Einstein Quart. J. Biol. and Med. (2001) 18(3):113-122
Chromatin Higher Order Structure and Chromosomal Positioning
in the Nucleus
Matthew Schmerer
Department of Developmental and Molecular Biology
Albert Einstein College of Medicine
Bronx, New York 10461
Abstract
While it has been known for some time that chromatin
is highly packaged, it has only recently been discovered
that chromatin is attached to the nuclear matrix. It has
also been recently shown that interphase chromosomes
remain separate from one another in so called, “chromosome domains.” This author suggests a functional
relationship between chromatin attachment to the
nuclear matrix and chromosomal domains in the interphase nucleus. Herein, the evidence for the presence of
matrix attachment sites and chromosomal domains is
presented, and it is proposed that chromatin attachment
to the nuclear matrix provides the physical foundation
for interphase chromosomes to be localized to chromosome domains.
Introduction
It has long been known, based on electron microscopy
studies, that chromatin is packaged into nucleosomes
(Olins and Olins, 1974), then into 10 nm fibers (Cameron
et al., 1979), and then into 30 nm solenoids (Finch and
Klug, 1976; Felsenfeld and McGhee, 1986). This knowledge raised, among other issues, the question of how
chromatin gets packaged into metaphase chromosomes.
Beyond the issue of chromatin packaging is the organization of the interphase nucleus and where chromatin,
as linear uncondensed chromosomes, fits in. This question was first addressed by David Comings in 1968 when
he described, “The Rationale for an Ordered
Arrangement of Chromatin in the Interphase Nucleus”
(Comings, 1968). In this paper, Comings described the
evidence at that time for the non-random localization of
chromatin in the nucleus. This includes some of the first
evidence that DNA replication occurs, to a greater
extent, at specific points on the nuclear membrane and
along the edge of the nucleolus, as well as some suggestive observations that chromatin is ordered in a specific
way in the nucleus. Since then we have learned a great
deal about the internal structure of the nucleus and how
it is compartmentalized. We now have evidence for
rRNA processing in the nucleolus (Sheer and Benavente,
1990) and for RNA Polymerase II transcription at certain
locations in the nucleus, but not the nucleolus (van Driel,
1995). It is also known that the nucleus has an internal
skeleton, called the nuclear matrix, where the machinery
for DNA replication and transcription is anchored
(Nelson, 1986). The nuclear matrix also seems to be a site
for chromatin attachment, which may provide a foundation for the higher order structure of the chromatin
(Nelson, 1986; van Driel, 1995). This concept of nuclear
compartmentalization alluded to the hypothesis that
whole chromosomes or parts of chromosomes may occupy certain regions of the nucleus (Manuelidis, 1985,
1990).
The existence of chromatin loops and chromosome
domains lend support to the contention that the nucleus is a highly structured organelle. The first level of chromatin organization is chromatin loops. The chromatin of
uncondensed chromosomes in interphase nuclei is
attached to the nuclear matrix at regular intervals making loops of 5 to 100 kb in length (Cook and Brazell,
1976; Paulson and Leammli, 1977). The second level of
chromatin organization is the location of the uncondensed chromosomes in the nucleus. Chromosomes
occupy unique domains in the nucleus such that individual chromosomes do not overlap (Manuelidis, 1985,
1990). This author would like to suggest that these two
levels of organization are linked together, such that
chromatin loops, which attach to the nuclear matrix provide a foundation for the localization of interphase
chromosomes to unique domains.
Chromatin Loops and SARs/MARs
The chromatin of interphase nuclei is attached to the
nuclear matrix at specific sites called matrix association
regions (MARs) or scaffold attachment regions (SARs).
These names refer to the same sequences that are associated with the nuclear matrix after high salt treatment
or ionic detergent treatment (Razin and Gromova, 1995)
and will be hereto referred to as MARs only (see Fig. 1).
The purpose of the high salt treatment is to remove the
histones from chromatin, so that the DNA will be more
accessible to nuclease treatment. Unfortunately, this
preparation method leaves behind actively transcribing
and replicating DNA as well as nuclear matrix-associated
DNA sequences (Razin and Gromova, 1995). This makes it
difficult to map specific sequences, but is sufficient for
the initial visualization of the chromatin loops themselves (Paulson and Laemmli, 1977; Hancock and Hughes,
1982).
The presence of chromatin loops was also suggested
based on ultra-centrifugation of high salt-extracted
nuclei (Cook and Brazell, 1976). Based on those initial
studies, a model was proposed where the chromatin
loops are composed of the 30 nm solenoid, which is constricted between MARs (see Figure 2) (Marsden and
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Figure 1: Mapping of relative positions of unique DNA sequences within the DNA loops by progressive detachment of DNA from the nuclear
matrix. Three extended DNA loops (after histone removal) are shown, one of them bearing three sequence elements (a, b and c) whose relative
positions in the loop are to be determined. After mild (A) or more intensive (B) treatment of the sample with a nuclease, the matrix-bound and
cleaved-off DNA are separated by differential centrifugation. The distantly located sequence element a and b, while element c will still be recovered in a matrix-bound DNA fraction. Knowing the average loop length and the percentage of DNA in the nuclear matrix preparations possessing different subsets of probed sequence elements, one can estimate the approximate positions of these elements within the loop. (Adapted from
Razin and Gromova, 1995).
Laemmli, 1979). Firm evidence for the existence of these
loops came from the identification of the MAR
sequences. These sequences were identified using
Exonuclease III (Exo III) sensitivity assays followed by
restriction mapping to identify the location of fragments
that are resistant to Exo III digestion. Blasquez et al.
(1989) used this technique to identify MAR sequences in
the mouse light chain immunoglobulin gene (see Figure
3). The numbered bands (1-11) in the matrix lanes indicate sequences that were protected from Exo III digestion, because they were bound to the nuclear matrix. It
is important to note the periodicity of the attachment
sites. It is this periodicity that leads to the loop structures. Other groups used similar approaches to determine the MAR sites within the Drosophila histone gene
cluster (Gasser and Laemmli, 1986) and the Drosophila
hsp70 gene (Mirkovitch et al., 1984). A fortuitous observation was that MARs often contain topoisomerase II
binding sites (Cockerill and Garrard, 1986a). Razin et al.
(1993) took advantage of the presence of topoisomerase
II binding sites as a way to map MAR sequences in the
chicken β-globin domain (see Figure 4 for a schematic of
the procedure). These data showing MAR sequences in
different genes of one organism and in genes from different organisms provide strong evidence for the evolutionary conservation of chromatin loops (see Cockerill
and Garrard, 1986b for further discussion).
What is the significance of these loops? Do they have
any function other than as a means of packaging DNA in
the nucleus? The observation that the nuclear matrix
and replication origins are bound to each other was first
made almost thirty years ago by David Comings (1968)
and was demonstrated more recently by Razin et al.
(1986). These chromatin loops and their attachment to
the nuclear matrix through MARs may be important for
initiating replication or for facilitating the process of
replication by bringing the DNA to the replication
machinery. They may also be important for localizing
certain sequences to specific compartments in the nucleus in order to silence them or bring them near the transcriptional machinery, but this is only speculation.
Chromatin Higher Order Structure
115
Figure 2: DNA domains in nuclei. Elements that organize chromosomal loops in nuclei, termed “matrix association regions” (MARs), are depicted as solid rectangles (DNA sequences) and circles (proteins) (Adapted from Blasquez et al., 1989).
Evidence for DNA localization to certain domains for the
purpose of silencing or activating stretches of the DNA is
present more at the level of chromosome domains.
tion in the nucleus, it is clear that interphase chromosomes remain separate entities and the domains they
form remain exclusive of the nucleolus.
Chromosome domains
The most popular techniques for determining chromosomal domains are in situ hybridization with biotin labeled
probes followed by electron microscopy, fluorescent in
situ hybridization (FISH), pulse labeling of cells with
BrdU (or similar analog) followed by confocal microscopy
(see Figure 5), or injection of a fluorescent nucleotide
analog (Cy3-AP3-dUTP) followed by video microscopy or
confocal microscopy (Leitch et al., 1990; Sadoni et al.,
1999; Zink et al., 1998). The in situ hybridization techniques are very powerful techniques for localizing DNA
in the nucleus, but they require fixation of the cells at
some point during the procedure, which could introduce
artifacts. These artifacts could distort the actual chromosomal positioning in vivo. The technique developed by
Zink et al. (1998) provides a much better approach,
because labeled chromatin can be observed in live cells,
which effectively removes the artifacts introduced from
fixation. This technique provides the added benefit of
video recording labeled chromatin in live nuclei to determine, in real time, if chromosome domains move
through the nucleus or remain fixed in place. In fact,
their results demonstrate that there is some shifting of
chromatin within domains, but the domains themselves
remain fairly stationary (see Figure 6). In contrast to this,
Brown et al. (1999) and Bridger et al. (2000) demonstrate
that when cells change from a proliferative state to a
Uncondensed chromosomes in the interphase nucleus do
not unravel and then randomly associate with each
other, but remain segregated from each other in their
own chromosomal domains (Manuelidis, 1990). The
domain organization of interphase chromosomes has
been observed in Drosophila (Mathog and Sedat, 1989),
plants (Leitch et al., 1990), and mammals (Zink et al.,
1998; Manuelidis and Borden, 1988). This demonstrates
that these domains have been conserved throughout
evolution in eukaryotic organisms. During metaphase,
the condensed chromosomes adopt what has become
known as the Rabl orientation, where the centromeres
are collected at one side of the cell and the telomeres
are at the other side (Rabl, 1885; Manuelidis and Borden,
1988). It has been suggested that during interphase, the
uncondensed chromosomes continue to adopt this orientation leaving a gap in the middle of the nucleus for
the nucleolus (Avivi and Feldman, 1980). Manuelidis and
Borden (1988) proposed that this may not be a ubiquitous feature. Their studies of human central nervous system cells using biotin labeled probes, in situ hybridization, and three-dimensional reconstruction of nuclei
indicate that some interphase chromosomes do not
adopt this Rabl orientation. Regardless of their orienta-
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Figure 3: Association of the X gene MAR with the nuclear matrix occurs through multiple specific binding sites. A 341-bp fragment encompassing the X gene MAR was 5' end labeled, in either the upper or lower strand, opposite to a 3' overhand. Exonuclease III digestion was performed after specific binding to plasmacytoma matrices (prepared using micrococcal nuclease instead of Dnase I digestion), and matrix-bound
DNA was purified for electrophoretic separation on a sequencing gel. Naked DNA was similarly digested as a control. Barriers toward the 5' side
(1-5) and the 3' side (6-11) are indicated. Four pairwise combinations of these barriers are shown, each of which leads to approximately 76 bp
segments (Adapted from Blasquez et al., 1989).
quiescent state, either certain sets of genes or whole
chromosomes change their position in the nucleus.
Brown et al. (1999) showed that a set of transcriptionally inactive genes associate with a centromeric heterochromatin region in the nucleus in dividing lymphocytes, but not in non-dividing lymphocytes. They also
demonstrated that when non-dividing cells are stimulated to divide, there is a shift of this set of genes back to
the centromeric heterochromatin. This demonstration of
a shift in nuclear chromosome positioning is at the gene
level; Bridger et al. (2000) demonstrated a similar phenomenon at the level of whole chromosomes. They
looked at the positions of chromosomes 18 and 19 in
proliferative and quiescent human fibroblasts. In proliferative fibroblasts, chromosome 18 is associated with the
nuclear periphery, and chromosome 19 is associated with
the nucleolus. As cells exit the cell cycle and enter a quiescent state after serum starvation, chromosomes 18 and
19 lose their associations with the nuclear periphery and
the nucleolus, respectively. Surprisingly, when serum is
added back followed by several passages, chromosomes
18 and 19 reestablish their associations with the nuclear
periphery and nucleolus, respectively. This suggests that
chromosomes may actively move around the nucleus,
and that this movement has a functional relationship to
the proliferitive state of the cell.
Comings (1968) suggested that chromosomes may be
localized to certain locations in the nucleus. There seems
to be some evidence that heterochromatic regions of
chromosomes are localized to the nuclear membrane or
the outer periphery of the nucleolus while euchromatic
regions are localized internally (Avivi and Feldman,
1980). The best example of a whole chromosome being
localized to a reproducible location in the nucleus is the
inactivated X chromosome in mammals. Dyer et al.
(1989) did in situ hybridization studies of human female
cell nuclei and showed that the inactive X chromosome
almost always associates with the heterochromatic sex
chromosome body (SCB). They also showed that in
hybrids between human and mouse nuclei, this close
association of the X chromosome with the SCB is lost.
This seems to suggest a breakdown of the heterochromatic region in these hybrids. Sanchez et al. (1997) did
localization studies of certain chromosomes, including
the X and Y chromosomes, in human neutrophil nuclei.
They determined that the X and Y chromosomes seemed
to localize to protrusions of the neutrophil nucleus (see
Chromatin Higher Order Structure
117
Figure 4: Excision of genomic DNA loops by topoisomerase II-mediated DNA cleavage at matrix attachment sites. (A) In living cells, the 30 nm
chromatin fibril is arranged into loops by periodic attachment to the nuclear scaffold. The latter contains topoisomerase II, which can cut DNA
at loop basements. In addition, soluble topoisomerase II can interact with DNA at a number of accessible sites, creating an apparently random
cleavage pattern (as far as the low-resolution analysis by PFGE is considered). The resulting cleavage pattern (panel I) will represent the superimposing of this random cleavage pattern over the regular pattern of cleavages at loop basements. (B) With the soluble topoisomerase II removed
by high-salt extraction, the cleavages can occur only within matrix attachment sites. Varying the concentration of VM-26, it is possible to cut DNA
at all loop basements (panel 3) or less frequently so that multimers of loops are also released (panel 2) (Adapted from Razin and Gromova, 1995).
Figure 7), while somatic chromosomes seem to localize in
a more random pattern. This argues against a general
theory of specific localization of chromosomes in the
nucleus. An interesting observation made by Borden and
Manuelidis (1988) was that in cells derived from human
females with epilepsy, the inactive X chromosome loses
its association with the heterochromatic region. Somatic
chromosomes and the Y chromosome were also tested,
and their heterochromatic regions remain associated
with the periphery of the nucleolus or the nuclear membrane. This seems to suggest a connection between the
disease state and the localization of certain chromosomes in the nucleus.
Certain domains within chromosomes seem to localize in
a non-random way within the nucleus. Even if the whole
chromosome is not specifically positioned, certain portions of it may be localized. Satellite sequences, which
cluster in certain portions of the chromosome, including
the centromere, non-randomly associate with certain
portions of the nucleus. Manuelidis (1984) demonstrated
this using biotin-labeled, centromere-specific satellite
DNA that was hybridized in situ to fixed tissue from
mouse cerebellum. She showed that individual neuronal
cell types display reproducible patterns of centromere
localization to the nuclear membrane or to areas adjacent to the nucleolus. However, no consistent pattern of
centromere localization was seen in all of the cell types
that were studied.
Ferreira et al. (1997) and Sadoni et al. (1999) did studies
of chromosomal replication patterns in multiple cell
types. They employed the use of BrdU replication-labeling alone, Cy3-dUTP replication-labeling alone, or a
unique double labeling procedure where they pulse
labeled during S phase with Cy3-dUTP and then labeled
in the next S phase with BrdU. The double labeling procedure allowed them to distinguish between DNA
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Figure 5: Metaphase spread with two differently sized (non-homologous), replication-labeled chromosomes. Note that only one of the two sister chromatids contains the label. Chromosomes were counterstained with DAPI (blue). Interphase nuclei in B counterstained with DAPI from the
same slide as the metaphase shown in A depict replication-labeled and segregated chromatid territories. Small patches of replication-labeled chromatin (arrow) most likely result from sister chromatid exchanges (Adapted from Zink et al., 1998).
Figure 6: Time-lapse recording of one nucleus from the neuroblastoma cell line. Cy3-AP3-dUTP-microinjected cells were grown for several cell
cycles. Each panel shows a projection of optical sections. 3D stacks were recorded every 20 minutes for 5 hours. The nucleus performed rotational and translational movements during the entire period. The framed segregated territory is clearly separated from the rest of the labeled territories. After a major initial rearrangement (indicated by large arrowheads), this territory maintained its overall shape. The inset (upper right)
within the first six panels shows an enlargement (factor 2.3) of subdomains (arrow). The subdomains display changing patterns of foldings and
extensions (some examples are indicated by arrowheads) (Adapted from Zink et al., 1998).
Chromatin Higher Order Structure
119
Figure 7: Deconvolved volumetric views of neutrophil nuclei from healthy females (top two panel pairs) and males (bottom two panel pairs).
The leftmost panel of each corresponds to a volumetric view of a nucleus stained for total DNA with DAPI (negative image). The rightmost panel
of each pair corresponds to a volumetric view of a nucleus hybridized for the X chromosome and the Y chromosome centromere sequences. In
these panels, the outline of the nuclei is indicated by a white dotted line and the asterisks mark the position of the sex-specific appendages.
Notice the presence of an X chromosome in the appendage of the female nucleus, a Y chromosome in the appendage of the male nucleus, and
the absence of chromosome pairing when the sex chromosomes reside in the same lobe. Bar represents 5 µm (Adapted from Sanchez et al., 1997).
labeled at a first S phase and DNA labeled at the next S
phase. Using these labeling techniques, Ferreira et al.
(1997) demonstrated that early replicating portions of
chromosomes (R bands that are identified by a light
Giemsa staining on chromosomes) preferentially localize
to interior regions of the nucleus, while late replicating
portions (G/Q bands that are identified as dark Giemsa
staining on chromosomes) preferentially localize to
peripheral regions. Sadoni et al. (1999) verified these
results (see Figure 8) and further showed, using their
double labeling technique, that S phase labeling patterns
are maintained from division to division. So, not only are
regions of chromosomes that replicate at different times
in S phase localized to different portions of the nucleus,
but that pattern is maintained in a cell type-specific manner with every cell division. These results provide a potential structural explanation for the tissue-specific expres-
sion patterns of different genes. It is noteworthy that
housekeeping genes are mostly localized in R bands,
whereas tissue specific genes are almost exclusively located in G/Q bands (Craig and Bickmore, 1993, 1994).
Conclusion
Two levels of chromatin structure have been presented,
that of chromatin loops and chromosome domains.
Marsden and Laemmli (1979) proposed that the 30 nm
solenoid chromatin filaments were attached at regular
intervals to the nuclear matrix forming chromatin loops.
The sequences that mediate the attachment of the DNA
to the nuclear matrix (MARs) have been extensively studied (Blasquez et al., 1989; Gasser and Laemmli, 1986;
Mirkovitch et al., 1984; Razin et al., 1993). The attachment of chromatin to the nuclear matrix has also been
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Figure 8: S phase replication labeling patterns are preserved. Cells ([a and b] Hv, human diploid fibroblasts; [c and d] SH-EP N14 human neuroblastoma cells; [e and f] HeLa cells; [g and h] CHO cells; and [i and j] C2C12 mouse myoblasts) were replication-labeled with BrdU (a-d, i, and
j; 30-min pulses) or Cy3-dUTP (e-h). Cells were fixed immediately after BrdU labeling or 30 minutes after microinjection of Cy3-dUTP to obtain
labeled S phase ells (left, a,c,e,g, and i). S phase cells display the typical replication labeling patterns (indicated on the left): (a) type I, (c) type II,
(e) type III, (g) type IV, and (i) type V. Similarly labeled cells were grown for one to five hours. after labeling and fixed after this growth period
(right, b,d,f,h, and j). The presence of similar patterns (types indicated on the right) several days after S phase labeling suggests that DNA replicating at a particular time during S phase always occupies similar nuclear positions. Arrowheads mark the nucleoli that are always unlabeled.
Arrowheads in c-e mark perinucleolar label. Arrows in b and d indicate unlabeled chromosome territories. The presence of unlabeled territories
demonstrates that cells do not display similar patterns because they stopped cycling, but rather that they went through at least two mitoses after
labeling. Note also the two very close nuclei in f displaying similar type III patterns, suggesting that similar patterns were conserved in sister nuclei
after cell division. All pictures display single light optical sections through midnuclear planes except i and j, which show epifluorescence images.
Bar in a is similar for all images (Adapted from Sadoni et al., 1999).
Chromatin Higher Order Structure
implicated in the process of replication initiation (Razin
et al., 1986). The suggestion that interphase chromosomes are specifically positioned in the nucleus was first
made by Comings in 1968. It has since been shown that
chromosomes occupy specific domains in the nucleus and
do not overlap in the interphase nucleus (Manuelidis,
1990; Leitch et al., 1990). There is also a great deal of evidence to suggest that certain portions of chromosomes
are non-randomly arranged in the nucleus (Manuelidis,
1984; Ferreira et al., 1997; Sadoni et al., 1999).
How do these two levels of chromatin organization connect with one another? What is their functional connection, if any? This author believes that the chromatin
attachments to the nuclear matrix in the form of chromatin loops may provide the “foundation” for the localization of, at least, specific domains of chromosomes if
not whole chromosomes. Some of the most striking evidence for a structural basis to this model comes from the
work of Ma et al. (1999). When they did a classical
extraction for nuclear matrix in the absence of DNaseI
followed by FISH, they showed that chromosome
domains remain intact. In contrast to this, when they did
a complete matrix extraction by doing RNase A treatment, to remove interactions with RNAs or RNPs, followed by 2.0 M NaCl, the chromosome domains are disrupted. This suggests that chromosome domains are
attached to the nuclear matrix most likely through the
attachment of chromatin loops to the matrix. This model
is also functionally supported by the observation that
chromatin is attached to the nuclear envelope at the site
of replication origins (Razin, 1986), which would bring
these regions of chromosomes near the nuclear pore
complexes. The proximity of the DNA and replication
complexes near the nuclear pore complexes means that
these complexes have ready access to nucleotide precursors and other cytoplasmic factors. Razin and Gromova’s
(1995) “channels model” of nuclear matrix structure
incorporates some of this idea. Their model postulates
that the nuclear matrix forms channels near the nuclear
pore complexes. These channels have large lateral extensions coming off of the main channel. The chromatin
loops are then attached to the matrix along these channels. The model provides a good explanation for the differential sensitivity of loop attachment sites to nuclease
treatment after the different extraction methods. It also
supports the observation that newly synthesized mRNAs
seem to track toward the nuclear pore complex and presumably out of the nucleus (Lawrence et al., 1989;
Kramer et al., 1994). These nuclear matrix channels may
be the means by which mRNAs exit the nucleus; this
remains to be seen. These channels may exist within
chromosome territories or, as Cremer et al. (1993) suggest, they may exist between chromosome territories.
They suggested that the interchromosomal spaces may
provide channels for newly synthesized mRNAs and
other precursor molecules to move in and out of the
nucleus. Both of these models provide a framework
121
under which the two levels of chromatin organization
can be understood. Undoubtedly, more work needs to
be done in order to fully understand the functional
organization of the nucleus and the functional relationships between chromatin loops, chromosome territories,
and the replication and transcription machinery.
Acknowledgments
The author would like to thank Rajarshi Ghosh, Nicole
Rempel, and Dr. U. Thomas Meier for their help and constructive criticism.
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