Organization of chromatin in the interphase mammalian cell

Micron 36 (2005) 95–108 www.elsevier.com/locate/micron

Review

Organization of chromatin in the interphase mammalian cell Hesam Dehghania, Graham Dellairea, David P. Bazett-Jonesa,b,* a

Programme in Cell Biology, The Research Institute, The Hospital for Sick Children, 555 University Avenue, Toronto, Ont., Canada M5G 1X8 b Department of Biochemistry, University of Toronto, Medical Sciences Building, 1 King’s College Circle, Toronto, Ont., Canada M5S 1A8 Received 20 August 2004; revised 11 October 2004; accepted 12 October 2004

Abstract The use of imaging techniques has become an essential tool in cell biology. In particular, advances in fluorescence microscopy and conventional transmission electron microscopy have had a major impact on our understanding of chromatin structure and function. In this review we attempt to chart the conceptual evolution of models describing the organization and function of chromatin in higher eukaryotic cells, in parallel with the advances in light and electron microscopy over the past 50 years. In the last decade alone, the application of energy filtered transmission electron microscopy (EFTEM), also referred to as electron spectroscopic imaging (ESI), has provided many new insights into the organization of chromatin in the interphase nucleus. Based on ESI imaging of chromatin in situ, we propose a ‘lattice’ model for the organization of chromatin in interphase cells. In this model, the chromatin fibers of 10 and 30 nm diameter observed by ESI, produce a meshwork that accommodates an extensive and distributed interchromosomal (IC) space devoid of chromatin. The functional implications of this model for nuclear activity are discussed. q 2004 Elsevier Ltd. All rights reserved. Keywords: Nucleus; Chromatin; Chromosome; Interphase; Models; Electron spectroscopic imaging; Imaging techniques

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

96

2. Pioneering ultrastructural studies of chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

96

3. Electron spectroscopic imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

98

4. Functional organization of DNA in the nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Chromosome territories . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Interchromosomal (IC) space and perichromatin structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Mind the [resolution] gap: the hazards of inferring chromatin ultrastructure from fluorescence microscopy 4.4. The ‘lattice’ model of interphase chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Regulation of gene transcription in the chromatin lattice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . .

. . . . . .

99 99 100 101 101 104

5. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106

* Corresponding author. Address: Department of Biochemistry, University of Toronto, Medical Sciences Building, 1 King’s College Circle, Toronto, Ont., Canada M5S 1A8. Fax: C1 416 813 2235. E-mail address: [email protected] (D.P. Bazett-Jones). 0968-4328/$ - see front matter q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.micron.2004.10.003

96

H. Dehghani et al. / Micron 36 (2005) 95–108

Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106

1. Introduction Close to a half century has passed since the first experiments were performed that addressed the question of how chromatin is organized in the nucleus. Much of the work was considered to be of peripheral interest because of the descriptive nature of microscopy at the time. In the last few years, however, the importance of chromatin organization and its remodeling during the regulation of nuclear events such as the transcription, replication and repair of DNA, has finally been recognized. As in all other aspects of cell biology, the study of chromatin has largely benefited from the use of imaging techniques. Consequently, our understanding of chromatin and nuclear organization has also benefited from advances in microscopy. In this review we will trace the development and conceptual evolution of models describing chromatin organization that is applicable to a large variety of eukaryotes. These models include those describing the filamentous structure of chromatin, its territorial subnuclear organization, and the perichromatin structures found in the interchromosomal (IC) space with which chromatin interacts. We will emphasize the impact that electron microscopy (EM) has had in this field as well as the advantages offered by electron spectroscopic imaging (ESI) over traditional transmission electron microscopy (TEM). We also propose a ‘lattice’ model to describe the organization of chromatin in situ within interphase cells based on observations using ESI. We conclude with a discussion of the functional implications of the lattice model and the potential of ESI for future work in chromatin and nuclear biology.

2. Pioneering ultrastructural studies of chromatin Before the 1950s, methods for the study of chromatin were limited to light microscopy, enzymatic digestion, and solvent extraction (Olins and Olins, 2003). Since that time, electron microscopy has revolutionized our understanding of much of cell biology, including how chromatin is organized in the cell nucleus. The filamentous nature of chromatin was first observed in the 1950s and 1960s. This paradigm was followed in the 1970s with the discovery of the nucleosome, the repeating chromatin subunit, based on

purified components analyzed by both biochemical and electron microscopy techniques. Finally, through the 1980s to the present, fluorescence microscopy and ultrastructural studies of higher order organization, particularly the observation of chromosome territories, have contributed to a greater appreciation of the importance of the structural compartmentalization of the nucleus. In the 1950s and 1960s, several pioneering electron microscopy studies were carried out, including work on lampbrush chromosomes of amphibian oocytes (Mirsky and Ris, 1951), grasshopper spermatocytes (De Robertis, 1956), frog erythrocytes (Davies and Spencer, 1962), and newt erythrocytes (Gall, 1963). Although these studies were revolutionary, the work suffered from limits in spatial resolution, the inability to obtain thin sections, and inadequate fixation techniques. Nonetheless, De Robertis (1956), for example, showed that chromosomes and chromatin in grasshopper spermatocytes are composed of a filamentous macromolecular component. Because ultramicrotomy had not yet advanced sufficiently, a ‘squash’ or ‘spread’ technique was used in most electron microscopy experiments. For example, blood cells were spread into a very thin layer on the surface of water and the remains of the hypotonically ruptured cells were picked upon carboncoated grids and air dried (Gall, 1963). With these methods, chromatin of interphase cells appeared as cylindrical fibers with an average diameter of 40–60 nm (Fig. 1A). Whereas digestion with ribonuclease did not have an effect, deoxyribonuclease digestion completely destroyed these fibers (Wolfe, 1965). Another study by Wolfe showed that 10 nm fibers were present in un-fixed nuclei. These fibers could be stabilized by fixation so that their diameter was not affected by spreading on water (Wolfe and Martin, 1968). In all, the results of close to two decades of research on the ultrastructure of chromatin could perhaps be summarized in ˚ fiber this sentence from Ris and Kubai (1970); “the 100 A demonstrated by both [these] approaches. must represent the basic unit of inactive chromatin”. The next decade of electron microscopy studies were centered on the structure of chromatin fibers and the repeating subunits along their length. Electron microscopy of thin sections of fixed nuclei revealed threads and granules (Davies and Small, 1968). In rat liver, nuclear regions with condensed and dispersed chromatin threads were observed, where the average width ranged from 10 to 19 nm (Olins "

Woodcock et al., 1976b). (D) EDTA-regressive staining of rat liver cells (reproduced with permission of Academic Press from Monneron and Bernhard, 1969). Ribosomes (r), nucleolus (nu), interchromatin granules (ig), perichromatin granules (/), perichromatin fibrils (–/ j ). (E) and (F) Chromosome painting in human lymphocyte nuclei shown for chromosome 1 (reproduced with permission from Lichter et al., 1988). (G) and (H) Chinese hamster ovary cells in G1, 1 h (G) and 2 h (H) after mitosis. Large arrows point to condensed chromatin and small arrows point to chromonema fibers (reproduced from Belmont and Bruce, 1994 with permission from The Rockefeller University Press).

H. Dehghani et al. / Micron 36 (2005) 95–108

97

Fig. 1. A gallery of chromatin images from key experiments between 1963 and 1994. (A) Fibers from red-cell nucleus of the newt spread on water surface ˚ in diameter prepared from isolated rat (reproduced with permission from Gall, 1963). (B) Linear arrays of spherical chromatin particles (n bodies) about 70 A thymus and chicken erythrocyte nuclei (reproduced with permission from Olins and Olins, 1974). (C) Chromatin fragments prepared from brief digestion with DNase II or micrococcal nuclease and separated on sucrose gradients and negatively stained with 1% uranyl formate. From left to right (3,4, and 5), single, double, and triple arrows point to monomer, dimer, and trimer peaks, respectively, from gradients containing 0.5 M NaCl (reproduced with permission from

98

H. Dehghani et al. / Micron 36 (2005) 95–108

and Olins, 1972). Woodcock (1973) described fibers that ranged from 10 to 30 nm in diameter, and variation in fiber diameter was the result of irregular coiling or folding of single chromatin fibers rather than through being multistranded. In 1974, Olins and Olins, studying purified chromatin, reported linear arrays of spherical chromatin particles (n bodies) about 7 nm in diameter separated by connecting 1.5 nm strands (Fig. 1B). These particles were later named ‘nucleosomes’ (Oudet et al., 1975). Woodcock and colleagues (1976a), also examining purified material, presented evidence suggesting that these particles are repeats of deoxyribonucleoprotein and are not preparation artifacts. This work was followed by the elegant demonstration that the linker DNA is DNase sensitive releasing the nucleosome subunits when digested (Fig. 1C). The first hints of hierarchy in the organization of chromatin would later come from work describing the divalent cation- and salt-dependent compaction of these bead-like nucleoprotein structures (Griffith and Christiansen, 1978), which we now refer to as 10 nm nucleosome fibers. The discovery of chromosome territories (reviewed in Section 4.1) would precede yet another decade of dramatic change in ultrastructural models of chromatin within cells. Fluorescence microscopy of chromosomes, probed by sequence-specific probes showed that DNA of a single chromosome is restricted to a discrete location within the interphase nucleus (Fig. 1E and F). At the resolution limit of the fluorescence microscope, an impression was created that chromosomes exist in a very condensed form and in a tightly enclosed volume. However, transmission electron microscopy revealed different levels of chromatin packing and substantial chromatin-free space throughout the nucleus. Belmont and Bruce in 1994 using non-ionic detergents in polyamine or divalent cation buffers (containing 80 mM KCl and 20 mM NaCl) and uranyl/lead staining, followed de-condensation of chromosomes during G1 of the cell cycle. In early G1 and late G1/early S phases, chromatin fibres, which they termed chromonema fibres, were observed; one class of which was 100–130 nm in diameter, and another of 60–80 nm in diameter (Fig. 1G and H, respectively). In this chromonema model, an approximately 100 nm diameter chromosome fiber folds into a 200–300 nm diameter prophase chromatid, which in turn coils into the metaphase chromosome structure (Belmont et al., 1999). Conventional TEM of these fibres, which was used to characterize the compaction of DNA from interphase to metaphase chromosomes, is facilitated by molecular stains that provide sufficient contrast between relatively thick fibres and the background of the nucleoplasm. Under the conditions used for chemical fixation, much of the soluble nucleoplasmic protein is extracted, thus providing sufficient contrast of the remaining chromatin fibers. In interphase, however, the majority of chromatin exists in a decondensed (euchromatic) state. As a result, euchromatin has little contrast in electron micrographs of samples prepared using the typical fixation and staining conditions with uranyl

acetate alone, or in combination with lead citrate. The use of energy filtered transmission electron microscopy (EFTEM), also referred to as electron spectroscopic imaging (ESI), can overcome this problem when imaging interphase chromatin by deriving contrast in electron micrographs from the elemental content (nitrogen and phosphorous) of the chromatin itself (Bazett-Jones and Hendzel, 1999; Dellaire et al., 2004; Nisman et al., 2004).

3. Electron spectroscopic imaging The principle behind electron spectroscopic imaging (ESI) is electron energy loss spectroscopy, where incident electrons lose specific amounts of energy based on excitations and ionizations of the specimen’s atoms (Fig. 2). To form energy-filtered images, an instrument that acts both as an electron spectrometer and as a lens is required. ESI images are formed with electrons that have lost specific amounts of energy as a result of inner electron shell ionization events. Since only electrons that have interacted with the specimen are used to create an image, the result is a dark field image. The contrast, therefore, is very high, even without the use of heavy atom contrast agents. Particularly when post-phosphorus edge images are recorded, the ‘endogenous stain’ effect of the phosphorus signal from the DNA backbone is responsible for the ability to readily visualize 10 nm chromatin (Dellaire et al., 2004). In spite of the ability of ESI to define the high-resolution biochemical composition of the nucleus, it has not been adopted as a widespread technique due to the increased cost of the imaging filter and the lack of expertise in obtaining and analyzing the energy loss data. Imaging of chromatin in situ with ESI has led us to propose an alternative to the chromonema model. This model and the impact of ESI on our understanding of nuclear organization in general, will be addressed later in this review. ESI is particularly well suited for structural studies of the cell nucleus. Phosphorus and nitrogen levels obtained from element-specific maps can be used to delineate protein— from nucleic acid-based structures (Fig. 2). Besides the information from these two elements, the continuing development of specific metal tags to detect endogenous or exogenously expressed proteins and antibodies will provide schemes for detecting multiple proteins simultaneously (i.e. multiplex detection) (Malecki et al., 2002; Nisman et al., 2004). Recently, we have applied quantum dots as probes for ESI. Quantum dots are luminescent nanocrystals that can be composed of a variety of chemical elements. Their fluorescent properties can be exploited in fluorescence microscopy, and they are ideally suited for imaging with ESI, since the unique elemental composition of quantum dots provides the opportunity for multiple detection of proteins by element-specific imaging. Even with fluorescence labels alone, that is without EM-specific tags, the identity of certain subnuclear structures can often

H. Dehghani et al. / Micron 36 (2005) 95–108

99

Fig. 2. Electron-specimen interactions in the electron microscope and the concept of ESI. To show the ability of ESI in detection of proteins and nucleic acids in biological specimens, the spectroscopic filtering of electrons from ionization of nitrogen and phosphorus are shown. Briefly, electrons collide with atoms in the specimen. This causes production of X-rays which could be used to provide analytical information. Some of the incident electrons scatter and are blocked by the objective aperture, providing bright-filed contrast. Some transmitted electrons that have lost energy enter the spectrometer, where they are separated according to their energy and used to create energy-filtered image.

be determined by correlative fluorescence microscopy and ESI. Correlative microscopy provides the advantage of being able to obtain, with a high success rate, the ultrastructural detail in complexes that are temporally or spatially rare (Ren et al., 2003).

4. Functional organization of DNA in the nucleus 4.1. Chromosome territories Biochemical models of nuclear function are primarily based on diffusion mechanisms. Consequently, the nucleus has been treated as little more than a membrane-bound bag containing a randomly organized genome in a nucleoplasm consisting of freely mobile regulatory factors. Such transacting factors seek out cis-regulatory elements in the DNA, whether for initiating transcription, replication, repair or recombination, through diffusion mechanisms. The simplicity of such models is attractive, but is now recognized to be naive. One of the first clues that the nucleus has a high degree of internal organization was the observation of chromosome territories. Early observations by Rabl in the 1880s (Rabl, 1885) hinted at a defined spatial organization of the genome, and the subject was re-visited in the 1960s

and 1970s (Comings, 1968; Wischnitzer, 1973; Stack, 1977). Interphase chromosomes appeared to be attached to the nuclear membrane at sites of initiation of DNA synthesis and occupied distinct subnuclear territories. Cremer and colleagues (1982) micro-irradiated certain regions of the nucleus of fibroblastoid Chinese hamster cells by laser UV resulting in the incorporate radioactive 3H at the sites of de novo DNA synthesis. These labeled regions in G1 phase could easily be followed by autoradiography after 20–60 h in later phases of the cell cycle. Although this technique was useful to study the dynamics of chromatin subdomains in the interphase nucleus (Cremer et al., 1993), it could not be used to define whole chromosomes in situ or to confirm the existence of chromatin territories. An elegant proof-concept for the territorial model would later come from key experiments by Lichter and colleagues (1988) (Fig. 1E and F). In this study, the application of fluorescence in situ hybridization (FISH) using chromosome-specific DNA probes coupled with fluorescence microscopy (also called ‘chromosome painting’) revealed that individual chromosomes in human fibroblast cells occupied focal territories in the interphase nucleus (Fig. 3), confirming Rabl’s model. Later using the same technique, chromosome territories (also called ‘chromatin domains’) were detected for all human chromosomes (Boyle et al., 2001).

100

H. Dehghani et al. / Micron 36 (2005) 95–108

Fig. 3. Timeline for development of models on the organization of chromatin in the interphase nucleus.

A second component of Rabl’s model, the anaphase– telophase orientation of individual chromosomes in the interphase nucleus (Rabl configuration), was later demonstrated to hold true in Drosophila (Marshall et al., 1996), Trypanosoma, fission yeast, as well as several plants including wheat, but does not occur in mammalian cells (Abranches et al., 1998). Fluorescence in situ hybridization has also been used to study the spatial organization and size of certain chromosome territories in different species (Greaves et al., 2003), during transcription (Croft et al., 1999), and throughout the cell cycle (Bridger et al., 2000). There have also been studies that relate the local changes in the structure of chromatin to changes in the localization of an individual chromosome, whole or in part. For example, Taddei and collaborators (2001) have shown that over-acetylation of histone H4 in pericentric heterochromatin causes the translocation of these domains to the periphery of nuclei in human cell lines. Furthermore, Tajbakhsh and colleagues (2000) have demonstrated that GC-rich/gene-rich segments of the genome display a much higher variability in their chromosomal territories than AT-rich/gene-poor segments, and Boyle and colleagues (2001) have shown that in human cells more generich chromosomes concentrate at the center of the nucleus, whereas the more gene-poor chromosomes localize towards the nuclear periphery. The use of multi-color FISH has also been used to localize genes within chromosome territories (Kurz et al., 1996; Dietzel et al., 1999; Nogami et al., 2000;

Volpi et al., 2000; Mahy et al., 2002a,b; Williams et al., 2002; Ragoczy et al., 2003; Weierich et al., 2003). By using two probes, one specific to the X chromosome and the other to the sequence of the dystrophin gene, for example, Lichter and Cremer (1992) were able to show that this gene is preferentially localized to the exterior of the chromosome (Fig. 3). Fluorescence in situ hybridization has also shown that the chromosomes within prometaphase chromosome rosettes of human fibroblasts and HeLa cells are consistently positioned on opposite sides of the rosette, which suggests that chromosomes are separated into two haploid sets, each derived from one parent (Nagele et al., 1995). 4.2. Interchromosomal (IC) space and perichromatin structures The homogenous appearance of the individual chromosome territories as observed by fluorescence microscopy seemed to imply that the chromatin within the territories is densely packed. Transcription, therefore, would have to take place mainly at the periphery of the territories (Fig. 3; Cremer et al., 2000). The space between the chromosome territories has been referred to as the interchromatin (IC) domain or space. Fine channels within the chromosome territories would connect to the IC domains. Though the fine channels would allow some diffusion of transcription or splicing factors into the interior of chromosome territories, nuclear activities such as transcription or RNA processing

H. Dehghani et al. / Micron 36 (2005) 95–108

would be favored to occur in the IC domain spaces on the periphery of the chromosome territories. Recent observations, however, do not support this model. Macromolecular enzyme complexes (Visser et al., 1998; Verschure et al., 2003) and nascent transcripts detected by thymidine analogues and FISH (Verschure et al., 1999) reveal that transcription is not confined to the periphery, but rather takes place throughout the territories. Also, Mahy and colleagues (2002a), using probes to detect four distal genes on human chromosome 11p13, have demonstrated that transcription is not always confined to the periphery of the chromosome and that transcriptional machinery can easily access the interior of this chromosome territory. As we describe later, high resolution EM and ESI confirm that chromosome territories are not densely packed and in fact are made of networks of chromatin fibers, which should be completely porous to regulatory complexes. The evidence from fluorescence microscopy that transcription does occur within chromosome territories and not only at the IC interface demands a re-interpretation of earlier electron microscopy studies that identified the socalled ‘perichromatin fibrils’. Perichromatin fibrils were first identified by Monneron and Bernhard (1969) using a staining protocol claimed to be specific for RNA (Fig. 1D). The technique, called EDTA-regressive staining, is performed by first over-staining sections with uranyl acetate, followed by chelation of the uranium with EDTA. Uranium bound to DNA and protein is supposedly chelated, whereas uranium bound to RNA is not chelated and therefore contrasts RNA specifically (Bernhard, 1969). Though the stain is not necessarily specific for RNA (Boisvert et al., 2000), the technique does label several nuclear substructures including the nucleolus, Cajal bodies, interchromatin granules, perichromatin granules, and perichromatin fibrils (Monneron and Bernhard, 1969) (Figs. 1D and 3). It was proposed that these fibrils represented RNA transcripts because the numbers of perichromatin fibrils increased in micrographs of liver cells taken from cortisoneinjected animals, and the staining of these fibrils was profoundly diminished by RNase treatment of the sections (Petrov and Bernhard, 1971). In 1975, Nash and colleagues (Nash et al., 1975), using [3H]uridine labelling, demonstrated that the perichromatin fibrils might be the extranucleolar sites of nascent RNA synthesis. With the advent of immuno-electron microscopy it was revealed that the core proteins of heterogeneous nuclear ribonucleoproteins (hnRNPs), small nuclear ribonucleoproteins (snRNPs), and SC-35, all of which are associated with spliceosomes, are also associated with the perichromatin fibrils (Fakan et al., 1984; Puvion et al., 1984; Spector et al., 1991). The fibres were called ‘perichromatin’ because they were observed on the surface of the densest blocks of chromatin (Fakan, 1994). A significant amount of chromatin, however, is not contrasted by this technique, and the chromatin that is negatively contrasted by the technique is condensed heterochromatin, predominantly clustered at the nuclear

101

envelope and peripheral to nucleolar regions. This is not euchromatin or potentially transcribed DNA. Therefore, perichromatin fibrils represent a subset of extra-nucleolar RNA transcripts that are peripheral to blocks of heterochromatin, but do not represent the transcripts that are synthesized throughout chromosome territories, which are not contrasted with this staining technique. The lack of specificity of EDTA-regressive staining for RNA indicates the need for caution in interpreting images derived from this technique. 4.3. Mind the [resolution] gap: the hazards of inferring chromatin ultrastructure from fluorescence microscopy In contrast to the interpretations of fluorescence micrographs (Mascetti et al., 2001), electron spectroscopic imaging of nuclei indicates that chromatin density (or compactness) is not directly related to higher order folding (or condensation) of the chromatin fiber. In fact, the integrated fluorescence intensity of nucleic acid binding dyes such as DAPI, can be misleading when used to infer the condensation level of chromatin for at least two reasons. Firstly, chromatin fibers of 10 and 30 nm diameter can be arranged in close proximity to each other, in such a way that the integrated level of fluorescence by light microscopy would give the perception of higher order folding of chromatin. This misperception of the level of chromatin condensation arises from the disparity in resolution between the light and electron microscope (as demonstrated in Fig. 4B). The lowest unit of resolution in light microscopy (i.e. one pixel with the area of approximately 200 nm! 200 nm) can be resolved by EM to accommodate the width of as few as one and as many as seven 30 nm fibers tightly packed beside each other. Thus, the density of the chromatin fibers within each pixel gives rise to the intensity of DAPI at that pixel and does not reflect the level of condensation of the chromatin fiber itself. Secondly, only a portion of double-helix DNA is accessible to the dye (Myc et al., 1992) and many dyes show sequence-specific preference for binding to nucleic acids (Santisteban et al., 1992). Furthermore, fluorescence from out-of-focus structures will also contribute to the total fluorescence signal. Another important point that needs to be emphasized is that in euchromatic regions of G1 cells, there are no fibres beyond 30 nm when examined at high resolution by EFTEM (Fig. 4C). At lower magnification and resolution, some parts of the de-condensed chromatin have the appearance of distinct thick fibers. At higher magnification (and resolution), however, imaging of the same regions reveals that what appeared to be fibers at low magnification are local concentrations of 30 and 10 nm fibers. 4.4. The ‘lattice’ model of interphase chromatin The application of ESI to the study of nuclear ultrastructure has led to many new insights regarding

102

H. Dehghani et al. / Micron 36 (2005) 95–108

Fig. 4. Disparity in resolution between the light, conventional and energy-filtered electron microscopy. (A) A schematic model of lattice chromatin in an interphase mammalian cell. This model represents a porous structure (right panel) in contrast to densely packed chromosome territories revealed by fluorescence microscopy (left panel). Ten and 30 nm fibers of two adjacent territories intermingle at inter-chromosomal borders. Intra-chromosomal and interchromosomal spaces are part of the unified intra-nuclear space. (B) Limitation of light microscopy to distinguish chromatin condensation (folding) from chromatin density. Whether the number of fibers (of the same diameter) increases (left panel), or an individual fiber enlarges by folding (right panel), the fluorescence microscope is only able to detect an increased level of fluorescence intensity. (C) Basis for misinterpretations of chromatin ultrastructure in electron micrographs. A low magnification, segmented high magnification (C-II), high magnification net phosphorus map (C-III), high magnification overlay of phosphorus and nitrogen maps (C-IV), segmented high magnification (C-V), and a high magnification net phosphorus map (C-VI) of a SK-N-SH cell (human bone marrow neuroblastoma cells) are shown. The two arrows appear to mark a fiber seen at low magnification (I). However, at intermediary (II) and high magnification (V), this structure can be seen to be a local accumulation of condensed chromatin that is discontinuous, and indeed is not a single fiber. This example shows that low magnification electron micrographs are as well limited to accurately represent fibers with low diameter. The ability to map proteins and nucleic acids within a specimen differentiates ESI from conventional electron microscopy. In panels C-III and C-VI net phosphorus maps (that represent nucleic acids) of the specimen have been imaged in different magnifications. Panels C-II, C-IV, and C-V show nitrogen maps (representing proteins) overlaid on the phosphorus maps. Comparing panels C-II with panel C-III and panels C-IV and C-V with panel C-VI clearly shows that there are non-chromosomal proteins that could only be differentiated in ESI images. In conventional electron microscopy these proteins could be accounted as part of the chromatin fiber and erroneously add to the thickness of the nucleic acid fibers. The scale bars are 500 nm (II, III), and 200 nm (IV–VI).

H. Dehghani et al. / Micron 36 (2005) 95–108

103

chromatin organization, as well as the spatial relationship of chromatin to ribonucleoprotein complexes (RNPs) and other protein-based structures. Within a nuclear region, phosphorus and nitrogen maps obtained by ESI can be used to delineate nucleic acid-based structures (red) and proteinbased structures (green) (Fig. 5B and C). This enables the distribution of nucleic acids both within and surrounding chromatin to be resolved without molecular stains that might occlude ultrastructural detail. Similarly, ESI also allows the observation of both chromosomal and nonchromosomal proteins within the nucleus (Figs. 4C, 5F and G). We define non-chromosomal proteins as nitrogen rich protein complexes that are distinct from histones and are primarily associated with dispersed chromatin and generally excluded from condensed chromatin. For example, proteinbased bodies like Promyelocytic leukemia (PML) nuclear bodies are considered accumulations of non-chromosomal proteins. ESI is perhaps the method of choice for the study of interphase chromatin in situ, as in contrast to traditional TEM, it can clearly delineate both the condensed and decondensed chromatin regions of the nucleus (Fig. 5D). In this figure, immunogold labeled RNA polymerase II is only visible in de-condensed chromatin and cannot be detected in heterochromatin or in the nucleolus. Based on the direct observation of chromatin fibers by ESI, which resolves with a much greater capacity the interand intra-chromosomal distances and spaces in the interphase nucleus, we have developed the lattice model of interphase chromatin organization. Using ESI, chromatin appears to be organized as an array or lattice of deoxyribonucleoprotein fibers of 10–30 nm fibers in diameter (Fig. 4A). Large channels between chromosome territories are not apparent. Although fibers belonging to one chromosome are still positioned in a specific subnuclear location and make a chromosome territory, they may intermingle with the fibers of the adjacent territories and form a larger lattice or network of fibers. Due to this specific arrangement of chromatin, the inter- and intra-chromosomal spaces are joined to form an almost contiguous nucleoplasmic space. In the development of this model we have considered the following experimental lines of evidence.

Fig. 5. Transcription of mRNA within chromosomal territories. The enzyme RNA polymerase II has been detected with an antibody that detects the phosphorylated form of the enzyme (phosphorylation at serine 2 of its C-terminal domain), when it is involved in the elongation of mRNA transcription. A rectangular region of a SK-N-SH cell in panel A (low magnification) has been imaged with higher (panels B–D, F and G) magnifications. Panels B, C, and E demonstrate phosphorus map, nitrogen map, and overlay of phosphorus and nitrogen maps, respectively. Panel D schematically demonstrates important structures in panel C. The lattice structure of euchromatin and its porous organization with enormous amounts of chromatin-free space can be clearly differentiated from nucleolus and heterochromatin. Panel B clearly shows that gold-tagged

(1) Macromolecular complexes freely diffuse throughout the nucleus. Chromosome territories are easily and readily accessible to large macromolecules. The fluorescence recovery after photo-bleaching (FRAP) of GFP fusion proteins fused to HMG-17 (a high mobility group protein that binds to nucleosomes), the pre-mRNA splicing factor SF2/ASF, and fibrillarin 3 RNA-polymerase II could only be detected in euchromatin. In panel F nonchromosomal proteins have been segmented and falsely colored as blue. A region of panel F has been imaged with higher magnification in panel G to show accumulation of a kind of non-chromosomal protein in PML nuclear bodies. White arrowheads in panels B and G point to gold-tagged RNA Polymerase II molecules. The scale bar in panels B-F is 500 and in panel G is 200 nm.

104

H. Dehghani et al. / Micron 36 (2005) 95–108

(a rRNA processing protein), have demonstrated that many nuclear proteins move easily and rapidly in an energy-independent manner in the nucleus (Phair and Misteli, 2000). In addition, it has been demonstrated that fluorescently labeled dextran molecules 3 and 10 kDa in size can access all compartments in the nucleus, and a significant fraction of chromatin can be accessed by dextrans as large as 70 kDa (Verschure et al., 2003). Similarly, GFP-tagged TFII-H and RNA polymerase II, which are components of large megadalton complexes, appear to be distributed throughout the nucleus (Verschure et al., 2003). (2) Transcription occurs within chromosomal territories. Transcription of nascent RNA occurs on the periphery and within chromosome territories. Detection of nascent RNA by incorporation of 5-bromouridine 5 0 -triphosphate (BrUTP) combined with FISH-labeling of X chromosomes and chromosome 19 territories revealed that transcription sites are scattered throughout the chromosome territories of active X and 19, but not inactive X (Verschure et al., 1999). Furthermore, transcription from ubiquitous or tissue-specific genes, which are not confined to the chromosome periphery, can occur inside a chromosome domain (Mahy et al., 2002b). As we have shown in Fig. 5, the serine 2phosphorylated form of RNA polymerase II (which is involved in elongation of mRNA transcription) is detected inside territories. Perichromatin fibrils could be ribonucleoproteins that are located in euchromatin. As we discussed, EDTAregressive staining followed by conventional (low resolution) TEM have shown that perichromatin fibrils are present in spaces in the immediate vicinity of the condensed chromatin (Fig. 1D). The high resolution ESI images of chromatin shows that in fact, non-condensed (euchromatic) chromatin always localizes at the immediate periphery of condensed chromatin (Figs. 4 and 5). Since in conventional electron microscopy, noncondensed chromatin was not readily observable, heterochromatin or condensed chromatin was assumed to be chromatin. Thus, perichromatin fibrils could be unique accumulations of RNP molecules associated with the edges of heterochromatin regions, but do not represent the majority of transcripts that are synthesized throughout the interior of chromosome territories (Fig. 5). Such RNA molecules are not well contrasted or resolved by conventional TEM. (3) The majority of chromatin exists as 10 and 30 nm fibres in situ. Ultrastructure of chromatin reveals fibers folded in different packaging orders. The diameter of chromonema fibers imaged by conventional transmission electron microscopy (Belmont and Bruce, 1994), change from 100–130 nm in early G1 nucleus to 60–80 nm in late G1. By ESI, however, we observe a more de-condensed state of chromatin,

consisting mostly fibers with 10 and 30 nm diameters (Bazett-Jones and Hendzel, 1999; Figs. 4 and 5). The lattice model and the model based on chromonema fibers are in agreement in that chromosome territories are not distinctly bounded regions separated by wide interchromatin channels (Cremer and Cremer, 2001). (4) The mammalian nucleus contains large areas of ‘chromosome-free’ space. Chromatin fibers do not have to fold extensively to fit into the interphase nucleus, though high levels of folding have been assumed to occur. For example, this topological problem was characterized by Ris and Kubai (1970) as similar to “[packing] a pipe cleaner 240 km long. [into] a cylinder 8 m long and 1 m in diameter”. Highresolution electron spectroscopic imaging of chromatin, however, reveals considerable amounts of chromatinfree space between the lattice of 10 and 30 nm chromatin fibers. Our calculation (Fig. 6) shows that if all of DNA is folded as a 10 nm fiber, approximately 83.5% of the volume of a typical fibroblast nucleus would be devoid of chromatin fibers. Thus, chromatin fibers do not have to pack as tightly as widely believed to fit into the nucleus. By thinking of chromatin as a lattice within the nucleus, we are also forced to readdress the concept of inter-chromatin domains. In the chromosome territory-interchromatin domain (CT-IC) model (Cremer and Cremer, 2001; Fig. 3), interchromatin domains are large spaces between chromosome territories that make small channels into the middle of densely packed chromosome territories. In the lattice model of chromatin, the inter-chromatin domains are indistinguishable from the intra-territorial spaces and in fact both spaces are part of the nucleoplasmic space (Fig. 4). 4.5. Regulation of gene transcription in the chromatin lattice In the lattice model, chromatin is mainly organized in an open configuration; some chromatin domains might and some might not be permissive to transcription. In fact, many recent lines of evidence emphasize that the control of transcription is not necessarily dependent on a shift between a ‘densely packed’ (higher order) and ‘highly open’ (lower order) structure of chromatin. The following experimental findings for the control of gene transcription provide mechanisms that could potentially work in the lowest order of chromatin packaging, 10 nm fibers. (1) DNA sequence-dependent regulation of transcription. Whether or not a gene is transcribed may depend on its DNA sequence or modifications. It is known that large stretches of repetitive DNA sequences in the genome are associated with condensed pericentric and telomeric chromatin. Also, it has recently been shown that

H. Dehghani et al. / Micron 36 (2005) 95–108

105

Fig. 6. Calculation of the volume of chromatin-free space in the nucleus. (A) The nucleus of a typical human fibroblast, with an average volume of 120 mm3 contains 6!109 bp DNA, which can be represented in 3.75!107, 10 nm fiber subunits. (B) The volume of a 10 nm fiber subunit is 580 nm3. (C) The volume of chromatin is 22 mm3. Thus it occupies only about 16.5% of the nuclear volume.

the existence of short triplet-repeats of DNA (CTG and GAA) in a given gene could confer silencing (Saveliev et al., 2003). It has also been shown in a number of studies that DNA methylation causes repression of transcription (for example in gene imprinting). (2) Regulation of gene expression by histone variants and the post-translational modification of histones. Wrapping of DNA around histones in nucleosomes provides a higher level of hierarchy in control of gene transcription. This level of control depends on enzymes that modify core histones, or are involved in the exchange of core histones by their homologue variants. It is well known that post-translational modification of core histones can change the structure of nucleosomes and facilitate transcription (Marmorstein, 2001). For example, it has been shown that modifications of histone H3 tails are necessary for spatial organization of pericentric heterochromatin (Maison, et al., 2002; Grewal and Moazed, 2003). On the other hand, in Drosophila the substitution of histone H3 with histone H3.3 would immediately activate genes at the site of incorporation (Ahmad and Henikoff, 2002b). The replication-independent deposition of these histone variants might label the active loci, which will inherit an active state in the next cell cycle (Ahmad and Henikoff, 2002a). Also, it has recently been demonstrated that in budding yeast, the incorporation of the H2A.Z variant into nucleosomes is catalyzed by an ATP-driven multi-subunit complex (Mizuguchi et al.,

2004) and protects euchromatin from heterochromatinization (Meneghini et al., 2003). (3) Regulation of gene silencing by both proteins and RNA. Proteins also exist that are involved in silencing and/or the nucleation of heterochromatin. Thiel and colleagues (2004) have categorized proteins that repress gene transcription into two groups of active and passive repressors. Passive repressors either compete with transcriptional activators for DNA binding or form inactive heterodimers with transcriptional activators. The Sp1-like zinc finger proteins, for instance, compete for a common GC-rich DNAbinding site, whereas members of bZIP family of transcription factors make inactive heterodimers. Transcriptional repressors such as Groucho (Fisher and Caudy, 1998) and Daxx (Hollenbach et al., 1999) are also able to bind to specific sets of DNA-binding transcription factors and repress their transcriptional activity. On the other hand, active repressors such as MeCP2 (methyl-CpG-binding protein), REST (repressor element-1 silencing transcription factor), and Rb (retinoblastoma protein) could recruit histone deactylases, or like HP1 (Heterochromatin Protein 1), participate in the structure of heterochromatin (Thiel et al., 2004). It has been shown that tethering of HP1 to euchromatic domains can silence a nearby reporter gene (Li et al., 2003). In mammalian cells, silencing of the X-chromosome is regulated by two RNA transcripts of Xist

106

H. Dehghani et al. / Micron 36 (2005) 95–108

(X chromosome inactive-specific transcript) (Brown et al., 1991) and Tsix (Lee et al., 1999). Besides these structural RNAs that are involved in silencing of large chromosomal stretches, short RNAs are also involved in gene silencing. Short RNAs that are produced by cleavage of dsRNA, bind to homologous transcripts, thereby activating the process of RNA interference (RNAi), which leads to degradation of the transcript. In Drosophila it has been shown that this process is involved in heterochromatin formation, and mutations in the genes encoding the components of this machinery would result in the loss of gene silencing (Pal-Bhadra et al., 2004). And, a RNAi effector complex, isolated from fission yeast, is involved in the assembly of heterochromatin (Verdel et al., 2004). (4) Chromosome/gene-positioning as a means of regulating gene-expression. Perhaps the highest hierarchical level of gene transcription can be defined by where the chromosomes and their constituent genes are located in the nucleus. Although no mechanism for control at this level has been elucidated, there are several studies that demonstrate that the subnuclear location of chromosomes determines the activity of their constituent genes. Several studies have shown an inverse correlation between the localization of genes to centromeric heterochromatin and their transcription. For example, Dernburg and colleagues (1996) in an elegant study inserted a stretch of heterochromatin into one allele of the Brown gene in Drosophila and showed that both the mutant and the wild-type alleles associate with the centromeric heterochromatin, leading to the gene’s transcriptional silencing. On the other hand, it has been shown that enhancer elements are able to keep genes away from centromeric heterochromatin, thereby keeping them active (Francastel et al., 1999). Also, the b-globin locus control region exhibits developmentally conserved nuclear compartments that are mainly dedicated to the transcription of these genes (Palstra et al., 2003). Of course, the preferential localization of genes and enhancer elements needs to be accompanied by the relevant transcription factors. It has also been shown that Ikaros for example, a protein involved in normal development of B, T, and NK cells, associates with centromeric heterochromatin, where several lymophocyte lineage-specific genes localize and become inactivated (Brown et al., 1997). Transcription factors of RUNX1 and RUNX2, which are involved in transcription of hematopoietic and skeletal genes, are also recruited to specific regions of the nucleus (Stein et al., 2004).

5. Concluding remarks Imaging of the nucleus by ESI has revealed new details of chromatin organization within the nucleus. In contrast to

the perception created by fluorescence microscopy of chromosome territories by FISH, ESI reveals that chromosome territories are not densely packed and are completely open to regulatory factors that participate in gene transcription. In light of the data provided by ESI analysis of chromatin in situ, we propose the ‘lattice’ model of chromatin. In this model, chromatin exists as 10 and 30 nm fibres that belong to individual chromosomes and that together create a loose meshwork of chromatin throughout the nucleus that intermingles at the periphery of chromosome territories. Inter- and intra-chromosomal spaces within this meshwork are essentially contiguous and together form the intra-nuclear space. A lattice of 10 and 30 nm chromatin fibers, obviates the need to invoke a shift from higher order organization to a lower condensation state upon activation or induction of a gene locus during interphase.

Acknowledgements We wish to acknowledge the skilled work of Ren Li in preparing and imaging samples by electron microscopy, and Reagan Ching for his aid on calculations of empty spaces in the nucleus. G.D. is a senior postdoctoral fellow of the Canadian Institutes of Health Research (CIHR). The work was funded by operating grants from the Canadian Institutes of Health and Research (FRN14311) and the Natural Sciences and Engineering Research Council of Canada to D.P.B.-J. D.P.B.-J. is the recipient of a Canada Research Chair in Molecular and Cellular Imaging.

References Abranches, R., Beven, A.F., Aragon-Alcaide, L., Shaw, P.J., 1998. Transcription sites are not correlated with chromosome territories in wheat nuclei. Journal of Cell Biology 143 (1), 5–12. Ahmad, K., Henikoff, S., 2002a. Epigenetic consequences of nucleosome dynamics. Cell 111 (3), 281–284. Ahmad, K., Henikoff, S., 2002b. The histone variant H3.3 marks active chromatin by replication-independent nucleosome assembly. Molecular Cell 9 (6), 1191–1200. Bazett-Jones, D.P., Hendzel, M.J., 1999. Electron spectroscopic imaging of chromatin. Methods 17 (2), 188–200. Belmont, A.S., Bruce, K., 1994. Visualization of G1 chromosomes: a folded, twisted, supercoiled chromonema model of interphase chromatid structure. Journal of Cell Biology 127 (2), 287–302. Belmont, A.S., Dietzel, S., Nye, A.C., Strukov, Y.G., Tumbar, T., 1999. Large-scale chromatin structure and function. Current Opinion in Cell Biology 11 (3), 307–311. Bernhard, W., 1969. A new staining procedure for electron microscopical cytology. Journal of Ultrastructure Research 27 (3), 250–265. Boisvert, F.M., Hendzel, M.J., Bazett-Jones, D.P., 2000. Promyelocytic leukemia (PML) nuclear bodies are protein structures that do not accumulate RNA. Journal of Cell Biology 148 (2), 283–292. Boyle, S., Gilchrist, S., Bridger, J.M., Mahy, N.L., Ellis, J.A., Bickmore, W.A., 2001. The spatial organization of human chromosomes within the nuclei of normal and emerin-mutant cells. Human Molecular Genetetics 10 (3), 211–219.

H. Dehghani et al. / Micron 36 (2005) 95–108 Bridger, J.M., Boyle, S., Kill, I.R., Bickmore, W.A., 2000. Re-modelling of nuclear architecture in quiescent and senescent human fibroblasts. Current Biology 10 (3), 149–152. Brown, C.J., Ballabio, A., Rupert, J.L., Lafreniere, R.G., Grompe, M., Tonlorenzi, R., Willard, H.F., 1991. A gene from the region of the human X inactivation centre is expressed exclusively from the inactive X chromosome. Nature 349, 38–44. Brown, K.E., Guest, S.S., Smale, S.T., Hahm, K., Merkenschlager, M., Fisher, A.G., 1997. Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91, 845–854. Comings, D.E., 1968. The rationale for an ordered arrangement of chromatin in the interphase nucleus. American Journal of Human Genetics 20 (5), 440–460. Cremer, T., Cremer, C., 2001. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nature reviews. Genetics 2 (4), 292–301. Cremer, T., Cremer, C., Schneider, T., Baumann, H., Hens, L., KirschVolders, M., 1982. Analysis of chromosome positions in the interphase nucleus of Chinese hamster cells by laser-UV-microirradiation experiments. Human Genetetics 62 (3), 201–209. Cremer, T., Kurz, A., Zirbel, R., Dietzel, S., Rinke, B., Schrock, E., Speicher, M.R., Mathieu, U., Jauch, A., Emmerich, P., 1993. Role of chromosome territories in the functional compartmentalization of the cell nucleus. Cold Spring Harbor Symposia on Quantitative Biology 58, 777–792. Cremer, T., Kreth, G., Koester, H., Fink, R.H., Heintzmann, R., Cremer, M., Solovei, I., Zink, D., Cremer, C., 2000. Chromosome territories, interchromatin domain compartment, and nuclear matrix: an integrated view of the functional nuclear architecture. Critical Reviews in Eukaryotic Gene Expression 10 (2), 179–212. Croft, J.A., Bridger, J.M., Boyle, S., Perry, P., Teague, P., Bickmore, W.A., 1999. Differences in the localization and morphology of chromosomes in the human nucleus. Journal of Cell Biology 145 (6), 1119–1131. Davies, H.G., Small, J.V., 1968. Structural units in chromatin and their orientation on membranes. Nature 217 (134), 1122–1125. Davies, H.G., Spencer, M., 1962. The variation in the structure of erythrocyte nuclei with fixation. Journal of Cell Biology 14, 445–458. Dellaire, G., Nisman, R., Bazett-Jones, D.P., 2004. Correlative light and electron spectroscopic imaging of chromatin in situ. Methods in Enzymology 375, 456–478. Dernburg, A.F., Broman, K.W., Fung, J.C., Marshall, W.F., Philips, J., Agard, D.A., Sedat, J.W., 1996. Perturbation of nuclear architecture by long-distance chromosome interactions. Cell 85, 745–759. De Robertis, E., 1956. Electron microscopic observations on the submicroscopic morphology of the meiotic nucleus and chromosomes. The Journal of Biophysical and Biochemical Cytology 2 (6), 785–796. Dietzel, S., Schiebel, K., Little, G., Edelmann, P., Rappold, G.A., Eils, R., Cremer, C., Cremer, T., 1999. The 3D positioning of ANT2 and ANT3 genes within female X chromosome territories correlates with gene activity. Experimental Cell Research 252 (2), 363–375. Fakan, S., 1994. Perichromatin fibrils are in situ forms of nascent transcripts. Trends in Cell Biology 4 (3), 86–90. Fakan, S., Leser, G., Martin, T.E., 1984. Ultrastructural distribution of nuclear ribonucleoproteins as visualized by immunocytochemistry on thin sections. Journal of Cell Biology 98 (1), 358–363. Fisher, A.L., Caudy, M., 1998. Groucho proteins: transcriptional corepressors for specific subsets of DNA-binding transcription factors in vertebrates and invertebrates. Genes and Development 12 (13), 1931–1940. Francastel, C., Walters, M.C., Groudine, M., Martin, D.I.K., 1999. A functional enhancer suppresses silencing of a transgene and prevents its localization close to centromeric heterochromatin. Cell 99, 259–269. Gall, J., 1963. Chromosome fibers from an interphase nucleus. Science 139, 120–121.

107

Greaves, I.K., Rens, W., Ferguson-Smith, M.A., Griffin, D., Marshall Graves, J.A., 2003. Conservation of chromosome arrangement and position of the X in mammalian sperm suggests functional significance. Chromosome Research 11 (5), 503–512. Grewal, S.I., Moazed, D., 2003. Heterochromatin and epigenetic control of gene expression. Science 301 (5634), 798–802. Griffith, J.D., Christiansen, G., 1978. Electron microscope visualization of chromatin and other DNA–protein complexes. Annual Review of Biophysics and Bioengineering 7, 19–35. Hollenbach, A.D., Sublett, J.E., McPherson, C.J., Grosveld, G., 1999. The Pax3-FKHR oncoprotein is unresponsive to the Pax3-associated repressor hDaxx. EMBO Journal 18 (13), 3702–3711. Kurz, A., Lampel, S., Nickolenko, J.E., Bradl, J., Benner, A., Zirbel, R.M., Cremer, T., Lichter, P., 1996. Active and inactive genes localize preferentially in the periphery of chromosome territories. Journal of Cell Biology 135 (5), 1195–1205. Lee, J.T., Davidow, L.S., Warshawsky, D., 1999. Tsix, a gene antisense to Xist at the X-inactivation centre. Nature Genetics 21, 400–404. Li, Y., Danzer, J.R., Alvarez, P., Belmont, A.S., Wallrath, L.L., 2003. Effects of tethering HP1 to euchromatic regions of the Drosophila genome. Development 130 (9), 1817–1824. Lichter, P., Cremer, T., 1992. Chromosome analysis by non-isotopic in situ hybridization. In: Rooney, D.E., Czepulkowski, B.H. (Eds.), Human Cytogenetics—A Practical Approach. IRL, Oxford, pp. 157–192. Lichter, P., Cremer, T., Borden, J., Manuelidis, L., Ward, D.C., 1988. Delineation of individual human chromosomes in metaphase and interphase cells by in situ suppression hybridization using recombinant DNA libraries. Human Genetics 80 (3), 224–234. Mahy, N.L., Perry, P.E., Bickmore, W.A., 2002a. Gene density and transcription influence the localization of chromatin outside of chromosome territories detectable by FISH. Journal of Cell Biology 159 (5), 753–763. Mahy, N.L., Perry, P.E., Gilchrist, S., Baldock, R.A., Bickmore, W.A., 2002b. Spatial organization of active and inactive genes and noncoding DNA within chromosome territories. Journal of Cell Biology 157 (4), 579–589. Maison, C., Bailly, D., Peters, A.H., Quivy, J.P., Roche, D., Taddei, A., Lachner, M., Jenuwein, T., Almouzni, G., 2002. Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component. Nature Genetics 30 (3), 329–334. Malecki, M., Hsu, A., Truong, L., Sanchez, S., 2002. Molecular immunolabeling with recombinant single-chain variable fragment (scFv) antibodies designed with metal-binding domains. Proceedings of the National Academy of Sciences of the United States of America 99 (1), 213–218. Marmorstein, R., 2001. Protein modules that manipulate histone tails for chromatin regulation. Nature reviews. Molecular Cell Biology 2 (6), 422–432. Marshall, W.F., Dernburg, A.F., Harmon, B., Agard, D.A., Sedat, J.W., 1996. Specific interactions of chromatin with the nuclear envelope: positional determination within the nucleus in Drosophila melanogaster. Molecular Biology of the Cell 7 (5), 825–842. Mascetti, G., Carrara, S., Vergani, L., 2001. Relationship between chromatin compactness and dye uptake for in situ chromatin stained with DAPI. Cytometry 44, 113–119. Meneghini, M.D., Wu, M., Madhani, H.D., 2003. Conserved histone variant H2A.Z protects euchromatin from the ectopic spread of silent heterochromatin. Cell 112 (5), 725–736. Mirsky, A.E., Ris, H., 1951. The composition and structure of isolated chromosomes. The Journal of General Physiology 34 (5), 475–492. Mizuguchi, G., Shen, X., Landry, J., Wu, W.H., Sen, S., Wu, C., 2004. ATP-driven exchange of histone H2AZ variant catalyzed by SWR1 chromatin remodeling complex. Science 303 (5656), 343–348. Monneron, A., Bernhard, W., 1969. Fine structural organization of the interphase nucleus in some mammalian cells. Journal of Ultrastructure Research 27 (3), 266–288.

108

H. Dehghani et al. / Micron 36 (2005) 95–108

Myc, A., Traganos, F., Lara, J., Melamed, M.R., Darzynkiewicz, Z., 1992. Unstainable DNA in cell nuclei. Cytometry 13, 389–394. Nagele, R., Freeman, T., McMorrow, L., Lee, H.Y., 1995. Precise spatial positioning of chromosomes during prometaphase: evidence for chromosomal order. Science 270 (5243), 1831–1835. Nash, R.E., Puvion, E., Bernhard, W., 1975. Perichromatin fibrils as components of rapidly labeled extranucleolar RNA. Journal of Ultrastructure Research 53 (3), 395–405. Nisman, R., Dellaire, G., Ren, Y., Li, R., Bazett-Jones, D.P., 2004. Application of quantum dots as probes for correlative fluorescence, conventional, and energy-filtered transmission electron microscopy. The Journal of Histochemistry and Cytochemistry: Official Journal of the Histochemistry Society 52 (1), 13–18. Nogami, M., Kohda, A., Taguchi, H., Nakao, M., Ikemura, T., Okumura, K., 2000. Relative locations of the centromere and imprinted SNRPN gene within chromosome 15 territories during the cell cycle in HL60 cells. Journal of Cell Science 113 (12), 2157–2165. Olins, D.E., Olins, A.L., 1972. Physical studies of isolated eucaryotic nuclei. Journal of Cell Biology 53 (3), 715–736. Olins, A.L., Olins, D.E., 1974. Spheroid chromatin units (n bodies). Science 183 (122), 330–332. Olins, D.E., Olins, A.L., 2003. Chromatin history: our view from the bridge. Nature reviews. Molecular Cell Biology 4 (10), 809–814. Oudet, P., Gross-Bellard, M., Chambon, P., 1975. Electron microscopic and biochemical evidence that chromatin structure is a repeating unit. Cell 4 (4), 281–300. Pal-Bhadra, M., Leibovitch, B.A., Gandhi, S.G., Rao, M., Bhadra, U., Birchler, J.A., Elgin, S.C., 2004. Heterochromatic silencing and HP1 localization in Drosophila are dependent on the RNAi machinery. Science 303 (5658), 669–672. Palstra, R.J., Tolhuis, B., Splinter, E., Nijmeijer, R., Grosveld, F., de Laat, W., 2003. The beta-globin nuclear compartment in development and erythroid differentiation. Nature Genetics 35 (2), 190–194. Petrov, P., Bernhard, W., 1971. Experimentally induced changes of extranucleolar ribonucleoprotein components of the interphase nucleus. Journal of Ultrastructure Research 35 (3), 386–402. Phair, R.D., Misteli, T., 2000. High mobility of proteins in the mammalian cell nucleus. Nature 404 (6778), 604–609. Puvion, E., Viron, A., Xu, F.X., 1984. High resolution autoradiographical detection of RNA in the interchromatin granules of DRB-treated cells. Experimental Cell Research 152 (2), 357–367. ¨ ber Zelltheilung Morphol. Jahrbuch (10), 214–330. Rabl, C., 1885. U Ragoczy, T., Telling, A., Sawado, T., Groudine, M., Kosak, S.T., 2003. A genetic analysis of chromosome territory looping: diverse roles for distal regulatory elements. Chromosome Research 11 (5), 513–525. Ren, Y., Kruhlak, M.J., Bazett-Jones, D.P., 2003. Same serial section correlative light and energy-filtered transmission electron microscopy. The Journal of Histochemistry and Cytochemistry: Official Journal of the Histochemistry Society 51 (5), 605–612. Ris, H., Kubai, D.F., 1970. Chromosome structure. Annual Review of Genetics 4, 263–294. Santisteban, M.S., Montmasson, M.P., Giroud, F., Ronot, X., Brugal, G., 1992. Fluorescence image cytometry of nuclear DNA content versus chromatin pattern: a comparative study of ten fluorochromes. The Journal of Histochemistry and Cytochemistry: Official Journal of the Histochemistry Society 40 (11), 1789–1797. Saveliev, A., Everett, C., Sharpe, T., Webster, Z., Festenstein, R., 2003. DNA triplet repeats mediate heterochromatin-protein-1-sensitive variegated gene silencing. Nature 422 (6934), 909–913. Spector, D.L., Fu, X.D., Maniatis, T., 1991. Associations between distinct pre-mRNA splicing components and the cell nucleus. EMBO Journal 10 (11), 3467–3481.

Stack, S.M., Brown, D.B., Dewey, W.C., 1977. Visualization of interphase chromosomes. Journal of Cell Science 26, 281–299. Stein, G.S., Lian, J.B., van Wijnen, A.J., Stein, J.L., Javed, A., Montecino, M., Zaidi, S.K., Young, D., Choi, J.Y., Gutierrez, S., Pockwinse, S., 2004. Nuclear microenvironments support assembly and organization of the transcriptional regulatory machinery for cell proliferation and differentiation. Journal of Cellular Biochemistry 91 (2), 287–302. Taddei, A., Maison, C., Roche, D., Almouzni, G., 2001. Reversible disruption of pericentric heterochromatin and centromere function by inhibiting deacetylases. Nature Cell Biology 3 (2), 114–120. Tajbakhsh, J., Luz, H., Bornfleth, H., Lampel, S., Cremer, C., Lichter, P., 2000. Spatial distribution of GC- and AT-rich DNA sequences within human chromosome territories. Experimental Cell Research 255 (2), 229–237. Thiel, G., Lietz, M., Hohl, M., 2004. How mammalian transcriptional repressors work. European Journal of Biochemistry 271, 2855–2862. Verdel, A., Jia, S., Gerber, S., Sugiyama, T., Gygi, S., Grewal, S.I., Moazed, D., 2004. RNAi-mediated targeting of heterochromatin by the RITS complex. Science 303 (5658), 672–676. Verschure, P.J., van der Kraan, I., Manders, E.M., van Driel, R., 1999. Spatial relationship between transcription sites and chromosome territories. Journal of Cell Biology 147 (1), 13–24. Verschure, P.J., van der Kraan, I., Manders, E.M., Hoogstraten, D., Houtsmuller, A.B., van Driel, R., 2003. Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules. EMBO Reports 4 (9), 861–866. Visser, A.E., Eils, R., Jauch, A., Little, G., Bakker, P.J., Cremer, T., Aten, J.A., 1998. Spatial distributions of early and late replicating chromatin in interphase chromosome territories. Experimental Cell Research 243 (2), 398–407. Volpi, E.V., Chevret, E., Jones, T., Vatcheva, R., Williamson, J., Beck, S., Campbell, R.D., Goldsworthy, M., Powis, S.H., Ragoussis, J., Trowsdale, J., Sheer, D., 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. Journal of Cell Science 113 (9), 1565–1576. Weierich, C., Brero, A., Stein, S., von Hase, J., Cremer, C., Cremer, T., Solovei, I., 2003. Three-dimensional arrangements of centromeres and telomeres in nuclei of human and murine lymphocytes. Chromosome Research 11 (5), 485–502. Williams, R.R.E., Board, S., Sheer, D., Ragoussis, J., 2002. Subchromosomal positioning of the epidermal differentiation complex (EDC) in keratinocyte and lymphoblast interphase nuclei. Experimental Cell Research 272, 163–175. Wischnitzer, S., 1973. The submicroscopic morphology of the interphase nucleus. International Review of Cytology 34, 1–48. Wolfe, S.L., 1965. The fine structure of isolated chromosomes. Journal of Ultrastructure Research 12, 104–112. Wolfe, S.L., Martin, P.G., 1968. The ultrastructure and strandedness of chromosomes from two species of Vicia. Experimental Cell Research 50 (1), 140–150. Woodcock, C.L., 1973. Ultrastructure of inactive chromatin. Journal of Cell Biology 59, 368a. Woodcock, C.L., Safer, J.P., Stanchfield, J.E., 1976a. Structural repeating units in chromatin. I. Evidence for their general occurrence. Experimental Cell Research 97, 101–110. Woodcock, C.L., Sweetman, H.E., Frado, L.L., 1976b. Structural repeating units in chromatin. II. Their isolation and partial characterization. Experimental Cell Research 97, 111–119.