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VERSION AUGUST 17, 2000
(CRITICAL REVIEWS IN EUKARYOTIC GENE EXPRESSION, IN PRESS)
CHROMOSOME TERRITORIES, INTERCHROMATIN DOMAIN COMPARTMENT AND
NUCLEAR MATRIX: AN INTEGRATED VIEW OF THE FUNCTIONAL NUCLEAR
ARCHITECTURE
T. Cremer* (1,2), G. Kreth (3,2+), H. Koester (4), R.H.A. Fink (5), R. Heintzmann (3), M. Cremer (1),
I. Solovei (1), D. Zink (1), C. Cremer* (3,2)
1) Institute of Anthropology and Human Genetics, Ludwig Maximilians University, Richard Wagner
Str. 10, D-80333 Munich, Germany
2) Interdisciplinary Center for Scientific Computing (+Graduate College), Ruprecht Karls University,
D-69120 Heidelberg, Germany
3) Applied Optics & Information Processing, Kirchhoff-Institute of Physics, Ruprecht Karls
University, Albert-Ueberle-Str. 3-5, D-69120 Heidelberg, Germany
4) Max-Planck-Institute for Biomedical Research, D-69120 Heidelberg, Germany
5) Physiological Institute, Ruprecht Karls University, D-69120 Heidelberg, Germany
*T. and C. Cremer are corresponding authors
email addresses for correspondence:
T. Cremer : [email protected]; FAX: ++49-89-2180-6719
C. Cremer : [email protected]; Fax: ++49-6221-54-9262
Keywords:
nuclear architecture – nuclear matrix – chromosome territories – interchromatin domain compartment
We dedicate this work to Professor Dr. Dr. h.c. Friedrich Vogel on the occasion of his 75th birthday.
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Summary
Advancements in the specific fluorescent labeling of chromatin in fixed and living human cells in
combination with 3D (three-dimensional) and 4D (four-dimensional, i.e. space plus time) fluorescence
microscopy and image analysis have opened the way for detailed studies of the dynamic, higher order
architecture of chromatin in the human cell nucleus and its potential role in gene regulation. Several
features of this architecture are now well established:
1. Chromosomes occupy distinct territories in the cell nucleus with preferred nuclear locations
although there is no evidence of a rigid suprachromosomal order.
2. Chromosome territories in turn contain distinct chromosome arm domains and smaller chromatin
foci or domains with diameters of some 300 to 800 nm and a DNA content in the order of 1 Mbp.
3. Gene dense, early replicating and gene poor, mid to late replicating chromatin domains exhibit
different higher order nuclear patterns which persist through all stages of interphase. In mitotic
chromosomes early replicating chromatin domains give rise to Giemsa light bands, while mid to
late replicating domains form Giemsa dark bands and C-bands.
In an attempt to integrate these experimental data into a unified view of the functional nuclear
architecture, we present a model of a modular and dynamic chromosome territory organization. We
propose that basically three nuclear compartments exist, an ”open” higher order chromatin
compartment with chromatin domains containing active genes, a ”closed” chromatin compartment
comprising inactive genes, and an interchromatin domain (ICD) compartment (Cremer et al. 1993;
Zirbel et al. 1993) which contains macromolecular complexes for transcription, splicing, DNA-
replication and repair. Genes in ”open”, but not in ”closed” higher order chromatin compartments have
access to transcription and splicing complexes located in the ICD compartment. Chromatin domains
which build the ”open” chromatin compartment are organized in a way which allows the direct contact
of genes and nascent RNA to transcription and splicing complexes, respectively, preformed in the ICD
compartment. In contrast, chromatin domains which belong to the ”closed” compartment are
topologically arranged and compacted in a way that precludes the accessibility of genes to
transcription complexes. We argue that the content of the ICD compartment is highly enriched in DNA
depleted biochemical matrix preparations. The ICD compartment may be considered as the structural
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and functional equivalent of the in vivo nuclear matrix. A matrix in this functional sense is compatible
with but does not necessitate the concept of a 3D nuclear skeleton existing of long, extensively
arborized filaments. In the abscence of unequivocal evidence for such a structural matrix in the nucleus
of living cells we keep an agnostic attitude about its existence and possible properties in maintaining
the higher order nuclear architecture. Quantitative modeling of the 3D and 4D human genome
architecture in situ shows that such an assumption is not necessary to explain presently known aspects
of the higher order nuclear architecture. We expect that the interplay of quantitative modeling and
experimental tests will result in a better understanding of the compartmentalized nuclear architecture
and its functional consequences.
Abbreviations 3D = three dimensional 4D = four dimensional (space plus time) BrdU = bromodeoxyuridine CHRAC = chromatin accessibility complex CldU = chlorodeoxyuridine CLSM = confocal laser scanning microscopy CT = chromosome territory CTAP = chromosome territory anchor protein EM = electron microscopy FEISEM = field emission in lens scanning electron microscopy FISH = fluorescence in situ hybridization GFP = green fluorescent protein hnRNPs = heterogeneous nuclear ribonucleoproteins ICD space or compartment = interchromatin domain IdU = iododeoxyuridine LM = light microscopy or microscopic MLS = multi loop subcompartment PcG = poly comb group PSF = point spread function RNPs = ribonucleoproteins RW/GL model = random walk/ giant loop model SCD = spherical chromatin domain SPDM = spectral precision distance microscopy trxG = trithorax group Xa = active X chromosome Xi = inactive X chromosome
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Introduction
The functional architecture of the cell nucleus is an unsolved enigma of cell biology (for reviews see
(Manuelidis 1985; Jackson 1995; van Driel et al. 1995; Lamond and Earnshaw 1998; Parks and De
Boni 1999; Stein and Berezney 1999; Stein and Berezney 2000). On the topological level, and the
level of over-all geometry, several lines of research have contributed to the study of this problem
during the last decades.
More than a hundred years ago Pfitzner and Balbiani claimed that chromosomes are composed of a
threadlike arrangement of chromatin globules (”Pfitzner-Balbiani Chromatinkugeln”; (Pfitzner 1881;
Waldeyer 1888). August Weismann hypothesized that each chromatin globule is composed of a
different quality of germ plasm, a so called Id, with an unknown, but definite molecular structure (for
review see (Cremer 1985). Carl Rabl and Theodor Boveri suggested that each chromosome in the cell
nucleus of animal cells occupies a distinct territory (Rabl 1885; Boveri 1909). This concept was
abandoned when electron microscopic (EM) studies performed in the fifties and sixties of the 20th
century revealed areas of compacted and dispersed chromatin, respectively, called hetero- and
euchromatin, but failed to distinguish chromosome territories (for review see (Wischnitzer 1973).
(Note: The term euchromatin is often used as a synonym for transcriptionally active chromatin.
However, one should be aware that euchromatin includes both active and potentially active genes.) At
this time only a minority of nuclear researchers argued for a functionally relevant, nonrandom higher
order chromatin architecture (for reviews see Comings, 1968; Vogel and Schroeder, 1974; Okada and
Comings, 1979). Compelling evidence for the existence of chromosome territories was obtained in the
eighties, employing newly developed chromosome painting procedures, and has spurred a growing
interest to study the implications of a territorial chromosome organization in nuclear architecture and
function (for review see (Cremer et al. 1993)). Recently, in vivo labeling of nuclear proteins, DNA and
RNA, have allowed insight into the dynamics of nuclear architecture in the living cell nucleus
(Robinett et al. 1996; Misteli et al. 1997; Kanda et al. 1998; Zink and Cremer 1998a; Zink et al. 1998b;
Sullivan and Shelby 1999; Phair and Misteli 2000).
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Electron microscopic (EM) studies have revealed a wealth of ultrastructural information on basic
features of nuclear architecture, such as the nuclear envelope and its pores, perichromatin fibrils,
interchromatin granules, interchromatin granule associated zones, coiled bodies etc. (Monneron and
Bernhard 1969; Fakan and Puvion 1980; Visa et al. 1993; Puvion and Puvion-Dutilleul 1996).
Unfortunately, chromatin compaction has made it difficult even at the EM resolution level to study
higher order chromatin structures above the level of the beads on a string conformation of nucleosomes
and DNA (Olins and Olins 1974; Kornberg and Lorch 1999). Recently, the introduction of new
techniques, such as cryo-ultra-microtomy or field emission in lens scanning electron microscopy
(FEISEM) allowed the ultrastructural visualization of a complex three-dimensional chromatin
architecture (Woodcock and Horowitz 1995; Ris 1997; Lattanzi et al. 1998; Gobbi et al. 1999). So far,
however, the details of higher order levels have remained a matter of speculation and controversy.
The nuclear matrix has provided a paradigm for a compartmentalized, functional nuclear
architecture (Berezney and Coffey 1974). The biochemical characterization of nuclear matrix
preparations has revealed a wealth of information on matrix associated complexes of proteins and
ribonucleoproteins (RNPs) involved in essential nuclear functions, such as transcription and splicing,
DNA-replication and repair (Stein and Berezney 1999). The generation of specific antibodies has
provided tools to elucidate the nuclear topology and over-all geometric arrangement of such
complexes, but has not yet led to the definition of proteins which constitute the basic network of
branched nuclear matrix core filaments described in EM studies (Nickerson et al. 1997). Notably, the
existence of a topologically/geometrically defined, structurally and/or functionally important in vivo
nuclear matrix has not generally been accepted to date (Hancock 2000; Pederson 2000)
Here, we attempt to integrate presently available experimental results into a unified view of the
functional nuclear architecture. We start with a brief summary of the chromosome territory-
interchromatin domain (ICD) compartment model including a suggested nomenclature. Thereafter, we
review the present experimental evidence for a compartmentalized nuclear architecture and discuss this
evidence in light of models of chromosome territory and nuclear architecture. Being aware of the
inherent challenges of any model which attempts to integrate a large body of experimental data,
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supporters of inductive research strategies have always recommended waiting for a secure factual basis
before proposing a model. However, the distinction between ”facts” and model views, it should be
noted, is necessarily much less clear cut in ongoing experimental research than undergraduate
textbooks often tend to make students believe. Speculative and controversial models that make clear,
experimentally falsifiable predictions are an indispensable part of scientific progress. They trigger the
mind to conceive of new experiments to substantiate or - more likely - to disprove a given model. We
hope that this article will serve this purpose.
Summary of the chromosome territory-interchromatin domain (ICD) compartment model
A summary of the chromosome territory-interchromatin domain (ICD) compartment model may be
helpful for readers to obtain a comprehensive view before looking into the details of experimental
findings and modeling aspects which are described thereafter. The uncertainties and controversies on
higher order chromatin structures in the cell nucleus are reflected by the lack of a generally agreed
upon nomenclature. In meeting discussions we have often noted that participants had different
structures in mind when using terms such as chromosomal or chromatin domains, foci, granules, loops
or fibers. Recently, focal chromatin aggregates with a DNA content in the order of 1 Mbp were
described as a basic feature of chromosome territories studied in nuclei of living cells (see below).
These larger aggregates are possibly built up from chromatin loops in the order of 100 kbp. Under
these circumstances we found it most appropriate to use the term chromatin domain together with an
indication of the order of DNA content in each case.
Chromatin structure modules
We propose a hierarchical order of chromatin structures (Fig. 1) consisting of
1. chromosome territories (ca. 50 - 230 Mbp for human chromosome territories in G1 nuclei and ca.
100 and 460 Mbp, respectively, for replicated chromosomes in G2 nuclei);
2. ~1-Mbp chromatin domains (ranging from a few hundred kbp to several Mbp; corresponding
names in the literature: subchromosomal foci, chromatin foci, chromatin granules, multiloop
subcompartments) connected by chromatin linkers; and
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3. ~100-kbp chromatin domains (range from ca. 30-200 kbp, comprising ca. 150-1000 nucleosomes
and one to several genes; corresponding names: chromosomal domains, chromatin loops).
This proposed view of higher order chromatin organization based on ~100-kbp and ~1-Mbp chromatin
domains is certainly strongly oversimplified. Other authors have argued for a hierarchy of thinner and
thicker chromatin fibers (Belmont and Bruce 1994). Such a hierarchy possibly co-exists with or is to
some extent equivalent to a hierarchy of chromatin domains. Our main reason to adopt, for the time
being, the proposed view (1.-3., above) emphasizing chromatin domains of these two sizes is a
pragmatic one: this view has provided a straightforward approach for the quantitative modeling of
chromosome territories in a way that is consistent with presently available experimental data (see
below).
A mammalian diploid cell nucleus in G1 contains a total of ca. 6350 Mbp DNA (Morton, 1991).
This means that such a nucleus harbors some six thousand ~1-Mbp chromatin domains and some sixty
thousand ~100-kbp chromatin domains. These chromatin structures together build up the chromatin
compartment of the cell nucleus and may persist during the cell cycle and in terminally differentiated
cells. While ~100-kbp chromatin loops have been suggested to represent structural blocks of the
eukaryotic genome (Razin 1999), we assume that still higher levels of a structure-function hierarchy
exist in the cell nucleus. We hypothesize that chromosome territory anchor proteins (CTAPs) required
to maintain the structural integrity of chromosome territories (Ma et al., 1999) are located in the
interior of ~1-Mbp chromatin domains and further that these chromatin domains can perform different
functions at different times by assembling different protein complexes for transcription, DNA
replication and repair. Depending on its association with specific protein complexes, a ~1-Mbp
chromatin domain can be functionally defined, e.g., as a replication domain (equivalent names:
replication site, replication focus), a transcription domain or a repair domain, although different
~100 kbp chromatin domains which constitute a given ~1-Mbp domain may be regulated
independently and functional processes may at least occasionally occur in a given chromatin domain
simultaneously.
Open and Closed Chromatin domains
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We distinguish ”open” from ”closed” ~100 kbp chromatin domains. Although the concept of
"closed” and "open” chromatin configurations has been widely discussed and is central to
topological/geometrical models of gene regulation (Eissenberg et al. 1985; Hebbes et al. 1994), the
structural basis for these configurations has remained elusive. To emphasize this point we will put the
terms ”closed” and ”open” in quotation marks throughout the text. ”Open” chromatin domains should
possess a configuration such that functionally relevant chromatin sites, e.g., transcription factor
binding sites, are easily accessible for large functional protein aggregates located in the inter-
chromatin domain (ICD) space. ”Closed” chromatin domains , in contrast, should maintain a
configuration such that binding sites located in the interior of these domains are not accessible for
large macromolecule complexes located in the ICD space. >From this, the ”functional surface” of a
chromatin domain may be defined as the smallest enveloping surface of the domain which cannot be
passed by a given macromolecular complex. Note that ”closed” chromatin domain and ”functional
surface” are defined as relative terms, i.e. with regard to the size of excluded macromolecules. Instead
of a two-dimensional surface one may also consider a three-dimensional border zone at the periphery
of a chromatin domain which is accessible in depth for a certain protein complex in contrast to the
chromatin domain interior. In the absence of detailed knowledge we use ”chromatin surface” for
simplicity, but imply the possibility of such a border zone as well. A chromatin domain which is
”closed” with regard, e.g., to a large, preformed transcription complex or to a spliceosome may still
allow individual proteins or small protein aggregates to penetrate into the chromatin domain interior,
e.g. a certain hormone receptor proteins and associated histone acetyltransferases required for
chromatin opening (Berger 1999). ”Open” and ”closed” chromatin domains, respectively, likely form
transcriptionally active and inactive higher order nuclear compartments (Brown et al. 1997; Wei et al.
1998; Brown et al. 1999; Sadoni et al. 1999; Wei et al. 1999).
Interchromatin domain compartment
A three-dimensionally interconnected nuclear space, called the interchromatin domain (ICD)
space, estends between higher order chromatin structures. This space together with its content and its
boundaries (provided by the surfaces of chromatin domains) is called the ICD compartment (Fig. 1). It
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was first assumed that the ICD compartment starts at nuclear pores and extends between chromosome
territories (Cremer et al. 1993; Zirbel et al. 1993). Later we considered that the ICD compartment has
branches that further extend into the interior of chromosome territories (Cremer et al. 1995) and
expand between neighbouring ~1-Mbp chromatin domains of a given chromosome territory and with
its finest branches between neighbouring ~100-kbp chromatin domains. We hypothesize that the
content of the ICD space is highly enriched in biochemical DNA-depleted matrix preparations and
comprises complex aggregates composed of many proteins that serve nuclear functions such as
transcription, splicing, DNA replication and repair. Protein aggregates of a size that precludes their
diffusion into the interior of ”closed” chromatin domains may serve as storage aggregates from which
individual components can be released and diffuse into the chromatin interior. Alternatively, functional
protein complexes, e.g. transcription complexes preformed in the ICD space, may attach to specific
DNA binding sites exposed at the surface of chromatin domains. For reasons detailed below, we
propose the term ”in vivo nuclear matrix” to designate both the content and dynamic, structural
organization of the ICD space and CTAPs in the nucleus of the living cell. Note that CTAPs are likely
not located in the ICD space but in the interior of chromatin domains (see below).
The matrix view of nuclear architecture
Since its discovery (Berezney and Coffey 1974) the nuclear matrix has been characterized as a highly
complex nuclear structure with a wealth of functionally important associations. The biochemical
analysis of the nuclear matrix revealed that hnRNP proteins are major protein components (Stratling
and Yu 2000). The nuclear matrix is associated with active genes and essentially involved in
transcription, splicing, DNA replication and DNA repair (for review see (Berezney et al. 1995). The
obvious, but still controversial conclusion is that this ”chromatin free” nuclear matrix is a principal
constituent of the functional nuclear architecture. Electron microscopy has provided evidence for a
three-dimensional network of branching 10 nm core filaments (Nickerson et al. 1997). These core
filaments may provide attachment sites for (ribonucleo-) protein complexes with distinct functional
properties, but are still biochemically ill-defined. It has not been possible to date to raise antibodies
that specifically and exclusively stain this network of core filaments.
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The existence of a permanent nuclear skeleton in the living cell nucleus and its possible role in the
organization of chromatin has become a focus of controversy for many years. It has often been argued
that biochemical nuclear matrix preparations represent an artifact comprising a wealth of proteins and
protein complexes freely diffusable in the nuclear sap. Components found adjacent to each other in an
insoluble nuclear matrix preparation may reflect chance aggregations that occurred only during
preparation steps but are irrelevant for the in vivo situation. This criticism is reasonable, if one views
the nuclear interior as a non-compartmentalized structure, where chromatin fibers float in largely
random arrangements in the nuclear sap. In contrast, we view the nucleus as a highly
compartmentalized structure. In our view DNA depleted biochemical nuclear matrix preparations do
not reflect a random aggregation of components from a nuclear sap, but comprise the partially intact
and functional content of the ICD space in the living cell nucleus (see previous discussions of a
dynamic in situ nuclear matrix in (Berezney 1984; Berezney et al. 1995). This view implicates a
specific organization of ”closed” and ”open” chromatin domains (or chromatin fiber aggregations)
which enables specific topological and geometrical relationships between the chromatin and the ICD
compartment (see below). The major functional purpose of this compartmentalization is to regulate the
access of genes to preformed functional protein complexes contained in the ICD space. Accordingly,
our view implicates the formation of higher order nuclear compartments comprising sets of actively
transcribed genes in ”open” chromatin domains, and inactive genes in ”closed” domains. Only genes in
”open” chromatin domains should be exposed to the ICD space and thus accessible for transcripton
and splicing complexes. Accordingly, we predict a reorganization of chromatin domains and their
topological relationships with the ICD compartment when gene expression patterns change. While
there is growing biochemical evidence for a gene activity-related remodelling process on the
nucleosomal level (Varga-Weisz et al. 1997; Wyrick et al. 1999), experimental proof for the functional
importance of a dynamic topological organization for gene regulation at the level of higher order
nuclear compartments is limited (Brown et al. 1997; Brown et al. 1999; Bridger et al. 2000). Thus, it
may seem prudent to concentrate research efforts on models of gene regulation at the level of
nucleosomes and individual chromatin loops. However, such a restriction could unduly postpone
studies of a topological level of gene regulation that depends on large scale nuclear architecture,
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studies which finally may turn out to be indispensable for the understanding of an orchestrated gene
regulation during cell cycle, cell differentiation and cellular senescence and the disturbances of this
regulation in malignant cells. Recent developments of DNA chips have allowed the study of
expression patterns simultaneously for thousands of genes. The results have further emphasized the
problem of an orchestrated gene regulation where the expression patters of hundreds or even thousands
of genes may be simultaneously affected. Neglect of the functional importance of over-all nuclear
geometry or nuclear topology for gene regulation may also be prohibitive for understanding the
circumstances which allow the adequate expression of transgenes.
The potential impact of nuclear architecture at large on gene regulation cannot be confirmed (or
rejected) without the development of appropriate experimental tools and their application by
interdisciplinary research teams. Presently available tools include multicolor fluorescence in situ
hybridization (FISH) combined with immunocytochemical detection of nuclear proteins for 3D studies
of fixed cell nuclei in cycling and terminally differentiated cells, in particular nuclei in tissue sections;
chromatin and protein labeling for 4D (three space coordinates plus time) studies of nuclei in cultured,
living cells; improved high resolution laser microscopy and bioinformatics for quantitative 3D and 4D
evaluations; and higher order chromatin and chromosome computer modeling.
Although it is safe to say that changes in chromatin structure are correlated with changes of gene
activity, we still lack experimental paradigms which show unequivocally whether changes in higher
order chromatin architecture are the cause (as we expect) or the consequence of an orchestrated gene
expression, for example by the formation of transcription and splicing complexes at sites of active
genes (Singer and Green 1997). As is often the case in the elucidation of complex problems, the
framework of a simple dichotomy - changes in nuclear and chromatin architecture are either the cause
or the consequence of changes in gene regulation - may turn out to be insufficient.
The chromosome territory view of nuclear architecture
State of evidence for distinct chromosome territories, their subchromosomal organization and
intranuclear distribution
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Compelling evidence for the existence of chromosome territories in both animal and plant nuclei was
established in the '70's and '80's (Zorn et al. 1976; Stack et al. 1977; Zorn et al. 1979; Cremer et al.
1984; Manuelidis 1985; Schardin et al. 1985; Pinkel et al. 1986; Cremer et al. 1988; Lichter et al.
1988; Pinkel et al. 1988; Heslop-Harrison and Bennett 1990). Fig. 2 shows a typical example of a
human cell nucleus with chromosome territories visualized by fluorescence in situ hybridization
(FISH) with chromosome specific DNA libraries, a procedure termed "chromosome painting". Present
evidence argues that each chromosome at least with regard to its major chromatin mass occupies its
own, separate territory with distinct arm and band domains (Cremer et al. 1996; Dietzel et al. 1998a;
Dietzel et al. 1998b; Visser and Aten 1999; Zink et al. 1999). Chromosome territories exhibit various
shapes with occasional finger like chromatin protrusions (Fig. 3). Such protrusions may expand into
the ICD space accessible between two neighbouring chromosome territories or into infoldings of an
adjacent chromosome territory. Accordingly, the topology between the surfaces of neighbouring
chromatin territories can be complex due to expansions and infoldings, but territories apparently
remain as mutually exclusive entities., i.e. there is no evidence that chromatin loops from two
differently colored (say green and red) chromosome territories could become intertwined to an extent
that resembles their full intermingling (yellow) at the light microscopic level (Cremer et al. 1996;
Visser and Aten 1999).
Several active and inactive genes visualized with specific DNA probes were preferentially
located in the periphery of painted chromosome territories in contrast to a noncoding sequence (Kurz
et al. 1996). Other studies regarding the spatial distribution of early replicating, GC-rich,
transcriptionally active chromatin as compared to mid to late replicating, AT-rich, transcriptionally
inactive chromatin, as well as the formation of nascent RNA have provided evidence that
transcriptionally active genes are also located in the chromosome territory interior (Visser et al. 1998;
Verschure et al. 1999; Tajbakhsh et al. 2000). One should note, however, that the interior location
within a chromosome territory is fully compatible with the location at the surface of a ∼1-Mbp or
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∼100-kbp chromatin domain (Fig. 1). Recently, it was reported that RNA is synthesized at the surface
of chromatin as defined by both fluorescence microscopy and EM morphological criteria (Cmarko et
al. 1999; Verschure et al. 1999). Dietzel et al. (1999) have noted that the topology of certain X-
chromosomal gene sites in the chromosome territory depends on their transcriptional activity. In
summary, these findings argue for a non-random distribution of genes in chromosome territories and
chromatin domains but generalized rules cannot be deduced at the moment.
From the viewpoint of the chromosome territory–ICD compartment model it makes sense if
active genes were located at the surface of chromatin domains. From there chromatin loops with one or
several active genes could expand into the ICD space, where (as we assume in this model) preformed
transcription and splicing complexes are located. Volpi et al. (2000) have recently studied the three-
dimensional large-scale chromatin organization of the major histocompatibility complex located on
human chromosome 6. Large chromatin loops containing several Mbp of DNA were observed
extending outwards from the surface of the chromosome 6 territory. The frequency with which a
genomic region was observed on an external chromatin loop appeared to be related to the number of
active genes in that region. Importantly, transcriptional up-regulation of genes in the major
histocompatibility complex led to an increase in the frequency with which these genes were found on
an external chromatin loop occasionally several microns away from the segmented surface of the
painted chromosome 6 territory. It is not yet clear whether such loops may penetrate neighbouring
chromosome territories, but as noted above we think it more likely that they expand into the ICD
space. These data further support an association between large-scale chromatin organization of specific
genomic regions and their transcriptional status.
The question of to what extent chromosomes or chromosomal subregions, such as centromeres
or telomeres, maintain specific positions and spatial arrangements during the cell cycle and cellular
differentiation has remained as a matter of speculation and controversy. Some authors have claimed a
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highly ordered positioning of chromosomes in prometaphase rosettes of human fibroblasts and HeLa
cells, including the separation of paternal and maternal chromosome sets (Nagele et al. 1995; Nagele et
al. 1998). Others have provided data clearly contrary to this claim (Allison and Nestor 1999). For
human interphase nuclei some authors have argued that chromosome arrangements do not deviate
significantly from a random model distribution (Lesko et al. 1995). Others noted the preferential
positioning of certain chromosome types (Popp et al. 1990; Nagele et al. 1999) and provided evidence
for a redistribution of chromatin during the cell cycle (Ferguson and Ward 1992; Weimer et al. 1992;
Vourc'h et al. 1993) and during cell differentiation (for reviews see (De Boni 1994; Manuelidis 1990).
Recently, Croft et al. (1999) reported the preferential localization of the (gene-poor) chromosome 18
territory in the nuclear periphery whereas the (gene-rich) chromosome 19 territory was mostly found in
the nuclear center. This positioning changed reproducibly when cells exit from G1 to G0 and vice
versa (Bridger et al. 2000). Our own unpublished data suggest a strong preference of small
chromosomes towards the center and of large chromosomes towards the periphery of nuclei from
different cell types and species (human and chicken; F. Habermann et al., in preparation). Systematic
studies using multiple color FISH with defined probe sets in various cell types from different species
are necessary in order to obtain a reliable answer to the question whether evolutionary conserved, cell
cycle and cell type dependent motifs of higher order chromatin arrangements exist.
Chromosome territories contain focal chromatin aggregates
Pulse labeling of cells during S-phase with halogenated thymidine analogs (BrdU, CldU, IdU)
revealed chromatin aggregations of ongoing DNA synthesis, termed replication sites or foci
(Nakamura et al. 1986; Nakayasu and Berezney 1989). These foci were found to have diameters
between some 300 and 800 nm with an estimated DNA content of about 1 Mbp (Jackson and Pombo
1998; Ma et al. 1999). Each replication focus consists of a cluster of active replicons together with
replication proteins and their auxiliary factors. This finding, however, is not unambiguously accepted,
since other groups performing short labeling assays with 3H-thymidine or BrdU were not able to see
distinct replication foci in EM studies (S. Fakan, personal communication).
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Recently, chromosome territory organization was studied after double-labeling of human cells
during early and mid to late S-phase with CldU and IdU (Visser et al. 1998; Zink et al. 1998b; Zink et
al. 1999). Several studies demonstrated the persistence of ~1-Mbp chromatin domains labeled during
S-phase at other stages of the cell cycle (Ferreira et al. 1997; Zink and Cremer 1998a; Zink et al.
1998b; Zink et al. 1999). Early and mid to late replicating chromatin domains form distinct higher
order nuclear compartments in early G1 and persist throughout interphase (Sadoni et al. 1999) and
references therein). Early replicating chromatin foci form R-bands in mitotic chromosomes, while mid-
to late replicating foci form G- and C-bands, respectively (Zink et al. 1999) and references therein). It
has been hypothesized that chromatin domains are capable of associating with or assemble different
sets of factors, e.g. for replication, transcription or repair, at different time points during the cell cycle
(Berezney and Wei 1998; Wei et al. 1998; Leonhardt et al. 2000).
When the double labeled cells were further grown for several cycles, segregation of replication
labeled and unlabeled chromatids was achieved during the second and subsequent mitotic events
resulting in nuclei with labeled chromosome territories side by side with unlabeled ones (Ferreira et al.
1997; Zink et al. 1998b; Zink et al. 1999). The overlap volume measured by confocal laser scanning
fluorescence microscopy (CLSM) between differently colored early and mid to late replicating foci in
segregated chromosome territories was small, typically a few percent only (Zink et al. 1999). In
agreement with the above mentioned views of the early cytologists, these data support the concept of
chromosome territories built up from ~1-Mbp chromatin domains.
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Dynamics of chromosome territory structure in living cell nuclei
It is difficult to deduce dynamic features of the large scale geometry and topology of nuclear
chromatin during the cell cycle and cell differentiation in fixed cell studies. To overcome this
limitation, methods that allow 4D (space + time) studies of large scale chromatin arrangements in
living cells were recently developed (Robinett et al. 1996; Shelby et al. 1996; Marshall et al. 1997;
Zink and Cremer 1998a; Manders et al. 1999). Zink et al. (1998b) performed microinjection of
fluorochrome labeled nucleotides into nuclei of human cells during S-phase. This approach resulted in
the live cell fluorescent labeling of nuclear chromatin. As noted above, further proliferation of labeled
cells for several cell cycles yielded the segregation of both labeled and unlabeled chromatids, making
possible for the first time the direct microsocopic observation of individual, fluorescent labeled
chromosome territories in nuclei of living cells. As in the fixed cell studies described above, a
composition of individual chromosome territories of chromatin foci with diameters between some 300
and 800 nm was observed. These foci were termed subchromosomal foci and are identical with the ~1-
Mbp domains observed in fixed cell nuclei. During observation periods of several hours we observed
movements of foci and territories typically on the order of a few hundred nanometers with occasional
maximum displacements up to several micrometers ((Bornfleth et al. 1999). 4D analyses suggested
that both Brownian-like movements, as well as occasional directed movements of chromatin foci occur
in living cell nuclei (Zink and Cremer 1998a; Bornfleth et al. 1999). Earlier studies indicated the
possibility of major chromatin movements during terminal differentiation of neuronal cells
(Manuelidis 1990; De Boni 1994). To allow for such movements, the nuclear matrix must have a
sufficiently dynamic structure. While a continuous matrix could provide sufficient elasticity to allow
smaller movements, major movements of chromatin territories may require - at least locally - a
reorganization or even dissolution of a continuous matrix. The formation of a continuous matrix fiber
network could help to stabilize a permanent 3D chromatin architecture in terminally differentiated
cells. However, as we will discuss below in more detail, our view of the in vivo nuclear matrix does
not necessarily implicate a continuous 3D-structure at any time. Instead, this view implicates that
chromosome territories and their tightly associated proteins govern the topology and overall geometry
of higher order chromatin architecture with other large macromolecular complexes necessarily located
17
in whatever space is left over. This space could be formed in the nucleus in a way that occasional
chromatin loops penetrate into this space. However, the mean chromatin density within chromatin
domains, though variable, should never drop below a certain minimum, while the mean chromatin
density within the interchromatin space over scales on the order of >50 nm should never get as big as
that minimum (Fig. 1, inset). This line of reasoning argues for a qualitative difference between the
chromatin and interchromatin compartments, based on quantitative differences in chromatin density.
Whether the data actually bear out such a dualistic behavior of chromatin density, rather than just a
continuum of values, is not clear, but the apparently chromatin "free" lacunes seen between chromatin
do suggest the possibility. An interchromatin space of this kind would not necessarily require a specif
organization of the surrounding chromatin. In other words, DNA sequences would be randomly
exposed towards the interchromatin space. In contrast to this view we hypothesize that chromatin
domains are specifically organized in such a way that transcriptionally active genes or at least their
promoter regions and actively transcribed parts at each given time point are exposed at chromatin
domain surfaces in direct contact with the interchromatin domain (ICD) compartment.
The interchromatin domain (ICD) compartment view of nuclear architecture
A space with a very variable width up to a few microns extending between aggregates of
chromatin has been noted in many electron microscopic studies of the nuclear architecture. This space
could also be demonstrated in confocal sections from living cell nuclei, in which green fluorescent
protein (GFP) tagged histone 2B allowed the visualization of the entire nuclear chromatin (Kanda et al.
1998). Fixation of nuclei with buffered formaldehyde leaves the distribution of the chromatin in the
living cell nucleus largely intact (Fig. 4A-C). The working hypothesis was established that the content
of this space together with the demarcating chromatin surfaces and possibly decondensed chromatin
looping out into this space constitutes a functionally important nuclear compartment, termed the ”inter
chromatin domain” (ICD) compartment. We assume that protein complexes for transcription, splicing,
DNA replication and repair are localized in this compartment (Cremer et al. 1993; Zirbel et al. 1993;
Cremer et al. 1995; Cremer et al. 1996). For example, Fig. 4D-F reveals that variously shaped nuclear
bodies that contain a variety of splicing factors and are known as speckles or interchromatin granules,
18
are located within the ICD space. Speckles are dynamic structures that can produce expansions to sites
of active genes (Misteli et al. 1997). Further, we assume that chromatin domain surfaces establish a
border zone between the ICD and the chromatin compartment of the cell nucleus which can only be
penetrated by factors or protein aggregates below a certain size. Note that this definition is relative,
e.g. a single protein may penetrate into the interior of a ”closed” chromatin domain, while a large,
preformed transcription factor complex cannot. In such a scenario, genes located inside of ”closed”
chromatin domains should become inaccessible to transcription protein aggregates. The opening of
chromatin domains could bring the regulatory and coding sequences of a given gene in direct contact
to the ICD-space and thus allow the specific binding, for example of large transcription complexes
formed in this space. This scenario is compatible with experimental evidence for RNA being
synthesized at chromatin surfaces (Cmarko et al. 1999; Verschure et al. 1999). RNA polymerase may
migrate along a chromatin loop with active genes sufficiently expanded within the ICD space (Volpi et
al. 2000). Alternatively, an immobilized transcription factor complex could spool the coding DNA
through the complex. Similar considerations have been proposed with regard to replication complexes
(Cook 1999). In a recent EM study it was observed that DNA replication, as well as transcription,
takes place at the surface of chromatin domains (S. Fakan, personal communication). The extent to which nuclear diffusion of macromolecules and macromolecule complexes is size
limited is not yet clear. Evidence based on experiments with fluorescein isothiocyanate (FITC) labeled
dextrans (Lukacs et al. 2000) indicates that such macromolecules with a size up to 580 kDa are fully
mobile in HeLa cell nuclei. In contrast, comparably sized DNA fragments of 250 bp and greater were
nearly immobile suggesting extensive binding to immobile obstacles. Pederson and coworkers have
developed an elegant method to visualize movements of endogenous polymerase II transcripts in the
nuclei of living cells(Politz et al. 1998; Politz et al. 1999; Politz and Pederson 2000). The authors
conclude that intranuclear trafficking of RNA occurs by diffusion throughout the ICD space of the
nucleus. Available evidence supports the possibility that diffusion coefficents are considerably lower
in certain chromatin compartments than in the ICD space leading to a differential accessibility of
macromolecules or macromolecule complexes.
19
Based on the hypothesis that the ICD compartment exists in vivo and possesses all the major
functional properties of biochemically DNA depleted nuclear matrix preparations, we propose that its
topology and content provides the in vivo equivalent of nuclear matrix preparations (Tan et al. 2000).
While the ICD compartment model is compatible with a continuous network of nuclear matrix core
filaments, such an assumption is not a necessary condition of this model (for a possible explanation of
the formation of a network in nuclear matrix preparations see below). If the concept of the ICD
compartment as the equivalent of a dynamic in vivo nuclear matrix holds true, the topology and
configuration of components in nuclear matrix preparations should reflect their topology and
configuration in the ICD compartment of the living cell to the extent that such features can be
preserved in nuclear matrix preparations. A test of this hypothesis requires bringing together light and
electron microscopical evidence of chromosome territories, chromatin domains (or higher order
chromatin fibers) and the in situ nuclear matrix.
Considerations on the role of the nuclear matrix in chromosome territory organization
Berezney and coworkers (Ma et al. 1999) have argued that a specific subset of proteins, termed
chromosome territory anchor proteins (CTAPs), is required to maintain the structural integrity of
chromosome territories. The authors noted that chromosome territories remained intact after high salt
extractions (2.0 M NaCl or 0.65 M (NH4)2SO4), although these high salt extractions removed the bulk
of histone proteins and other soluble nuclear proteins. RNAse A treatment of the cells alone also did
not visibly affect the structural integrity of chromosome territories. A disruption of chromosome
territories was, however, noted after RNAse plus 2.0 M NaCl treatment.
In the light of the multiloop subcompartment (MLS) model (see below) the structural integrity
of chromosome territories depends on anchor proteins in the interior of each ~1-Mbp chromatin
domain necessary to keep together some ten ~100-kbp chromatin loop domains. Notably, these anchor
proteins do not form a continuous backbone (see below). We predict that the CTAPs described by (Ma
20
et al. 1999) are the equivalent of the anchoring proteins predicted by the MLS model. Since CTAPs are
not affected by RNAse treatment, it is clear that RNAse treatment alone should not disturb the
integrity of chromosome territories. However, one should expect that high salt extraction should be
able to remove the CTAPs and thus be sufficient to destroy the chromosome territory structure. Why is
this not the case? Why is the combined effect of high salt extraction plus RNAse necessary for
chromosome territory disruption? We consider two explanations for this surprising result. Both
explanations argue that RNAse attacks the integrity of RNPs contained in the ICD space. The first
explanation takes into account the possibility that in addition to CTAPs, a continuous RNP containing
matrix in the ICD space is of importance and in fact sufficient to maintain chromosome territories in
vivo even in the absence of CTAPs. This continuous matrix should resist high salt treatment and
become only destroyed after RNAse treatment. The second explanation argues that the ICD space in
the living cell nucleus contains protein and RNP aggregates only in a discontinuous, soluble form.
High salt treatment of cell nuclei results in the formation of an insoluble in situ matrix. Whether or not
one is willing to accept present evidence for a network of branched core filaments in the nucleus of the
living cell, it seems likely that high salt treatment leads to the aggregation of additional proteins and
RNPs onto this network. What matters in this context is the assumption that the in situ nuclear matrix
present in high salt extracted nuclei holds together the chromosome territory structure even after the
removal of CTAPs.
We wish to reemphasize here that we consider the in vivo nuclear matrix as a highly dynamic
structure (for relevant discussions of a dynamic view of the in situ nuclear matrix see (Berezney 1984;
Berezney et al. 1995). The longstanding controvery of whether or not such a matrix is based on a
permanent 3D network of branched core filaments in spite of its importance has distracted from such a
dynamic view. This dynamic view implicates the possibility (but not necessity) that a continuous 3D
network may be fully or partially established at some time, become dissolved at another time (e.g. to
enable chromosome territory movements or the reordering of early and mid to late replicating
chromatin domains), and again become reestablished later (e.g. to help to preserve a distinct 3D
chromatin architecture of a terminally differentiated cell type). Most relevant in the present context is
the assumption that the network contains RNA which can be cleaved by RNase A digestion. RNase A
21
digestion then would either destroy the continuous in vivo nuclear matrix network (if it exists) or
prevent the aggregation of RNPs to form a continuous network during high salt treatment. In each case
the nuclear matrix network surrounds chromatin domains and holds these structures together after the
extraction of CTAPs. In summary, we hypothesize that the effects of RNase A and high salt treatments
are directed to distinct, geometrically separated structures. The RNPs (and other proteins) possibly
form a 3D nuclear matrix network in the ICD space surrounding the chromatin domains, while the
CTAPs possibly provide an anchoring structure in the interior of each chromatin domain (Cremer et al.
1995). Both these distinct structures need to be dissolved in order to destroy the territorial organization
of chromosomes.
Computer simulation and virtual microscopy of chromosome territory organization and of the
ICD compartment
Several quantitative models of nuclear genome structure have recently been developed, which explain
the observed compartmentalization of nuclear chromatin with and without the assumption of a
continuous nuclear or chromosomal matrix/ chromosome scaffold. These models show that
chromosome territories with defined substructures can be obtained starting with a few basic
assumptions concerning the ultrastructural chromatin folding motifs. Computer modeling allows one to
translate chromatin folding motifs into "virtual" microscopical images. Virtual microscopical sections
and 3D images of various models of chromosome territory structure were generated taking into
account experimentally measured confocal point spread functions to simulate a given optical
resolution. Virtual microscopy allows a direct comparison with experimental images and 3D
reconstructions of chromosome territories (Münkel et al. 1999; Kreth et al. 2000 (in Press)). Virtual
microscopy also helps the experimentalist to realize the limitations of certain light microscopic
procedures. Neglect of these limitations can result in misinterpretations and misconceptions.
22
Random walk/giant loop (RW/GL) model
The RW/GL model proposed by Sachs et al. (1995) suggests giant chromatin loops with a size
of about 3 Mbp. Each loop is created by a random walk of chromatin fibers and held together at its
basis by the attachment to a continuous backbone structure (Sachs et al. 1995; Yokota et al. 1995). A
continuous backbone structure is a possibility that cannot be excluded by present experimental data.
However, it is not an a priori necessity to explain the formation of chromosome territories, since they
can be modeled without such an assumption. It is sufficient to assume protein ”clamps” possibly
equivalent to CTAPs (Ma et al. 1999), see above) that hold together a chromatin loop at its base
(Münkel and Langowski 1998).
Multi Loop Subcompartment (MLS) model
The MLS model (Münkel and Langowski 1998; Münkel et al. 1999) was based on the well
established finding of ca. 30 – 150 kbp sized chromatin loops. This model assumes that chromatin foci
in the order of 1 Mbp are formed by rosettes of several chromatin loops (Okada and Comings 1979)
each containing ~100 kbp of DNA. Rosettes are connected by chromatin linkers comprising only a
small part of the total nuclear chromatin (Fig. 5). For the simulation of single chromosome territories a
potential energy barrier was assumed, which in the absence of neighbouring chromosome territories
provided a hindrance for individual chromatin segments to leave the territory volume. The simulation
of diploid human cell nuclei with 46 chromosome territories was successfully performed without the
use of such a barrier for each individual chromosome territory. The nuclear envelope was considered
the equivalent of a rigid spherical shell modeled by a hard sphere potential.
Comparison of the RW/GL model and MLS model with experimental findings
Calculation of the overlap volumes of early and mid to late replicating chromatin foci predicted
by the RW/GL model (using a special computer simulation of this model) revealed a high degree of
overlap, which was not consistent with the experimental data, demonstrating only a very minor overlap
of these foci (Zink et al. 1998b; Münkel et al. 1999; Zink et al. 1999). In contrast, in the MLS model
(Münkel et al. 1999) the size and mutual exclusivity of individual chromatin rosettes agreed well with
23
the size and mutual exclusivity of the light microscopically observed early and mid to late replicating
chromatin foci (Zink et al. 1998b; Zink et al. 1999).
Spherical ~1-Mbp Chromatin Domain (SCD) model and modeling of the interchromatin domain (ICD)
space For extended computer simulations of large scale chromatin organization in entire human
diploid nuclei the SCD model was developed (Kreth et al., 2000; Kreth et al., in preparation). This
model is based on the key parameters used in the MLS model (for a justification of these parameters
see (Münkel and Langowski 1998; Münkel et al. 1999). However, taking into account the limited
knowledge concerning the actual folding of the chromatin fiber at the ultrastructural level, it makes no
assumptions on the ultrastructural chromatin topology inside the ~1-Mbp chromatin domains.
Furthermore, this simplification results in a drastic reduction (several orders of magnitude) in
computer time and allows for the computing of human model nuclei with 46 chromosome territories
within one day using a personal computer. This has already allowed to calculate several hundred
model nuclei and to study the resulting arrangements of model chromosome territories as compared to
experimentally observed chromosome territory arrangements ( our unpublished data). Fig. 6 shows a
spherical human model nucleus with a 10 microns (µm) diameter and a volume of about 500 µm3
calculated according to the SCD model. Each color visualizes a distinct chromosome territory. For
each chromosome territory, we assumed a chain of spherical 1-Mbp chromatin domains with 500
nanometer (nm) diameter. Between different spheres, repulsive forces were assumed (see below for a
justification of this assumption). Modeling was done in such a way that it allowed for slight volume
overlaps between neigbouring 1-Mbp chromatin domains. This was achieved by an increasing
potential energy with a half width of ~250 nm. Chromatin linkers connecting adjacent chromatin
spheres were modeled by a spring potential in order to allow a thermodynamic equilibrium distance of
about 600 nm between the centers of connected chromatin domains at 37°C. In addition, a spherical
24
potential energy barrier was applied around each territory with a size corresponding to its DNA
content. This potential energy barrier is essential to maintain a certain compactness of the modeled
chromosome territories. It prohibits chromatin segments from leaving the territory volume and
accounts in a drastically simplified way for forces which in real nuclei may arise from a variety of
parameters, including the rigidity of higher order chromatin segments, chromosome territory anchor
proteins and possibly a nuclear matrix network formed in the ICD space (see above). Using the
parameters specified above, we achieved a good correlation of the SCD model with present
experimental data concerning the variability of chromosome territories in size and shape, as well as the
experimentally measured angle distribution between the centromer and the two telomers of X-
chromosome territories (Dietzel et al. 1998a); G. Kreth and C. Cremer, unpublished data). Clearly, the
examples of modeling chromosme territory structure and nuclear arrangements presented above are
only a beginning. Multifactorial modeling needs to be performed, in which a whole range of variable
quantities is assigned to the key parameters. Only then it will become possible to see which model
comes closest to the observed data. However, such an approach is rendered difficult not only because
of the extensive computer time required but also due to the present limitation of quantitative 3D and
4D experimental data on the higher order nuclear architecture. Improvements on both sides,
quantitative modeling and quantitative, experimental tests, are necessary to achieve a better
understanding of the compartmentalized nuclear architecture and its functional consequences.
A certain rigidity of higher order chromatin structures is essential for tensegrity models of
nuclear architecture (see below). The variable shape and rigidity of higher order chromatin structures
also contributes to the formation of an ICD compartment with a variable width of space. We have
started to use this approach to model the effects of geometrical constraints of differently sized human
chromosome territories on their intranuclear distribution and the frequencies of translocations between
pairs of individual chromosome territories (data not shown). The present version of the SCD model
25
neglects Brownian motion of chromatin domains and chromosome territories, respectively.
Observations of the ICD-space in confocal sections of nuclei from living HeLa cells with GFP-labeled
H2B histons (Fig. 4) suggest a relatively stable ICD space. Depending on the efficient viscosity of
large scale chromatin structures, it seems possible that interchromatin channels can open and close due
to Brownian motion (R. Sachs, personal communication, and our own preliminary results of computer
simulations of Brownian dynamics of the SCD model; Kreth et al., in preparation) Possible time
scales of such an opening or closure, however, are presently not known.
To create images of virtual ”low resolution” ultrastructural sections through SCD model nuclei,
the voxel data of 156 nm thick sections were calculated. As an example, Fig. 7A shows a virtual
section in one color only to reflect the fact that chromosome territories in EM sections cannot be
differently colored as in multiple color FISH experiments. Fig. 7B shows the same virtual section
taking into account the maximum resolution of a commercial confocal fluorescence microscope.
Notably, the borders of individual chromosome territories cannot be detected in these images,
emphasizing the importance of specific labeling approaches which can distinguish individual
chromosome territories. In contrast to the virtual EM section, individual 1-Mbp chromatin domains are
no longer distinguishable in the ”light microscopical” section (compare Fig. 4). Both virtual sections,
however, reveal an ICD space extending between chromosome territories and ~1-Mbp chromatin
domains with diameters up to the micrometer range, while the finest branches of the virtual ICD space
extending between modeled ~100-kbp chromatin domains have diameters in the nanometer range (Fig.
7C-E). The density of the ”chromatin” varies in various parts of the model sections. It is obvious that
the chromatin-free space ascribed to the ICD compartment space fills up a considerable part of the
”virtual microscopic section”. A minimum of 20% of the nuclear volume was calculated for the ICD
space in SCD model nuclei under the assumption that the ICD space is given by the entire spherical
nuclear volume minus the total volume of the 6350 ~1-Mbp chromatin domains (each with a diameter
of 500 nm) reflecting the total DNA content of a diploid mammalian cell nucleus in G1 (Morton 1991).
26
Note that the ICD space obtained in the SCD model is a consequence of a few simple assumptions of
chromatin domain organization including a certain rigidity of higher order chromatin structures.
According to the SCD model, the ICD space is formed as a consequence of chromosome territory
architecture and its interaction with other chromosome territories and does not require the assumption
of a primary, chromatin organizing role of a filamentous nuclear matrix network possibly formed
within this space. This view eliminates the problem of how a chromatin organizing nuclear matrix
consisting of a continuous 3D network of branched core filaments is replicated during S-phase together
with chromatin to organize the two sister chromatids properly. The SCD model, however, does not
exclude a secondary role of a filamentous nuclear network (if it exists in vivo) in the maintainance of a
higher order chromatin architecture (see also the considerations on a role of tensegrity forces described
below).
In spherically shaped model nuclei with a diameter of 10 µm and a DNA content of 6350 Mbp,
the volume calculated for all nucleosomes and the DNA chain with zero space in between makes up
only a few percent of the entire nuclear volume (500 µm3). In some tissues, however, nuclei with
much smaller diameters can be observed (4 – 5 µm) resulting in a total nuclear volume of only about
50 µm3 (Mayhew and Astle 1997). In these particularly small nuclei, the volume required for most
densely packed nucleosomes occupies almost half of the nuclear volume and much more for ordinary
packaged chromatin. Compared to large nuclei, small nuclei with full chromatin content necessitate a
higher compactness of chromatin and/or a much smaller ICD space. Small diploid nuclei possibly
carry a much larger fraction of inactive genes than large diploid nuclei and provide a particularly
interesting case to test the predicted topological and geometrical relationships between the ICD
compartment and active or inactive genes (Fig. 1, insert).
Modeling and virtual microscopy of the ultrastructure of ~1-Mbp and ~100-kbp chromatin domains
In SCD model nuclei sections through individual ~1-Mbp chromatin domains appear as small
”chromatin circles” resembling the chromatin globules or granules described by the early cytologists.
Attempts were undertaken to model the possible ultrastructure of 1-Mbp and 100-kbp chromatin
domains at the nucleosome level. Fig. 8 shows several three-dimensional computer models of the
27
interior spatial architecture of 1-Mbp chromatin domains exhibiting different degrees of condensation
of some or all 100-kbp domains. In accordance with the MLS model we assumed that each 1-Mbp
domain was built up by ten 100-kbp chromatin domains. For the purpose of modeling, we further
assumed that the nucleosome fiber of each domain was held together by central elastic, tensegrity
forces acting on the respective reference points of the 100-kbp domains (where the modelling process
starts; Kreth et al., in preparation). The term "tensegrity" was first introduced in architecture to
describe the property of skeleton structures developed by the architect R. Buckminster-Fuller.
Tensegrity employs continuous tension parts and discontinuous compression parts, i.e. parts of
different rigidity. These parts are interconnected in the whole structure in a way that transient
mechanical stresses applied at a given site result in either reversible, or irreversible geometrical
rearrangements of all other - even remote - parts, depending on the special conditions of the system.
Recently, a major role of tensegrity has been proposed for cellular architecture and function (Ingber
1997; Beliakova et al. 1999; Chen and Ingber 1999). Application of the tensegrity concept to nuclear
architecture requires sufficient rigidity of higher order chromatin structures to generate tensegrity
forces by the interaction of these chromatin segments with each other and possibly with a nuclear
matrix network. In addition to chromosome terrritory anchor proteins (CATPs), a contribution to
tensegrity forces might result directly from interactions between DNA segments, e.g. via the
interaction of polypurine/poly pyrimidine sequences (Wells et al. 1988; Vogt 1990; Sinden 1994).
Between neighbouring, condensed 100-kbp domains shown in Fig. 8A-D, spaces can be noted,
suggesting that the ICD-space with its finest branches expands between the surfaces of condensed 100-
kbp chromatin domains. Within these condensed domains the nucleosome density appears to be so
high that the diffusion of large macromolecular complexes (with diameters in the range of, say, 20-30
nm) into the interior of these structures should be prohibited. Here, the interaction between chromatin
and ICD-compartment could indeed be limited to the chromatin surface. The role of the 3D chromatin
nanostructure on the diffusion of macromolecule complexes needs to be simulated more exactly. A
decrease in the tensegrity force of a given 100-kbp chromatin domain is equivalent to a more ”open”
28
configuration (Fig. (8E,F), while an increase leads to a more compacted or ”closed” configuration.
Notably, due to geometrical constraints the opening of a single 100-kbp chromatin domain in a 1-Mbp
chromatin domain containing other 100-kbp domains in a ”closed” configuration leads to the
expansion of the chromatin from the opening 100-kbp domain at the periphery of the remaining
”closed” 100-kbp domains (Fig. 8D). Such an expansion could help to expose the chromatin of the
”open” domain to a branch of the ICD compartment and facilitate the access to protein aggregates
contained within this compartment.
In contrast to virtual electron microscopic images, virtual light optical confocal sections
through modeled chromatin domains do not reveal any details of an ”open” or a ”closed” configuration
(Fig. 7C-E). This trivial consequence of the difference in resolution is explicitly mentioned here to
reinforce the necessity to take into account the limits of light microscopic observations. Otherwise,
conclusions based on light optical approaches, concerning e.g. the ”randomness” of chromatin
arrangements or the ”diffuse” intranuclear migration of RNA, can be grossly misleading. Note that the
”practical” 3D resolution of a conventional laser scanning microscope is ca. 250 x 250 x 700 nm. New
approaches of far field fluorescence microscoopy have recently been developed which have the
potential to obtain 3D-images of entire cell nuclei at a resolution corresponding to a smaller voxel size.
These include certain types of point spread function (PSF) engineered microscopy (Hell and
Wichmann 1994; Schneider et al. 1998) or X-ray tomographic microscopy (Lehr 1997). In
combination with spectral precision distance microscopy (SPDM; (Bornfleth et al. 1998; Edelmann et
al. 1999), a further resolution of topological and geometrical details is expected. The SPDM approach
is based on the principle, well known in astronomy, that distances between given object sites much
smaller than the conventional optical resolution (as given by the point spread function) can be
measured if these object sites are labeled with different spectral signatures. Recently, confocal SPDM
has been applied to approach the internal topology of the BCR-ABL fusion gene in bone marrow cell
nuclei of leukemia patients at a 3D ”resolution equivalent” of about 50 nm, corresponding to
29
approximately one tenth of the ”conventional” optical 3D resolution. (Esa et al. 2000 (in press)). These
developments in optical engineering and image analysis, combined with multicolor labeling of specific
chromatin sites, should become highly applicable in the future to test predictions of computer models
of 1-Mbp and 100-kbp chromatin domains described below.
Physical forces of potential influence in maintaining large scale chromatin and interchromatin
domain organization
In the models described above, both attractive and repulsive forces were assumed to act
between chromatin domains. In the following we will argue that such forces are not just a convenient
assumption for computer modeling but are closely correlated to physical forces existing in the cell
nucleus. In our view, these forces are essential for the maintainance of distinct, mutually exclusive
chromosome territories with a considerable degree of dynamic flexibility (but also a considerable
amount of rigidity), the formation of an ICD space, and the opening/closing of chromatin domains
(Kreth et al., 2000; Kreth et al., in preparation)
Attractive forces
Tensegrity forces will be found in all networks between elements of different elasticity/rigidity (Ingber
1993). Note that in a tensegrity network, ”long range” forces (e.g. elastic forces between opposing
sites of the structure) are implied although the actual physical forces act on a short range basis between
closely neighboring elements only. The reason for this is the multiple connections given in a tensegrity
network. Thus, it is possible that ”short range” forces can combine to create ”long range” interactions.
Furthermore, simple physical tensegrity model systems show that a change in the ”macrostructure” of
the system (e.g. changing its overall shape) may produce subtle local changes at specific sites, i.e.
influence its ”microstructure” in a site specific manner depending on the special construction of the
system. In the case of chromatin, the existence of elastic forces and of stretches of different rigidity is
experimentally well established (Houchmandzadeh et al. 1997). To produce the network typical for
tensegrity forces, attractive forces have to exist not only between neighbouring chromatin domains
30
built up from a continuous stretch of DNA, but also between domains the DNA of which does not form
linearly adjacent sequences. Note that chromatin domains can be neighbors in the nuclear space
although they are genomically well separated, e.g. by many Mbp along a chromosome. Attractive
forces acting between chromatin domains (or rosettes in the MLS model) coming very close to each
other include van der Waals forces and hydrogen bonds between neighboring chromatin domains. In
addition to hydrogen bonds between proteins, the possibility of such bonds acting between three bases
(e.g. Hogsteen pairing), may be taken into account: polypurine/polypyrimidine sequences can give rise
to triple-stranded DNA stretches plus a single stranded sequence; the single stranded sequence may
form a triple-stranded stretch with a polypurine/polypyrimidine sequence of the DNA of a neighboring
chromatin domain (Vogt 1990), or a double stranded stretch with a single stranded sequence from
another polypurine/polypyrimidine site. If so, this would provide an additional mechanism for the
formation of highly complex, sequence dependent chromatin structures. A computer analysis of the
human genome data bank revealed a large number of highly repetitive as well as low repetitive/unique
sequence motifs which might serve for such a function (Winkler 1999). Another important source of
the site specific short range interactions characteristic for the formation of long range tensegrity forces
might be sequence specific non-histone proteins. In Drosophila melanogaster, nuclear factors encoded
by the Polycomb group (PcG) and trithorax group (trxG) genes were discovered. These factors act on
specific chromosomal elements and play an important role in chromatin remodeling processes, which
direct specific chromatin structures into either a ”closed” or ”open” conformation (Cavalli and Paro,
1998, and references therein). Other relevant protein candidates for control of tensegrity might be
chromatin accessibility complexes (CHRAC) which induce the sliding of nucleosomes on DNA
(Varga-Weisz et al. 1997). Tensegrity forces may be correlated to genetic activity (Ingber 1993;
Singhvi et al. 1994). Assuming different tensegrity forces, it was possible to perform computer
simulations (Kreth et al. 2000 (in Press)) of the different morphologies experimentally observed for the
active (Xa) and the inactive (Xi) X chromosome territories in human amniotic fluid cell nuclei (Eils et
al. 1996).
Repulsive forces resulting from Lenard-Jones and Donnan potentials
31
The other major forces assumed in modeling of nuclear genome structure are repulsive short range
forces acting between neighbouring chromatin domains. Such forces may originate from different
sources:
a) Lenard-Jones potentials (declining with the sixth power of the distance) are widely used in solid
state physics to describe short range repulsive forces. If the chromatin surfaces of two neighboring
chromatin domains come very close together (less than one nm), the electron shells of their atoms will
start to strongly repulse each other. These short range repulsive forces inhibit the penetration of
neighbouring ~1-Mbp and ~100-kbp domains when these domains are pushed together, e.g. by
Brownian chromatin movements..
b) Electrostatic interactions: Here, we want to consider especially a possible role of Donnan potentials.
These may be generated as a result of negatively charged phosphate groups of very low electric
mobility in the DNA, and counter ions with high electric mobility in the entire interchromatin space.
Donnan potentials were expected to be formed between negatively charged chromatin domain surfaces
and the ICD space (Cremer et al. 1993; Cremer et al. 1995; Cremer et al. 1996) and to result in
repulsive forces between opposite chromatin domain surfaces. Although the range of repulsive forces
created by Donnan potentials is much larger than the range of Lenard-Jones potentials, it is limited to a
few nm. Since the width of the ICD space may be in the range of several hundred nm and some sites in
the µm range (compare Fig. 4 with Fig.7 and 8), it appears at first glance unlikely that Donnan
potentials play any role in the maintenance of the ICD compartment. However, we have to consider
that the ICD compartment is a dynamic structure. In spite of its enormous width at some locations it
becomes very narrow at many other sites. The ICD space in ”closed” higher order chromatin
compartments should be largely or entirely devoid of macromolecular complexes for transcription and
splicing and therefore much more narrow than the ICD space in transcriptionally active higher order
chromatin compartments. As a result of statistical fluctuations of the width of the ICD-space, e.g. as a
result of Brownian chromatin movements, chromatin surfaces from neighboring chromatin domains
32
(belonging even to different chromosome territories) occasionally approach each other very closely
resulting in the local compression of the space between opposite chromatin domain surfaces. We argue
that Donnan potentials in the order of a few mV are sufficient (see calculations and measurements
described below) to maintain a minimum width of the ICD-space at these narrow sites. For the case of
a zero Donnan potential, ”long range” (i.e. in the order of a few nanometers) repulsive forces should
no longer exist. Consequently, opposite chromatin surfaces can be driven so close towards each other
by Brownian movements that attractive forces, such as van der Waals forces and hydrogen bonds, may
become predominant. This could lead to the (even irreversible) clumping together of chromatin
domains (belonging even to different chromosome territories) and result in permanent loss of fine
branches of the ICD space.
For several reasons we believe that the maintenance of a minimum width of the ICD space
(larger than the range of attractive forces between chromatin domains) is an essential feature of the
dynamic, functional nuclear architecture predicted by the chromosome territory–ICD compartment
model. Let us consider the predicted structural change of a higher order chromatin architecture from a
genetically inactive to a transcriptionally highly active state. In the case that short range attractive
forces lead to the attachment of chromatin domains as soon as they come close enough in an inactive
higher order chromatin architecture with an empty ICD space, it would become necessary to overcome
these attractive forces in order to regain a genetically active chromatin architecture. Clumping of
chromatin domains from different chromatids and chromosome territories could interfere with
chromosome and chromatid separation during mitosis and interfere with decondensation processes
during the telophase/G1 transition. These functionally adverse consequences could be avoided, if an
ICD space of a few nanometers could be maintained as a result of long range repulsive forces between
chromatin domain surfaces even in genetically inactive chromatin, safeguarding a permanent 3D
interconnectivity of the ICD space both in genetically active and inactive higher order nuclear
compartments. When inactive chromatin compartments become active, narrow ICD compartments
could expand as a result of transcription and splicing complexes which form de novo or enter as
preformed complexes from the surrounding ICD space segments, pushing opposing chromatin surfaces
of narrow ICD spaces apart.
33
In addition to their contribution to the dynamic ICD compartment structure, Donnan potentials
in the suggested order of a few mV could enforce the electrostatic (Coulombic) ”trapping” of
negatively charged macromolecules and (not too large) macromolecule complexes within the ICD
space. Electrostatic trapping may result in an enrichment of such macromolecules, e.g. RNPs, in the
ICD space and play a role in their channeled diffusion. If each individual component of functional
macromolecular complexes would only show a moderate enrichment within the ICD space, it would
follow from thermodynamic considerations that these complexes should form preferentially in the ICD
compartment. In contrast, the ICD compartment should be depleted to a certain extent of positively
charged proteins, such as the strongly basic histones.
Theoretical considerations on the size of Donnan potentials expected in nuclei of living cells
The following calculations are based on the Debye-Hückel and Donnan potential theory and were
performed under highly simplified assumptions (Fig. 9). They are provided here solely with the
intention to show that the order of magnitude of the required physical effects seems plausible enough
to justify further theoretical and experimental efforts. Repulsive forces between the opposing surfaces
of two chromatin domains A and B arise as a consequence of negative charges on both surfaces. This
assumption is based on the following rationale. Electrophysiological measurements showed that the
interior of the interphase nucleus is negatively charged (Kanno and Loewenstein 1963; Oberleitner et
al. 1993). DNA and chromatin under physiological conditions carries a small negative net charge. For
RNA, DNA, as well as for entire human mitotic chromosomes in suspension the electric mobility at pH
7 was observed to be in the same order of magnitude, i.e. approx. –1x10-8m2V-1s-1. This finding
suggests a negative electric net charge for entire chromosomes in the order of one to several thousand
negative elementary charges (Bier et al. 1989). For simplicity, in the following model calculations we
take into account negative electric charges of such an order and further assume that chromatin surfaces
that demarcate the ICD-space have everywhere the same negative electric mobility. In reality, of
course, the mobility depends on the viscosity and the viscosity is different for the actual cell nucleus
and for the preparations where the mobility was measured. In any case, chromatin domains should
repel each other. In case of an electrolyte in ICD space with monovalent ions of each sign and bulk
34
concentration c, the size of the repulsive force can be estimated as follows. According to the Debye-
Hückel theory the electric potential U at a distance x from the surface of chromatin domain A can be
described by
equation 1:
e0 = negative elementary charge; ε0 = electric field constant; εr = relative dielectric constant; kB =
Boltzmann constant; c= mobile counter ion concentration; T = absolute temperature of the ICD-space
(310 Kelvin); DL = Debye length; (all values in SI-units) this length gives the distance from the
surface of chromatin domain A where the potential is reduced to 1/e (37%) of ∆ϕ (see below). Fig. 9A
shows the behavior of DL in dependence on the mobile counterion concentration according to eq. (1).
For example, assuming two types of monovalent, mobile counter ions, e.g. Na+ and Cl-, with a
concentration of each 150 mM and a dielectric constant εr = 80 (water), K = 1.2x109m-1 follows. In
this case the Debye length is 0.8 nm. ∆ϕ is the maximum potential difference between the surface of
chromatin domain A (x=0) and the ICD fluid at a sufficiently large distance. Usually ∆ϕ is given as a
function of the surface charge density σ:
( ) KxexU −⋅∆= ϕ where
K2
1
0
201 2 ⎟⎟
⎠
⎞⎜⎜⎝
⎛== c
Tke
BrDL εε
35
equation 2:
The charge density σ represents the charge load per unit area in the absence of small mobile
counterions. Since reliable values for σ are not available, we estimated ∆ϕ using the Donnan theory in
its simplest form. This theory gives an estimate of the electric potential difference (Donnan potential)
between a phase of fixed ions and a fluid phase with mobile counterions of bulk concentration c of
each sign assuming a constant concentration [Y] of negatively charged, fixed phosphate groups within
the chromatin domain.
equation 3:
where F is the Faraday constant.
In Fig. 9C the expected Donnan potential ∆ϕ is plotted as a function of the mobile counterion
concentration for various fixed concentrations [Y]. In a scenario where all phosphate groups are
exposed we obtain a Donnan potential estimate of ~2.7mV for the case of a nuclear concentration of
monovalent counterions c ~ 150 mM (measurements by Century et al. (1970) yielded a value of c =
145 mM) and of fixed ion concentrations Y ≈ 30 mM. On the average, a large fraction of negatively
charged phosphate groups will be already compensated by histones, other nuclear proteins and
polyamines. However, this shielding effect might be compensated to an unknown extent by the
presence of large amounts of negatively charged nonhistone proteins bound to chromatin.
Kr 0εεσϕ =∆
⎥⎥
⎦
⎤
⎢⎢
⎣
⎡
⎟⎟
⎠
⎞
⎜⎜
⎝
⎛⎟⎠⎞
⎜⎝⎛++−=∆
22112
lnYc
cY
FRTϕ
36
In the following we consider the dependence of the expected repulsive force between two
adjacent chromatin domains A and B as a function of the mobile counter ion concentration. For
simplicity, it is assumed that chromatin domain A is fixed, while chromatin domain B can be moved.
The repulsive force Frep which is exerted by chromatin domain A on chromatin domain B in a
distance x can then be calculated (using equation (1) by
equation 4:
gradU: gradient of the electric potential U
Q: negative net charge on the surface of chromatin domain B; see above for explanation of other
symbols.
The maximum repulsive force F0 between the two territories expected for x = 0 is given by
equation 5:
Frep decreases exponentially with increasing distance x between the two chromatin domain surfaces.
For x = DL, i.e. the Debye length, Frep decreases to 1/e F0. Fig. 9B shows the distance dependence
Frep/F0 for different concentrations c of mobile counterions in the ICD space. The repulsive force
declines to 1/e F0 at 3.2nm for a low ion concentration (10mM). This range decreases to 0.8 nm for
150mM (expected order of mean monovalent ion concentration in the cell nucleus, see above). The
( ) Kxrep KeQQgradUxF −⋅∆=−= ϕ
KQF ϕ∆=0
37
maximum repulsive force F0 between two chromatin domains A and B can be estimated for a given
mobile ion concentration, if the net charge Q on the surface of chromatin domain B is known.
For a rough estimate of the distance x between a fixed chromatin domain A and an adjacent, movable
chromatin domain B produced by the repulsive electric forces Frep discussed above, we neglect
Brownian movements and assume that x = 0 for t = 0. (In reality, we assume that the process will be
the opposite: following mitosis, the territories approach each other until they are repelled. Due to the
repulsive force Frep, the chromatin domain surface of B moves away from A with a given velocity v(t)
= dx/dt. This velocity will be slowed down due to ”frictional” forces Ffric exerted mainly by other
surrounding territories. For a quantitative treatment, Stokes law
equation 6:
is assumed, where a is the radius of the chromatin domain to be moved and Ffric the effective nuclear
friction factor exerted on the chromatin domain, using a pseudo viscosity ηnucl). Neglect of
acceleration terms (Ffric = Frep) and integration using equ. (4) results in
equation 7:
Since Q = 6πηau (u electric mobility of a chromatin domain with net charge Q in a medium with
viscosity η),
( ) ( )tatF nuclfric v6πη=
( ) ⎟⎟⎠
⎞⎜⎜⎝
⎛+⋅
∆= 1
6ln1
.
2
ta
KQK
txnuclηπ
ϕ
38
equation 8:
follows. Note that in this equation the radius a is eliminated. Thus, under the assumptions made the
width of the ICD-space is independent of the size of the chromatin domain. For an estimate of the
apparent pseudo viscosity ηnucl”, calculations based on optical tweezer experiments suggested ηnucl
to be in the order of 50 kgm-1s-1 (Cremer et al. 1993). Recently, the movement of individual
chromosome territories was quantitatively measured in undisturbed, living human cell nuclei (Zink et
al. 1998). From these data, ηnucl was estimated to be in the order of 2.5-20 kgm-1s-1. Note that ηnucl is
a global parameter describing the overall movement of large chromatin structures within the nucleus.
Thus, it is expected to be strongly dependent on the type of structures considered. In Fig. 9d, a
numerical example for equ. (8) is shown, assuming as constant values ∆ϕ = -2.7 mV, K = 1.2 x 109m-
1, u = 1 x 10-8m2V-1sec-1, η = 0.7 x 10-3kgm-1sec-1, and two values for ηnucl, 50 kgm-1sec-1 and
2.5 kgm-1sec-1. After t = 100 sec the average ICD-compartment width is around 8 to 10 nm and
increases to 13 to 15 nm after three hours. Due to the logarithmic dependence (equation 8), the width
changes very little, when u∆ϕK2η/ηnucl changes by one order magnitude.
Measurements of Donnan potentials
In the previous section we have argued that the existence of a Donnan potential in the range of a few
mV could account for repulsive electric forces between chromatin domain surfaces sufficient to
maintain a minimum width of the ICD space in the range of a few nanometers. The existence of an
electrical potential difference between the nuclear interior and the cytoplasm has been shown by a
variety of reports (Bustamante 1994; Kanno and Loewenstein 1963; Oberleitner et al. 1993). It was not
known, however, whether these differences were due to a Donnan potential. Only a few measurements
could be made so far with nuclei from living T-cell-tumor-lymphocytes (JURKAT). These
( ) ⎟⎟⎠
⎞⎜⎜⎝
⎛+⋅
∆= 1ln1
.
2
tKuK
txnuclη
ηϕ
39
measurements yielded a mean potential difference of -15 mV. For technical reasons, however, we did
not succeed to measure Donnan potential curves in nuclei of living cells. We therefore measured the
dependence of the nuclear voltage on the charge concentration of the bath solution surrounding fixed
cell nuclei which were prepared from osmotically extracted lymphocytes, human tumor T-Cell
lymphocytes (JURKAT), cultured diploid human lymphocytes, and HeLa cells. Cells were fixed with
either 4% buffered formaldehyde or a 3 : 1 (v,v) mixture of methanol and acetic acid. For voltage
recording, a cell nucleus was held with a patch pipette and entered with a KCl-filled microelectrode.
The nuclear voltage was monitored as a change of the potential and was negative in amplitude. The
penetration of the cell nuclei was frequently successful as could be seen as a step in the monitored
voltage (Fig. 10). The recordings were mechanically very sensitive as even very little vibrations
produced a short circuit which could be seen as a drop in the voltage. The charge concentration was
varied from about 10 mM to 1300 mM, covering the physiological range at about 124 mM up to very
high charge concentrations. Recorded voltages were corrected for liquid junction potentials. At
physiological conditions (charge concentration 124 mM) the average nuclear voltage was -17.4 mV +/-
1.6 mV for the formaldehyde fixed JURKAT cell nuclei and -16.8 mV+/-1.3 mV for formaldehyde
fixed human lymphocyte cell nuclei. The nuclear voltages followed a curve predicted for a Donnan
potential (Fig. 11). The fit with a Donnan curve yielded a concentration of fixed negative charges of
177 mM. For methanol/ acetic acid fixed JURKAT cell nuclei the concentration of fixed negative
charges for the fitted Donnan curve was 76 mM. The nuclear voltage at a charge concentration similar
to intracellular conditions (124 mM) was -5.2 +/- 0.4 mV for the JURKAT cell nuclei. Methanol/acetic
acid fixed diploid lymphocytes were not applicable for comparison. Cell nuclei fixed with
methanol/acetic acid were smaller in size than paraformaldehyde fixed nuclei. This effect was
particularly notable for lymphocyte nuclei. Since it was not feasible to pick these cells and penetrate
them with the microelectrode, we used methanol/ acetic acid fixed HeLa cells instead. The nuclear
voltage measured at 124 mM was -5.2 mV+/-0.2 mV for HeLa cell nuclei. This value is about the
magnitude reported by other authors (Oberleitner et al. 1993). Our results indicate that the measured
nuclear voltage is similar for different human cell types but is strongly dependent on the fixation
method. The methanol/acetic acid fixed cell nuclei showed a steady increase of the concentration of
40
fixed charges as calculated from nuclear voltages assuming a Donnan distribution with the mobile
charge concentration of the external solution. This change of the concentration of fixed negative
charges can be caused by several effects. Either volume changes or changes in the composition of the
cell nucleus like loss of histones at high salt concentrations may increase the net charge of the nuclei.
Further, a short circuit will affect higher voltage differences more than smaller ones.
The measured relation between the nuclear voltage and the charge concentration of the bath
solution clearly followed that of a Donnan potential. This result rules out the possibility that changes in
the tip potential of the microelectrodes or liquid-junction potentials are responsible for the results of
the electrical measurements, since those artefacts show a completely different dependence on the
charge concentration. We have not yet examined the role of the nuclear membrane in maintaining a
diffusional resting potential; the nuclear membrane, however, is an unlikely source for a Donnan
potential. This is in accordance with results published by others (Oberleitner et al. 1993). The
experiments showed the same nuclear voltage despite chemical (methanol/acetic acid fixation) or
mechanical (several penetrations with the micropipette) destruction of the nuclear membrane. The
results of these preliminary electrophysiological measurements of fixed cell nuclei support the
existence of a Donnan potential in the predicted order of several millivolts, but it remains to be seen
whether this order can be confirmed for nuclei of living cells as well. In summary, a decisive role of
repulsive forces due to charge in the maintainance of a minimum width of the ICD-space, when
neighbouring chromatin domains are driven towards each other by Brownian forces, has not been
proven so far, but requires further investigations.
Consequences of Donnan potentials for nuclear matrix formation
At high salt concentrations (used in standard nuclear matrix preparations) the Donnan potential
declined to almost zero values. As a consequence the repulsive forces between negatively charged
(ribonucleo-) proteins of the ICD compartment may be reduced to such an extent that they aggregate.
Such aggregations may or may not contain a continuous in vivo fiber network of matrix core filaments.
Finally, it should be reemphasized that the lack of such an in vivo network does NOT mean that
41
nuclear matrix preparations are a meaningless "artefact" in the sense of a random aggregation of
factors distributed in the nuclear sap. On the contrary, we propose that biochemical nuclear matrix
preparations represent the content and the functional properties of a specific, topologically and
functionally defined subcompartment of the living cell nucleus, called the ICD compartment or in vivo
nuclear matrix.
Acknowledgements
This work was supported by the Bundesminister für Bildung und Forschung (BMBF; German Human
Genome Project) and the Deutsche Forschungsgemeinschaft. T.C. and C.C. are indebted to Friedrich
Vogel for his strong encouragement of our chromosome territory studies over many years. The ICD
model was originally developed in 1993 together with P. Lichter. We thank him and the colleagues in
his and our groups for many discussions, which helped to shape the views presented here. Stimulating
discussions on quantitative higher order chromatin modeling with J. Langowski, Ch. Münkel and T.
Knoch are also gratefully acknowledged. We thank R. Berezney, P. Becker, R. van Driel, S. Fakan,
W. Hörz, T. Pederson, R. Sachs, A.P. Wolffe and L. Zech for helpful comments on earlier stages of
this manuscript. In particular, we greatly appreciate discussions with Ronald Berezney concerning his
views of a dynamic structure of the in situ nuclear matrix. Finally, the authors thank P. Edelmann and
B. Schädler for support in image analysis and L. Hildenbrand for literature research.
Fig. 1
Chromosome territory - interchromatin domain (ICD) compartment model of nuclear architecture.
Each chromosome in a cell nucleus occupies a distinct territory with a variable shape and complexly
folded surface. We propose that chromosome territories (CT) are built up from ~1-Mbp chromatin
domains connected by chromatin linkers. These ~1-Mbp chromatin domains in turn are composed of
linker connected ~100-kbp chromatin (loop) domains (Münkel and Langowski 1998; Münkel et al.
1999). An inter-chromatin domain (ICD) space starts at the nuclear pores and extends between the
mutually exclusive chromosome territories where it can form large lacunes with a width up to several
micrometers. Finer branches extend into infoldings of territory surfaces and between ~1-Mbp
42
chromatin domains located in the chromosome territory interior, the finest branches possibly end
between ~100-kbp chromatin domains. The ICD-space together with its demarcating chromatin
domain surfaces (and chromatin loops not shown here that may expand into this space) forms the ICD-
compartment of the cell nucleus. It contains macromolecular complexes for transcription, splicing,
DNA-replication and repair. For a rough representation of size, higher order chromatin structures and
the ICD-space were arbitrarily drawn onto the confocal section through a HeLa cell nucleus with GFP-
labeled histone 2B chromatin structures (white background, compare Fig. 4). The inset shows a purely
hypothetical example of topological and geometrical relationships between the ICD-compartment and
active and inactive genes of ~100-kbp chromatin domains. This drastically simplified scenario assumes
that factor complexes act at chromatin domain surfaces and consequently that active genes (white dots)
are exposed at chromatin domain surfaces and thus have direct access to preformed factor complexes
contained in the ICD-compartment, while inactive genes (black dots) are located in the chromatin
domain interior and not accessible by these factor complexes. This scenario, however, does not take
into account decondensed ("open") and condensed ("closed") chromatin domain configurations. For
more detailed 3D models of "open" and "closed" ~100-kbp chromatin domains compare Fig. 8. We
wish to emphasize that small changes in the location of genes (less than 100 nm) can possibly make a
decisive difference with regard to the availability of transcription and splicing complexes located in the
ICD-compartment. While positional changes of this magnitude cannot be detected by conventional
light microscopic approaches, new developments of spectral precision distance microscopy and other
types of high resolution laser microscopy provide the means to test this model (see text).
Fig. 2
Visualization and 3D-reconstruction of chromosome territories X and 8 in a human female lymphocyte
nucleus
A. Light optical serial sections through the nucleus were obtained with a laser confocal scanning
microscope. From a total of 48 (250 nm) equidistant sections every fourth section is shown. FISH was
performed with specific paint probes for chromosomes X (detected with Cy3, shown in green) and 8
(detected with Cy5, shown in red). The nucleus was counterstained with YOYO (shown in blue).
43
B. 3D reconstruction of the chromosome territories by surface rendering (Amira TM, TGS Inc.) is
shown in combination with one median optical section of the counterstained nucleus.
Fig. 3
Optical sections at high magnification through a painted human chromosome 2 territory (A) and
chromosome 18 territory (B).
Light optical sections of painted chromosome territories recorded from two diploid human fibroblast
nuclei. Both chromosome territories were visualised by FISH with the respective chromosome paint
probes. Note the irregular shape and focal substructure of the chromosome territories showing regions
with different chromatin density. Arrowheads in (B) point to finger like chromatin protrusions
extending from a chromosome territory core. ICD-channels expanding into the territory interior
(examples (indicated by arrows in (A)) occasionally open up to small ICD-lacunes (marked by asterix).
This interpretation is presented with the caveat that structures designated as ICD-channels and lacunes
do not represent areas free of chromatin but rather of non-painted chromatin due to a limited DNA
complexity of chromosome paint probes and/ or the fact that the visualization of repetitive sequences
was suppressed by an excess of unlabeled Cot1 DNA. In favor of this interpretation one should note
that light optical serial sections through nuclei where the entire chromatin is labeled in vivo by GFP-
tagged histone 2B consistently demonstrate the presence of ICD-channels and lacunes (compare
Fig. 4).
Fig. 4
Chromatin and interchromatin domain (ICD) compartment in HeLa cell nuclei
with GFP-tagged histone 2B
A. Laser confocal section through the nucleus of a living cell exhibits chromatin of variable density
visualized by GFP-tagged histone 2B (Kanda et al. 1998). Three nucleoli (marked by asterix) and
the ICD-compartment (black) contrast sharply to the labeled chromatin.
44
B. Laser confocal section corresponding to A obtained after 3D fixation of the nucleus with freshly
prepared, PBS-buffered 4% paraformaldehyde (10 min) and permeabilization with 0.5% Triton X
100 (5 min). GFP-tagged chromatin visualized in red.
C. The overlay of nuclear sections shown in A and B indicates that the topology/geometry of the
chromatin and the ICD-compartment seen in the living cell nucleus is maintained to a large extent
in the fixed cell nucleus, although shifts of chromatin domains up to several hundred nm can be
noted. For example, arrowheads indicate a rim of condensed chromatin around a large nucleolus,
which has slightly widened during fixation.
D-E. Laser confocal sections through a fixed HeLa cell nucleus.
D. Section showing GFP-tagged chromatin, two nucleoli (marked by asterix) and the ICD-
compartment (black).
E. Speckles visualized immunocytochemically in the same section with with SC35 antibodies (Cy3;
red).
F. Overlay of the sections (D) and (E) demonstrates that all speckles are positioned in expanded
regions (lacunes) of the ICD-compartment. Notably, the ICD-compartment is only partially filled
by the speckles leaving space for other macromolecular complexes.
Fig. 5
Scheme of the multiloop subcompartment (MLS) model
(adapted from: Münkel and Langowski (1998), with permission).
Three ~100 kbp chromatin loop domains (with a contour length of approximately 1.2 µm) are
exemplified. Each domain is modeled as random walk chromatin fiber held together by a loop base
spring formed by a harmonic potential. About ten chromatin loop domains form a higher order
subcompartment representing a ~1Mbp chromatin domain. These domains are linked by small
chromatin fibers. Loop base springs in the MLS model of chromosome territory organization may
reflect the function of chromosome territory anchor proteins (CTAP: Ma et al. 1999).
Fig. 6.
45
Spherical 1 Mbp chromatin domain (SCD) model of a diploid human cell nucleus with 46 chromosome
territories.
All 46 chromosome territories of a simulated diploid, spherical human male cell model nucleus
(46,XY) are visualized with 24 different colors (using Persistence of Vision (TM) Ray-Tracer Pov-
Ray(TM))). The model nucleus is shown as a relaxed configuration after 150000 Monte Carlo steps
(for further details see Kreth et al., in preparation) with statistically independently distributed
chromosome territories.
Fig. 7
Virtual sections through a diploid human cell model nucleus with 46 chromosome territories simulated
according to the SCD model
A. Simulated ”low resolution” ultrastructural section through a digitized three dimensional SCD model
of a diploid human cell nucleus (compare Fig. 6). This virtual section was obtained by an extended
view projection of two median sections (thickness 156 nm) of the digitized data stack with 78 nm axial
and 39 nm lateral voxel sizes. The encirclement indicates two neighboring 1-Mbp domains shown at
higher magnification in C-E.
B. The simulated median section shown in A. was filtered with a 250nm gaussian kernel. The result
corresponds to a light optical (laser confocal) section. Note the loss of information compared to the
virtual ”low resolution” ultrastructural image. Note that in both (A) and (B) an interchromatin domain
space (black) is clearly recognizable. Borders between virtual chromosome territories, however, cannot
be distinguished in the absence of specific coloring of chromatin domains belonging to different
territories (compare Fig. 4 and 6).
C-E. Virtual sections through the two encircled spherical 1-Mbp chromatin domains from 7A
correspond to ”high resolution electron microscopic” (EM) images. The finest dots represent
individual nucleosomes.
C. The two 1-Mbp chromatin domains were simulated under the assumption that each of the ten 100-
kbp domains contributing to their formation was highly compacted and formed a mutually distinct
46
compartment. In the virtual section four out of ten highly compacted 100-kbp domains contributing to
each 1-Mbp chromatin domain are seen (compare Fig. 8B).
D-E. The nucleosome chain configuration within each of the ten 100-kbp chromatin domains was
modeled according to Fig. 8E resulting in a moderate compartmentalization (D), or to Fig. 8F resulting
in a weak compaction (E) and pronounced loss of compartmentalization.
Below the virtual high resolution EM images (C-E), virtual ” light microscopical” (LM) images of the
same model structures are shown. The assumed LM resolution of 250 nm was simulated by filtering
the digitized model sections with a 250 nm gaussian kernel. Note that in the virtual LM images no
internal structure can be deciphered.
Fig. 8
High resolution simulation of the internal ultrastructure of a 1-Mbp chromatin domain.
Computer simulations of 1-Mbp domains were performed under the assumption that each domain is
built up from ten 100-kbp domains (indicated by 10 pseudo colors). The six simulations (A – F) were
performed according to different assumptions on the organization of the nucleosome chain within the
1-Mbp chromatin domain. All Visualizations were done using Persistence of Vision (TM) Ray-Tracer
Pov-Ray(TM).
A. 1-Mbp chromatin domain model with its nucleosome chain compacted into a 30nm chromatin fiber
(visualized by cylinder segments). The 30 nm fiber is folded into ten 100-kbp sized loop domains
according to the MLS model (compare Fig. 5) and occasionally interrupted by short regions of a
”beads-on-a-string” chain of individual nucleosomes (10 nm; small white dots, one indicated by an
asterix) (compare Alberts et al. (1994). A transcription factor complex bound to an exposed
chromatin site is shown as a brown sphere with a diameter of 30 nm (arrow).
B. Each of the 100-kbp chromatin domains was modeled under the assumption of a zig-zag (random
walk) nucleosome chain with volume exclusion, held together by ”tensegrity” forces directed to the
barycenters of the 100-kbp domains. These tensegrity forces result in ”closed” 100-kbp domains
(compare Fig. 7C). In these ”closed” 100-kbp chromatin domains, the accessibility to preformed
factor complexes built up in the ICD-space is limited to the surface of each domain, while most of
47
the chromatin becomes inaccessible in the chromatin domain interior. Each dot represents an
individual nucleosomes.
C. ”Beads on a string” nucleosome chains building up 100-kbp chromatin domains are interspersed by
several short segments of 30 nm chromatin fibers (four 30 nm fiber segments with 12.4 kbp are
visualized as short cylinders).
D. Simulation of a 1-Mbp chromatin domain was performed as in (B) with the exception that for one
of the 100-kbp chromatin domains (denoted in yellow), a lower tensegrity force was applied. Note
that the resulting relaxed chain structure of the yellow 100-kbp domain expands in the periphery of
the 1-Mbp domain along the surface of other compacted 100-kbp domains. This peripheral
expansion of decondensed chromatin domains would represent an ”open” 100-kbp chromatin
domain configuration that allows accessibility to transcription complexes preformed in the ICD
space.
E. Simulation was performed as in (B) with the difference that a lower tensegrity force was applied to
all ten 100-kbp domains resulting in moderately compacted 100-kbp domains (compare Fig. 7B).
As a consequence of their decreased compaction, these domains expose more chromatin at the
chromatin domain surface. Furthermore, factor complexes which are excluded from the interior of
highly compacted, ”closed” chromatin domains (compare B) may now penetrate into the interior.
F. Here, for all ten 100-kbp domains only a very low tensegrity force was applied corresponding to a
random walk behaviour of the 10 nm nucleosome chains. The effects of decreasing tensegrity
forces on chromatin organization noted in (E) is still more pronounced. In addition to a low
compaction of all 100-kbp chromatin domains that facilitates the penetration of large factor
complexes in the domain interior (compare Fig. 7E), also the compartmentalization the 100-kbp
domains into distinct entities is lost (compare Fig. 7A and 8B).
48
Fig. 9
Numerical calculations to estimate the size of Donnan potentials expected in nuclei of living cells.
The calculations are based on the Debye-Hückel and Donnan potential theory and were performed
under highly simplified assumptions.
A. Dependence of the Debye Length DL on the concentration of mobile counterions.
B. Distance dependence of the normalized repulsive force Frep/F0 of two chromatin domains for
different concentrations c of mobile counterions in the ICD space.
C. Donnan potential plotted as a function of the concentration of mobile counterions c in the presence
of various concentrations of fixed ions [Y].
D. Estimate of the distance x (t) between a fixed chromatin domain A and an adjacent, mobile
chromatin domain B produced by the repulsive electric forces Frep (compare B). For this rough
estimate we neglected Brownian movements and assumed that x = 0 for t = 0.
Fig. 10
Shift in the recorded electric potential at the microelectrode penetration of a fixed human cell nucleus.
Ordinate: potential in mV, abscissa: time in s. Cell nuclei from diploid human lymphocytes were
isolated and fixed using standard procedures. Briefly, human blood was mixed with Heparin and
Ringer solution 1:1 to prevent coagulation. Lymphocytes were separated from the plasma and
erythrocytes by density centrifugation using UNI-SEP tubes (WAK-Chemie, Homburg, FRG) by
centrifugation for 20 min at 1000g. The lymphocytes were extracted and washed with PBS-Buffer. For
isolation of cell nuclei, cells were treated 5 min. with hypotonic solution composed of 50 mM KCl at
37°C in order to rupture the cell membranes. Cell nuclei were separated from the cytosolic fraction by
repeated centrifugation at 100g for 15 min and removal of the supernatant. Methanol/acetic acid (3:1)
or 4% formaldehyde was dropped on a cell nuclei pellet. Cell nuclei were washed and stored in the
quasi cytosolic solution (see legend of Fig. 11) in which the cells were bathed during the experiments.
The graph shows an electrophysiological recording using a sharp microelectrode filled with 3M KCl
(82 M Ω resistance) in this quasi cytosolic solution (see legend of Fig. 11). Repeated impalements of a
given nucleus resulted in reproducible shifts of the recorded electric potential.
49
Fig. 11
Electric Potential (mV) of formaldehyde fixed tumor T-Cells measured at various charge
concentrations.
The graph shows the result of electrical measurements at different charge concentrations. The quasi
cytosolic solutions used contained 2 mM MgCl2, 0.15 mM CaCl2 (pCa = 8), 1.1mM EGTA, 10 mM
HEPES, pH = 7.2, and various concentrations of K and Na at the ratio of 11:1. Six different solutions
with charge concentrations of 10, 72, 124, 240, 394 and 1300 mM were used. Error bars show ± 1 S.D.
The continous line shows a calculated curve assuming a Donnan-potential.
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