Sagai T, Tanabe H, Mizushina Y, Nakazawa H, Shiroishi T. 2009. Chromosomal dynamics at the Shh locus: Limb bud-specific differential regulation of competence and active transcription. Dev Cell 16: 47–57. Andrulis ED, Neiman AM, Zappulla DC, Sternglanz R. 1998. Perinuclear localization of chromatin facilitates transcriptional silencing. Nature 394: 592 –595. Apostolou E, Thanos D. 2008. Virus Infection Induces NF-kB-dependent interchromosomal associations mediating monoallelic IFN-bgene expression. Cell 134: 85–96. Bacher CP, Guggiari M, Brors B, Augui S, Clerc P, Avner P, Eils R, Heard E. 2006. Transient colocalization of X-inactivation centres accompanies the initiation of X inactivation. Nat Cell Biol 8: 293 –299. Bernardi R, Pandolfi PP. 2007. Structure, dynamics and functions of promyelocytic leukaemia nuclear bodies. Nat Rev Mol Cell Biol 8: 1006–1016. Bolzer A, Kreth G, Solovei I, Koehler D, Saracoglu K, Fauth C, Muller S, Eils R, Cremer C, Speicher MR, et al. 2005. Three-dimensional maps of all chromosomes in human male fibroblast nuclei and prometaphase rosettes. PLoS Biol 3: e157. Borden J, Manuelidis L. 1988. Movement of the X chromosome in epilepsy. Science 242: 1687–1691. Boyle S, Gilchrist S, Bridger JM, Mahy NL, Ellis JA, Bickmore WA. 2001. The spatial organization of human chromosomes within the nuclei of normal and emerinmutant cells. Hum Mol Genet 10: 211– 219. Branco MR, Pombo A. 2006. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol 4: e138. Branco MR, Branco T, Ramirez F, Pombo A. 2008. Changes in chromosome organization during PHA-activation of resting human lymphocytes measured by cryo-FISH. Chromosome Res 16: 413 –426. Brianna Caddle L, Grant JL, Szatkiewicz J, van Hase J, Shirley BJ, Bewersdorf J, Cremer C, Arneodo A, Khalil A, Mills KD. 2007. Chromosome neighborhood composition determines translocation outcomes after exposure to high-dose radiation in primary cells. Chromosome Res 15: 1061–1073. Brown KE, Baxter J, Graf D, Merkenschlager M, Fisher AG. 1999. Dynamic repositioning of genes in the nucleus of

lymphocytes preparing for cell division. Mol Cell 3: 207 –217. Brown JM, Green J, das Neves RP, Wallace HA, Smith AJ, Hughes J, Gray N, Taylor S, Wood WG, Higgs DR, et al. 2008. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J Cell Biol 182: 1083–1097. Brown KE, Guest SS, Smale ST, Hahm K, Merkenschlager M, Fisher AG. 1997. Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91: 845–854. Brown JM, Leach J, Reittie JE, Atzberger A, Lee-Prudhoe J, Wood WG, Higgs DR, Iborra FJ, Buckle VJ. 2006. Coregulated human globin genes are frequently in spatial proximity when active. J Cell Biol 172: 177– 187. Cairns BR. 2005. Chromatin remodeling complexes: Strength in diversity, precision through specialization. Curr Opin Genet Dev 15: 185– 190. Chambeyron S, Bickmore WA. 2004. Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription. Genes Dev 18: 1119–1130. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215– 2223. Chubb JR, Boyle S, Perry P, Bickmore WA. 2002. Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol 12: 439–445. Cook PR. 1999. The organization of replication and transcription. Science 284: 1790–1795. Cosma MP. 2002. Ordered recruitment: Gene-specific mechanism of transcription activation. Mol Cell 10: 227 –236. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Cremer M, Kupper K, Wagler B, Wizelman L, von Hase J, Weiland Y, Kreja L, Diebold J, Speicher MR, Cremer T. 2003. Inheritance of gene density-related higher order chromatin arrangements in normal and tumor cell nuclei. J Cell Biol 162: 809 –820. Dean A. 2006. On a chromosome far, far away: LCRs and gene expression. Trends Genet 22: 38–45. Dechat T, Pfleghaar K, Sengupta K, Shimi T, Shumaker DK, Solimando L, Goldman RD. 2008. Nuclear lamins: Major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22: 832– 853. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Dillon N. 2006. Gene regulation and large-scale chromatin organization in the nucleus. Chromosome Res 14: 117 –126. Dorigo B, Schalch T, Kulangara A, Duda S, Schroeder RR, Richmond TJ. 2004. Nucleosome arrays reveal the two-start organization of the chromatin fiber. Science 306: 1571– 1573.

C. Ferrai et al. Drissen R, Palstra RJ, Gillemans N, Splinter E, Grosveld F, Philipsen S, de Laat W. 2004. The active spatial organization of the b-globin locus requires the transcription factor EKLF. Genes Dev 18: 2485–2490. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095– 1103. Eivazova ER, Aune TM. 2004. Dynamic alterations in the conformation of the Ifng gene region during T helper cell differentiation. Proc Natl Acad Sci U S A 101: 251–256. Fay FS, Taneja KL, Shenoy S, Lifshitz L, Singer RH. 1997. Quantitative digital analysis of diffuse and concentrated nuclear distributions of nascent transcripts, SC35 and poly(A). Exp Cell Res 231: 27– 37. Ferrai C, Xie SQ, Luraghi P, Munari D, Ramirez F, Branco MR, Pombo A, Crippa MP. 2010. Poised transcription factories prime silent uPA gene prior to activation. PLoS Biol 8: e1000270. Finch KA, Fonseka KG, Abogrein A, Ioannou D, Handyside AH, Thornhill AR, Hickson N, Griffin DK. 2008. Nuclear organization in human sperm: Preliminary evidence for altered sex chromosome centromere position in infertile males. Hum Reprod 23: 1263–1270. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Frey MR, Matera AG. 1995. Coiled bodies contain U7 small nuclear RNA and associate with specific DNA sequences in interphase human cells. Proc Natl Acad Sci U S A 92: 5915– 5919. Frey MR, Matera AG. 2001. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J Cell Biol 154: 499–509. Frey MR, Bailey AD, Weiner AM, Matera AG. 1999. Association of snRNA genes with coiled bodies is mediated by nascent snRNA transcripts. Curr Biol 9: 126 –135. Fullwood MJ, Liu MH, Pan YF, Liu J, Xu H, Mohamed YB, Orlov YL, Velkov S, Ho A, Mei PH, et al. 2009. An oestrogen-receptor-a-bound human chromatin interactome. Nature 462: 58–64. Gall JG. 2003. The centennial of the Cajal body. Nat Rev Mol Cell Biol 4: 975–980. Grimaud C, Bantignies F, Pal-Bhadra M, Ghana P, Bhadra U, Cavalli G. 2006. RNAi components are required for nuclear clustering of Polycomb group response elements. Cell 124: 957– 971. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. 2008. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453: 948 –951. Hadjur S, Williams LM, Ryan NK, Cobb BS, Sexton T, Fraser P, Fisher AG, Merkenschlager M. 2009. Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 460: 410–413. Hall LL, Smith KP, Byron M, Lawrence JB. 2006. Molecular anatomy of a speckle. Anat Rec A Discov Mol Cell Evol Biol 288: 664– 675.

Handwerger KE, Gall JG. 2006. Subnuclear organelles: New insights into form and function. Trends Cell Biol 16: 19– 26. Harmon B, Sedat J. 2005. Cell-by-cell dissection of gene expression and chromosomal interactions reveals consequences of nuclear reorganization. PLoS Biol 3: e67. Hewitt SL, High FA, Reiner SL, Fisher AG, Merkenschlager M. 2004. Nuclear repositioning marks the selective exclusion of lineage-inappropriate transcription factor loci during T helper cell differentiation. Eur J Immunol 34: 3604–3613. Hiragami-Hamada K, Xie SQ, Saveliev A, Uribe-Lewis S, Pombo A, Festenstein R. 2009. The molecular basis for stability of heterochromatin-mediated silencing in mammals. Epigenetics Chromatin 2: 14. Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009. Large-scale chromatin structure of inducible genes: Transcription on a condensed, linear template. J Cell Biol 185: 87– 100. Iborra FJ, Pombo A, Jackson DA, Cook PR. 1996. Active RNA polymerases are localized within discrete transcription “factories” in human nuclei. J Cell Sci 109: 1427–1436. Jackson DA, Hassan AB, Errington RJ, Cook PR. 1993. Visualization of focal sites of transcription within human nuclei. Embo J 12: 1059– 1065. Jackson DA, Iborra FJ, Manders EM, Cook PR. 1998. Numbers and organization of RNA polymerases, nascent transcripts, and transcription units in HeLa nuclei. Mol Biol Cell 9: 1523– 1536. Jacobs EY, Frey MR, Wu W, Ingledue TC, Gebuhr TC, Gao L, Marzluff WF, Matera AG. 1999. Coiled bodies preferentially associate with U4, U11, and U12 small nuclear RNA genes in interphase HeLa cells but not with U6 and U7 genes. Mol Biol Cell 10: 1653– 1663. Johnson C, Primorac D, McKinstry M, McNeil J, Rowe D, Lawrence JB. 2000. Tracking COL1A1 RNA in osteogenesis imperfecta. splice-defective transcripts initiate transport from the gene but are retained within the SC35 domain. J Cell Biol 150: 417– 432. Jolly C, Vourc’h C, Robert-Nicoud M, Morimoto RI. 1999. Intron-independent association of splicing factors with active genes. J Cell Biol 145: 1133– 1143. Kim SH, McQueen PG, Lichtman MK, Shevach EM, Parada LA, Misteli T. 2004. Spatial genome organization during T-cell differentiation. Cytogenet Genome Res 105: 292 –301. Kimura H, Tao Y, Roeder RG, Cook PR. 1999. Quantitation of RNA polymerase II and its transcription factors in an HeLa cell: Little soluble holoenzyme but significant amounts of polymerases attached to the nuclear substructure. Mol Cell Biol 19: 5383– 5392. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158 –162. Kumaran RI, Spector DL. 2008. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol 180: 51–65. Kuroda M, Tanabe H, Yoshida K, Oikawa K, Saito A, Kiyuna T, Mizusawa H, Mukai K. 2004. Alteration of

Gene Positioning chromosome positioning during adipocyte differentiation. J Cell Sci 117: 5897– 5903. Lanzuolo C, Roure V, Dekker J, Bantignies F, Orlando V. 2007. Polycomb response elements mediate the formation of chromosome higher-order structures in the bithorax complex. Nat Cell Biol 9: 1167– 1174. Lawrence JB, Clemson CM. 2008. Gene associations: True romance or chance meeting in a nuclear neighborhood? J Cell Biol 182: 1035– 1038. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –293. Lin C, Yang L, Tanasa B, Hutt K, Ju BG, Ohgi K, Zhang J, Rose DW, Fu XD, Glass CK, et al. 2009. Nuclear receptorinduced chromosomal proximity and DNA breaks underlie specific translocations in cancer. Cell 139: 1069– 1083. Ling JQ, Li T, Hu JF, Vu TH, Chen HL, Qiu XW, Cherry AM, Hoffman AR. 2006. CTCF mediates interchromosomal colocalization between Igf2/H19 and Wsb1/Nf1. Science 312: 269– 272. Liu Z, Garrard WT. 2005. Long-range interactions between three transcriptional enhancers, active Vk gene promoters, and a 30 boundary sequence spanning 46 kilobases. Mol Cell Biol 25: 3220–3231. Liu JL, Murphy C, Buszczak M, Clatterbuck S, Goodman R, Gall JG. 2006. The Drosophila melanogaster Cajal body. J Cell Biol 172: 875 –884. Lomvardas S, Barnea G, Pisapia DJ, Mendelsohn M, Kirkland J, Axel R. 2006. Interchromosomal interactions and olfactory receptor choice. Cell 126: 403– 413. Luco RF, Maestro MA, Sadoni N, Zink D, Ferrer J. 2008. Targeted deficiency of the transcriptional activator Hnf1a alters subnuclear positioning of its genomic targets. PLoS Genet 4: e1000079. Malhas A, Lee CF, Sanders R, Saunders NJ, Vaux DJ. 2007. Defects in lamin B1 expression or processing affect interphase chromosome position and gene expression. J Cell Biol 176: 593–603. Mani RS, Tomlins SA, Callahan K, Ghosh A, Nyati MK, Varambally S, Palanisamy N, Chinnaiyan AM. 2009. Induced chromosomal proximity and gene fusions in prostate cancer. Science 326: 1230. Masny PS, Bengtsson U, Chung SA, Martin JH, van Engelen B, van der Maarel SM, Winokur ST. 2004. Localization of 4q35.2 to the nuclear periphery: Is FSHD a nuclear envelope disease? Hum Mol Genet 13: 1857–1871. Matarazzo MR, Boyle S, D’Esposito M, Bickmore WA. 2007. Chromosome territory reorganization in a human disease with altered DNA methylation. Proc Natl Acad Sci U S A 104: 16546– 16551. Mathas S, Kreher S, Meaburn KJ, Johrens K, Lamprecht B, Assaf C, Sterry W, Kadin ME, Daibata M, Joos S, et al. 2009. Gene deregulation and spatial genome reorganization near breakpoints prior to formation of translocations in anaplastic large cell lymphoma. Proc Natl Acad Sci U S A 106: 5831–5836.

Meaburn KJ, Misteli T. 2008. Locus-specific and activity-independent gene repositioning during early tumorigenesis. J Cell Biol 180: 39– 50. Meaburn KJ, Cabuy E, Bonne G, Levy N, Morris GE, Novelli G, Kill IR, Bridger JM. 2007. Primary laminopathy fibroblasts display altered genome organization and apoptosis. Aging Cell 6: 139– 153. Meaburn KJ, Gudla PR, Khan S, Lockett SJ, Misteli T. 2009. Disease-specific gene repositioning in breast cancer. J Cell Biol 187: 801 –812. Meaburn KJ, Levy N, Toniolo D, Bridger JM. 2005. Chromosome positioning is largely unaffected in lymphoblastoid cell lines containing emerin or A-type lamin mutations. Biochem Soc Trans 33: 1438–1440. Misteli T. 2007. Beyond the sequence: Cellular organization of genome function. Cell 128: 787– 800. Moen PTJr, Johnson CV, Byron M, Shopland LS, de la Serna IL, Imbalzano AN, Lawrence JB. 2004. Repositioning of muscle-specific genes relative to the periphery of SC-35 domains during skeletal myogenesis. Mol Biol Cell 15: 197 –206. Morey C, Kress C, Bickmore WA. 2009. Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to up-regulate gene expression. Genome Res 19: 1184– 1194. Morey C, Da Silva NR, Perry P, Bickmore WA. 2007. Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134: 909 –919. Murrell A, Heeson S, Reik W. 2004. Interaction between differentially methylated regions partitions the imprinted genes Igf2 and H19 into parent-specific chromatin loops. Nat Genet 36: 889 –893. Neves H, Ramos C, da Silva MG, Parreira A, Parreira L. 1999. The nuclear topography of ABL, BCR, PML, and RARa genes: Evidence for gene proximity in specific phases of the cell cycle and stages of hematopoietic differentiation. Blood 93: 1197–1207. Nikiforova MN, Stringer JR, Blough R, Medvedovic M, Fagin JA, Nikiforov YE. 2000. Proximity of chromosomal loci that participate in radiation-induced rearrangements in human cells. Science 290: 138– 141. Noordermeer D, Branco MR, Splinter E, Klous P, van Ijcken W, Swagemakers S, Koutsourakis M, van der Spek P, Pombo A, de Laat W. 2008. Transcription and chromatin organization of a housekeeping gene cluster containing an integrated b-globin locus control region. PLoS Genet 4: e1000016. O’Sullivan JM, Tan-Wong SM, Morillon A, Lee B, Coles J, Mellor J, Proudfoot NJ. 2004. Gene loops juxtapose promoters and terminators in yeast. Nat Genet 36: 1014–1018. Orlando V. 2003. Polycomb, epigenomes, and control of cell identity. Cell 112: 599–606. Osborne CS, Chakalova L, Brown KE, Carter D, Horton A, Debrand E, Goyenechea B, Mitchell JA, Lopes S, Reik W, et al. 2004. Active genes dynamically colocalize to shared sites of ongoing transcription. Nat Genet 36: 1065–1071. Osborne CS, Chakalova L, Mitchell JA, Horton A, Wood AL, Bolland DJ, Corcoran AE, Fraser P. 2007. Myc

C. Ferrai et al. dynamically and preferentially relocates to a transcription factory occupied by Igh. PLoS Biol 5: e192. Palstra RJ, Tolhuis B, Splinter E, Nijmeijer R, Grosveld F, de Laat W. 2003. The b-globin nuclear compartment in development and erythroid differentiation. Nat Genet 35: 190 –194. Parada L, Misteli T. 2002. Chromosome positioning in the interphase nucleus. Trends Cell Biol 12: 425– 432. Pombo A, Jackson DA, Hollinshead M, Wang Z, Roeder RG, Cook PR. 1999. Regional specialization in human nuclei: Visualization of discrete sites of transcription by RNA polymerase III. Embo J 18: 2241– 2253. Ragoczy T, Bender MA, Telling A, Byron R, Groudine M. 2006. The locus control region is required for association of the murine b-globin locus with engaged transcription factories during erythroid maturation. Genes Dev 20: 1447– 1457. Reddy KL, Zullo JM, Bertolino E, Singh H. 2008. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452: 243–247. Rodley CD, Bertels F, Jones B, O’Sullivan JM. 2009. Global identification of yeast chromosome interactions using Genome conformation capture. Fungal Genet Biol 46: 879–886. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287 –291. Rozycka M, Sawicki W, Traczyk Z, Bem W, Strojny P. 1988. Analysis of chromatin pattern in blood lymphocytes of healthy donors and in lymphoid cells of patients with chronic lymphocytic leukaemia. J Clin Pathol 41: 504–509. Sabbattini P, Lundgren M, Georgiou A, Chow C, Warnes G, Dillon N. 2001. Binding of Ikaros to the lambda5 promoter silences transcription through a mechanism that does not require heterochromatin formation. Embo J 20: 2812–2822. Sadoni N, Langer S, Fauth C, Bernardi G, Cremer T, Turner BM, Zink D. 1999. Nuclear organization of mammalian genomes. Polar chromosome territories build up functionally distinct higher order compartments. J Cell Biol 146: 1211–1226. Sadoni N, Targosz BS, Englmann A, Fesser S, Koch J, Schindelhauer D, Zink D. 2008. Transcription-dependent spatial arrangements of CFTR and conserved adjacent loci are not conserved in human and murine nuclei. Chromosoma 117: 381 –397. Saurin AJ, Shiels C, Williamson J, Satijn DP, Otte AP, Sheer D, Freemont PS. 1998. The human polycomb group complex associates with pericentromeric heterochromatin to form a novel nuclear domain. J Cell Biol 142: 887– 898. Scaffidi P, Gordon L, Misteli T. 2005. The cell nucleus and aging: Tantalizing clues and hopeful promises. PLoS Biol 3: e395. Schoenfelder S, Sexton T, Chakalova L, Cope NF, Horton A, Andrews S, Kurukuti S, Mitchell JA, Umlauf D, Dimitrova DS, et al. 2010. Preferential associations between co-regulated genes reveal a transcriptional interactome in erythroid cells. Nat Genet 42: 53–61.

Schuettengruber B, Chourrout D, Vervoort M, Leblanc B, Cavalli G. 2007. Genome regulation by polycomb and trithorax proteins. Cell 128: 735– 745. Schul W, van Der Kraan I, Matera AG, van Driel R, de Jong L. 1999. Nuclear domains enriched in RNA 30 -processing factors associate with coiled bodies and histone genes in a cell cycle-dependent manner. Mol Biol Cell 10: 3815–3824. Sexton T, Bantignies F, Cavalli G. 2009. Genomic interactions: Chromatin loops and gene meeting points in transcriptional regulation. Semin Cell Dev Biol. 20: 849–55. Shiels C, Islam SA, Vatcheva R, Sasieni P, Sternberg MJ, Freemont PS, Sheer D. 2001. PML bodies associate specifically with the MHC gene cluster in interphase nuclei. J Cell Sci 114: 3705– 3716. Shopland LS, Johnson CV, Lawrence JB. 2002. Evidence that all SC-35 domains contain mRNAs and that transcripts can be structurally constrained within these domains. J Struct Biol 140: 131– 139. Shopland LS, Byron M, Stein JL, Lian JB, Stein GS, Lawrence JB. 2001. Replication-dependent histone gene expression is related to Cajal body (CB) association but does not require sustained CB contact. Mol Biol Cell 12: 565– 576. Shopland LS, Johnson CV, Byron M, McNeil J, Lawrence JB. 2003. Clustering of multiple specific genes and gene-rich R-bands around SC-35 domains: Evidence for local euchromatic neighborhoods. J Cell Biol 162: 981– 990. Simonis M, Klous P, Splinter E, Moshkin Y, Willemsen R, de Wit E, van Steensel B, de Laat W. 2006. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat Genet 38: 1348–1354. Skok JA, Brown KE, Azuara V, Caparros ML, Baxter J, Takacs K, Dillon N, Gray D, Perry RP, Merkenschlager M, et al. 2001. Nonequivalent nuclear location of immunoglobulin alleles in B lymphocytes. Nat Immunol 2: 848 –854. Smith KP, Lawrence JB. 2000. Interactions of U2 gene loci and their nuclear transcripts with Cajal (coiled) bodies: Evidence for PreU2 within Cajal bodies. Mol Biol Cell 11: 2987–2998. Smith KP, Byron M, Johnson C, Xing Y, Lawrence JB. 2007. Defining early steps in mRNA transport: Mutant mRNA in myotonic dystrophy type I is blocked at entry into SC-35 domains. J Cell Biol 178: 951 –964. Smith KP, Moen PT, Wydner KL, Coleman JR, Lawrence JB. 1999. Processing of endogenous pre-mRNAs in association with SC-35 domains is gene specific. J Cell Biol 144: 617–629. Solovei I, Kreysing M, Lanctot C, Kosem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137: 356 –368. Spector DL. 1993. Macromolecular domains within the cell nucleus. Annu Rev Cell Biol 9: 265– 315. Spilianakis CG, Flavell RA. 2004. Long-range intrachromosomal interactions in the T helper type 2 cytokine locus. Nat Immunol 5: 1017–1027. Spilianakis CG, Lalioti MD, Town T, Lee GR, Flavell RA. 2005. Interchromosomal associations between alternatively expressed loci. Nature 435: 637–645.

Gene Positioning Splinter E, Heath H, Kooren J, Palstra RJ, Klous P, Grosveld F, Galjart N, de Laat W. 2006. CTCF mediates long-range chromatin looping and local histone modification in the b-globin locus. Genes Dev 20: 2349– 2354. Sproul D, Gilbert N, Bickmore WA. 2005. The role of chromatin structure in regulating the expression of clustered genes. Nat Rev Genet 6: 775– 781. Stanek D, Neugebauer KM. 2006. The Cajal body: A meeting place for spliceosomal snRNPs in the nuclear maze. Chromosoma 115: 343 –354. Sun Y, Durrin LK, Krontiris TG. 2003. Specific interaction of PML bodies with the TP53 locus in Jurkat interphase nuclei. Genomics 82: 250–252. Szutorisz H, Dillon N, Tora L. 2005. The role of enhancers as centres for general transcription factor recruitment. Trends Biochem Sci 30: 593 –599. Takizawa T, Meaburn KJ, Misteli T. 2008b. The meaning of gene positioning. Cell 135: 9 –13. Takizawa T, Gudla PR, Guo L, Lockett S, Misteli T. 2008a. Allele-specific nuclear positioning of the monoallelically expressed astrocyte marker GFAP. Genes Dev 22: 489–498. Tan-Wong SM, Wijayatilake HD, Proudfoot NJ. 2009. Gene loops function to maintain transcriptional memory through interaction with the nuclear pore complex. Genes Dev 23: 2610–2624. Tiwari VK, Cope L, McGarvey KM, Ohm JE, Baylin SB. 2008a. A novel 6C assay uncovers Polycomb-mediated higher order chromatin conformations. Genome Res 18: 1171– 1179. Tiwari VK, McGarvey KM, Licchesi JD, Ohm JE, Herman JG, Schubeler D, Baylin SB. 2008b. PcG proteins, DNA methylation, and gene repression by chromatin looping. PLoS Biol 6: 2911–2927. Tolhuis B, Palstra RJ, Splinter E, Grosveld F, de Laat W. 2002. Looping and interaction between hypersensitive sites in the active b-globin locus. Mol Cell 10: 1453–1465. Towbin BD, Meister P, Gasser SM. 2009. The nuclear envelope–a scaffold for silencing? Curr Opin Genet Dev 19: 180–186. Tsukamoto T, Hashiguchi N, Janicki SM, Tumbar T, Belmont AS, Spector DL. 2000. Visualization of gene activity in living cells. Nat Cell Biol 2: 871–878. Vakoc CR, Letting DL, Gheldof N, Sawado T, Bender MA, Groudine M, Weiss MJ, Dekker J, Blobel GA. 2005. Proximity among distant regulatory elements at the b-globin locus requires GATA-1 and FOG-1. Mol Cell 17: 453 –462. Verschure PJ, van der Kraan I, Manders EM, Hoogstraten D, Houtsmuller AB, van Driel R. 2003. Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules. EMBO Rep 4: 861– 866. Verschure PJ, van Der Kraan I, Manders EM, van Driel R. 1999. Spatial relationship between transcription sites and chromosome territories. J Cell Biol 147: 13–24. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J,

et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–1576. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. 2003. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol 160: 685–697. Wang J, Shiels C, Sasieni P, Wu PJ, Islam SA, Freemont PS, Sheer D. 2004. Promyelocytic leukemia nuclear bodies associate with transcriptionally active genomic regions. J Cell Biol 164: 515– 526. Wansink DG, Schul W, van der Kraan I, van Steensel B, van Driel R, de Jong L. 1993. Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol 122: 283–293. Webster M, Witkin KL, Cohen-Fix O. 2009. Sizing up the nucleus: Nuclear shape, size and nuclear-envelope assembly. J Cell Sci 122: 1477– 1486. Wiech T, Timme S, Riede F, Stein S, Schuricke M, Cremer C, Werner M, Hausmann M, Walch A. 2005. Human archival tissues provide a valuable source for the analysis of spatial genome organization. Histochem Cell Biol 123: 229 –238. Williams RR, Azuara V, Perry P, Sauer S, Dvorkina M, Jorgensen H, Roix J, McQueen P, Misteli T, Merkenschlager M, et al. 2006. Neural induction promotes large-scale chromatin reorganisation of the Mash1 locus. J Cell Sci 119: 132–140. Xie SQ, Martin S, Guillot PV, Bentley DL, Pombo A. 2006. Splicing speckles are not reservoirs of RNA polymerase II, but contain an inactive form, phosphorylated on serine2 residues of the C-terminal domain. Mol Biol Cell 17: 1723–1733. Xing Y, Johnson CV, Moen PTJr, McNeil JA, Lawrence J. 1995. Nonrandom gene organization: Structural arrangements of specific pre-mRNA transcription and splicing with SC-35 domains. J Cell Biol 131: 1635– 1647. Xu M, Cook PR. 2008. Similar active genes cluster in specialized transcription factories. J Cell Biol 181: 615 –623. Xu N, Tsai CL, Lee JT. 2006. Transient homologous chromosome pairing marks the onset of X inactivation. Science 311: 1149– 1152. Zhao Z, Tavoosidana G, Sjolinder M, Gondor A, Mariano P, Wang S, Kanduri C, Lezcano M, Sandhu KS, Singh U, et al. 2006. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat Genet 38: 1341–1347. Zink D, Amaral MD, Englmann A, Lang S, Clarke LA, Rudolph C, Alt F, Luther K, Braz C, Sadoni N, et al. 2004. Transcription-dependent spatial arrangements of CFTR and adjacent genes in human cell nuclei. J Cell Biol 166: 815 –825.

Chromatin Higher-order Structure and Dynamics Christopher L. Woodcock1,2 and Rajarshi P. Ghosh2 1

Biology Department, University of Massachusetts, Amherst, Massachusetts 01003

2

Program in Molecular and Cellular Biology, University of Massachusetts, Amherst, Massachusetts 01003

Correspondence: [email protected]

The primary role of the nucleus as an information storage, retrieval, and replication site requires the physical organization and compaction of meters of DNA. Although it has been clear for many years that nucleosomes constitute the first level of chromatin compaction, this contributes a relatively small fraction of the condensation needed to fit the typical genome into an interphase nucleus or set of metaphase chromosomes, indicating that there are additional “higher order” levels of chromatin condensation. Identifying these levels, their interrelationships, and the principles that govern their occurrence has been a challenging and much discussed problem. In this article, we focus on recent experimental advances and the emerging evidence indicating that structural plasticity and chromatin dynamics play dominant roles in genome organization. We also discuss novel approaches likely to yield important insights in the near future, and suggest research areas that merit further study.

s the cell’s primary information storage, retrieval, and duplication organelle, the nucleus, by analogy with human-created information repositories (libraries and computer discs), might be expected to be highly structured. However, chromatin within the living nucleus appears to be organized in a quite unstructured manner, and the rules that define or encode the principles of chromatin organization have been difficult to decipher. These frustrations are reflected in some reviews of chromatin organization with titles such as “Chromatin higher-order structure: chasing a mirage?” (van Holde and Zlatanova 1995), “Higher-order structures of chromatin: the elusive 30 nm fiber” (Tremethick 2007), and “Chromatin fiber structure: where is the problem now?” (van Holde and Zlatanova 2007). In this article, we discuss what is known about chromatin structure, and

A

consider the prospects of improving our understanding in the near future. In recent years, work on chromatin organization has increasingly focused on the many dynamic aspects of chromatin, which contribute to its structural and functional plasticity, and it is becoming clear that dynamics plays a crucially important functional role, perhaps contributing to nuclear selforganization (Misteli 2001). CHROMATIN HIGHER-ORDER STRUCTURE

In the context of chromatin, “higher-order structure” may be defined as any assemblage of nucleosomes that assumes a reproducible conformation in 3D space. The most obvious chromatin higher-order structure is the mitotic/ meiotic chromosome in which the DNA is compacted some 10,000- to 20,000-fold. Metaphase

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000596 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000596

C.L. Woodcock and R.P. Ghosh

chromosomes have characteristic shapes, banding patterns, and locations of specific genes. Although chromosomes have a consistent structure at the light microscope level in terms of longitudinal positioning of bands and genes, this consistency is evidently modulated by an intrinsic variability in longitudinal position of up to 0.3 mm (Strukov and Belmont 2009). However, these authors reported that there was no consistency in the axial positioning of loci, suggesting that chromosome architecture involves considerable plasticity at some level or levels of folding. Understanding how chains of nucleosomes are folded in the creation of mitotic chromosomes and how they are arranged in interphase continues to be an exciting yet technically challenging endeavor. The concept of primary, secondary, tertiary, and quaternary structures used for proteins can also be usefully applied to chromatin structural hierarchies (Woodcock and Dimitrov 2001), with the beads-on-a-string organization of nucleosomes and linker DNA constituting the primary structure, and arrangements resulting from interactions between nucleosomes giving rise to secondary structures. Thus, the chromatin equivalent of protein secondary structure might involve interactions analogous to those leading to a-helices and b-sheets. Unlike proteins that consist of sequences of 20 amino acids, chromatin consists of a repeating chain of more-or-less identical nucleosomes, and thus might be predicted to form highly “ordered” secondary structures. Note that the term “ordered” can be used in the sense that a crystal is highly ordered and also used to describe hierarchical levels where the concept is more akin to its meaning in “orders of magnitude.” Although in one sense, the basic nucleosome-linker DNA unit comprising the primary structure of chromatin is simple, there are a number of potentially variable parameters that contribute to complexity. There is structural uncertainty even at the level of the nucleosome core particle (NCP). Although the structure of the NCP is well established at the atomic level (Luger et al. 1997), portions of the four core histones are not seen in crystals because of their evident mobility. For example, the unstructured

terminal regions that are extremely important in modulating chromatin structure are not seen in X-ray data. The length of linker DNA between NCPs varies not only between species, but also between tissues of the same organism, and within a single nucleus (van Holde 1989). Mean linker lengths (defined as the nucleosome repeat length minus the 145 bp of DNA protected in NCPs) range from a low value of 20 bp in budding yeast to 75 bp in echinoderm sperm. For a typical vertebrate nucleus the mean linker length is 35 bp. The rich literature concerning the locations of nucleosome positions on specific DNA sequences, reveals instances, especially in the upstream control regions of genes, in which nucleosomes are preferentially located or excluded, and this positioning is often important in regulating transcription (Simpson 1991; Jiang and Pugh 2009). For some DNA sequences, the preference for nucleosomes to be present at a particular location is retained when the nucleosomes are reconstituted on the DNA in vitro. In addition to linker DNA length, nucleosomes vary in their complement of core histone variants and histone postsynthetic modifications. All these variations are likely to impact chromatin secondary structures. The obvious experimental approaches to determine chromatin secondary and higherorder structures using light and electron microscopy yield disappointingly little information, largely because nucleosomes and linker DNA cannot be adequately resolved in the compact chromatin that occurs in the nucleus. A recent study using cryo-electron microscopy of thin sections of chromosomes vitrified in vivo, and therefore expected to faithfully reflect the native structure, provides an instructive example. Despite careful image processing of the micrographs, there was no evidence for any higher-order structure, leading the authors to suggest that the chromatin forms a “molten” mass similar to that assumed by certain polymers, and that in this form, the size, shape and trajectories of any higher-order structures would not be resolved (Eltsov et al. 2008). This may well be the case, but nevertheless, the arrangements of arrays of nucleosomes in

Chromatin Higher-order Structure and Dynamics

3D space are to some extent constrained and nonrandom, and important information would be obtained if it were possible to trace the paths of chains of nucleosomes through the nucleus. The generally amorphous appearance of chromatin in thin sections contrasts sharply with the appearance of cytoskeletal elements such as microtubules, which can be resolved in cells even when in compact bundles. This striking difference supports the concept that tightly packed chains of nucleosomes readily interdigitate with each other, leading to the rather featureless appearance of chromatin in EM images (Woodcock and Horowitz 1995; Woodcock 2006). A new technique for preparing frozen hydrated material that involves thinning of samples by milling with a focused ion beam (Marko et al. 2007, 2008) that does not involve the compression and possible local heating of cryosectioning is currently under development in several laboratories, and it will be most interesting to see whether this method provides improved structural information when applied to nuclear structures. The 30-nm Fiber

In the absence of clear conclusions from the examination of nuclei and chromosomes in situ, many investigators have focused on isolated chromatin. Through the use of nucleases that cut the linker DNA between nucleosomes, it is possible to isolate polynucleosomes and study their properties under defined conditions in solution. Early work clearly established that the compaction state of polynucleosomes is highly dependent on the ionic milieu. Naked DNA, being highly charged, is self-repulsive in low ionic strength buffers—the self-repulsion can be reduced or eliminated in the presence of cations, with divalent and polyvalent cations being especially effective. Chromatin behaves similarly to DNA in this respect because the DNA negative charge is not fully neutralized by histones. The response of chains of nucleosomes to changes in the ionic environment can be satisfactorily modeled (Clark and Kimura 1990; Arya and Schlick 2006), suggesting that our basic understanding of the response of

chromatin to changes in the ionic environment is robust. Electron microscopy of isolated polynucleosomes clearly reveals an open beads-on-a-string conformation in low salt and a progressive compaction that occurs as the ionic strength is raised. There is general agreement that an early stage in compaction is the formation of a fiber 30 nm in diameter, and that, at least for polynucleosomes in vitro, this constitutes a bona fide chromatin secondary structure. However, the arrangement of nucleosomes and linker DNA within isolated 30 nm fibers has been difficult to study and remains controversial. Careful examination of isolated 30 nm fibers has failed to reveal any consistent clear-cut arrangement of nucleosomes. The realization that native nucleosomes are not identical, but, as described earlier, vary in several parameters, suggested that these could be the source of the equivocal findings regarding 30-nm fiber organization. To eliminate these potential problems, investigators turned to fully defined artificial polynucleosomes reconstituted onto DNA containing regularly spaced nucleosome localization sequences (Simpson et al. 1985). A major achievement using this approach was the successful crystallization and structure determination by X-ray diffraction of a tetranucleosome (Schlach et al. 2005). This showed unequivocally that, at least for this particular construct, exposed to crystallization-promoting conditions that included 90-mM Mgþþ , the complex assumed a zigzag arrangement, with contacts between nucleosomes 1 and 3 and between 2 and 4, with the linker extending between them (Fig. 1). The work clearly established the predominant internucleosome interactions that result in a specific secondary structure for this particular chromatin construct in this ionic environment. It is not yet clear how far the zigzag structure can be extrapolated to long chromatin with heterogeneity in histone content and linker length. Nevertheless, it does provide an important context in which to view earlier results. For example, the interpretation of the first EM studies of isolated compact 30-nm fibers in terms of a simple solenoidal organization in

C.L. Woodcock and R.P. Ghosh

N2

N1'

N1

N2'

Figure 1. The DNA path of a tetranucleosome as

determined by X-ray diffraction. The structure consists of two stacks of nucleosomes, with linker DNA passing back and forth between them. Thus, the primary interactions occur between alternate rather than adjacent nucleosomes along the DNA strand, creating a zigzag architecture. From Schlach et al. 2005.

which linker DNA followed the helical trajectory established in the nucleosome is probably incorrect, whereas subsequent suggestions involving some species of zigzag are probably closer to the mark. THE ROLE OF HISTONE H1

In addition to the four “core” histones present as an octameric unit in the nucleosomes, a fifth histone, H1 or linker histone, is present in most nuclei. As its name implies linker histone is associated with linker DNA, and provides partial nuclease protection for 20 bp of linker DNA. H1 contains a conserved globular region and extended amino- and carboxy-termini, the latter being rich in lysines and able to interact strongly with DNA. Although the exact location of H1 in chromatin remains controversial, it is clear that it plays a crucial role in promoting chromatin higher-order structure. H1-containing chromatin shows a distinct structural motif in which the entering and exiting linker DNA segments are brought together, perhaps promoting an overall zigzag arrangement (Bednar et al. 1998). Arrays of nucleosomes depleted in H1 do not readily form 30-nm fibers in vitro, and it is likely that loss of the charge neutralization effected by H1 is at least partly responsible. The crystals of tetranucleosomes discussed above were created without H1—and to date, no publications of the X-ray structure of

H1-containing mono- or oligo-nucleosomes have appeared. Modest depletion of H1 from cells (achieved for the mouse by knocking out several of the six somatic H1 variants) results in reductions in nucleosome repeat length in some tissues whereas severe depletion is fatal for both mice and Drosophila (Fan et al. 2003; Lu et al. 2009). H1 depletion also inhibits the proper folding of chromosomes at mitosis (Maresca and Heald 2006). Interestingly, there is a strong linear relationship between H1 content and nucleosome repeat length in vivo (Woodcock et al. 2006), a feature that will act to maintain the electrostatic balance between DNA and histones. CHROMATIN FIBERS IN NUCLEI

Thin sections of typical nuclei reveal a coarse differentiation between strongly staining heterochromatin (see the following for more on this term) adjacent to the nuclear periphery and nucleoli, and more weakly staining euchromatin dispersed throughout. Little internal structural detail can be resolved in either component. There are, however, a few special cases that are more informative. Echinoderm sperm nuclei contain uniformly distributed and highly compact chromatin, but on controlled reduction in ionic strength, the chromatin swells, revealing a mass of 30-nm fibers. Although the fibers appear to be quite uniform in diameter, tomographic 3D reconstructions reveal an internal structure that consists of an irregular arrangement of nucleosomes and linker DNA but with a distinctive underlying zigzag motif (Horowitz et al. 1994). Sections of the transcriptionally inert chicken erythrocyte nucleus also reveal a uniform compaction of chromatin, which resolves into irregularly structured 30-nm fibers when allowed to swell slightly by, for example, lowering the ionic strength. In contrast, 30-nm fibers are rarely seen in more typical nuclei from cycling cells, even after controlled swelling (Woodcock and Horowitz, 1995). These special cases in which the 30-nm organization is revealed on swelling have several common features, including uniform core histone modifications, a greater

Chromatin Higher-order Structure and Dynamics

abundance of linker histone (H1) relative to core histones, the presence of H1 variants with more positive charges and unusually long linker DNA. It is not clear why these properties lead to the appearance of distinct 30-nm fibers in nucleo, but an attractive possibility is that these conditions favor intrafiber over interfiber interactions. NUCLEOSOME-NUCLEOSOME INTERACTIONS IN ISOLATED CHROMATIN FIBERS

One approach to understanding chromatin fiber structure is to identify nucleosome-nucleosome interactions within the compact fiber. As noted, this cannot be achieved by direct EM observation of compact fibers, but ways to examine this have been devised. One study used the premise that the face-to-face nucleosome contacts that occur in crystals of mononucleosomes (Luger et al. 1997) would likely be seen in compact chromatin. The contacts involved the positively charged N-terminus of histone H4 and the “acidic patch” that includes a portion of histone H2A. Dorigo et al. (2004) created H4 and H2A constructs with a cysteine in the putative contact region and reconstituted them into nucleosomal arrays. The arrays were then placed in conditions that promoted compaction and formation of S-S crosslinks between close cysteines. If the basic architecture of the array were solenoidal, crosslinking would result in a simple stack-of-coins conformation. In contrast, a zigzag arrangement would result in the creation of a ladder-like structure with the linker DNA segments forming the rungs. When cross-linking was initiated, the arrays clearly adopted the ladder-like structure, strongly supporting a zigzag architecture, albeit under somewhat artificial conditions. An approach that does not restrict internucleosome interactions to a specific histone-histone contact involves exposing nucleosomal arrays or whole cells to controlled formaldehyde crosslinking such that a few of the nucleosome-nucleosome contacts become covalently linked. Subsequently, the arrays are allowed to disperse in low salt and imaged by electron microscopy. With

his technique, termed EMANIC (EM-assisted nucleosome interaction capture) (Grigoryev et al. 2009), decondensed arrays appear in the open beads-on-a-string conformation (Fig. 2A), whereas crosslinked sites are seen as touching nucleosomes. Analysis of the crosslinking pattern of defined nucleosome arrays with H1 present once again reveals a predominantly zigzag pattern (Fig. 2 D, E). With H1 and 1 mM MgCl2 together, some crosslinking is also seen between adjacent nucleosomes, suggesting these conditions favor a mixed, heteromorphic fiber architecture (Fig. 2F, G) in which the zigzag dominates, but is interrupted by the type of nucleosome-nucleosome interaction expected for a solenoidal architecture. Interestingly, computer modeling predicts heteromorphic fibers, and also shows that this arrangement allows greater compaction as it reduces the crowding of linker DNAs in the fiber interior (Grigoryev et al. 2009), and is more consistent with the higher packing ratios seen in compact chromatin (Daban 2000; 2003). Although there is substantial support for a basic propensity of chromatin to form zigzag higher-order structures, results leading to alternate architectures have been presented. EM examination of long reconstituted nucleosomal arrays containing linker histone reveals remarkably uniform structures interpreted as consisting of solenoids in which adjacent gyres interdigitate (Robinson et al. 2006). This arrangement also increases the nucleosome packing density, and may thus better match the calculated in vivo packing. Additional studies of long nucleosomal arrays have led to the conclusion that the architecture is strongly influenced by linker length (Routh et al. 2008), increasing the potential complexity of chromatin structure in vivo, where varied linker DNA lengths are the norm. MECHANICAL PROPERTIES OF ISOLATED CHROMATIN FIBERS

The development of sophisticated instruments using “optical tweezers” has made it possible to take an isolated chromatin fiber, anchor one end, and pull on the other end while

C.L. Woodcock and R.P. Ghosh

A

Crosslinking

Unfolding and EM

i+/–1

i+/–2

B

C

D

D’ 2 8

E

F

G

H

I

I’ 2

2

Figure 2. EMANIC analysis of internucleosomal interactions. (A) Scheme of the EMANIC procedure. The two

models for the structure of the chromatin 30-nm fiber, namely solenoid (Upper) and zigzag (Lower), lead to dominant i + 1 and i + 2 internucleosome interactions, respectively. (B – D) EM of nucleosome reconstitutes crosslinked in low salt without linker histone show few crosslinks. (E-I) With H1 present, +2 interactions predominate. D0 and I0 diagram the nucleosome arrays corresponding to the adjacent EM images. From Grigoryev et al. 2009.

measuring the force/length relationship. The applied force is expected first to break the weak internucleosomal bonds holding the fiber together, and ultimately, the much stronger intranucleosomal bonds. For a fiber with a zigzag arrangement, the breaking of individual nucleosome-nucleosome interactions is predicted to produce jumps in length with a consistent increment which would not be observed with solenoid type structures. Analysis of force/length curves for isolated chromatin fibers from chicken erythrocyte nuclei led to the conclusion that a zigzag arrangement predominated (Cui and Bustamante 2000). However, force/length data from a more recent study using reconstituted nucleosomal arrays (Kruithof et al. 2009) were interpreted as supporting a solenoidal architecture. The differences in starting material and ionic conditions used in these studies makes it difficult to evaluate the findings. In the future, studies encompassing a broader range of conditions and

chromatin substrates should provide more definitive information from this potentially valuable technique. INSIGHTS FROM COMPUTER MODELING

As noted earlier, computer modeling was consistent with the empirical finding of a heteromorphic fiber architecture. Current “mesoscale” modeling programs are supplied with sophisticated structural and chemical models of the nucleosome, H1 (if desired) and linker DNA, as well as the selected ionic environment (Arya and Schlick 2006). The changes in structure are then followed over a large number of small increments of time, and the final structures analyzed. As expected, the stochastic nature of the interactions involved lead to different final structures with each run of the program. However, consistent motifs emerge: the final structures are fibers, rather than spherical aggregates of nucleosomes, and, although zigzag

Chromatin Higher-order Structure and Dynamics

arrangements of nucleosomes and linker DNA predominate, they may be interspersed with the bent linker motif characteristic of simple helical arrangements of nucleosomes. Modeling results are dependent on the accuracy of the input data, and computational limitations still preclude modeling at the atomic level. Nevertheless, this approach has the advantage of being able to sample a wide range of conditions, informing the selection of conditions for empirical examination with “real” chromatin. As computational speed continues to increase, modeling is likely to lead to more penetrating insights into chromatin architecture. LOCAL EVENTS THAT MAY MODULATE CHROMATIN FIBER ARCHITECTURE

Much of recent work on chromatin secondary structure has featured fully defined nucleosomal arrays in which recombinant histones are assembled in vitro onto DNA containing strong nucleosome positioning sequences. Yet, as noted, even with highly uniform arrays, results have led to differing conclusions regarding the underlying architecture. In the nucleus, local effects of linker DNA length variability, core histone variants, and postsynthetic histone modifications are likely to contribute even more irregularity to secondary structures. Additional complexity is introduced by the presence of nonhistone proteins known collectively as Chromatin Architectural Proteins (CAPs) that influence chromatin conformation (Luger and Hansen 2005). In addition to the “major” core histones, synthesized primarily in S phase and deposited at replication forks, there are numerous histone variants encoded by separate genes, which are often synthesized constitutively at low levels and incorporated differently. Some of these have been shown to play a critical role in establishing the local properties of chromatin in which they are embedded (Santenard and Torres-Padilla 2009). An important concept in this regard is the distinction between the major variant that is synthesized during S phase and deposited as newly replicated chromatin, and minor variants. It has been proposed that the

incorporation of different H3 variants serves as a “bar code” (Hake and Allis 2006) contributing to the establishment of the local functional potential. Of the several variants of H2A, H2A.Z has been particularly well studied and shown to modulate chromatin higher-order structure. The X-ray structure of nucleosomes containing H2A.Z (Suto et al. 2000) reveals an enlargement of the ‘acidic patch’ at the nucleosome surface that appears to strengthen nucleosome-nucleosome interactions, and contribute to the highly compact chromatin surrounding centromeres (Greaves et al. 2007). Another major source of chromatin variation is the large number of postsynthetic modifications of the core histones and H1 (Kouzarides 2007). They fall into two major categories, those that alter the charge on the histone, and those that establish binding sites for other proteins. Acetylation of lysines results in a reduction of the histone positive charge and weakens the DNA-histone interaction leading to a less compact chromatin secondary structure. The balance in activity between histone acetyl transferases (HATs) and histone deacetylases (HDACs) governs the acetylation status of a given region of chromatin. In general, hyperacetylation is the hallmark of active chromatin, whereas hypoacetylation is seen in repressed chromatin. A well-studied example is the acetylation of H4 at lysine 16. The loss of a positive charge at this site is likely to weaken the internucleosome interaction between the amino-terminus of H4 and the acidic patch on H2A discussed above. Indeed, the effect of this modification is to loosen the compaction of chromatin secondary structures (ShogrenKnaak et al. 2006; Fischle et al. 2003). An example of a modification that establishes a binding site is the methylation of H3 at lysine 9. The modification creates a binding site for a domain of the HP1 protein (Jacobs and Khorasanizadeh 2002). As discussed below, HP1 binding is associated with the formation and propagation of compact heterochromatin. Work on postsynthetic histone modifications has led to the widely discussed concept that, collectively, they constitute a “histone code” that defines the local structural and functional potential

C.L. Woodcock and R.P. Ghosh

of a region of chromatin (Jenuwein and Allis 2001). Chromatin architectural proteins, as their name implies, do impact chromatin structure, mostly by increasing the compaction state (Luger and Hansen 2005). This appears to be their sole common feature, because otherwise they are highly diverse in both structure and in chromatin binding mechanism. Binding may be restricted to specific DNA sequences (Polycomb), methylated DNA (MeCP2), or specific histone modifications (HP1). Only in a few cases, such as the SIR family of DNA-binding regulatory proteins is a detailed account of their molecular interactions with chromatin and with each other emerging (Buchberger et al. 2008). Some CAPs contain more than one DNA/chromatin binding sites, allowing them to act as inter- and/or intrafiber bridges, and thus promote chromatin compaction. Chromatin remodeling complexes may be considered as a distinct class of CAPs. Several families of these large multicomponent complexes have been identified. In vitro, they have the ability to restructure, eject, or move nucleosomes in an ATP-dependent manner (Clapier and Cairns 2009). This type of activity in vivo will likely influence chromatin secondary structure, and, by altering the local positioning of nucleosomes, modulate chromatin folding. An interesting case has been reported of the combined effect of a histone variant (H2A.Z), a histone modification (H3K9Me), and a CAP (HP1), all of which act to increase chromatin compaction (Fan et al. 2004). Thus, the proposed histone code based on histone modifications may need to be expanded to include synergistic effects of histone variants. A novel function in maintaining the pluripotential “open” state of chromatin for the remodeling factor Chd1 (Gaspar-Maia et al. 2009) may presage new insights into the roles of these complexes. A scenario of chromatin secondary structure that, in our opinion, appears to fit all the current data involves chains of nucleosomes that are sufficiently close packed in vivo to interdigitate, effectively merging with each other and preventing individual fibers being seen by

microscopy. Nevertheless, the overall trajectory of a chain of nucleosomes traces out an irregular 30-nm fiber within the nucleus. Fragmentation of chromatin by limited nuclease digestion followed by elution from nuclei results in individual fibers becoming separated, and available for microscopic and biochemical characterization. Irregularity dominates the architecture of isolated native chromatin fibers, reflecting the heterogeneity in linker DNA length, and composition of histone variants, histone postsynthetic modification, and CAPs. The end result is likely a hybrid (heteromorphic) organization with a predominantly zigzag architecture interspersed with other structural motifs. Most of these sources of irregularity are dynamic, changing at different time scales. Thus, it is probably futile to expect that chromatin secondary structure can be defined with the precision expected from studies of other biological fibers such as microtubules or Tobacco Mosaic Virus. Thus, seeking a defined secondary structure for chromatin may indeed be “Chasing a Mirage” (van Holde and Zlatanova 1995). Structural Order Beyond the 30-nm Fiber— Electron Microscopy Results

The lack of definitive information on chromatin secondary structure complicates the question of whether there is a bona fide set of hierarchical structures above the 30-nm fiber. Although there is no doubt that some higher levels of organization are present—the metaphase chromosome being an obvious example—precisely what form they take and whether they are truly hierarchical is still a matter of intensive study and debate. Much of this work uses the light microscope and has been limited in resolution to 250 nm by the properties of light. The impact of a 250-nm resolution limit is well illustrated in a study that considers the possible structural arrangement of large genetic loci (400 kb) by matching the predictions of a number of models for large-scale chromatin structures with actual observations (Muller et al. 2004). Somewhat surprisingly, at the resolution attainable with a high quality confocal

Chromatin Higher-order Structure and Dynamics

microscope, all models fitted the data equally well! The higher resolution of the EM, coupled with tomography, and applied to thin sections has revealed short regions of fiber-like structures with a variety of diameters, none particularly consonant with a hierarchical series based on a 30-nm organization (Konig et al. 2007; Belmont et al. 1989, 1994). More recently, however, significant improvements in technique have been made using an in vivo immuno-gold technique that promises to lead to important insights (Kireev et al. 2008). The introduction by Belmont and colleagues of techniques for constructing cell lines containing large foreign DNA segments that are “self identifying” through an ingenious use of the lac operator/repressor system (Belmont et al. 1999) has had a major impact within the field and is widely used. A recent adaptation of the method involving the intracellular injection of antibodies specific for the introduced chromatin followed by gold decoration of the antibodies, thin sectioning and EM observation reveals clear-cut cylindrical fibers as can be seen in the stereo pair in Figure 3 (Kireev et al. 2008). The fibers ranged in diameter from 120 nm to 170 nm with different mean values in two cell types examined. Only rarely were individual 30-nm-like fibers seen, and, as the authors note, it remains to be seen whether the thick fibers constitute one level in a hierarchical folding series. Hopefully, further exploitation of the technique will lead to long-awaited progress in our understanding of chromatin organization in situ. Structural Order Beyond the 30-nm fiber— Light Microscopy Results

The ability to identify specific segments of chromatin using light microscopy has provided important insights into interphase chromatin organization. However, in evaluating the impressive body of information obtained with the light microscope, it is important to bear in mind the preparative method used in each case, because this may significantly modulate the outcome of an investigation, especially when close to the resolution limit. Studies

Figure 3. Stereo pair of a section of nucleus in which a

large region of chromosome is decorated with gold particles. Clear fiber-like structures of the order of 100 nm are seen. Scale is 500 nm. From Kireeva et al., 2008.

with living cells are likely to provide the most reliable data, whereas the use of fluorescence in situ hybridization (FISH) requires a denaturation step which may introduce structural perturbations. Muller et al. (2004) reported that 400 kb segments of nontranscribed chromatin occupy roughly spherical volumes with a diameter not much greater than the resolution limit of the microscope. A 400 kb length of DNA translates into 2000 nucleosomes, and, if present as a linear 30 nm fiber with 10 nucleosomes per 10 nm length would have a length of 2 mm. Thus, in this case, an untranscribed region of chromatin must be extremely tightly packed. However, an underlying fiber-like organization below the resolution of the light microscope cannot be excluded (Tumbar et al. 1999). It will be very interesting to see if the higher resolution offered by subdiffraction light microscopy will reveal any internal structure in these compact regions. The situation changes dramatically on transcription, when loci typically expand, often becoming linear and adopting a beaded form. In an informative study, Shopland et al. (2006) examined the intranuclear distribution of a 4 Mb segment of mouse chromosome 14 that included four gene clusters separated by gene “deserts.” By labeling the gene clusters and deserts with different colored fluorescent probes, it was possible to trace the conformation of this large segment of chromatin (Fig. 4). Four patterns occurred in roughly

C.L. Woodcock and R.P. Ghosh

Figure 4. (A) A large locus consisting of 4-Mbp region containing regions of gene “deserts” (red fluorescence)

and gene clusters (green fluorescence) is seen in the nucleus in multiple configurations (C ). In general, gene deserts are more closely associated with the heterochromatin at the nuclear periphery (B). Scale is 1 mm. From Shopland et al. 2006.

equal proportions, ranging from linear “striped” arrangements to more compact clusters. Interestingly, the inactive segments were often clustered and oriented closer to the nuclear periphery than the gene-rich segments, in keeping with the preferential location of inactive chromatin in peripheral heterochromatin. HETEROCHROMATIN AND EUCHROMATIN

The terms heterochromatin and euchromatin refer to states of compaction and transcriptional potential rather than categories of chromatin higher-order structure per se. Nevertheless, they are used ubiquitously in the literature and merit a brief mention. Heterochromatin was originally defined well before the discovery of DNA as regions of nuclei that stained strongly with basic dyes (Heitz, 1928) and, together with its counterpart euchromatin, has provided a useful qualitative indication of chromatin compaction state. In general, heterochromatin tends to be located at the nuclear periphery, where specific interactions with the envelope may occur and often forms blocks surrounding the nucleolus. These distinctions can be clearly seen in light and electron micrographs. Transcription is largely

confined to euchromatin, and it is interesting to note that Heitz presciently suggested a functional difference between the two forms. Today, the term heterochromatin is more loosely applied, and is often extended to include transcriptionally silent regions of chromatin regardless of their staining properties. An important distinction is made between constitutive and facultative heterochromatin. Constitutive heterochromatin is always compact, and tends to be enriched in repetitive, gene-poor, and late replicating DNA sequences, whereas facultative heterochromatin can reversibly undergo transitions from a compact, transcriptionally inactive state to become more open, and transcriptionally competent. In a recent review, Trojer and Reinberg (2007) suggested that facultative heterochromatin be molecularly defined as condensed, transcriptionally silent chromatin regions that decondense and allow transcription within temporal, spatial, or parental/heritable contexts. During embryogenesis, for example, the amount of facultative heterochromatin increases as unwanted sets of genes are progressively shut down until at maturity, a cell expresses only the genes appropriate for that tissue. The reverse occurs when,

Chromatin Higher-order Structure and Dynamics

for example, differentiated cells are reprogrammed to become stem cells. These events are typically accompanied by profound changes in histone variants, histone modifications, and the presence of CAPS. An important feature of heterochromatin is its propensity to “spread” to adjacent regions of chromatin (Liaw and Lustig 2006; Hines et al. 2009). A recent comparison of properties of heterochromatin and euchromatin as diffusion barriers has yielded interesting and provocative results (Bancaud et al. 2009). Measurements of diffusion constants of large polymers within these nuclear compartments confirmed that heterochromatin constitutes a more crowded environment, leading to the more efficient trapping of chromatin binding proteins such as histone H1. The crowding effect was also suggested to assist in the maintenance of heterochromatin. Further, kinetic analyses indicated an anomalous component of diffusion that was interpreted in terms of a fractal chromatin organization at spatial scales below 100 nm. This indicates a physical arrangement more similar to the “molten globule” state suggested by the lack of structure in cryo sections than to the classic 30-nm fiber. The ubiquitous location of heterochromatin at the nuclear periphery and association with the nuclear lamina and nuclear envelope suggests that this location is both structurally and functionally important. It was therefore surprising to read that, in the rod photoreceptor cells of nocturnal, but not diurnal, mammals, heterochromatin is concentrated in the center of nuclei (Solovei et al. 2009). The authors postulate that with this arrangement, nuclei act as collecting lenses, providing additional sensitivity in very low light environments. This unexpected finding underscores the plasticity of chromatin organization. CHROMATIN LOOPS

The concept of large scale chromatin loops is important in many scenarios of chromatin function. The generic term “chromatin loop” is used for a variety of phenomena that play important roles in nuclear organization and

function, but otherwise may have little in common. Further, it is not at all clear whether the various loop phenomena that have been reported constitute distinct level(s) of chromatin higher-order structure (Kadauke and Blobel 2009). It is clear, however, that loop phenomena vary in terms of their stability. For example, enhancer-promoter loops that facilitate transcription are transitory, dynamic events, whereas other types of loops appear to be more stable. An early observation that has stood the test of time is the formation of a DNA “halo” around isolated nuclei exposed to a mild treatment that releases histones (Cook and Brazell 1975). The halos were shown to consist of supercoiled DNA, suggesting that the DNA twisting that occurs when histones are removed from nucleosomes is preserved by being anchored in the residual nuclear structure. This finding is also consistent with the concept of an insoluble proteinaceous nuclear matrix or karyoskeleton to which chromatin loops are anchored by DNA sequences referred to as matrix attachment regions (MARs). However, because it has not been possible to define its composition and structure, the matrix remains a useful working concept rather than a well-accepted structure in the same sense as the cytoskeleton (Pederson 2000). A somewhat different perspective involves the “looping out” from chromosome territories of large segments of genome that are related in terms of transcriptional activity. The supposition that these different loop phenomena reflect the same underlying chromatin organization was called into question by a surprising recent finding that halo diameter was related to the spacing of origins of replication during the previous S phase (Courbet et al. 2008). Under conditions of rapid replication, origins were widely spaced, leading to large halos, whereas slow replication triggered the firing of additional replication origins, and led to smaller halos. Halos can also be generated from metaphase chromosomes, and giant meiotic lampbrush chromosomes provide a particularly compelling example of large chromatin loops and their relation to transcriptionally coupled genes. Large-scale chromatin loops are featured in some models of chromatin and

C.L. Woodcock and R.P. Ghosh

chromosome organization, but how they are integrated into chromatin/nuclear structure and whether they constitute a distinct hierarchical level of chromatin organization remains to be established. The interaction between enhancers and promoters probably represents an unrelated manifestation of chromatin looping that is critical for transcriptional activity. Enhancer sequences are often many Kb distant from promoters, and may be located upstream, downstream, or on a different chromosome. There seems now to be a consensus that physical interaction between enhancer and promoter is necessary to initiate transcription (Visel et al. 2009), but how widely separated loci are brought into contact has remained an open question. One possibility, supported by recent data (Nolis et al. 2009) is that sequence-specific transcription factors bind to both enhancer and promoter, and also bind strongly with each other. In support of this hypothesis, “decoy” transcription factor binding sites placed between a promoter and its enhancer blocked transcription, suggesting that they trapped the enhancer in unproductive loops (Nolis et al. 2009). Finally, the recently introduced technique of chromosome conformation capture (3C) and related methods, which allow mapping of physical chromatin interactions in vivo, is providing growing evidence for physical interactions between distant loci other than enhancerpromoter juxtapositions (Gondor and Ohlssen 2009). Hints that there must be mechanisms for bringing specific loci together has come from the common occurrence of some chromosomal translocations, especially those leading to human diseases. For example, Roix et al. (2003) showed that genes associated with chromosome translocations leading to human lymphomas tend to be in physical proximity and located toward the nuclear interior. The data also suggested that the phenomenon was dependent on the “higherorder spatial organization” of the genome rather than the sequences of the genes involved. Human prostate cancer offers a system for tracking the physical proximity of two loci. The TMPRSS and ERG genes are 3MB apart on human chromosome 21, but become fused in 50% of

prostate cancer cases, apparently in response to hormone levels. Using FISH to follow the locations of the two genes, Mani et al. (2009) showed that, upon hormone treatment, the two genes became physically close in a significant proportion of cells. The mechanism(s) involved in this large-scale motion remain unknown. Another recent study of intranuclear chromatin associations capitalizing on massively-parallel sequencing has yielded important insights into the large scale organization of chromatin (LiebermanAiden et al. 2009). Although the technique in its current form is limited to megabase scale resolution, the data clearly indicated that active and inactive chromatin occupy different domains within chromosome territories. Further, at this level of resolution, chromatin conformation was consistent with a knot-free, fractal globule organization. In the representation of their conclusions, Lieberman-Aiden et al. show modeled nuclei with chromatin modeled as worm-like structures of uniform diameter, and it is important to appreciate that these structures are not intended to correspond to a specific level of chromatin higher-order structure. It is interesting to note that whereas this study indicated a fractal-like organization at very large scales, unrelated work on the accessibility of differentially packed chromatin suggested a fractal conformation for chromatin below 10 nm (Bancaud et al. 2009). Hopefully, these two sets of findings obtained through widely divergent experimental approaches, augur a future in which chromatin organization can be studied effectively at many different levels. The study of chromatin long range organization using 3C and related techniques (Rusk, 2009) is clearly at an early stage in development, and promises to provide important insights into the mechanisms that bring distant loci together and their relationship to chromosome territories and chromatin dynamics. The Metaphase Chromosome

The metaphase chromosome, in which the DNA is compacted some 10,000- to 20,000fold, is the one consistent manifestation of chromatin higher-order structure. It is therefore sobering to reflect that despite a great deal of

Chromatin Higher-order Structure and Dynamics

research, there is still no widely accepted model for the internal organization of this critical cellular component. Andrew Belmont, who has contributed seminal work in the field, has aptly applied the quotation “a riddle, wrapped in a mystery, inside an enigma” to illustrate the complexities of metaphase chromosome structure. There are several features that must be accounted for in any viable model. For a given species, the diameter of chromosome arms at metaphase is quite constant, although there are wide variations in chromosome diameter and number from species to species. The width constancy appears to be independent of DNA sequence. This is seen clearly in cases in which a chromosome bears a large insert, either as a response of the cell to a genetic defect that is overcome by accumulating multiple tandem repeats of the mutated gene (e.g., Sullivan and Bickmore 2000), or by insertion of large tandemly repeated loci. Typically, these inserts, which appear at metaphase as homogeneously staining regions (HSRs) on account of their lack of band/interband organization, have a uniform diameter that closely matches that of the rest of the chromosomes. This indicates that something other than DNA sequence is establishing the underlying folding pattern that governs chromosome diameter. It is well-established that, with the exception of polytene systems, chromosomes are unineme, in that a single strand of DNA, albeit multiply folded, extends from one end (telomere) to the other (reviewed in Gall 1981). Although for a particular chromosome, metaphase banding patterns and gene locations are consistent from cell to cell, the spatial resolution of light microscopy provides very limited structural information in the context of a chromosome arm that may have a diameter of 500 nm. There is, however, a great deal of information relevant to the overall architecture of metaphase chromosomes. An early and striking result was derived from treating chromosomes to remove histones, which, like similar treatments of nuclei, revealed a “halo” of DNA loops surrounding a denser core with a size and shape comparable to the starting chromosome (Earnshaw and Laemmli

1983). The core structures were termed “scaffolds” and it was postulated that they formed the structural basis of chromosome architecture. DNase treatment could be used to isolate the scaffolds, which contained specific DNA sequences (scaffold-attachment regions or SARS) and were enriched for a few proteins thought to be essential for scaffold, and hence chromosome structural integrity. Avery fruitful approach to chromosome structure that allows experimental manipulation uses amphibian egg extracts, which can be manipulated to recapitulate, in sperm nuclei, the essential cell cycle events of DNA replication, chromosome formation and mitosis (Lohka and Masui 1983). Importantly, the system allows the roles of individual components to be explored by their addition to or removal from the extract. Support for the scaffold hypothesis was provided by the observation that, in chromosomes formed in vitro in egg extracts, some components isolated from scaffolds were distributed axially at metaphase rather than uniformly. However, subsequent experiments established that the situation was more complex. For example, topoisomerase II could be removed from chromosomes assembled in vitro without compromising their structure (Hirano and Mitchison 1993), but was essential for proper chromosome assembly and separation of sister chromatids at anaphase. Further, genetic screens for chromosome segregation mutants revealed a family of “structural maintenance of chromosomes” (SMC) proteins (Hirano 2006). Also identified were two complexes, cohesin and condensin that contain SMCs and other proteins and, in additiontootherfunctions,appear tobeinvolved in sister chromatin adhesion and chromosome condensation (Hagstrom and Meyer 2003). Condensin is consistently seen to have an axial distribution in chromosomes, and in its absence or mutation, proper condensation fails. A completely different approach, in which micromechanical properties of chromosomes are measured, has introduced a dramatically different aspect of their structure. Importantly, these results have had the salutatory effect of requiring fresh thinking to reconcile the seemingly disparate sets of data. The strategy is to

C.L. Woodcock and R.P. Ghosh

isolate chromosomes either from mitotic cells or egg extracts, hold each end with micropipettes, pull on one end, and measure the force needed for a given length increase (Houchmandzadeh et al. 1997; Poirier and Marko 2003). The environment of the chromosome can be altered by modulating the buffer with cations etc., and the effects of nucleases or proteases can be determined. The results show that chromosomes exhibit a remarkable degree of elasticity (Fig. 5), repeatedly returning to the original length after being stretched fivefold. The general conclusion from several studies is that mitotic chromosomes consists of a network or gel in which individual chromatin fibers are connected by crosslinking elements, presumably proteins (Fig. 6). It was also possible to estimate a mean distance between crosslinks of 15 Kbp by applying restriction enzymes with different cutting frequencies to chromosomes before stretching (Poirier and Marko, 2002). The estimated distance between crosslinks is close to that calculated for the chromatin loops associated with chromosome scaffolds (Gasser et al. 1986). One surprising feature of the work was the large difference in flexibility between “native” chromosomes extracted from cells by micromanipulation and those assembled in egg extracts in vitro. The latter were almost two orders of magnitude more flexible, suggesting that their non-DNA components were thin, rigid, and highly extensible (Houchmandzadeh et al. 1997) In its simplest form, the network model seems to be consistent with the lack of consistent large-scale structures seen in cryo-sections of HeLa cell chromosomes (Eltsov et al. 2008) or in EM tomograms of chromosomes formed in Xenopus egg extracts (Ko¨nig et al. 2007), but at odds with hierarchical models of chromosome structure, which would predict that stretching a chromosome would result in the unfolding of hierarchical levels with stepwise reductions in chromosome diameter. The network model also seems to contradict the many light microscopic observations of an axial localization of key proteins, especially condensins known to be involved in chromosome compaction.

Figure 5. Time sequence showing the extension of a metaphase chromosome in which the two ends are anchored by micropipettes. The diameter remains quite constant throughout. From Marko 2008.

Recently, however, models that account for most of the apparently disparate observations have been put forward. An architecture based on observations of early chromosome condensation that involves hierarchical folding with an axial “glue” provided by condensin (Fig. 7) has been proposed by Kireeva et al. (2004), and a more elaborate system with specific locations and roles for topoisomerase II, cohesin and condensin has been put forward by Marko (2007). In the case of the model in Figure 7, a delicate balance between the forces maintaining the 200– 250 nm and 500 – 750 nm fibers would Linker protein

30 nm chromatin fiber

Mitotic chromosome

Extensible net model of mitotic chromosome structure derived from force-extension measurements. From Poirier and Marko 2002.

Figure

6.

Chromatin Higher-order Structure and Dynamics

A

30 nm

B

100–130 nm

200–250 nm 500–750 nm

Figure 7. Model of chromosome formation that incorporates the concept of a central axis enriched in condensins (red dots) and irregularly folded chromatin fibers. From Kireeva et al. 2004.

be needed to produce the stretching effect shown in Figure 5. As noted by Belmont (2006), many questions remain unsolved, and there are probably additional proteins, levels of complexity, and redundant mechanisms to be discovered before the complete story of chromosome architecture, formation and maintenance is uncovered. Indeed, a new activity, RCA (regulator of chromosome architecture), that can condense chromosomes in the absence of condensins has recently been proposed (Vagnarelli et al. 2006). To date, no satisfactory explanation of the constant diameter of chromosomes has emerged. Thus, the simple-appearing metaphase chromosome is far from simple, and fully understanding its composition, formation, maintenance and architecture will likely remain a challenge for some time. LAMPBRUSH AND POLYTENE CHROMOSOMES

These two special and quite rare manifestations of chromatin provide some insight into the

surprisingly wide range of nuclear organizations that “work.” Lampbrush chromosomes (LBCs) are largely confined to meiosis in oocytes, especially avian and amphibian, but are also present in the large unicellular alga Acetabularia. They appear to be an adaptation for maximizing transcriptional output to serve thousands of cells in the developing embryo and the macroscopic cell body in Acetabularia. Light microscopy reveals an axial core from which loops of varying size extend. Loops contain transcription units that can be seen in the electron microscope to be loaded with RNA polymerases and continuously transcribing. Although the loops are the most conspicuous features of LBCs, they represent only a small fraction of the genome, the bulk forming the compact chromatin of the axial cores. The fact that sperm nuclei exposed to extract from oocytes at the lampbrush state develop into haploid LBCs (Gall and Murphey 1998) indicates that the signals needed to transform a chromosome into the lampbrush state evidently reside in the oocyte. A key question is whether this loop organization is unique to lampbrush chromosomes or whether it offers an especially clear example of a general organizing principle for transcriptionally active chromatin. Although the growing evidence for the ubiquitous organization of chromatin into large scale loops containing functionally related genes is consistent with the latter interpretation, there is also compelling evidence transcription of “normal” chromatin is organized quite differently (Hu et al. 2009). Polytene chromosomes are formed by repeated mitoses without nuclear division and consist of genomes precisely aligned in trans. They are especially prominent in dipteran larva, with 1024 copies of the genome being present in the third instar of Drosophila. In the light microscope, a unique pattern of bands and interbands is seen, allowing the locations of individual genes to be mapped cytologically. It has long been recognized that at the sites of transcriptionally active genes, “puffs” appear as the chromatin is locally decondensed. Disappointingly, polytene chromosomes have not yielded much insights into chromatin organization in general.

C.L. Woodcock and R.P. Ghosh

In the electron microscope, their alignment appears far from regular, and it is as difficult to recognize and follow individual chromatin fibers as in normal diploid nuclei. Polytene chromosomes may also be used to infer the structural organization of interphase chromatin. For example, a component that, when depleted or mutated, results in disorganization of polytene chromosomes, may be involved in sister chromatid adhesion. Polyteny is also seen in the first stages of the development of the nurse cells surrounding the Drosophila ovary. As nurse cells mature, the polytene alignment breaks down, and the nuclei become polyploid. Surprisingly, in flies with mutations of putative condensin 2 components, the polytene chromosomes fail to disperse at the appropriate time, whereas the same mutations induce dispersion of salivary gland chromosomes (Hartl et al. 2008). These findings may lead to further insights into the complex events controlling polytene chromosome integrity. Both lampbrush and polytene chromosomes dispel the idea that transcription is obligatorily confined to “factories” that contain RNA polymerases and the multiple other factors necessary for RNA synthesis and processing (Cook 1999). Observation of lampbrush loops clearly indicates that it is the polymerases that move along the loops rather than vice versa. Similarly, it seems very unlikely that DNA replication in polytene chromosomes occurs at replication factories through which DNA is translated (Cook 1999). These considerations do not lessen the potential importance of transcription and replication factories in general, but again emphasize the remarkable plasticity of nuclear organization. CHROMATIN DYNAMICS

The profound structural rearrangements that chromatin must undergo during the cell cycle underscores one aspect of the dynamic nature of chromatin. There is, in addition, compelling evidence of an important role for local structural changes in compaction and/or position of specific genetic loci likely to be of critical importance in their transcriptional regulation.

In fact, most of the functionally important characteristics of chromatin show dynamic behavior in the sense of time-dependent changes. These include the mobility of many chromatinbinding components, including chromatin architectural proteins (Phair and Misteli, 2000), and the status of posttranslational modifications, especially of the core histones. Many of these phenomena are likely to impact local and global chromatin conformation and thus modulate higher-order structure. The study of chromatin dynamics in all of its manifestations has opened up exciting new perspectives—indeed, a literature search reveals over 60 reviews with titles containing the terms “chromatin” and “dynamics” (e.g., Huebner and Spector 2010). Here, we focus on aspects of dynamics that are likely to impact higher-order structures. All involve changes in location and/ or shape of specific genetic loci that are thought to have important functional implications. As the field of chromatin dynamics has developed, it has become increasingly clear that a number of underlying physical and molecular phenomena are involved, and that to understand them fully, information at many different levels is required. For a given locus these may include the temporal and spatial scales of observed movements, their energy dependence, and their location, both within the nucleus and within their specific chromosome territory. One important goal of current research is to be able to relate these changes in location/shape to the underlying physical changes in chromatin higher-order structure. For movements that are correlated with changes in transcriptional activity, it is also important to determine whether a change in location is a prerequisite for, or a consequence of, the altered level of RNA synthesis. Advances in light microscopy techniques over the past 25 years have allowed chromatin dynamics to be examined with steadily improving temporal and spatial resolution, and has enabled three principal types of dynamic change to be recognized (Soutoglou and Misteli 2007). These differ in the extent of motion, their energy dependence, and the time scales involved (Fig. 8). Early studies tracking changes in

Chromatin Higher-order Structure and Dynamics

location of large segments of chromatin and whole chromosome territories indicated that, when corrected for nuclear rotation, the regions were essentially immobile over distances .0.4 mm (Cremer et al. 1982, Diboni and Mintz 1986, Shelby et al. 1996). However, more recent work combining higher spatial resolution with in vivo labeling of smaller defined segments of chromatin, revealed a more complex scenario with different regions of chromatin showing very different mobilities. For example, a segment of budding yeast chromatin located near a centromere was found to show constrained random walk diffusive motion with a confinement radius of 0.3 mm (Marshall et al. 1997), that was independent of the metabolic state of the cell. The rate of mobility of this region of chromatin was calculated to be approximately three orders of magnitude lower than expected for free DNA of similar length (Marshall et al. 1997), leading to the suggestion that the locus was tethered within the nucleus, perhaps to the nuclear envelope or some internal structure. A detailed study of chromatin

mobility in Drosophila spermatocytes using loci labeled with the lac repressor system recorded relatively large movements during short time intervals in early nuclei. Movements of this magnitude would be expected to result in displacements of over 4 mm in 1 h and cover the entire 11 – 12 mm diameter of a nucleus in 6– 7 h (Vazquez et al. 2001). However, when long time periods were examined, it was clear that loci were constrained within a volume approximately equivalent to a chromosome territory. Another important finding was that spermatocytes in late G2 stage and approaching meiosis showed greatly reduced chromatin mobility, suggesting that some form of additional tethering precedes chromosome condensation. A comparison of the mobility of different chromosomal sites in yeast also revealed striking locus- and cell cycle-dependent differences in chromatin dynamics (Heun et al. 2001). Some chromatin regions showed occasional large (0.5 mm) movements over time periods as short as 10 seconds that were inhibited in ATPdepleted cells and thus dependent on the

Short-range motion -Locally constrainemd within ~1μm -Fast (seconds) -Random direction -ATP dependent -Frequent -Occurs by default

Long-range motion -Several micrometers -Slow (minutes) -Directed -ATP/motor dependent -Rare -In response to physiological cues (cell cycle)

Genome Reorganization -Several micrometers -Slow (hours, days) -Global changes -Likely requires cell division -In response to physiological cues (differentiation, development)

Figure 8. The different forms of large-scale motion that contribute to chromatin dynamics. From Soutoglou and

Misteli 2007.

C.L. Woodcock and R.P. Ghosh

metabolic state of the cell. The fact that the magnitude of these large movements is similar for organisms with widely different nuclear volumes has important structural and functional implications. For example, in budding yeast, which has a nuclear diameter of 2 mm, movement within a radius of 0.5 mm would allow access to a large portion of the nuclear volume. In contrast, in a typical mammalian nucleus of 10 mm in diameter this motion would explore only a thousandth of the nuclear volume. In yeast, the effectively large displacements of loci within nuclei may promote the observed high recombination frequency, which may require substantial intermingling among different regions of the genome (Gasser 2002). Observations of “free” ectopic regions of chromatin, exemplified by large (15 kb) circular plasmids in yeast underscore the context dependence of chromatin motion. An early study using a centromere-containing plasmid reported very limited movements, similar to those observed for centromeric regions in intact chromosomes (Marshall et al. 1997). However, more recently, Gartenberg et al. (2004) using plasmids designed to be either transcriptionally silent or competent, found that the active plasmids showed unconstrained movement, whereas the silent ones were strongly constrained and preferentially located near the nuclear envelope. Also relevant to the factors controlling intranuclear mobility is the finding that whereas double-stranded DNA breaks tend to stay together through the repair process (Kruhlak et al. 2006), the absence of a critical repair factor leads to long-range movements of the broken ends (Downs et al. 2004). A general pattern emerging from numerous studies using different organisms is that all chromatin loci show constrained Brownian motion with rather similar diffusion constants, but a wide range of confinement volumes. A strong relationship between the confinement volume, and distance from the nuclear envelope has also been established, with loci closer to the envelope being more constrained. The periphery of the nucleus proximal to the nuclear envelope is considered a transcriptionally repressive environment, establishing a correlation between

transcriptional silencing and limited intranuclear mobility (Marshall 2002). The underlying mechanism for this correlated behavior is not clear, but there is evidence that the constrained motion results from physical tethering to the nuclear lamina (Heun et al. 2001; Chubb et al. 2002; Gartenberg et al. 2004). The higher spatial resolution (20 nm) of two-photon microscopy, together with a temporal resolution of 30 msec has revealed new levels of complexity for constrained diffusive motion of chromatin (Levi et al. 2005). In these experiments, GFP-labeled repeats inserted in CHO cells showed periods of rapid constrained diffusive motion alternating with occasional energy-dependent curvilinear leaps of 150 nm that lasted only 0.3 – 2.0 second. The energy-dependence of these leaps suggested that they may need the activity of chromatin remodeling complexes or other ATP-requiring changes in chromatin organization that could lead to events such as the decondensation of compact 30-nm fibers resulting in the rapid changes in location. Another important outcome of this study pertinent to the extent of physical coupling between neighboring loci on the same chromosome was the finding that loci 1– 2 mm apart in the nucleus moved independently (Levi et al. 2005). A similar conclusion was reached by Hu et al. (2009) who showed that for a 2 Mb region of chromatin that appeared as a linear cluster of beads in the light microscope, the individual beads moved independently. These findings suggest that neighboring loci are not structurally tied together by, for example, being anchored to an underlying nonchromatin structure. As it becomes possible to study yet closer regions of chromatin, there will be a point at which movement of the neighbors is correlated, and defining this point will be very useful in being able to infer the effective chromatin compaction level. Thus, although the accumulating data clearly demonstrate that chromatin in the interphase nucleus undergoes constant dynamic reorganization through constrained diffusive motion, the level(s) of chromatin higher-order structure that participate in this phenomenon remain to be determined.

Chromatin Higher-order Structure and Dynamics

Some hints of the mechanism whereby loci undergo directed movement have emerged from work on an inserted transgene that typically resides in peripheral heterochromatin, but tends to relocate toward the center of the nucleus within 1– 2 hour of transcriptional up-regulation (Chuang et al. 2006). Importantly, this movement was blocked by the expression of a mutant nuclear myosin I and by a nonpolymerizable mutant of actin. This intriguing finding should be viewed in the context that several families of actin and actinrelated proteins (ARPs) are found in the nucleus, many of which are components of chromatin remodeling and histone modification complexes (Chen and Shen 2007). Also, it has been shown that the BAF remodeling complex binds to the ends and branch points of actin filaments in a PIP2-dependent manner (Rando et al. 2002). Unlike their cytoplasmic counterparts, nuclear actins and ARPs tend to occur as monomers rather than the filamentous structures, and it is therefore curious that it was an actin polymerization mutant that was defective in long range movement (Chuang et al. 2006). Determining the mechanism by which actin/myosin influences the mobility and directionality of loci is clearly an important future goal. CHROMATIN DYNAMICS AND TRANSCRIPTION

Studies of the spatial locations and transcriptional competence of loci with respect to their chromosome territories has provided some important insights. For example, the ANT2 gene located on the X chromosome is found in the interior of the territory of the inactive homolog, but peripherally on the active homolog (Dietzel et al. 1999). In contrast, the related ANT3 gene, which is located on the pseudoautosomal region of the X chromosome was peripherally located on the territory of both active and inactive homologs. This pattern of inactive genes located inside chromosome territories, and relocating to the periphery, perhaps by a “looping out” mechanism requiring a dramatic relaxation of local chromatin structure,

has been widely observed (Chambeyron and Bickmore 2005; Chambeyron et al. 2005; Volpi et al. 2000). Other studies have documented large scale movements of loci undergoing changes in transcriptional status as part of a developmental program. Loci were found to relocate to specific regions of the nucleus rather than locations within territories. An informative example is provided by the nuclear reorganization that accompanies differentiation of mouse lymphocytes (Brown et al. 1997, 1999). Here, lymphoid genes targeted for shutdown become relocated close to the heterochromatic clusters of pericentromeric chromatin characteristic of murine cells. Importantly, in this case, transcriptional repression appears to precede movement, suggesting that relocation is not a prerequisite for silencing. The relocation of genes destined for transcriptional up-regulation away from centromeric chromatin (Schu¨beler et al. 2000) or the nuclear periphery (Kosak et al. 2002) has also been documented. Although there is a consensus regarding the correlation of intranuclear and intraterritory location with transcription, the chromatin conformational changes that accompany switches in transcriptional status are still far from clear. On the one hand, it seems unlikely that the bulky transcription machinery could access DNA and operate in compact chromatin, and in this respect, the concept of the chromatin looping out from compact domains to facilitate transcription is attractive. Indeed, transcriptional activity in LBC loops and polytene puffs is demonstrably unfolded, in some cases to the beads-on-a-string primary structure. Although there is general agreement that segments of chromatin are more compact when transcriptionally active than when silenced (e.g., Muller et al. 2001), the extent of this decondensation is unlikely to be as great as in LBC loops. A recent study designed to examine the compaction state of active and inactive chromatin in interphase nuclei has indicated quite small transcription-related changes in higher-order structure, implying that transcription can proceed in quite compact chromatin (Hu et al. 2009). Labeled 2 Mb chromatin domains designed to support a level of transcriptional

C.L. Woodcock and R.P. Ghosh

activity typical of nuclear genes were inserted into cells, and tracked over time. In the uninduced state, the arrays assumed a linear, often beaded conformation, with lengths indicative of DNA packing ratios of over 1000:1. On transcriptional induction, the arrays retained their linear, beaded conformation, and showed only a modest 1.5- to threefold increase in length. No evidence was seen for large-scale looping or unfolding to the primary beads-on-a-string level of chromatin structure. Can transcription proceed in a compact structure that is over 10-times more compact than a 30-nm fiber? In terms of access, several studies have shown that large macromolecules can penetrate quite compact chromatin (Verschure et al. 2003; Chen et al. 2005; Bancaud et al. 2009), perhaps as a function of the dynamic mobility of chromatin. Also, in a simple in vitro system, conformational fluctuations allowed direct access to the central nucleosome in a compact trinucleosome (Poirier et al. 2009). It is also possible that the binding of transcription factors could be followed by extensive, local unfolding, undetectable at the resolution of the light microscope. The many facets of chromatin dynamics span orders of magnitude both in the spatial and temporal domains. Placing this complexity in the context of the level(s) of chromatin higher-order structure that are modulated during changes in transcriptional status remains an important challenge in understanding nuclear function. The difficulty is compounded by the current uncertainty regarding chromatin organization above the 30-nm fiber level. Nevertheless, continuing improvements in the spatial resolution with which specific loci can be imaged should soon begin to narrow the possible interpretations in terms of the level(s) of chromatin structure that could be involved. For example, as it becomes possible to image closer and closer neighboring loci on the same chromatin and follow their individual dynamics, there should be a point at which coupled motion will be observed. Once that point is reached, it will become possible to combine knowledge of the physical properties of arrays of nucleosomes with the observed dynamics

and decide whether, for example, the data are consistent with a change from a compact to a decondensed 30-nm fiber. New strategies that allow “super” resolution in light microscopy will soon be in common use and may open the way to this type of correlative work. CONCLUDING REMARKS

The past few years have witnessed remarkable progress in our understanding of chromatin higher-order structure, and this rapid trajectory can be expected to continue. There are some areas of research that seem especially poised to generate significant new insights in the near future, and others that have been overly neglected. Approaches that are likely to provide important new information in the near future include the application of “super resolution” light microscopy, and further exploration of the role of RNAi and other small and long nuclear RNAs (for a review see Wilusz et al. 2009). A plethora of strategies for extending the resolution limitation of the light microscope well beyond the “diffraction limit” is maturing and some have been implemented commercially (Huang et al. 2009). Techniques capable of generating high resolution 3D images of the nuclear interior using principles such as PALM and STED will be especially suitable for studies of nuclei and chromatin. However, as experience with electron microscopy shows, high resolution alone may not be informative, especially for compact chromatin. Progress in understanding the paths of putative 30-nm fibers in nuclei will require both high resolution and a contrasting technique that highlights a single stretch of DNA/chromatin. One application of an interference-based subdiffraction technique that was focused on the nuclear envelope showed the remarkable improvement that can be expected (Schermelleh et al. 2008), but little additional detail was seen in peripheral heterochromatin. It is also important that, as resolution improves further, preparative methods that provide a level of preservation consistent with the spatial resolution of the imaging system. Such methods should be capable of

Chromatin Higher-order Structure and Dynamics

preserving membranous and cytoskeletal elements, as judged by electron microscopy. Since the discovery of RNAi and other small nuclear RNAs, their importance in a host of nuclear functions has been shown. Among these functions is a role in the formation of the constitutive heterochromatin that flanks centromeres (Verdel et al. 2004; Moazed et al. 2006; Li et al. 2009). The process involves an RNAi-containing complex (RNA-induced transcriptional silencing, RITS) that binds to specific DNA sequences via siRNA-directed base pairing, and also to histone H3 methylated at lysine 9, a heterochromatin-specific histone modification. In combination, these processes promote the spreading of heterochromatin. It seems very likely that, as the roles of small nuclear RNAs become more fully understood, they will be found to impact other aspects of chromatin higher-order structure. In addition, the identification and characterization of long noncoding RNAs that are retained in the nucleus is beginning to emerge as an important field that will add to our understanding of gene expression (see Wilusz et al. 2009 for a recent review). In the category of neglected areas that merit further study are the ionic composition of nuclei, chromosomes, and chromatin, and the perhaps related issue of the impact of intranuclear molecular “crowding” (Hancock 2004a). Given the dramatic effects of changes in conformation that isolated chromatin undergoes in response to changes in the ionic environment, it is surprising that so little is known about the intranuclear ionic environment. It is generally assumed that “physiological” salt includes 100 mM monovalent ions, and a smaller concentration (1 – 2 mM) of divalent cations. Buffers for work with isolated nuclei and chromatin in solution typically include Naþ or Kþ for the monovalent cation, Mgþþ for the divalent cation, and Cl2 for the anion. It is also well established that polyamines, especially spermidine (4þ) and spermine (8þ) are ubiquitous nuclear components with potent chromatin compaction properties. Very little is known about the intranuclear distributions and concentrations of these charged species, making it

difficult to have confidence that in vitro experiments and modeling studies are based on milieus that accurately reflect conditions in the nucleus. An isolated report of the distribution of cations in nuclei and isolated mitotic chromosomes using secondary ion mass spectrometry (Strick et al. 2001) concluded that during the transition from interphase to mitosis, Caþþ and Mgþþ concentrations increased three- to four-fold, reaching 18 mM and 11 mM respectively, whereas isolated chromosomes contained almost twice these amounts. Interestingly, Caþþ , but not Mgþþ showed an axial enrichment in chromosomes. Of the monovalent ions, nuclear Naþ concentrations were close to those measured in the cytosol, whereas Kþ was clearly enriched in both interphase and mitosis. It is not clear what the effect would be of using these remarkably high divalent ion levels in nuclear buffers, and it is unfortunate that the technique has not been further exploited using a wider range of ionic conditions. An area of research that has received little attention until recently, but may have a farreaching impact on our understanding of chromatin function and dynamics relates to the highly crowded nuclear milieu (Hancock 2004a; Richter et al. 2008; Miyoshi and Sigimoto 2008; Bancaud et al. 2009). It is clear that the extremely high concentrations of macromolecules (of the order of 100 mg/mL) that are present in the nucleus have important thermodynamic implications (Minton, 2001). These concentrations cannot be effectively recapitulated in vitro, with the consequence that experiments in solution may not reflect the events that occur in nucleo. In one set of experiments, when isolated nuclei were allowed to swell in low ionic strength medium, nucleoli and PML bodies disassembled and nucleolar transcription was dramatically reduced (Hancock 2004b). However, after adding inert macromolecules ( polyethylene glycol or dextran) to 12% w/v, the compartments reassembled, and nucleolar transcription recovered. Interestingly, this experiment also showed that it was not the lack of ions per se that caused disassembly. A fuller understanding of these effects, as well as the

C.L. Woodcock and R.P. Ghosh

intranuclear levels and roles of ions is likely to expand our understanding of chromatin and nuclear organization. This snapshot in time of chromatin higherorder structure and dynamics reveals an exciting period in research, with rapid advances being made on many fronts. As the different interweaving threads begin to coalesce, we can anticipate a much more comprehensive understanding of the structural and functional organization of chromatin in the context of its nuclear location. An overriding theme is that chromatin and its higher-order structures feature plasticity and dynamics rather than rigidity. How this is possible in what must at some level be a self-organizing system (Misteli 2001) remains a key open question.

REFERENCES Arya G, Schlick T. 2006. Role of histone tails in chromatin folding revealed by a mesoscopic oligonucleosome model. Proc Natl Acad Sci 103: 16236– 16241. Bancaud A, Huet S, Daigle N, Mozziconacci J, Beaudouin J, Ellenberg J. 2009. Molecular crowding affects diffusion and binding of nuclear proteins in heterochromatin and reveals the fractal organization of chromatin. EMBO J doi:10.1038/emboj.2009.340. Bednar J, Horowitz RA, Grigoryev SA, Carruthers LM, Hansen JC, Koster AJ, Woodcock CL. 1998. Nucleosomes, linker DNA, and linker histone form a unique structural motif that directs the higher-order folding and compaction of chromatin. Proc Natl Acad Sci 95: 14173–14178. Belmont AS. 2006. Mitotic chromosome structure and condensation. Curr Opin Cell Biol 18: 632 –638. Belmont AS, Bruce K. 1994. Visualization of G1 chromosomes: A folded, twisted, supercoiled chromonema model of interphase chromatid structure. J Cell Biol 127: 287– 302. Belmont AS, Braunfeld MB, Sedat JW, Agard DA. 1989. Large-scale chromatin structural domains within mitotic and interphase chromosomes in vivo and in vitro. Chromosoma 98: 129–43. Belmont AS, Li G, Sudlow G, Robinett C. 1999. Visualization of large-scale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol 58: 203–222. Brown KE, Baxter J, Graf D, Merkenschlager M, Fisher AG. 1999. Dynamic repositioning of genes in the nucleus of lymphocytes preparing for cell division. Mol Cell 3: 207–17. Brown KE, Guest SS, Smale ST, Hahm K, Merkenschlager M, Fisher AG. 1997. Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91: 845– 54.

Buchberger JR, Onishi M, Li G, Seebacher J, Rudner AD, Gygi SP, Moazed D. 2008. Sir3-nucleosome interactions in spreading of silent chromatin in Saccharomyces cerevisiae. Mol Cell Biol 28: 6903–6918. Chambeyron S, Bickmore WA. 2004. Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription. Genes Dev 18: 1119–1130. Chambeyron S, Da Silva NR, Lawson KA, Bickmore WA. 2005. Nuclear re-organisation of the Hoxb complex during mouse embryonic development. Development 132: 2215–2223. Chen M, Shen X. 2007. Nuclear actin and actin-related proteins in protein dynamics. Curr Opin Cell Biol 19: 326 –330. Chen D, Dundr M, Wang C, Leung A, Lamond A, Misteli T, Huang S. 2005. Condensed mitotic chromatin is accessible to transcription factors and chromatin structural proteins. J Cell Biol 168: 41–54. Cremer T, Cremer C, Baumann H, Luedtke EK, Sperling K, Teuber V, Zorn C. 1982. Rabl’s model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Hum Genet 60: 46– 56. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 31. Chubb JR, Boyle S, Perry P, Bickmore WA. 2002. Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol 12: 439–445. Clapier CR, Cairns BR. 2009. The biology of chromatin remodeling complexes. Annu Rev Biochem 78: 273– 304. Clark DJ, Kimura T. 1990. Electrostatic mechanism of chromatin folding. J Mol Biol 211: 883– 96. Cook PR. 1999. The organization of transcription and replication. Science 284: 1790–1795. Cook PR, Bazell IA. 1975. Supercoils in human DNA. J Cell Sci 19: 261– 279. Courbet S, Gay S, Arnoult N, Wronka G, Anglana M, Brison O, Debatisse M. 2008. Replication fork movement sets chromatin loop size and origin choice in mammalian cells. Nature 455: 557–560. Cui Y, Bustamante C. 2000. Pulling a single chromatin fiber reveals the forces that maintain its higher-order structure. Proc Natl Acad Sci 97: 127– 32. Daban J-R. 2000. Physical constraints in the condensation of eukaryotic chromosomes. Local concentration of DNA versus linear packing ratio in higher order chromatin structures. Biochemistry 39: 3861–3866. Daban J-R. 2003. High concentration of DNA in condensed chromatin. Biochem Cell Biol 81: 91–99 DiBoni U, Mintz AH. 1986. Curvilinear, 3-dimensional motion of chromatin domains and nucleoli in neuronal interphase nuclei. Science 234: 863– 866. Dietzel S, Schiebel K, Little G, Edelmann P, Rappold GA, Eils R, Cremer C, Cremer T. 1999. The 3D positioning of ANT2 and ANT3 genes within female X chromosome territories correlates with gene activity. Exp Cell Res 252: 363–375.

Chromatin Higher-order Structure and Dynamics Dorigo B, Schlach T, Kulangara A, Duda S, Schroeder RR, Richmond TJ. 2004. Nucleosome arrays reveal the two-start organization of the chromatin fiber. Science 306: 1571–1573. Downs JA, Jackson SP. 2004. A means to a DNA end: The many roles of Ku. Nature Rev Mol Cell Biol 5: 367 –378. Earnshaw WC, Laemmli UK. 1983. Architecture of metaphase chromosomes and chromosome scaffolds. J Cell Biol 96: 84–93. Eltsov M, Maclellan KM, Maeshima K, Frangakis AS, Dubochet J. 2008. Analysis of cryo-electron microscopy images does not support the existence of 30-nm chromatin fibers in mitotic chromosomes in situ. Proc Natl Acad Sci 105: 19732–19737. Fan Y, Nikitina T, Morin-Kensicki EM, Zhao J, Magnuson TR, Woodcock CL, Skoultchi AI. 2003. H1 linker histones are essential for mouse development and affect nucleosome spacing in vivo. Mol Cell Biol 23: 4559–72. Fan JY, Rangasamy D, Luger K, Tremethick DJ. 2004. H2A.Z alters the nucleosome surface to promote HP1alphamediated chromatin fiber folding. Mol Cell 16: 655– 61. Fischle W, Wang Y, Jacobs SA, Kim Y, Allis CD, Khourasanizadeh S. 2003. Molecular basis for the discrimination of repressive methyl-lysine marks in histone H3 by Polycomb and HP1 chromodomains. Genes Dev 17: 1870– 1881. Gall JG. 1981. Chromosome structure and the C-value paradox. J Cell Biol 91: 3s– 14s. Gall J, Murphy C. 1998. Pictures in cell biology. Lampbrush chromosomes from sperm chromatin. Trends Cell Biol 8: 207. Gartenberg MR, Neumann FR, Laroche T, Blaszczyk M, Gasser SM. 2004. Sir-mediated repression can occur independently of chromosomal and subnuclear contexts. Cell 119: 955– 67. Gaspar-Maia A, Alajem A, Polesso F, Sridharan R, Mason MJ, Heidersbach A, Ramalho-Santos J, McManus MT, Plath K, Meshorer E, et al. 2009. Chd1 regulates open chromatin and pluripotency of embryonic stem cells. Nature 460: 863–8. Gasser SM, 2002, Visualizing chromatin dynamics in interphase nuclei. Science 296: 1412– 1416. Gasser SM, Laroche T, Falquet J, Boy de la Tour E, Laemmli UK. 1986. Metaphase chromosome structure. Involvement of topoisomerase II. J Mol Biol 188: 613 –29. Gondor A, Ohlsson R. 2009. Chromosome cross-talk in three dimensions. Nature 461: 199 –205. Greaves IK, Rangasamy D, Ridgway P, Tremethick DJ. 2007. H2A.Z contributes to the unique 3D structure of the centromere. Proc Natl Acad Sci 104: 525–530. Grigoryev SA, Arya G, Correll S, Woodcock CL, Schlick T. 2009. Evidence for heteromorphic chromatin fibers from analysis of nucleosome interactions. Proc Natl Acad Sci 106: 13317– 22. Hagstrom KA, Meyer BJ. 2003. Condensin and cohesin: More than chromosome compactor and glue. Nat Rev Genet 4: 520 –534. Hake SB, Allis CD. 2006. Histone H3 variants and their potential role in indexing mammalian genomes: The “H3 barcode hypothesis.” Proc Natl Acad Sci 103: 6428– 35.

Hancock R. 2004a. A role for molecular crowding effects in the assembly and function of compartments in the nucleus. J Struct Biol 146: 281 –290. Hancock R. 2004b. Internal organisation of the nucleus: Assembly of compartments by macromolecular crowding and the nuclear matrix model. Biol Cell 96: 595–601. Hartl TA, Smith HF, Bosco G. 2008. Chromosome alignment and transvection are antagonized by condensin II. Science. 322: 1384–7. Heitz E. 1928. Das Heterochromatin der Moose, 1, Jahrb Wiss Bot 69: 762– 818. Heun P, Laroche T, Shimada K, Furrer P, Gasser SM. 2001. Chromosome dynamics in the yeast interphase nucleus. Science 294: 2181– 2186. Hines KA, Cryderman DE, Flannery KM, Yang H, Vitalini MW, Hazelrigg T, Mizzen CA, Wallrath L. 2009. Domains of HP1 Required for Drosophila melanogaster heterochromatin spreading. Genetics. 182: 967– 77. Hirano T. 2006. At the heart of the chromosome: SMC proteins in action. Nat Rev Mol Cell Biol 7: 311 –322. Hirano T, Mitchison. 1993. Topoisomerase II does not play a scaffolding role in the organization of mitotic chromosomes assembled in Xenopus egg extracts. J Cell Biol 120: 601–612. Horowitz RA, Agard DA, Sedat JM, Woodcock CL. 1994. The three dimensional archeitecture of chromatin in situ: Electron tomography reveals fibers composed of a continuously variable zigzag nucleosomal ribbon. J Cell Biol 125: 1 –10. Houchmandzadeh B, Marko JF, Chatenay D, Libchaber A. 1997. Elasticity and structure of eukaryote chromosomes studied by micromanipulation and micropipette aspiration. J Cell Biol 139: 1– 12. Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009. Large-scale chromatin structure of inducible genes: Transcription on a condensed, linear template. J Cell Biol 185: 87– 100. Huang B, Bates M, Zhuang X. 2009. Super-resolution fluorescence microscopy. Annu Rev Biochem 78: 993– 1016. Huebner MR, Spector DL. 2010. Chromatin dynamics. In Press. Jacobs SA, Khorasanizadeh S. 2002. Structure of the HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science 295: 2080–2083. Jenuwein T, Allis CD. 2001. Translating the histone code. Science 293: 1074– 80. Jiang C, Pugh BF. 2009. Nucleosome positioning and gene regulation: Advances through genomics. Nat Rev Genet 10: 161– 72. Kadauke S, Blobel GA. 2009. Chromatin loops in gene regulation. Biochim Biophys Acta 1789: 17–25. Kireev I, Lakonishok M, Liu W, Joshi VN, Powell R, Belmont AS. 2008. In vivo immunogold labeling confirms largescale chromatin folding motifs. Nat Methods 5: 311– 313. Kireeva N, Lakonishok M, Kireev I, Hirano T, Belmont AS. 2004. Visualization of early chromosome condensation: A hierarchical folding, axial glue model of chromosome structure. J Cell Biol 166: 775– 785. Ko¨nig P, Braunfeld MB, Sedat JW, Agard DA. 2007. The three-dimensional structure of in vitro reconstituted

C.L. Woodcock and R.P. Ghosh Xenopus laevis chromosomes by EM tomography. Chromosoma 116: 349 –372. Kosak ST, Skok JA, Medina KL, Riblet R, Le Beau MM, Fisher AG, Singh H. 2002. Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296: 158– 62. Kouzarides T. 2007. Chromatin modifications and their function. Cell 128: 693–705. Kruhlak MJ, Celeste A, Nussenzweig A. 2006. Spatiotemporal dynamics of chromatin containing DNA breaks. Cell Cycle 5: 1910–1912. Kruithof M, Chien FT, Routh A, Logie C, Rhodes D, van Noort J. 2009. Single-molecule force spectroscopy reveals a highly compliant helical folding for the 30-nm chromatin fiber. Nat Struct Mol Biol 16: 534– 40. Li H, Motamedi MR, Yip C K, Wang Z, Waltz T, Patel DJ, Moazed D. 2009. An Alpha Motif at Tas3 C Terminus Mediates RITS cis Spreading and Promotes Heterochromatic Gene Silencing. Mol Cell 34: 155– 167. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. 2009. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326: 289 –93. Levi V, Ruan Q, Plutz M, Belmont AS, Gratton E. 2005, Chromatin dynamics in interphase cells revealed by tracking in a two-photon excitation microscope. Biophys J 89: 4275–85. Liaw H, Lustig AJ. 2006. Sir3 C-terminal domain involvement in the initiation and spreading of heterochromatin. Mol Cell Biol 26: 7616–7631. Lohka MJ, Masui Y. 1983. Formation in vitro of sperm pronuclei and mitotic chromosomes induced by amphibian ooplasmic components. Science 220: 719– 721. Lu X, Wontakal SN, Emelyanov AV, Morcillo P, Konev AY, Fyodorov DV, Skoultchi AI. 2009. Linker histone H1 is essential for Drosophila development, the establishment of pericentric heterochromatin, and a normal polytene chromosome structure. Genes Dev 23: 452– 65. Luger K, Hansen JC. 2005. Nucleosome and chromatin fiber dynamics. Curr Opin Struct Biol 15: 188 –196. Luger K, Mader AW, Richmond RK, Sargent DF, Richmond TJ. 1997. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389: 251–60. Mani RS, Tomlins SA, Callahan K, Ghosh A, Nyati MK, Varambally S, Palanisamy N, Chinnaiyan AM. 2009. Induced chromosomal proximity and gene fusions in prostate cancer. Science 326: 1230. Marko JF. 2008. Micromechanical studies of mitotic chromosomes. Chromosome Res 16: 469– 497. Marko M, Hsieh C, Salmon N, Rodriguez M, Frank J, Manella C. 2008. FIB milling of frozen hydrated cells and tissues for TEM cryo-tomography. Microsc Microanal 13: 968CD. Marko M, Hsieh C, Schalek R, Frank J, Mannella C. 2007. Focused-ion-beam thinning of frozen-hydrated biological specimens for cryo-electron microscopy. Nat Methods 4: 215 –7. Maresca TJ, Heald R. 2006. The long and the short of it: Linker histone H1 is required for metaphase chromosome compaction. Cell Cycle 5: 589– 91.

Marshall WF. 2002. Order and disorder in the cell nucleus. Curr Biol 12: R185–R192. Marshall WF, Straight A, Marko JF, Swedlow J, Dernberg A, Belmont A, Murray AW, Agard DA, Sedat J. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930–939. Minton AP. 2001. The influence of macromolecular crowding and macromolecular confinement on biological reactions in physiological media. J Biol Chem 276: 10577– 10580. Misteli T. 2001. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Miyoshi D, Sigimoto N. 2008. Molecular crowding effects on structure and stability of DNA. Biochimie 90: 1040–1051. Moazed D, Bu¨hler M, Buker SM, Colmenares SU, Gerace EL, Gerber SA, Hong EJ, Motamedi MR, Verdel A, Ville´n J, Gygi SP. 2006. Studies on the mechanism of RNAidependent heterochromatin assembly. Cold Spring Harb Symp Quant Biol 71: 461–471. Muller WG, Rieder D, Kreth G, Cremer C, Trajanoski Z, McNally JG. 2004. Generic features of tertiary chromatin structure as detected in natural chromosomes. Mol Cell Biol 24: 9359– 9370. Muller WG, Walker D, Hager G, McNally J. 2001. Large-scale chromatin decondensation and recondensation regulated by transcription from a natural promoter. J Cell Biol 154: 33– 48. Nolis IK, McKay DJ, Mantouvalou E, Lomvaradas S, Merika M, Thanos D. 2009. Transcription factors mediate longrange enhancer-promoter interactions. Proc Natl Sci 106: 20222–7. Pederson T. 2000. Half a century of “the nuclear matrix.” Mol Biol Cell 11: 799– 805. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian nucleus. Nature 404: 604–609. Poirier MG, Marko JF. 2002. Mitotic chromosomes are chromatin networks without a mechanically contiguous protein scaffold. Proc Natl Acad Sci 99: 15393– 15397. Poirier MG, Marko JF. 2003. Micromechanical studies of mitotic chromosomes. Curr Top Dev Biol 55: 75–141. Poirier MG, Oh E, Tims HS, Widom J. 2009. Dynamics and function of compact nucleosome arrays. Nat Struct Mol Biol 16: 938 –44. Rando OJ, Zhao K, Janmey P, Crabtree GR. 2002. Phosphatidylinositol-dependent actin filament binding by the SWI/SNF-like BAF chromatin remodeling complex. Proc Natl Acad Sci 99: 2824– 2829. Richter K, Nessling M, Lichter P. 2008. Macromolecular crowding and its potential impact on nuclear function. Biochim Biophys Acta 1783: 2100–2107. Robinson PJ, Fairall L, Huynh VA, Rhodes D. 2006. EM measurements define the dimensions of the “30-nm” chromatin fiber: Evidence for a compact, interdigitated structure. Proc Natl Acad Sci 103: 6506– 6511. Roix JJ, McQueen PG, Munson PJ, Parada LA, Misteli T. 2003. Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34: 287– 291. Routh A, Sandin S, Rhodes D. 2008. Nucleosome repeat length and linker histone stoichiometry determine chromatin fiber structure. Proc Natl Acad Sci 105: 8872–8877.

Chromatin Higher-order Structure and Dynamics Rusk N. 2009. When ChIA PETs meet Hi-C. Nature Methods 6: 863. Santenard A, Torres-Padilla ME. 2009. Epigenetic reprogramming in mammalian reproduction: Contribution from histone variants. Epigenetics 4: 80–4. Schermelleh L, Carlton PM, Haase S, Shao L, Winoto L, Kner P, Burke B, Cardoso MC, Agard DA, Gustafsson MG, et al. 2008. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320: 1332– 1336. Schlach T, Duda S, Sargent DF, Richmond TJ. 2005. X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature 436: 138– 141. Schu¨beler D, Francastel C, Cimbora DM, Reik A, Martin DI, Groudine M. 2000. Nuclear localization and histone acetylation: A pathway for chromatin opening and transcriptional activation of the human beta-globin locus. Genes Dev 14: 940 –50. Shelby RD, Hahn KM, Sullivan KF. 1996. Dynamic elastic behavior of alpha satellite domains visualized in living cells. J Cell Biol 135: 545–557. Simpson RT. 1991. Nucleosome positioning: Occurrence, mechanisms, and functional consequences. Prog Nucleic Acid Res Mol Biol 40: 143 –84. Shogren-Knaak M, Ishii H, Sun JM, Pazin MJ, Davie JR, Peterson CL. 2006. Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science 311: 844– 847. Shopland LS, Lynch CR, Peterson KA, Thornton K, Kepper N, Hase J, Stein S, Vincent S, Molloy KR, Kreth G, et al. 2006. Folding and organization of a contiguous chromosome region according to the gene distribution pattern in primary genomic sequence. J Cell Biol 174: 27–38. Simpson RT, Thoma F, Brubaker J. 1985. Chromatin reconstituted from tandemly repeated cloned DNA fragments and core histones: A model system for study of higher order structure. Cell 42: 799 –808. Solovei I, Kreysing M, Lanctoˆt C, Ko¨sem S, Peichl L, Cremer T, Guck J, Joffe B. 2009. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell. 137: 356 –68. Soutoglou E, Misteli T. 2007. Mobility and immobility of chromatin in transcription and genome stability. Curr Opin Genet Dev 17: 435 –4. Strick R, Strissel PL, Gavrilov K, Levi-Setti R. 2001. Cation-chromatin binding as shown by ion microscopy is essential for the structural integrity of chromosomes. J Cell Biol 155: 899 –910. Strukov YG, Belmont AS. 2009. Mitotic chromosome structure: Reproducibility of folding and symmetry between sister chromatids. Biophys J 96: 1617–28. Sullivan BA, Bickmore WA. 2000. Unusual chromosome architecture and behaviour at an HSR. Chromosoma 109: 181– 9.

Suto RK, Clarkson MJ, Tremethick DJ, Luger K. 2000. Crystal structure of a nucleosome core particle containing the histone variant H2A.Z. Nature Struct Biol 7: 1121–1124. Tremethick DJ. 2007. Higher-order structures of chromatin: The elusive 30 nm fiber. Cell 128: 651– 654. Trojer P, Reinberg D. 2007. Facultative heterochromatin: Is there a molecular signature. Mol Cell 28: 1 –12. Tumbar T, Sudlow G, Belmont AS. 1999. Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain. J Cell Biol 145: 1341– 54. Vagnarelli P, Hudson DF, Ribeiro SA, Trinkle-Mulcahy L, Spence JM, Lai F, Farr CJ, Lamond AI, Earnshaw WC. 2006. Condensin and Repo-Man-PP1 co-operate in the regulation of chromosome architecture during mitosis. Nat Cell Biol 8: 1133– 1142. van Holde K. 1989. Chromatin. Springer Verlag, New York. van Holde K, Zlatanova J. 1995. Chromatin structure: Chasing a mirage? J Biol Chem 270: 8373–8376. van Holde K, Zlatanova J. 2007. Chromatin fiber structure: Where is the problem now? Sem Cell Dev Biol 18: 651– 658. Vazquez J, Bemont AS, Sedat JW. 2001. Multiple regimes of constrained chromosome motion are regulated in the interphase Drosophila nucleus. Curr Biol 11: 1227–1239. Verdel A, Jia S, Gerber S, Sugiyama T, Gygi S, Grewal SI, Moazed D. 2004. RNAi-mediated targeting of heterochromatin by the RITS complex. Science 303: 672– 676. Verschure PJ, van der Kraan I, Manders EM, Hoogstraten D, Houtsmuller AB, van Driel R. 2003. Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules. EMBO Rep 4: 861 –866. Visel A, Rubin EM, Pennacchio LA. 2009. Genomic views of distant-acting enhancers. Nature 461: 199– 205. Volpi EV, Chevret E, Jones T, Vatcheva R, Williamson J, Beck S, Campbell RD, Goldsworthy M, Powis SH, Ragoussis J, et al. 2000. Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113: 1565–2576. Wilusz JE, Sunwoo H, Spector DL. 2009. Long non-coding RNAs: Functional surprises from the RNA world. Genes Dev 23: 1494–1504. Woodcock CL. 2006. Chromatin architecture. Curr Opin Struct Biol 16: 213 –220. Woodcock CL, Dimitrov S. 2001. Higher-order structure of chromatin and chromosomes. Curr Opin Genet Dev 11: 130 –135. Woodcock CL, Horowitz RA. 1995. Chromatin organization re-viewed. Trends Cell Biol 5: 272– 277. Woodcock CL, Skoultchi AI, Fan Y. 2006. Role of linker histone in chromatin structure and function: H1 stoichiometry and nucleosome repeat length. Chromosome Res 14: 17–25.

Nuclear Organization and Dosage Compensation Jennifer C. Chow and Edith Heard Mammalian Developmental Epigenetics Group, Institut Curie, CNRS UMR3215, INSERM U934, Paris, F-75248 France Correspondence: [email protected]

Dosage compensation is a strategy to deal with the imbalance of sex chromosomal gene products relative to autosomes and also between the sexes. The mechanisms that ensure dosage compensation for X-chromosome activity have been extensively studied in mammals, worms, and flies. Although each entails very different mechanisms to equalize the dose of X-linked genes between the sexes, they all involve the co-ordinate regulation of hundreds of genes specifically on the sex chromosomes and not the autosomes. In addition to chromatin modifications and changes in higher order chromatin structure, nuclear organization is emerging as an important component of these chromosome-wide processes and in the specific targeting of dosage compensation complexes to the sex chromosomes. Preferential localization within the nucleus and 3D organization are thought to contribute to the differential treatment of two identical homologs within the same nucleus, as well as to the chromosome-wide spread and stable maintenance of heterochromatin.

pecies that have evolved a chromosomally based sex determination system face the problem of dosage compensation of sex chromosomes between males and females. As the non-recombining chromosome degenerates, the heterogametic sex is left with only one functional copy of genes on the sex chromosomes compared with the double dose that exists in the homogametic sex. Diverse dosage compensation strategies have evolved in different species to deal with this problem and have been most extensively studied in flies (D. melanogaster), worms (C. elegans), and mammals. In flies, males (XY) up-regulate their single X to achieve expression levels equivalent to the two

S

copies present in females (reviewed in Gelbart and Kuroda 2009). Increased transcriptional output of the X chromosome is also observed in worms (Gupta et al. 2006) and mammals (Nguyen and Disteche 2006; Lin et al. 2007). However, in these latter cases, hyper-expression is not specific to the heterogametic sex, as it occurs in both males and females, necessitating secondary compensation mechanisms in females who would otherwise over-express Xlinked genes. In C. elegans, hermaphrodites (XX) down-regulate transcription from both their X chromosomes by half, whereas in female mammals, one of the two X chromosomes is chosen for inactivation (reviewed in Vicoso

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000604 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000604

J.C. Chow and E. Heard

and Bachtrog 2009). Therefore, it is through the co-ordinate regulation of hundreds of genes across the chromosome that these compensation mechanisms restore not only the balance between sexes but also the ratio of gene dosage between the sex chromosomes and the autosomes. How dosage compensation complexes are targeted specifically to the sex chromosomes on a chromosome-wide scale is still not fully understood in any of these organisms. Changes in chromatin structure and higher-order chromosome architecture clearly participate, but it is also becoming increasingly clear that in all of these dosage compensation systems, nuclear organization may play a role.

NUCLEAR ORGANIZATION AND DOSAGE COMPENSATION

The eukaryotic nucleus is a highly organized organelle consisting of multiple compartments. In addition to more stable structures, such as the nuclear envelope and the nucleolus, it also contains numerous specialized functional compartments, where processes such as RNA transcription and processing, as well as DNA replication and repair, take place (reviewed in Spector 2003; Misteli 2007). Chromosomes themselves also exhibit a distinct 3D organization and can adopt non-random positions within the nucleus, both with respect to each other and with respect to specific nuclear landmarks. It is becoming clear that this spatial organization likely plays an important role in the regulation and expression of the genome and is believed to be a key factor in the chromosomewide regulation that occurs during dosage compensation. In particular, the interphase organization of X-chromosome sequences may facilitate their differential activity compared to autosomes or between the sexes. Similarly, preferential localization in specialized nuclear subcompartments may contribute to specific processes that affect the sex chromosomes more than the autosomes. In the case of mammals, nuclear compartmentalization is likely to facilitate the differential treatment of the two X chromosomes within the same nucleus.

Drosophila melanogaster

In Drosophila, dosage compensation is mediated by a ribonucleoprotein complex that binds hundreds of sites on the single X chromosome in males, and induces a two-fold up-regulation of X-chromosomal genes (Fig. 1A). The complex, known as the male-specific lethal (MSL) complex, is made up of five protein subunits (MSL1, MSL2, MSL3, MOF, MLE) and two non-coding RNAs (roX1, roX2) (reviewed in Gelbart and Kuroda 2009; Hallacli and Akhtar 2009). It can only form in male cells, due to the male-specific expression of the male-specificlethal 2 (MSL2) protein. MSL2 is required to stabilize components of the complex such as MSL1 and roX RNAs. MOF and MLE are enzymatic components, MLE being an RNA/DNA helicase and MOF a histone acetyltransferase, responsible for acetylation of histone H4 at lysine 16 (H4K16Ac). The H4K16Ac chromatin modification is believed to be a key component in the transcriptional up-regulation across the X chromosome in males (Bone et al. 1994). MSL1 and MSL2 form the core of the complex and are thought to be the main targeting components; however, the full complex is required for complete localization to the X chromosome (Gelbart and Kuroda 2009). Targeting to the X chromosome is thought to occur through co-transcriptional assembly of the complex at the sites of roX1 and roX2 transcription on the X chromosome (Kelley et al. 1999). From these assembly sites, the complex then spreads to other high affinity binding sites on the X or chromatin entry sites (CES), and is thought to occur, at least partially, through the recognition of specific DNA sequence motifs (Alekseyenko et al. 2008; Straub et al. 2008) (Fig. 1A). Nevertheless, the identified DNA motifs alone are unable to unequivocally predict MSL complex binding since many motifs are not associated with MSL binding; therefore, other factors must also be involved. It has been proposed that nuclear organization and higher order chromatin structure may play a role in this regard, perhaps facilitating the spread of the complexes to the actively transcribed portions of the chromosome. In fact, the X chromosome has been found

Nuclear Organization and Dosage Compensation

A

D. melanogaster

Males: Two-fold up-regulation of single X chromosome

Females

mRNA more rapidly processed or exported?

Nuclear pore

Nuclear pore

MSL complex

X

roX1

Clustering of MSL complexbound high affinity sites may aid in the spread to lower affinity sites

roX2

Y High affinity sites Low affinity sites Lowest affinity sites

roX RNA

roX 1,2 loci

Transcriptionally active genes roX 1 and 2 high affinity chromatin entry sites High affinity sites Low affinity sites Lowest affinity sites MSL complex H4K16 acetylated chromatin

B

C. elegans

Hermaphrodites: Down-regulation of both X chromosomes by half Nuclear membrane

Males Nuclear membrane

DCC complex

X X High affinity sites (rex sites) Low affinity sites (dox sites)

C

DCC-binding to high affinity sites and spread to low affinity sites leads to global chromosome-wide repression

Mammals

Females Inactive X

Xi Xa

Xic Xic

High affinity sites (rex sites) Low affinity sites (dox sites) Genes Dosage compensation complex

Inactivated genes are recruited into the compartment

Unknown Xist RNA “entry” sites (some genes?)

Genes that Waystations/booster elements (repeats, including LINEs - facilitators of repetitive escape remain Active LINE1 core formation) elements facilitate external Waystations/booster elements (young, active LINEs) silencing

Nuclear membrane

Active X

Xist RNA Xist RNA creates a silent compartment made up of silent LINE1 elements and other repeats

RNA Pol II

Figure 1. Diverse strategies for dosage compensation in different organisms. (A) In D. melanogaster, males (XY)

up-regulate two-fold their single X chromosome. The dosage compensation complex has been found to be associated with nuclear pore proteins, which may indicate a participation of nuclear localization in this upregulation. (B) C. elegans hermaphrodites (XX) down-regulate both X chromosomes by one half. This may be linked to a particular organization of the two Xs with the dosage compensation machinery. (C) Female mammals inactivate one of their two X chromosomes. The X chromosomes often reside at the nuclear periphery or at the nucleolus. The inactive X is organized as a silent, repetitive compartment by Xist RNA, into which genes are located as they become silenced.

to adopt a male-specific 3D conformation with “clustering” of the high affinity MSL complex binding sites occurring in males, but not females (Grimaud and Becker 2009). In addition, MSL proteins co-purify with components of the nuclear pore complex, Nup153 and Megator (Mtor), and RNAi knockdowns have shown

that these factors are essential for Msl-mediated dosage compensation in male cells (Mendjan et al. 2006). Although the nuclear periphery is often associated with heterochromatin and transcriptional silencing, there are sub-regions at the nuclear envelope, particularly at nuclear pores, that are associated with transcriptional

J.C. Chow and E. Heard

activity (Taddei et al. 2004; Akhtar and Gasser 2007). Recently, the genome-wide chromatin binding profiles of Nup153 and Megator have been reported (Vaquerizas et al. 2010). These Nucleoporin-Associated Regions (NARs) represent regions of open chromatin as they are enriched in markers for active transcription such as RNA polymerase II and histone H4K16 acetylation. Interestingly, the male hyperactive X chromosome is particularly enriched in NARs and relies on the interaction with nucleoporins for its peripheral localization (Vaquerizas et al. 2010). The link between the MSL complex, nuclear pore components, and the nuclear periphery suggests that the X chromosome in males specifically localizes to nuclear sub-domains through its interactions with the nuclear pore and that this might contribute to its two-fold up-regulation. Thus, the enhanced expression of the X chromosome in males may be not only transcriptional via chromatin-mediated effects, but also posttranscriptional, perhaps through the more rapid processing and export of messenger RNA, thanks to the association with the nuclear pore. Caenorhabditis elegans

In C. elegans, dosage compensation involves a two-fold reduction in X-chromosomal gene activity in hermaphrodites (Fig. 1B). The worm dosage compensation complex (DCC) consists of a hermaphrodite-specific core of SDC (sex determination and dosage compensation) proteins, required for the stabilization and targeting of the complex to the X chromosome, as well as a condensin subcomplex, which somehow induces the chromatin changes necessary for dosage compensation. How the DCC imposes a precise two-fold down-regulation of X-linked gene expression in a chromosomewide manner remains unclear. The DCC is targeted to X chromosomes by initial binding to a number of recruiting elements, followed by dispersal or spreading to secondary sites. However, DCC binding does not correlate with DCCmediated repression, implying that the complex acts in a chromosome-wide manner, rather than on a gene-by-gene basis. Indeed, it has been

proposed that the DCC does not repress genes by direct binding, but rather acts at a distance and may involve looping of chromatin fibers to bring regulated genes to the vicinity of DCC bound sites (Jans et al. 2009). The implication of condensins suggests that X-chromosome down-regulation may involve similar mechanisms to mitotic chromosome condensation (Chuang et al. 1996; Lieb et al. 1996; Lieb et al. 1998; Tsai et al. 2008; Csankovszki et al. 2009). The DCC may alter higher order chromosome organization by inducing partial chromosome condensation at interphase, and this might in turn lead to decreased transcription. This is supported by the fact that the DCC condensin subunit differs from the mitotic condensin I complex by only a single subunit. Although the exact molecular mechanisms of condensin-mediated X chromosome down-regulation are still unclear, the consensus is that this process must involve some higher-order reorganization of the X chromosomes within the nucleus. Whether nuclear compartmentalization or localization also plays a role remains to be seen. NUCLEAR ORGANIZATION OF THE INACTIVE X CHROMOSOME IN FEMALE MAMMALS

In mammals, the presence of an inactive X (Xi) chromosome in female cells was first noted at the cytological level by Barr in 1949 as a heteropycnotic body that was often at the nuclear periphery or within the perinucleolar region (Barr and Bertram 1949). These initial observations were already suggestive of a possible role for nuclear compartmentalization in the formation/maintenance of the inactive state (Comings 1968). The nuclear and perinucleolar compartments are indeed often associated with heterochromatin (de Wit and van Steensel 2009), and in the case of the mammalian X chromosome, this might promote the inactive state by facilitating access to heterochromatic factors or restricting access to transcription factors. The nucleolar localization of the Xi (Zhang et al. 2007; Rego et al. 2008) has been proposed to be important for faithful replication of the Xi’s epigenetic state, as the nucleolar

Nuclear Organization and Dosage Compensation

periphery is enriched in factors required for replication of heterochromatin, such as Snf2h (Zhang et al. 2007). In addition to its preferential locations in the nucleus, the inactive X chromosome also appears to adopt a very distinctive 3D organization. In both mouse and humans, fluorescence in situ hybridization (FISH) studies have shown that the inactive X chromosome is coated by the non-coding Xist RNA, which triggers the initiation of X inactivation. Xist induces global changes in chromatin structure, and also mediates changes in the global organization of the chromosome, both of which are thought to play roles in chromosome-wide silencing. Xist RNA initially coats a central core of the X-chromosome territory that is made up largely of repetitive DNA (Chaumeil et al. 2006; Chow et al. 2010). X-linked genes are mostly excluded from this inner region, particularly at the early stages of inactivation and instead reside outside, or at the periphery, of this repetitive compartment (Chaumeil et al. 2006). However, as X inactivation proceeds and genes become silenced, they move into the Xist RNA coated, silent compartment. There is thus a progressive nuclear reorganization of the X chromosome as it becomes inactivated and heterochromatic, ultimately resulting in a condensed structure with a repetitive core and silent genes embedded into its periphery (Clemson et al. 2006) (Figs. 1C, 2A). The degree to which this particular 3D organization of the Xi is due to a compartmentalizing function of Xist RNA, or to the chromatin changes that are induced during X chromosome inactivation (XCI) or to the repetitive nature of the mammalian X chromosome, still remains to be seen. HETEROGENEOUS HETEROCHROMATIN ON THE INACTIVE X CHROMOSOME

Inactivation of the X chromosome involves the acquisition of many chromatin changes that clearly distinguish it from its active homolog. Immunofluorescence studies in mitotic and/or interphase cells show that several histone marks typical of heterochromatin appear to be enriched on the Xi, including H3K27me3, H3K9me2, H3K9me3, H4K20me1, H2AK119Ub, and the

histone variant macroH2A (reviewed in Heard and Disteche 2006). Conversely, euchromatic marks such as H3K4me2/3 and H3 and H4 acetylated lysines are largely depleted. However, these modifications are not uniformly distributed across the chromosome, suggesting that the inactive X chromosome is made up of several different types of heterochromatin. In human somatic cell lines, at least two different heterochromatic states have been characterized by immunofluorescence: One is defined by the presence of XIST RNA, macroH2A, H3K27me3, H4K20me1, and H2AK119Ub, and the other is defined by features more reminiscent of constitutive heterochromatin, including later replication timing, H3K9me3, H4K20me3, and association with HP1. These different heterochromatin domains appear to remain spatially distinct from each other, both during metaphase and interphase (Chadwick and Willard 2004; Chadwick 2007). Ultra-structural analyses of the H3K27me3-enriched portion of the Xi by light and electron microscopy in mouse and human fibroblasts (Rego et al. 2008) have shown that this portion of the heterochromatic X has a distinctive structure, being made up of densely packed fibers with intervening spaces and a mean chromatin compaction higher than euchromatin, but less than constitutive, H3K9me3-enriched heterochromatin (Fig. 2B, C). The significance of these different types of heterochromatin and whether they reflect differences in the epigenetic mechanisms underlying the initiation and/or maintenance of the inactive state is still unknown. The relative proportions of these two heterochromatic signatures varies considerably between species. In mouse, enrichment of HP1 and H3K9me3 on the Xi is much less distinctive than has been described in humans (Peters et al. 2002; Kohlmaier et al. 2004). Nevertheless, it should be noted that in both of these species, the inactive X appears to be similarly organized in the nucleus, with genes surrounding a silent repetitive core (Chaumeil et al. 2006; Clemson et al. 2006). In marsupials, although the Xi is clearly depleted for active marks as on the mouse and human Xi (Koina et al. 2009; Mahadevaiah et al. 2009), the degree of enrichment on the Xi

J.C. Chow and E. Heard

A

5 μm

Xi

Xi

Xi

Xi *Xa

*Xa

1 μm

*Xa

LINE1 DNA Lamp2,G6pdX, AtrX,Fmr1, Huwe1 DNA

LINE1 DNA Xist RNA

LINE1 DNA Lamp2,G6pdX, AtrX,Fmr1, Huwe1 DNA Xist RNA

B

nu

1 μm

2 μm 2 μm

C Nu

1 μm S11

S11

Figure 2. Nuclear organization of the inactive X chromosome in female mammals. (A) Combined RNA-DNA

fluorescence in-situ hybridization in day 8 differentiated female ES cells shows the distinctive organization of the silent inactive X (Xi) compared to the active X (Xa). Both the Xa and the Xi often reside at the periphery of the nucleus. However, on the Xi, Xist up-regulation and coating results in the formation of a transcriptionally silent compartment made up of a central core of repetitive sequences, including LINE1 elements. Genes transcriptionally silenced at this stage (Lamp2, G6pdX, AtrX, Fmr1, and Huwe1) have been recruited into the compartment. All images shown are single slices taken from the same Z plane. (B,C ) Electron micrographs of single 200nm sections from female WI-38 human fibroblasts. The inactive X chromosome (arrow) has an ultrastructure distinct from the chromocenters (arrowheads), adopting a looser, less compact organization. Adapted, with permission, from Rego et al. 2008, Journal of Cell Science.

for histone modifications characterizing heterochromatin(H3K9me2,H3K27me3,H3K9me3, H4K20me3) is less clear-cut, with some studies reporting no enrichment on the Xi at metaphase (Koina et al. 2009) and others reporting some enrichment of H3K27me3 at interphase (Mahadevaiah et al. 2009). Given that marsupials lack the XIST gene (Duret et al. 2006), differences in histone modification patterns on the

Xi might be expected, since Xist is required for the recruitment of many of the chromatin marks in mouse. The situation in monotremes appears to be even more complex with females having five pairs of XX chromosomes and males having five XY pairs. Although there does appear to be some degree of dosage compensation, no systematic analysis has been performed to determine the heterochromatic status of the

Nuclear Organization and Dosage Compensation

X chromosomes in females, and dosage compensation appears to be largely incomplete and variable between genes (Grutzner et al. 2004; Rens et al. 2004; Deakin et al. 2008). A more detailed view of the chromatin content of the eutherian inactive X chromosome is starting to be obtained by higher resolution epigenomic analyses, such as ChIP-chip and allele-specific ChIP (Brinkman et al. 2006; Valley et al. 2006; Valley and Willard 2006; Chadwick 2007; Marks et al. 2009; Mietton et al. 2009). ChIP-chip for macroH2A1 in mouse liver cells reveals an approximately 1.5-fold enrichment in females versus males. As this enrichment is distributed uniformly across the entire X chromosome, this implies that macroH2A1 may influence global chromatin structure, rather than directly inhibiting transcription at promoters of genes (Mietton et al. 2009). The distribution of H3K27me3 has also been examined on the inactivating X chromosome in differentiating mouse ES cells by ChIP-Seq (Marks et al. 2009). Allele-specific mapping of ChIP-seq tags showed that the H3K27me3 mark is deposited on the inactive X chromosome in female ES cells, particularly in gene-rich, active regions, which is in agreement with the results observed in human somatic cells by immunofluorescence (Chadwick 2007; Marks et al. 2009). DNA methylation of promoters of X-linked genes is another hallmark of XCI in eutherian somatic cells. Global microarray analyses involving methylated DNA immunoprecipitation (MeDIP) were used to assess the DNA methylation status of the Xi relative to the active X (Xa) in human primary cells. This study revealed that although CpG islands are hypermethylated on the Xi, the overall levels of methylation on the X chromosome are in fact lower in females compared to males, especially in gene-poor regions (Weber et al. 2005). Hellman and Chess (2007) analyzed the DNA methylation status of more than 1000 X-linked loci and found that on the active X, DNA methylation is concentrated in gene bodies, confirming previous studies (for review, see Heard et al. 1997). Although the function of methylation within transcribed genes is unknown, it is a general feature of active

genes in many organisms (for example, Zhang et al. 2006). The lack of gene-body DNA methylation on the inactive X chromosome at least partly explains its globally hypomethylated status, with only promoters of genes being hypermethylated. In summary, the inactive X chromosome contains many different epigenetic marks, with some variations in distribution between different mammals. The degree to which the different signatures on the Xi reflect differences in the higher-order structure and/or nuclear organization of the inactive X should become more clear with the advent of techniques such as chromosome conformation capture, which measures physical interaction within the chromatin fiber (Simonis et al. 2007; van Berkum and Dekker 2009). INITIATING X INACTIVATION: XIST RNA REGULATION AND FUNCTION

The evolution of the Xist gene is believed to have been a key event in the evolution of the stable, chromosome-wide X-inactivation process that exists in eutherian mammals. Xist RNA coating of the prospective inactive X chromosome appears to be the key trigger for the X-inactivation cascade. Deletion analyses have shown that Xist is essential for silencing and likely has a role at multiple levels in the process (Penny et al. 1996; Marahrens et al. 1997; Csankovszki et al. 1999; Wutz et al. 2002; Hoki et al. 2009). The regulation of Xist during development is a complex process, ensured by multiple long-range cis elements and trans-acting factors. Xist is in fact located in a region of the X chromosome known as the X-inactivation center (Xic) that ensures the appropriate initiation of X inactivation in female cells (reviewed in Barakat et al. 2010) (Fig. 3). The Xic contains sequence elements that ensure that Xist is only up-regulated in XX and not XY cells. The Xic also ensures that in XX cells, Xist up-regulation occurs on only one and not both X chromosomes. Recent insights into this process have revealed that the presence of two or more copies of certain X-linked loci can activate Xist. This may work at several levels. For example, a double dose of the X-linked Rnf12

J.C. Chow and E. Heard Xic mediated initiation of XCI: • Sensing and counting X chromosome number: trigger Xist up-regulation • Choosing one X for inactivation

Undifferentiated female ES cells

Monoallelic Xist up-regulation

Xic

Xa Xa Pluripotency factors Rnf12

Xpr

+

Xist Tsix Xite

Rnf12

Xpr

?+

+

Pluripotency factors

= trans interactions

Xist

Xist

Tsix Xite

Rnf12

Xpr

Tsix Xite

?–

–

= Xist RNA

Differentiated female ES cells

• Xist RNA accumulation and coating of the X in cis: exclusion of RNA pol II, silencing of repeats

Xa

Xi

• Appearance of macro H2A on Xi • Smchd1 recruitment to Xi? • DNA methylation of promoters of X-linked genes (hypomethylation of gene bodies)

• Histone modifications enriched on Xi (H3K9me2, H3K27me3, H4K20me1, etc) • Shift to late replication timing • Young LINE expression on Xi • Progressive spread of silencing onto all parts of the X (facilitated by LINEs?)

• Xist RNA silent compartment • Loss of active histone marks (H3K4me2/me3, H3Ac, H4Ac) • Gene silencing and relocation into Xist RNA compartment (via SATB1/2?) • Recruitment of PcG complexes

Spread and progressive locking in of heterochromatic state

Figure 3. Random X chromosome inactivation is a multi-stage process. During the initiation stage, develop-

mental factors as well as a complex interplay of elements within the X inactivation center (Xic) ensure that monoallelic Xist up-regulation occurs only when more than one X chromosome is present per diploid autosome set. Once up-regulated, Xist RNA coats the chromosome in cis and triggers chromosome-wide gene silencing. The spread of silencing occurs progressively with the acquisition of many different epigenetic changes that together serve to create an extremely stable, heterochromatic state.

gene, which encodes a ubiquitin ligase, can trigger Xist up-regulation (Jonkers et al. 2009). Trans interactions ( pairing) between Xics may also be involved in triggering the expression of Xist and in ensuring its monoallelic regulation in female cells (Bacher et al. 2006; Xu et al. 2006; Augui et al. 2007). These transient Xic pairing events may facilitate the coordination of reciprocal Xist expression patterns on

the future active and inactive X chromosomes during the initiation of X inactivation (Augui et al. 2007; Nicodemi and Prisco 2007; Xu et al. 2007). Thus, the initiation of monoallelic Xist expression in female cells involves a dynamic series of events at the Xic loci, integrating developmental triggers, dosage-dependent proteins factors, interactions in trans, and noncoding RNAs (Fig. 3).

Nuclear Organization and Dosage Compensation

Once Xist RNA has become monoallelically up-regulated (at around day 2 in differentiating female ES ells), it begins the process of chromosome-wide gene silencing. Xist RNA induces X inactivation during an early differentiation time window, and recent studies have implicated the SATB1 and SATB2 (Special AT-rich Binding Protein) proteins in enabling Xist function during this early differentiation time window and also in some cancer cells (Agrelo et al. 2009). The 17,000–19,000nt-long, spliced, poly-adenylated Xist transcript consists of multiple different functional domains (Wutz et al. 2002). The most conserved region of Xist is a set of tandem repeats at the 50 end of the gene, called the Arepeats, that are required for its gene silencing function (Wutz et al. 2002; Hoki et al. 2009). Inducible Xist transgenes deleted for A-repeats are unable to induce gene silencing, but can still coat the chromosome in cis and recruit polycomb group proteins, which are involved in the recruitment of repressive chromatin changes including H3K27me3 to the X chromosome (Wutz et al. 2002; Plath et al. 2003; Kohlmaier et al. 2004). Thus, other portions of Xist RNA are required for its capacity to coat the X chromosome in cis and to recruit polycomb and macroH2A (Wutz et al. 2002). The Xist A-repeat region also produces an independent 1.6-kb transcript, RepA, and although its function is not entirely clear, this transcript may play a role in regulating Xist itself (Zhao et al. 2008; Hoki et al. 2009). Although Xist RNA may be involved in directly or indirectly recruiting chromatin-modifying enzymes, including polycomb group proteins, so far there is no evidence that this accounts for its gene-silencing role. However, there is increasing evidence pointing to Xist RNA having an architectural role that may be important for the specific 3D-organization of the inactive X (Chaumeil et al. 2006; Clemson et al. 2006; Rego et al. 2008), which may impact both the initiation and maintenance of the inactive state. One of the earliest events following Xist up-regulation and coating of the future inactive X chromosome is the formation of a nuclear compartment that is depleted for RNA polymerase, transcription factors, and

euchromaticmarks(Chaumeiletal.2006).Based on the analysis of Xist and PcG mutant ES cell lines, gene silencing and the recruitment of polycomb group proteins can be functionally dissociated from the formation of the Xist silent nuclear compartment (Plath et al. 2003; Kohlmaier et al. 2004; Chaumeil et al. 2006; Schoeftner et al. 2006). The silent compartment initially encompassed by Xist RNA is in fact largely constituted of X-chromosome repeat sequences, rather than genes. Prior to their inactivation, X-linked genes actually reside outside or at the periphery of this Xist RNA compartment. As differentiation proceeds and X inactivation occurs, genes move into this Xist RNA domain as they become silenced (Chaumeil et al. 2006). The mechanisms underlying this relocalization of genes are currently unknown. The Xist A-repeat region, which is essential for gene silencing, may have a role since genes do not become relocated into the silent repetitive compartment in Xist A-repeat mutants. Furthermore, given the implication of the SATB1 and SATB2 factors in Xist RNA’s competence to induce gene silencing during early differentiation (Agrelo et al. 2009), and the fact that these DNA-binding proteins facilitate the organization of chromatin structure during T-cell development (Cai et al. 2006), they may similarly play a role in gene relocation into the silent Xist repetitive compartment during X inactivation. So far, no physical association between Xist A-repeats and SATB1/2 has been demonstrated, but their window of action and their known function in nuclear organization make them key candidates for a role in Xist-mediated gene silencing and relocation. Importantly, recent findings demonstrate that Xist RNA can actually induce gene silencing outside an early developmental context, in some cancer cells (Hall et al. 2002; Chow et al. 2007), and in adult progenitor stem cells (Savarese et al. 2006). Agrelo et al. (2009) show that this reacquired capacity for Xist-induced silencing in tumor cells is associated with the re-expression of the SATB1 protein. This finding reinforces the notion that the nuclear and epigenetic plasticity found in some cancer cells may be similar to that found during early embryogenesis.

J.C. Chow and E. Heard

SPREADING INACTIVATION ACROSS THE X CHROMOSOME

Early studies using inducible Xist genes in male ES cells demonstrated that Xist had a developmentally restricted function, limited to the first few days of differentiation (Wutz and Jaenisch 2000). X-inactivation was thus originally thought to be a rather concerted process, occurring in the space of just a few cell divisions. However, recent studies in mouse pre-implantation embryos (Patrat et al. 2009) and differentiating female ES cells (Lin et al. 2007; Chow et al. 2010) have revealed that inactivation of the X chromosome in its entirety can take much longer. Some genes are silenced very quickly (in a matter of days or even hours after Xist RNA accumulation), while others are silenced much later (days, or .1 week), well outside of the time window in which Xist RNA is thought to act (Lin et al. 2007; Patrat et al. 2009). This suggests either that Xist’s silencing function is not limited to early differentiation for some loci, or else that the changes induced by Xist are propagated more slowly into some regions compared to others. Such heterogeneity suggests that the kinetics of inactivation are controlled by regionspecific processes, presumably due to different sequence environments. In addition, several X-linked genes are known to escape the X-inactivation process either partly or fully (Yang et al. 2010). Some of these genes (e.g., Jarid1c/Kdm5c) have the capacity to escape from X inactivation, whatever their location on the X chromosome (Li and Carrel 2008), implying that they have specific features preventing them from being efficiently silenced. Importantly, all escapees tend to be located outside the Xist RNA compartment, suggesting that part of the mechanism enabling escape may be linked to their capacity to resist internalization into the silent, repetitive compartment. However, the exact sequence and/or chromatin signatures that underlie the capacity of some genes to escape and those that underlie the variability in X-inactivation kinetics between genes, are still not well understood. Identifying such genomic and/ or chromatin features has turned out to be a challenge since multiple actors are likely to be

at work and account for the diversity of X-inactivation profiles. Genes that can escape from X inactivation may be flanked by insulator elements that help form domains that prevent the spread of heterochromatinization and/or recruitment of a gene into the silent repetitive Xist compartment. CTCF-binding sites have been hypothesized to play such a role as they have been identified between inactivation and escape domains (Filippova et al. 2005). Sequences that facilitate the propagation of X inactivation have also been proposed to exist, termed “way-stations” or “booster elements” (Gartler and Riggs 1983). Lyon proposed that possible candidates for such booster elements could include non-LTR retrotransposon LINE1 repeats, as they are enriched on the X chromosome and their density correlates well with the spread of silencing into autosomal sequences in X: autosome translocations (Lyon 1998). Furthermore, LINE1 elements are depleted in the immediate vicinity of escape genes (Bailey et al. 2000; Carrel et al. 2006). More recently, it has been shown that LINEs may facilitate the propagation of X inactivation at at least two different levels. Truncated, ancient LINEs may participate in nucleating the heterochromatic repetitive compartment induced by Xist RNA early in X inactivation (Chow et al. 2010). On the other hand, younger, more active LINEs, which are actually transcribed from the inactive X during development, may participate in the local propagation of X inactivation into regions that would otherwise be prone to escape (Chow et al. 2010). One such region that has been studied in detail contains the Huwe1 gene, which has a full length LINE at its 30 end. Huwe1 displays very slow X-inactivation kinetics, initiating only after day 2 of differentiation (Patrat et al. 2009; Chow et al. 2010). The slow silencing of Huwe1 may be due to its proximity to an escapee, Jarid1c, which might render its surrounding region, including Huwe1, more resistant to X inactivation. Huwe1 is, however, eventually inactivated, possibly due to the presence of nearby fulllength LINE1 elements. Intriguingly, these full length LINEs are transcribed from the inactive X chromosome during the exact window of time that X inactivation is occurring and their

Nuclear Organization and Dosage Compensation

transcription is accompanied by the production of small RNAs (Chow et al. 2010) (Fig. 4). In the case of Huwe1, it has been proposed that transient LINE1 transcription may drive antisense transcription into the gene and this may help to induce its silencing. Antisense transcription has increasingly emerged as an important regulator of gene expression in a diverse number of ways (Su et al. 2010). Active LINE1 elements/ promoters may therefore represent a significant source of antisense transcription and play regulatory roles in the genome, particularly on the X chromosome, where they are enriched, at least in eutherian mammals. In summary, LINEs may participate in generating heterochromatin in more than one way, with young, full-length

LINEs acting as local boosters to facilitate silencing of certain regions of the X chromosome; for example, by driving the transient production of natural antisense transcripts on the X chromosome undergoing inactivation; while older, truncated LINEs may facilitate creation of the silent repetitive core of the X chromosome, into which genes are recruited as they are inactivated (Chaumeil et al. 2006; Chow et al. 2010). Although it is clear that Xist RNA is essential for the recruitment and propagation of chromatin modifications as well as the specific reorganization of the chromosome, how these changes result in later changes, such as the shift to late replication timing, macroH2A enrichment,

Day 2 differentiation

Undifferentiated ES cells (Active X)

Xist up-regulation triggers the formation of a transcriptionally silent repetitive compartment

Genes

Day 4-8 differentiation Repeats Full length active L1 elements Recruitment of silencing factors

Huwe1

Xist RNA

Heterochromatinization? Methylation?

Active L1 element

kdm5C/Jarid1C

Spread of silencing? Small RNA pathway? Spread?

Huwe1

Active L1 element

Some regions are more resistant to silencing and are subject to slower silencing kinetics (e.g., near escape genes?) Differentiated ES cells (Inactive X)

Boundary elements

Kdm5C/Jarid1C Full length L1 elements may help to spread silencing into these more resistant regions.

Figure 4. Diverse, region-specific elements contribute to the spread and stability of silencing and result in

the heterogeneous inactivation kinetics observed across the chromosome. The X-linked Huwe1 gene is Xinactivated relatively late during development. This resistance to silencing may be due to the influence of the nearby escapee domain containing the Jarid1C/Kdm5c gene. The presence of a full length LINE1 element 30 of Huwe1 may help to eventually spread silencing into the Huwe1 gene.

J.C. Chow and E. Heard

and DNA methylation at promoters, remains unclear. A recent new player in X inactivation that may be involved in some of the downstream changes is the Smchd1 protein (Blewitt et al. 2008). Smchd1 mutant female embryos display both placental and extra-embryonic defects and die prior to stage E13.5. This relatively late lethality is suggestive of a role in the maintenance, rather than the initiation of XCI. Indeed, promoter DNA hypomethylation on the Xi was detected in mutants, suggesting that Smchd1 may affect the DNA methylation deposition step (Blewitt et al. 2008). Smchd1 is an SMC-like protein with homology to components of cohesin and condensin complexes. Interestingly, condensins are implicated in the down-regulation of both X chromosomes in C. elegans hermaphrodites (Chuang et al. 1994), providing an exciting potential link between these two forms of dosage compensation.

and chromatin patterns found for different regions of the inactive X chromosome suggest that there may be more than one way of inducing heterochromatin during X inactivation. Furthermore, different mammals exploit different mechanisms to achieve chromosome-wide silencing, with variations in chromatin marks. The extent to which nuclear organization and spatial segregation of the X chromosomes represent a universal feature of dosage compensation strategies will remain an interesting question for the future. ACKNOWLEDGMENTS

Our work is supported by grants from the CNRS, FRM (Equipe FRM), ANR, and Canceropole (GepiG), as well as HEROIC (Highthroughput Epigenetic Regulatory Organization in Chromatin), an integrated project funded by the European Union, and the ERC.

CONCLUSIONS AND PERSPECTIVES

Although very different mechanisms have evolved in different organisms to ensure chromosomewide gene regulation during dosage compensation, a common theme is the necessity for recruitment of dosage compensation complexes across the length of the X chromosome in a concerted fashion. In flies and worms, specific recruitment or entry sites are beginning to be identified, and should help to unravel the exact mechanisms by which the dosage compensation complexes are recruited and spread in these organisms. However, chromatin structure, noncoding RNAs, and nuclear organization may also play important roles. Mammals present the unusual situation whereby two genetically identical X chromosomes have to be differentially treated within the same nucleus and nuclear organization may contribute to this process. Coating of one of the two X chromosomes by the non-coding Xist RNA triggers formation of a silent nuclear compartment, into which genes are recruited as they become inactivated, as well as the recruitment of repressive complexes. The cis-limited action of Xist RNA still remains mysterious, as do the proteins or sequences to which it binds. The diversity of X-inactivation kinetics

REFERENCES Agrelo R, Souabni A, Novatchkova M, Haslinger C, Leeb M, Komnenovic V, Kishimoto H, Gresh L, Kohwi-Shigematsu T, Kenner L, et al. 2009. SATB1 defines the developmental context for gene silencing by Xist in lymphoma and embryonic cells. Dev Cell 16: 507 –516. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507– 517. Alekseyenko AA, Peng S, Larschan E, Gorchakov AA, Lee OK, Kharchenko P, McGrath SD, Wang CI, Mardis ER, Park PJ, et al. 2008. A sequence motif within chromatin entry sites directs MSL establishment on the Drosophila X chromosome. Cell 134: 599 –609. Augui S, Filion GJ, Huart S, Nora E, Guggiari M, Maresca M, Stewart AF, Heard E. 2007. Sensing X chromosome pairs before X inactivation via a novel X-pairing region of the Xic. Science 318: 1632– 1636. Bacher CP, Guggiari M, Brors B, Augui S, Clerc P, Avner P, Eils R, Heard E. 2006. Transient colocalization of Xinactivation centres accompanies the initiation of X inactivation. Nat Cell Biol 8: 293–299. Bailey JA, Carrel L, Chakravarti A, Eichler EE. 2000. Molecular evidence for a relationship between LINE-1 elements and X chromosome inactivation: the Lyon repeat hypothesis. Proc Natl Acad Sci U S A 97: 6634– 6639. Barakat TS, Jonkers I, Monkhorst K, Gribnau J. 2010. X-changing information on X inactivation. Exp Cell Res 316: 679–687. Barr ML, Bertram EG. 1949. A morphological distinction between neurones of the male and female, and the behaviour of the nucleolar satellite during accelerated nucleoprotein synthesis. Nature 163: 676.

Nuclear Organization and Dosage Compensation Blewitt ME, Gendrel AV, Pang Z, Sparrow DB, Whitelaw N, Craig JM, Apedaile A, Hilton DJ, Dunwoodie SL, Brockdorff N, et al. 2008. SmcHD1, containing a structuralmaintenance-of-chromosomes hinge domain, has a critical role in X inactivation. Nat Genet 40: 663 –669. Bone JR, Lavender J, Richman R, Palmer MJ, Turner BM, Kuroda MI. 1994. Acetylated histone H4 on the male X chromosome is associated with dosage compensation in Drosophila. Genes Dev 8: 96–104. Brinkman AB, Roelofsen T, Pennings SW, Martens JH, Jenuwein T, Stunnenberg HG. 2006. Histone modification patterns associated with the human X chromosome. EMBO Rep 7: 628 –634. Cai S, Lee CC, Kohwi-Shigematsu T. 2006. SATB1 packages densely looped, transcriptionally active chromatin for coordinated expression of cytokine genes. Nat Genet 38: 1278–1288. Carrel L, Park C, Tyekucheva S, Dunn J, Chiaromonte F, Makova KD. 2006. Genomic environment predicts expression patterns on the human inactive X chromosome. PLoS Genet 2: e151. Chadwick BP. 2007. Variation in Xi chromatin organization and correlation of the H3K27me3 chromatin territories to transcribed sequences by microarray analysis. Chromosoma 116: 147 –157. Chadwick BP, Willard HF. 2004. Multiple spatially distinct types of facultative heterochromatin on the human inactive X chromosome. Proc Natl Acad Sci U S A 101: 17450–17455. Chaumeil J, Le Baccon P, Wutz A, Heard E. 2006. A novel role for Xist RNA in the formation of a repressive nuclear compartment into which genes are recruited when silenced. Genes Dev 20: 2223–2237. Chow JC, Ciaudo C, Fazzari MJ, Mise N, Servant N, Glass JL, Attreed M, Avner P, Wutz A, Barillot E, et al. 2010. LINE-1 activity in facultative heterochromatin formation during X chromosome inactivation. Cell 141: 956 –969. Chow JC, Hall LL, Baldry SE, Thorogood NP, Lawrence JB, Brown CJ. 2007. Inducible XIST-dependent X-chromosome inactivation in human somatic cells is reversible. Proc Natl Acad Sci U S A 104: 10104– 10109. Chuang PT, Albertson DG, Meyer BJ. 1994. DPY-27:a chromosome condensation protein homolog that regulates C. elegans dosage compensation through association with the X chromosome. Cell 79: 459– 474. Chuang PT, Lieb JD, Meyer BJ. 1996. Sex-specific assembly of a dosage compensation complex on the nematode X chromosome. Science 274: 1736–1739. Clemson CM, Hall LL, Byron M, McNeil J, Lawrence JB. 2006. The X chromosome is organized into a gene-rich outer rim and an internal core containing silenced nongenic sequences. Proc Natl Acad Sci U S A 103: 7688– 7693. Comings DE. 1968. The rationale for an ordered arrangement of chromatin in the interphase nucleus. Am J Hum Genet 20: 440– 460. Csankovszki G, Collette K, Spahl K, Carey J, Snyder M, Petty E, Patel U, Tabuchi T, Liu H, McLeod I, et al. 2009. Three distinct condensin complexes control C. elegans chromosome dynamics. Curr Biol 19: 9 –19.

Csankovszki G, Panning B, Bates B, Pehrson JR, Jaenisch R. 1999. Conditional deletion of Xist disrupts histone macroH2A localization but not maintenance of X inactivation. Nat Genet 22: 323– 324. de Wit E, van Steensel B. 2009. Chromatin domains in higher eukaryotes: insights from genome-wide mapping studies. Chromosoma 118: 25–36. Deakin JE, Hore TA, Koina E, Marshall Graves JA. 2008. The status of dosage compensation in the multiple X chromosomes of the platypus. PLoS Genet 4: e1000140. Duret L, Chureau C, Samain S, Weissenbach J, Avner P. 2006. The Xist RNA gene evolved in eutherians by pseudogenization of a protein-coding gene. Science 312: 1653–1655. Filippova GN, Cheng MK, Moore JM, Truong JP, Hu YJ, Nguyen DK, Tsuchiya KD, Disteche CM. 2005. Boundaries between chromosomal domains of X inactivation and escape bind CTCF and lack CpG methylation during early development. Dev Cell 8: 31– 42. Gartler SM, Riggs AD. 1983. Mammalian X-chromosome inactivation. Annu Rev Genet 17: 155– 190. Gelbart ME, Kuroda MI. 2009. Drosophila dosage compensation: a complex voyage to the X chromosome. Development 136: 1399–1410. Grimaud C, Becker PB. 2009. The dosage compensation complex shapes the conformation of the X chromosome in Drosophila. Genes Dev 23: 2490–2495. Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O’Brien PC, Jones RC, Ferguson-Smith MA, Marshall Graves JA. 2004. In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z and mammal X chromosomes. Nature 432: 913 –917. Gupta V, Parisi M, Sturgill D, Nuttall R, Doctolero M, Dudko OK, Malley JD, Eastman PS, Oliver B. 2006. Global analysis of X-chromosome dosage compensation. J Biol 5: 3. Hall LL, Byron M, Sakai K, Carrel L, Willard HF, Lawrence JB. 2002. An ectopic human XIST gene can induce chromosome inactivation in postdifferentiation human HT-1080 cells. Proc Natl Acad Sci U S A 99: 8677–8682. Hallacli E, Akhtar A. 2009. X chromosomal regulation in flies: when less is more. Chromosome Res 17: 603–619. Heard E, Clerc P, Avner P. 1997. X-chromosome inactivation in mammals. Annu Rev Genet 31: 571– 610. Heard E, Disteche CM. 2006. Dosage compensation in mammals: fine-tuning the expression of the X chromosome. Genes Dev 20: 1848– 1867. Hellman A, Chess A. 2007. Gene body-specific methylation on the active X chromosome. Science 315: 1141–1143. Hoki Y, Kimura N, Kanbayashi M, Amakawa Y, Ohhata T, Sasaki H, Sado T. 2009. A proximal conserved repeat in the Xist gene is essential as a genomic element for X-inactivation in mouse. Development 136: 139 –146. Jans J, Gladden JM, Ralston EJ, Pickle CS, Michel AH, Pferdehirt RR, Eisen MB, Meyer BJ. 2009. A condensin-like dosage compensation complex acts at a distance to control expression throughout the genome. Genes Dev 23: 602 –618. Jonkers I, Barakat TS, Achame EM, Monkhorst K, Kenter A, Rentmeester E, Grosveld F, Grootegoed JA, Gribnau J. 2009. RNF12 is an X-Encoded dose-dependent activator of X chromosome inactivation. Cell 139: 999– 1011.

J.C. Chow and E. Heard Kelley RL, Meller VH, Gordadze PR, Roman G, Davis RL, Kuroda MI. 1999. Epigenetic spreading of the Drosophila dosage compensation complex from roX RNA genes into flanking chromatin. Cell 98: 513– 522. Kohlmaier A, Savarese F, Lachner M, Martens J, Jenuwein T, Wutz A. 2004. A chromosomal memory triggered by Xist regulates histone methylation in X inactivation. PLoS Biol 2: E171. Koina E, Chaumeil J, Greaves IK, Tremethick DJ, Graves JA. 2009. Specific patterns of histone marks accompany X chromosome inactivation in a marsupial. Chromosome Res 17: 115– 126. Li N, Carrel L. 2008. Escape from X chromosome inactivation is an intrinsic property of the Jarid1c locus. Proc Natl Acad Sci U S A 105: 17055– 17060. Lieb JD, Albrecht MR, Chuang PT, Meyer BJ. 1998. MIX-1: an essential component of the C. elegans mitotic machinery executes X chromosome dosage compensation. Cell 92: 265 –277. Lieb JD, Capowski EE, Meneely P, Meyer BJ. 1996. DPY-26, a link between dosage compensation and meiotic chromosome segregation in the nematode. Science 274: 1732– 1736. Lin H, Gupta V, Vermilyea MD, Falciani F, Lee JT, O’Neill LP, Turner BM. 2007. Dosage compensation in the mouse balances up-regulation and silencing of X-linked genes. PLoS Biol 5: e326. Lyon MF. 1998. X-chromosome inactivation: a repeat hypothesis. Cytogenet Cell Genet 80: 133– 137. Mahadevaiah SK, Royo H, VandeBerg JL, McCarrey JR, Mackay S, Turner JM. 2009. Key features of the X inactivation process are conserved between marsupials and eutherians. Curr Biol 19: 1478–1484. Marahrens Y, Panning B, Dausman J, Strauss W, Jaenisch R. 1997. Xist-deficient mice are defective in dosage compensation but not spermatogenesis. Genes Dev 11: 156 –166. Marks H, Chow JC, Denissov S, Francoijs KJ, Brockdorff N, Heard E, Stunnenberg HG. 2009. High-resolution analysis of epigenetic changes associated with X inactivation. Genome Res 19: 1361– 1373. Mendjan S, Taipale M, Kind J, Holz H, Gebhardt P, Schelder M, Vermeulen M, Buscaino A, Duncan K, Mueller J, et al. 2006. Nuclear pore components are involved in the transcriptional regulation of dosage compensation in Drosophila. Mol Cell 21: 811– 823. Mietton F, Sengupta AK, Molla A, Picchi G, Barral S, Heliot L, Grange T, Wutz A, Dimitrov S. 2009. Weak but uniform enrichment of the histone variant macroH2A1 along the inactive X chromosome. Mol Cell Biol 29: 150–156. Misteli T. 2007. Beyond the sequence: cellular organization of genome function. Cell 128: 787 –800. Nguyen DK, Disteche CM. 2006. Dosage compensation of the active X chromosome in mammals. Nat Genet 38: 47–53. Nicodemi M, Prisco A. 2007. Symmetry-breaking model for X-chromosome inactivation. Phys Rev Lett 98: 108104. Patrat C, Okamoto I, Diabangouaya P, Vialon V, Le Baccon P, Chow J, Heard E. 2009. Dynamic changes in paternal Xchromosome activity during imprinted X-chromosome inactivation in mice. Proc Natl Acad Sci U S A 106: 5198– 5203.

Penny GD, Kay GF, Sheardown SA, Rastan S, Brockdorff N. 1996. Requirement for Xist in X chromosome inactivation. Nature 379: 131 –137. Peters AH, Mermoud JE, O’Carroll D, Pagani M, Schweizer D, Brockdorff N, Jenuwein T. 2002. Histone H3 lysine 9 methylation is an epigenetic imprint of facultative heterochromatin. Nat Genet 30: 77–80. Plath K, Fang J, Mlynarczyk-Evans SK, Cao R, Worringer KA, Wang H, de la Cruz CC, Otte AP, Panning B, Zhang Y. 2003. Role of histone H3 lysine 27 methylation in X inactivation. Science 300: 131– 135. Rego A, Sinclair PB, Tao W, Kireev I, Belmont AS. 2008. The facultative heterochromatin of the inactive X chromosome has a distinctive condensed ultrastructure. J Cell Sci 121: 1119– 1127. Rens W, Grutzner F, O’Brien PC, Fairclough H, Graves JA, Ferguson-Smith MA. 2004. Resolution and evolution of the duck-billed platypus karyotype with an X1Y1X2Y2X3Y3X4Y4X5Y5 male sex chromosome constitution. Proc Natl Acad Sci U S A 101: 16257–16261. Savarese F, Flahndorfer K, Jaenisch R, Busslinger M, Wutz A. 2006. Hematopoietic precursor cells transiently reestablish permissiveness for X inactivation. Mol Cell Biol 26: 7167–7177. Schoeftner S, Sengupta AK, Kubicek S, Mechtler K, Spahn L, Koseki H, Jenuwein T, Wutz A. 2006. Recruitment of PRC1 function at the initiation of X inactivation independent of PRC2 and silencing. EMBO J 25: 3110–3122. Simonis M, Kooren J, de Laat W. 2007. An evaluation of 3C-based methods to capture DNA interactions. Nat Methods 4: 895– 901. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72: 573–608. Straub T, Grimaud C, Gilfillan GD, Mitterweger A, Becker PB. 2008. The chromosomal high-affinity binding sites for the Drosophila dosage compensation complex. PLoS Genet 4: e1000302. Su WY, Xiong H, Fang JY. 2010. Natural antisense transcripts regulate gene expression in an epigenetic manner. Biochem Biophys Res Commun 396: 177–181. Taddei A, Hediger F, Neumann FR, Gasser SM. 2004. The function of nuclear architecture: a genetic approach. Annu Rev Genet 38: 305 –345. Tsai CJ, Mets DG, Albrecht MR, Nix P, Chan A, Meyer BJ. 2008. Meiotic crossover number and distribution are regulated by a dosage compensation protein that resembles a condensin subunit. Genes Dev 22: 194– 211. Valley CM, Pertz LM, Balakumaran BS, Willard HF. 2006. Chromosome-wide, allele-specific analysis of the histone code on the human X chromosome. Hum Mol Genet 15: 2335–2347. Valley CM, Willard HF. 2006. Genomic and epigenomic approaches to the study of X chromosome inactivation. Curr Opin Genet Dev 16: 240– 245. van Berkum NL, Dekker J. 2009. Determining spatial chromatin organization of large genomic regions using 5C technology. Methods Mol Biol 567: 189–213. Vaquerizas JM, Suyama R, Kind J, Miura K, Luscombe NM, Akhtar A. 2010. Nuclear pore proteins nup153 and megator define transcriptionally active regions in the Drosophila genome. PLoS Genet 6: e1000846.

Nuclear Organization and Dosage Compensation Vicoso B, Bachtrog D. 2009. Progress and prospects toward our understanding of the evolution of dosage compensation. Chromosome Res 17: 585 –602. Weber M, Davies JJ, Wittig D, Oakeley EJ, Haase M, Lam WL, Schubeler D. 2005. Chromosome-wide and promoter-specific analyses identify sites of differential DNA methylation in normal and transformed human cells. Nat Genet 37: 853– 862. Wutz A, Jaenisch R. 2000. A shift from reversible to irreversible X inactivation is triggered during ES cell differentiation. Mol Cell 5: 695 –705. Wutz A, Rasmussen TP, Jaenisch R. 2002. Chromosomal silencing and localization are mediated by different domains of Xist RNA. Nat Genet 30: 167 –174. Xu N, Donohoe ME, Silva SS, Lee JT. 2007. Evidence that homologous X-chromosome pairing requires transcription and Ctcf protein. Nat Genet 39: 1390– 1396.

Xu N, Tsai CL, Lee JT. 2006. Transient homologous chromosome pairing marks the onset of X inactivation. Science 311: 1149– 1152. Yang F, Babak T, Shendure J, Disteche CM. 2010. Global survey of escape from X inactivation by RNA-sequencing in mouse. Genome Res 20: 614 –622. Zhang LF, Huynh KD, Lee JT. 2007. Perinucleolar targeting of the inactive X during S phase: evidence for a role in the maintenance of silencing. Cell 129: 693– 706. Zhang X, Yazaki J, Sundaresan A, Cokus S, Chan SW, Chen H, Henderson IR, Shinn P, Pellegrini M, Jacobsen SE, et al. 2006. Genome-wide high-resolution mapping and functional analysis of DNA methylation in arabidopsis. Cell 126: 1189– 1201. Zhao J, Sun BK, Erwin JA, Song JJ, Lee JT. 2008. Polycomb proteins targeted by a short repeat RNA to the mouse X chromosome. Science 322: 750– 756.

The Budding Yeast Nucleus Angela Taddei1,2, Heiko Schober3, and Susan M. Gasser3 1

UMR 218, Centre National de la Recherche Scientifique, 26 rue d’Ulm, 75231 Paris Cedex 05, France

2

Institut Curie-Section de Recherche, 26 rue d’Ulm, 75231 Paris Cedex 05, France

3

Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, CH-4058 Basel, Switzerland

Correspondence: [email protected]

The budding yeast nucleus, like those of other eukaryotic species, is highly organized with respect to both chromosomal sequences and enzymatic activities. At the nuclear periphery interactions of nuclear pores with chromatin, mRNA, and transport factors promote efficient gene expression, whereas centromeres, telomeres, and silent chromatin are clustered and anchored away from pores. Internal nuclear organization appears to be function-dependent, reflecting localized sites for tRNA transcription, rDNA transcription, ribosome assembly, and DNA repair. Recent advances have identified new proteins involved in the positioning of chromatin and have allowed testing of the functional role of higher-order chromatin organization. The unequal distribution of silent information regulatory factors and histone modifying enzymes, which arises in part from the juxtaposition of telomeric repeats, has been shown to influence chromatin-mediated transcriptional repression. Other localization events suppress unwanted recombination. These findings highlight the contribution budding yeast genetics and cytology have made to dissecting the functional role of nuclear structure.

ith 16 chromosomes and a closed mitosis, the budding yeast nucleus provides an ideal model system for understanding structure– function relationships within one of the cell’s most complex compartments. Eukaryotic chromatin has an intrinsic higher-order structure based on the folding of the nucleosomal fiber. Yet beyond the intrinsic folding of the 10- and 30-nm fibers, sequence-determined subchromosomal domains and entire chromosomes assume non-random positions in the nucleus. Here we discuss a few general principles of nuclear organization. The first is an order imposed by enzymatic function. Examples include replication foci, splicing centers,

W

or sites of DNA repair by homologous recombination (Lamond and Spector 2003). The second is a global territorial organization that stems from a nonoverlapping positioning of entire chromosomes within the nucleus (Cremer et al. 2006). The third principle reflects binding characteristics of particular sequence elements, such as simple repeats, and the factors they recruit to assemble unique higher-order levels of chromatin folding (Gasser et al. 2004). The most straightforward way to show the functional importance of nuclear and chromosomal structure is to identify the proteins and sequences involved and to mutate those elements. Any resulting alteration in nuclear

Editors: David L. Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000612 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000612

A. Taddei, H. Schober, and S.M. Gasser

function might stem from the change in organization. These loss-of-function approaches then need to be coupled with gain-of-function assays that restore or promote a particular function or spatial organization. For these manipulations budding yeast is an ideal organism. Yeast has extremely efficient homologous recombination machinery that allows for targeted gene disruption, mutagenesis, chromosomal modification, and in vivo tagging of both DNA and proteins for live imaging. These techniques can be coupled with powerful biochemical and molecular genetic techniques, many of which give genome-wide read-outs. Such approaches have provided basic insights into the relationship of nuclear structure and function (e.g., Andrulis et al. 1998; Taddei et al. 2004; Taddei et al. 2006). Unfortunately, some elements of nuclear organization present in multicellular organisms are missing in yeast. Having a closed mitosis means that disassembly and reassembly of the yeast nuclear envelope does not occur during the cell cycle, although yeast has less pronounced changes in its nuclear membrane during cell division. Coupled with the yeast cell’s closed mitosis is a lack of nuclear intermediate filament proteins or lamins, which form a rigid network underlying the inner nuclear membrane (INM). The nuclearlamina stabilizes nuclear shape and help anchor interphase chromatin (Gruenbaum et al. 2005). Yeast cells also lack many of the well-characterized subnuclear “bodies” with the exception of a nucleolar domain that functions in snoRNA maturation (Verheggen et al. 2001) and which appears to be equivalent to Cajal bodies (Gall, 2000). Despite lacking detectable splicing compartments (Misteli, 2000) or PML ( promyelocytic leukemia) (Zhong et al. 2000) bodies, yeast supports many of the activities coordinated by these compartments, and recent evidence suggests that the yeast nuclear pore provides a platform for events related to gene expression and double strand break processing (Akhtar and Gasser 2007; Taddei 2007; Nagai et al. 2008). Other aspects of subnuclear organization, such as the distribution of chromatin within the nucleus, are conserved from yeast to man.

NUCLEAR ORGANIZATION Nuclear Envelope Associated Proteins and the Nuclear Pore Complex

The nucleus is delimited by a double membrane called the nuclear envelope (NE) that is contiguous with the endoplasmic reticulum. The NE separates chromatin from the cytoplasm and provides anchorage for various nuclear and chromosomal structures, including the spindle pole body, and separate clusters of centromeres and telomeres. Trafficking between the nucleoplasm and the cytoplasm occurs through approximately 200 nuclear pore complexes (NPCs), which allow the free diffusion of small molecules, whereas regulating the transport of macromolecules. NPCs also provide a platform for mRNA transcription and quality control, as well as its export. These proteinaceous assemblies of approximately 50 MDa contain 456 nucleoporins of 30 different types (D’Angelo and Hetzer 2008). Each pore is a doughnut-shaped structure with eightfold symmetry around a central channel. Flexible protein filaments emanate from the core into both the cytoplasm and nucleoplasm, providing binding sites for both transport proteins and chromatin. A detailed map for the relative position of each nucleoporin was calculated based on multiple molecular, biochemical, and structural data revealing a strongly modular structure (Alber et al. 2007). Beside nucleoporins, many yeast proteins were shown by imaging techniques to be associated with the nuclear envelope (Huh et al. 2003). Of particular interest are the integral proteins of the inner nuclear membrane (INM) (Lusk et al. 2007) including: Doa10, a RING domain containing protein that targets nuclear proteins for degradation, Mps3, a member of the SUN (Sad1, UNC-84) family that is a shared component of the INM and of the spindle pole body (SPB; Jaspersen et al. 2002), and Helix – extension – helix-1 and -2 (Src1 or Heh1 and Heh2) (King et al. 2006), which are orthologs of the mammalian lamin associated protein MAN1. These proteins interact with the nuclear lamina in mammals, as well as with chromatin through a small protein called BAF1. Although yeast lack the lamins, the roles of SUN-domain

The Budding Yeast Nucleus

and Man1 family proteins in chromosome anchorage seem to be conserved (Grund et al. 2008). In addition, an INM-associated protein called Esc1 (Enhancer of silent chromatin 1) shares features with lamina components: Esc1 binds but does not span the inner nuclear membrane, and anchors silent chromatin through a component of the silencer complex, Sir4 (Taddei et al. 2004). Intriguingly, overexpression of Esc1 induces INM expansion (Hattier et al. 2007). Given their intimate proximity, it is not surprising that functional cross-talk among INM proteins such as Esc1 and the nuclear pore basket proteins Nup60, Mlp1 and Mlp2 (Therizols et al. 2006; Lewis et al. 2007; Palancade et al. 2007) has been detected. Chromosome Folding

Observations of chromosome folding were originally recorded by one of the founders of the field of cytology, Carl Rabl (Rabl 1885), who described the fold-back conformation of anaphase chromosomes in spotted salamander larvae. In the so-called Rabl conformation a chromosome folds back at its centromere, such that telomeres are juxtaposed (Fig. 1). Today we know that this conformation results from the attachment of microtubules to kinetochores, which assemble at centromeric chromatin, and which lead the movement of chromosomes as chromosomes are pulled by microtubules into daughter cells. The telomeres, on the other hand, lag passively behind. Evidence that yeast chromosomes assume a Rabl conformation came initially from the work of Loidl and coworkers, who used fluorescent in situ hybridization (FISH) to identify chromosomal landmarks such as centromeres and telomeres (Jin et al. 2000). Using chromosome conformation capture (3C) to model chromosome conformation, the Kleckner lab predicted that yeast Chr III folds as a contorted ring, with a strong bend near the centromere and the telomeres in close proximity to each other (Dekker et al. 2002). Gasser and co-workers confirmed and extended these observations showing that not only Chr III, but other small metacentric chromosomes, such as Chr VI,

form small loops in vivo with closely juxtaposed right and left telomeres. In contrast, telomeres on chromosome arms of vastly different lengths did not interact (Bystricky et al. 2005; Schober et al. 2008). Importantly, these studies showed that the Rabl configuration is not unique to anaphase in yeast, but persists throughout the cell cycle. Using a genome-wide conformation capture approach, in which spatial chromosome interactions can be mapped, O’Sullivan and co-workers have recently shown that yeast chromosome placement is not random, i.e., some chromosomes were found never to interact, whereas others showed favored interactions (Rodley et al. 2009). Furthermore, this method detected intra-chromosomal loops, much like those described by microscopic analysis for chromosomes 3 and 6 (Bystricky et al. 2005; Schober et al. 2008). Consistently, a new imaging and data analysis approach revealed a strong confinement of several loci into “gene territories” reflecting the architectural constraints imposed on chromosomes by nuclear structures (i.e., centromere attachments to the SPB and telomere anchoring to the NE). Importantly, this organization can be significantly remodeled upon transcriptional activation (Berger et al. 2008). Chromatin Dynamics

Chromatin in living cells is subjected to constant motion, which can be best described as a constrained random walk (Marshall et al. 1997) (Fig. 2). Rapid time-lapse imaging led to the distinction of at least two types of motion in yeast: small random movements (,0.2 mm within 1.5 s) that occur constantly, as well as larger, more directional movements (i.e., .0.5 mm in a 10.5 s interval) (Heun et al. 2001). The smaller movements are observed for internal loci as well as for peripheral silent domains, although the active loci in the nuclear lumen make large movements more frequently (i.e., a large movement is .0.5mm within 10.5 s) (Heun et al. 2001). Intriguingly, changes in cellular energy levels because of depletion of glucose or the addition of protonophores to deplete membrane potential, abolish large movements

A. Taddei, H. Schober, and S.M. Gasser

A

B Telophase 23

Nucleolus

Interphase TEL

CEN

TEL

C a

Telomeres + HM

b

SBP

c TelR, TelL

SBP

Figure 1. Chromosomes have distinct orientation within the yeast interphase nucleus, which results from their

Rabl configuration at anaphase: centromeres cluster and lead the way into daughter nuclei, with telomeres following behind. Telomeres of equal-length chromosome arms tend to cluster together in yeast interphase nuclei. (A) In 1885, Carl Rabl drew the folded conformation of anaphase chromosomes in spotted salamander larvae, in which anaphase chromosomes fold back on themselves because of attachment of the centromere to microtubules through the kinetochore. Centromeres lead the way as chromosomes are actively pulled into daughter cells. (B) The Rabl organization persists in the interphase yeast nucleus. (C ) Confocal fluorescence images show (a) the clustering of yeast centromeres (green by FISH) near the spindle pole body (SPB, red) or in the larger image the nucleolus (red) opposite the SPB (anti-Spc42, white); (b) yeast telomeric foci are labeled with anti-Sir4 protein and FISH for the silent HM loci in yeast; (c) visualization of the centromere and right and left telomeres of Chr VI through Tel6R-CFP-lacO, Tel6L-YFP-TetR, and centromere staining (white).

(Marshall et al. 1997; Heun et al. 2001). This suggests that interphase chromatin movement at least responds to ATP levels. Recent work suggests that these most likely stem from the action of chromatin remodelers, rather than RNA or DNA polymerases. Indeed, highly transcribed galactose-inducible genes that become tethered at nuclear pores show less mobility, despite their high level of transcription (Taddei et al. 2006; Cabal et al. 2006). Moreover, chromatin movement decreases, rather than increases, as cells enter S phase and DNA polymerases are active (Heun et al. 2001). Given that the mobility of a genomic locus is constrained by its continuity

with the rest of the chromosome (Gartenberg et al. 2004), it is likely that the S-phase associated drop in mobility reflects the association of DNA in replication factories (Kitamura et al. 2006). DNA BASED COMPARTMENTS: NUCLEOLUS, TELOMERES, tRNA The Nucleolus

The most evident subnuclear compartment is the nucleolus, a crescent-shaped structure abutting the nuclear envelope and occupying roughly one third of the nucleus opposite the spindle pole body (Yang et al. 1989; Bystricky

The Budding Yeast Nucleus Active loci, mid-chromosome XIV

Induced HXK1 gene at pore

Figure 2. Rapid time-lapse imaging of GFP-lacI tagged chromosomal loci in yeast cells bearing GFP-Nup49,

allows 3D tracking of an internal locus (Gartenberg et al. 2004). 3D stacks are taken at 1.5 s intervals and the spatial rendering and alignment is performed with Imaris (Bitplane, Zu¨rich). Tracks show movement over 7.5 min (300 stacks). To the left is a region of Chr IVX that shows low level transcription upon activation by glucose. To the right is shown the movement of the induced HXK1, a subtelomeric gene that is strongly induced in the absence of glucose (Taddei et al. 2006). Reductions in radius of constraint and diffusion coefficients are monitored for the pore-associated gene.

et al. 2005). This subcompartment is the site of RNA pol I-mediated rDNA transcription and ribosome subunit assembly and can be seen as a factory dedicated to ribosome biogenesis. Its morphology is strongly influenced by the cell growth rate, probably as a result of adapting the rate of ribosome production to the needs of the cell (Oakes et al. 1993; Powers and Walter, 1999). The nucleolus is a prime example of a functional organization principle, for it compartmentalizes different steps of ribosome biogenesis although the nucleolar machineries are in permanent exchange with the nucleoplasm and other nuclear bodies. The nucleolus further generates a domain of retention/sequestration of molecules that are normally active only outside the nucleolus (Sirri et al. 2008). In budding yeast, rRNA is encoded in 100– 200 tandem repeats. Each repeat unit is 9.1 kb in size and yields a 35S precursor rRNA, transcribed by RNA polymerase I and a 5S rRNA, transcribed by RNA polymerase III. The 5S unit is surrounded by two intergenic spacers, IGS1 and IGS2 (Fig. 3). Within this spacer there is a so-called polar replication fork barrier (RFB), a recombination enhancer (RE), a RNA

polymerase I transcription initiation region (TIR) and an origin of replication (ARS, from autonomously replicating sequence). This extended array of tandem repeats serves as an ideal template for homologous recombination, which is highly efficient in budding yeast. However, suppression of this recombination and the ensuing stability of the yeast rDNA array, are absolutely critical for cell growth and survival. By stabilizing the rDNA repeats the cell avoids or delays replicative senescence (Sinclair and Guarente 1997). The mechanisms that suppress recombination within the rDNA array involve both local nucleosomal organization mediated by Sir2 (Gottlieb and Esposito 1989; Bryk et al. 1997; Fritze et al. 1997; Smith and Boeke 1997) and long-range chromatin structure that involves tethering at the nuclear envelope. tRNA genes

Although the 274 Pol III-transcribed tRNA genes are scattered throughout the yeast genome, many of these are clustered close to the nucleolus (Thompson et al. 2003; Wang et al.

A. Taddei, H. Schober, and S.M. Gasser 9.1 kb 35S

35S

RE

35S

5S

35S

ARS

RFB IGS1

TIR IGS2

Figure 3. Organization of the rDNA repeats. The 9.1 kb unit encodes the 35S precursor for all the 25S, 18S, and

5.8S rRNA and the 5S rRNA. The 35S is transcribed by RNA polI whereas the 5S is transcribed by RNA pol III. The 9.1 kb unit is repeated up to 200 times on chr XII. Other abbreviations are as follows: IGS, intergenic spacer; RE recombination enhancer; RFB, Replication fork block, TIR, PolI transcription initiation region, open circle—origin of DNA replication.

2005). This association could create an environment that fosters the coregulation of transcription by RNA pol III and RNA pol I, given that the other major RNA pol III transcript is the 5S rRNA, which is part of the rDNA repeat (Fig. 3). Intriguingly, some tRNA genes remain associated with the nucleolus throughout the cell cycle, although they are not present on Chr XII, which contains the rDNA. This unusual positioning depends both on microtubules and on the action of condensin (Haeusler et al. 2008). Given the high number of RNA pol III-transcribed genes, their positioning is likely to affect the spatial organization of a large portion of the genome. Intriguingly, RNA pol II genes in the vicinity of tRNA genes become silenced through a phenomenon known as tRNA-gene mediated gene silencing (tgm) (Wang et al. 2005). Consistent with a link between nucleolar tethering and tRNAassociated silencing, it was shown that conditional ablation of condensin subunits leads to a coordinate loss of both nucleolar tRNA clustering and tgm (Haeusler et al. 2008). We note, however, that not all active tRNA genes relocate to the nucleolus. It remains to be determined what characterizes those that do show nucleolar association. Telomere Foci: Assemblies of Repetitive DNA and Silencing Factors

Another type of subnuclear compartment stems from the clustering of the 32 yeast telomeres

into three to eight foci, which are mainly found at the nuclear periphery. These foci are unfavorable for RNA polII-driven transcription. The budding yeast telomeric repeat consist of 250 – 300 base pairs of irregular tandem repeats (called TG1-3) (Shampay et al. 1984). A crucial feature of telomeric DNA is the 30 overhang of the G-rich strand, which is 10 – 15bp in length (Larrivee et al. 2004). Toward the end of S phase after completion of replication, an extended G-rich overhang is generated, providing a template for the action of telomerase, a conserved ribonucleoprotein complex with reverse transcriptase activity (reviewed in Zakian 1996; Hug and Lingner 2006). The protein subunits of this complex were identified in a screen for “ever shorter telomeres” and are called Est1, Est2, and Est3 (Lundblad and Szostak 1989; Lendvay et al. 1996), whereas the RNA moiety that templates the extension of TG repeats is called Tlc1 (Singer and Gottschling 1994). Tlc1 and Est2 form the catalytic core of telomerase, whereas Est1 and Est3 are auxiliary subunits that are dispensable for in vitro activity (Lingner et al. 1997). The end-binding factor yKu interacts with telomerase and helps recruit it to telomeres (Stellwagen et al. 2003), as does the ssDNA binding factor Cdc13 (reviewed in Fisher et al. 2004). Unlike this association with ssDNA, the Repressor Activator protein 1 binds the doublestranded telomeric repeat (Rap1) (Shore and Nasmyth 1987). Rap1 is a conserved factor with a double Myb-like domain that mediates

The Budding Yeast Nucleus

high affinity binding to its consensus within the TG1-3 repeat. The Rap1 carboxyl terminus is a binding site for silencing factors Sir3 and Sir4 (Moretti et al. 1994; Marcand et al. 1997; Wotton and Shore 1997), which form a stochiometric complex with the NAD-dependent histone deacetylase Sir2, to mediate transcriptional repression in subtelomeric zones (Aparicio et al. 1991; Martino et al. 2009). The same site binds Rif1, which together with Rif2, antagonizes Sir4 interaction, and feeds back to limit telomerase activity (Wotton and Shore 1997). To prevent spreading of silent chromatin, subtelomeric regions contain boundaries, which are characterized by the presence of acetylated histone variant Htz1 (or H2A.Z) (Meneghini et al. 2003; Babiarz et al. 2006). In addition to the immediate zone of SIR-mediated repression, characterized by nucleosomes that lack detectable acetylation or methylation, there is a subtelomeric region called HAST for (Hda1affected subtelomeric) that is characterized by a continuous stretch of Hda1-deacetylated chromatin that extends 10 – 25 kb inwards from the telomeric repeat (Robyr et al. 2002). Thus telomeres are not only designed to ensure end replication, but they nucleate special domains with respect to SIR protein spreading and histone modifications.

MECHANISMS UNDERLYING NUCLEAR COMPARTMENTATION Anchoring of DNA at the Nuclear Envelope

The positioning of chromosomes within the nucleus depends on reversible interactions of chromosomal landmarks with structural features of the nucleus, such as the spindle pole body (SPB) and the inner nuclear envelope. Centromeres are held in a cluster near the SPB through short microtubules that persist through interphase (Jin et al. 2000; Bystricky et al. 2004). The tethering of telomeres in perinuclear foci is achieved by two redundant pathways that require Sir4, a silencing factor, and the yKu70/yKu80 heterodimer (Hediger et al. 2002; Taddei et al. 2004). Sir4 anchors repressed chromatin to the NE through its

partitioning and anchoring domain (PAD, aa 950 to 1262). PAD specifically binds Esc1, a low-abundance acidic protein associated exclusively with the inner face of the NE (Andrulis et al. 2002; Gartenberg et al. 2004; Taddei et al. 2004). By electron microscopy it was shown that Esc1 localizes in patches along the nuclear membrane independently of Sir4 and is excluded from nuclear pores (Taddei et al. 2004). Although yKu80 and Sir4 interact, a mutant allele of yKu80 that loses interaction with Sir4 (yKu80-4) (Taddei et al. 2004; Roy et al. 2004) can still tether chromatin to the NE (Taddei et al. 2004). Given that this occurs in strains deleted for the genes encoding Sir4 and Esc1, it was proposed that yKu should bind at least one additional membrane-bound factor. This is achieved indirectly through the ability of yKu to bind telomerase (Schober et al. 2009). Intriguingly, the requirements of the Sir4independent yKu tethering pathway varied with phases of the cell cycle: yKu80 tethering was dependent on yKu70 in G1-phase but not in S-phase cells (Taddei et al. 2004). Conversely, the domain of yKu80 that binds a stem-loop in the telomerase RNA Tlc1, was found to be necessary for telomere anchoring in S-phase, but not in G1-phase cells (Schober et al. 2009). The S-phase specific anchoring by yKu not only required Tlc1, but Est1 as well, which forms a complex with the catalytic subunit of telomerase, Est2. It was shown that a targeted Est2 fusion protein can anchor DNA to the nuclear perimeter in an Est1-dependent manner, and that Est1 interacts with an integral nuclear membrane protein, Mps3 (Fig. 4) (Schober et al. 2009; Jaspersen et al. 2002; Antoniacci et al. 2007). An acidic amino-terminal domain of this SUN-domain family member extends into the nucleoplasm where it contacts Est1 to tether telomerase-bound telomeres. Yet Mps3 also appears to contribute to an alternative telomere anchoring pathway mediated by Sir4 (Bupp et al. 2007). In both cases Mps3dependent anchoring is specific to S-phase cells. The budding yeast Mps3 also organizes the SPB (Jaspersen et al. 2002; Nishikawa et al.

A. Taddei, H. Schober, and S.M. Gasser Nuclear envelope

Nuclear pore

Mps3

Esc1 X?

G1

G1

S phase

G1 S phase

Est1 Sir4

Est3

Rap1

yKu

Tlc1 TG(1–6)

yKu Est2

Nucleosomes

Figure 4. Parallel mechanisms lead to yeast telomere attachment at the nuclear envelope. At different stages of the

cell cycle the telomere associated proteins mediate different contacts with inner nuclear membrane components. The Sir4-PAD domain binds the Esc1 C terminus, as well as yKu80 and Mps3. yKu80 binds telomerase, which also associates with Mps3 in S phase through Est1. There is an unidentified anchor for yKu in G1 phase that is neither Esc1- nor Mps3-dependent.

2003), yet by deletion of its N-terminal domain, the role of Mps3 in chromatin tethering could be separated from its essential role in spindle pole body organization (Bupp et al. 2007). Importantly, antagonism of the yKu-telomerase-Mps3 interaction led to hyper-recombination among telomeres, suggesting that this mechanism protects replicating ends from unequal strand invasion (Schober et al. 2009). This may be particularly relevant in S phase. Finally, it has been shown that the yKu-mediated peripheral positioning is switched off after DNA replication, leading to the dislodgment of telomeres from the nuclear envelope after replication (Ebrahimi and Donaldson 2008). Such release may facilitate mitotic separation of replicated telomeres. Besides these pathways other proteins such as Asf1, Rtt109, Esc2, and Ctf18 have been implicated yeast telomers anchoring, although it could not be determined whether their effects were direct or indirect (Hiraga et al. 2006, 2008). Indeed, mutations in proteins of the nuclear pore such as Mlp1, Mlp2, or Nup133 were shown to interfere with mRNA export and cell cycle progression, which could indirectly affect perinuclear organization.

Trans-Association of Chromatin Loci

The high concentration of macromolecules within the nucleoplasm (between 100 and 400 mg/ml) produces volume exclusion effects that enhance attractive interactions among macromolecules, a phenomenon known as macromolecular crowding. This effect has been proposed to favor the formation of compartments by driving bulky components into structurally compact organizations (Richter et al. 2007). Combination of these effects with specific intermolecular interactions and attachment to nuclear structures such as the NE could account for the formation of subnuclear compartments. One example of a functional compartment is provided by telomere clusters onto which silencing factors concentrate. The bifunctional role of Sir4 as a mediator of repression and an anchor for silent chromatin provides a mechanism for the self-organization of repressive compartments. In vivo, a nonsilent telomere can be localized at the NE through interaction with yKu, and thus be brought into proximity of other telomeres which generate a zone enriched for Sir proteins. Recruitment to this

The Budding Yeast Nucleus

compartment results in a higher probability that a subtelomeric gene gets repressed. Once Sir-dependent repression is established, the Sir4 anchoring pathway ensures that the silent chromatin stays perinuclear. In this way, the formation of silent chromatin is self-reinforcing, thanks to its ability to bind to the same sites as telomeres. Intriguingly, telomeres are not evenly distributed at the nuclear periphery but rather form discrete foci. We cannot rule out that telomere clustering arises in part from volume exclusion effects and molecular crowding (Iborra 2007), but it has also been shown that Sir proteins interact with themselves and with each other, establishing trans-interactions between telomeres. Recent work has shown that interactions between silent domains depends not only on silencing proteins Sir2, Sir3, and Sir4, but also on Sir1 and Esc2, two proteins involved in establishment of silencing at HM loci (Miele et al. 2009). Trans-interactions were not dependent on yKu or Esc1, suggesting that the mechanisms of NE-tethering and telomere clustering can be at least partially separated. Nonetheless, the self-organization of heterochromatin into perinuclear foci illustrates a self-perpetuating mechanism that applies to other chromatin-based domains of epigenetic character.

FUNCTIONAL CONSEQUENCES OF NUCLEAR ORGANIZATION ON GENE TRANSCRIPTION Nuclear Organization and Gene Silencing

Chromatin-based compartments like those discussed earlier not only concentrate factors where they are needed, but also keep factors away from sites where they would interfere with other types of regulation (Taddei et al. 2009). Specifically, the clustering of telomeres leads to the sequestration of SIRs, which was shown both to favor subtelomeric repression and to prevent promiscuous effects on a distinct subset of promoters (Taddei et al. 2009). Essential to such a phenomenon is the fact that the Sir3 protein level, estimated at 1400 copies per cell

(Ghaemmaghami et al. 2003), is both tightly regulated and limiting for the spread of silent chromatin (Renauld et al. 1993). Sir4 abundance is also tightly regulated, and low level overexpression of Sir4 actually disrupts telomeric repression. It has been shown that each Rap1 tail and each nucleosome in the silent subtelomeric domain binds one SIR complex comprising Sir2, Sir3 and Sir4 in precisely equal molar ratio (Martino et al. 2009). This means that each telomere of 16 Rap1 binding sites (Gilson et al. 1993) and 18 subtelomeric nucleosomes (Renauld et al. 1993; Strahl-Bolsinger et al. 1997), provides binding sites for 30 –40 SIR complexes. Because each haploid yeast cell has 32 telomeres clustered in three to eight telomeric foci, a focus contains several hundred potential SIR binding sites. The clustering of telomeric repeats could thus create a “sink” for limiting amounts of Sir proteins, and could account for the strikingly unequal distribution of this repressive complex within the nucleoplasm (Gotta et al. 1996). Confirming the functional consequences of this unequal distribution of Sir proteins, it was found that silencer-nucleated repression is highly sensitive to the distance of the reporter from a telomere (Renauld et al. 1993; Stavenhagen and Zakian 1994; Thompson et al. 1994; Maillet et al. 1996; Marcand et al. 1996). Reciprocally, the tethering of a weakened HMR silencer to the NE favored the repression of a reporter gene (Andrulis et al. 1998). Importantly, this effect depends on the ability of telomere clusters to concentrate SIR factors at the nuclear envelope (Mondoux et al. 2007; Taddei et al. 2009). Thus, it appears that transcriptional silencing is not inherent to position, but rather requires facilitated access to a local high concentration of SIR proteins. Confirming this, when telomere anchoring is impaired, by deletion of YKU70 and ESC1, transcription is affected differently at different loci: genes at internal loci flanked by silencers show enhanced repression, whereas telomere-proximal repression is lost (Maillet et al. 2001; Taddei et al. 2009). In conclusion, the efficiency with which silent chromatin is formed depends on spatial concern,

A. Taddei, H. Schober, and S.M. Gasser

such as the local concentration of SIR proteins, and the strength of cis-acting nucleation elements. In apparent contradiction with this interpretation is the absence of a correlation between the colocalization of a subtelomeric gene in telomere clusters (visualized as Rap1-GFP foci) and the efficiency of TPE (Mondoux et al. 2007). This may be because of the dynamic behavior of individual telomeres, which move into and out of Rap1 and SIR foci rapidly, presumably without loss of their epigenetic status, because they retain SIR factors (Schober et al. 2008). Indeed, a repressed ring of chromatin maintains a stable state of repression, even when it is excised from its subtelomeric chromosomal context, suggesting that a high local concentration of SIR factors is dispensable once silencing is established (Gartenberg et al. 2004). The promiscuous repression of nontelomeric genes by released SIR factors, was analyzed genome-wide using a computational program that maps putative transcriptional factor sites. The affected promoters carry specific transcription factor motifs for either Abf1 or the PAC factors, Pdf1 and Pdf2, which are part of the Rpd3L histone deacetylase complex (RNA Polymerase A and C promoters; Dequard-Chablat et al. 1991; Zhu et al. 2009). A related motif, RRPE (ribosomal RNA processing element; Hughes et al. 2000) also correlated weakly with promiscuous SIR regulation. Intriguingly, these elements are primarily bound to promoters of genes expressing enzymes and proteins involved in ribosome biogenesis. Down-regulation may be seen as a global attempt to suppress growth, and perhaps favor a stress survival pathway. These results make a forceful argument that changes in the spatial distribution of repetitive sequences that bind silencing factors can regulate patterns of gene expression genome-wide. The cell may exploit such a mechanism by controlling SIR complex release, an event that can respond rapidly to environmental insult (Martin et al. 1999; Ai et al. 2002). Consistently, SIR dispersion, or modulation of TPE, have been observed in the presence of various forms of stress (Stone and Pillus 1996; Martin et al.

1999; McAinsh et al. 1999; Mills et al. 1999; Ray and Runge 1999; Bi et al. 2004; Mercier et al. 2005), stress-induced redistribution of SIR proteins may derepress subtelomeric genes required for use of alternative carbon sources, and simultaneously contribute to the downregulation of genes involved in ribosome biogenesis. Nuclear Organization and Inducible Gene Expression

Over the last years nuclear pore complexes (NPCs) emerged as a major player in organizing gene activity. First, the nucleoporin Nup2 was shown to exert a strong boundary activity that can block the spread of heterochromatin when targeted on both sides of a reporter gene (Ishii et al. 2002). In this case, NPC tethering was proposed to establish a protected chromatin domain by creating a small DNA loop. Subsequently, a series of genome-wide studies identified nuclear pore components to be associated with highly active genes (Casolari et al. 2004, 2005; Schmid et al. 2006). Furthermore, specific inducible genes (INO1, HXK1, GAL1, GAL2, HSP104) were shown to associate with the nuclear periphery upon activation (Akhtar and Gasser 2007; Taddei 2007) (Fig. 5). Although many active genes seem to be associated with the NE, a stable interaction is not an obligate feature of gene activity (Casolari et al. 2004) and activation of the same promoter by different pathways or with different 30 UTR can alter a gene’s position within the nucleus (Abruzzi et al. 2006; Taddei et al. 2006). How specific genes associate with the NPC is still unclear and each step of mRNA production, maturation and export has been implicated at some level (Taddei 2007 for review). If multiple steps contribute to the stable association of active genes to NPCs, the importance of each individual step to NPC anchoring is probably specific for each gene. One possible scenario is that promoter-bound factors such as SAGA and Sus1 promote an initial, transient contact of the gene with pore proteins. This may be later stabilized by factors recruited for transcript termination, processing, quality

The Budding Yeast Nucleus Telomere clusters

A

Transcription compartments On Off

Off

Nucleolus

On

On

SPB

Foci of silent information regulators stem from anchored telomeres (Gotta et al. 1996) and favor repression (Andrulis et al. 1998) Boundary elements and stress-induced genes bind nuclear pores (Ishii et al. 2002; Casolari et al., 2004), in some cases increasing mRNA level (Brickner and Walter, 2004; Taddei et al. 2006)

On/Off Normal promoter-based Pol IIdependent transcription RNA Pol III (tRNA) transcription at the nucleolar edge (Thompson et al. 2003)

NPC Boundary

NPC

DNA Repair compartments

B Mps3

Mps3 binds Est1-telomerase to suppress tel-tel recombination (Oza et al. 2009; Schober et al. 2009)

NPC Heh1

Canonical HR-mediated repair (Bystricky et al. 2009) Nucleolus : exclusion of recombination proteins (Torres-Rosell et al. 2007) Nup84/Slx5/Slx8 regulates a switch to alternative repair pathways and telomerase-independent telomere maintenance (Azam et al. 2006; Nagai et al. 2008)

Heh1 NPC

Heh1 (Man1 homologue) suppresses rDNA recombination by tethering (Mekhail et al. 2008)

Figure 5. Nuclear subcompartments of yeast. (A) Transcription subcompartments within the yeast nucleus.

Telomere clustering favors repression through SIR factors whereas pore association correlates with induced expression of certain genes and boundary function. (B) DNA repair compartments within the yeast nucleus. The nucleoplasm is the site of Rad52-mediated recombination whereas sequestration at Mps3 or nuclear pores either suppresses recombination between telomeres or processes a DSB for alternative repair pathways. For the rDNA, binding to the INM protein Heh1 (Src1) prevents rDNA recombination.

control and export. These multiple anchors could favor the formation of active gene loops connecting promoters and 30 sequences, thus improving the recycling of polymerases and elongation efficiency (O’Sullivan et al. 2004). Gene-NPC association might be particularly important for inducible genes such as

galactose- and heat-shock controlled promoters for which NPC association involves the histone acetyltransferase SAGA complex. This complex is connected to the mRNA export machinery by one of its components, Sus1, which is also part of the Sac3-Thp1-Cdc31 complex that binds the NPC through Nup1. The SAGA

A. Taddei, H. Schober, and S.M. Gasser

complex is generally required for the induction of stress-responsive genes, suggesting a rationale for gene relocalization: Stress-induced genes are likely to require rapid and high level expression and export, which could be facilitated by their positioning at pores. Although, mutations affecting gene-NPC association show normal activation level for GAL1 (Cabal et al. 2006) it is still possible that the kinetics of induction are affected in these mutants. Importantly, association with the nuclear periphery has been shown to enhance the activation of at least some genes (Brickner and Walter 2004; Menon et al. 2005; Taddei et al. 2006) and could provide an additional layer of regulation for fine-tuning expression levels. Gene-NPC association has also been proposed to reflect an epigenetic mark that enables past events to be “remembered” (Brickner et al. 2007). It was reported that both the GAL1 and INO1 genes remain associated with the NPC for several generations after the inducing agent is removed. These genes appeared to be more rapidly re-activated in a pore-bound state than after a long-term repression. However, this mechanism has been called into question because other mechanisms involving cytoplasmic factors also can account for the rapid reactivation of these genes (Zacharioudakis et al. 2007). The mechanism through which NPC-association can influence expression is still unclear. In 1985, Blobel proposed the “gene gating” hypothesis according to which a “circumscribed space subjacent to the NPC and extending into the interior of the nucleus was envisioned to serve as the locale where transcription and much of the co- and posttranscriptional processing would occur” (Blobel 1985). Thus, the NPC was proposed to be a scaffold to build an assembly line favoring the coordination of the different processes that occur at an active gene. Because of their eightfold symmetry, each pore may accommodate multiple genes, forming a factory for highly efficiency transcription, in which factors that promote initiation or elongation can concentrate. The existence of internal transcription factories has been proposed in differentiation mammalian nuclei or in cultured

cells to coordinate expression of coregulated genes (Sexton et al. 2007). Such internal transcription factories might also exist in yeast although it remains unclear what provides the scaffolding for promoter recruitment. The Dual Role of the Nuclear Envelope

Given its role in the creation of repressive telomeric compartments and its ability to support, if not promote, high-level gene induction, one can conclude that the nuclear periphery has a dual role in regulating gene expression. This dual role is corroborated by the structural organization of the nuclear envelope. Indeed, microscopy studies show that repressive and activating compartments at the nuclear periphery coexist but do not overlap, as visualized by the positioning of subtelomeric domains in-between nuclear pores (Taddei et al. 2004). This proximity between repressive (telomere clusters) and activating compartments (NPCs) could favor the efficiency and reversibility of gene induction. This might be especially relevant for subtelomeric genes, which are mainly involved in the usage of alternative carbon sources and have to be induced only under specific growth conditions (Fabre et al. 2005). Consistently, increasing the association of the HXK1 subtelomeric gene with the nuclear periphery improves both its repression on glucose medium and its activation in the absence of glucose (Taddei et al. 2006). FUNCTIONAL CONSEQUENCES OF NUCLEAR ORGANIZATION ON GENOME STABILITY

Although these studies indicate important functions for yeast nuclear structure in the regulation of gene expression, recent work suggests that DNA repair may also respond to spatial cues in the nucleus. The most obvious elements that organize DNA repair are foci that form at double strand breaks (DSBs) as they are processed for homologous recombination (HR). This can be followed through the binding of fluorescently tagged Rad52, a protein that facilitate Rad51-dependent strand invasion, preceding

The Budding Yeast Nucleus

the formation of a Holliday junction (New et al. 1998). A Rad52-containing HR center is able to recruit more than one DSB (Lisby et al. 2003), suggesting that there is a self-recognition event that allows breaks to congregate. Foci of Rad52, whether spontaneous or induced by DNA damage, are strongly enriched in the nuclear interior (Bystricky et al. 2009). Given that sister chromatid exchange is the dominant pathway for homologous recombination in mitotically dividing cells (Kadyk and Hartwell 1992), the internal enrichment of Rad52 foci argues that most sister chromatid exchange occurs in the nuclear interior. Indeed, when the HMR or HML sequences are available as donors for the cleaved MATa locus, MAT stays internal and recruits the appropriate donor, e.g., HMLa, from a more peripheral position for gene conversion (Nagai et al. 2008; Bystricky et al. 2009). The inward shift is only seen for the appropriate donor locus, and thus is matingtype dependent. Pore Association of Persistent DSBs, Collapsed Forks and Critically Short Telomeres

If a donor sequence is not available, and the HO endonuclease is continuously expressed, the cut at the MAT locus persists. Such irreparable breaks are seen to shift to the nuclear periphery in a process dependent on Mec1 kinase and the histone variant Htz1 (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Several lines of evidence implicate a nuclear pore subcomplex as the binding site, although the INM protein Mps3 has also been implicated transiently in the relocation process (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Alternatively one might argue that the breaks move nonspecifically to the NE because crosslinking to both nuclear pores and Mps3, was reported (Nagai et al. 2008; Kalocsay et al. 2009; Oza et al. 2009). Links between DNA damage and nuclear pores was supported by an epistasis-miniarray profile (E-MAP) analysis (Collins et al. 2007b), in which the Nup84 subcomplex of the nuclear pore, containing Nup133, Nup120 and

Nup84, was clustered with a conserved SUMOdependent ubiquitin ligase complex, Slx5/Slx8. The Slx5/Slx8 complex plays an important role in the maintenance of genome integrity particularly during DNA replication (Perry et al. 2008). In addition, Slx5, which bears two SUMO interacting motifs, interacts physically with components of the proteasome (Collins et al. 2007a). Tellingly, there is an accumulation of SUMOylated proteins in cells lacking either Slx5 or Slx8 (Wang et al. 2006; Xie et al. 2007), suggesting that the Slx5/Slx8 complex normally helps the proteasome degrade SUMOylated proteins through its ubiquitin ligase activity (Wang et al. 2006; Xie et al. 2007). Indeed, an accumulation of SUMO conjugates has been shown to be responsible for the DNA damage sensitivity of the slx8 mutant in fission yeast (Prudden et al. 2007). Mutation of any component of the Nup84 complex, i.e., Nup84, Nup120, or Nup133 nucleoporins, renders cells hypersensitive to DNA damage, such as irradiation, MMS, bleomycin and HU (Bennett et al. 2001; Loeillet et al. 2005; Therizols et al. 2006). The same is true for mutation of Slx5 or Slx8 (Nagai et al. 2008). Furthermore, mutants of the Nup84 complex have been shown to be synthetically lethal with mutants of DNA repair, recombination and replication genes (Loeillet et al. 2005; Pan et al. 2006; Collins et al. 2007b; Nagai et al. 2008). A direct involvement of nuclear pores in DNA repair was supported by the fact that Nup84 and Slx5/Slx8 colocalize and coimmunoprecipitate (Nagai et al. 2008). ChIP assays have shown that irreparable DSBs and collapsed, but not stalled, replication forks, becomes associated with the Nup84 – Slx5/Slx8 complex. Moreover, the rate of spontaneous gene conversion could be enhanced by tethering the recipient site of recombination to the nuclear periphery. The enhancement of DNA repair was dependent on Slx8 and Nup84 proteins, suggesting that Slx5/Slx8 - Nup84 complex facilitates at least one kind of strand invasion event (Nagai et al. 2008). Short telomeres have also been reported to associate with the nuclear pore complex (Khadaroo et al. 2009).

A. Taddei, H. Schober, and S.M. Gasser

Importantly, pore binding is not detected if DSBs can be readily repaired by HR (Nagai et al. 2008). This suggests that only lesions that are refractive to DNA repair by canonical HR pathways associate stably with pores. At the pore a repair intermediate may be shunted to alternative recombination and repair pathways through steps that are controlled by SUMO recognition, ubiquitylation, and degradation, in other words through Slx5/Slx8 and the proteasome. The yeast proteasome was also shown by ChIP to be recruited to irreparable DSBs, and was genetically shown to be required for recombination-mediated modes of DNA repair (Krogan et al. 2004). In mammals, the proteasome has been implicated in the differential choice of recombination pathways; namely the up-regulation of gene-conversion at the expense of single-strand annealing (Gudmundsdottir et al. 2007). Consistently, Slx5/Slx8 has been shown to counteract Rad51-independent recombination, such as single-strand annealing and the break induced replication (Burgess et al. 2007). An increased dependence on Rad51-independent recombination pathways favors deleterious genomic rearrangements, such as gene amplifications, deletions and translocations (Haber and Debatisse 2006). Thus, we suggest the possibility that the pore-associated DNA repair pathway mediated by Slx5/Slx8 complex helps counteract error-prone Rad51-independent recombination, whereas up-regulating alternative pathways for recovery. Alternative pathways may be particularly important at collapsed replication forks and at telomeres. Which pathways are used, will depend on the targets of Slx5/Slx8 action. Based on E-MAP data likely pathways will include Pol32, Rad27, Est1, and Srs2 (Nagai et al. 2008). Intriguingly, the outgrowth of type II survivors of telomerase inhibition, which requires TG repeat amplification by recombination, requires not only Rad52 and Sgs1, but Slx5 and Slx8 (Azam et al. 2006; McEachern and Haber 2006). Given the involvement of the Slx5/8 E3 ligase and proteasome, it seems likely that shifting the mode of DNA repair at these loci may require the degradation of an inhibitor, one that could be

the target of SUMO-directed ubiquitination by Slx5/Slx8 (Collins et al. 2007a; Wang et al. 2006; Xie et al. 2007). Finally, there may be crosstalk between Mps3 and these pathways, because the mps3D75-150 mutant, which fails to bind DSBs and telomeres, delayed recombinational repair of DSBs and decreased gross chromosomal rearrangements in either a slx5 deletion or in the pif1-m2 mutant, which abrogates the negative regulation of telomerase activity by the Pif1 helicase (Oza et al. 2009). Consistently, overexpression of the Mps3N domain increased telomere exchange (Schober et al. 2009). Regulation of Recombination at the rDNA Locus

In budding yeast, the maintenance of rDNA repeat number is important for cell growth as well as for preventing premature senescence. It has been proposed that HR helps to maintain rDNA copy number (Smith 1974). Indeed, expansion or contraction of rDNA repeats appears to stem from HR, which follows on Fob1induced replication fork blocking (Kobayashi et al. 1998). Although several sequence elements that stimulate recombination are found within the rDNA (Keil and Roeder 1984; Kobayashi et al. 1998), the frequency of HR is nonetheless significantly lower than what one would expect for a repetitive array in yeast. This argues that recombination is actively suppressed at the rDNA locus to maintain rDNA repeat homeostasis. Interestingly, components of the HR machinery, namely Rad52, Rad51, Rad55, and Rad59, are excluded from the nucleolus, although sensors of DSBs, such as Mre11 and Rfa1 are present (Torres-Rosell et al. 2007). Furthermore, a single DSB in the rDNA induced by the I-SceI endonuclease shifts away from the nucleolus when Rad52 was recruited, suggesting repair of rDNA by HR can only take place outside of the nucleolus. Nucleolar exclusion of Rad52 foci requires the Smc5/Smc6 complex, which harbors E3 SUMO ligase activity, and requires SUMOylation of Rad52. Given that mutants that impair nucleolar exclusion of

The Budding Yeast Nucleus

HR, i.e., mutants of Smc5 or Smc6, show rDNA hyper-recombination and unequal sister chromatid exchange, one can conclude that the extrusion of HR proteins from the nucleolus is functionally important for genome stability (Burgess et al. 2007; Torres-Rosell et al. 2007). Although it is well-established that the yeast nucleolus abuts the nuclear periphery (Oakes et al. 1998), the functional significance of this localization has been poorly understood. Recent work shows that the perinuclear anchoring of rDNA repeats by the inner nuclear membrane proteins Heh1(also called Src1) and Nur1, contributes to the maintenance of rDNA repeat stability (Mekhail et al. 2008). Heh1 shares homology with human Man1 (King et al. 2006), an INM protein containing a conserved LEM (LAP-Emerin-Man1) domain. Deletion of Heh1 or Nur1 releases rDNA from the nuclear periphery, and like loss of Smc5/ Smc6, this release correlates with increase unequal sister chromatid exchange. Importantly, unlike Sir2, which also functions to reduce unequal sister chromatid exchange (Gottlieb and Esposito 1989), neither Heh1 nor Nur1 is required for rDNA silencing. The artificial tethering of repeats to the INM through a Sir2-Heh1 fusion suppresses rDNA instability in the absence of Lrs4, a protein required for peripheral tethering of the rDNA. Thus, the peripheral tethering appears to limit access of the repetitive arrays to the machinery mediating recombination and sister chromatid exchange (Mekhail et al. 2008).

adapts to changes in carbon source or encounters other conditions of metabolic stress. These changes reinforce the notion of “cell type-” or “condition-” specific levels of nuclear organization. Most importantly, yeast cells allow us to test for subtle changes in the efficiency of replication, DNA repair and transcription, as a result of changes in nuclear structure. Although the field is still at an early stage, the number of genes recognized as regulating aspects of nuclear structure increases constantly, and both microscopy and genomic assays improve at staggering rates. These are forceful arguments for pursuing yeast as a model in which to test the relationship between nuclear structure and function. ACKNOWLEDGMENTS

We would like to acknowledge critical reading by V. Dion and H. Ferreira, and many fruitful discussions with the Gasser laboratory. We thank S. Nagai for the DNA repair figure and K. Bystricky images in Figure 1. S.M.G. acknowledges support of the Novartis Research Foundation, the National Center for Competence in Research, “Frontiers-in-Genetics” and the EU NOE Epigenome. A.T. is supported by the French “Agence Nationale pour la Recherche” (ANR), and received funding from the European Research Council under the European Community’s Seventh Framework Programme (FP7/2007-2013 Grant Agreement no. [210508]). REFERENCES

CONCLUDING REMARKS

The yeast nucleus provides a readily manipulated genetic system for the analysis of nuclear organization. Its main advantage is that yeast provides ample means to test both gain and loss of function mutations on nuclear organization and genetic function. Data show clearly that there are zones that regulate DNA repair (Fig. 5B) just as there are zones that influence transcription (Fig. 5A). The one drawback of yeast is that it does not differentiate into highly differentiated cell types. It does, however, show changes in nuclear organization as the culture

Abruzzi KC, Belostotsky DA, Chekanova JA, Dower K, Rosbash M. 2006. 30 -end formation signals modulate the association of genes with the nuclear periphery as well as mRNP dot formation. EMBO J 25: 4253–4262. Ai W, Bertram PG, Tsang CK, Chan TF, Zheng XF. 2002. Regulation of subtelomeric silencing during stress response. Mol Cell 10: 1295–1305. Akhtar A, Gasser SM. 2007. The nuclear envelope and transcriptional control. Nat Rev Genet 8: 507– 17. Alber F, Dokudovskaya S, Veenhoff LM, Zhang W, Kipper J, Devos D, Suprapto A, Karni-Schmidt O, Williams R, Chait BT, et al. 2007. The molecular architecture of the nuclear pore complex. Nature 450: 695– 701. Andrulis ED, Neiman AM, Zappulla DC, Sternglanz R. 1998. Perinuclear localization of chromatin facilitates transcriptional silencing. Nature 394: 592– 595.

A. Taddei, H. Schober, and S.M. Gasser Andrulis ED, Zappulla DC, Ansari A, Perrod S, Laiosa CV, Gartenberg MR, Sternglanz R. 2002. Esc1, a nuclear periphery protein required for Sir4-based plasmid anchoring and partitioning. Mol Cell Biol 22: 8292– 8301. Antoniacci LM, Kenna MA, Skibbens RV. 2007. The nuclear envelope and spindle pole body-associated Mps3 protein bind telomere regulators and function in telomere clustering. Cell Cycle 6: 75– 79. Aparicio OM, Billington BL, Gottschling DE. 1991. Modifiers of position effect are shared between telomeric and silent mating-type loci in S. cerevisiae. Cell 66: 1279– 1287. Azam M, Lee JY, Abraham V, Chanoux R, Schoenly KA, Johnson FB. 2006. Evidence that the S.cerevisiae Sgs1 protein facilitates recombinational repair of telomeres during senescence. Nucleic Acids Res 34: 506 –516. Babiarz JE, Halley JE, Rine J. 2006. Telomeric heterochromatin boundaries require NuA4-dependent acetylation of histone variant H2A.Z in Saccharomyces cerevisiae. Genes Dev 20: 700 –710. Bennett CB, Lewis LK, Karthikeyan G, Lobachev KS, Jin YH, Sterling JF, Snipe JR, Resnick MA. 2001. Genes required for ionizing radiation resistance in yeast. Nat Genet 29: 426–434. Berger AB, Cabal GG, Fabre E, Duong T, Buc H, Nehrbass U, Olivo-Marin JC, Gadal O, Zimmer C. 2008. Highresolution statistical mapping reveals gene territories in live yeast. Nature Methods 5: 1031–1037. Bi X, Yu Q, Sandmeier JJ, Elizondo S. 2004. Regulation of transcriptional silencing in yeast by growth temperature. J Mol Biol 344: 893 –905. Blobel G. 1985. Gene gating: A hypothesis. Proc Natl Acad Sci 82: 8527–8529. Brickner JH, Walter P. 2004. Gene recruitment of the activated INO1 locus to the nuclear membrane. PLoS Biol 2: e342. Brickner DG, Cajigas I, Fondufe-Mittendorf Y, Ahmed S, Lee PC, Widom J, Brickner JH. 2007. H2A.Z-mediated localization of genes at the nuclear periphery confers epigenetic memory of previous transcriptional state. PLoS Biol 5: e81. Bryk M, Banerjee M, Murphy M, Knudsen KE, Garfinkel DJ, Curcio MJ. 1997. Transcriptional silencing of Ty1 elements in the RDN1 locus of yeast. Genes Dev 11: 255–269. Bupp JM, Martin AE, Stensrud ES, Jaspersen SL. 2007. Telomere anchoring at the nuclear periphery requires the budding yeast Sad1-UNC-84 domain protein Mps3. J Cell Biol 179: 845 –854. Burgess RC, Rahman S, Lisby M, Rothstein R, Zhao X. 2007. The Slx5-Slx8 complex affects sumoylation of DNA repair proteins and negatively regulates recombination. Mol Cell Biol 27: 6153–6162. Bystricky K, Heun P, Gehlen L, Langowski J, Gasser SM. 2004. Long-range compaction and flexibility of interphase chromatin in budding yeast analyzed by highresolution imaging techniques. Proc Natl Acad Sci U S A 101: 16495– 16500. Bystricky K, Laroche T, van Houwe G, Blaszczyk M, Gasser SM. 2005. Chromosome looping in yeast: telomere pairing and coordinated movement reflect anchoring

efficiency and territorial organization. J Cell Biol 168: 375 –387. Bystricky K, Van Attikum H, Montiel MD, Dion V, Gehlen L, Gasser SM. 2009. Regulation of nuclear positioning and dynamics of the silent mating type loci by the yeast Ku70/80 complex. Mol Cell Biol 29: 835 –848. Cabal GG, Genovesio A, Rodriguez-Navarro S, Zimmer C, Gadal O, Lesne A, Buc H, Feuerbach-Fournier F, OlivoMarin JC, Hurt EC, et al. 2006. SAGA interacting factors confine sub-diffusion of transcribed genes to the nuclear envelope. Nature 441: 770– 773. Casolari JM, Brown CR, Drubin DA, Rando OJ, Silver PA. 2005. Developmentally induced changes in transcriptional program alter spatial organization across chromosomes Genes Dev 19: 1188–98. Casolari JM, Brown CR, Komili S, West J, Hieronymus H, Silver PA. 2004. Genome-wide localization of the nuclear transport machinery couples transcriptional status and nuclear organization. Cell 117: 427– 439. Collins SR, Kemmeren P, Zhao XC, Greenblatt JF, Spencer F, Holstege FC, Weissman JS, Krogan NJ. 2007a. Toward a comprehensive atlas of the physical interactome of Saccharomyces cerevisiae. Mol Cell Proteomics 6: 439–450. Collins SR, Miller KM, Maas NL, Roguev A, Fillingham J, Chu CS, Schuldiner M, Gebbia M, Recht J, Shales M et al. 2007b. Functional dissection of protein complexes involved in yeast chromosome biology using a genetic interaction map. Nature 446: 806 –810. Cremer T, Cremer M, Dietzel S, Mu¨ller S, Solovei I, Fakan S. 2006. Chromosome territories – a functional nuclear landscape. Curr Opin Cell Biol 18: 307 –316. D’Angelo MA, Hetzer MW. 2008. Structure, dynamics and function of nuclear pore complexes. Trends Cell Biol 18: 456 –466. Dequard-Chablat M, Riva M, Carles C, Sentenac A. 1991. RPC19, the gene for a subunit common to yeast RNA polymerases A (I) and C (III). J Biol Chem 266: 15300– 15307. Dekker J, Rippe K, Dekker M, Kleckner N. 2002. Capturing chromosome conformation. Science 295: 1306– 1311. Ebrahimi H, Donaldson AD. 2008. Release of yeast telomeres from the nuclear periphery is triggered by replication and maintained by suppression of Ku-mediated anchoring. Genes Dev 22: 3363–3374. Fabre E, Muller H, Therizols P, Lafontaine I, Dujon B, Fairhead C. 2005. Comparative genomics in hemiascomycete yeasts: Evolution of sex, silencing, and subtelomeres. Mol Biol Evol 22: 856–873. Fisher TS, Taggart AK, Zakian VA. 2004. Cell cycledependent regulation of yeast telomerase by Ku. Nat Struct Mol Biol 11: 1198– 1205. Fritze CE, Verschueren K, Strich R, Easton Esposito R. 1997. Direct evidence for SIR2 modulation of chromatin structure in yeast rDNA. EMBO J 16: 6495–6509. Gall JG. 2000. Cajal bodies: The first 100 years. Annu Rev Cell Dev Biol 16: 273 –300. Gasser SM, Hediger F, Taddei A, Neumann FR, Gartenberg MR 2004. The function of telomere clustering in yeast: The Circe effect. Cold Spring Harb Symp Quant Biol 69: 327 –337.

The Budding Yeast Nucleus Gartenberg MR, Neumann FR, Laroche T, Blaszczyk M, Gasser SM. 2004. Sir-mediated repression can occur independently of chromosomal and subnuclear contexts. Cell 119: 955– 967. Ghaemmaghami S, Huh WK, Bower K, Howson RW, Belle A, Dephoure N, O’Shea EK, Weissman JS. 2003. Global analysis of protein expression in yeast. Nature 425: 737–741. Gilson E, Roberge M, Giraldo R, Rhodes D, Gasser SM. 1993. Distortion of the DNA double helix by RAP1 at silencers and multiple telomeric binding sites. J Mol Biol 231: 293–310. Gotta M, Laroche T, Formenton A, Maillet L, Scherthan H, Gasser SM. 1996. The clustering of telomeres and colocalization with Rap1, Sir3, and Sir4 proteins in wild-type Saccharomyces cerevisiae. J Cell Biol 134: 1349–1363. Gottlieb S, Esposito RE. 1989. A new role for a yeast transcriptional silencer gene, SIR2, in regulation of recombination in ribosomal DNA. Cell 56: 771 –776. Gruenbaum Y, Margalit A, Goldman RD, Shumaker DK, Wilson KL. 2005. The nuclear lamina comes of age. Nat Rev Mol Cell Biol. 6: 21–31. Grund SE, Fischer T, Cabal GG, Antunez O, Perez-Ortin JE, Hurt E. 2008. The inner nuclear membrane protein Src1 associates with subtelomeric genes and alters their regulated gene expression. J Cell Biol 182: 897– 910. Gudmundsdottir K, Lord CJ, Ashworth A. 2007. The proteasome is involved in determining differential utilization of double-strand break repair pathways. Oncogene 26: 7601–7606. Haber JE, Debatisse M. 2006. Gene amplification: Yeast takes a turn. Cell 125: 1237–1240. Haeusler RA, Pratt-Hyatt M, Good PD, Gipson TA, Engelke DR. 2008. Clustering of yeast tRNA genes is mediated by specific association of condensin with tRNA gene transcription complexes. Genes Dev 22: 2204– 2214. Hattier T, Andrulis ED, Tartakoff AM. 2007. Immobility, inheritance and plasticity of shape of the yeast nucleus. BMC Cell Biol 8: 47. Hediger F, Neumann FR, Van Houwe G, Dubrana K, Gasser SM. 2002. Live imaging of telomeres: yKu and Sir proteins define redundant telomere-anchoring pathways in yeast. Curr Biol 12: 2076– 2089. Heun P, Laroche T, Shimada K, Furrer P, Gasser SM. 2001. Chromosome dynamics in the yeast interphase nucleus. Science 294: 2181–2186. Hiraga S, Botsios S, Donaldson AD. 2008. Histone H3 lysine 56 acetylation by Rtt109 is crucial for chromosome positioning. J Cell Biol 183: 641–651. Hiraga S, Robertson ED, Donaldson AD. 2006. The Ctf18 RFC-like complex positions yeast telomeres but does not specify their replication time. EMBO J 25: 1505– 1514. Hug N, Lingner J. 2006. Telomere length homeostasis. Chromosoma. 115: 413– 25. Hughes JD, Estep PW, Tavazoie S, Church GM. 2000. Computational identification of cis-regulatory elements associated with groups of functionally related genes in Saccharomyces cerevisiae. J Mol Biol 296: 1205–1214.

Huh WK, Falvo JV, Gerke LC, Carroll AS, Howson RW, Weissman JS, O’Shea EK. 2003. Global analysis of protein localization in budding yeast. Nature 425: 686 –691. Iborra FJ. 2007. Can visco-elastic phase separation, macromolecular crowding and colloidal physics explain nuclear organisation? Theor Biol Med Model 4: 15. Ishii K, Arib G, Lin C, Van Houwe G, Laemmli UK. 2002. Chromatin boundaries in budding yeast: The nuclear pore connection. Cell 109: 551 –562. Jaspersen SL, Giddings THJr, Winey M. 2002. Mps3p is a novel component of the yeast spindle pole body that interacts with the yeast centrin homologue Cdc31p. J Cell Biol 159: 945– 956. Jin QW, Fuchs J, Loidl J. 2000. Centromere clustering is a major determinant of yeast interphase nuclear organization. J Cell Sci 113: 1903–1912. Kadyk LC, Hartwell LH. 1992. Sister chromatids are preferred over homologs as substrates for recombinational repair in Saccharomyces cerevisiae. Genetics 132: 387 –402. Kalocsay M, Hiller NJ, Jentsch S. 2009. Chromosome-wide Rad51 spreading and SUMO-H2A.Z-dependent chromosome fixation in response to a persistent DNA doublestrand break. Mol Cell 33: 335 –343. Keil RL, Roeder GS. 1984. Cis-acting, recombinationstimulating activity in a fragment of the ribosomal DNA of S. cerevisiae. Cell 39: 377– 386. Khadaroo B, Teixeira MT, Luciano P, Eckert-Boulet N, Germann SM, Simon MN, Gallina I, Abdallah P, Gilson E, Ge´li V, et al. 2009. The DNA damage response at eroded telomeres and tethering to the nuclear pore complex. Nat Cell Biol Jul 13. [Epub ahead of print] King MC, Lusk CP, Blobel G. 2006. Karyopherin-mediated import of integral inner nuclear membrane proteins. Nature 442: 1003– 1007. Kitamura E, Blow JJ, Tanaka TU. 2006. Live-cell imaging reveals replication of individual replicons in eukaryotic replication factories. Cell, 125: 1297– 1308. Kobayashi T, Heck DJ, Nomura M, Horiuchi T. 1998. Expansion and contraction of ribosomal DNA repeats in Saccharomyces cerevisiae: Requirement of replication fork blocking (Fob1) protein and the role of RNA polymerase I. Genes Dev 12: 3821–3830. Krogan N, Lam MHY, Fillingham J, Keogh MC, Gebbia M, Li J, Datta N, Cagney G, Buratowski S, Emili A, et al. 2004. Proteasome involvement in the repair of DNA doublestrand breaks. Molecular Cell 16: 1027– 1034. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Mol Cell Biol 4: 605–612. Larrivee M, LeBel C, Wellinger RJ. 2004. The generation of proper constitutive G-tails on yeast telomeres is dependent on the MRX complex. Genes Dev 18: 1391–1396. Lendvay TS, Morris DK, Sah J, Balasubramanian B, Lundblad V. 1996. Senescence mutants of Saccharomyces cerevisiae with a defect in telomere replication identify three additional EST genes. Genetics 144: 1399– 1412. Lewis A, Felberbaum R, Hochstrasser M. 2007. A nuclear envelope protein linking nuclear pore basket assembly, SUMO protease regulation, and mRNA surveillance. J Cell Biol 178: 813– 27.

A. Taddei, H. Schober, and S.M. Gasser Lingner J, Cech TR, Hughes TR, Lundblad V. 1997. Three ever shorter telomere (EST) genes are dispensable for in vitro yeast telomerase activity. Proc Natl Acad Sci 94: 11190–11195. Lisby M, Mortensen UH, Rothstein R. 2003. Colocalization of multiple DNA double-strand breaks at a single Rad52 repair centre. Nat Cell Biol 5: 572– 577. Loeillet S, Palancade B, Cartron M, Thierry A, Richard GF, Dujon B, Doye V, Nicolas A. 2005. Genetic network interactions among replication, repair and nuclear pore deficiencies in yeast. DNA Repair (Amst) 4: 459– 468. Lundblad V, Szostak JW. 1989. A mutant with a defect in telomere elongation leads to senescence in yeast. Cell 57: 633 –643. Lusk CP, Blobel G, King MC. 2007. Highway to the inner nuclear membrane: rules for the road. Nat Rev Mol Cell Biol 8: 414– 420. Maillet L, Boscheron C, Gotta M, Marcand S, Gilson E, Gasser SM. 1996. Evidence for silencing compartments within the yeast nucleus: A role for telomere proximity and Sir protein concentration in silencer-mediated repression. Genes Dev 10: 1796– 1811. Maillet L, Gaden F, Brevet V, Fourel G, Martin SG, Dubrana K, Gasser SM, Gilson E. 2001. Ku-deficient yeast strains exhibit alternative states of silencing competence. EMBO Rep 2: 203 –210. Marcand S, Gilson E, Shore D. 1997. A protein-counting mechanism for telomere length regulation in yeast. Science 275: 986– 990. Marcand S, Buck SW, Moretti P, Gilson E, Shore D. 1996. Silencing of genes at nontelomeric sites in yeast is controlled by sequestration of silencing factors at telomeres by Rap 1 protein. Genes Dev 10: 1297– 1309. Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A, Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7: 930– 939. Martin SG, Laroche T, Suka N, Grunstein M, Gasser SM. 1999. Relocalization of telomeric Ku and SIR proteins in response to DNA strand breaks in yeast. Cell 97: 621–633. Martino F, Kueng S, Robinson P, Tsai-Pflugfelder M, van Leeuwen F, Ziegler M, Cubizolles F, Cockell MM, Rhodes D, Gasser SM. 2009. Reconstitution of yeast silent chromatin: Multiple contact sites and O-AADPR binding load SIR complexes onto nucleosomes in vitro. Mol Cell 33: 323 –334. McAinsh AD, Scott-Drew S, Murray JA, Jackson SP. 1999. DNA damage triggers disruption of telomeric silencing and Mec1p-dependent relocation of Sir3p. Curr Biol 9: 963–966. McEachern MJ, Haber JE. 2006. Break-induced replication and recombinational telomere elongation in yeast. Annu Rev Biochem 75: 111–135. Mekhail K, Seebacher J, Gygi SP, Moazed D. 2008. Role for perinuclear chromosome tethering in maintenance of genome stability. Nature 456: 667 –670. Meneghini MD, Wu M, Madhani HD. 2003. Conserved histone variant H2A.Z protects euchromatin from the ectopic spread of silent heterochromatin. Cell 112: 725–736.

Menon BB, Sarma NJ, Pasula S, Deminoff SJ, Willis KA, Barbara KE, Andrews B, Santangelo GM. 2005. Reverse recruitment: the Nup84 nuclear pore subcomplex mediates Rap1/Gcr1/Gcr2 transcriptional activation. Proc Natl Acad Sci 102: 5749–5754. Mercier G, Berthault N, Touleimat N, Kepes F, Fourel G, Gilson E, Dutreix M. 2005. A haploid-specific transcriptional response to irradiation in Saccharomyces cerevisiae. Nucleic Acids Res 33: 6635–6643. Miele A, Bystricky K, Dekker J. 2009. Yeast silent mating type loci form heterochromatic clusters through silencer protein-dependent long-range interactions. PLoS Genet 5: e1000478. Epub 2009 May 8. Mills K, Sinclair D, Guarente L. 1999. MEC1-dependent redistribution of the Sir3 silencing protein from telomeres to DNA double-strand breaks. Cell 97: 609 –620. Misteli T. 2000. Cell biology of transcription and pre-mRNA splicing: Nuclear architecture meets nuclear function. J Cell Sci 113: 1841– 1849. Mondoux MA, Scaife JG, Zakian VA. 2007. Differential nuclear localization does not determine the silencing status of Saccharomyces cerevisiae telomeres. Genetics 177: 2019–2029. Moretti P, Freeman K, Coodly L, Shore D. 1994. Evidence that a complex of SIR proteins interacts with the silencer and telomere-binding protein RAP1. Genes Dev 8: 2257–2269. Nagai S, Dubrana K, Tsai-Pflugfelder M, Davidson MB, Roberts TM, Brown GW, Varela E, Hediger F, Gasser SM, Krogan NJ. 2008. Functional targeting of DNA damage to a nuclear pore-associated SUMO-dependent ubiquitin ligase. Science 322: 597– 602. New JH, Sugiyama T, Zaitseva E, Kowalczykowski SC. 1998. Rad52 protein stimulates DNA strand exchange by Rad51 and replication protein A. Nature 391: 407– 410. Nishikawa S, Terazawa Y, Nakayama T, Hirata A, Makio T, Endo T. 2003. Nep98p is a component of the yeast spindle pole body and essential for nuclear division and fusion. J Biol Chem 278: 9938–9943. Oakes M, Aris JP, Brockenbrough JS, Wai H, Vu L, Nomura M. 1998. Mutational analysis of the structure and localization of the nucleolus in the yeast Saccharomyces cerevisiae. J Cell Biology 143: 23– 34. Oakes M, Nogi Y, Clark MW, Nomura M. 1993. Structural alterations of the nucleolus in mutants of Saccharomyces cerevisiae defective in RNA polymerase I. Mol Cell Biol 13: 2441–2455. O’Sullivan JM, Tan-Wong SM, Morillon A, Lee B, Coles J, Mellor J, Proudfoot NJ. 2004. Gene loops juxtapose promoters and terminators in yeast. Nat Genet 36: 1014–1018. Oza P, Jaspersen SL, Miele A, Dekker J, Peterson CL. 2009. Mechanisms that regulate localization of a DNA doublestrand break to the nuclear periphery. Genes Dev 23: 912 –927. Palancade B, Liu X, Garcia-Rubio M, Aguilera A, Zhao X, Doye V. 2007. Nucleoporins prevent DNA damage accumulation by modulating Ulp1-dependent sumoylation processes. Mol Biol Cell 18: 2912– 2923.

The Budding Yeast Nucleus Pan X, Ye P, Yuan DS, Wang X, Bader JS, Boeke JD. 2006. A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell 124: 1069–1081. Perry JJ, Tainer JA, Boddy MN. 2008. A SIM-ultaneous role for SUMO and ubiquitin. Trends Biochem Sci 33: 201–208. Powers T, Walter P. 1999. Regulation of ribosome biogenesis by the rapamycin-sensitive TOR-signaling pathway in Saccharomyces cerevisiae. Mol Biol Cell 10: 987–1000. Prudden J, Pebernard S, Raffa G, Slavin DA, Perry JJ, Tainer JA, McGowan CH, Boddy MN. 2007. SUMO-targeted ubiquitin ligases in genome stability. EMBO J 26: 4089– 4101. ¨ ber Zelltheilung. Morphologisches Jahrbuch Rabl C. 1885. U 10: 214 –330. Ray A, Runge KW. 1999. Varying the number of telomerebound proteins does not alter telomere length in tel1D cells. Proc Natl Acad Sci U S A 96: 15044–15049. Renauld H, Aparicio OM, Zierath PD, Billington BL, Chhablani SK, Gottschling DE. 1993. Silent domains are assembled continuously from the telomere and are defined by promoter distance and strength, and by SIR3 dosage. Genes Dev 7: 1133– 1145. Richter K, Nessling M, Lichter P. 2007. Experimental evidence for the influence of molecular crowding on nuclear architecture. J Cell Sci 120: 1673–1680. Robyr D, Suka Y, Xenarios I, Kurdistani SK, Wang A, Suka N, Grunstein M. 2002. Microarray deacetylation maps determine genome-wide functions for yeast histone deacetylases. Cell 109: 437 –446. Rodley CD, Bertels F, Jones B, O’Sullivan JM. 2009. Global identification of yeast chromosome interactions using Genome conformation capture. Fungal Genet Biol 46: 879–886. Roy R, Meier B, McAinsh AD, Feldmann HM, Jackson SP. 2004. Separation-of-function mutants of yeast Ku80 reveal a Yku80p-Sir4p interaction involved in telomeric silencing. J Biol Chem 279: 86–94. Schmid M, Arib G, Laemmli C, Nishikawa J, Durussel T, Laemmli UK. 2006. Nup-PI: the nucleopore-promoter interaction of genes in yeast. Mol Cell 21: 379 –91. Schober H, Ferreira H, Kalck V, Gehlen LR, Gasser SM. 2009. Yeast telomerase and the SUN domain protein Mps3 anchor telomeres and repress subtelomeric recombination. Genes Dev 23: 928– 938. Schober H, Kalck V, Vega-Palas MA, Van Houwe G, Sage D, Unser M, Gartenberg MR, Gasser SM. 2008. Controlled exchange of chromosomal arms reveals principles driving telomere interactions in yeast. Genome Res 18: 261 –271. Sexton T, Umlauf D, Kurukuti S, Fraser P. 2007. The role of transcription factories in large-scale structure and dynamics of interphase chromatin. Semin Cell Dev Biol 18: 691 –697. Shampay J, Szostak JW, Blackburn EH. 1984. DNA sequences of telomeres maintained in yeast. Nature 310: 154–157. Shore D, Nasmyth K. 1987. Purification and cloning of a DNA binding protein from yeast that binds to both silencer and activator elements. Cell 51: 721– 732. Sinclair DA, Guarente L. 1997. Extrachromosomal rDNA circles –a cause of aging in yeast. Cell 91: 1033– 1042.

Singer MS, Gottschling DE. 1994. TLC1: template RNA component of Saccharomyces cerevisiae telomerase. Science 266: 404–409. Sirri V, Urcuqui-Inchima S, Roussel P, Hernandez-Verdun D. 2008. Nucleolus: The fascinating nuclear body. Histochem Cell Biol 129: 13– 31. Smith GP. 1974. Unequal crossover and the evolution of multigene families. Cold Spring Harb Symp Quant Biol 38: 507– 513. Smith JS, Boeke JD. 1997. An unusual form of transcriptional silencing in yeast ribosomal DNA. Genes Dev 11: 241 –254. Stavenhagen JB, Zakian VA. 1994. Internal tracts of telomeric DNA act as silencers in Saccharomyces cerevisiae. Genes Dev 8: 1411–1422. Stellwagen AE, Haimberger ZW, Veatch JR, Gottschling DE. 2003. Ku interacts with telomerase RNA to promote telomere addition at native and broken chromosome ends. Genes Dev 17: 2384– 2395. Stone EM, Pillus L. 1996. Activation of an MAP kinase cascade leads to Sir3p hyperphosphorylation and strengthens transcriptional silencing. J Cell Biol 135: 571– 583. Strahl-Bolsinger S, Hecht A, Luo K, Grunstein M. 1997. SIR2 and SIR4 interactions differ in core and extended telomeric heterochromatin in yeast. Genes Dev 11: 83– 93. Taddei A. 2007. Active genes at the nuclear pore complex. Curr Opin Cell Biol 19: 305– 310. Taddei A, Hediger F, Neumann FR, Bauer C, Gasser SM. 2004. Separation of silencing from perinuclear anchoring functions in yeast Ku80, Sir4 and Esc1 proteins. EMBO J 23: 1301–1312. Taddei A, Van Houwe G, Hediger F, Kalck V, Cubizolles F, Schober H, Gasser SM. 2006. Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature 441: 774–778. Taddei A, Van Houwe G, Nagai S, Erb I, van Nimwegen E, Gasser SM. 2009. The functional importance of telomere clustering: global changes in gene expression result from SIR factor dispersion. Genome Res 19: 611– 625. Therizols P, Fairhead C, Cabal GG, Genovesio A, OlivoMarin JC, Dujon B, Fabre E. 2006. Telomere tethering at the nuclear periphery is essential for efficient DNA double strand break repair in subtelomeric region. J Cell Biol 172: 189– 199. Thompson JS, Johnson LM, Grunstein M. 1994. Specific repression of the yeast silent mating locus HMR by an adjacent telomere. Mol Cell Biol 14: 446 –455. Thompson M, Haeusler RA, Good PD, Engelke DR. 2003. Nucleolar clustering of dispersed tRNA genes. Science 302: 1399– 1401. Torres-Rosell J, Sunjevaric I, De Piccoli G, Sacher M, EckertBoulet N, Reid R, Jentsch S, Rothstein R, Aragon L, Lisby M. 2007. The Smc5-Smc6 complex and SUMO modification of Rad52 regulates recombinational repair at the ribosomal gene locus. Nat Cell Biol 9: 923– 931. Verheggen C, Mouaikel J, Thiry M, Blanchard JM, Tollervey D, Bordonne R, Lafontaine DL, Bertrand E. 2001. Box C/ D small nucleolar RNA trafficking involves small nucleolar RNP proteins, nucleolar factors and a novel nuclear domain. EMBO J 20: 5480–5490.

A. Taddei, H. Schober, and S.M. Gasser Wang Z, Jones GM, Prelich G. 2006. Genetic analysis connects SLX5 and SLX8 to the SUMO pathway in Saccharomyces cerevisiae. Genetics 172: 1499– 1509. Wang L, Haeusler RA, Good PD, Thompson M, Nagar S, Engelke DR. 2005. Silencing near tRNA genes requires nucleolar localization. J Biol Chem 280: 8637–8639. Wotton D, Shore D. 1997. A novel Rap1p-interacting factor, Rif2p, cooperates with Rif1p to regulate telomere length in Saccharomyces cerevisiae. Genes Dev 11: 748– 760. Xie Y, Kerscher O, Kroetz MB, McConchie HF, Sung P, Hochstrasser M. 2007. The yeast Hex3.Slx8 heterodimer is a ubiquitin ligase stimulated by substrate sumoylation. J Biol Chem 282: 34176–34184. Yang CH, Lambie EJ, Hardin J, Craft J, Snyder M. 1989. Higher order structure is present in the yeast nucleus: autoantibody probes demonstrate that the nucleolus

lies opposite the spindle pole body. Chromosoma 98: 123 –128. Zacharioudakis I, Gligoris T, Tzamarias D. 2007. A yeast catabolic enzyme controls transcriptional memory. Curr Biol 17: 2041–2046. Zakian VA. 1996. Structure, function, and replication of Saccharomyces cerevisiae telomeres. Annu Rev Genet 30: 141 –72. Zhong S, Salomoni P, Pandolfi PP. 2000. The transcriptional role of PML and the nuclear body. Nat Cell Biol 2:, E85–90. Zhu C, Byers KJ, McCord RP, Shi Z, Berger MF, Newburger DE, Saulrieta K, Smith Z, Shah MV, Radhakrishnan M et al. 2009. High-resolution DNA-binding specificity analysis of yeast transcription factors. Genome Res 19: 556 –566.

Nuclear Functions of Actin Neus Visa1 and Piergiorgio Percipalle2 1

Department of Molecular Biology & Functional Genomics, Stockholm University, SE-106 91, Stockholm, Sweden

2

Department of Cell and Molecular Biology, Karolinska Institutet, SE-171 77, Stockholm, Sweden

Correspondence: [email protected] and [email protected]

Actin participates in several essential processes in the cell nucleus. Even though the presence of actin in the nucleus was proposed more than 30 years ago, nuclear processes that require actin have been only recently identified. Actin is part of chromatin remodeling complexes; it is associated with the transcription machineries; it becomes incorporated into newly synthesized ribonucleoproteins; and it influences long-range chromatin organization. As in the cytoplasm, nuclear actin works in conjunction with different types of actin-binding proteins that regulate actin function and bridge interactions between actin and other nuclear components.

ctin is a highly conserved protein of approximately 42 kDa found in all eukaryotic cells. It is a major component of the cytoskeleton and plays fundamental roles in essential biological processes such as determining cell shape, cell migration, and intracellular trafficking. The functions of actin in the cytoplasm are intrinsically coupled to the dynamics of actin polymerization, which is a tightly regulated process that responds to extracellular signals (reviewed by Moustakas and Heldin 2008; Papakonstanti and Stournaras 2008). Early studies raised the possibility that actin is also present in the cell nucleus and is implicated in the expression of protein-coding genes (Scheer et al. 1984; Egly et al. 1984). However, the existence of the so-called “nuclear actin” was

A

initially met with massive skepticism (reviewed by Pederson and Aebi 2002). Biochemists could not rule out contamination artifacts in nuclear preparations because of the high abundance of actin in the cytoplasm, and microscopists could not visualize in the cell nucleus the conspicuous actin filaments that are commonly observed in the cytoplasm. Nevertheless, research performed in the last 10 years has provided convincing evidence for the existence of actin in the cell nucleus and for the involvement of actin in fundamental nuclear processes. Actin is part of the chromatin remodeling complex; it is associated with the transcription machineries; it associates with newly synthesized ribonucleoproteins; and it influences long-range chromatin organization.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000620 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000620

N. Visa and P. Percipalle

ACTIN AND CHROMATIN REMODELING

Actin participates in gene expression as a component of chromatin-modifying complexes. Early findings by Crabtree and coworkers revealed that actin interacts with Brg1, the ATPase subunit of the BAF (Brg or Brm Associated Factors) SWI/SNF-like chromatin remodeling complex (Zhao et al. 1998). Since then, b-actin and a considerable number of actin-related proteins (ARPs) have been identified as components of different types of chromatin remodeling and histone acetyltransferase (HAT) complexes in a wide range of organisms from yeast to man (reviewed by Olave et al. 2002; Chen and Shen 2007; Farrants 2008). A central question that has not been fully answered concerns the mechanism(s) by which actin and ARPs contribute to chromatin remodeling. Not all chromatin remodeling complexes contain actin or ARPs, which indicates that these proteins are not essential for chromatin remodeling per se. The fact that actin and ARPs often bind to the ATPase subunit of the chromatin remodeling complexes suggests that actin and ARPs work as allosteric regulators of chromatin remodeling (Blessing et al. 2004). In agreement with this possibility, Zhao et al. (1998) showed that actin is needed for the full ATPase activity of the BAF complex. Actin also mediates or facilitates the binding of BAF to chromatin (Zhao et al. 1998). The ability of Brg1 to bind actin filaments in vitro (Rando et al. 2002) and the association of actin with the RNA polymerase (see the following discussion) suggest that actin recruits BAF to transcribed genes by bridging the interaction between the chromatin remodeling complex and the transcription machinery. In other cases, the binding of ARPs to core histones might bridge the interaction (reviewed by Blessing et al. 2004). Independently of its function in chromatin remodeling, actin participates in the regulation of a subset of genes that code for components of the actin cytoskeleton and that are activated by the serum response factor (SRF) (reviewed by Farrants 2008; Vartiainen 2008). This function of actin is mediated by MAL, a transcriptional coactivator of SRF that can sense variations in

the cellular concentration of G-actin (Miralles et al. 2003). MAL is an actin-binding protein that shuttles continuously between the nucleus and the cytoplasm, and only the actin-bound form of MAL is efficiently exported from the nucleus. Actin regulates both the nucleocytoplasmic transport of MAL and the ability of the MAL-SRF complex to activate SRF target genes. Serum stimulation reduces the rate of MAL nuclear export. Under these conditions, MAL and SRF associate with the SRF target promoters, and the SRF target genes become activated upon disruption of the MAL-actin interaction (Vartiainen et al. 2007). ACTIN IN ASSOCIATION WITH THE TRANSCRIPTION MACHINERY

Actin is associated with all three RNA polymerases, Pol I, II, and III (Fomproix and Percipalle 2004; Philimonenko et al. 2004; Hofmann et al. 2004; Hu et al. 2004; Kukalev et al. 2005). The actin – polymerase interaction has not been characterized at the structural level. Three Pol III subunits have been detected in association with actin in coimmunoprecipitation experiments (Hu et al. 2004). It has been proposed that two of them, RPABC2 and RPABC3, build the actin-binding site, because these subunits are common to all three polymerases and are located close to each other at the surface of the polymerase (Cramer et al. 2001; Hu et al. 2004). Coimmunoprecipitation experiments also showed that actin can interact with the largest Pol I subunit and with the carboxy-terminal domain (CTD) of the largest Pol II subunit (Fomproix and Percipalle 2004; Kukalev et al. 2005), which provides additional actin-binding sites specific for these polymerases. The interaction of actin with the RNA polymerases is functionally relevant. Anti-actin antibodies inhibit transcription by Pol I and Pol II in vivo and in vitro (Hofmann et al. 2004; Philimonenko et al. 2004), and actin restores the full activity of partially purified, inactive Pol III preparations in vitro (Hu et al. 2004). These observations indicate that actin associates with RNA polymerases that are engaged in transcription.

Nuclear Functions of Actin

The inhibitory effect of anti-actin antibodies on Pol I and II, as well as the ability of actin to activate Pol III, have been observed in vitro on naked DNA (Hofmann et al. 2004; Philimonenko et al. 2004; Hu et al. 2004), which suggests that actin is involved in basal transcription independently of its role in chromatin regulation. These observations have led to the proposal that actin is required for the activity of the basal transcription machinery. However, we are far from understanding how actin contributes to the transcription process and whether all three RNA polymerases use actin in the same manner. A relevant question is whether actin is required for transcription initiation or elongation. Actin occupancy at promoters of Pol I and Pol II genes and copurification of actin with preinitiation complexes (PIC) suggest that actin is required for the assembly of transcription-competent polymerases (Hofmann et al. 2004; Philimonenko et al. 2004; Percipalle et al. 2006; Obrdlik et al. 2008; Louvet and Percipalle 2009). However, in vitro rDNA transcription assays that discriminate between transcription initiation (defined operationally by the synthesis of the initial trinucleotide) and elongation show that actin and nuclear myosin 1 (NM1) are needed for Pol I transcription elongation (Philimonenko et al. 2004; Percipalle et al. 2006). An important contribution toward understanding the role of actin in association with the transcription machinery was provided by the finding that NM1 is also intimately associated with Pol I and Pol II (Pestic-Dragovich et al. 2000; Fomproix and Percipalle 2004; Hofmann et al. 2006a). NM1 is a short-tailed monomeric myosin that acts as an actindependent ATPase. NM1 was found on active nucleolar transcription sites of both interphase and mitotic HeLa cells (Fig. 1) (Fomproix and Percipalle 2004), and microinjection of anti-NM1 antibodies into living cells inhibited rRNA synthesis (Philimonenko et al. 2004). Furthermore, depletion of either actin or NM1 from transcription extracts repressed the ability of the extracts to support transcription in vitro (Hofmann et al. 2004; Philimonenko et al.

2004). The facts that endogenous actin and NM1 associate with promoter and coding regions of rRNA genes, that actin and NM1 are associated with the transcription machinery, and that impairment of the myosin ATPase activity down-regulates Pol I transcription led to the proposal that an actomyosin activity is required for Pol I transcription (Fomproix and Percipalle 2004; Philimonenko et al. 2004; Percipalle et al. 2006). A recent study by Ye and coworkers (2008) provided further experimental support to this hypothesis. Actin mutants that are deficient in polymerization failed to associate with Pol I and did not support Pol I transcription. NM1 mutants that were defective in ATP-binding, actin-binding, or calmodulinbinding failed to associate with the rDNA and with Pol I and finally, the association of actin and NM1 with Pol I was regulated by ATP hydrolysis (Ye et al. 2008). These observations have enabled a model to be proposed in which the actin-NM1 complex acts as a molecular motor that helps the transcription machinery slide along the rDNA (Ye et al. 2008). Such a motor should interact with the transcription machinery itself and with the DNA, directly or indirectly (Fig. 2A). The actin-Pol I interaction is well-documented. The direct interaction between NM1 and DNA in vitro (Hofmann et al. 2006b) might explain how the actin-NM1 complex fastens to the DNA. ACTIN IN NASCENT TRANSCRIPTS

Actin not only interacts with the transcription machineries, it also associates with the nascent transcripts (Percipalle et al. 2001; Percipalle et al. 2002). Immunohistochemistry experiments performed in the dipteran C. tentans provided initial insights into the cotranscriptional binding of actin to nascent transcripts, because antibodies to actin labeled many loci in polytene chromosome preparations but failed to label after treatment of the chromosomes with RNase A (Percipalle et al. 2001). Immunoelectron microscopy experiments on ultrathin sections of C. tentans salivary glands showed that actin was preferentially associated with the distal region of the active transcription unit,

N. Visa and P. Percipalle

Actin

Fibrillarin

Merge

NM1

Fibrillarin

Merge

NM1 pre-serum

Fibrillarin

Merge

Figure 1. Actin and NM1 localize to mammalian nucleoli. Homogeneous preparations of nucleoli from HeLa

cells were analyzed by immunofluorescence labeling and confocal microscopy using rabbit polyclonal antibodies to actin (scale bar, 2 mm) and NM1 (scale bar, 2 mm) or corresponding preimmune sera (scale bar, 1 mm). All the specimens were coimmunostained with a mouse monoclonal antibody to fibrillarin as marker for nucleoli.

away from the promoter (Percipalle et al. 2001). Later on, anti-actin antibodies coprecipitated coding regions of Pol I and Pol II genes in chromatin immunoprecipitation experiments (Hofmann et al. 2004; Philimonenko et al. 2004; Obrdlik et al. 2008), emphasizing the presence of actin along active genes. The connection between actin and RNAwas further shown at the molecular level by chromatin RNA immunoprecipitation assays, in which it was possible to analyze the cotranscriptional association of protein factors with nascent RNA (Obrdlik and Percipalle 2009; Obrdlik et al. 2008). Specific heterogeneous nuclear ribonucleoproteins (hnRNPs) bind actin and mediate its association with RNA. Actin-associated hnRNP proteins were discovered mainly by resolving

nuclear extracts or RNP preparations with DNase I affinity chromatography. In both C. tentans and mammals, a significant fraction of actin-associated hnRNPs belong to the A/B type family. The hnRNP A/B proteins contain two conserved RNA recognition motifs (RRMs) flanked at the amino terminus by an acidic domain and at the carboxyl terminus by a divergent module for protein– protein interactions, termed the “auxiliary domain” (Krecic and Swanson 1999; Dreyfuss et al. 2002). In C. tentans, actin is associated with the shuttling mRNA-binding protein hrp36 (Visa et al. 1996), whereas in mammals, in which the repertoire of A/B-type hnRNPs is somewhat larger (Krecic and Swanson 1999; Dreyfuss et al. 2002), actin associates with several proteins, including

Nuclear Functions of Actin

A

5¢

5¢

NM 1

Chromatin remodeling complex Actin Actin Actin Actin

Pol Actin

NM 1

Actin

Pol

Actin Actin

B

5¢

hnRNP proteins

5¢

Chromatin Actin remodeling complex 65/U HAT

5¢

Pol

Pol

Pol

Figure 2. Two possible ways by which actin contributes to transcription elongation. (A) An actomyosin motor

helps the transcription machinery slide along the DNA. The architecture of the motor is not well understood but we envisage that the actin-NM1 complex makes contact with both the transcription machinery and the DNA/ chromatin. Two possible configurations are depicted in the figure. (B) Actin participates in the recruitment of histone modifiers to protein-coding genes. Actin binds to hnRNP proteins and becomes incorporated into the nascent pre-mRNPs. Actin forms a complex with specific hnRNP adaptor proteins such as Hrp65 in Chironomus tentans and hnRNP U in mammals, and recruits HATs that acetylate histones and facilitate transcription elongation. The HATs might be components of larger chromatin remodeling complexes that might also establish direct contacts with actin.

hnRNP A2, hnRNPA3, and the transport mediator CArG-box binding factor (CBF-A) (Percipalle et al. 2002; Raju et al. 2008). Another actin-associated hnRNP protein of C. tentans, hrp65, shares strong homology with the human proteins PSF (Patton et al. 1993), p54nrb/NonO (Dong et al. 1993), PSP1 (Fox et al. 2002), and with the Drosophila NonA/BJ6 (Jones and Rubin 1990; von Besser et al. 1990). These proteins do not belong to the A/B-type and they are characterized by a central domain of about 320 amino acids

termed “DBHS” (Drosophila behavior and human splicing). This domain is evolutionarily conserved. The DBHS domain comprises two RRMs and an additional carboxy-terminal stretch of approximately 100 amino acids that mediates dimerization or oligomerization among DBHS proteins (Kiesler et al. 2003). The association of DBHS proteins with actin is conserved in mammals; the human PSF and NonO proteins have been identified as components of a large nuclear protein complex together with actin, Pol II, and N-WASP, a key

N. Visa and P. Percipalle

regulator of cytoplasmic microfilaments that is also present in the cell nucleus (Wu et al. 2006). Actin associates with hnRNP U/SAF-A (Scaffold attachment factor A), a large 120kDa tripartite protein with ATP-binding activity in mammals. The amino terminus of this protein displays DNA-binding activity, the carboxyl terminus contains two RRMs for RNA binding, and the central domain displays Pol II-binding activity (Kukalev et al. 2005). Some of the actin-associated hnRNP proteins identified so far interact directly with actin, whereas others are part of actin-containing complexes and are not in direct contact with actin. The C. tentans hrp65-2 (hrp65 isoform 2) and the human hnRNP U proteins interact with actin through a conserved motif (Percipalle et al. 2003; Kukalev et al. 2005). Remarkably, inhibition of the actin-hrp65-2 or actin-hnRNP U interactions in living cells inhibits transcription elongation by Pol II as monitored by incorporation of labeled nucleotide precursors in run-on assays (Percipalle et al. 2003; Kukalev et al. 2005). Based on these results, we proposed a regulatory mechanism in which actin facilitates the cross talk between chromatin, elongating polymerase, and nascent RNA (Louvet and Percipalle 2009). How do actin-hnRNP complexes facilitate transcription elongation? Experiments in insects and mammals suggest that specific actinhnRNP complexes facilitate the elongation phase by recruiting transcriptional coactivators to remodel the nucleosome barrier imposed by chromatin (Percipalle and Visa 2006; Miralles and Visa 2006; Louvet and Percipalle 2009). Actin-hrp65-2 and actin-hnRNP U complexes mediate the recruitment of the HATs p2D10 and PCAF, respectively, to transcribed genes (Fig. 2B) (Sjo¨linder et al. 2005; Obrdlik et al. 2008). The disruption of these actin-hnRNP interactions in vivo reduced the level of histone acetylation in chromatin, blocked Pol II transcription, and affected HAT occupancy along active genes (Sjo¨linder et al. 2005; Kukalev et al. 2005; Obrdlik et al. 2008). These results indicate that the integrity of specific actin-hnRNP

interactions is required for coactivator recruitment to active genes. The topology of the actin complex is not known. However, an antibody specific for the actin-binding site on hnRNP U coprecipitates endogenous hnRNP U and PCAF but fails to precipitate actin in mammals (Obrdlik et al. 2008). This leads us to favor the idea that hnRNP U is able to interact simultaneously with both PCAF and actin. There is also recent evidence that the HAT PCAF and BRG1, the ATPase subunit of the SWI/ SNF chromatin remodeling complex, are sequentially recruited to the promoter of the myogenin gene and are both involved in gene activation (Li et al. 2007). Actin is also a component of the SWI/SNF complex (reviewed in Farrants 2008). We speculate, therefore, that during elongation of nascent pre-mRNA, the actin-RNP complexes work as modules that facilitate recruitment of HATs and chromatin remodeling complexes to active genes (see Fig. 2B). Studies by Obrdlik and coworkers (2008) revealed that the interplay between actin and the nascent RNP also involves interaction with the RNA polymerase in some cases. During elongation, the carboxy-terminal domain (CTD) of the largest subunit of Pol II is subjected to a series of covalent modifications that modulate recruitment of transcriptional coactivators. The amino acid residues Ser2 and Ser5 within the conserved CTD heptapeptide repeats become combinatorially phosphorylated and the different phosphorylation states represent hallmarks for individual transcription steps (Meinhart et al. 2005). Therefore, if specific actin-hnRNP complexes are required for efficient elongation, it is likely that they are cotranscriptionally associated with specific CTD phosphorylation states. Recent evidence shows that this is the case. Actin together with hnRNP U associates only with Ser2-phosphorylated heptapeptide repeats or with heptapeptide repeats that are phosphorylated on both Ser5 and Ser2, and fails to interact with unphosphorylated heptapeptide repeats or with those repeats that are phosphorylated on Ser5 only (Obrdlik et al. 2008). These findings

Nuclear Functions of Actin

support the idea that specific actin-hnRNP complexes physically associate with the elongating Pol II, a view that is corroborated by evidence that PSF and p54nrb, mammalian homologs of the C. tentans hrp65 protein, also interact with actin and with the CTD (Emili et al. 2002; Wu et al. 2006). As discussed earlier, actin and NM1 are required for transcription elongation by Pol I, and NM1 also appears to mediate the association of specific chromatin-modifying components to the rDNA genes. NM1 is a core component of the B-WICH chromatin remodeling complex together with the Williams syndrome transcription factor (WSTF) and the ATPase SNF2 h (Percipalle et al. 2006), and this complex turned out to also contain precursor rRNA (Cavellan et al. 2006). Remarkably, inhibition of endogenous NM1 or WSTF in abortive initiation assays did not affect the formation of initial pre-rRNA transcripts but affected the synthesis of the entire transcript when chromatin templates were used in run-on transcription assays (Philimonenko et al. 2004; Percipalle et al. 2006). These results suggest that the core components of the B-WICH complex, including NM1, are required after transcription initiation, and a model has been proposed in which the dynamic interaction between actin and NM1 is required for local recruitment of the B-WICH complex to active ¨ stlund Farrants 2006; genes (Percipalle and O Louvet and Percipalle 2009). The previous considerations suggest that actin modulates the elongation of nascent pre-mRNAs and pre-rRNAs by controlling the cotranscriptional association of chromatinmodifying components with the active genes. This view is of particular interest in the case of Pol I transcription, where active rRNA genes are believed to be devoid of nucleosomes. For protein-coding genes, it is intriguing that actin together with hnRNP U and the HAT PCAF associate with the CTD in a phosphorylated Ser2dependent manner (Obrdlik et al. 2008), an observation that strengthens the hypothesis that the coupling of actin to the elongating Pol II facilitates the modifications of chromatin structure.

ACTIN BEYOND TRANSCRIPTION

The studies performed in C. tentans showed that actin is cotranscriptionally added to nascent pre-mRNA, remains incorporated in mature RNPs released into the nucleoplasmic milieu, and accompanies the mRNA all the way to polysomes (Percipalle et al. 2001). There is no evidence of nuclear actin-based motors promoting directional movement of RNP particles or even preribosomal subunits (Singh et al. 1999; Politz et al. 2003; Siebrasse et al. 2008). However, actin is associated with mature mRNPs and with pre-60S subunits, it decorates pore-linked filaments that project inward into the nucleoplasm from the nuclear pore basket, and it has been implicated in controlling the export of viral mRNA transcripts (Percipalle et al. 2001, 2002; Hofmann et al. 2001; Kiseleva et al. 2004; Oeffinger et al. 2007). Also, NM1 is loaded into preribosomes and involved in the export-competent pre-60S subunits (Obrdlik et al. 2009). In addition, NM1 accompanies preribosomal subunits in transit to the nuclear envelope (Cisterna et al. 2007; Obrdlik et al. 2009), is present at the nuclear envelope (Holaska et al. 2007), and has been recently incorporated in the export of small ribosomal subunits (Cisterna et al. 2009). Finally, there is recent evidence that a complex containing rRNA and NM1 is associated with the RNA-binding protein Nup153 at the nuclear pore (Obrdlik et al. 2009), which suggests that NM1 accompanies the rRNA from the gene to the nuclear pore complex. These observations suggest that the synergy between actin and NM1 plays a more general role, and is not limited to the gene. We favor the idea that actin and NM1 contribute to the maturation and assembly of export-competent mRNPs and rRNPs by promoting the remodeling of RNP structures through dynamic protein–protein interactions (reviewed in Percipalle et al. 2009).

REGULATION OF ACTIN POLYMERIZATION

The presence of actin in the cell nucleus raises the possibility that there are nuclear-based mechanisms regulating actin polymerization

N. Visa and P. Percipalle

(Pederson 2008; Louvet and Percipalle 2009). It has been suggested that nuclear actin coexists in monomeric (G) or short oligomeric forms and in a polymeric form (Louvet and Percipalle 2009; Gieni & Hendzel 2009). It is unclear how transitions between these forms occur, but dynamic alterations in actin polymerization may be instrumental in modulating the function of actin in RNA biogenesis. Canonical actin filaments, which are commonly seen in the cytoplasm by immunofluorescence methods, have not been convincingly visualized in the cell nucleus. However, many of the factors that are known to control cytoplasmic actin polymerization and stabilize filamentous F-actin structures have been detected in the cell nucleus. N-WASP and the ARP2/3 complexes cooperate in the nucleation and branching of new actin filaments. These proteins are present in the cell nucleus, where they are implicated in the elongation of nascent mRNA transcripts (Wu et al. 2006; Yoo et al. 2007). More recently, other proteins that facilitate cytoplasmic actin nucleation independently of N-WASP and ARP2/3 have been discovered in the nucleus, including the forminlike mDia proteins and JMY (Miki et al. 2008; Zuchero et al. 2009). These findings indeed suggest the presence of polymeric actin in the nucleus, and it is likely that there are several independent mechanisms that regulate nuclear actin polymerization, as is the case in the cytoplasm. It is possible that the need to have several mechanisms for actin polymerization reflects the multiple roles that actin plays in the cell nucleus. This may or may not be the case, but it is clear that nuclear actin assembled into higher order structures is rather dynamic with a high turnover (McDonald et al. 2006), which indicates that a pool of monomeric actin is also present in the nucleus. This pool is probably necessary to feed dynamic polymeric actin structures. The existence of monomeric actin was indirectly suggested by the discovery of G-actin binding proteins such as cofilin, profilin, b-thymosin, and a gelsolin-like protein in the cell nucleus (Prendergast et al. 1991; Pendleton et al. 2003; Skare et al. 2003; Huff et al. 2004). Actin can be copurified with hnRNP U and

the Pol II machinery using DNase I affinity chromatography (Kukalev et al. 2005; Obrdlik et al. 2008), which has higher affinity for monomeric than for filamentous actin (Zechel 1980). In addition, monoclonal antibodies designed to recognize epitopes exclusively available in monomeric actin or actin dimers revealed significant immunostaining of the cell nucleus (reviewed in Jokusch et al. 2006). There is also evidence that monomeric or short oligomeric actin is directly involved in nuclear functions such as the role of monomeric actin in the regulation of SRF-dependent transcription in complex with the coactivator MAL (Vartiainen et al. 2007). The G-actin binding protein profilin affects transcription of rRNA genes, and drugs that inhibit actin polymerization, such as cytochalasin D and latrunculin B, inhibit the transcription of rDNA genes in vivo and in vitro, whereas drugs that favor actin assembly do not influence rDNA transcription (Ye et al. 2008). Furthermore, a recent study indicated that actin polymerization takes place in retinoic acid (RA)-induced HoxB transcription (Ferrai et al. 2009). These observations suggest that polymeric actin is functional in the cell nucleus. Because N-WASP and ARP2/3 are important during premRNA elongation, and an actin-NM1 complex is clearly implicated in transcription elongation by Pol I, we favour the idea that both monomeric and polymeric actin are present along active genes, and that cotranscriptional transitions in the actin polymerization states result in the existence of different actin forms associated with different parts of the gene. What is the polymerization state of actin in RNPs? Ultrastructural analysis has not revealed canonical filamentous actin structures attached to mature RNPs (reviewed in Percipalle 2009), which hints at an intrinsic mechanism that keeps actin from polymerizing as soon as it is assembled into RNPs. An equally intriguing possibility is that actin undergoes covalent modifications that do not favor the establishment of polymers. Some of these ideas are speculative, but they emphasize the importance of regulating actin polymerization for nuclear function.

Nuclear Functions of Actin

LONG-RANGE MOVEMENT

Most of our current knowledge of nuclear actin originates from research in the field of chromatin regulation and transcription. However, recent studies point to the possibility that actin plays a role also in the large-scale organization of the genome. The eukaryotic nucleus is highly compartmentalized and, in spite of the lack of membrane-bound organelles, most nuclear components show specific distributions restricted to certain nuclear domains (reviewed by Lamond and Spector 2003; Schneider and Grosschedl 2007). The functional compartmentalization of the nucleus is reflected in the fact that different types of sequences occupy different positions in the nucleus. For example, gene-rich chromosome regions tend to occupy internal positions, whereas gene-poor sequences are associated with the nuclear periphery, and repressed genes are often associated with heterochromatic regions, whereas active genes are usually located near nuclear speckles (reviewed by Spector 2003). What is most interesting is that the position of a gene locus can change in response to transcriptional activation (Parada and Misteli 2002) and that the transcriptional activity of a gene is affected by its position in the nucleus (Finlan et al. 2008). Chuang and coworkers (2006) visualized the dynamics of gene repositioning by constructing cell lines that carry tandem gene arrays under the control of an inducible promoter. When the expression of the array was activated, the entire locus relocated from the cell periphery to the nuclear interior. Unidirectional trajectories were recorded over distances of 1 – 5 mm, which suggests that an active, directed mechanism for long-range chromatin movement is present. Interestingly, the expression of mutant actin or NM1 with defects in polymerization and actin-binding inhibited the relocation of the array on transcriptional activation (Chuang et al. 2006). Dundr and coworkers (2007) used a similar experimental design based on the use of synthetic inducible arrays to study the association of Cajal bodies (CBs) with U2 snRNA genes in human cells. After transcription had been induced, the U2 snRNA gene array moved

over long distances inside the nucleus (2 – 3 mm) and became stably associated with CBs. Also in this case, the expression of a nonpolymerizable actin mutant inhibited the movement of the U2snRNA locus (Dundr et al. 2007). In a third study, Hu and coworkers identified long-range, estrogen-induced interactions among genes located in different chromosomes. The interactions were lost after treatment of the cells with drugs that either blocked actin polymerization or inhibited actin depolymerization. Depletion of NM1 by RNAi or nuclear injection of antibodies against NMI also blocked the estrogen-induced interactions (Hu et al. 2008). Moreover, the inhibitory effect of the anti-NM1 antibodies could be reversed by the expression of wild-type NM1, but not by the expression of NM1 mutants that were defective in actin binding or lacked ATPase activity (Hu et al. 2008). The results reveal that the dynamics of actin in the cell nucleus affect gene positioning and large-scale chromatin organization, in a direct or indirect manner. The molecular mechanisms underlying these long-range chromatin movements are still unclear. The type of movement observed could be explained by active transport mechanisms based on actomyosin motors. This would imply the existence of mechanisms for the regulation of actin dynamics at micrometer scale inside the nucleus. If this is the case, further work must characterize such an actin large-scale regulatory network and elucidate the mechanisms that impose directionality inside the nucleus. Alternatively, and considering the fact that the observed relocations take place after transcription activation, the actin dependence could be at the transcriptional level and the chromatin movements could be a consequence of chromatin decondensation. This model, however, does not provide a satisfactory explanation of the fact that the U2 snRNA genes are targeted to relatively stable positions in the nucleus, nor does it provide any basis for the directionality of the movements. Further work is needed to understand the molecular mechanisms of large-scale chromatin relocation and to answer the questions raised by the discovery of gene relocation events. We anticipate that the

N. Visa and P. Percipalle

answers to these questions will provide a fundamental advance in our understanding of the dynamics of the eukaryotic genome.

activity of actin in gene expression is also sensitive to extracellular stimuli.

ACKNOWLEDGMENTS CONCLUDING REMARKS

There is an increasing body of evidence that supports the idea that actin plays a role in controlling multiple phases of gene transcription as a component of chromatin-remodeling complexes and RNP particles, and that it is closely associated with all RNA polymerases. Two possible models by which actin may regulate transcription elongation are illustrated in Figure 2. One model proposes the existence of an actin-NM1 motor, whereas the other one emphasizes the role of actin-RNP complexes in recruiting chromatin modifiers to transcribed genes. Both models are based on solid experimental data and are not mutually exclusive. We do not know whether both mechanisms act in all types of genes. Support for the existence of an actin-NM1 motor comes mostly from studies of Pol I transcription, whereas the recruitment of HATs mediated by RNPs has been mostly documented for protein-coding genes. It is possible that the way in which actin acts in transcription is different for different types of genes. After many years of debate about the existence of actin in the nucleus, it is now clear that actin is an abundant nuclear protein and many independent studies support the idea that it is involved in gene expression. An interesting question is whether the nuclear functions of actin are linked to—or regulated by—the dynamics of actin in the cytoplasm. The actin cytoskeleton is highly dynamic and reacts to a variety of extracellular signals (for recent reviews, see Moustakas and Heldin 2008; Papakonstanti and Stournaras 2008), and there is evidence that the cytoplasmic and actin pools are not independent of each other (Vartiainen et al. 2007). It is therefore tempting to suggest that actin acts as a sensor of extracellular signals with the ability to transduce the signals to the genome and modulate gene expression. The question remains whether the overall

¨ stlund Farrants for We thank Ann-Kristin O critical reading of the manuscript and George Farrants for language editing. Our work is supported by grants from the Swedish Research Council (Vetenskapsra˚det), the Swedish Cancer Society (Cancerfonden), and the European Science Foundation (Eurocores Programme RNAQuality).

REFERENCES Blessing CA, Ugrinova GT, Goodson HV. 2004. Actin and ARPs: Action in the nucleus. Trends Cell Biol 14: 435 –442. ¨ stlund Farrants AK. 2006. Cavellan E, Asp P, Percipalle P, O The chromatin remodelling complex WSTF-SNF2 h interacts with several nuclear proteins in transcription. J Biol Chem 281: 16264– 16271. Chen M, Shen X. 2007. Nuclear actin and actin-related proteins in chromatin dynamics. Curr Opin Cell Biol 19: 326 –330. Chuang CH, Carpenter AE, Fuchsova B, Johnson T, de Lanerolle P, Belmont AS. 2006. Long-range directional movement of an interphase chromosome site. Curr Biol 16: 825– 831. Cisterna B, Malatesta M, Dieker J, Muller S, Prosperi E, Biggiogera M. 2009. An active mechanism flanks and modulates the export of the small ribosomal subunits. Histochem Cell Biol 131: 743– 753. Cisterna B, Necchi D, Prosperi E, Biggiogera M. 2006. Small ribosomal subunits associate with nuclear myosin and actin in transit to the nuclear pores. FASEB J 20: 1901–1903. Cramer P, Bushnell DA, Kornberg RD. 2001. Structural basis of transcription: RNA polymerase II at 2.8 angstrom resolution. Science 292: 1863– 1876. Dong B, Horowitz DS, Kobayashi R, Krainer AR. 1993. Purification and cDNA cloning of HeLa cell p54nrb, a nuclear protein with two RNA recognition motifs and extensive homology to human splicing factor PSF and Drosophila NONA/BJ6. Nucl Acids Res 21: 4085–4092. Dreyfuss G, Kim VN, Kataoka N. 2002. Messenger-RNAbinding proteins and the messages they carry. Nat Rev Mol Cell Biol 3: 195–205. Dundr M, Ospina JK, Sung M-H, John S, Upender M, Reid T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo, J. Cell Biol 179: 1095–1103. Egly JM, Miyamoto NG, Moncollin V, Chambon P. 1984. Is actin a transcription initiation factor for RNA polymerase B? EMBO J 3: 2363– 2371.

Nuclear Functions of Actin Emili A, Shales M, McCracken S, Xie W, Tucker PW, Kobayashi R, Blencowe BJ, Ingles CJ. 2002. Splicing and transcription-associated proteins PSF and p54nrb/nonO bind to the RNA polymerase II CTD. RNA 8: 1102– 1111. Farrants AK. 2008. Chromatin remodelling and actin organisation. FEBS Lett 582: 2041–2050. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. 2008. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet 4: e1000039. Fomproix N, Percipalle P. 2004. An actin-myosin complex on actively transcribing genes. Exp Cell Res 294: 140 –148. Fox AH, Lam YW, Leung AK, Lyon CE, Andersen J, Mann M, Lamond AI. 2002. Paraspeckles: A novel nuclear domain. Curr Biol 12: 13–25. Ferrai C, Naum-Onganı´a G, Longobardi E, Palazzolo M, Disanza A, Diaz VM, Crippa MP, Scita G, Blasi F. 2009. Induction of HoxB transcription by retinoic acid requires actin polymerization. Mol Biol Cell May 28. [Epub ahead of print]. Gieni RS, Hendzel MJ. 2009. Actin dynamics and functions in the interphase nucleus: Moving toward an understanding of nuclear polymeric actin. Biochem Cell Biol 87: 283 –306. Harata M, Oma Y, Mizuno S, Jiang YW, Stillman DJ, Wintersberger U. 1999. The nuclear actin-related protein of Saccharomyces cerevisiae, Act3p/Arp4, interacts with core histones. Mol Biol Cell 10: 2595–2605. Hofmann WA, Johnson T, Klapczynski M, Fan J-L, de Lanerolle P. 2006b. From transcription to transport: Emerging roles for nuclear myosin I. Biochem Cell Biol 84: 418 –426. Hofmann WA, Reichart B, Ewald A, Muller E, Schmitt I, Stauber RH, Lottspeich F, Jockusch BM, Scheer U, Hauber J, et al. 2001. Cofactor requirements for nuclear export of Rev response element (RRE)- and constitutive transport element (CTE)-containing retroviral RNAs. An unexpected role for actin. J. Cell Biol 152: 895– 910. Hofmann WA, Stojiljkovic L, Fuchsova B, Vargas GM, Mavrommatis E, Philimonenko V, Kysela K, Goodrich JA, Lessard JL, Hope TJ, et al. 2004. Actin is part of preinitiation complexes and is necessary for transcription by RNA polymerase II. Nat Cell Biol 6: 1094–1101. Hofmann WA, Vargas GM, Ramchandran R, Stojiljkovic L, Goodrich JA, de Lanerolle P. 2006a. Nuclear myosin I is necessary for the formation of the first phosphodiester bond during transcription initiation by RNA polymerase II. J Cell Biochem 99: 1001–1009. Holaska JM, Wilson KL. 2007. An emerin “proteome”: Purification of distinct emerin-containing complexes from HeLa cells suggests molecular basis for diverse roles including gene regulation, mRNA splicing, signaling, mechanosensing, and nuclear architecture. Biochemistry 46: 8897–8908. Hu P, Wu S, Hernandez N. 2004. A role for b-actin in RNA polymerase III transcription. Genes Dev 18: 3010– 3005. Hu Q, Kwon YS, Nunez E, Cardamone MD, Hutt KR, Ohgi KA, Garcia-Bassets I, Rose DW, Glass CK, Rosenfeld MG, et al. 2008. Enhancing nuclear receptor-induced transcription requires nuclear motor and LSD1-dependent gene networking in interchromatin granules. Proc Natl Acad Sci 105: 19199– 19204.

Huff T, Rosorius O, Otto AM, Muller CS, Ballweber E, Hannappel E, Mannherz HG. 2004. Nuclear localisation of the G-actin sequestering peptide thymosin b4. J Cell Sci 117: 5333– 5341. Jockusch BM, Schoenenberger CA, Stetefeld J, Aebi U. 2006. Tracking down the different forms of nuclear actin. TRENDS Cell Biol 16: 391–396. Jones KR, Rubin GM. 1990. Molecular analysis of no-on-transient A, a gene required for normal vision in Drosophila. Neuron 4: 711 –723. Kiesler E, Miralles F, Ostlund Farrants AK, Visa N. 2003. The Hrp65 self-interaction is mediated by an evolutionarily conserved domain and is required for nuclear import of Hrp65 isoforms that lack a nuclear localization signal. J Cell Sci 116: 3949– 3956. Kiseleva E, Drummond SP, Goldberg MW, Rutherford SA, Allen TD, Wilson KL. 2004. Actin- and protein-4.1containing filaments link nuclear pore complexes to subnuclear organelles in Xenopus oocyte nuclei. J Cell Sci 117: 2481– 2490. Krecic AM, Swanson MS. 1999. hnRNP complexes: Composition, structure, and function. Curr Opin Cell Biol 11: 363 –371. Kukalev A, Nord Y, Palmberg C, Bergman T, Percipalle P. 2005. Actin and hnRNP U cooperate for productive transcription by RNA polymerase II. Nat Struct Mol Biol 12: 238 –244. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605– 612. Li ZY, Yang J, Gao X, Lu JY, Zhang Y, Wang K, Cheng MB, Wu NH, Zhang Y, Wu Z, et al. 2007. Sequential recruitment of PCAF and BRG1 contributes to myogenin activation in 12-O-tetradecanoylphorbol-13-acetate-induced early differentiation of rhabdomyosarcoma-derived cells. J Biol Chem 282: 18872– 18878. Louvet E, Percipalle P. 2009. Actin and myosin in gene transcription. Int Rev Cell Mol Biol 272: 107–147. McDonald D, Carrero G, Andrin C, de Vries G, Hendzel MJ. 2006. Nucleoplasmic b-actin exists in a dynamic equilibrium between low-mobility polymeric species and rapidly diffusing populations. J Cell Biol 172: 541– 552. Meinhart A, Kamenski T, Hoeppner S, Baumli S, Cramer P. 2005. A structural perspective of CTD function. Genes Dev 19: 1401–1415. Miki T, Okawa K, Sekimoto T, Yoneda Y, Watanabe S, Ishizaki T, Narumiya S. 2008. mDia2 shuttles between the nucleus and the cytoplasm through the importin-{a}/ {b}- and CRM1-mediated nuclear transport mechanism. J Biol Chem 284: 5753–5762. Miralles F, Posern G, Zaromytidou AI, Treisman R. 2003. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113: 329 –342. Moustakas A, Heldin CH. 2008. Dynamic control of TGF-b signaling and its links to the cytoskeleton. FEBS Lett 582: 2051–2065. Obrdlik A, Kukalev A, Louvet E, Farrants AK, Caputo L, Percipalle P. 2008. The histone acetyltransferase PCAF associates with actin and hnRNP U for RNA polymerase II transcription. Mol Cell Biol 28: 6342–6357. Obrdlik A, Louvet E, Kukalev A, Naschekin D, Kiseleva E, Fahrenkrog B, Percipalle P. 2009. Nuclear myosin 1 is in

N. Visa and P. Percipalle complex with mature rRNA transcripts and associates with the nuclear pore complex. FASEB J, 2009 Sept 3 [Epub ahead of print] PMID: 19729515. Oeffinger M, Wei KE, Rogers R, DeGrasse JA, Chait BT, Aitchison JD, Rout MP. 2007. Comprehensive analysis of diverse ribonucleoprotein complexes. Nat Methods 4: 951–956. Olave IA, Reck-Peterson SL, Crabtree GR. 2002. Nuclear actin and actin-related proteins in chromatin remodelling. Annu Rev Biochem 71: 755–781. Papakonstanti EA, Stournaras C. 2008. Cell responses regulated by early reorganization of actin cytoskeleton. FEBS Lett 582: 2120– 2127. Parada L, Misteli T. 2002. Chromosome positioning in the interphase nucleus. Trends Cell Biol 12: 425– 432. Patton JG, Porro EB, Galceran J, Tempst P, Nadal-Ginard B. 1993. Cloning and characterization of PSF, a novel pre-mRNA splicing factor. Genes Dev 7: 393– 406. Pederson T. 2008. As functional nuclear actin comes into view, is it globular, filamentous, or both? J Cell Biol 180: 1061–1064. Pederson T, Aebi U. 2002. Actin in the nucleus: what form and what for? J Struct Biol 140: 3– 9. Pendleton A, Pope B, Weeds A, Koffer A. 2003. Latrunculin B or ATP depletion induces cofilin-dependent translocation of actin into nuclei of mast cells. J Biol Chem 278: 14394–14400. Percipalle P. 2009. The long journey of actin and actin-associated proteins from gene to polysomes. Cell Mol Life Sci 66: 2151–2165. Percipalle P, Obrdlik A. 2009. Analysis of nascent RNA transcripts by chromatin RNA immunoprecipitation. Meth Mol Biol 567: 215– 235. Percipalle P, Visa N. 2006. Molecular functions of nuclear actin in transcription. J Cell Biol 172: 967– 971. ¨ stlund Farrants AK. 2006. Chromatin remodPercipalle P, O elling and transcription: Be-WICHed by nuclear myosin 1. Curr Opin Cell Biol 18: 267 –274. Percipalle P, Fomproix N, Cavella´n E, Voit R, Reimer G, Kru¨ger T, Thyberg J, Scheer U, Grummt I, Farrants AK. 2006. The chromatin remodelling complex WSTFSNFh interacts with nuclear myosin 1 and has a role in RNA polymerase I transcription. EMBO Rep 7: 525 –530. Percipalle P, Fomproix N, Kylberg K, Miralles F, Bjorkroth B, Daneholt B, Visa N. 2003. An actin-ribonucleoprotein interaction is involved in transcription by RNA polymerase II. Proc Natl Acad Sci 100: 6475–6480. Percipalle P, Jonsson A, Nashchekin D, Karlsson C, Bergman T, Guialis A, Daneholt B. 2002. Nuclear actin is associated with a specific subset of hnRNP A/B-type proteins. Nucleic Acids Res 30: 1725– 1734. Percipalle P, Zhao J, Pope B, Weeds A, Lindberg U, Daneholt B. 2001. Actin bound to the heterogeneous nuclear ribonucleoprotein hrp36 is associated with Balbiani ring mRNA from the gene to polysomes. J Cell Biol 153: 229–236. Pestic-Dragovich L, Stojiljkovic L, Philimonenko AA, Nowak G, Ke Y, Settlage RE, Shabanowitz J, Hunt DF, Hozak P, de Lanerolle P. 2000. A myosin I isoform in the nucleus. Science 290: 337– 341.

Philimonenko VV, Zhao J, Iben S, Dingova H, Kysela K, Kahle M, Zentgraf H, Hofmann WA, de Lanerolle P, Hozak P, et al. 2004. Nuclear actin and myosin I are required for RNA polymerase I transcription. Nat Cell Biol 6: 1165– 1171. Politz JC, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805–4812. Prendergast GC, Ziff EB. 1991. Mbh 1: a novel gelsolin/ severin-related protein which binds actin in vitro and exhibits nuclear localization in vivo. EMBO J 10: 757– 766. Raju CS, Go¨ritz C, Nord Y, Hermanson O, Lopez-Iglesias C, Visa N, Castelo-Branco G, Percipalle P. 2008. In cultured oligodendrocytes the A/B-type hnRNP CBF-A accompanies MBP mRNA bound to mRNA trafficking sequences. Mol Biol Cell 19: 3008– 3018. Rando OJ, Zhao K, Janmey P, Crabtree GR. 2002. Phospatidylinositol-dependent actin filament binding by the SWI/SNF-like BAF chromatin remodelling complex. Proc Natl Acad Sci 99: 2824– 2829. Scheer U, Hinssen H, Franke WW, Jockusch BM. 1984. Microinjection of actin-binding proteins and actin antibodies demonstrates involvement of nuclear actin in transcription of lampbrush chromosomes. Cell 39: 111 –122. Schneider R, Grosschedl R. 2007. Dynamics and interplay of nuclear architecture, genome organization, and gene expression. Genes Dev 21: 3027–3043. Siebrasse JP, Veith R, Dobay A, Leonhardt H, Daneholt B, Kubitscheck U. 2008. Discontinuous movement of mRNP particles in nucleoplasmic regions devoid of chromatin. Proc Natl Acad Sci 105: 20291–20296. Singh OP, Bjo¨rkroth B, Masich S, Wieslander L, Daneholt B. 1999. The intranuclear movement of Balbiani ring premessenger ribonucleoprotein particles. Exp Cell Res 251: 135–146. ¨ stlund Farrants Sjo¨linder M, Bjo¨rk P, Soderberg E, Sabri N, O AK, Visa N. 2005. The growing pre-mRNA recruits actin and chromatin-modifying factors to transcriptionally active genes. Genes Dev 19: 1871– 1884. Skare P, Kreivi JP, Bergstro¨m A, Karlsson R. 2003. Profilin I colocalizes with speckles and Cajal bodies: A possible role in pre-mRNA splicing. Exp Cell Res 286: 12–21. Spector DL. 2003. The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72: 573–608. Vartiainen MK. 2008. Nuclear actin dynamics– from form to function. FEBS Lett 582: 2033–2040. Vartiainen MK, Guettler S, Larijani B, Treisman R. 2007. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science 316: 1749–1752. Visa N, Alzhanova-Ericsson AT, Sun X, Kiseleva E, Bjo¨rkroth B, Wurtz T, Daneholt B. 1996. A pre-mRNA-binding protein accompanies the RNA from the gene through the nuclear pores and into polysomes. Cell 84: 253– 264. von Besser H, Schnabel P, Wieland C, Fritz E, Stanewsky R, Saumweber H. 1990. The puff-specific Drosophila protein Bj6, encoded by the gene no-on transient A, shows homology to RNA-binding proteins. Chromosoma 100: 37– 47.

Nuclear Functions of Actin Wu X, Yoo Y, Okuhama NN, Tucker PW, Liu G, Guan JL. 2006. Regulation of RNA-polymerase-II-dependent transcription by N-WASP and its nuclear-binding partners. Nat Cell Biol 8: 756– 763. Ye J, Zhao J, Hoffmann-Rohrer U, Grummt I. 2008. Nuclear myosin I acts in concert with polymeric actin to drive RNA polymerase I transcription. Genes Dev 22: 322–330. Yoo Y, Wu X, Guan JL. 2007. A novel role of the actinnucleating Arp2/3 complex in the regulation of RNA polymerase II-dependent transcription. J Biol Chem 282: 7616– 7623.

Zhao K, Wang W, Rando OJ, Xue Y, Swiderek K, Kuo A, Crabtree GR. 1998. Rapid and phosphoinositoldependent binding of the SWI/SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95: 625–636. Zechel K. 1980. Isolation of polymerization competent cytoplasmic actin by affinity chromatography on immobilized DNase I using formamide as eluant. Eur J Biochem 110: 343–348. Zuchero JB, Coutts AS, Quinlan ME, Thangue NB, Mullins RD. 2009. p53-cofactor JMY is a multifunctional actin nucleation factor. Nat Cell Biol 11: 451–459.

The Nucleolus Thoru Pederson Program in Cell and Developmental Dynamics, Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts 01605 Correspondence: [email protected]

When cells are observed by phase contrast microscopy, nucleoli are among the most conspicuous structures. The nucleolus was formally described between 1835 and 1839, but it was another century before it was discovered to be associated with a specific chromosomal locus, thus defining it as a cytogenetic entity. Nucleoli were first isolated in the 1950s, from starfish oocytes. Then, in the early 1960s, a boomlet of studies led to one of the epochal discoveries in the modern era of genetics and cell biology: that the nucleolus is the site of ribosomal RNA synthesis and nascent ribosome assembly. This epistemologically repositioned the nucleolus as not merely an aspect of nuclear anatomy but rather as a cytological manifestation of gene action—a major heuristic advance. Indeed, the finding that the nucleolus is the seat of ribosome production constitutes one of the most vivid confluences of form and function in the history of cell biology. This account presents the nucleolus in both historical and contemporary perspectives. The modern era has brought the unanticipated discovery that the nucleolus is plurifunctional, constituting a paradigm shift.

FIRST SIGHTING

t is likely that some of the few lucky enough to have a microscope in the 18th century saw the nucleolus if they examined thin specimens of tissue in the mode of illumination that later became known as bright field, a century before phase contrast was discovered, for which a Nobel Prize, rare in microscopy, was conferred on Frits Zernike in 1953 (Fig. 1). The first properly documented accounts of the nucleolus were made independently by Wagner (1835) and Valentin (1836, 1839). Beyond occasional studies in which nucleoli were mentioned in passing in the context of broader cytological work, no significant literature on the nucleolus ensued for another half century. Then,

I

a monumental monograph on the nucleolus was published by Montgomery (1898), with an astonishing 346 hand-drawn color figures of nuclei and nucleoli from a vast array of biological material. But as comprehensive and elegant a piece of scholarship as it was, this treatise is primarily known to us today not because of what it revealed the nucleolus to be at the time, but because of what the nucleolus later became. THE NUCLEOLUS BECOMES PART OF THE KARYOTYPE

For three decades after Montgomery’s treatise, there was little momentum until the discovery that the nucleolus arises at a specific chromosomal locus (Heitz 1931; McClintock 1934). While

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved. Advanced Online Article. Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a000638

1

T. Pederson

Figure 1. Looking at the nucleolus. Nucleoli observed in HeLa cells (left) and isolated HeLa cell nuclei (right) by differential interference microscopy. (Images courtesy of David L. Spector [left] and Thoru Pederson [right].)

the intimacy of associations between nucleoli and heterochromatin had previously been noted, the independent findings of Heitz and McClintock established that the nucleolus actually forms at a discrete locus, termed the nucleolus organizer by McClintock. The importance of this advance cannot be overstated because it meant that rather than being mere nuclear anatomy, nucleoli are cytogenetic entities. From this perspective, the astonishing diversity of nucleolar shapes and sizes recorded by Montgomery began to be sensed as reflecting differences in the chromosomal sites at which they arise. The discovery of the nucleolus organizer meant that, whatever its function(s), the nucleolus is stationed at a genetic locus, although it would still be another decade before the discovery that the genetic material is DNA. MONTEVIDEO: A HIGH ALTITUDE VIEW OF THE NUCLEOLUS

We have all attended meetings, and the luckiest of us know that sometimes, when the muses smile, we are witnesses to a renaissance. It just happened that leading up to a meeting on the nucleolus in Montevideo, Uruguay, in December 1965, several new findings had been made or were just about to be submitted for publication. Donald Brown presented a brilliant

experiment, done with John Gurdon (Brown and Gurdon 1964), showing that anucleoate Xenopus embryos arrest in development, plausibly due to the inability of these embryos to make new ribosomes when the maternal stockpile becomes limiting (Fig. 2). Brown also reported the existence of amplified nucleoli in the germinal vesicle (nucleus) of Xenopus oocytes (a finding that had been made contemporaneously [Brown and Dawid 1968; Gall 1968]). Papers on the isolation of nucleoli were given by Walter Vincent (from starfish oocytes), and by Rachele Maggio and Harris Busch (from guinea pig and rat liver, respectively). The biosynthesis of rRNA via large precursor molecules was reported in talks by Joseph Gall, Sheldon Penman, Georgii Georgiev, and Robert Perry. But the most important discovery announced at the meeting was that reported in the talks by Max Birnstiel and Ferrucio Ritossa; namely, that nucleic acid hybridization revealed that DNA complementary to rRNA resides in the nucleolus, which, together with the results of Brown and Gurdon (1964), ushered in a new age in the history of the nucleolus. The proceedings of this conference, including the lively discussion exchanges after each talk (Vincent and Miller 1966), together with an exceptionally insightful synopsis of the meeting (Perry 1966), constitute a definitive archive of this exciting moment in the field.

The Nucleolus

Figure 2. A portal to the modern era: the dependence of ribosomal RNA synthesis on the nucleolus. Anucleolate

(left) or wild-type (right) Xenopus laevis embryos at the neurula stage were incubated with C14-labeled carbon dioxide and RNA was extracted 20 hours later, when both groups of embryos were still morphologically and physiologically indistinguishable. Shown are sucrose gradient sedimentation profiles of 28S and 18S rRNA and transfer RNA. This finding was a keystone in establishing the role of the nucleolus in the biosynthesis of ribosomes. (Reproduced from Brown DC and Gurdon JB 1964. Proc Natl Acad Sci USA 51: 139– 146, by kind permission of Donald D. Brown, Carnegie Institution for Science, Washington, D.C.)

COMING DOWN FROM THE MOUNTAIN

Oscar Miller had presented EM pictures of spread nucleolar “cores” and stretched nucleolar DNA at the meeting, but they were his first attempts and not particularly revealing. But later, he and his talented research assistant Barbara Beatty showed the world what these genes really look like in full transcriptional action (Miller and Beatty 1969). These pictures earned their rightful place as among the most iconic of any in the history of cytology and cell biology. Incisive studies on the synthesis and processing of ribosomal RNA introduced by Sheldon Penman at the Montevideo meeting were subsequently refined by him and independently by the laboratory of James Darnell (reviewed by Lewin 1980). In contrast, the isolation of nucleoli presented at the meeting was to await several decades for further advances. In his summary of the Montevideo conference, the Edinburgh embryologist C.H.

Waddington said: “The nucleolus probably should not be considered a relatively simple organelle with a single function, comparable to a machine tool turning out a particular part of an automobile. It is not just ‘the organelle where the cell manufactures ribosomes.’ It is rather a structure through which materials of several different kinds are flowing, comparable more to a whole production line than to a single machine tool.” One cannot imagine a more prescient view. As we shall see, every atom of his statement has been borne out in subsequent research on the nucleolus.

INTO THE MODERN ERA

In the 1970s and 1980s, the nucleolus field addressed the details of ribosome biosynthesis. One axis was a troubled one: the goal of reconciling the stages of ribosome synthesis with the classically defined subcompartments of the

T. Pederson

nucleolus. Just as one commentator famously described (to every Latin student centuries later) the Roman conquest territory that would later become France, “Gallia est omnis divisa in partes tres” (Caesar, 40– 50s BC), the nucleolus is also tripartite. Its three classical regions are defined by the different appearance of intranucleolar regions when viewed by electron microscopy (Fig. 3). These are the fibrillar centers, the surrounding dense fibrillar component, and the granular component. Few in the field anticipated the controversies that would ensue when several labs tried, logically enough, to link the sites of rRNA transcription and processing and assembly into nascent ribosomes with these three EM-defined nucleolar zones. These debates were waged with intense controversy and, in a few cases, frank boorishness on the part of some in this raging controversy on the floor at international meetings. In hindsight, this had to do with how short a pulse label one could introduce to tag nascent rRNA. Thankfully, recent advances have settled the issue. A key step in this resolution was a reconciliation of the “Christmas tree” EM images of spread nascent ribosomal RNA (Miller and Beatty 1969) with the high resolution detection of these genes and

Figure 3. The tripartite organization of the nucleolus.

Electron micrograph of a mouse fibroblast nucleolus. (f ) fibrillar center, (d) dense fibrillar component, (g) granular component, (arrows) perinucleolar heterochromatin, ( ) denotes the presence of dense fibrillar component material within the fibrillar center, which is occasionally observed. (Reprinted from Trends in Cell Biol 13: 517 –525, Rasˇka, I. # 2003, with permission from Elsevier.)

their transcripts in nucleoli in situ (Koberna et al. 2002; for further discussion of the earlier controversy and its resolution, see Rasˇka et al. 2006). Recent work has indicated that one of the three nucleolar regions, the granular component, is itself composed of at least two distinct molecular domains. Meanwhile, the covalent steps in rRNA processing continued to be explored, revealing different nuances in yeast versus higher eukaryotes superimposed on a conserved foundation (reviewed by Fatica and Tollervey 2002). But another important axis was also emerging in the 1970s. The laboratory of Harris Busch, notably Ramachandra Reddy, came upon and sequenced several small nucleolar RNAs. One of these, U3 small nucleolar RNA, was subsequently shown to be base-paired with nucleolar rRNA in vivo (Calvet and Pederson 1981) and later was demonstrated to function in the initial cleavage of the 45S rRNA precursor (Kass et al. 1990). All of these studies were done, of course, with deproteinized nucleolar RNA. Soon thereafter, isolated nucleoli were subjected to various extraction conditions designed to preserve nascent rRNA– protein complexes, resulting in the characterization of the ribonucleoprotein forms of the rRNA precursor and processing intermediates that had previously been defined with deproteinized RNA (Warner and Soeiro 1967; Pederson and Kumar 1971; Kumar and Warner 1972). Jumping ahead, we now enjoy a far more detailed picture of the covalent steps of rRNA maturation (Fatica and Tollervey 2002) as well as numerous accessory factors that support this pathway but are not part of the final ribosomes (reviewed by Granneman and Baserga 2004). Among the latter are more than 100 small nucleolar RNAs that guide the extensive site-specific nucleotide modifications (ribose 20 -O-methylation and pseudouridine formation) of rRNA during its processing (reviewed by Bachellerie et al. 2002). Like the initial rRNA precursor, processing intermediates, and products themselves, all these small nucleloar RNAs are complexed with proteins, thus instilling in the nucleolar landscape a high concentration of both RNA and protein, a point to which we shall return.

The Nucleolus

A PARADIGM SHIFT

By the 1960s and into the 1970s, the notion that the nucleolus is the seat of ribosome synthesis was as cogently established as anything in the modern canon of the molecular and cell biology of eukaryotes. Then, something unexpected happened. In my laboratory, we had been studying the traffic of fluorescently labeled small nucleolar RNAs after microinjection into the nuclei of cultured mammalian cells and finding them to move into nucleoli, as expected, but extending this to define nucleolar targeting elements in these RNAs (Jacobson et al. 1995; 1997; Jacobson and Pederson 1998a). In the case of U3 and U8 (Jacobson and Pederson 1998a), we planned to expand the findings but first decided to submit an initial report to convey our novel experimental approach (reviewed in Pederson 2001a), something quite new at the time for the nucleolus, though we had previously applied it to study pre-mRNA traffic to nucleoplasmic domains (Wang et al. 1991). While writing this paper (Jacobson and Pederson 1998a), it occurred to us that we had only used a single RNA as a negative control. Although it did not display nucleolar localization after microinjection, we decided to try a second small RNA that presumably would also not localize in nucleoli, thus amplifying our case that the observed nucleolar localizations of U3 and U8 were specific. My post-doc Marty Jacobson suggested that we try the signal recognition particle RNA, a 300-nt pol III transcript, not only to fulfill the need for another negative control but also because, to his credit, he realized nothing was known about the nuclear phase of SRP biosynthesis. To our surprise, SRP RNA displayed as rapid and as quantitative a nucleolar localization as U3 and U8 snoRNAs. But looking into this further, we saw that SRP RNA trafficked into nucleoli only transiently. U3 and U8 on the other hand remained stably localized in nucleoli, as did other small nucleolar RNAs we had begun to study. SRP RNA then moved out into the cytoplasm, where it became associated with the endoplasmic reticulum, its expected final localization (Jacobson and Pederson 1998b).

These findings on SRP RNA, together with a number of other contemporaneous results that were stirring, led to the formulation of the “plurifunctional nucleolus” hypothesis (Pederson 1998a). Subsequently, the link between the nucleolus and signal recognition particle biosynthesis was made in yeast (Ciufo and Brown 2000; Grosshans et al. 2001), and confirmed and extended in mammalian cells (Politz et al. 2000) (Fig. 4) and in Xenopus oocytes (Sommerville et al. 2005). A speculative essay suggested how a common nucleolar assembly site might have co-evolved for the four translational ribonucleoproteins (the two ribosomal subunits, the 5S rRNA– protein complex, and the SRP) (Pederson and Politz 2000). Notwithstanding how it came about in the evolution of eukaryotes, the fact that the nucleolus is a site of more than ribosome synthesis had clearly arrived once its role in the biosynthesis of the SRP had been discovered. Also embedded in the plurifunctional nucleolus hypothesis were provocative results linking the nucleolus to cellcycle progression (Pederson 1998a,b). Both the SRP and cell-cycle aspects of the hypothesis were welcomed into reviews on the nucleolus (Scheer and Hock 1999; Olson et al. 2000; Olson et al. 2002; Visintin and Amon 2000), and the concept has continued to be well received (Alberts et al. 2002; Rasˇka et al. 2006; Boisvert et al. 2007). THE PARTS LIST, AND SEGUEING INTO THE CELL CYCLE

When nucleoli were first isolated (Vincent 1952), only assays for total protein and RNA, as well as tests for various enzymatic activities, were available (the latter scoring only one—acid phosphatase). Jumping ahead four and a half decades, we now speak of specific nucleolar molecules. As mentioned earlier, progress on chronicling all of the small RNAs present in nucleoli had proceeded through the 1980s and 1990s and on into the 2000 decade. As for nucleolar proteins (beyond the ribosomal structural proteins), early accounts (e.g., Soeiro and Basile 1973) were followed by numerous reports of individual, non-ribosomal proteins observed

T. Pederson

Figure 4. The nucleolus and signal recognition particle biosynthesis. Nucleolar localization of GFP-tagged

SRP19 (left) and SRP68 (right) proteins in rat NRK cells. Cytoplasmic localization is also observed, as expected from the known steady-state localization of the signal recognition particle. Similar results were obtained with GFP-tagged SRP72 protein, whereas the SRP54 protein assembles with the nascent signal recognition particle in the cytoplasm (Politz et al. 2000; Sommerville et al. 2005). Reproduced from Politz et al. 2000, with permission from the Proceedings of the National Academy of Sciences.

either in isolated nucleoli or, more typically, by immunofluorescence or GFP-tagged protein localization. These were useful findings on each protein’s individual merit but obviously did not reveal the molecular breadth of the nucleolar protein landscape. In 2002, two groups refined the classical nucleolar isolation methods (notably that developed by Maggio et al. 1963) and prepared high purity nucleoli from HeLa cells followed by proteomics (Andersen et al. 2002; Scherl et al. 2002; reviewed by Pederson 2002 and Leung et al. 2003). These two contemporaneous proteomics studies revealed, in combination, a total of approximately 350 proteins. Excluding ribosomal structural proteins, there were many that had been or would be later linked to ribosome biosynthesis as chaperone-like mediators not ending up in mature ribosomes. This finding expanded and put more molecular definition on earlier research, showing that nucleolar preribosomal particles have a higher protein:RNA mass ratio than mature ribosomes (Liau and Perry 1969; Pederson and Kumar 1971; Kumar and Warner 1972). Many others of the observed nucleolar proteins were ones involved in DNA replication or repair, cell-cycle progression, or its control, validating this aspect of the initial

plurifunctional nucleolus hypothesis (Pederson 1998a) and findings being made at the time that had suggested such a link (reviewed by Pederson 1998b; Garcia and Pillus 1999; Vinistin and Amon 2000). Similar findings were later made in a proteomics analysis of purified nucleoli from Arabidopsis (Pendle et al. 2005). Beyond proteomic analysis, and its emphasis on nonribosomal proteins, there has been recent work on ribosomal proteins themselves that has brought surprises. The first of these implicated ubiquitinylation, including polyubiquitinylation, of ribosomal proteins in the pathway of ribosome synthesis (Stavreva et al. 2006), a most unanticipated, counterintuitive result and one not immediately reconcilable with the “standard model” of ribosome biosynthesis. The second study revealed a surprisingly large pool of unassembled ribosomal proteins, exceeding the level needed to support ribosome assembly on the number of rRNA transcripts being produced (Lam et al. 2007). Obviously, one senses the likely possibility of a link between these two sets of observations. These findings also lead us to bear in mind the possibility that there may be further surprises ahead regarding the established lore of the nucleolar ribosome biosynthetic pathway, just as

The Nucleolus

has been the case with respect to the nonribosomal aspects of nucleolar organization and function in the 12 years since the plurifunctional nucleolus hypothesis was advanced (Pederson 1998a). THE NUCLEOLUS IS NEITHER AS STATIC NOR AS COMPACT AS FIRST THOUGHT

Between 2000 and 2001, a number of studies based on fluorescence recovery after photobleaching (FRAP), in one case employing as well the complementary method of fluorescence loss in photobleaching (FLIP), brought about a complete revision in thinking about the stasis of several nuclear proteins (for reviews, see Pederson 2000; 2001b; Leung and Lamond 2003). In the case of the nucleolar proteins studied, namely fibrillarin (Phair and Misteli 2000) and selected ribosomal structural proteins (Chen and Huang 2001), the dynamics were faster than would have been anticipated by many (perhaps most) nucleolus investigators. Fibrillarin functions as an rRNA modification enzyme in the dense fibrillar component and therefore might have been assumed to either be stably associated with the nuclelous or at least display a long residence time. A nucleolar stasis of ribosomal structural proteins was an even more entrenched notion, as they would have been assumed to enter nucleoli in stoichiometric amounts to assemble with nascent rRNA and yet they were found to be rapidly shuttling in and out of nucleoli. As mentioned above, later work revealed that ribosomal structural proteins undergo ubiquitinylation and degradation on the assembly pathway (Stavreva et al. 2006) and also enter the nucleus in suprastoichiometric levels (Lam et al. 2007). Clearly, the FRAP results (Chen and Huang 2001) reflected, or at least were significantly influenced by, these later-recognized aspects of ribosomal protein instability, whereas the fast exchange of fibrillarin (Phair and Misteli 2000) was, and remains, even more surprising. It has been more difficult to study the dynamics of nucleolar RNAs. The fact that fluorescent snoRNAs accumulate in nucleoli after microinjection into the nucleus does not address this

question since the initial localization may either be filling unoccupied sites or replacing those formerly occupied by egressing snoRNAs. Stated differently, snoRNAs are always at high concentration in the nucleolus so the concentration of the putative molecules that have moved into the nucleoplasm in any experiment that examines the steady-state would be very low. Only methods like FRAP or FLIP can capture this and these are not applicable to RNA. However, endogenous RNAs can be tagged with fluorescent probes (Politz et al. 1998), including photoactivatable ones (Politz 1999; Politz et al. 2005). In a study using this approach to track 28S rRNA out of the nucleolus (Politz et al. 2003), we noted that some signal occasionally returned to nucleoli. We pondered that there is probably no barrier to a completed ribosomal subunit occasionally gaining access back into the nucleolus (which is surprisingly porous, vide infra) before it engages the nuclear export machinery. Notwithstanding this observation, how dynamic or static nucleolar RNAs are remains an open and important question. How compact a bundle of mass is the nucleolus? Its appearance in differential interference microscopy (Fig. 1) indicates that it has higher mass per unit volume than the surrounding nucleolplasm, but how much higher? Fortunately, differential interference microscopy can be used in a quantitative mode. Its application to the nucleus of Xenopus oocytes revealed that the nucleoli are only about twice as dense as the surrounding nucleoplasm (Handwerger et al. 2005; discussed in Pederson 2010a.) In retrospect, it may be that a preferentially high affinity of some nucleolar proteins for the heavy metal atoms (uranium, osmium, tungsten) used in electron microscopy has led to a false perception of a higher ( protein) density in the nucleolus, or in some of its regions, than is actually the case. The use of basic dyes in classical cytological work on the nucleolus that are taken preferentially by polyanions such as RNA, or silver stains that bind avidly to certain nucleolar proteins, may have contributed to this notion from cytochemistry. In any case, similar studies to those done in the nucleus of Xenopus oocyte nuclei are now needed in mammalian cells, where

T. Pederson

there are more data on nucleolar protein dynamics that can be used as parameters in computational models. A first step has been taken in this direction, in which the percolation of ovalbumin (molecular weight 43,000) in the nucleolus and nucleoplasm was measured in a mammalian cell (Spell and Kubitscheck 2009), revealing that this protein had the same mobility in both compartments. One is also reminded here that the use of photo-activatable GFP-tagged nucleolar proteins in such studies could be very revealing, as has been their use in studying another protein’s dynamics in the Xenopus oocyte nucleus (Deryusheva and Gall 2004). A GROWING FAMILY OF CELL-CYCLE REGULATORS IN THE NUCLEOLUS

The nucleolus proteomics studies revealed numerous cell-cycle-related proteins among the residents (or visitors). A great many studies over the past decade have brought further insights into the nucleolar transit of cell-cycle regulatory proteins. These good endeavors have now engendered a large literature, much of which is more pertinent to cell-cycle progression control in general, whereas only a subset of this emerging work relates to the nucleolus; hence the brief, selective coverage that follows. Before embarking on the relationship between nucleolar activity and cell-cycle progression, a caveat needs to be registered. There is no doubt that when ribosome production is impaired or limiting, cell growth slows or ceases. The obligatory relationship between ribosome biosynthesis and cell growth rate was established ´´ le Maaloe and Karl for bacteria in studies by O Lark in the 1960s, and later was confirmed for eukaryotes. We can plausibly surmise that new ribosomes do count for a cell’s (or organism’s) future because selection would disfavor cells, embryos, or adult organisms that are slow to build new proteins for either cell division (a necessary inheritance for G1 progeny) or for sustained vigor of tissues and organs. That said, we here take up the question of whether the nucleolus might play roles in cell-cycle progression beyond ribosome biosynthesis (see also

Pederson 2007). This is not an idea that was pondered in the classical era, when ribosome production, protein synthesis rates, and cell growth were seen as the operation of a continuous, forward- and back-regulated production logic circuit. The point of departure was the finding that the action of two cell-cycle regulators, p53 and Mdm2, are modulated by nucleolar sequestration (Weber et al. 1999; Kulikov et al. 2010). Another key finding was that p53 is stabilized upon DNA damage-triggered nucleolar disruption or elicited by other cellular stressors (Rubbi and Milner 2003). Subsequent work has defined an extensive interactome of nucleolus-nucleoplasmic shuttling cell-cycle progression proteins and it now appears that the action of several is based on their dynamic interplay between the nucleolus and the nucleoplasm. An instructive example is nucleostemin (reviewed by Ma and Pederson 2008a; Pederson and Tsai 2009). This protein was discovered in a study of neural stem cells in the subventricular zone of the adult rat brain (Tsai and McKay 2002). Nucleostemin is a p53-interactive protein (Tsai and McKay 2002; Ma and Pederson 2007) and the initial idea (Tsai and McKay 2005; reviewed by Misteli 2005) was that when it shuttles from the nucleolus into the nucleoplasm, it binds to and disables p53, and thus drives the cell cycle. But the continuing challenge in this field is to reconcile a protein’s intranuclear location with function. For example, although p53 is mostly nucleoplasmic, the small nucleolar fraction of p53 has different binding partners, measured by bimolecular fluorescence complemenatation (see Ma and Pederson 2008b), than those with which it consorts in the nucleoplasm (Pederson, unpubl.). It is usually assumed that the highest intranuclear concentration of a protein is where it executes its function. This assumption may be suspect in some cases, and in the case of nucleostemin, the jury is out as to whether it enacts its cell-cycledriving function in the nucleolus (at high concentration) or in the nucleoplasm (at low concentration). A protein’s concentration in a region of the cell may not be a quantitative signal for its action, which should be kept in mind

The Nucleolus

as systems biology approaches now arrive in cell (and nuclear) biology. THE PERINUCLEOLAR COMPARTMENT: ANOTHER LINK BETWEEN THE NUCLEOLUS AND GROWTH CONTROL?

In some cells, there is a discoid structure situated on the nucleolus, covering a portion of its surface as a “cap.” Termed the perinucleolar compartment (PNC), it was discovered on the basis of its high concentration of a particular hnRNP protein (Ghetti et al. 1992; Matera et al. 1995; for reviews, see Huang 2000 and Pollock and Huang 2010). The PNC accretes, but is not the transcription site of, a number of small RNAs (Matera et al. 1995; Wang et al. 2003). The fact that PNCs are not seen in all cells but are noted particularly in mammalian transformed cell lines, tumor-derived cell lines, and tumor biopsy specimens suggests a key link between their appearance and tumor initiation and/or progression (see Pollock and Huang 2010). Although this is an encouraging new tool for the pathologist, the PNC’s harboring of certain proteins and RNAs presently offers no clues whatsoever as to why it appears more frequently in tumor cells. Clearly, further work on the PNC is warranted. In fact, it is the only presently known link between the nucleolus (in the form of a nucleolus-vicinal cytological entity) and a very high morbidity/mortality human disease, breast carcinoma. THE NUCLEOLUS AND VIRAL REPLICATION, STEM CELL BIOLOGY, AND CELLULAR SENESCENCE: ONLY ZEPHYRS SO FAR

Beyond cell-cycle progression and growth control, the past decade has seen a surge of attention to the nucleolus in three other fields: viral replication, stem cell biology, and cellular senescence. These studies include a number of provocative findings, briefly discussed below, but the objective summary statement is that a mechanistic link to the nucleolus has not been made. Regarding viral replication, some viruses display a nucleolar tropism, but the functional

significance of this is generally not well understood. A surprising recent finding was that the nucleolus and Cajal bodies are required for the establishment of a systemic infection by a plant virus (Kim et al. 2007). In an intriguing retroviral therapeutic development, the fact that unspliced HIV RNA exits the nucleus via the nucleolus was exploited to design a nucleolustargeting inhibitor of viral replication (Michienzi et al. 2002). The nucleolus has also been implicated in the replication of herpes viruses (Boyne and Whitehouse 2006.) Yet another recent case is the finding that the replication of the Dependovirus AAV2 (adenovirus-associated virus 2) takes place in the nucleolus (Sonntag et al. 2010). AAV2 is among the smallest animal cell viruses known (25 nm in diameter) and its assembly might thus be feasible within the relatively porous nucleolus, yet the biological significance of this nucleolar tropism is presently unclear (reviewed by Pederson 2010a). The connection of the nucleolus to stem cell biology has received some attention, with a possible link hinted at both by classical and contemporary studies (e.g., Gonda et al. 2003; reviewed by Misteli 2003; Fle´chon 2006). Thus, when a somatic nucleus from an adult vertebrate animal is placed into an enucleated egg, development can proceed up to a tadpole in the case of amphibians or even birth in the case of mammals. Much attention has been placed on the effects of the egg cytoplasm on the implanted nucleus in terms of epigenetic marks and the redirected program of gene expression. But, something else also happens. In the nucleolus of the adult, somatic nucleus transiently disappears and then comes back into view a few hours later. This intriguing behavior of the nucleolus in somatic nucleus transfer experiments has not been explored to the extent it should be. If properly investigated, this is likely to reveal profound clues as to what happens when newly combined nuclei and cytoplasm become molecularly familiar with themselves. Another potential link between the nucleolus and stem cell biology has arisen from studies of the transcription factor Hand1, whose localization within and outside the nucleolus acts as a trigger for cell fate determination (Martindill

T. Pederson

et al. 2007). Another engaging recent study has implicated a U3 small nucleolar RNA-bound protein, Wcd, in a neural stem cell niche of Drosophila embryos (Fichelson et al. 2009). Wcd displays asymmetric distribution to daughter cells and thus has “stem cellness” written all over it. One hopes that these groups and/or other nucleolus investigators are pursuing these two findings, for they are among the most intriguing links between the nucleolus and stem cell biology to have been reported. Studies linking the nucleolus to cellular senescence round out this body of “conceptually nucleolus-related” research in that, once again, some provocative clues have been uncovered, but there is no sense at present of a unifying concept or mechanism. For example, an influential study in yeast reported an acceleration of aging (whatever this means in fungi) in mutants displaying nucleolar fragmentation (Sinclair et al. 1997; reviewed in Guarente 1997). And, a number of studies have made a connection between the nucleolus and Werner syndrome, an inherited human-aging-like condition (Marciniak et al. 1998; Szekely et al. 2000; von Kobbe and Bohr 2002; Kyng et al. 2003), and this seems the best case so far for a role of the nucleolus in cellular senescence, though still far from being linked to aging of a higher eukaryotic organism. A QUADRIPARTITE NUCLEOLUS, OR MORE?

After discovering that the SRP assembly occurs in the nucleolus (Jacobson and Pederson 1998b; Politz et al. 2000; Sommerville et al. 2005), the question arose as to the intranucleolar site. Probing for both SRP RNA and 28S rRNA in dual color in situ hybridization experiments revealed incomplete overlap, although both signals resided in the granular component (Politz et al. 2002). Given this evidence for molecular segregation even within a given nucleolar subcompartment, we reasoned that probing for a molecule not involved in either ribosome or SRP biosynthesis would be the logical next step and chose nucleostemin. The results conformed and amplified the notion that the nucleolar granular component is molecularly territorialized (Politz et al. 2005). But, the most important

finding in this study came from the application of electron spectroscopic imaging, which revealed the granular component to be an interspersed array of RNA-rich and RNA-deficient particles, rather than a uniform lawn of nascent ribosomes, as would have been anticipated (Fig. 5). It remains to be seen how many subdivisions exist within the granular component, or within the dense fibrillar component for that matter. The probes and optical resolution are now at hand to address this further. TO RESIST OR REVISIT HERESY? DOES THE NUCLEOLUS PARTICIPATE IN MESSENGER RNA BIOSYNTHESIS?

Beyond the possible roles of the nucleolus in cell-cycle control, viral replication, stem cell biology, and cellular senescence, there is another potential nucleolus function that warrants consideration, both for its historical precedence and its now reasonable plausibility based on a number of studies. This is the possibility that the nucleolus is involved, somehow, in the production of messenger RNA. In the 1960s and 1970s, Henry Harris at the University of Oxford posited that the nucleolus was a way station for mRNA export from the nucleus (see Harris 1974). He had been a pioneer of somatic cell genetics, unsullied as such, but was also a constitutional contrarian. He had become something of an enfant terrible outside his immediate field by challenging the thenemerging (and prevailing) views on the meaning of high molecular weight nuclear RNA as regards the pathway of eukaryotic messenger RNA biosynthesis (at a time when the connection was admittedly still wobbly). But, in contrast to his mRNA biosynthesis contrarianism, Harris’ idea that the nucleolus might be involved in mRNA export was based on results, not mere speculation. He had fused chicken erythrocytes, containing a condensed, inactive nucleus, with HeLa cells and found that chicken-specific proteins appeared in these heterokaryons only after the erythrocyte nucleus had undergone chromatin decondensation and the appearance of nucleoli. From this, he elaborated, and for years promulgated, the idea that messenger RNA must

The Nucleolus

Figure 5. The nucleolar granular component contains both RNA-rich and RNA-deficient particles. Electron spectroscopic imaging (ESI) is a mode of energy loss measurement in electron microscopy that can be tuned to record beam collisions with orbital electrons of nitrogen or phosphorus in the specimen. Using the empirical chemical formulas of protein and nucleic acid with respect to these two elements, ESI allows one to construct a map of protein-rich versus nucleic-acid-rich objects. Shown is an ESI image from a portion of a human neuroblastoma cell nucleus spanning a region of the nucleoplasm and part of a nucleolus. The protein-rich and nucleicacid-rich regions have been colored blue and yellow, respectively. (DCh) decondensed chromatin, (CCh) condensed chromatin, (PML) promyelocytic leukemia body, (GC) granular component of the nucleolus. The PML body (entirely blue, i.e., protein) serves as a valuable internal control, as it is known not to contain appreciable nucleic acid. The granular component of the nucleolus, once thought to be a uniform zone of nascent ribosomes, contains an interspersed landscape of nucleic-acid-rich versus nucleic-acid-deficient particles. ESI cannot distinguish between DNA and RNA, but since several independent methods have detected no DNA in the nucleolar granular component, the nucleic-acid-rich particles are plausibly nascent ribosomes, perhaps with a contribution from assembling signal recognition particles, whereas the nucleic-acid-deficient, protein-rich particles may be heterotypic complexes of cell-cycle regulatory proteins, as discussed. (Reprinted from Politz JC et al. 2005, Mol Biol Cell 16: 3401–3410.)

obligatorily pass through the nucleolus. The contemporaneous work of Penman, Darnell, and other groups dispelled this notion for mRNA as a population, en masse, but in a few corners of the field the possibility lingered that

a subset of mRNAs might visit the nucleolus prior to export. Subsequently, there were reports of specific mRNAs in the nucleolus (reviewed in Pederson 1998a) and a recent study (Kim et al. 2009) has rekindled these earlier observations and carried them into the modern era of methodology. In addition, several microRNAs have now been localized in the nucleolus (Politz et al. 2006; Politz et al. 2009) and there is increasing evidence that some microRNAs may be derived from snoRNAs (discussed in Politz et al. 2009; see also Taft et al. 2009). There is also recent evidence for transfer RNA-derived microRNAs (reviewed by Pederson 2010b), and given that the nucleolus has been implicated in tRNA biosynthesis, this is yet another fascinating possibility for a link between the nucleolus and mRNA regulation via microRNAs. Although it is counterintuitive that negative regulators of mRNA function would be associating with nascent mRNAs before the latter even leave the nucleus, intuition has often been an unreliable docent in the field of gene expression. The possible role of the nucleolus in mRNA export and/or microRNA function and dynamics certainly warrants further investigation. NEW FRONTIERS: WHAT LIES BENEATH

Among the remaining issues, the classical phenomenon of nucleolar dominance stands ready for renewed analysis, as has been happening productively (Earley et al. 2010; reviewed by Costa-Nunes et al. 2010). The role of intergenic transcripts in the control of ribosomal RNA transcription (Mayer et al. 2006; Santoro et al. 2010) is another fertile area of current investigation (and likely intersects with nucleolar dominance). Moreover, there is growing evidence that a gene’s positioning or repositioning near or at the nucleolus can regulate its expression (e.g., Royo et al. 2009) and that the rDNA can itself impact the genome globally (e.g., Parades and Maggert 2009). Understanding the process of nucleolar assembly is also a high priority and although there has been substantial progress on this from a cell biological perspective (reviewed by Hernandez-Verdun et al. 2002), a new system

T. Pederson

(Prieto and McStay 2007, 2008) promises molecular detail. A role for the nucleolus has recently emerged in studies of how a postsynaptic density protein of dendritic spines shuttles back to the cell body after neurotransmitter activation (reviewed by Richer and Fallon 2007), raising the possibility of an entire new field linking the nucleolus to neurobiology. Also foremost on the agenda is continued investigation of the extent to which the nucleolus directs the cell cycle (Pederson 2007), as well as the clinical utility of the PNC. A link between the nucleolus, p53, and innate immunity in C. elegans (Fuhrman et al. 2009) is also among the most provocative recent findings and warrants immediate exploration both in nematodes and in a wider phyletic range. Also standing before us is the entire question of mRNA and microRNA traffic through the nucleolus and its functional meaning. Yet, other new vistas on the nucleolus horizon will almost certainly appear in due course, but meanwhile, the agenda is both full and promising. CONCLUSION

The nucleolus is unique in that the discovery of one of its functions took far longer, namely more than a century, than was the case for most other cell components. Another half-century has brought the realization that the nucleolus is also the site of signal recognition particle biosynthesis, serves as a regulatory zone of cell-cycle progression mediators, and is a locus of mRNA and microRNA traffic, the functional meaning of which remains to be discovered. The nucleolus is now known to be a more dynamic domain of nuclear organization than once thought, and with functions even beyond those first grandly recognized atop a Uruguay mountain in 1965. ACKNOWLEDGMENTS

Cited work from the author’s laboratory was supported by grants R01 GM-21595 and R01 GM60551 from the National Institutes of Health, grant MCB-0445841 from the National Science Foundation, and grant RGP0031 from the Human Frontier Scientific Program Organization

(with co-investigators Jan Ellenberg, Angus Lamond, and Matthias Mann). Marty Jacobson in the author’s lab played a key role in the signal recognition particle work that was a catalyst in formulating the plurifunctional nucleolus hypothesis. Joan Politz creatively advanced and confirmed this hypothesis in her SRP work, amidst other pioneer research she led on mRNA and rRNA intranuclear dynamics in the author’s lab. I am greatly indebted to them both. I am also grateful to Joseph Gall, Susan Gerbi, and Masayasu Nomura for their longstanding support of my research on the nucleolus, encouraging my efforts into unsettled and unexplored terrain. REFERENCES Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. 2002. Molecular Biology of the Cell, 4th edition. Garland Science, New York. p. 332. Andersen JS, Lyon CE, Fox AH, Leung AK, Lam YW, Steen H, Mann M, Lamond AI. 2002. Directed proteomic analysis of the human nucleolus. Curr Biol 12: 1 –11. Bachellerie JP, Cavaille´ J, Hu¨ttenhofer A. 2002. The expanding RNA world. Biochimie 84: 775–790. Boisvert FM, van Koningsbruggen S, Navascue´s J. Lamond AI. 2007. The multifunctional nucleolus. Nat Rev Mol Cell Biol 8: 574– 585. Boyne JR, Whitehouse A. 2006. Nucleolar trafficking is essential for nuclear export of intronless herpesvirus mRNA. Proc Natl Acad Sci USA 103: 15190– 15195. Brown DD, Gurdon JB. 1964. Absence of rRNA synthesis in the anucleolate mutant of X. laevis. Proc Natl Acad Sci USA 51: 139 –146. Brown DD, Dawid IG. 1968. Specific gene amplication in oocytes. Oocyte nuclei contain extrachromosomal replicas of the genes for ribosomal RNA. Science 160: 272– 280. Caesar J. (40-50s BC). De Bello Gallico I: 1. Calvet JP, Pederson T. 1981. Base-pairing interactions between snRNAs and nuclear RNA precursors as revealed by psoralen cross-linking in vivo. Cell 26: 363 –370. Chen D. Huang S. 2001. Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells. J Cell Biol 153: 169– 176. Ciufo LF, Brown JD. 2000. Nuclear export of yeast signal recognition particle lacking Srp54p by the Xpo1p/Crm1p NES-dependent pathway. Curr Biol 10: 1256– 1264. Costa-Nunes P, Pontes O, Preuss SB. Pikaard CS. 2010. Extra views on RNA-dependent DNA methylation and MBD6dependent heterochromatin formation in nucleolar dominance. Nucleus 1: 1– 6. Deryusheva S, Gall JG. 2004. Dynamics of coilin in Cajal bodies of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 101: 4810–4814.

The Nucleolus Earley KW, Pontvianne F, Wierzbicki AT, Blevins T, Tucker S, Costa-Nunes P, Pontes O, Pikaard CS. 2010. Mechanisms of HDA6-mediated rRNA gene silencing; suppression of intergenic pol II transcription and differential effects on maintenance versus siRNA-directed cytosine methylation. Genes Dev 24: 1119–1132. Fatica A, Tollervey D. 2002. Making ribosomes. Curr Opin Cell Biol 14: 313 –318. Fichelson P, Moch C, Ivanovitch K, Martin C, Sidor CM, Lepesant J-A, Bellaiche Y. Huynh J-R. 2009. Live-imaging of single stem cells within their niche reveals that a U3 snoRNP component segregates asymmetrically and is required for self-renewal in Drosophila. Nat. Cell Biol 11: 685–693. Fle´chon JE. 2006. Analysis of the nucleolar compartment of the nucleus as an indicator of nuclear reprogramming after nuclear transfer. Meth Mol Biol 325: 225 –238. Fuhrman LE, Goel AK, Smith J, Shianna KV, Aballay A. 2009. Nucleolar proteins suppress Caenorhabditis elegans innate immunity by inhibiting p53/CEP-1. PLoS Genetics 5: e1000657 Gall JG. 1968. Differential synthesis of the genes for ribosomal RNA during amphibian oogenesis. Proc Natl Acad Sci USA 60: 553– 560. Garcia SN, Pillus L. 1999. Net results of nucleolar dynamics. Cell 97: 825– 828. Ghetti A, Pinol-Roma S, Michael WM, Morandi C. Dreyfuss G. 1992. hnRNP I, the polypyrimidine tract-binding protein: distinct nuclear localization and association with hnRNAs. Nucleic Acids Res 20: 3671– 3678. Gonda K, Fowler J, Katoku-Kikyo N, Haroldson J, Wudel J, Kikyo N. 2003. Reversible disassembly of somatic nucleoli by the germ cell proteins FRGY2a and FRGY2b. Nat Cell Biol 5: 205–210. Granneman S, Baserga SJ. 2004. Ribosome biogensis: of knobs and RNA processing. Exp Cell Res 296: 43– 50. Grosshans H, Deinhert K, Hurt E, Somos G. 2001. Biogenesis of the signal recognition particle (SRP) involves import of SRP proteins into the nucleolus, assembly with the SRP RNA, and Xpo1p-mediated export. J Cell Biol 153: 745– 761. Guarente L. 1997. Link between aging and the nucleolus. Genes Dev 11: 2449–2455. Handwerger KE, Cordero JA, Gall JG. 2005. Cajal bodies, nucleoli and speckles have a low-density, sponge-like structure. Mol Biol Cell 16: 202– 211. Harris H. 1974. Nucleus and Cytoplasm 3rd edition. Clarendon Press, Oxford. Heitz E. 1931. Nukleolar und chromosomen in der gattung. Vicia Planta 15: 495–505. Hernandez-Verdun D, Roussel P. Ge´brane-Youne`s J. 2002. Emerging concepts of nucleolar assembly. J Cell Sci 115: 2265– 2270. Huang S. 2000. Perinucleolar structures. J Struct Biol 129: 233–240. Jacobson MR, Cao LG, Wang YL, Pederson T. 1995. Dynamic localization of RNase MRP RNA in the nucleolus observed by fluorescent RNA cytochemistry in living cells. J Cell Biol 131: 1649– 1658.

Jacobson MR, Cao LG, Taneja K, Singer RH, Wang YL, Pederson T. 1997. Nuclear domains of the RNA subunit of RNase P. J Cell Sci 110: 829 –837. Jacobson MR, Pederson T. 1998a. A 7-methylguanosine cap commits U3 and U8 small nuclear RNAs to the nucleolar localization pathway. Nucleic Acids Res 26: 756–760. Jacobson MR, Pederson T. 1998b. Localization of signal particle RNA in the nucleolus of mammalian cells. Proc Natl Acad Sci USA 95: 7981– 7986. Kass S, Tyc K, Steitz JA, Sollner-Webb B. 1990. The U3 small nucleolar ribonucleoprotein functions in the first step of pre-ribosomal RNA processing. Cell 60: 897–908. Kim SH, Koroleva OA, Lewandowska D, Pendle AF, Clark GP, Simpson CG, Shaw PJ, Brown JW. 2009. Aberrant mRNA transcripts and the nonsense-mediated decay proteins UPF2 and UPF3 are enriched in the Arabidopsis nucleolus. Plant Cell 21: 2045– 2057. Kim SH, Ryabov EV, Kalinina NO, Rakitina DV, Gillespie T, MacFarlane S, Haupt S, Brown JWS, Taliansky M. 2007. Cajal bodies and the nucleolus are required for a plant virus systemic infection. EMBO J 26: 2169–2179. Koberna K, Malinsky J, Pliss A, Masata M, Vecerova J, Fiolova M, Bednar J, Rasˇka I. 2002. Ribosomal genes in focus: new transcripts label the dense fibrillar components and form clusters indicative of “Christmas trees”. J Cell Biol 157: 743– 748. Kulikov R, Letienne J, Kaur M, Grossman SR, Arts J, Blattner C. 2010. Mdm2 facilitates the association of p53 with the proteasome. Proc Natl Acad Sci USA 107: 10038– 10043. Kumar A, Warner JR. 1972. Characterization of ribosomal precursor particles from HeLa cell nucleoli. J Mol Biol 63: 233– 246. Kyng KJ, May A, Kø´vraa S, Bohr VA. 2003. Gene expression profiling in Werner syndrome closely resembles that of normal aging. Proc Natl Acad Sci USA 100: 12259– 12264. Lam YW, Lamond AI, Mann M, Andersen JS. 2007. Analysis of nucleolar protein dynamics reveals the nuclear degradation of ribosomal proteins. Curr Biol 17: 749 –760. Leung AKL, Andersen JS, Mann M, Lamond AI. 2003a. Bioinformatic analysis of the nucleolus. Biochem J 376: 553 –569. Leung AKL, Lamond AI. 2003. The dynamics of the nucleolus. Crit Rev Euk Gene Expr 13: 39– 54. Lewin B. 1980. Gene Expression. Volume 2. Eukaryotic Chromosomes. 2nd edition. John Wiley & Sons, New York. Liau MC, Perry RP. 1969. Ribosome precursor particles in nucleoli. J Cell Biol 42: 272– 283. Ma H, Pederson T. 2007. Depletion of the nucleolar protein nucleostemin causes G1 cell cycle arrest via the p53 pathway. Mol Biol Cell 18: 2630–2635. Ma H, Pederson T. 2008a. Nucleostemin: a multiplex regulator of cell-cycle progression. Trends Cell Biol 18: 575– 579. Ma H, Pederson T. 2008b. Nucleophosphmin is a binding partner of nucleostemin in human osteosarcoma cells. Mol Biol Cell 19: 2870– 2875. Maggio R, Siekevitz P, Palade GE. 1963. Studies on isolated nuclei. I. Isolation and chemical composition of nucleolar and nucleoplasmic subfractions. J Cell Biol 18: 293 –312.

T. Pederson Marciniak RA, Lombard DB, Johnson FB, Guarente L. 1998. Nucleolar localization of the Werner syndrome protein in human cells. Proc Natl Acad Sci USA 95: 6887–6892. Martindill DMJ, Risebro CA, Smart N, Franco-Viseras MDM, Rosario CO, Swallow CJ, Dennis JW, Roliry PR. 2007. Nucleolar release of Hand1 acts as a molecular switch to determine cell fate. Nat Cell Biol 9: 1131– 1141. Matera AG, Frey MR, Margelot K, Wolin SL. 1995. A perinucleolar compartment contains several RNA polymerase III transcripts as well as the polypyrimidine tract-binding protein. J Cell Biol 129: 1181– 1193. Mayer C, Schmitz KM, Li J, Grummt I, Santoro R. 2006. Intergenic transcripts regulate the epigenetic state of rRNA genes. Mol Cell 22: 351– 361. McClintock B. 1934. The relationship of a particular chromosomal element to the development of the nucleoli in Zea mays. Z. Zellforsch Mikrosk 21: 294– 398. Michienzi A, Li S, Zaia JS, Rossi JJ. 2002. A nucleolar TAR decoy inhibitor of HIV-1 replication. Proc Natl Acad Sci USA 99: 14047–14052. Miller OL Jr, Beatty BR. 1969. Visualization of nucleolar genes. Science 164: 955– 957. Misteli T. 2003. A nucleolar disappearing act in somatic cloning. Nat Cell Biol 5; 183 –184. Misteli T. 2005. Going in GTP cycles in the nucleolus. J Cell Biol 168: 177–178. Montgomery TH. 1898. Comparative cytological studies, with especial regard to the morphology of the nucleolus. J Morphol 15: 265– 582. Olson MO, Dundr M, Szebeni A. 2000. The nucleolus: an old factory with unexpected capabilities. Trends Cell Biol 10: 189– 196. Olson MO, Hingorani K, Szebeni A. 2002. Conventional and nonconventional roles of the nucleolus. Int Rev Cytol 219: 199– 266. Parades S, Maggert KA. 2009. Ribosomal DNA contributes to global chromatin regulation. Proc Natl Acad Sci USA 106: 17829– 17834. Pederson T. 1998a. The plurifunctional nucleolus. Nucleic Acids Res 26: 3871–3876. Pederson T. 1998b. Growth factors in the nucleolus? J Cell Biol 143: 279–281. Pederson T. 2000. Diffusional protein transport within the nucleus: a message in the medium. Nat Cell Biol 2: E73– E74. Pederson T. 2001a. Fluorescent RNA cytochemistry: tracking gene transcripts in living cells. Nucleic Acids Res 29: 1013– 1016. Pederson T. 2001b. Protein mobility within the nucleuswhat are the right moves? Cell 104: 635–638. Pederson T. 2002. Proteomics of the nucleolus: more proteins, more functions? Trends Biochem Sci 27: 111– 112. Pederson T. 2007. Ribosomal protein mutations in Diamond-Blackfan anemia: might they operate upstream of protein synthesis? The FASEB J 21: 3442– 3445. Pederson T. 2010a. “Compact” nuclear domains: reconsidering the nucleolus. Nucleus 1: 444– 445. Pederson T. 2010b. Regulatory derived from transfer RNA? RNA 16: 1865– 1869.

Pederson T, Kumar A. 1971. Relationship between protein synthesis and ribosome assembly in HeLa cells. J Mol Biol 61: 655 –668. Pederson T, Politz JC. 2000. The nucleolus and the four ribonucleoproteins of translation. J Cell Biol 148: 1091–1095. Pederson T, Tsai RYL. 2009. In search of nonribosomal protein function and regulation. J Cell Biol 184: 771– 776. Pendle AF, Clark GP, Boon R, Lewandowska D, Lam YW, Andersen J, Mann M, Lamond AI, Brown JW, Shaw PJ. 2005. Proteomic analysis of the Arabidopsis nucleolus suggests novel nucleolar functions. Mol Biol Cell 16: 260– 269. Perry RP. 1966. Nucleolus: Structure and function. Science 153: 214–219. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604 –609. Politz JC. 1999. Use of caged fluorochromes to track macromolecular movement in living cells. Trends Cell Biol 9: 284 –287. Politz JC, Browne ES, Wolf DE, Pederson T. 1998. Intranuclear diffusion and hybridization state of oligonucleotides measured by fluorescence correlation spectroscopy. Proc Natl Acad Sci USA 95: 6043– 6048. Politz JCR, Hogan EM, Pederson T. 2009. MicroRNAs with a nucleolar location. RNA 15: 1705– 1715. Politz JC, Lewandowski LB, Pederson T. 2002. Signal recognition particle RNA localization within the nucleolus differs from the classical sites of ribosome synthesis. J Cell Biol 159: 411 –418. Politz JCR, Polena I, Trask I, Bazett-Jones DP, Pederson T. 2005. A nonribosomal landscape in the nucleolus revealed by the stem cell protein nucleostemin. Mol Biol Cell 16: 3401– 3410. Politz JC, Tuft RA, Pederson T. 2003. Diffusion-based transport of nascent ribosomes in the nucleus. Mol Biol Cell 14: 4805–4812. Politz JCR, Tuft RA, Pederson T. 2005. Photoactivationbased labeling and in vivo tracking of RNA molecules in the nucleus. Live Cell Imaging: A Laboratory Manual Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Politz JC, Yarovoi S, Kilroy S, Gowda K, Zwieb C, Pederson T. 2000. Signal recognition particle components in the nucleolus. Proc Natl Acad Sci USA 97: 55–60. Politz JCR, Zhang F, Pederson T. 2006. MicroRNA-206 colocalizes with ribosome-rich regions in both the nucleolus and cytoplasm of rat myogenic cells. Proc Natl Acad Sci USA 103: 18957– 18962. Pollock C, Huang S. 2010. The perinucleolar compartment. Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect. a000679. Prieto J-L, McStay B. 2007. Recruitment of factors linking transcription and processing of pre-rRNA to NOR chromatin is UBF-dependent and occurs independent of transcription in human cells. Genes Dev 21: 2041–2054. Prieto J-L, McStay B. 2008. Pseudo-NORs; a novel method for studying nucleoli. Biochim et Biophys Acta 1783: 2116–2123. Rasˇka I. 2003. Oldies but goldies: searching for Christmas trees within the nucleolar architecture. Trends Cell Biol 13: 517– 525.

The Nucleolus Rasˇka I, Shaw PJ, Cmarko D. 2006. Structure and function of the nucleolus in the spotlight. Curr Opin Cell Biol 18: 325–334. Richter J.D. Fallon JR. 2007. Synapses go nucle(ol)ar. Nat Neurosci 10: 399–400. Royo F, Paz N, Espinosa L, McQueen PG, Vello´n L, Parada LA. 2009. Spatial link between nucleoli and expression of the ZacI gene. Chromosoma 118: 711– 722. Rubbi CP, Milner J. 2003. Disruption of the nucleolus mediates stabilization of p53 in response to DNA damage and other stressors. EMBO J 22: 6068– 6077. Santoro R, Schmitz K-M, Sandoval J, Grummt I. 2010. Intergenic transcripts originating from a subclass of ribosomal DNA repeats silence ribosomal RNA genes in trans. EMBO rep 11: 52– 58. Scheer UHock R. 1999. Structure and function of the nucleolus. Curr Opin Cell Biol 11: 385 –390. Scherl A, Coute´ Y, De´on C, Calle´ A, Kindbeiter K, Sanchez JC, Greco A, Hochstrasser D, Diaz JJ. 2002. Functional proteomic analysis of human nucleolus. Mol Biol Cell 13: 4100–4109. Sinclair DA, Milles K, Guarente L. 1997. Accelerated aging and nucleolar fragmentation in yeast sgs1 mutants. Science 277: 1313– 1316. Soeiro R, Basile C. 1973. Non-ribosomal nucleolar proteins in HeLa cells. J Mol Biol 79: 507 –519. Sommerville J, Brumwell CL, Politz JCR, Pederson T. 2005. Signal recognition particle assembly in relation to the function of amplified nucleoli of Xenopus oocytes. J Cell Sci 118: 1299– 1307. Sonntag F, Schmidt K, Kleinschmidt JA. 2010. A viral assembly factor promotes AAV2 capsid assembly in the nucleolus. Proc Natl Acad Sci USA 107: 10220– 10225. Spell J, Kubitscheck U. 2009. Single ovalbumin molecules exploring nucleoplasm and nucleoli of living cell nuclei. Biochim Biophys Acta 1803: 396 –404. Stavreva DA, Kawasaki M, Dundr M, Koberna K, Muller WG, Tsujimura-Takahashi T, Komatsu W, Hayano T, Isobe T, Rasˇka I, et al. 2006. Potential roles for ubiquitin and the proteasome during ribosome biogenesis. Mol Cell Biol 26: 5131– 5145.

Szekely AM, Chen Y-H, Zhang C, Oshima J, Weissman SM. 2000. Werner protein recruits DNA polymerase d to the nucleolus. Proc Natl Acad Sci USA 97: 11365– 11370. Taft RJ, Glazov EA, Lassmann T, Hayashizaki Y, Carninci P, Mattick JS. 2009. Small RNAs derived from snoRNAs. RNA 15: 1233–1240. Tsai RY, McKay RD. 2002. A nucleolar mechanism controlling cell proliferation in stem cells and cancer cells. Genes Dev 16: 2991–3003. Tsai RY, McKay RD. 2005. A multistep, GTP-driven mechanism driving the dynamic cycling of nucleostemin. J Cell Biol 168: 179 –184. Valentin G. 1836. Repertorium fu¨r Anatomie und Physiologie 1: 1 –293. Verlag von Veit und Comp. Berlin. Valentin G. 1839. Repertorium fu¨r Anatomie und Physiologie 4: 1 –275. Verlag von Veit und Comp. Berlin. Vincent WS. 1952. The isolation and chemical properties of the nucleoli of starfish oocytes. Proc Natl Acad Sci USA 38: 139– 145. Vincent WS, Miller OL. 1966. International symposium on the nucleolus. Its structure and function. Montevideo, Uruguay. J Natl Cancer Inst Monogr 23: 1– 610. Vinistin R, Amon A. 2000. The nucleolus: the magician’s hat for cell cycle tricks. Curr Opin Cell Biol 12: 372– 377. von Kobbe C, Bohr VA. 2002. A nucleolar targetting sequence in the Werner syndrome protein resides within residues 949-1092. J Cell Sci 115: 3901–3907. Wagner R. 1835. Einige Bemerkungen und Fragen u¨ber das Keimbla¨schen (vesicular germinativa). Mu¨ller’s Archiv Anat Physiol Wissenschaft Med 373– 377. Wang J, Cao LG, Wang YL, Pederson T. 1991. Localization of pre-messenger RNA at discrete nuclear sites. Proc Natl Acad Sci 88: 7391–7395. Wang C, Politz JC, Pederson T, Huang S. 2003. RNA polymerase III transcripts and the PTB protein are essential for the integrity of the perinucleolar compartment. Mol Biol Cell 14: 2425–2435. Warner JR, Soeiro R. 1967. Nascent ribosomes from HeLa cells. Proc Natl Acad Sci USA 58: 1984– 1990. Weber JD, Taylor LJ, Roussel MF, Sherr CJ, Bar-Sagi D. 1999. Nucleolar Arf sequesters Mdm2 and activates p53. Nat Cell Biol 1: 20– 26.

Nuclear Speckles David L. Spector1 and Angus I. Lamond2 1

Cold Spring Harbor Laboratory, One Bungtown Road, Cold Spring Harbor, New York 11724

2

Wellcome Trust Centre for Gene Regulation and Expression, College of Life Sciences, University of Dundee, MSI/WTB/JBC Complex Dow Street, Dundee DD1 5EH, United Kingdom

Correspondence: [email protected], [email protected]

Nuclear speckles, also known as interchromatin granule clusters, are nuclear domains enriched in pre-mRNA splicing factors, located in the interchromatin regions of the nucleoplasm of mammalian cells. When observed by immunofluorescence microscopy, they usually appear as 20– 50 irregularly shaped structures that vary in size. Speckles are dynamic structures, and their constituents can exchange continuously with the nucleoplasm and other nuclear locations, including active transcription sites. Studies on the composition, structure, and dynamics of speckles have provided an important paradigm for understanding the functional organization of the nucleus and the dynamics of the gene expression machinery.

he mammalian cell nucleus is a highly compartmentalized yet extremely dynamic organelle (reviewed in Misteli 2001a; Spector 2006; Zhao et al. 2009). Many nuclear factors are localized in distinct structures, such as speckles, paraspeckles, nucleoli, Cajal bodies, polycomb bodies, and promyelocytic leukemia bodies and show punctate staining patterns when analyzed by indirect immunofluorescence microscopy (reviewed in Lamond et al. 1998; Spector 2001; Spector 2006). In mammalian cells the pre-mRNA splicing machinery, including small nuclear ribonucleoprotein particles (snRNPs), spliceosome subunits, and other non-snRNP protein splicing factors, shows a punctate nuclear localization pattern that is usually termed “a speckled pattern” but has also been referred to as “SC35 domains (Wansink et al. 1993)” or “splicing factor

T

compartments (Phair et al. 2000)” (Figs. 1 and 2). The first detailed description of the nuclear domains that we presently refer to as nuclear speckles was reported by Santiago Ramo´n y Cajal in 1910 (Ramo´n y Cajal 1910; reviewed in Lafarga et al. 2009). Ramo´n y Cajal used acid aniline stains to identify structures he referred to as “grumos hialinas” (literally “translucent clumps”). The term “speckles” was first put forth in 1961 by J. Swanson Beck (Beck 1961) upon examination of rat liver sections immunolabeled with the serum of individuals with autoimmune disorders. Although the connection was not made at the time, these speckles had been identified two years earlier by Hewson Swift (Swift 1959) at the electron microscopic level and called interchromatin particles. Swift observed that these particles were not randomly distributed but that they occurred in localized

Editors: Tom Misteli and David L. Spector Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2011 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000646 Cite this article as Cold Spring Harb Perspect Biol 2011;3:a000646

D.L. Spector and A.I. Lamond

Figure 1. Speckles form in the interchromatin space.

HeLa cells showing splicing factors localized in a speckled pattern as well as being diffusely distributed throughout the nucleoplasm. Bar ¼ 5 mm.

“clouds,” and cytochemical analysis indicated that they contained RNA (Swift 1959). However, the first link between pre-mRNA splicing and nuclear speckles or interchromatin granule clusters came from an examination of the distribution of snRNPs using anti-splicing factor-specific antibodies, demonstrating a speckled distribution pattern of snRNPs in cell nuclei (Lerner et al. 1981; Perraud et al. 1979; Spector et al. 1983). It is now clear that much of the punctate localization of splicing factors observed by immunofluorescence microscopy corresponds to the presence of these factors in nuclear speckles of variable size and irregular shape that are revealed by electron microscopy as interchromatin granule clusters (IGCs) (Fig. 3). IGCs range in size from one to several micrometers in diameter and are composed of 20 – 25 nm granules that are connected in places by a thin fibril resulting in a beaded chain appearance (Thiry 1995b). These structures can be observed by electron microscopy without antibody labeling (Thiry 1995b). We will define “speckles” here specifically as the IGC component of the

splicing factor labeling pattern, and distinguish this from other nuclear structures, including perichromatin fibrils and Cajal bodies, which also contain splicing factors (reviewed in Fakan 1994; Spector 1993). For some of the speckle components a speckle targeting signal has been identified. The arginine/serine-rich domain (RS domain) of some SR pre-mRNA splicing factors has been shown to be necessary and sufficient for the targeting of these factors to nuclear speckles (Caceres et al. 1997; Hedley et al. 1995; Li et al. 1991). In addition, the threonine-proline repeats of SF3b1 (Eilbracht et al. 2001) and the forkhead-associated domain in NIPP1 (Jagiello et al. 2000) have also been implicated in speckle-targeting. Most recently, Salichs et al. (Salichs et al. 2009) performed a genomewide analysis of homopolymeric histidine tracts and identified 86 human proteins that contain stretches of five or more histidines. Of the 22

Figure 2. Structured illumination microscopy, using the OMX system (Applied Precision, Issaqua, Washington), of a HeLa cell expressing SC35-EYFP. At 100 nm resolution substructure can be observed within speckles. In addition, the diffuse population of SC35-EYFP is resolved as a granular distribution. Projection of twelve 0.125 mm optical sections through the center of a nucleus encompassing 1.5 mm. Image provided by Zsolt Lazar and R. Ileng Kumaran. Bar ¼ 2 mm.

Nuclear Speckles

studies using an anti-splicing factor antibody to confirm the localization of any novel factors to nuclear speckles. STRUCTURE AND LOCATION OF SPECKLES

Figure 3. Nuclear speckles are equivalent to inter-

chromatin granule clusters. Immunoelectron microscopy using a primary antibody against SC35 and a secondary antibody conjugated to 15 nm colloidal gold. IGCs are composed of a series of particles measuring 20 –25 nm in diameter that are connected in places by a thin fibril resulting in a beaded chain appearance. Bar ¼ 500 nm.

that were nuclear localized, 15 were shown to be present in nuclear speckles. Based on these data, the polyHis-repeats were proposed to act as a speckle-targeting signal that functions by acting as an interaction surface for resident nuclear speckle constituents. Interestingly, these targeting signals rely mostly on charge effects, being basic protein regions. Interestingly, structures similar to nuclear speckles have been identified in the amphibian oocyte nucleus (Gall et al. 1999) and in Drosophila melanogaster embryos when transcription increases upon cellularization during cycle 14 (Segalat et al. 1992), but not in yeast (Potashkin et al. 1990). Importantly, not all nuclear proteins that show a speckle-like labeling pattern by immunofluorescence microscopy localize to IGCs. For example, the ER repeat protein YT521-B localizes in a speckled-like distribution that corresponds to YT bodies (Nayler et al. 2000), whereas PSPC1 localizes in approximately 5– 20 punctate interchromatin structures termed “paraspeckles,” which are also distinct from nuclear speckles (Fox et al. 2002; Fox and Lamond 2010). Therefore, it is essential to perform double-label immunofluorescence

As determined by both light and electron microscopy, the clusters of interchromatin granules that constitute speckles form throughout the nucleoplasm in regions containing little or no DNA (Thiry 1995b). Although they apparently contain few, if any, genes speckles are often observed close to highly active transcription sites. This suggests that they likely have a functional relationship with gene expression, and some specific genes have been reported to preferentially localize near speckles (Brown et al. 2008; Huang et al. 1991; Johnson et al. 2000; Moen et al. 2004; Smith et al. 1999; Xing et al. 1993; Xing et al. 1995), although this does not appear to be obligatory for transcription/premRNA splicing. Interestingly, Shopland et al. (Shopland et al. 2003) found that gene-rich chromosomal regions (R-bands) are more frequently found along the edge of nuclear speckles than gene-poor regions (G-bands). In addition, coordinately expressed active genes can be found in association with the same nuclear speckle. Based on these findings, Shopland et al. (2003) suggested that nuclear speckles act as functional centers that organize active genes on their periphery to form euchromatic neighborhoods. Several lines of evidence point to speckles acting as storage/assembly/modification compartments that can supply splicing factors to active transcription sites (reviewed in Lamond and Spector 2003). For example, a series of high resolution pulse-labeling experiments analyzed at the electron microscopic level, studying the incorporation of either tritiated uridine or Br-UTP after short pulses, have shown that nascent pre-mRNA is predominantly localized outside of nuclear speckles (IGCs) in fibrillar structures, 3– 5 nm in diameter, which are termed perichromatin fibrils (PFs) (Cmarko et al. 1999; Fakan et al. 1971; Fakan et al. 1978; Monneron et al. 1969). It is likely that most of the cotranscriptional splicing

D.L. Spector and A.I. Lamond

is associated with these PFs, rather than within IGCs. PFs can occur both on the periphery of IGCs and in nucleoplasmic regions away from IGCs (Fakan 1994). Some apparent discrepancies in the literature concerning the possible direct role of speckles as splicing sites may have arisen because the PFs can show a close topological relationship with the periphery of IGCs. Using the fluorescence microscope it is difficult to distinguish these PFs from the IGCs. In addition, as highly expressed genes will recruit a significant amount of pre-mRNA splicing factors (Huang et al. 1996), these regions of highly active transcription will be indistinguishable from IGCs at the fluorescence microscopy level. Although many in the field do not view speckles as direct transcription/pre-mRNA splicing centers, others suggest that they may have a more direct role relating to the splicing and transport of pre-mRNA (reviewed in Hall et al. 2006; Melcak et al. 2000; Shopland et al. 2002; Wei et al. 1999). COMPOSITION OF SPECKLES

Many pre-mRNA splicing factors, including snRNPs and SR proteins (Fu 1995), have been localized to nuclear speckles by either immunofluorescence, fluorescent protein-tagging, and/ or immunoelectron microscopy. In fact, this speckled localization pattern is highly diagnostic for proteins involved in pre-mRNA splicing. In addition, several kinases (Clk/STY, hPRP4, and PSKHI) (Brede et al. 2002; Colwill et al. 1996; Ko et al. 2001; Kojima et al. 2001; SaccoBubulya et al. 2002) and phosphatases (PP1) (Trinkle-Mulcahy et al. 1999; Trinkle-Mulcahy et al. 2001) that phosphorylate/dephosphorylate components of the splicing machinery have also been localized to nuclear speckles. This supports the idea that speckles may be involved in regulating the pool of factors that are accessible to the transcription/pre-mRNA processing machinery (reviewed in Misteli et al. 1997b). The protein composition of nuclear speckles has been assessed by proteomic analysis of an enriched IGC fraction purified from mouse liver nuclei. This approach identified 146 known proteins, as well as numerous uncharacterized

proteins (Mintz et al. 1999; Saitoh et al. 2004). The proteomic information, together with additional localization studies, has revealed that speckles contain many other proteins apart from pre-mRNA splicing factors. Of particular interest is the localization of transcription factors (Larsson et al. 1995; Mortillaro et al. 1996; Zeng et al. 1997), 30 -end RNA processing factors (Krause et al. 1994; Schul et al. 1998), eukaryotic translation initiation factor eIF4E (Dostie et al. 2000), eif4AIII, a protein involved in translation inhibition (Li et al. 1999), and structural proteins (Jagatheesan et al. 1999; Nakayasu et al. 1984; Sharma et al. 2010). Consistent with these findings, proteomic analyses of in vitro assembled spliceosomes indicate that they may also contain transcription and 30 -end RNA processing factors, together with splicing factors, in a higher order complex (Rappsilber et al. 2002; Zhou et al. 2002). However, they do not contain factors primarily involved in ribosome subunit biogenesis or tRNA production, and the protein composition of speckles underlines their close relationship with mechanisms of gene expression by RNA Pol II. Although transcription does not take place within the majority of nuclear speckles (Cmarko et al. 1999) and DNA is not localized to these nuclear regions (reviewed in Thiry 1995b), a population of the serine-2-phosphorylated form of the RNA polymerase II (RNAPII) large subunit (LS) that is involved in elongation has been localized to these regions by immunofluorescence microscopy (Bregman et al. 1995; Mortillaro et al. 1996). In addition, biochemical characterization of the IGC proteome has identified several subunits of RNAPII (Mintz et al. 1999; Saitoh et al. 2004), supporting the presence of a pool of RNAPII in speckles. However, other studies have not observed an enrichment of RNAPII in speckles (Grande et al. 1997; Kimura et al. 2002; Zeng et al. 1997) and it is not present in B snurposomes (Doyle et al. 2002). The Cdk9-cyclin T1 complex, also known as TAK/P-TEFb, is thought to be involved in transcriptional elongation via phosphorylation of the RNAPII LS (reviewed in Price 2000). This complex was found diffusely distributed throughout the nucleoplasm, but not in nucleoli

Nuclear Speckles

(Herrmann et al. 2001). In addition, a significant overlap between cyclin T1 and nuclear speckles was observed. However, although Cdk9 was present in the vicinity of nuclear speckles, the degree of overlap was limited (Herrmann et al. 2001; Matera et al. 1993). Further evidence for a link between transcription and speckles comes from observations on FBI-1, a cellular POZ-domain-containing protein that binds to the HIV-1 long terminal repeat and associates with the HIV-1 transactivator protein Tat (Pessler et al. 1997). FBI-1 has been found to partially colocalize with Tat and its cellular cofactor, P-TEFb at nuclear speckles (Pendergrast et al. 2002). In addition, the nucleosome binding protein HMG-17, which can alter the structure of chromatin and enhance transcription, has been localized in a similar pattern to FBI-1 (Hock et al. 1998). Therefore, although little or no transcription takes place in nuclear speckles, a subset of proteins involved in this process are associated with these nuclear regions in addition to being present at transcription sites. Although it is currently unclear what determines the subset of transcription factors that are localized to nuclear speckles, their presence may relate to the assembly of higher-order complexes and/or to regulatory steps affecting either the modification state or accessibility of specific transcription factors. In addition to transcription factors, a population of poly(A)þ RNA has been localized to nuclear speckles (Carter et al. 1991; Huang et al. 1994; Visa et al. 1993). This population of poly(A)þ RNA does not chase to the cytoplasm when transcription is blocked with aamanitin, as would be expected if these species represented nascent mRNA (Huang et al. 1994). Interestingly, Hutchinson et al. (Hutchinson et al. 2007) identified MALAT1 (metastasis-associated lung adenocarcinoma transcript 1), a long nuclear retained noncoding RNA to be enriched in nuclear speckles (reviewed in Wilusz et al. 2009). Recent studies have implicated MALAT1 in the recruitment of SR splicing factors from nuclear speckles to sites of transcription (Bernard et al. 2010) and in the regulation of alternative splicing by modulating SF2/ ASF phosphorylation (Tripathi et al. 2010).

Although an underlying scaffold that would serve as a platform on which to organize IGCs has thus far not been identified (Sacco-Bubulya et al. 2002), several proteins with possible structural roles in the nucleus, such as a population of lamin A (Jagatheesan et al. 1999) and snRNP-associated actin (Nakayasu et al. 1984), have been detected in nuclear speckles. However, another study failed to detect an alteration in nuclear speckles in LMNA-/- cells (Vecerova et al. 2004). In addition to actin, phosphatidylinositol (4,5)-bisphosphate [PtdIns(4,5)P2], a lipid that regulates actin-binding proteins (Zhao et al. 1998), as well as multiple phosphatidylinositol phosphate kinase (PIPK) isoforms, have also been localized to nuclear speckles (Boronenkov et al. 1998). Recently, Sharma et al. (Sharma et al. 2010) have implicated the 2530amino acid Son protein in the organization of nuclear speckles. Son contains a concentrated region of multiple tandem repeat sequences including multiple serine-rich repeats and an RS domain. Based on RNAi depletion experiments, Son was proposed to act as a scaffolding protein for RNA processing factors in nuclear speckles (Sharma et al. 2010). DYNAMICS OF NUCLEAR SPECKLES

Speckles are dynamic structures; their size, shape, and number can vary, both between different cell types and within a cell type, according to the levels of gene expression and in response to signals that influence the pools of active splicing and transcription factors available. When transcription is halted, either by the use of inhibitors, or as a result of heat shock, splicing factors accumulate predominantly in enlarged, rounded speckles (Melcak et al. 2000; Spector et al. 1991; Spector et al. 1983). The fact that nuclear speckles become round and increase in size upon transcriptional inhibition supports the view that speckles may function in the storage/assembly/modification of splicing factors, and that they are not direct sites of splicing. Furthermore, when expression of introncontaining genes increases (Huang et al. 1996; Misteli et al. 1997a), or during viral infection when transcription levels are high (Bridge et al.

D.L. Spector and A.I. Lamond

1995; Jime´nez-Garcı´a et al. 1993), the accumulation of splicing factors in speckles is reduced, and they redistribute to nucleoplasmic transcription sites. Individual speckle components can therefore shuttle continually between speckles and active gene loci. Speckles are also dynamically regulated during mitosis. The movement of factors into and out of speckles can be directly visualized by fluorescence microscopy as fluctuations in the shape and intensity of speckles in live cells expressing splicing factor/fluorescent protein fusions (Misteli et al. 1997a). Speckles in such cells show transcription-dependent peripheral movements, although individual speckles remain in their neighborhoods. Photobleaching techniques have also been used to measure the flux of some speckle components and have shown that their exchange rate is very rapid (Kruhlak et al. 2000; Phair et al. 2000). Complete recovery for GFP-SF2/ASF (a member of the SR-family of pre-mRNA splicing factors), after photobleaching of the fluorescence signal in speckles, was apparent in approximately 30 seconds with half recovery in approximately 3–5 seconds. The movement rates for splicing factors through the nucleoplasm were measured to be slow compared with free GFP; this reduction in movement was proposed to result from multiple transient interactions of splicing factors with nuclear binding sites, both within and outside of speckles. Kinetic modeling indicated that the maximal mean residence time for GFP-SF2/ASF in speckles was less than 50 seconds (Phair et al. 2000). It is a remarkable feature of nuclear organization that at steady-state the overall structure of speckles, as well as other nuclear domains, persists despite the large flux of their components. THE SPECKLE CELL CYCLE

Upon entry into mitosis and following breakdown of the nuclear envelope/lamina, proteins associated with nuclear speckles become diffusely distributed throughout the cytoplasm (Ferreira et al. 1994; Reuter et al. 1985; Spector et al. 1986; Thiry 1995a). During metaphase, these proteins continue to localize in a diffuse cytoplasmic pattern and also accumulate within

one to three small structures called mitotic interchromatin granules (MIGs) (Ferreira et al. 1994; Leser et al. 1989; Prasanth et al. 2003; Verheijen et al. 1986). MIGs appear to be structurally analogous to IGCs (Leser et al. 1989; Thiry 1993; Thiry 1995a). As mitosis progresses from anaphase to early telophase, the MIGs increase in number and size. During mid- to late-telophase and after re-formation of the nuclear envelope/lamina, pre-mRNA splicing factors enter daughter nuclei and, concomitantly, their localization in MIGs decreases, demonstrating that these factors are recycled from the cytoplasmic MIGs into daughter nuclei (Prasanth et al. 2003). Live-cell studies have indicated that the majority of these factors enter daughter nuclei within 10 minutes (Prasanth et al. 2003). Although MIGs have been proposed to be the mitotic equivalent of nuclear speckles (Ferreira et al. 1994; Leser et al. 1989; Thiry 1995a; Thiry 1995b), their function in mitotic cells is unclear. In telophase cells, some MIGs were found to be in close proximity to the newly formed nuclear envelope (Prasanth et al. 2003; Thiry 1995a). This close proximity of MIGs to the nuclear periphery and the disappearance of MIGs in late telophase cells with the concomitant appearance of IGCs in daughter nuclei have suggested that the MIGs might be directly transported into the nuclei (Leser et al. 1989; Thiry 1995a). However, colocalization of SF2/ ASF and a hyperphosphorylated form of RNAPII LS in MIGs of late telophase cells has suggested that this may not be the case. For example, SF2/ASF and other pre-mRNA processing factors were shown to enter daughter nuclei while a subpopulation of SC35 and RNAPII LS remained in MIGs until G1, demonstrating that various components of MIGs are differentially released for subsequent entry into daughter nuclei (Prasanth et al. 2003). Further support for differential release of factors from MIGs comes from an earlier study that reported the nuclear import of snRNPs while cytoplasmic MIGs were still labeled with anti-SR protein and anti-SC35 antibodies (Ferreira et al. 1994). Based on these findings, it was suggested that MIGs may play important roles

Nuclear Speckles

either in the modification of the components of the splicing machinery before their nuclear entry, or as enriched populations of these factors, allowing for protein-protein interactions to occur between subsets of proteins before their nuclear entry (Prasanth et al. 2003). Interestingly, splicing factors were shown to be competent for pre-mRNA splicing immediately upon entry into daughter nuclei (Prasanth et al. 2003), supporting the possibility that MIGs may be responsible for splicing factor modification, allowing for immediate targeting of modified ( phosphorylated) pre-mRNA processing complexes to transcription sites in telophase nuclei. Because daughter nuclei late in telophase have not yet assembled nuclear speckles, cytoplasmic MIGs are likely to function as their counterparts to provide competent premRNA splicing factors to the initial sites of transcription in newly formed nuclei (Prasanth et al. 2003). Perhaps splicing factors are released from MIGs via hyperphosphorylation, as has been shown for their release from nuclear speckles in interphase nuclei. SPECKLE BIOGENESIS

Nuclear speckles are one of the most prominent nuclear compartments, and their study has served as a paradigm for understanding the biogenesis of nuclear bodies. Most evidence points to the fact that nuclear speckles form through a process of self-assembly (reviewed in Misteli 2001b) whereby transient macromolecular interactions form the basis of speckle morphogenesis. Under steady-state conditions, the respective rates of association and disassociation of individual speckle components will define their exchange rates and the sizes of their bound and soluble pools in the nucleus. Regulatory mechanisms can influence these association and/or disassociation rates, thereby changing the fraction of bound and soluble speckle components in response to specific cellular signals. In this view, the entry of splicing factors into late-telophase nuclei results in an association of a subset of these factors with initial transcription/pre-mRNA processing sites (Prasanth et al. 2003). As the population of factors

increases, there is an increased probability of protein-protein interactions among those factors not engaged in transcription/pre-mRNA processing, resulting in the formation of nuclear speckles. These initial speckles appear to form predominantly in nucleoplasmic regions that are devoid of chromosome territories and/or other nuclear organelles. They may initiate either at random locations, or in the vicinity of genes that are transcribed at high levels during the telophase/G1 transition. Interestingly, in this regard Brown et al. (Brown et al. 2008) examined the position of various erythroid genes in erythroblasts and found that the majority of associations between erythroid genes occurred at a nuclear speckle. Interestingly, the associations were predominantly observed in regard to active genes. Based on their findings, the authors proposed that active genes can be brought into close proximity by the nucleation of splicing factors into nuclear speckles. The size and shape of interphase speckles is a reflection of the steady-state dynamics of protein constituents that are both arriving at and leaving from these structures (Kruhlak et al. 2000; Phair et al. 2000). Although photobleaching analyses have indicated rapid recovery kinetics of splicing factors in speckles, consistent with a diffusion-based process (Kruhlak et al. 2000; Phair et al. 2000), the relative size of speckles remains constant throughout interphase. In fact, the incubation of permeabilized cells with a nuclear extract containing an ATPregenerating system maintains transcriptional activity and does not result in a loss of speckles (Misteli et al. 1996), nor does simple treatment of unfixed cells with detergent (Spector et al. 1992). The observed basal exchange rate may be directly related to the maintenance of speckles, rather than indirectly related to an involvement in transcriptional/pre-mRNA processing events. In addition, the irregular shape of individual nuclear speckles in interphase nuclei may result from a nonuniform release and/or delivery of factors, related to the location of active genes in their vicinity (Misteli et al. 1997a). Consistent with this possibility, either upon inhibition of RNAPII transcription by a-amanitin

D.L. Spector and A.I. Lamond

(Spector et al. 1993), or inhibition of pre-mRNA splicing using an antisense approach (O’Keefe et al. 1994), speckles tend to round-up, suggesting a uniform exchange rate of factors in all directions. This exchange rate of speckle factors may be regulated through phosphorylation/dephosphorylation events. For example, phosphorylation of the RS domain of SR splicing factors has been shown to be necessary for recruitment of SR proteins from nuclear speckles to sites of transcription/pre-mRNA processing (Misteli et al. 1998) and for their association with the forming spliceosome (Mermoud et al. 1994). Several kinases (i.e., Clk/STY [Colwill et al. 1996; Sacco-Bubulya et al. 2002] and hPRP4 [Kojima et al. 2001]) involved in this phosphorylation, as well as a kinase proposed to be involved in the phosphorylation of the carboxyterminal domain of RNAPII LS in vitro (Ko et al. 2001), have been localized to nuclear speckles, leaving open the possibility that phosphorylation/dephosphorylation plays a role in determining the basal rate of factor exchange. However, in addition to the basal activities, an additional level of control can be exerted by modulating phosphorylation events. For example, the rapid induction either of a gene (Huang et al. 1996), or group of genes such as during viral infection (Bridge et al. 1995; Jime´nezGarcı´a et al. 1993), can result in an increased outward flow of factors from speckles. An extreme example of this can be observed upon overexpression of Clk/STY kinase, or addition of SRPK1 kinase to permeabilized cells (Gui et al. 1994a; Gui et al. 1994b), which results in the complete redistribution of splicing factors from speckles to the diffuse nuclear pool (Colwill et al. 1996; Sacco-Bubulya et al. 2002). Interestingly, expression of a mutant form of Clk/STY that lacks its catalytic activity resulted in an increased accumulation of factors in highly concentrated foci on the periphery of speckles, possibly a reflection of their inability to be released (Sacco-Bubulya et al. 2002). Consistent with this observation, the addition of kinase inhibitors to cells resulted in an inhibition of the dynamic movements on the periphery of speckles (Misteli et al. 1997a). Also,

protein phosphatase 1 (PP1) inhibitors resulted in enlarged irregularly shaped speckles with less well defined edges, probably resulting from the inability of factors to be released from PFs, on the periphery of IGCs, also consistent with a modulating effect on the exchange rate (Misteli et al. 1996). In summary, a basal exchange rate of factors, coupled with a mechanism to modulate this rate (that is, providing a stimulus-induced burst), ensures that the needed factors, in the correct phosphorylation state, are available to pre-mRNA transcripts at the sites of transcription. In addition, such a mechanism ensures that a significant population of factors, which are not functionally needed, are sequestered out of the soluble nuclear pool, in this case in nuclear speckles, representing a basic mechanism for the organization of nonmembranebound nuclear organelles. CONCLUSIONS

Nuclear speckles are organelles located in the interchromatin nuclear space and are among the most widely studied nuclear domains. They are best known for accumulating high local concentrations of snRNPs and other non-snRNP protein splicing factors. However, the presence in speckles of many other factors involved in mRNA production by RNA polymerase II further supports their intimate relationship with gene expression. Although most speckles apparently do not contain DNA in an analogous way to the rRNA gene repeats within nucleoli, nonetheless highly expressed genes can be found associated with speckles, consistent with an important role for speckles in coordinating the supply and/or recycling of pre-mRNA processing and transcription factors. The analysis of nuclear speckles has helped to establish some key paradigms and principles for the dynamic assembly of membrane-free organelles in the nucleus. We anticipate that future work will refine our understanding of speckle composition and of mechanisms involved in targeting proteins to speckles and regulating their formation. It is likely that building a detailed model of gene expression in vivo will require further

Nuclear Speckles

characterization of nuclear speckles to define how the complex events required for transcription and RNA processing are efficiently coordinated within the nucleus.

ACKNOWLEDGMENTS

We thank members of the Spector and Lamond groups for helpful comments. Angus Lamond is a Wellcome Trust Principal Research Fellow. David L. Spector is funded by NIGMS/NIH 42694, NIH/NCI 5PO1CA013106-38, and NIH/ EY 18244.

REFERENCES Beck JS. 1961. Variations in the morphological patterns of “autoimmune” nuclear fluorescence. Lancet 1: 1203– 1205. Bernard D, Prasanth KV, Tripathi V, Colasse S, Nakamura T, Xuan Z, Zhang MQ, Sedal F, Jourdren L, Coulpier F, et al. 2010. A long nuclear retained no-coding RNA regulates synaptogenesis by modulating gene expression. EMBO J doi:10.1038/emboj.2010.199. Boronenkov IV, Loijens JC, Umeda M, Anderson RA. 1998. Phosphoinositide signaling pathways in nuclei are associated with nuclear speckles containing pre-mRNA processing factors. Mol Biol Cell 9: 3547– 3560. Brede G, Solheim J, Prydz H. 2002. PSKH1, a novel splice factor compartment-associated serine kinase. Nucleic Acids Res 30: 5301–5309. Bregman DB, Du L, van der Zee S, Warren SL. 1995. Transcription-dependent redistribution of the large subunit of RNA polymerase II to discrete nuclear domains. J Cell Biol 129: 287 –298. Bridge E, Xia D-X, Carmo-Fonseca M, Cardinali B, Lamond AI, Pettersson U. 1995. Dynamic organization of splicing factors in adenovirus-infected cells. J Virol 69: 281 –290. Brown JM, Green J, das Neves RP, Wallace HA, Smith AJ, Hughes J, Gray N, Taylor S, Wood WG, Higgs DR, et al. 2008. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J Cell Biol 182: 1083–1097. Caceres JF, Misteli T, Screaton GR, Spector DL, Krainer AR. 1997. Role of the modular domains of SR proteins in subnuclear localization and alternative splicing specificity. J Cell Biol 138: 225 –238. Carter KC, Taneja KL, Lawrence JB. 1991. Discrete nuclear domains of poly(A) RNA and their relationship to the functional organization of the nucleus. J Cell Biol 115: 1191– 1202. Cmarko D, Verschure PJ, Martin TE, Dahmus ME, Krause S, Fu XD, van Driel R, Fakan S. 1999. Ultrastructural analysis of transcription and splicing in the cell nucleus after bromo-UTP microinjection. Mol Biol Cell 10: 211–223.

Colwill K, Pawson T, Andrews B, Prasad J, Manley JL, Bell JC, Duncan PI. 1996. The Clk/Sty protein kinase phosphorylates splicing factors and regulates their intranuclear distribution. EMBO J 15: 265 –275. Dostie J, Lejbkowicz F, Sonenberg N. 2000. Nuclear eukaryotic initiation factor 4E (eIF4E) colocalizes with splicing factors in speckles. J Cell Biol 148: 239– 247. Doyle O, Corden JL, Murphy C, Gall JG. 2002. The distribution of RNA polymerase II largest subunit (RPB1) in the Xenopus germinal vesicle. J Struct Biol 140: 154 –166. Eilbracht J, Schmidt-Zachmann MS. 2001. Identification of a sequence element directing a protein to nuclear speckles. Proc Natl Acad Sci 98: 3849–3854. Fakan S. 1994. Perichromatin fibrils are in situ forms of nascent transcripts. Trends Cell Biol 4: 86–90. Fakan S, Bernhard W. 1971. Localisation of rapidly and slowly labelled nuclear RNA as visualized by high resolution autoradiography. Exp Cell Res 67: 129 –141. Fakan S, Nobis P. 1978. Ultrastructural localization of transcription sites and of RNA distribution during the cell cycle of synchronized CHO cells. Exp Cell Res 113: 327 –337. Ferreira JA, Carmo-Fonseca M, Lamond AI. 1994. Differential interaction of splicing snRNPs with coiled bodies and interchromatin granules during mitosis and assembly of daughter cell nuclei. J Cell Biol 126: 11– 23. Fox AH, Lam YW, Leung AK, Lyon CE, Andersen J, Mann M, Lamond AI. 2002. Paraspeckles: A novel nuclear domain. Curr Biol 12: 13– 25. Fox AH, Lamond AI. 2010. Paraspeckles. Cold Spring Harb Perspect Biol doi:10.1101/cshperspect.a000687. Fu X-D. 1995. The superfamily of arginine/serine-rich splicing factors. RNA 1: 663– 680. Gall JG, Bellini M, Wu Z, Murphy C. 1999. Assembly of the nuclear transcription and processing machinery: Cajal bodies (coiled bodies) and transcriptosomes. Mol Biol Cell 10: 4385– 4402. Grande MA, van der Kraan I, de Jong L, van Driel R. 1997. Nuclear distribution of transcription factors in relation to sites of transcription and RNA polymerase II. J Cell Sci 110: 1781– 1791. Gui JF, Lane WS, Fu X-D. 1994a. A serine kinase regulates intracellular localization of splicing factors in the cell cycle. Nature 369: 678 –682. Gui JF, Tronchere H, Chandler SD, Fu X-D. 1994b. Purification and characterization of a kinase specific for the serine- and arginine-rich pre-mRNA splicing factors. Proc Natl Acad Sci 91: 10824–10828. Hall LL, Smith KP, Byron M, Lawrence JB. 2006. Molecular anatomy of a speckle. Anat Rec A Discov Mol Cell Evol Biol 288: 664–675. Hedley ML, Amrein H, Maniatis T. 1995. An amino acid sequence motif sufficient for subnuclear localization of an arginine/serine rich splicing factor. Proc Natl Acad Sci 92: 11524–11528. Herrmann CH, Mancini MA. 2001. The Cdk9 and cyclin T subunits of TAK/P-TEFb localize to splicing factor-rich nuclear speckle regions. J Cell Sci 114: 1491– 1503. Hock R, Wilde F, Scheer U, Bustin M. 1998. Dynamic relocation of chromosomal protein HMG-17 in the nucleus

D.L. Spector and A.I. Lamond is dependent on transcriptional activity. EMBO J 17: 6992– 7001. Huang S, Spector DL. 1991. Nascent pre-mRNA transcripts are associated with nuclear regions enriched in splicing factors. Genes Dev 5: 2288– 2302. Huang S, Spector DL. 1996. Intron-dependent recruitment of pre-mRNA splicing factors to sites of transcription. J Cell Biol 131: 719 –732. Huang S, Deerinck MH, Ellisman MH, Spector DL. 1994. In vivo analysis of the stability and transport of nuclear poly(A)þ RNA. J Cell Biol 126: 877 –899. Hutchinson JN, Ensminger AW, Clemson CM, Lynch CR, Lawrence JB, Chess A. 2007. A screen for nuclear transcripts identifies two linked noncoding RNAs associated with SC35 splicing domains. BMC Genomics 8: 39. Jagatheesan G, Thanumalayan S, Muralikrishna B, Rangaraj N, Karande AA, Parnaik VK. 1999. Colocalization of intranuclear lamin foci with RNA splicing factors. J Cell Sci 112: 4651–4661. Jagiello I, Van Eynde A, Vulsteke V, Beullens M, Boudrez A, Keppens S, Stalmans W, Bollen M. 2000. Nuclear and subnuclear targeting sequences of the protein phosphatase-1 regulator NIPP1. J Cell Sci 21: 3761–3768. Jime´nez-Garcı´a LF, Spector DL. 1993. In vivo evidence that transcription and splicing are coordinated by a recruiting mechanism. Cell 73: 47– 59. Johnson C, Primorac D, McKinstry M, McNeil J, Rowe D, Lawrence JB. 2000. Tracking COL1A1 RNA in osteogenesis imperfecta. splice-defective transcripts initiate transport from the gene but are retained within the SC35 domain. J Cell Biol 150: 417 –432. Kimura H, Sugaya K, Cook PR. 2002. The transcription cycle of RNA polymerase II in living cells. J Cell Biol 159: 777– 782. Ko TK, Kelly E, Pines J. 2001. CrkRS: A novel conserved Cdc2-related protein kinase that colocalises with SC35 speckles. J Cell Sci 114: 2591– 2603. Kojima T, Zama T, Wada K, Onogi H, Hagiwara M. 2001. Cloning of human PRP4 reveals interaction with Clk1. J Biol Chem 276: 32247–32256. Krause S, Fakan S, Weis K, Wahle E. 1994. Immunodetection of Poly(A) Binding Protein II in the Cell Nucleus. Exp Cell Res 214: 75–82. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazett-Jones DP, Hendzel MJ. 2000. Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150: 41–51. Lafarga M, Casafont I, Bengoechea R, Tapia O, Berciano MT. 2009. Cajal’s contribution to the knowledge of the neuronal cell nucleus.. Chromosoma 118: 437–443. Lamond AI, Earnshaw WC. 1998. Structure and function in the nucleus. Science 280: 547– 553. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nat Rev Mol Cell Biol 4: 605 –612. Larsson SH, Charlieu JP, Miyagawa K, Engelkamp D, Rassoulzadegan M, Ross A, Cuzin F, van Heyningen V, Hastie ND. 1995. Subnuclear localization of WT1 in splicing or transcription factor domains is regulated by alternative splicing. Cell 81: 391– 401. Lerner EA, Lerner MR, Janeway CA, Steitz JA. 1981. Monoclonal antibodies to nucleic acid-containing cellular

constituents: Probes for molecular biology and autoimmune disease. Proc Natl Acad Sci 78: 2737–2741. Leser GP, Fakan S, Martin TE. 1989. Ultrastructural distribution of ribonucleoprotein complexes duirng mitosis. snRNP antigens are contained in mitotic granule clusters. Eur J Cell Biol 50: 376–389. Li H, Bingham PM. 1991. Arginine/Serine-rich domains of the su(wa) and tra RNA processing regulators target proteins to a subnuclear compartment implicated in splicing. Cell 67: 335– 342. Li Q, Imataka H, Morino S, Rogers GW Jr, Richter-Cook NJ, Merrick WC, Sonenberg N. 1999. Eukaryotic translation initiation factor 4AIII (eIF4AIII) is functionally distinct from eIF4AI and eIF4AII. Mol Cell Biol 19: 7336– 7346. Matera AG, Ward DC. 1993. Nucleoplasmic organization of small nuclear ribonucleoproteins in cultured human cells. J Cell Biol 121: 715 –727. Melcak I, Cermanova S, Jirsova K, Koberna K, Malinsky J, Raska I. 2000. Nuclear pre-mRNA compartmentalization: Trafficking of released transcripts to splicing factor reservoirs. Mol Biol Cell 11: 497– 510. Mermoud JE, Cohen PTW, Lamond AI. 1994. Regulation of mammalian spliceosome assembly by a protein phosphorylation mechanism. EMBO J 13: 5679– 5688. Mintz PJ, Patterson SD, Neuwald AF, Spahr CS, Spector DL. 1999. Purification and biochemical characterization of interchromatin granule clusters. Embo J 18: 4308–4320. Misteli T. 2001a. Protein dynamics: Implications for nuclear architecture and gene expression. Science 291: 843– 847. Misteli T. 2001b. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Misteli T, Spector DL. 1996. Serine/threonine phosphatase 1 modulates the subnuclear distribution of pre-mRNA splicing factors. Mol Biol Cell 7: 1559– 1572. Misteli T, Spector DL. 1997b. Protein phosphorylation and the nuclear organization of pre-mRNA splicing. Trends Cell Biol 7: 135– 138. Misteli T, Ca´ceres JF, Spector DL. 1997a. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387: 523 –527. Misteli T, Caceres JF, Clement JQ, Krainer AR, Wilkinson MF, Spector DL. 1998. Serine phosphorylation of SR proteins is required for their recruitment to sites of transcription in vivo. J Cell Biol 143: 297– 307. Moen PT Jr, Johnson CV, Byron M, Shopland LS, de la Serna IL, Imbalzano AN, Lawrence JB. 2004. Repositioning of muscle-specific genes relative to the periphery of SC-35 domains during skeletal myogenesis. Mol Biol Cell 15: 197 –206. Monneron A, Bernhard W. 1969. Fine structural organization of the interphase nucleus in some mammalian cells. J Ultrastruct Res 27: 266– 288. Mortillaro MJ, Blencowe BJ, Wei X, Nakayasu H, Du L, Warren SL, Sharp PA, Berezny R. 1996. A hyperphosphorylated form of the large subunit of RNA polymerase II is associated with splicing complexes and the nuclear matrix. Proc Natl Acad Sci 93: 8253–8257. Nakayasu H, Ueda K. 1984. Small nuclear RNA-protein complex anchors on the actin filaments in bovine lymphocyte nuclear matrix. Cell Struct Funct 9: 317 –325.

Nuclear Speckles Nayler O, Hartmann AM, Stamm S. 2000. The ER repeat protein YT521-B localizes to a novel subnuclear compartment. J Cell Biol 150: 949 –962. O’Keefe RT, Mayeda A, Sadowski CL, Krainer AR, Spector DL. 1994. Disruption of pre-mRNA splicing in-vivo results in reorganization of splicing factors. J Cell Biol 124: 249– 260. Pendergrast PS, Wang C, Hernandez N, Huang S. 2002. FBI-1 Can Stimulate HIV-1 Tat Activity and Is Targeted to a Novel Subnuclear Domain that Includes the Tat-P-TEFb-containing Nuclear Speckles. Mol Biol Cell 13: 915 –929. Perraud M, Gioud M, Monier JC. 1979. Intranuclear structures recognized by autoantibodies against ribonucleoproteins: Study on monkey kidney cells in culture using immunofluorescent techniques and immunoelectron microscopy [in French]. Ann Immunol 130: 635 –647. Pessler F, Pendergrast PS, Hernandez N. 1997. Purification and characterization of FBI-1, a cellular factor that binds to the human immunodeficiency virus type 1 inducer of short transcripts. Mol Cell Biol 17: 3786–3798. Phair RD, Misteli T. 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404: 604–609. Potashkin JA, Derby RJ, Spector DL. 1990. Differential distribution of factors involved in pre-mRNA processing in the yeast cell nucleus. Mol Cell Biochem 10: 3524– 3534. Prasanth KV, Sacco-Bubulya P, Prasanth SG, Spector DL. 2003. Sequential entry of components of gene expression machinery into daughter nuclei. Molec Biol Cell 14: 1043– 1057. Price DH. 2000. P-TEFb, a cyclin-dependent kinase controlling elongation by RNA polymerase II. Mol Cell Biol 20: 2629– 2634. Ramo´n y Cajal S. 1910. El nucleo de las celulas piramidales del cerebro humano y de algunos mamiferos. Trab Lab Invest Biol 8: 27–62. Rappsilber J, Ryder U, Lamond AI, Mann M. 2002. Largescale proteomic analysis of the human spliceosome. Genome Res 12: 1231– 1245. Reuter R, Appel B, Rinke J, Lu¨hrmann R. 1985. Localization and structure of snRNPs during mitosis. Immunofluorescent and biochemical studies. Exp Cell Res 159: 63– 79. Sacco-Bubulya P, Spector DL. 2002. Disassembly of interchromatin granule clusters alters the coordination of transcription and pre-mRNA splicing. J Cell Biol 156: 425–436. Saitoh N, Spahr CS, Patterson SD, Bubulya P, Neuwald AF, Spector DL. 2004. Proteomic analysis of interchromatin granule clusters. Mol Biol Cell 15: 3876–3890. Salichs E, Ledda A, Mularoni L, Alba MM, de la Luna S. 2009. Genome-wide analysis of histidine repeats reveals their role in the localization of human proteins to the nuclear speckles compartment. PLoS Genet 5: e1000397. Schul W, van Driel R, de Jong L. 1998. A subset of poly(A) polymerase is concentrated at sites of RNA synthesis and is associated with domains enriched in splicing factors and poly(A) RNA. Exp Cell Res 238: 1 –12. Segalat L, Lepesant JA. 1992. Spatial distribution of the Sm antigen in Drosophila early embryos. Biol Cell 75: 181– 185.

Sharma A, Takata H, Shibahara K, Bubulya A, Bubulya PA. 2010. Son is essential for nuclear speckle organization and cell cycle progression. Mol Biol Cell 21: 650– 663. Shopland LS, Johnson CV, Lawrence JB. 2002. Evidence that all SC-35 domains contain mRNAs and that transcripts can be structurally constrained within these domains. J Struct Biol 140: 131– 139. Shopland LS, Johnson CV, Byron M, McNeil J, Lawrence JB. 2003. Clustering of multiple specific genes and generich R-bands around SC-35 domains: Evidence for local euchromatic neighborhoods. J Cell Biol 162: 981– 990. Smith KP, Moen PT, Wydner KL, Coleman JR, Lawrence JB. 1999. Processing of endogenous pre-mRNAs in association with SC-35 domains is gene specific. J Cell Biol 144: 617–629. Spector DL. 1993. Macromolecular domains within the cell nucleus. Annu Rev Cell Biol 9: 265– 315. Spector DL. 2001. Nuclear bodies. J Cell Sci 114: 2891– 2893. Spector DL. 2006. SnapShot: Cellular bodies. Cell 127: 1071. Spector DL, Smith HC. 1986. Redistribution of U-snRNPs during mitosis. Exp Cell Res 163: 87–94. Spector DL, Fu X-D, Maniatis T. 1991. Associations between distinct pre-mRNA splicing components and the cell nucleus. EMBO J 10: 3467–3481. Spector DL, Lark G, Huang S. 1992. Differences in snRNP localization between transformed and nontransformed cells. Mol Biol Cell 3: 555– 569. Spector DL, O’Keefe RT, Jime´nez-Garcı´a LF. 1993. Dynamics of transcription and pre-mRNA splicing within the mammalian cell nucleus. Cold Spring Harb Symp Quant Biol 58: 799 –805. Spector DL, Schrier WH, Busch H. 1983. Immunoelectron microscopic localization of snRNPs. Biol Cell 49: 1 –10. Swift H. 1959. Studies on nuclear fine structure. Brookhaven Symp Biol 12: 134– 152. Thiry M. 1993. Differential location of nucleic acids within interchromatin granule clusters. Eur J Cell Biol 62: 259 –269. Thiry M. 1995a. Behavior of interchromatin granules during the cell cycle. European J Cell Biol 68: 14–24. Thiry M. 1995b. The interchromatin granules. Histol Histopathol 10: 1035– 1045. Trinkle-Mulcahy L, Sleeman JE, Lamond AI. 2001. Dynamic targeting of protein phosphatase 1 within the nuclei of living mammalian cells. J Cell Sci 114: 4219– 4228. Trinkle-Mulcahy L, Ajuh P, Prescott A, Claverie-Martin F, Cohen S, Lamond AI, Cohen P. 1999. Nuclear organisation of NIPP1, a regulatory subunit of protein phosphatase 1 that associates with pre-mRNA splicing factors. J Cell Sci 112: 157 –168. Tripathi V, Ellis J, Shen Z, Song D, Freier SM, Bennett CF, Sharma A, Bubulya PA, Blencowe BJ, Prasanth SG, et al. 2010. Nuclear-retained non-coding RNA regulates alternative splicing by modulating SR splicing factor phosphorylation. Mol Cell advance online publication doi:101016/jmolcel201008011. Vecerova J, Koberna K, Malinsky J, Soutoglou E, Sullivan T, Stewart CL, Raska I, Misteli T. 2004. Formation of nuclear

D.L. Spector and A.I. Lamond splicing factor compartments is independent of lamins A/C. Mol Biol Cell 15: 4904– 4910. Verheijen R, Kuijpers H, Vooijs P, Van Venrooij W, Ramaekers F. 1986. Distribution of the 70K U1 RNA-associated protein during interphase and mitosis. Correlation with other U RNP particles and proteins of the nuclear matrix. J Cell Sci 86: 173– 190. Visa N, Puvion-Dutilleul F, Harper F, Bachellerie J-P, Puvion E. 1993. Intranuclear distribution of poly A RNA determined by electron microscope in situ hybridization. Exp Cell Res 208: 19– 34. Wansink DG, Schul W, van der Kraan I, van Steensel B, van Driel R, de Jong L. 1993. Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol 122: 283– 293. Wei X, Somanathan S, Samarabandu J, Berezney R. 1999. Three-dimensional visualization of transcription sites and their association with splicing factor-rich nuclear speckles. J Cell Biol 146: 543– 558. Wilusz JE, Sunwoo H, Spector DL. 2009. Long noncoding RNAs: Functional surprises from the RNA world. Genes Dev 23: 1494–1504.

Xing Y, Johnson CV, Dobner PR, Lawrence JB. 1993. Higher level organization of individual gene transcription and RNA splicing. Science 259: 1326–1330. Xing Y, Johnson CV, Moen PT, McNeil JA, Lawrence JB. 1995. Nonrandom gene organization: Structural arrangements of specific pre-mRNA transcription and splicing with SC-35 domains. J Cell Biol 131: 1635–1647. Zeng C, Kim E, Warren SL, Berget SM. 1997. Dynamic relocation of transcription and splicing factors dependent upon transcriptional activity. EMBO J 16: 1401– 1412. Zhao R, Bodnar MS, Spector DL. 2009. Nuclear neighborhoods and gene expression. Curr Opin Genet Dev 19: 172 –179. Zhao K, Wang W, Rando OJ, Xue Y, Swiderek K, Kuo A, Crabtree GR. 1998. Rapid and phosphoinositol-dependent binding of the SWI/SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95: 625 –636. Zhou Z, Licklider LJ, Gygi SP, Reed R. 2002. Comprehensive proteomic analysis of the human spliceosome. Nature 419: 182–185.

The Cajal Body and Histone Locus Body Zehra Nizami, Svetlana Deryusheva, and Joseph G. Gall Department of Embryology, Carnegie Institution for Science, Baltimore, Maryland 21218 Correspondence: [email protected]

The Cajal body (CB) is a nuclear organelle present in all eukaryotes that have been carefully studied. It is identified by the signature protein coilin and by CB-specific RNAs (scaRNAs). CBs contain high concentrations of splicing small nuclear ribonucleoproteins (snRNPs) and other RNA processing factors, suggesting that they are sites for assembly and/or posttranscriptional modification of the splicing machinery of the nucleus. The histone locus body (HLB) contains factors required for processing histone pre-mRNAs. As its name implies, the HLB is associated with the genes that code for histones, suggesting that it may function to concentrate processing factors at their site of action. CBs and HLBs are present throughout the interphase of the cell cycle, but disappear during mitosis. The biogenesis of CBs shows the features of a self-organizing structure.

he CB and the HLB can be considered together for several reasons. First, the canonical marker for the CB, coilin, occurs at high concentration in some HLBs. Second, CBs and HLBs are often physically associated, suggesting some type of interaction between the two. Finally, the very large CBs of the amphibian oocyte, long a favorite for studies of CB function, share properties of both HLBs and CBs.

T

CAJAL BODIES

As the current name suggests, the CB was first described by Ramon y Cajal, the great Spanish neuroanatomist, who in 1906 shared the Nobel Prize with Camillo Golgi for studies on the cellular architecture of the nervous system. Using a silver impregnation technique, Cajal found a small round body within the nuclei of various nerve cells, which he called the accessory body

(cuerpo accessorio) (Cajal 1903; Cajal 1910). His remarkable morphological studies also included observations on splicing speckles, nucleoli, and the nuclear envelope (Lafarga et al. 2009). Bodies that we now know are related to Cajal’s cuerpo accessorio were discovered independently in organisms as diverse as mammals, amphibians, insects, and plants. They were given equally diverse names—coiled bodies in mouse, rat, and human cells (Monneron and Bernhard 1969), Binnenko¨rper or endobodies in insects (Bier et al. 1967), and nucleolus associated bodies in plants (Chamberland and Lafontaine 1993). Order was brought to this somewhat chaotic field with the discovery of a protein named coilin, after its discovery in the coiled bodies of HeLa cells (Andrade et al. 1991; Raska et al. 1991). Antibodies against coilin turned out to be good markers for coiled bodies in vertebrate cells and

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000653 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000653

Z. Nizami, S. Deryusheva, and J.G. Gall

even cross-reacted with the nucleolus associated bodies of the pea, Pisum sativum (Beven et al. 1995). It thus became clear that homologous nuclear organelles existed in a wide range of eukaryotes (Figs. 1 and 2). To recognize this commonality and to regularize the terminology, the name Cajal body was adopted as a general term for a nuclear body that contains coilin (Gall et al. 1999).

Despite its usefulness as a marker for the CB, relatively little is known about coilin as a protein, particularly what biochemical function(s) it might play in the CB. Coilin binds to the survival motor neuron (SMN) protein and to various Sm and Lsm proteins, suggesting that it may function in assembly or modification of snRNPs (Hebert and Matera 2000; Hebert et al. 2001; Xu et al. 2005). There is strong genetic evidence from the mouse, Arabidopsis, and Drosophila that coilin is required for formation of CBs. Knockout of the coilin gene in the mouse leads to a semi-lethal phenotype. Some homozygous individuals die as embryos, and those adults that do survive have significant fertility and fecundity defects (Walker et al. 2009). Cultured cells derived from the knockouts do not have typical CBs. Instead, they show three types of “residual” bodies, each of which contains a subset of CB components (Tucker et al. 2001; Ja´dy et al. 2003). In Arabidopsis the no cajal body 1 (ncb-1) mutant involves a single base substitution in the coilin gene, although it is not certain that the mutant is completely null for coilin (Collier et al. 2006). Homozygous ncb-1 plants are fully viable, but CBs are not detectable with antibodies against other CB components (U2B00 and fibrillarin) or by electron microscopy. In Drosophila two independent null mutants for coilin are fully

Coilin

After its discovery in HeLa cells, coilin quickly became the signature marker for CBs in mammalian cells. Human and mouse coilin are similar in size, MW ¼ 62.6 and 62.3 kDa respectively, and their overall sequence identity is high (Andrade et al. 1991; Tucker et al. 2000). Xenopus coilin is slightly smaller, 59.6 kDa, and its sequence diverges considerably from that of the two mammalian proteins (Tuma et al. 1993). Outside the vertebrates it becomes increasingly difficult to recognize coilin by sequence comparison. What are clearly orthologs of coilin have been described from Arabidopsis (Collier et al. 2006) and Drosophila (Liu et al. 2009), but so far coilin has not been identified in Caenorhabditis, Saccharomyces, or other important nonvertebrate model organisms.

A

B

Figure 1. CBs in a HeLa cell nucleus. (A) DIC image showing four prominent nucleoli and three faintly

detectable CBs (arrowheads). (B) The same nucleus immunostained with antibodies against coilin (green) and fibrillarin (red). The three CBs in this nucleus appear yellow because they stain with both antibodies. The nucleoli stain only for fibrillarin. Fibrillarin is the methyltransferase for 20 -O-methylation of rRNA in the nucleoli and snRNAs in the CBs. Modified from Gall (2003) with permission from Nature Reviews. Bar ¼ 10 mm.

Cajal Body and Histone Locus Body

A

B

Figure 2. A cell from a protoplast of Arabidopsis after

transfection with a construct of U2B00 -mRFP, a component of the U2 snRNP that is enriched in the CB. (A) Fluorescence image showing two brightly labeled CBs in the nucleus. Each is associated with a large, unstained nucleolus. (B) DIC image of the same cells. The nucleoli are now visible (arrowheads) but the CBs are not. Bar ¼ 10 mm. Image kindly provided by Zdravko Lorkovic, Medical University of Vienna, Austria.

viable as homozygotes (Beumer et al. 2008; Liu et al. 2009). Cells from coilin-null flies lack CBs detectable by immunostaining or in situ hybridization for several typical CB components. Thus, in the three organisms in which coilin mutations have been studied, coilin is required for normal CB formation, but neither coilin nor a typical CB is essential for viability. Despite the long-standing use of coilin as a unique marker for CBs, and the critical role that coilin plays in maintaining structural integrity of CBs, we now know that coilin can also occur in HLBs, as discussed later in this article. Small Nuclear Ribonucleoproteins (SnRNPs)

As soon as CBs were identified by immunostaining with antibodies against coilin, it became easy to use other antibodies and in situ hybridization probes to make a catalog of common CB components. It was quickly realized that CBs contain a variety of proteins and RNAs involved in RNA processing, particularly the splicing snRNAs (U1, U2, U4, U5, and U6) and their associated proteins (Lamond and CarmoFonseca 1993; Matera and Ward 1993; Spector 1993). Because splicing itself does not occur in CBs, it was suggested that CBs play some role in the assembly or modification of the splicing snRNPs before their recruitment to the chromosomes. The biogenesis of splicing snRNPs

is a complex process involving both nuclear and cytoplasmic steps (Will and Lu¨hrmann 2001; Patel and Bellini 2008). In brief, transcription of the snRNAs occurs in the nucleus followed by their export to the cytoplasm. In the cytoplasm the monomethyl guanosine cap at the 50 end becomes trimethylated, and each snRNA is assembled into a complex with seven conserved Sm proteins. Finally, the assembled snRNPs are imported back into the nucleus. Because snRNAs in the CB are associated with snRNP proteins and have a trimethylguanosine cap, it is assumed that they have already returned from the cytoplasm. This conclusion is reinforced by kinetic studies showing that newly imported snRNPs travel first to the CB (Sleeman and Lamond 1999) and only later appear in the speckles (interchromatin granule clusters) and finally on the chromosomes, where splicing itself takes place. Modification of specific nucleotides on the snRNAs probably occurs in the CB. It is less clear how much assembly of the splicing machinery takes place in the CB. A role for CBs has been suggested in the final steps of U2 snRNP biogenesis (Nesic et al. 2004) and a particularly compelling case has been made that assembly of the U4/U6-U5 tri-snRNP occurs in the CB (Stanek et al. 2003; Schaffert et al. 2004). Evidence has also been presented that snRNPs recycle through CBs (Stanek et al. 2008). The splicing snRNPs very probably pass from CBs to the nuclear speckles (interchromatin granule clusters) on their way to the sites of RNA synthesis and splicing on the chromosomes. However, the extent to which individual snRNPs are organized into higher complexes in the speckles is not known (Lamond and Spector 2003). A recent study on the amphibian oocyte suggests that snRNPs may be recruited to lampbrush chromosomes independently of their assembly into mature spliceosomes (Patel et al. 2007). If this is generally true for other cells, then CBs may have only a limited role in assembly of snRNPs into higher order complexes. Small CB-associated RNAs (scaRNAs)

An important advance in the understanding of CB function came with the discovery of

Z. Nizami, S. Deryusheva, and J.G. Gall

CB-specific RNAs (scaRNAs). Sca-RNAs are closely related to small nucleolar RNAs (snoRNAs) in both structure and function. Both share the so-called box C/D and box H/ ACA motifs and both are involved in the posttranscriptional modification of other RNAs. Box C/D snoRNAs act as guides for the insertion of 20 -O-methyl groups on specific ribose moieties of rRNA, whereas the box H/ACA class mediates the conversion of specific uridines to pseudouridine (Bachellerie and Cavaille 1997; Tollervey and Kiss 1997). Fibrillarin is the methyl transferase and dyskerin/NAP57/CBF5 is the pseudouridine synthase; each is complexed with three additional proteins to form the active enzyme. ScaRNAs carry out similar reactions on snRNAs. The first discovered and the best-studied scaRNA is U85 (Darzacq et al. 2002; Kiss et al. 2002). It is an unusual guide RNA that mediates two modifications: 20 O-methylation of C45 and pseudouridylation of U46 in human U5 snRNA (C46 and U47 in Drosophila). Cell fractionation and in situ hybridization experiments demonstrate that U85 scaRNA is strongly localized in CBs in both HeLa cells and Drosophila (Figs. 3A and 4B). This localization contrasts with that of its substrate, U5 snRNA, which is concentrated in CBs, but like other snRNAs is widely distributed throughout the nucleus. The localization of U85 and other scaRNAs in the CB differs from the localization of most box C/D and box H/ ACA guide RNAs, which concentrate in the nucleolus. It has been shown that CB localization in vertebrate cells is dependent on a relatively short consensus sequence termed the CAB box (Richard et al. 2003). A related but somewhat different motif has been described for Drosophila scaRNAs. The CAB box of both human and Drosophila scaRNAs binds a conserved WD40-repeat protein, which is required for their CB localization (Tycowski et al. 2009). The specific localization of scaRNAs in the CB suggests that methylation and pseudouridylation of snRNAs occur in the CB, after import of the assembled snRNPs into the nucleus. This hypothesis is strongly supported by experiments in cultured cells showing that artificial substrates of scaRNAs became modified when

U5 + U85

SDMA + Lsm11

A

C

B

D

Figure 3. Malpighian tubule nuclei of Drosophila after

fluorescent in situ hybridization (A and B) or immunostaining (C and D) for CB components. (A) A nucleus from a wild type fly showing two CBs. The CBs appear yellow because they contain both U85 scaRNA (red) and U5 snRNA (green). There is a high level of U5 throughout the nucleoplasm. (B) A nucleus from a coilin-null fly hybridized as in A. A CB is no longer detectable, although U5 label in the nucleoplasm is unaffected. (C) A nucleus from a wild type fly showing a CB immunostained for symmetric dimethylarginine, a marker for snRNP proteins (green), and an HLB labeled for Lsm11, a component of the U7 snRNP (red). (D) A nucleus from a coilin-null fly labeled as in C. A CB is no longer detectable, but the HLB is apparently unaffected. Modified from Liu et al. (2009) with permission from Molecular Biology of the Cell. Bar ¼ 10 mm in A and B, 5 mm in C and D.

targeted to the CB but not when targeted to the nucleolus (Ja´dy et al. 2003). This hypothesis is also consistent with the well-known concentration of fibrillarin in the CB (Fig. 1). At the same time, it is unlikely that modification of snRNAs is limited to CBs, because coilin-null flies, which lack CBs (Fig. 3 B,D; Fig. 4C), have normal levels of scaRNAs, and all of their snRNAs are correctly modified (Deryusheva and Gall 2009). It seems probable that scaRNAs and other CB components normally exist in the nucleoplasm as macromolecular complexes that are too small to be resolved individually by conventional light microscopy. Coilin is essential

Cajal Body and Histone Locus Body

A scaRNA of unusual interest is the RNA component of telomerase, the enzyme responsible for maintaining the ends or telomeres of eukaryotic chromosomes. Telomerase RNA has been shown by in situ hybridization in CBs in human cancer cell lines (Zhu et al. 2003; Ja´dy et al. 2004), but it is at low or undetectable levels in noncancer cells. Telomerase RNA has a box H/ACA motif as well as a CAB box. As with other scaRNAs, the CAB box is required for CB localization (Theimer et al. 2007). The same WD40-repeat protein that binds other scaRNPs is part of the human telomerase holoenzyme and is required for telomere synthesis in HeLa cells (Venteicher et al. 2009).

A

B

D

C

E

F

CBs, Gems, and the SMN Protein

Figure 4. CBs and HLBs in Drosophila. (A) Each

nucleus of the ejaculatory duct of the male fly usually has a single CB, immunostained green with an antibody against coilin, and a single HLB, immunostained red with an antibody against Lsm11. Often the two bodies are close to one another or touching. Bar ¼ 5 mm. (B and C) In Drosophila and many other insects, highly polyploid nurse cells supply proteins and RNAs to the transcriptionally silent oocyte. Their nuclei display prominent CBs and HLBs, here shown after double in situ hybridization for U85 scaRNA (red) and U7 snRNA (green). (B) shows a stage 8 egg chamber of a wild type fly. C shows the same stage from a coilin-null fly. CBs are not detectable in the mutant, whereas HLBs appear unaffected. Bar ¼ 5 mm. (D–F) Nurse cell nuclei from a stage 11 egg chamber showing prominent HLBs that contain coilin. Immunostained with antibodies against Lsm11 (D, red) and coilin (E, green). The overlay in F shows colocalization of Lsm11 and coilin. Modified from Liu et al. (2009) with permission from Molecular Biology of the Cell. Bar ¼ 20 mm.

for these complexes to assemble into detectable CBs, but assembly of such bodies is not required for function, at least not for the scaRNAdependent modification of splicing snRNAs.

An unusually interesting component of CBs is the survival motor neuron protein (SMN). When the intracellular localization of SMN was first studied by immunofluorescence, the protein was seen throughout the cytoplasm and in a nuclear body similar in size to the CB and often close to it. For this reason the body was called the “Gemini of the CB” or simply gem (Liu and Dreyfuss 1996). By chance the HeLa line in which the discovery was made is unusual: In other human cell lines, including different HeLa strains, in primary neurons, and in Drosophila, SMN is colocalized with coilin in the CB (Matera and Frey 1998; Carvalho et al. 1999; Liu et al. 2006). Hence SMN can generally be treated as an important component of the CB, not as a marker for a separate nuclear body. As the name implies, SMN is required in mammals for proper functioning of motor neurons, especially those of the spinal cord. In the mouse and Drosophila, null mutations in the single-copy smn gene are lethal (Miguel-Aliaga et al. 2000; Chan et al. 2003; Rajendra et al. 2007). The situation in humans is somewhat different, because there are two copies of the gene, one of which has an altered splice site that leads to inefficient processing of its transcript. Without going into the somewhat complex genetics of the human smn gene, suffice it to say that mutations often lead to the condition

Z. Nizami, S. Deryusheva, and J.G. Gall

known as spinal muscular atrophy (SMA). SMA occurs in approximately 1 in 6000 newborns and results in early death. Biochemical studies show that SMN in vertebrate cells occurs in a macromolecular complex called the assemblysome, consisting of SMN itself, seven gemins, and several other factors. This complex functions in the cytoplasm as the chaperone for assembly of the splicing snRNAs with the seven-member Sm ring (Meister et al. 2002; Pellizzoni et al. 2002; Shpargel and Matera 2005; Battle et al. 2006). SMN accompanies the assembled snRNPs on their return to the nucleus (Narayanan et al. 2004), but whether SMN has a specific nuclear function in the CB is not known. Expression of GFP-tagged human SMN in budding yeast led to specific localization of the probe in a small structure inside the nucleolus, which the authors called the nucleolar body (Verheggen et al. 2001; Verheggen et al. 2002). Some steps in the maturation of U3 snoRNA also occur in this body. The association with the nucleolus, the targeting of SMN, and the maturation of U3 all suggest that the yeast nucleolar body is equivalent to the CB of more complex eukaryotes. Association of CBs with Specific Gene Loci

Because nucleoli are associated with a specific locus or loci on chromosomes, it is natural to ask whether similar associations exist for CBs and other nuclear organelles. In the case of the CB, there is no evidence that transcription occurs in the body itself and so there is no reason to suppose that a CB corresponds to a highly active gene locus like the nucleolus. Nevertheless, CBs might form at specific loci or travel there by some unknown mechanism as a means of bringing needed factors to those loci. Such a relationship is suggested by the fact that CBs in cultured vertebrate cells show preferential association with snRNA gene loci. CBs in these cells associate not only with U1, U2, and U4 gene clusters but also with the loci of the minor U11 and U12 snRNAs (Smith et al. 1995; Frey et al. 1999; Jacobs et al. 1999; Frey and Matera 2001). Matera suggested that snRNAs in the

CB somehow exert a feedback regulation on snRNA transcription at these gene loci. Whatever the reason for the association, the relationship between CBs and snRNA loci is both dynamic and dependent on transcription, as shown in a recent experimental analysis (Dundr et al. 2007). An array of inducible U2 snRNA genes was cotransfected into cultured cells along with fluorescently labeled coilin. So long as the U2 array was transcriptionally silent, the array showed no specific relationship to CBs in the same nucleus. However, upon induction of transcription, the U2 array moved toward the closest CB and eventually came in contact with it. This remarkable movement was disrupted in the presence of a dominant negative mutant of b-actin, suggesting a role for nuclear actin in repositioning chromosomal loci in response to transcriptional activation. Another special relationship exists between CBs and the telomeres. During most of the cell cycle telomerase RNA is demonstrable only in CBs. However, during S phase, when telomeres are elongated, telomerase RNA can also be detected in a subset of telomeres (Ja´dy et al. 2006; Tomlinson et al. 2006; Tomlinson et al. 2008). These telomeres contain other components of the telomerase holoenzyme, which do not normally accumulate there. In addition, CBs were seen to make transient associations with telomeres during the S phase. These results suggest that specific interactions occur between CBs and telomeres during telomere elongation. The functional relevance remains to be determined. HISTONE LOCUS BODIES

The HLB was described several times in Drosophila and human cells before a clear distinction was made between it and the CB (Frey and Matera 1995; Calvi et al. 1998; Liu et al. 2000; Ma et al. 2000). The existence of two separate bodies within the same nucleus was shown in experiments designed to identify the Drosophila CB in tissues of the fly (Liu et al. 2006). Because Drosophila coilin was not available at that time, our laboratory decided to use two other probes that we thought would serve equally well: the U7

Cajal Body and Histone Locus Body

snRNP and U85 scaRNA. Based on studies of the Xenopus oocyte, the U7 snRNP seemed to be a good marker for the CB (Wu and Gall 1993), and U85 scaRNA had been shown to label the CB in Drosophila S2 cells (Darzacq et al. 2002). To our surprise we found that the U7 snRNP and U85 scaRNA detected two independent nuclear organelles in various tissues of the fly (Fig. 3C,D; Fig. 4A,B). The organelle that contained U85 scaRNA was designated the CB, based on the presence of other common CB components. Because the organelle that contained the U7 snRNP was invariably associated with the histone genes on chromosome 2, we called it the HLB. The CB and HLB lie close together or actually touch one another in many Drosophila nuclei, suggesting some still unknown functional relationship between the two. Studies from other laboratories soon showed that HLBs could be identified as organelles distinct from CBs in the nuclei of cultured mammalian cells (Bongiorno-Borbone et al. 2008; Ghule et al. 2008). Most of the factors that have been shown in HLBs are known to be involved in processing of histone pre-mRNAs. The genes coding for the major histones do not contain introns and hence their pre-mRNAs are not spliced. However, the pre-mRNAs carry a 30 extension that is cut off before export to the cytoplasm (reviewed in Marzluff 2005). The site of cleavage is determined by the U7 snRNP acting in concert with a variety of other factors (Dominski et al. 2005; Kolev and Steitz 2005; Sullivan et al. 2009). In addition to U7 snRNA and the two U7 specific snRNP proteins Lsm10 and Lsm11, a number of other proteins have recently been detected in HLBs, including stem-loop binding protein (SLBP), FLASH, p220NPAT, negative elongation factor (NELF), symplekin, and the unknown protein recognized by mAb MPM-2 (Narita et al. 2007; White et al. 2007; Bongiorno-Borbone et al. 2008; Ghule et al. 2008; Ghule et al. 2009; Sullivan et al. 2009; Yang et al. 2009). The behavior of CBs and HLBs during oogenesis in Drosophila turned out to be particularly interesting. The polyploid nurse cell nuclei in the youngest egg chambers usually

display a single CB and a single HLB. As oogenesis proceeds, the number of HLBs increases until about stage 8, when there are usually 8 – 16 HLBs. As the nurse cells continue to grow, their CBs break up into tiny fragments that eventually disappear. In contrast, the HLBs retain their size and number and begin to accumulate coilin. After the nurse cells have dumped their cytoplasm into the oocyte and their nuclei begin to degenerate, the HLBs are especially prominent. They display the usual HLB markers like the U7 snRNP, but now stain for coilin as brightly as typical CBs in other cells (Fig. 4D – F). The net result is that at the end of oogenesis nurse cell nuclei lack typical CBs but have prominent coilin-containing HLBs. AMPHIBIAN OOCYTE NUCLEI: CBs OR HLBs?

The giant oocyte nucleus or germinal vesicle (GV) of amphibians such as Xenopus houses several thousand extrachromosomal bodies. Among these are 50 – 100 bodies originally called spheres on the basis of their shape, but now referred to as CBs, because they contain a high concentration of coilin (Fig. 5). Although most of these bodies “float” freely in the nucleoplasm, a few are attached to the lampbrush chromosomes at the histone gene loci (Gall et al. 1981; Callan et al. 1991). There is no question that the free and attached bodies are equivalent structures, because they are identical in all features of morphology and molecular composition, including a high concentration of the U7 snRNP and other factors involved in histone pre-mRNA processing (Wu and Gall 1993; Abbott et al. 1999). Thus, by composition and association with the histone genes, these amphibian oocyte bodies are HLBs. It is tempting to equate them with the late-stage HLBs of the Drosophila ovary, which also have a high concentration of coilin. If this analogy is carried further, one might expect to find both CBs and HLBs in early amphibian oocytes, with loss of the CBs during oogenesis. The distribution of scaRNAs should shed light on this issue, because they are not in the HLBs of Drosophila, and hence may provide a more

Z. Nizami, S. Deryusheva, and J.G. Gall

A

B

C

D

Figure 5. A single large nuclear body from a Xenopus

germinal vesicle after immunostaining for coilin (green) and trimethylguanosine (red). Because of their high concentration of coilin, these bodies have been described as CBs. However, they contain the U7 snRNP and some are attached to the histone gene loci, making them HLBs by definition. (A) DIC image showing one speckle inside the CB/HLB and three speckles attached to its periphery. The irregular object to the lower left is an extrachromosomal nucleolus. (B) Green channel, showing that coilin is limited to the CB/HLB, being absent from the speckles. (C) Red channel showing trimethylguanosine at high concentration in the CB/HLB and lower concentration in the speckles. (D) Overlay of B and C. Although trimethylguanosine occurs in the nucleolus (caps on U3 and U8 snoRNAs), its concentration is too low to detect in this image. Modified from Bellini and Gall (1998) with permission from Molecular Biology of the Cell. Bar ¼ 5 mm.

definitive marker to distinguish the two types of bodies. We know that the Xenopus early oocyte contains several types of nuclear bodies (besides the amplified nucleoli), but it is not yet clear what happens to them during the midstages of oogenesis.

DYNAMICS AND ASSEMBLY OF CBs AND HLBs

The rate at which components of a cellular structure turn over can be a useful indicator of metabolic activity and ultimately of function.

Broadly speaking, one can distinguish storage particles, whose components are expected to be relatively immobile, and metabolically active bodies that exchange molecules with their environment on a short time scale. Various studies, especially those involving fluorescence recovery after photobleaching (FRAP), suggest that CBs and HLBs fall into the latter category. FRAP studies performed on CBs in cultured cells show rapid exchange of coilin, fibrillarin, and other CB components between the body and the nucleoplasm, with half-times for recovery of the order of a few minutes (Snaar et al. 2000; Sleeman et al. 2003; Dundr et al. 2004). Somewhat slower kinetics were found for coilin, U7 snRNA, and the TATA-binding protein in the large CBs/HLBs of Xenopus oocytes, of the order of 30 min (Handwerger et al. 2003; Deryusheva and Gall 2004). These studies suggest that macromolecules have relatively short residence times inside CBs and HLBs, consistent with these bodies having metabolic functions independent of simple storage. All subcellular organelles show a certain degree of Brownian movement, depending primarily on their size and the viscosity of the surrounding medium. CBs show Brownian motion, but careful time-lapse measurements made on CBs in both mammalian and plant cells show more complex behavior as well, including anomalous diffusion and associations with chromatin (Boudonck et al. 1999; Platani et al. 2000; Carmo-Fonseca et al. 2002; Platani et al. 2002). Occasionally CBs have been seen to move in a directed manner, even traversing a nucleolus (Boudonck et al. 1999). Despite considerable knowledge about the composition of CBs and HLBs, it remains unclear what function is served by the concentration of their components in structures large enough to be seen by conventional microscopy. Because coilin null mutants either lack CBs entirely (Drosophila and Arabidopsis) or show unusual distribution of CB components, one cannot argue that CBs are absolutely essential for nuclear physiology. A possible scenario is that CBs bring macromolecular components together at higher concentrations than

Cajal Body and Histone Locus Body

ordinarily found in the nucleus, thus favoring reactions that otherwise might proceed at unacceptably slow rates. Alternatively, one could argue that CBs might serve to sequester splicing components and hence act to regulate the rate of RNA processing in the cell. In a sense, it is easier to speculate about the function of HLBs. In this case, factors that process histone pre-mRNAs are concentrated at the sites where the pre-mRNAs are transcribed. Whether the bodies are the actual sites of processing is not known. HLBs are found throughout the cell cycle in Drosophila (Liu et al. 2006; White et al. 2007), even though histone pre-mRNA transcription and processing is limited to the S phase. Without knowing the biological role of nuclear bodies in detail, one can still ask how such bodies assemble and disassemble during the cell cycle. Two broad categories of explanation can be distinguished, one being that a body must be built step by step, using cellular machinery in an energy-dependent fashion, the other being that a body will nucleate spontaneously when and if its macromolecular components occur at high enough concentration (Misteli 2001). A recent study provides strong support for this second self-assembly mechanism. In an experiment designed to ask whether coilin can act as a nucleating site for CB formation, Dundr and his colleagues made a construct in which GFP-coilin was linked to the Lac repressor and then transfected into cells that carried an array of 250 repeats of the Lac operator at a specific chromosome site (Kaiser et al. 2008). As expected, an accumulation of GFP-coilin appeared at the site of the Lac operator array. Remarkably, however, this accumulation not only had the size and shape of a typical CB but it contained a variety of other CB components, including SMN, snRNP proteins, and U85 scaRNA. Thus a genuine CB had formed at the chromosomal locus, morphologically and compositionally identical to other CBs found elsewhere in the same nucleus. To test whether coilin was required for the nucleation process, 17 other CB components were targeted one by one to the Lac operator. In each case a CB was formed essentially

identical to the one formed by coilin. In fact, several other components were more efficient than coilin in terms of the fraction of transfected cells that displayed a CB. These experiments demonstrate clearly that formation of a CB is not a step-wise process involving the hierarchical addition of components starting, for instance, with coilin. Instead assembly of the CB has the hallmarks of a self-organizing system, taking place whenever the concentration of one of the major macromolecules, or macromolecular complexes, is high enough. The process continues, sweeping up other components until some still undefined end point is reached.

CONCLUDING REMARKS

In the 18 years since the identification of coilin as a marker for the CB, this nuclear organelle has gone from an almost completely neglected curiosity to a well-studied nuclear body whose involvement in snRNP biogenesis has been clearly shown. Because CBs are found in all eukaryotes that have been carefully examined, it is probable that new and unexpected roles for them will be uncovered in the near future, when additional genetic and molecular studies are undertaken. The relationship of the newly recognized HLB to the CB remains to be worked out. As additional organisms are examined, it seems likely that the HLB, like the CB, will prove to have a wide phylogenetic distribution. Because of its intimate relationship with the histone genes, the HLB promises to give new insight into the control of histone pre-mRNA processing during the cell cycle.

ACKNOWLEDGMENTS

We thank former members of our laboratory for their contributions to the understanding of CB and HLB structure and function. Original research cited here was supported by National Institutes of Health grant GM 33397 from the National Institute for General Medical Studies. JGG is American Cancer Society Professor of Developmental Genetics.

Z. Nizami, S. Deryusheva, and J.G. Gall

REFERENCES Abbott J, Marzluff WF, Gall JG. 1999. The stem loop binding protein (SLBP1) is present in coiled bodies of the Xenopus germinal vesicle. Mol Biol Cell 10: 487 –499. Andrade LEC, Chan EKL, Raska I, Peebles CL, Roos G, Tan EM. 1991. Human autoantibody to a novel protein of the nuclear coiled body: Immunological characterization and cDNA cloning of p80-coilin. J Exp Med 173: 1407– 1419. Bachellerie J-P, Cavaille J. 1997. Guiding ribose methylation of rRNA. Trends Biochem Sci 22: 257– 261. Battle DJ, Kasim M, Yong J, Lotti F, Lau CK, Mouaikel J, Zhang Z, Han K, Wan L, Dreyfuss G. 2006. The SMN complex: An assembly machine for RNPs. Cold Spring Harb Symp Quant Biol 71: 313– 320. Bellini M, Gall JG. 1998. Coilin can form a complex with the U7 small nuclear ribonucleoprotein. Mol Biol Cell 9: 2987– 3001. Beumer KJ, Trautman JK, Bozas A, Liu J-L, Rutter J, Gall JG, Carroll D. 2008. Efficient gene targeting in Drosophila by direct embryo injection with zinc-finger nucleases. Proc Natl Acad Sci USA 105: 19821– 19826. Beven AF, Simpson GG, Brown JWS, Shaw PJ. 1995. The organization of spliceosomal components in the nuclei of higher plants. J Cell Sci 108: 509–518. Bier K, Kunz W, Ribbert D. 1967. Struktur und Funktion der Oocytenchromosomen und Nukleolen sowie der ExtraDNS wa¨hrend der Oogenese panoistischer und meroistischer Insekten. Chromosoma 23: 214– 254. Bongiorno-Borbone L, De Cola A, Vernole P, Finos L, Barcaroli D, Knight RA, Melino G, De Laurenzi V. 2008. FLASH and NPAT positive but not Coilin positive Cajal Bodies correlate with cell ploidy. Cell Cycle 7: 2357– 2367. Boudonck K, Dolan L, Shaw PJ. 1999. The movement of coiled bodies visualized in living plant cells by the green fluorescent protein. Mol Biol Cell 10: 2297–2307. Cajal SRy. 1903. Un sencillo metodo de coloracion seletiva del reticulo protoplasmatico y sus efectos en los diversos organos nerviosos de vertebrados e invertebrados. Trab Lab Invest Biol Univ Madrid 2: 129 –221. Cajal SRy. 1910. El nu´cleo de las ce´lulas piramidales del cerebro humano y de algunos mamiferos. Trab Lab Invest Biol Univ Madrid 8: 27– 62. Callan HG, Gall JG, Murphy C. 1991. Histone genes are located at the sphere loci of Xenopus lampbrush chromosomes. Chromosoma 101: 245– 251. Calvi BR, Lilly MA, Spradling AC. 1998. Cell cycle control of chorion gene amplification. Genes Dev 12: 734– 744. Carmo-Fonseca M, Platani M, Swedlow JR. 2002. Macromolecular mobility inside the cell nucleus. Trends Cell Biol 12: 491– 495. Carvalho T, Almeida F, Calapez A, Lafarga M, Berciano MT, Carmo-Fonseca M. 1999. The spinal muscular atrophy disease gene product, SMN: A link between snRNP biogenesis and the Cajal (coiled) body. J Cell Biol 147: 715–727. Chamberland H, Lafontaine JG. 1993. Localization of snRNP antigens in nucleolus-associated bodies: Study of plant interphase nuclei by confocal and electron microscopy. Chromosoma 102: 220– 226.

Chan YB, Miguel-Aliaga I, Franks C, Thomas N, Trulzsch B, Sattelle DB, Davies KE, van den Heuvel M. 2003. Neuromuscular defects in a Drosophila survival motor neuron gene mutant. Hum Mol Genet 12: 1367– 1376. Collier S, Pendle A, Boudonck K, van Rij T, Dolan L, Shaw P. 2006. A distant coilin homologue is required for the formation of Cajal bodies in Arabidopsis. Mol Biol Cell 17: 2942–2951. Darzacq X, Ja´dy BE, Verheggen C, Kiss AM, Bertrand E, Kiss T. 2002. Cajal body-specific small nuclear RNAs: A novel class of 20 -O-methylation and pseudouridylation guide RNAs. EMBO J 21: 2746–2756. Deryusheva S, Gall JG. 2004. Dynamics of coilin in Cajal bodies of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 101: 4810–4814. Deryusheva S, Gall JG. 2009. Small Cajal body-specific RNAs of Drosophila function in the absence of Cajal bodies. Mol Biol Cell 20: 5250–5259. Dominski Z, Yang XC, Marzluff WF. 2005. The polyadenylation factor CPSF-73 is Involved in histone-pre-mRNA processing. Cell 123: 37– 48. Dundr M, Hebert MD, Karpova TS, Stanek D, Xu H, Shpargel KB, Meier UT, Neugebauer KM, Matera AG, Misteli T. 2004. In vivo kinetics of Cajal body components. J Cell Biol 164: 831 –842. Dundr M, Ospina JK, Sung MH, John S, Upender M, Ried T, Hager GL, Matera AG. 2007. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J Cell Biol 179: 1095–1103. Frey MR, Matera AG. 1995. Coiled bodies contain U7 small nuclear RNA and associate with specific DNA sequences in interphase human cells. Proc Natl Acad Sci USA 92: 5915–5919. Frey MR, Matera AG. 2001. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J Cell Biol 154: 499 –509. Frey MR, Bailey AD, Weiner AM, Matera AG. 1999. Association of snRNA genes with coiled bodies is mediated by nascent snRNA transcripts. Current Biol 9: 126– 135. Gall JG. 2003. The centennial of the Cajal body. Nature Rev Mol Cell Biol 4: 975–980. Gall JG, Bellini M, Wu Z, Murphy C. 1999. Assembly of the nuclear transcription and processing machinery: Cajal bodies (coiled bodies) and transcriptosomes. Mol Biol Cell 10: 4385– 4402. Gall JG, Stephenson EC, Erba HP, Diaz MO, BarsacchiPilone G. 1981. Histone genes are located at the sphere loci of newt lampbrush chromosomes. Chromosoma 84: 159 –171. Ghule PN, Dominski Z, Lian JB, Stein JL, van Wijnen AJ, Stein GS. 2009. The subnuclear organization of histone gene regulatory proteins and 3’ end processing factors of normal somatic and embryonic stem cells is compromised in selected human cancer cell types. J Cell Physiol 220: 129–135. Ghule PN, Dominski Z, Yang XC, Marzluff WF, Becker KA, Harper JW, Lian JB, Stein JL, van Wijnen AJ, Stein GS. 2008. Staged assembly of histone gene expression machinery at subnuclear foci in the abbreviated cell cycle of human embryonic stem cells. Proc Natl Acad Sci USA 105: 16964–16969.

Cajal Body and Histone Locus Body Handwerger KE, Murphy C, Gall JG. 2003. Steady-state dynamics of Cajal body components in the Xenopus germinal vesicle. J Cell Biol 160: 495 –504. Hebert MD, Matera AG. 2000. Self-association of coilin reveals a common theme in nuclear body localization. Mol Biol Cell 11: 4159–4171. Hebert MD, Szymczyk PW, Shpargel KB, Matera AG. 2001. Coilin forms the bridge between Cajal bodies and SMN, the spinal muscular atrophy protein. Genes Dev 15: 2720– 2729. Jacobs EY, Frey MR, Wu W, Ingledue TC, Gebuhr TC, Gao L, Marzluff WF, Matera AG. 1999. Coiled bodies preferentially associate with U4, U11 and U12 small nuclear RNA genes in interphase HeLa cells but not with U6 and U7 genes. Mol Biol Cell 10: 1653–1663. Ja´dy BE, Bertrand E, Kiss T. 2004. Human telomerase RNA and box H/ACA scaRNAs share a common Cajal bodyspecific localization signal. J Cell Biol 164: 647–652. Ja´dy BE, Darzacq X, Tucker KE, Matera AG, Bertrand E, Kiss T. 2003. Modification of Sm small nuclear RNAs occurs in the nucleoplasmic Cajal body following import from the cytoplasm. EMBO J 22: 1878–1888. Ja´dy BE, Richard P, Bertrand E, Kiss T. 2006. Cell cycledependent recruitment of telomerase RNA and Cajal bodies to human telomeres. Mol Biol Cell 17: 944– 954. Kaiser TE, Intine RV, Dundr M. 2008. De novo formation of a subnuclear body. Science 322: 1713– 1717. Kiss AM, Ja´dy BE, Darzacq X, Verheggen C, Bertrand E, Kiss T. 2002. A Cajal body-specific pseudouridylation guide RNA is composed on two box H/ACA sno-RNA-like domains. Nucleic Acids Res 30: 4643–4649. Kolev NG, Steitz JA. 2005. Symplekin and multiple other polyadenylation factors participate in 3’-end maturation of histone mRNAs. Genes Dev 19: 2583– 2592. Lafarga M, Casafont I, Bengoechea R, Tapia O, Berciano MT. 2009. Cajal’s contribution to the knowledge of the neuronal cell nucleus. Chromosoma 118: 437– 443. Lamond AI, Spector DL. 2003. Nuclear speckles: A model for nuclear organelles. Nature Rev Mol Cell Biol 4: 604– 612. Lamond AI, Carmo-Fonseca M. 1993. Localisation of splicing snRNPs in mammalian cells. Mol Biol Reports 18: 127–133. Liu Q, Dreyfuss G. 1996. A novel nuclear structure containing the survival of motor neurons protein. EMBO J 15: 3555– 3565. Liu J-L, Hebert MD, Ye Y, Templeton DJ, Kung H-J, Matera AG. 2000. Cell cycle-dependent localization of the CDK2-cyclin E complex in Cajal (coiled) bodies. J Cell Sci 113: 1543–1552. Liu J-L, Murphy C, Buszczak M, Clatterbuck S, Goodman R, Gall JG. 2006. The Drosophila melanogaster Cajal body. J Cell Biol 172: 875 –884. Liu J-L, Wu Z, Nizami Z, Deryusheva S, Rajendra TK, Gao H, Beumer KJ, Carroll D, Matera AG, Gall JG. 2009. Coilin is essential for Cajal body organization in Drosophila melanogaster. Mol Biol Cell 20: 1661–1670. Ma T, Van Tine BA, Wei Y, Garrett MD, Nelson D, Adams PD, Wang J, Qin J, Chow LT, Harper JW. 2000. Cell cycle-regulated phosphorylation of p220NPAT by cyclin

E/Cdk2 in Cajal bodies promotes histone gene transcription. Genes Dev 14: 2298– 2313. Marzluff WF. 2005. Metazoan replication-dependent histone mRNAs: A distinct set of RNA polymerase II transcripts. Curr Opin Cell Biol 17: 274–280. Matera AG, Frey MR. 1998. Coiled bodies and gems: Janus or Gemini? Amer J Hum Gen 63: 317–321. Matera AG, Ward DC. 1993. Nucleoplasmic organization of small nuclear ribonucleoproteins in cultured human cells. J Cell Biol 121: 715 –727. Meister G, Eggert C, Fischer U. 2002. SMN-mediated assembly of RNPs: A complex story. Trends Cell Biol 12: 472 –478. Miguel-Aliaga I, Chan YB, Davies KE, van den Heuvel M. 2000. Disruption of SMN function by ectopic expression of the human SMN gene in Drosophila. FEBS Lett 486: 99– 102. Misteli T. 2001. The concept of self-organization in cellular architecture. J Cell Biol 155: 181– 185. Monneron A, Bernhard W. 1969. Fine structural organization of the interphase nucleus in some mammalian cells. J Ultrastruct Res 27: 266– 288. Narayanan U, Achsel T, Lu¨hrmann R, Matera AG. 2004. Coupled in vitro import of U snRNPs and SMN, the spinal muscular atrophy protein. Mol Cell 16: 223 –234. Narita T, Yung TM, Yamamoto J, Tsuboi Y, Tanabe H, Tanaka K, Yamaguchi Y, Handa H. 2007. NELF interacts with CBC and participates in 30 end processing of replicationdependent histone mRNAs. Mol Cell 26: 349 –365. Nesic D, Tanackovic G, Kramer A. 2004. A role for Cajal bodies in the final steps of U2 snRNP biogenesis. J Cell Sci 117: 4423– 4433. Patel SB, Bellini M. 2008. The assembly of a spliceosomal small nuclear ribonucleoprotein particle. Nucleic Acids Res 36: 6482– 6493. Patel SB, Novikova N, Bellini M. 2007. Splicing-independent recruitment of spliceosomal small nuclear RNPs to nascent RNA polymerase II transcripts. J Cell Biol 178: 937 –949. Pellizzoni L. 2007. Chaperoning ribonucleoprotein biogenesis in health and disease. EMBO Rep 8: 340– 345. Pellizzoni L, Yong J, Dreyfuss G. 2002. Essential role for the SMN complex in the specificity of snRNP assembly. Science 298: 1775– 1779. Platani M, Goldberg I, Lamond AI, Swedlow JR. 2002. Cajal body dynamics and association with chromatin are ATPdependent. Nature Cell Biol 4: 502–508. Platani M, Goldberg I, Swedlow JR, Lamond A. 2000. In vivo analysis of Cajal body movement, separation, and joining in live human cells. J Cell Biol 151: 1561–1574. Rajendra TK, Gonsalvez GB, Walker MP, Shpargel KB, Salz HK, Matera AG. 2007. A Drosophila melanogaster model of spinal muscular atrophy reveals a function for SMN in striated muscle. J Cell Biol 176: 831 –841. Raska I, Andrade LEC, Ochs RL, Chan EKL, Chang C-M, Roos G, Tan EM. 1991. Immunological and ultrastructural studies of the nuclear coiled body with autoimmune antibodies. Exp Cell Res 195: 27– 37. Richard P, Darzacq X, Bertrand E, Ja´dy BE, Verheggen C, Kiss T. 2003. A common sequence motif determines the

Z. Nizami, S. Deryusheva, and J.G. Gall Cajal body-specific localisation of box H/ACA scaRNAs. EMBO J 22: 4283– 4293. Schaffert N, Hossbach M, Heintzmann R, Achsel T, Lu¨hrmann R. 2004. RNAi knockdown of hPrp31 leads to an accumulation of U4/U6 di-snRNPs in Cajal bodies. EMBO J 23: 3000– 3009. Shpargel KB, Matera AG. 2005. Gemin proteins are required for efficient assembly of Sm-class ribonucleoproteins. Proc Natl Acad Sci USA 102: 17372–17377. Sleeman JE, Lamond AI. 1999. Newly assembled snRNPs associate with coiled bodies before speckles, suggesting a nuclear snRNP maturation pathway. Current Biol 9: 1065– 1074. Sleeman JE, Trinkle-Mulcahy L, Prescott AR, Ogg SC, Lamond AI. 2003. Cajal body proteins SMN and coilin show differential dynamic behaviour in vivo. J Cell Sci 116: 2039–2050. Smith KP, Carter KC, Johnson CV, Lawrence JB. 1995. U2 and U1 snRNA gene loci associate with coiled bodies. J Cell Biochem 59: 473– 485. Snaar S, Wiesmeijer K, Jochemsen A, Tanke H, Dirks R. 2000. Mutational analysis of fibrillarin and its mobility in living human cells. J Cell Biol 151: 653– 662. Spector DL. 1993. Nuclear organization of pre-mRNA processing. Curr Opin Cell Biol 5: 442 –447. Stanek D, Pridalova-Hnilicova J, Novotny I, Huranova M, Blazikova M, Wen X, Sapra AK, Neugebauer KM. 2008. Spliceosomal small nuclear ribonucleoprotein particles repeatedly cycle through Cajal bodies. Mol Biol Cell 19: 2534– 2543. Stanek D, Rader SD, Klingauf M, Neugebauer KM. 2003. Targeting of U4/U6 small nuclear RNP assembly factor SART3/p110 to Cajal bodies. J Cell Biol 160: 505–516. Sullivan KD, Steiniger M, Marzluff WF. 2009. A core complex of CPSF73, CPSF100, and Symplekin may form two different cleavage factors for processing of poly(A) and histone mRNAs. Mol Cell 34: 322– 332. Theimer CA, Ja´dy BE, Chim N, Richard P, Breece KE, Kiss T, Feigon J. 2007. Structural and functional characterization of human telomerase RNA processing and Cajal body localization signals. Mol Cell 27: 869–881. Tollervey D, Kiss T. 1997. Function and synthesis of small nucleolar RNAs. Curr Opin Cell Biol 9: 337 –342. Tomlinson RL, Abreu EB, Ziegler T, Ly H, Counter CM, Terns RM, Terns MP. 2008. Telomerase reverse transcriptase is required for the localization of telomerase RNA to Cajal bodies and telomeres in human cancer cells. Mol Biol Cell 19: 3793–3800. Tomlinson RL, Ziegler TD, Supakorndej T, Terns RM, Terns MP. 2006. Cell cycle-regulated trafficking of human telomerase to telomeres. Mol Biol Cell 17: 955 –965.

Tucker KE, Berciano MT, Jacobs EY, LePage DF, Shpargel KB, Rossire JJ, Chan EKL, Lafarga M, Conlon RA, Matera AG. 2001. Residual Cajal bodies in coilin knockout mice fail to recruit Sm snRNPs and SMN, the spinal muscular atrophy gene product. J Cell Biol 154: 293 –307. Tucker KE, Massello LK, Gao L, Barber TJ, Hebert MD, Chan EKL, Matera AG. 2000. Structure and characterization of the murine p80 coilin gene, Coil. J Struct Biol 129: 269 –277. Tuma RS, Stolk JA, Roth MB. 1993. Identification and characterization of a sphere organelle protein. J Cell Biol 122: 767 –773. Tycowski KT, Shu MD, Kukoyi A, Steitz JA. 2009. A conserved WD40 protein binds the Cajal body localization signal of scaRNP particles. Mol Cell 34: 47– 57. Venteicher AS, Abreu EB, Meng Z, McCann KE, Terns RM, Veenstra TD, Terns MP, Artandi SE. 2009. A human telomerase holoenzyme protein required for Cajal body localization and telomere synthesis. Science 323: 644– 648. Verheggen C, Lafontaine DLJ, Samarsky D, Mouaikel J, Blanchard J-M, Bordonne´ R, Bertrand E. 2002. Mammalian and yeast U3 snoRNPs are matured in specific and related nuclear compartments. EMBO J 21: 2736–2745. Verheggen C, Mouaikel J, Thiry M, Blanchard J-M, Tollervey D, Bordonne´ R, Lafontaine DLJ, Bertrand E. 2001. Box C/D small nucleolar RNA trafficking involves small nucleolar RNP proteins, nucleolar factors and a novel nuclear domain. EMBO J 20: 5480–5490. Walker MP, Tian L, Matera AG. 2009. Reduced viability, fertility and fecundity in mice lacking the Cajal body marker protein, coilin. PLoS One 4: e6171. White AE, Leslie ME, Calvi BR, Marzluff WF, Duronio RJ. 2007. Developmental and cell cycle regulation of the Drosophila histone locus body. Mol Biol Cell 18: 2491–2502. Will CL, Lu¨hrmann R. 2001. Spliceosomal UsnRNP biogenesis, structure and function. Curr Opin Cell Biol 13: 290 –301. Wu C-HH, Gall JG. 1993. U7 small nuclear RNA in C snurposomes of the Xenopus germinal vesicle. Proc Natl Acad Sci USA 90: 6257–6259. Xu H, Pillai RS, Azzouz TN, Shpargel KB, Kambach C, Hebert MD, Schu¨mperli D, Matera AG. 2005. The Cterminal domain of coilin interacts with Sm proteins and U snRNPs. Chromosoma 114: 155 –166. Yang XC, Burch BD, Yan Y, Marzluff WF, Dominski Z. 2009. FLASH, a proapoptotic protein involved in activation of caspase-8, is essential for 30 end processing of histone premRNAs. Mol Cell 36: 267– 278. Zhu Y, Tomlinson RL, Lukowiak AA, Terns RM, Terns MP. 2003. Telomerase RNA accumulates in Cajal bodies in human cancer cells. Mol Biol Cell 15: 81– 90.

PML Nuclear Bodies Vale´rie Lallemand-Breitenbach1 and Hugues de The´1,2 1

INSERM/CNRS/Universite´ Paris Diderot/Institut Universitaire He´matologie U944/ UMR7212, Laboratoire associe´ de la Ligue Nationale contre le Cancer, Hoˆpital St. Louis, 1, Av. C. Vellefaux 75475 Paris Cedex 10, France

2

Service de Biochimie, Hoˆpital St. Louis, 1, Av. C. Vellefaux 75475 Paris Cedex 10, France

Correspondence: [email protected]

PML nuclear bodies are matrix-associated domains that recruit an astonishing variety of seemingly unrelated proteins. Since their discovery in the early 1960s, PML bodies have fascinated cell biologists because of their beauty and their tight association with cellular disorders. The identification of PML, a gene involved in an oncogenic chromosomal translocation, as the key organizer of these domains drew instant interest onto them. The multiple levels of PML body regulation by a specific posttranslational modification, sumoylation, have raised several unsolved issues. Functionally, PML bodies may sequester, modify or degrade partner proteins, but in many ways, PML bodies still constitute an enigma.

PML BODIES, AN INTRODUCTION

ML nuclear bodies (NBs) are spheres of 0.1 – 1.0 mm in diameter found in most cell-lines and many tissues. They belong to the nuclear matrix, an ill-defined super-structure of the nucleus proposed to anchor and regulate many nuclear functions, including DNA replication, transcription, or epigenetic silencing (Stuurman et al. 1990). The PML protein is the key organizer of these domains that recruits an ever-growing number of proteins, whose only common known feature to date is their ability to be sumoylated (Bernardi and Pandolfi 2007). PML and NBs were proposed to fine-tune a wide variety of processes, through facilitation of partner protein posttranslational modifications (notably sumoylation itself ) resulting in partner sequestration, activation, or degradation. Several

P

NBs subtypes have been defined on morphological bases, which all contain an electron-dense shell and an inner core. PML NBs came to the forefront with the observation that the oncogenic PML/RARA protein disrupts them in a treatment-reversible manner (Daniel et al. 1993; Dyck et al. 1994; Koken et al. 1994; Weis et al. 1994; Zhu et al. 1997). PML NBs are regulated by cellular stress: viral infection (Everett 2006), DNA-damage, transformation (Koken et al. 1995; Terris et al. 1995; Gurrieri et al. 2004), and oxidative stress (Yamada et al. 2001a; Villagra et al. 2006). Moreover, transcription of PML and several genes encoding partner proteins is dramatically enhanced by interferons. Yet, pml-/- mice, which cannot assemble NBs, develop normally and live well, demonstrating that NBs are dispensable for most basic biological functions.

Editors: David Spector and Tom Misteli Additional Perspectives on The Nucleus available at www.cshperspectives.org Copyright # 2010 Cold Spring Harbor Laboratory Press; all rights reserved; doi: 10.1101/cshperspect.a000661 Cite this article as Cold Spring Harb Perspect Biol 2010;2:a000661

V. Lallemand-Breitenbach and H. de The´

Nevertheless, recent data has implicated PML in the control of cellular senescence and stem cell self-renewal, extending the fields of investigation of PML function (Ferbeyre et al. 2000; Pearson et al. 2000; Salomoni and Pandolfi 2002; Ito et al. 2008).

HISTORICAL OVERVIEW

As for several other nuclear structures, electron microscopy and auto-antibodies were the two founding fathers of the field. Work from several pioneers in the early 1960s noted the presence of dense spherical objects by electron microscopy (de The´ et al. 196