Chromosoma (2004) 112: 410–423
DOI 10.1007/s00412-004-0287-3
RESEARCH ARTICLE
Irina Solovei . Lothar Schermelleh . Klaus Düring .
Andrea Engelhardt . Stefan Stein . Christoph Cremer .
Thomas Cremer
Differences in centromere positioning of cycling and postmitotic
human cell types
Received: 10 March 2004 / Revised: 14 April 2004 / Accepted: 15 April 2004 / Published online: 9 June 2004
# Springer-Verlag 2004
Abstract Centromere positioning in human cell nuclei
was traced in non-cycling peripheral blood lymphocytes
(G0) and in terminally differentiated monocytes, as well as
in cycling phytohemagglutinin-stimulated lymphocytes,
diploid lymphoblastoid cells, normal fibroblasts, and
neuroblastoma SH-EP cells using immunostaining of
kinetochores, confocal microscopy and three-dimensional
image analysis. Cell cycle stages were identified for each
individual cell by a combination of replication labeling
with 5-bromo-2′-deoxyuridine and immunostaining of
pKi67. We demonstrate that the behavior of centromeres
is similar in all cell types studied: a large fraction of
centromeres are in the nuclear interior during early G1; in
late G1 and early S phase, centromeres shift to the nuclear
periphery and fuse in clusters. Peripheral location and
clustering of centromeres are most pronounced in noncycling cells (G0) and terminally differentiated monocytes. In late S and G2, centromeres partially decluster and
migrate towards the nuclear interior. In the rather flat
nuclei of adherently growing fibroblasts and neuroblastoma cells, kinetochores showed asymmetrical distributions
with preferential kinetochore location close either to the
bottom side of the nucleus (adjacent to the growth surface)
or to the nuclear upper side. This asymmetrical distribution of centromeres is considered to be a consequence of
chromosome arrangement in anaphase rosettes.
Communicated by E.A. Nigg
I. Solovei (*) . L. Schermelleh . K. Düring . A. Engelhardt .
T. Cremer
Department of Biology II, Humangenetik, Ludwig
Maximillians University (LMU),
Richard Wagner Str. 10,
80333 Munich, Germany
e-mail: [email protected]
S. Stein . C. Cremer
Kirchhoff Institute of Physics, University of Heidelberg,
69120 Heidelberg, Germany
Introduction
The interphase nucleus has emerged as a dynamic
organelle with rapid movements of many nuclear proteins
and RNA molecules. In contrast, interphase chromatin
appears relatively immobile (Abney et al. 1997; Bornfleth
et al. 1999; Manders et al. 1999; Dundr and Misteli 2001;
Gerlich et al. 2003; Walter et al. 2003). Nevertheless, some
chromosome subregions demonstrate more extensive mobility. Giant chromatin loops containing gene clusters have
been reported to extend outwards from the surface of
corresponding chromosome territories upon gene(s) activation (Volpi et al. 2000; Mahy et al. 2002; Williams et al.
2002). In mouse lymphocytes, dynamic repositioning of
genes towards the chromocenter clusters or away from
them has been described in association with gene silencing
and activation (Brown et al. 1997, 1999). Extensive rapid
movements of certain chromosome loci have also been
reported for nuclei of budding yeast and of Drosophila
embryos (Csink and Henikoff 1998; Gasser 2002; Marshall 2002).
Centromeres provide another example where major
movements have been observed in nuclei of both postmitotic cells (Manuelidis 1985; Alcobia et al. 2000; Martou
and De Boni 2000; Solovei et al. 2004a) and cycling cells
(Bartholdi 1991; Ferguson and Ward 1992; Weimer et al.
1992; Vourc’h et al. 1993; Hulspas et al. 1994). Previous
studies of the distribution of centromeres/kinetochores in
cycling and non-cycling cell types had important technical
limitations, including the unequivocal discrimination of
non-cycling cells (G0) from cells at different stages and
substages of interphase (see Discussion). As already
emphasized by Ferguson and Ward (1992) “the morphological preservation of specimens through the various
techniques of isolation, fixation, and hybridization is of
the utmost concern.” Preservation of the three-dimensional
(3D) structure of cells and their nuclei during fixation is a
prerequisite for reliable morphological studies using
immunostaining and/or fluorescence in situ hybridization
(FISH) techniques in combination with confocal microscopy (Bridger and Lichter 1999; Solovei et al. 2002a). In
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FISH studies, the detrimental effects of the heat denaturation step have to be taken into account (Solovei et al.
2002a,b). While this step apparently does not lead to a
major distortion of higher order chromatin arrangements, it
seems advantageous, whenever possible, to use protocols
that avoid this step.
Considering these limitations we decided to perform
studies of 3D centromere arrangements in five human cell
types (normal diploid fibroblasts, normal lymphocytes and
diploid lymphoblastoid cells, normal peripheral blood
monocytes and neuroblastoma SH-EP cells) employing
protocols for centromere/kinetochore visualization that
keep the 3D nuclear architecture intact to the best possible
extent. Precise identification of the quiescent or cycling
stage of each individual cell was based on the nuclear
patterns of pKi67 (Kill 1996; Starborg et al. 1996; Bridger
et al. 1998) and replication labeling with 5-bromo-2′deoxyuridine (BrdU) (O’Keefe et al. 1992; van Driel et al.
1998). Following fixation of cells that preserved the 3D
structure, centromeres were visualized either by kinetochore immunostaining or by 3D-FISH with a pancentromeric probe. The 3D centromere/kinetochore arrangements were assessed in light optical image stacks from
nuclei recorded by a confocal microscope.
Materials and methods
Cell types and 3D fixation
Human lymphocytes (G0) and monocytes from peripheral blood
were isolated in a Ficoll gradient, and incubated for 3 days in RPMI
1640 supplemented with 20% FCS and 1% phytohemagglutinin
(PHA). A diploid lymphoblastoid cell line was established in our
laboratory from a healthy male donor; these cells were grown in
RPMI 1640 supplemented with 15% FCS. For 3D-preserving
fixation, lymphoblastoid cells and lymphocytes were harvested and
resuspended in fresh medium with 50% FCS at a final concentration
of ca. 1×106 cells/ml. Three hundred microliter aliquots of the
suspension were placed on coverslips coated with polylysine (1 mg/
ml). Cells were allowed to attach for about 1 h at 37°C, and then
fixed in 4% paraformaldehyde in 0.3×PBS. To prevent shrinkage of
spherical lymphocyte and lymphoblastoid nuclei, cells were briefly
(1 min) incubated in 0.3×PBS before fixation. Human skin
fibroblasts were grown directly on coverslips in DMEM supplemented with 10% FCS. Neuroblastoma SH-EP cells [47, XX, der
(1;14) t (1;14)/der (7) t (7;8)/der (8) t (7;8)/der (22) t (17;22)] were
kindly donated by Prof. W.W. Franke, DKFZ, Heidelberg) and
subcultured on coverslips in RPMI supplemented with 10% FCS.
The two adherently growing cell types were fixed in 4% paraformaldehyde in 1×PBS. In all cases, special coverslips (26×76 mm)
with an even thickness of 0.17±0.01 mm (Assistent, Germany) were
used for cell attachment and growth. These coverslips are
recommended for optimized confocal microscopy measurements.
After fixation, the cell membrane and the nuclear envelope were
permeabilized by incubation in 0.5% Triton X-100. The 3D fixed
and permeabilized cells were stored in PBS with 0.04% sodium
azide at 4°C until use.
Synchronization of neuroblastoma SH-EP cells
SH-EP cells were subcultured on coverslips in full medium (RPMI
1640, 10% FCS) until 40–50% confluency. Effective synchroniza-
tion of SH-EP cells was obtained with the following protocol.
Coverslips with subconfluent SH-EP cells were transferred to
serum-deprived medium (RPMI 1640 without FCS) for 14–24 h.
Thereafter cells were transferred to full medium supplemented with
aphidicolin (1 μg/ml) to block DNA polymerase (Ikegami et al.
1978). After 12 h cells were released from the block by washing
three times and transferred to medium with 10% FCS. The degree of
synchronization and duration of the cell cycle stages were
determined using BrdU incorporation and pKi67 staining (see
below). About 95% of cells were in early S phase in 30–60 min after
release from the block; in mid-S phase after 4.5 h; in late S phase
after 6.5 h; in G2 after 7 h; most of the cells entered mitosis 8–9 h
after release. The majority of the cells reached the G1 stage of the
next cell cycle 10–11 h after release. The degree of synchronization
in the next cell cycle was still high: about 85% of the cells could be
simultaneously labeled by BrdU in the next early S.
Identification of cell cycle stages and kinetochore staining
Replication labeling with BrdU was used to identify S phase cells
and to distinguish between cells in early S, mid-S, and late S phase
(O’Keefe et al. 1992). For pulse labeling, cells were incubated in
culture medium with 10 μg/ml BrdU (Sigma) for 30 min. To avoid
morphological alterations caused by standard methods of BrdU
detection due to the DNA denaturation step (van Driel et al. 1998),
detection of BrdU in fixed cells was performed following the 3Dpreserving protocol described by Tashiro et al. (2000). Cells were
incubated with mouse anti-BrdU antibodies (Roche) in a solution
composed of 0.5% BSA, 0.5×PBS, 30 mM TRIS, 0.3 mM MgCl2,
0.5 mM 2-mercaptoethanol, and 10 μg/ml DNase I (Roche); then
sheep anti-mouse Cy3-conjugated (Jackson ImmunoResearch Laboratories) secondary antibody was applied. Cells at G0 and early
G1 stages were identified by staining with antibodies against protein
Ki67 (Dianova) as described by Bridger et al. (1998). For
kinetochore staining, rabbit anti-CENP-B and rabbit anti-CENP-C
antibodies (kindly donated by Prof. W.C. Earnshaw, University of
Edinburgh, UK) were used either separately or as a 1:1 mixture with
identical results. Alexa 488-conjugated goat anti-rabbit antibody
(Molecular Probes) was applied as the secondary antibody. All
antibodies (except anti-BrdU) were diluted in blocking solution
containing 1×PBS, 0.05% Triton X-100, and 3% BSA; all washings
were performed in 1×PBS with 0.05% Tween at 37°C. Nuclear
DNA was counterstained with TO-PRO-3 (Molecular Probes) and
4’,6-diamidino-2-phenylindole (DAPI) (Sigma). Cells were
mounted in Vectashield (Vector Laboratories) antifade medium.
Three-dimensional fluorescence in situ hybridization with a
pancentromeric probe
Normal human diploid fibroblasts were fixed and prepared for threedimensional fluorescence in situ hybridization (3D-FISH) according
to standard protocols (Solovei et al. 2002a). Briefly, cells were fixed
in 4% paraformaldehyde in 1×PBS, permeabilized with 0.5% Triton
X-100, incubated in 20% glycerol, repeatedly frozen-thawed using
liquid nitrogen, incubated in 0.1 N HCl for 5 min, and stored in 50%
formamide, 2×SSC at 4°C until hybridization. A probe for αsatellite sequences contained in centromere regions of all human
chromosomes (Choo 1997), referred to as a pancentromeric probe,
was generated and labeled by the polymerase chain reaction using
5′-CAT CAC AAA GAA GTT TCT GAG GCT TC and 5′-TGC
ATT CAA CTC ACA GAG TTG AAC CTT CC primers and human
placenta DNA as a template. Following labeling with fluorescein
isothiocyanate-12-dUTP (Roche Molecular Biochemicals) the
pancentromeric probe was shortened to 100–300 bp by DNase
digestion. The probe was dissolved in hybridization mixture (50%
formamide, 10% dextran sulfate, 1×SSC), loaded on a coverslip
with cells, covered with a smaller coverslip, and sealed with rubber
cement. Cell and probe DNA were denatured simultaneously on a
hot-block at 75°C for 2 min. Hybridization was performed for
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2 days at 37°C in humid boxes. Post-hybridization washes were
done in 2×SSC at 37°C and 0.1×SSC at 60°C. After BrdU and/or
pKi67 detection, nuclear DNA was counterstained with TO-PRO-3
and DAPI, and cells were mounted in Vectashield. To preserve 3D
nuclear morphology, air-drying of cells was carefully avoided at all
steps from fixation to mounting in the antifade (Solovei et al. 2002a,
2002b).
Confocal microscopy and image processing
Series of confocal sections through whole nuclei were collected
using a Leica TCS SP equipped with Plan Apo 100×/1.4 NA oil
immersion objectives. For each optical section, images were
collected sequentially for two or three fluorochromes. Fluorochromes were excited using an argon laser with excitation wavelengths of
488 nm (for Alexa 488) and 513 nm (for Cy3), or a helium–neon
laser with excitation wavelength of 633 nm (for TO-PRO-3). Stacks
of 8-bit gray-scale images were recorded with an axial distance
between optical sections of 300 nm and a pixel size of 50 nm.
Galleries of RGB confocal images were assembled using NIH Image
and Adobe Photoshop software packages. Three-dimensional
reconstructions of nuclei and kinetochore or centromere signals
were performed by volume and surface rendering of image stacks
using Amira 2.3 TGS software (http://www.amiravis.com).
Quantitative assessment of the 3D positioning and clustering of
kinetochore signals
Positions of kinetochore signals within the nucleus were classified
as shown in Fig. 1a. Kinetochore signals that abutted the nuclear
border or were separated from it by a distance not exceeding the
diameter of the kinetochore signal were classified as peripheral; all
other signals were scored as internal. For spherical nuclei of
lymphocytes and lymphoblastoid cells, RGB galleries of serial XYoptical sections were used for visual tracing and scoring of
kinetochore signals. Owing to the limited Z resolution of the
confocal microscope, this approach was not applicable to the flat
nuclei of fibroblasts and SH-EP cells. For scoring kinetochore
signals in these cells, image stacks were loaded into the Amira 2.3
program and data were viewed as ZY and/or ZX optical sections
(Fig. 4a). In late G2, most kinetochore signals are duplicated and
appear as doublets. However, since not all kinetochores are seen as
doublets and not all close pairs of signals are true doublets, we
counted all visible kinetochore signals separately.
To analyse the distribution of kinetochores with regard to
different surfaces of the nucleus, peripheral signals were subclassified as top, bottom, and lateral (Fig. 1b). Each cell could therefore be
characterized by proportions of top (Pt) and bottom (Pb) signals to
total number of signals in this cell. Since scatter diagrams with Pb
and Pt axes indicated the presence of two groups of cells in some
stages of the cell cycle, relative numbers of top and bottom signals
in individual nuclei were analyzed in more detail. Regression
equations for Pt and Pb were calculated and regression lines were
used as new x-axes with zero in the point Pb=Pt (R-axes). Positive
Table 1 Cell cycle identification based on intranuclear localization of incorporated 5bromo-2′-deoxyuridine (BrdU),
pKi67 pattern, and kinetochore
signal morphology
Fig. 1a, b Scheme for scoring the centromere signals. a Spherical
nuclei of lymphocytes and lymphoblastoid cells and nuclei of
monocytes. b Flat nuclei of fibroblasts and SH-EP cells. The
position of a kinetochore signal in the nucleus was considered as
peripheral when it was touching the border of the nucleus defined by
the counterstain (1) or was separated from the border by a distance
not exceeding the size of the signal itself (2). Other signals,
including those adjacent to the nucleoli (n), were scored as internal
(3, 4). Kinetochore signals on the surfaces of the flat nuclei facing
the substrate and free surface of the cell were classified as bottom
(1b) and top (1t), respectively; signals adjacent to the lateral surface
of the nucleus were classified as side signals (1s)
and negative values on the R-axis represent nuclei with kinetochores
preferentially located at the top and bottom side, respectively (see
Fig. 6).
In addition to visual tracing and scoring of kinetochore signals in
spherical nuclei of lymphocytes, a special three-dimensional relative
radius distribution (3D-RRD) computer program was used (see
Cremer et al. 2001 for detailed description of the program). This
program finds borders of nuclei based on the DNA counterstain and
calculates the gravity centers of the nuclei. Then the length of the
nuclear radius in any direction is assigned to 100% and the nuclear
space is divided into 25 shells of equal relative thickness (4%). In
this way the average distribution of kinetochore signals can be
presented for a set of nuclei as a function of the their average relative
distances from the center of a normalized nucleus. Signals
distributed randomly in the nuclear DNA should have the same
distribution as the DNA counterstain; deviations from the counterstain curve would indicate a non-random distribution.
Results
Cell cycle stage identification
The definition of cell cycle stages for individual cells was
based on three criteria: the distribution of BrdU replication
label (O’Keefe et al. 1992), the pattern of pKi67
immunostaining (Bridger et al. 1998), and the morphology
of the kinetochore signal (Table 1). Figure 2 presents
examples of lymphocyte nuclei with typical BrdU or
pKi67 staining patterns used for this purpose (see also
Phase
BrdU
Ki67
Kinetochore signal
G1 early
G1 late
S early
S mid
S late
G2 early/mid
G2 late
G0
−
−
+ (nuclear interior)
+ (nuclear and nucleolar periphery)
+ (few internal and external foci)
−
−
−
+
+
+
+
+
+
+
−
Small, single
Small, single
Single
Single
Large, single
Large, single
Large, doublets
Large clusters
(multiple granules)
(nucleoli)
(nucleoli)
(nucleoli)
(nucleoli)
(nucleoli)
(nucleoli)
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Fig. 2a–f Simultaneous detection of kinetochores and cell cycle
stage in nuclei of normal lymphocytes from peripheral blood
stimulated with phytohemagglutinin (PHA). a Four representative
confocal sections (1–4) through the nucleus of a non-stimulated
lymphocyte. Note the large size of the kinetochore signals (green)
produced by clusters of kinetochores on the periphery of the
nucleus. The TO-PRO-3 counterstain is shown in blue. b–f Mid-
sections through nuclei of stimulated lymphocytes at different cell
cycle stages: early G1 (b), G2 (c), early S (d), mid-S (e), late S (f).
Overlay of kinetochore immunostaining (green), immunostaining
with pKi67 (b, c) or 5-bromo-2′-deoxyuridine (BrdU) (d–f) (red),
counterstaining with TO-PRO-3 (blue), and corresponding gray
scale images are shown for each stage. Bar represents 5 μm
Solovei et al. 2004b). The same patterns were also
observed in the other three cell types included in our study.
In early G1, pKi67 is accumulated in granules of various
sizes distributed throughout the nuclear volume, but
mainly associated with heterochromatin regions
(Fig. 2b). In late G1, S, and G2, pKi67 is localized in
the nucleoli (Fig. 2c). Nuclei of postmitotic and quiescent
(G0) cells lack a pKi67 signal (Fig. 2a). Nuclear
incorporation of BrdU marks S phase cells. Early
S phase is characterized by replication of chromatin in
most of the nuclear volume except for chromatin domains
surrounding the nucleoli and adjacent to the nuclear
periphery (Fig. 2d). In mid-S phase, replication is mostly
restricted to the domains located around the nucleolus and
at the nuclear periphery (Fig. 2e). In late S phase, a few
large chromatin clusters are labeled with both peripheral
and internal nuclear locations (Fig. 2f). The morphology
of kinetochore signals, as observed after immunostaining
with antibodies against CENP-B/CENP-C, helped to
distinguish between late G1 (single, small signals) and
late G2 (larger signals, some still appearing as single,
others as doublets) (Table 1).
Clustering of kinetochores varies in cycling and noncycling cells
In all cell types, the number of kinetochore signals per cell
changed during interphase and upon exit from the cell
cycle (Fig. 3). In G1 cells, the number of signals was
always close to the expected 46 for diploid cells and 47 for
SH-EP cells, i.e., most of the kinetochores were separate.
The number of signals in S phase cells was lower than in
G1, indicating kinetochore clustering. The difference
between G1 and S phase was more pronounced in
fibroblasts and stimulated lymphocytes, than in SH-EP
and lymphoblastoid cells. In late G2, most of the
kinetochores were represented by doublets. Each doublet
was counted as two signals, explaining the high number of
signals scored at this cycle stage. The number of signals
observed in G0 nuclei (Fig. 3) was generally smaller than
in nuclei from cycling cells and most G0 signals were
considerably larger than signals observed in cycling cells
(Fig. 2a). This finding indicates a strong clustering of
kinetochores upon exit from the cell cycle.
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Fig. 3 Changes in the mean number of kinetochore signals per
nucleus (numbers above the histogram columns) with cell cycle
stage in the nuclei of fibroblasts, SH-EP neuroblastoma cells,
lymphocytes, and lymphoblastoid cells. Numbers in parentheses are
the numbers of nuclei studied
Kinetochore distribution and dynamics in flat nuclei of
adherently growing cells
Nuclei of normal diploid fibroblasts and SH-EP cells were
relatively flat. Their dimensions changed to some degree
depending on the cell cycle stage, but their ellipsoidal
(fibroblast nuclei) or oval shape (SH-EP cell nuclei)
persisted (Figs. 4a–c, 8a,d,g). The average X, Y, Z
diameters for nuclei of fibroblasts in early S were about
17×12×4 μm. The same parameters for nuclei of early
S phase SH-EP were 15×12×5 μm. For both these
adherently growing cell types we studied six cell cycle
stages (G1 early, G1 late, S early, S mid, S late and midlate G2). To locate the positions of kinetochore signals, 3D
reconstructed nuclei were virtually sectioned in the XY,
ZY and ZX directions (for example see Fig. 4a). Each
nucleus was screened section by section in at least two
(ZX, ZY) directions and signals were classified as
peripheral or internal according to the scheme shown in
Fig. 1a. Kinetochore signals that abutted the nuclear
border or were separated from it by a distance not
exceeding the kinetochore signal size in the xy-plane (or
half the kinetochore size in the z-direction, because, due to
the low resolution of the confocal microscope in the zdirection, the observed z-size of the signals was about
three times greater than the size in the xy-plane), were
classified as peripheral, all other signals were scored as
internal (Fig. 1).
The height of nuclei (size along the z-axis) is,
depending on cell cycle stage, 3–5 μm in fibroblasts and
5–7 μm in SH-EP cells. The observed z-size of kinetochore signals was about 0.6 μm. Random distribution of
kinetochores along the z-axis means in this case an even
distribution. Using the counting rule described above, in
the case of random distribution only 12–20% of kinetochore signals would be classified as peripheral in fibroblasts and 10–15% in SH-EP cells. In actuality at least
65% of kinetochores have peripheral localization at any
stage in both cell types (Fig. 5). Hence, the distribution is
markedly non-random.
Figure 5 provides the percentages of internal and
peripheral kinetochore signals in fibroblast and SH-EP cell
nuclei. Both cell types showed the same trend in
kinetochore distribution: at all interphase stages most
kinetochores were situated peripherally. The proportions
of peripheral kinetochores, however, increased from early
G1 to early S phase but decreased in late S phase and G2.
G0 fibroblasts contained very few internal kinetochores
(SH-EP cells were not studied at this stage). Most of the
internal kinetochores observed in S phase and G0 cells
abutted the nucleolus, which was identified either by its
more intense TO-PRO-3 staining (note: this fluorochrome
stains RNA as well as DNA) compared with the
surrounding chromatin or by staining with anti-pKi67
antibodies (see above). In early G1 and late G2 cells, a
significant proportion of the internal kinetochores were not
associated with the nucleolus.
Peripheral signals were additionally classified with
respect to their location close to the nuclear envelope at
the bottom of the nucleus (the side facing the glass slide on
which cells were grown) or top (the other side of the
nucleus). Signals located at the nuclear edge were scored
as lateral signals (Fig. 1b). The latter were rather few and
excluded from further consideration. With regard to top
and bottom signals, the question arises as to whether
kinetochores are evenly distributed between the top and
bottom sides of the nucleus or not. The first aspect of this
problem is comparison of average proportions of kinetochore signals at the top and bottom sides of the nucleus at
different stages of the cell cycle. In fibroblast nuclei at G1,
S and G2, the mean proportion of kinetochores on the
415
Fig. 4a–g Evaluation of the
spatial distribution of kinetochore signals using Amira software (a–f) and a three-dimensional relative radius distribution (3D-RRD) computer program (g). a SH-EP cell nucleus
in early S phase: XY, XZ, and
YZ maximum intensity projections combined with an XZ
ortho-section. Scrolling through
XZ or YZ ortho-sections allowed counting of the peripheral
and internal kinetochore signals.
b–f Three-dimensional reconstructions (by volume rendering)
combined with XZ ortho-section
through nuclei of fibroblast (b)
and SH-EP cell in early S (c),
lymphocyte in G0 (d), lymphoblastoid cell in G0 (e), and
postmitotic monocyte (f). Note
predominantly peripheral location of the kinetochore signals
(green) in counterstained (red)
nuclei. Bars represent 5 μm. g
Quantitative 3D evaluation of
radial kinetochore distribution in
nuclei of lymphocytes in G0 and
at different cell cycle stages. n
number of evaluated nuclei.
Error bars indicate standard
deviation of the mean for each
shell
bottom side was higher than on the top side, and all
differences were significant (0.43/0.29, p<0.001 for G1;
0.45/0.33, p<0.006 for S; 0.42/0.31, p<0.007 for G2). In
G0 fibroblasts, the difference was smaller, but still
significant (0.46/0.39, p<0.005). Surprisingly, in SH-EP
cells the proportions of kinetochores at the top and bottom
sides of the nucleus were equal (0.31/0.32 for G1;
0.33/0.35 for S; 0.36/0.40 for G2).
Another aspect of this problem is the distribution of
kinetochores in individual nuclei. Images and kinetochore
counts clearly showed that in some nuclei the majority of
kinetochores were located on the bottom side, while in
other nuclei most kinetochores were gathered at the top
side. To address this matter, we made scatter diagrams as
shown in Figs. 6a, 7a,b,c. Each dot on these diagrams
represents an individual nucleus. The coordinates of a dot
show the proportions of kinetochores situated at top
(abscissa) and bottom (ordinate) sides of the nucleus (the
sum of abscissa and ordinate is less than 1, because there
are some internal and a few lateral kinetochores). If
kinetochores had been distributed evenly between the top
and bottom sides, the dots would have formed one cluster
on these scatter diagrams. However, at least in some cases
two clear clusters with more kinetochores at the top or
bottom sides, respectively, were observed (Fig. 7a,b).
Simple histograms do not allow these two clusters to be
demonstrated because their projections on the x-axis
overlap (Fig. 6a,b). To cope with this difficulty, regression
equations for proportions of kinetochores at the top and
bottom sides (Pt, Pb) were calculated and used as a new x-
416
Fig. 5 Changes in the percentage of peripheral (light gray)
and internal kinetochore signals
(dark gray) depending on the
cell cycle stage in nuclei of
fibroblasts, neuroblastoma SHEP cells, lymphocytes, and
lymphoblastoid cells. Numbers
in parentheses are the numbers
of nuclei studied
axis (R-axis). The zero of the R-axis was always set to the
point Pt=Pb. This allowed minimization of overlapping of
clusters, and observation of them as two maxima of a
bimodal distribution (Fig. 6c). Positive and negative
values show predominant localization of kinetochore
signals in a nucleus at the top and bottom sides,
respectively. For example, a nucleus with 59% kinetochore
signals adjacent to the bottom surface and 11% of
kinetochore signals adjacent to the top surface (Fig. 6a,
arrow) has a value of −0.33 on the histogram on Fig. 6c.
Figure 7a–j shows the distribution of kinetochores in
both cell types studied at different stages of the cell cycle
using the same histograms as in Fig. 6a,c, respectively.
Figure 7d,e shows a clearly bimodal distribution for G1
and S phase SH-EP cells with the minimum not far from
zero. This means that nuclei with similar proportions of
kinetochores at the top and bottom sides are the least
frequent, although one would expect them to prevail.
Instead, there are two groups of cells: (1) with the majority
of kinetochores gathered at the bottom side and (2) with
the majority of kinetochores at the top side. On the
histograms for G1 phase and S phase fibroblasts (Fig. 7g,
h), a similar bimodality is seen, although it is less obvious.
The left peak is much higher than the right one, which
means that the majority of cells have most peripheral
kinetochores on the bottom side of the nucleus (correspondingly, there are more bottom centromeres, than top
centromeres; see above). In G0 fibroblasts (Fig. 7j), we
observed essentially one group of cells with a moderate
prevalence of kinetochore signals localized at the nuclear
bottom compared with the nuclear top. The distribution for
G2 fibroblasts (Fig. 7i) had an intermediate character
between those for G1-S and G0. In G2 SH-EP cells the
bimodality of the kinetochore signal distribution was also
much less pronounced, than at G1 and S phase (Fig. 7f).
This means that the proportion of cells with kinetochores
more or less equally divided between the top and bottom
surfaces increased notably in G2 and G0 (Fig. 7k–n).
Correspondingly, the standard deviation of the distributions described above decreased from G1 and S to G2 and
G0 (Fig. 7o).
The kinetochore provides the core structure of the
centromere to which microtubules attach during mitosis;
however, it is only a small part of the whole human
centromere, which comprises a relatively large (1–5%)
chromosome region consisting of α-satellite DNA (Choo
1997). To trace the behavior of centromeres as a whole
during the cell cycle, we performed 3D-FISH with a
pancentromeric α-satellite probe in nuclei of normal
diploid fibroblasts in G0, early G1, and S phase. Compatibility of FISH and BrdU/pKi67 detection (Bridger and
Lichter 1999; Solovei et al. 2002a) allowed us to
determine the stage of the cell cycle of individual cells.
The shape of the centromere signals was strikingly
different between cycling and non-cycling cells. Early
G1 centromeres were mostly spherical compact bodies;
417
Fig. 6a–c Approach to the analysis of distribution of kinetochores
in individual nuclei. a A scatter diagram showing individual nuclei
as dots with coordinates corresponding to proportions of signals at
the top (Pt) and bottom (Pb) sides of the nucleus (SH-EP cells,
S phase). Note that dots form two clusters. b A histogram showing
number of nuclei (n) depending on Pt. Such straightforward
histograms do not allow demonstratation of the presence of two
clusters because their projection on the abscissa overlaps. c A
regression line is used as a new abscissa (R-axis); zero is set to the
point Pt=Pb. Nuclei with prevalence of kinetochores on the bottom
side (e.g. the one marked by an arrow) have negative values on the
R-axis and vice versa. The histogram on c is clearly bimodal: nuclei
with similar proportions of kinetochores at the top and bottom sides
are least common
S phase centromeres looked similar but were notably
larger than those in G1 (Fig. 8b,c,e,f). Centromere signals
in G0 fibroblasts had elongated and highly irregular
shapes (Fig. 8h,i).
Kinetochore distribution and dynamics in spherical
nuclei of cells growing in suspension
In contrast to the flat shapes of fibroblast and SH-EP cell
nuclei, lymphocytes and lymphoblastoid cells possess
nuclei with a spherical shape (Fig. 4d,e). Kinetochore
signals were scored on RGB galleries of serial optical
sections, as well as on XZ/YZ sections through 3Dreconstructed nuclei using Amira 2.3. Positions of kinetochore signals within the nucleus were classified as
peripheral or internal according to the scheme in Fig. 1a.
As in the case of flat cells, the distribution of
kinetochores is clearly not random. The diameter of nuclei
in lymphocytes and lymphoblastoid cells is 7–11 μm
depending on the cell cycle stage. The kinetochore signal
was 0.1–0.3 μm in size. In the case of random distribution,
only 13–20% of kinetochores would be situated in the
outer 0.5 μm thick shell and would be classified as
peripheral. In reality, at least ca. 40% of kinetochores
occupy a peripheral position (Fig. 5). Scores proved to be
very similar for early, mid, and late S phase, and therefore
these data were summarized for the whole S phase
(Fig. 5). In G0 lymphocytes about 90% of the kinetochores had a peripheral location. In early G1, the
frequency of kinetochores with internal localization
reached 50% (Fig. 5). These kinetochores were distributed
throughout the whole nuclear volume, often being in
contact with pKi67-positive granules (Fig. 2b) represented
by small forming nucleoli and heterochromatin granules.
In S phase, about 70% of signals were found peripherally,
while about 30% were internally located and predominantly adjacent to the nucleoli (identified by TO-PRO-3
counterstain or anti-Ki67 immunostaining). In late G2, the
proportion of internal kinetochores was about 50% as in
G1 nuclei. Interestingly, the majority of internal signals at
this cell cycle stage were located adjacent to the nucleoli
(Fig. 2c). The intranuclear distribution of kinetochores in
cycling and non-cycling lymphoblastoid cells was similar
to those described above for the lymphocytes (Fig. 5). The
proportion of internal signals was slightly higher but their
localization with regard to pKi67 immunofluorescence
followed the same pattern.
In addition to the visual examination, the radial 3D
positioning of kinetochores at different cell cycle stages
and in non-cycling lymphocytes was evaluated using the
3D-RRD computer program (see Materials and methods).
Figure 4g shows the radial distribution of fluorescence
signal intensities as a function of the relative nuclear
radius for G0, early S, mid-S, late S, late G2, and early G1
lymphocytes. The radial intensity distribution in G0 cell
nuclei shows a single, relatively narrow peak with a modal
value at about 85% of the relative nuclear radius and thus
confirms the mostly peripheral location of kinetochore
signals. All curves for cycling cells also show a major
peak at about 85%, but in addition a “plateau” at a relative
radius of ca. 45–65%. This small second peak apparently
represents the internal signals associated with nucleoli.
The fraction of internally located signals increases in this
series from S early → S mid → S late → late G2 → early
G1. The curve for early G1 nuclei is rather similar to that
for the DNA counterstain, reflecting the particularly
pronounced variability of kinetochore locations at this
cell cycle stage.
Monocytes from peripheral blood
One of our blood samples was taken from a person with
acute inflammatory disease. This sample was strongly
enriched in monocytes. Monocytes are the largest of the
418
Fig. 7a–r Relative numbers of
kinetochores at the top and
bottom surfaces of individual
nuclei in fibroblasts and SH-EP
cells at different stages of the
cell cycle. a–c Scatter diagrams
representing individual nuclei as
dots with coordinates corresponding to the proportion of
kinetochore signals at the top
(abscissa) and bottom (ordinate)
surfaces of the nucleus (c.f.
Fig. 6a). d–j Histograms showing number of nuclei (d–f SHEP cells, g–j fibroblasts) with
regard to preferential top or
preferential bottom location of
kinetochores. The ordinate
shows the number of nuclei, the
abscissa the R-axis (c.f. Fig. 6c).
k–n Partial projections of XZ
ortho-sections through characteristic fibroblast nuclei at the
stages corresponding to those
for histograms g–j. k, l Nuclei
with predominantly bottom signals; m, n nuclei with more or
less equally abundant signals at
the top and bottom surfaces. o
Standard deviations of distributions shown in d–j. p Successive cell cycle stages of SH-EP
cells (projections of confocal
stacks) after kinetochore immunostaining (green) and chromatin counterstain (red). Note
concentration of kinetochores at
one pole of anaphase rosettes,
which is inherited by the following telophase. r Projections
of XZ ortho-sections through
the nuclei of early G1 SH-EP
cells showing preferential top (t)
and preferential bottom (b) localization of kinetochores. Bars
represent 5 μm
circulating leukocytes and possess a characteristic kidneyshaped nucleus (Fig. 4f). Anti-pKi67 staining showed that
all monocytes were in G0. Immunostaining of CENP-B/
CENP-C revealed on average 27 kinetochore signals per
nucleus (n=14) with strict peripheral intranuclear positioning (90% of peripheral signals). The few internally located
signals (ca. three per nucleus) were all adjacent to the two
nucleoli found in these cells.
Discussion
Kinetochore arrangements were studied in four diploid
human cell types (diploid fibroblasts, lymphocytes,
lymphoblastoid cells, monocytes) and in a tumor cell
line (SH-EP N14 neuroblastoma cells).
Clustering of kinetochores
Our data demonstrate that clustering of kinetochore
regions in all cell types changes with the cell cycle
stage. Very little or no clustering was observed immediately after mitosis (early G1) as indicated by signal
numbers close to the expected chromosome number. The
number of kinetochore signals decreased in late G1/S and
even more so upon exit from the cell cycle (G0). This
change could be explained either as a result of kinetochore
antigen masking or as the result of kinetochore clustering.
Several arguments support the latter conclusion. Signals in
late G1/S and G0 were generally larger and more intense
than in early G1. Furthermore, the average number of 13
kinetochore signals in G0 lymphocytes observed in the
present study corresponds to the number of 11 and 13
419
Fig. 8a–i Shape of fibroblast
nuclei and centromere signals in
G1 stage (a–c), mid-S stage (d–
f), and in G0 (g–i). a, d, g
Volume rendering of the nuclei
(red) and centromere signals
(green); b, e, h maximum intensity projections of image
stacks; c, f, i reconstruction of
centromere signals by surface
rendering. Note elongated irregular shape of the centromere
signals in G0 cells in contrast to
their more or less spherical
shape in cycling cells. Bars
represent 10 μm
centromere signals reported in two FISH studies performed with a pancentromeric α-satellite probe (Alcobia et
al. 2000; Weierich et al. 2003, respectively). Notably,
kinetochore clustering during S phase was less pronounced in flat nuclei (fibroblasts, SH-EP cells) than in
spherical nuclei (lymphocytes, lymphoblastoid cells). The
reason for this difference is unknown, but topological
constraints depending on the different relationships
between nuclear surface and nuclear volume in flat and
spherical nuclei may be considered. Strong centromere
clustering was observed in particular in terminally
differentiated cells, such as monocytes (present study)
and neurons (Manuelidis 1984; Martou and De Boni 2000;
Solovei et al. 2004a).
The possible mechanisms and reasons for centromere
clustering remain unclear. Chromocenters formed by
clustered centromeres are typical of many cell types
(Haaf and Schmid 1991; Manuelidis 1984). Chromocenters consist mainly of highly repetitive heterochromatin
(pericentric hetrochromatin) and are known to have certain
epigenetic “marks”, such as DNA methylation, histone H3
methylation at lysine 9 (H3-K9), and enrichment in
heterochromatin protein-1 (HP1) (reviewed in Bird 2002;
Richards and Elgin 2002; Lachner and Jenuwein 2002;
Lachner et al. 2003; Maison and Almouzni 2004). One of
the putative functions of chromocenters is linked to
regulation of transcriptional activity by silencing genes
situated in the vicinity of a chromocenter (Brown et al.
1997, 1999; Baxter et al. 2002). Therefore, the clustering
of centromeres that takes place during G1 and early S
probably manifests the establishment of epigenetically
controlled “silencing” zones in the nucleus. This assumption corresponds to the fact that centromere clustering is
especially strong in quiescent and terminally differentiated
cells.
Kinetochore movements in cycling cells and upon exit
from the cell cycle
The present study demonstrates for the first time that
kinetochore topology in all investigated cell types differs
markedly in G0 and G1 cells and changes from early to
late G1, as well as from early to mid-late S and G2. In
quiescent cells (G0), most kinetochores showed a peripheral localization (80–95% depending on the cell type),
while the few internally located kinetochores were found
mostly adjacent to the nucleoli. In all cell types, except for
monocytes, which were only studied in a terminally
differentiated state, we found distinct changes of kinetochore arrangements during interphase. In early G1, a
considerable fraction of kinetochores (about 30–60%
depending on cell type) were located in the nuclear
interior, clearly remote from the nuclear envelope. From
early to late G1, the internal fraction of signals decreased
(to about 20–50%) with the result that most kinetochores
420
now attained a peripheral localization (about 50–80%). In
early S phase the fraction of peripherally located
kinetochores became even higher (about 50–90%). In
fibroblasts and SH-EP cells, the fraction of internal
kinetochore signals started to increase again (about 20–
25%) from mid to late S phase, while it remained roughly
constant in lymphocytes and lymphoblastoid cells. During
G2, the internal and peripheral kinetochore fractions did
not change further in fibroblasts and SH-EP cells, while
the internal fraction increased significantly in lymphocytes
and lymphoblastoid cells. Fibroblasts and PHA-stimulated
lymphocytes showed a higher frequency of peripherally
located kinetochores compared with lymphoblastoid cells
and neuroblastoma cells; also the fraction of centromeres
that switched from a peripheral to an internal localization
during mid-S/G2 was smaller in the latter two cell types.
In conclusion, our data confirm cell type and cell cycle
dependent differences in kinetochore arrangements. Possible functional implications and mechanisms involved in
the dynamic centromere patterns have not yet been
elucidated.
Changes in kinetochore arrangements during early G1
are consistent with a live cell imaging study performed in
HeLa cells (Walter et al. 2003). This study indicated
considerably higher chromatin mobility during early G1
compared with subsequent interphase stages, from mid G1
to late G2, when movements of chromosome territories/
chromatin domains were strongly restricted. Furthermore,
replication foci during S phase were found to be rather
immobile except for some constrained Brownian movements (Leonhardt et al. 2000). These findings suggest that
centromere regions may undergo more pronounced movements from mid G1 to late G2 than other chromosomal
subregions. We do not know, however, whether all
centromeres may occasionally perform large-scale movements (i.e., movements over a range of several micrometers), or whether the capability for such movements is
restricted to the centromeres of certain chromosome
territories.
Results from previous studies are in partial or complete
disagreement with our present findings. Bartholdi (1991)
investigated 3D centromere arrangements in a growing
population of human fibroblasts, employing laser confocal
scanning microscopy after staining with anti-kinetochore
antibodies. In this study cell cycle staging was based on
fluorescence measurements of propidium iodide in situ.
For G1 nuclei, Bartholdi reported that many centromeres
were located in association with nucleoli or fused in
chromocenters that expanded from nuclear top to nuclear
bottom. No specific association of kinetochores was
observed with the nuclear envelope. During S phase,
chromocenters dispersed often forming patterns of rings or
lines. In contrast, our present study revealed that most
centromeres were located at the 3D nuclear periphery both
in G0 and cycling fibroblasts, while only a minor fraction
of centromeres moved from the nuclear periphery to the
nuclear interior and vice versa.
Weimer et al. (1992) also employed anti-kinetochore
antibodies to study centromere arrangements in non-
stimulated (G0) and stimulated human lymphocytes (G1,
S, G2). They performed a 2D evaluation based on
conventional epifluorescence microscopy and found a
distinct tendency for a peripheral position of centromeres
during G0 and G1, which became weaker during S phase.
While we agree with the general conclusions of Weimer et
al. concerning the dynamics of centromere topology
during interphase, the difference in kinetochore topology,
in particular between G0 and early G1 nuclei demonstrated
in the present study, escaped their notice. This may be due
to a less precise staging of cells. Weimer et al. (1992)
synchronized stimulated lymphocytes by a thymidine
block and then fixed cells at different times after release
from the block. The time of harvest was used for a rough
discrimination of G0, G1, S and G2, as well as G1 and
early S of the subsequent cell cycle. In an attempt to
confirm the cycle stage of the cells studied in situ after
anti-kinetochore antibody staining, a fraction of cells was
stained in parallel with Hoechst 33258 and subjected to
flow cytometry. In our present study we took great care to
establish a protocol that enabled us to discriminate
unequivocally between G0 and early and late substages
of G1, as well as early and late S and G2 phase at the level
of single cells in situ.
Ferguson and Ward (1992) and Vourc’h et al. (1993)
performed FISH and confocal microscopy experiments
with stimulated human and mouse lymphocytes, respectively. For cell cycle staging, cells were flow sorted after
DNA/RNA staining with propidium iodide. Probes for
major and minor satellite DNA were used for murine cells.
Ten chromosome-specific centromere probes were employed for human cells but detailed data were only
presented for centromeres of chromosomes 1, 7, 11, and
17. Consistent with our present findings, the authors
observed a centromere repositioning from the nuclear
periphery to the nuclear interior during S/G2. For G1
lymphocyte nuclei they state that most centromeres were
located in the nuclear periphery, although they were not
able to discriminate cells in G1 from cells in G0. In the
present study we found that about 90% of centromeres
were located in the periphery of human lymphocyte nuclei
in G0, while about half of centromeres were located in the
periphery and half in the nuclear interior, including
adjacent to the nucleoli, of lymphocyte nuclei in early G1.
Hulspas et al. (1994) carried out FISH with a human
chromosome 11 centromere-specific probe in both nonstimulated lymphocytes harvested directly from peripheral
blood and in cultured, PHA-stimulated human lymphocytes. Laser confocal microscopy using the nuclear center
as a reference point revealed that the distribution of the
chromosome 11 centromeres appeared to be random
during the G0 stage, while most, if not all, centromeres
were found at the nuclear periphery in G1. The peripheral
topology of kinetochores described in the present study for
G0 lymphocytes strongly differs from the apparently
random topology described by Hulspas et al. for chromosome 11 centromeres. This result needs to be confirmed by
independent experiments in order to demonstrate that
chromosome 11 centromere topology is an exceptional
421
case. It should be noted that PHA stimulates only
T lymphocytes and, accordingly, non-cycling nuclei from
B lymphocytes and T lymphocytes were assessed together
with a fraction of stimulated T cells. In the present study
we were able to distinguish cycling T cells unequivocally
from non-cycling T cells and B cells.
Asymmetry of kinetochore distribution in nuclei of
adherently growing cycling cells may reflect the
behavior of the preceding anaphase rosettes
More than a century ago Carl Rabl described a polar
orientation of chromosomes in the interphase nuclei of
Salamandra maculata with centromeres and telomeres
positioned at opposite sides (Polfeld and Gegen-Polfeld)
of the nucleus (Rabl 1885). According to Rabl, this polar
orientation results from anaphase chromosome movements
(see for example, Abranches et al. 1998). In mammalian
cells such a Rabl orientation is rarely observed (Weierich
et al. 2003; reviewed in Parada and Mistelli 2002) likely
due to additional chromatin movements that occur during
telophase/early G1 (Walter et al. 2003). In the present
study we observed an asymmetric distribution of kinetochores in the rather flat nuclei of adherently growing
fibroblast and SH-EP cells. Kinetochores were located
predominantly at the nuclear bottom in fibroblast nuclei,
while in SH-EP cells the fraction of nuclei with
centromeres predominantly located at the nuclear top
was similar to the fraction of nuclei with centromeres
preferentially located at the bottom (Fig. 7).
For an explanation of this obvious nuclear asymmetry
of centromere arrangements we propose the following
hypothesis, which we call the “fallen rosette” scenario
(Fig. 9). In fibroblasts both the metaphase plate and the
two resulting anaphase rosettes are arranged perpendicular
to the substratum, while the spindle is arranged parallel to
the substratum (our unpublished observations). During
anaphase the centromeres are located on the external side
of the chromatin mass facing their respective centriole
(Fig. 7p for SH-EP cells; see also Habermann et al. 2001,
Fig. 5e for human fibroblasts). At the end of anaphase the
two anaphase rosettes fall over, resulting in a parallel
arrangement with respect to the growth surface. The
preferential location of centromeres at the bottom side of
G1 fibroblast nuclei suggests that each anaphase rosette
should fall over preferentially with their centromeres
downwards. Correspondingly, the two anaphase rosettes
should usually fall over in opposite directions yielding
symmetrical arrangements of chromosome territories in
the two daughter nuclei. We noted that the spindles in
anaphase rosettes of SH-EP cells were located at various
angles to the growth surface (our unpublished observations) and the direction to which a given anaphase rosette
falls over should depend on this angle. Since the angle is
the same for both anaphase rosettes of a given cell, while
their centromeres are located at opposite sites of the two
rosettes, we should expect that one anaphase rosette falls
over with the centromeres facing the substratum, while the
Fig. 9 Changes in the localization of kinetochores in cell cycle: the
“fallen rosettes” scenario. At the late anaphase stage kinetochores
are concentrated on one side of the rosette. The rosette falls on one
side, as a result of which nuclei with kinetochores gathered at the top
side (left) or at the bottom side (right) arise. Judging from the
relative amounts of top and bottom kinetochores in the two cell
types studied, the bottom (right) variant strongly prevails in
fibroblasts, while in SH-EP cells both variants are equally probable.
After late G1, certain rearrangements of kinetochores take place
owing to which the numbers of kinetochores at the top and bottom
sides become more even in G2 or G0
other falls over with an upward centromere positioning. In
G0 and G2 fibroblasts most nuclei still showed their
centromeres preferentially at the nuclear bottom, but the
proportion of nuclei with kinetochores at the nuclear top
was higher compared with nuclei in G1 and S phase.
Similarly, in G2 SH-EP cells we observed a decreased
fraction of nuclei with centromeres predominantly concentrated at either the nuclear top or bottom compared
with G1 and S cells, while the fraction of nuclei showing
similar numbers of centromeres in top and bottom
positions increased. By and large, the further cells had
departed from mitosis, the more uniformly their centromeres were distributed at the lower and upper nuclear
periphery (Fig. 9). To explain these changes we must
invoke some additional centromere migration during
interphase and in postmitotic cells (Manuelidis 1985;
Martou and De Boni 2000; Solovei et al. 2004a). The
importance of the fallen rosettes mechanism for determination of the nuclear chromosomal arrangement has earlier
been suggested by us based on the data on relative
422
positions of small and large chromosomes in interphase
nuclei (Habermann et al. 2001). The fact that data of a
very different kind, the distribution of centromeres, also
correspond to the fallen rosettes scenario strongly supports
this hypothesis.
The dynamics and highly reproducible changes of
centromere topology during the cell cycle, upon exit from
the cell cycle and during terminal differentiation cannot be
explained as a result of Brownian movements. The
question as to whether these changes play a role in
epigenetic gene regulation and require energy dependent
mechanism(s) is a matter for future studies.
Acknowledgements We are grateful to W.C. Earnshaw (University of Edinburgh) for the generous gift of anti-CENP-B and antiCENP-C antibodies. B. Joffe (Technical University of Munich) has
suggested and greatly helped with the analysis of the asymmetrical
distribution of peripheral kinetochores in individual cells. This work
was supported by grants from the Deutsche Forschungsgemeinschaft
to T. Cremer (Cr 59/20, 1–3).
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