Chromatin domains and the interchromatin compartment form structurally deﬁned … ·...

Chromatin domains and the interchromatin compartment form structurally

defined and functionally interacting nuclear networks

Heiner Albiez1, Marion Cremer1, Cinzia Tiberi2, Lorella Vecchio2, Lothar Schermelleh1, Sandra Dittrich1,

Katrin Kupper1, Boris Joffe1, Tobias Thormeyer1, Johann von Hase3, Siwei Yang4, Karl Rohr4,

Heinrich Leonhardt1, Irina Solovei1, Christoph Cremer3, Stanislav Fakan2 & Thomas Cremer1*1Department of Biology II, LMU Biozentrum, Großhaderner Straße 2, 82152 Planegg-Martinsried, Germany;Tel: +49-89-218074329; Fax: +49-89-218074331; E-mail: [email protected]; 2Centre ofElectron Microscopy, University of Lausanne, 1005 Lausanne, Switzerland; 3Kirchhoff Institute of Physics andBiophysics of Genome Structure, IPMB, University of Heidelberg, 69120 Heidelberg, Germany; 4Department ofBioinformatics and Functional Genomics, IPMB, University of Heidelberg and DKFZ Heidelberg,69120 Heidelberg, Germany* Correspondence

Received 10 July 2006. Received in revised form and accepted for publication by Hans Lipps 4 August 2006

Key words: chromatin condensation, chromosome territories, nuclear architecture, interchromatin compartment

Abstract

In spite of strong evidence that the nucleus is a highly organized organelle, a consensus on basic principles of the

global nuclear architecture has not so far been achieved. The chromosome territoryYinterchromatin compartment

(CT-IC) model postulates an IC which expands between chromatin domains both in the interior and the periphery

of CT. Other models, however, dispute the existence of the IC and claim that numerous chromatin loops expand

between and within CTs. The present study was undertaken to resolve these conflicting views. (1) We demonstrate

that most chromatin exists in the form of higher-order chromatin domains with a compaction level at least 10 times

above the level of extended 30 nm chromatin fibers. A similar compaction level was obtained in a detailed analysis

of a particularly gene-dense chromosome region on HSA 11, which often expanded from its CT as a finger-like

chromatin protrusion. (2) We further applied an approach which allows the experimental manipulation of both

chromatin condensation and the width of IC channels in a fully reversible manner. These experiments, together

with electron microscopic observations, demonstrate the existence of the IC as a dynamic, structurally distinct

nuclear compartment, which is functionally linked with the chromatin compartment.

Introduction

Chromosomes in both animal and plant cell nuclei

are organized in territories, but the internal structure

of chromosome territories (CTs) and their interac-

tions with neighboring CTs has remained a matter of

controversy (reviewed in Cremer et al. 2006). The

present plurality of models reflects the complexity of

nuclear architecture and highlights the still-unresolved

role that this architecture may play in epigenetic

gene regulation. Initially, it was proposed that CTs

are objects with a rather smooth surface, separated

by an interchromosome domain (ICD) which con-

tains nuclear speckles and bodies (Zirbel et al. 1993).

Further studies of CTs and their substructure using

state-of-the-art light microscopy, combined with

results obtained by electron microscopy, led to the

formulation of the CT-IC model (IC = interchromatin

Chromosome Research (2006) 14:707–733# Springer 2006

DOI: 10.1007/s10577-006-1086-x

compartment) (for reviews see Cremer & Cremer

2001, Cremer et al. 2000, 2006, Williams 2003). This

model states that CTs are built up from higher-order

chromatin structures of several hundred kb to several

Mb, called ~ 1 Mb chromatin domains. The notation

F~ 1 Mb_ refers to the order of DNA content, which

may range for individual replication foci from a few

hundred kb to several Mb (Jackson & Pombo 1998,

Berezney et al. 2000, Koberna et al. 2005). Originally

these domains were detected in light microscopic

studies as replication foci, when cells during S-phase

were pulse-labeled with thymidine analogs (Nakamura

et al. 1986, Ma et al. 1998). It was found that foci,

labeled during S-phase, can be detected as higher-

order chromatin structures at any stage of interphase,

including subsequent cell cycles (for review see

Berezney et al. 2000). The internal structure of

replication foci or ~ 1 Mb chromatin domains is still

unknown, although we think it possible that they are

built up from a series of ~ 100 kb chromatin loop

domains, each comprising DNA in the order of

50Y200 kb. Both individual ~ 100 kb domains and

entire ~ 1 Mb chromatin domains may adopt a

different configuration depending on the transcrip-

tional status of their genes. However, direct evidence

for a strict causal connection between chromatin

compaction, transcription and silencing of individual

genes in living cells has not so far been provided.

According to the CT-IC model the IC consists of a

contiguous three-dimensional (3D) network of channels

and lacunas starting at nuclear pores and permeating the

nuclear interior as a functionally indispensable nuclear

compartment. It expands between CTs, but also pen-

etrates into the CT interior with its finest branches

(G100 nm) separating ~ 100 kb chromatin domains. In

transcriptionally inactive zones the IC channels can

become very narrow, but never collapse entirely

(Cremer et al. 2000). The IC contains nuclear speckles

and bodies, such as Cajal (coiled) bodies, PML bodies

or Rad51 foci. Nuclear speckles and bodies are

dynamic structures formed through transient associa-

tion of proteins which roam the nuclear space (Misteli

et al. 1997, Phair & Misteli 2000, Tashiro et al. 2000,

Pederson 2002). The IC is separated from the interior

of compact higher-order chromatin domains by the

perichromatin region, which was structurally defined

by EM studies as a narrow border zone of decondensed

chromatin at the surface of higher-order chromatin

domains. It represents a functionally important nuclear

compartment, where DNA and RNA synthesis, as well

as co-transcriptional splicing take place (for reviews see

Fakan 2004a,b).

The random walk/giant loop model predicts that

CTs are built up from chromatin loops in the low Mb

range (Sachs et al. 1995). Several groups presented

evidence to show that giant loops with a length of

several microns expand from compact CT cores and

that this extensive looping-out results in zones of

intermingling chromatin fibers between neighboring

CTs (for review see Foster & Bridger 2005). For

example, Bickmore and colleagues (Mahy et al.2002) reported that a locus-specific FISH signal cor-

responding to a gene dense segment on HSA 11p15.5

was often located about 1Y2 2m away from the

border of the painted HSA 11 CT core and postulated

that Fthe organization of chromosomes within the

nucleus is probably somewhere in between the com-

plete decondensation of chromatin fibers like spa-

ghetti on a plate suggested 30 years ago and the

model of a discrete territorial organization._Although the chromatin region between the spe-

cific signal and the CT was not visualized in these

experiments, it was proposed that this unstained region

typically forms a largely extended chromatin thread

meandering in the neighborhood of the corresponding

CT (Chubb & Bickmore 2003). Recently, Branco and

Pombo described extensive intermingling of chroma-

tin loops both within a given CT and between

neighboring CTs, and proposed the interchromosomal

network (ICN) model of chromatin organization

(Branco & Pombo 2006). These observations and

their interpretation by the cited groups have chal-

lenged the two major postulates of the CT-IC model,

namely that CTs are built up from ~ 1 Mb chromatin

domains and that a mostly DNA-free IC space

expands between them.

The present study was undertaken with the aim of

rigorously testing conflicting postulations of current

models. Our results argue against the contribution of

a large fraction of giant loops to higher-order

chromatin arrangements. In support of the CT-IC

model we demonstrate that a dynamic network of IC

channels and lacunas exists, and describe their

topographical relationships with nuclear speckles

and bodies, nascent DNA and nascent RNA, as well

as with RNA polymerase II. The width of the IC

could be strongly and fully reversibly manipulated in

nuclei of living cells under conditions which induced

a transient shift from normally condensed chromatin

(NCC) to hyper-condensed chromatin (HCC). Global

708 H. Albiez et al.

higher-order chromatin arrangements remained sta-

ble in nuclei of single living cells, which were

subjected to several NCCYHCCYNCC cycles, and

cells continued to proliferate and divide.

Material and methods

Cell culture

Early passages of primary human fibroblasts (46,XX)

(provided by T. Meitinger, Technical University

Munich), CHO cells (provided by J. Broers, Univer-

sity of Maastricht, The Netherlands) and the neuro-

blastoma cells LAN and Kelly (Solovei et al. 2000)

were grown in DMEM supplemented with 10% FCS,

penicillin (100 I.E.) and streptomycin (100 2g/ml) at

37-C in an atmosphere of 5% CO2. HeLa cells stably

transfected with histone H2B-GFP (Kanda et al.1998), MCF-7 cells (provided by P. Meltzer, NIH,

Bethesda, USA), amniotic fluid cells (provided by M.

Speicher, Technical University of Munich) and periph-

eral blood lymphocytes were grown in RPMI 1640

medium with the same supplements. Cells were grown

on coverslips to 50Y70% confluence before use.

ATP-depletion experiments

ATP-depletion was achieved by incubating cells for

30 min in a glucose-free culture medium containing

10 mM sodium azide and 6 mM 2-deoxyglucose

(Dingwall et al. 1987, Phair & Misteli 2000).

Formation of hypercondensed chromatin (HCC)

HCC formation was induced by incubating the cells

in a hyper-osmolar medium at osmolarities above

380 mOsm. A maximum effect was noted for osmo-

larities from 570 to 750 mOsm. If not stated

otherwise, this range was used in the present experi-

ments. Osmolarities were measured with an osmom-

eter (Osmomat 030, Gonotec, Germany). As a

standard protocol, 1 ml 20 � PBS (2.8 M NaCl,

54 mM KCl, 130 mM Na2HPO4, 30 mM KH2PO4 in

H2O, pH adjusted with HCl to 7.4) was diluted with

19 ml standard culturing medium (290 mOsm) to yield

an osmolarity of 570 mOsm. To reverse the effect the

cells were again incubated in their physiological

medium (290 mOsm). Hypo-osmolar medium was

obtained by diluting standard culture medium with

ddH2O in a ratio of 1:1 (~ 140 mOsm). Since

chromatin condensation and decondensation processes

started within seconds, washing steps in physiological

salt solutions, such as 1 � PBS (290 mOsm) were

strictly avoided prior to the fixation of cells.

Replication labeling

Replication foci were pulse-labeled in S-phase cells

either with bromodeoxyuridine (BrdU) or with

cyanin-3-deoxyuridine triphosphate (Cy3-dUTP)

(Ferreira et al. 1997, Zink et al. 1998). BrdU incor-

poration was achieved by incubating cells for 10 min

in 20 2M BrdU (nascent DNA) or up to 7 h in 50 2M

BrdU for extensive DNA labeling. Following incor-

poration of Cy3-dUTP, cells were grown for 6 days,

allowing the completion of three or more subsequent

cell cycles. This approach resulted in the random

segregation of labeled and unlabeled chromatids and

the generation of nuclei with clusters of fluorescent

foci, also termed ~ 1 Mb chromatin domains

(Cremer & Cremer 2001), representing labeled CTs

(Schermelleh et al. 2001).

Labeling of nascent RNA

Scratch-labeling of nascent RNA transcripts in living

cells was performed with bromouridine triphosphate

(BrUTP, 5 mM, Sigma-Aldrich, Germany). The spec-

ificity was either confirmed by labeling in presence of

the inhibitor !-Amanitin (2 2g/ml, Calbiochem,

Germany) or by RNase treatment after fixation

(0.02%, Roche, Germany), which completely removed

any labeling within the nucleus (data not shown).

Permeabilization of living cells

Permeabilization of the cell membrane of living cells

was performed by incubation in medium containing

40 2g/ml digitonin for 2 min. This treatment resulted

in the permeabilization of 95% of cells and did not

lead to an apparent change of nuclear chromatin

structure at the light microscopic level.

Cell fixation and immunofluorescence staining

Cells were fixed with 4% formaldehyde in 1� PBS for

10 min and washed several times with 1 � PBS

containing 0.01% Tween. For immunostaining the cells

Chromatin domains and the interchromatin compartment 709

were permeabilized with 0.5% Triton X100 in PBS for

5 min. Cells were incubated for 10 min in 0.1 N HCl

and blocked thereafter in PBS containing 2% BSA and

0.01% Tween for 10 min. DNase was added to the

primary antibody solution to a final concentration of

20 2g/ml and the incubations with primary (45 min)

and secondary antibodies (30 min) were performed in a

humidified chamber at 37-C in blocking solution. The

HCl and DNase treatments were obligate to ensure the

accessibility of HCC bundles, which was confirmed by

successful detection of incorporated BrdU. Primary

antibodies used were: mouse-anti-PML (1:100, St Cruz

Biotech, USA), mouse-anti-SC-35 (1:1000, Sigma-

Aldrich, Germany), rabbit-anti-Rad51 (1:1000, gift of

Satoshi Tashiro, Hiroshima, Japan), mouse-anti-RNA

polymerase II LS (FPOL 3/3_, 1:10; gift of Dirk Eick,

GSF, Munich, Germany) and mouse-anti-BrdU (1:200,

Roche, Germany). Secondary antibodies used were:

Cy3-conjugated donkey-anti-goat (1:500, Rockland,

USA), Alexa 488-conjugated goat-anti-mouse (1:400,

Molecular Probes, USA), Cy3-conjugated sheep-anti-

mouse and Cy3-conjugated goat-anti-rabbit (both 1:500,

Dianova, Germany). Cells were mounted in Vectashield

antifade medium (Vector Laboratories, Canada).

DNA probes and FISH experiments

Human paint probes for chromosomes 7, 8 and 11

(kindly donated by M. Ferguson-Smith and J. Wienberg,

University of Cambridge, UK), were amplified and

Biotin-, Cy3 or FITC labeled by a DOP-PCR using the

6 MW primer as described (Schermelleh et al. 1999).

BAC probes were purchased from BACPAC Resour-

ces Center (Oakland, CA): RP11-240G10, RP11-

326C3, RP13-46H24, RP11-412M16, RP11-496I9,

RP11-1391J7, RP11-1335O1, RP11-371C18, RP13-

25N22, RP11-295K3, RP11-534I22, RP5-998N23,

RP11-889I17, RP5-1075F20, RP11-847E17. Genomic

DNA of single BACs or of pooled BACs was amplified

and labeled either with the haptens DIG-dUTP, DNP-

dUTP or by dUTP-TexasRed by a modified DOP-

PCR using two different primers DOP-2 and DOP-3

as described (Fiegler et al. 2003). Multicolor FISH on

morphologically preserved nuclei, detection of labeled

probes by fluorochrome-conjugated antibodies or

fluorochrome-conjugated avidin was performed

according to protocols described in detail elsewhere

(http://www.epigenome-noe.net/researchtools/proto-

col.php? protid=23). Nuclear DNA was counterstained

with DAPI (0.05 2g/ml, Sigma, Germany) for 5 min.

Epifluorescence and confocal microscopyand live-cell imaging

Epifluorescence images were acquired using a Zeiss

Axiophot 2 microscope equipped with a 100�/1.3 oil

objective and a Coolview CCD camera. For live-cell

observations a perfusable FCS2 live-cell chamber

system (Bioptechs, USA) was used in combination

with an objective heater to provide stable tempera-

ture conditions (Walter et al. 2003). Confocal micros-

copy was performed using either a Zeiss LSM 410

(z-step size 200 nm) or a Leica SP2 (z-step size

120 nm) both equipped with 63�/1.4 plan-apochromat

oil objectives. For bleaching experiments the power of

the Ar-Laser (488 nm, 15 mW) at the Zeiss LSM 410

was set to 100% and stripes were irradiated till

sufficient bleaching was obtained.

Electron microscopy

For electron microscopic investigations the cells grow-

ing on microgrid coverslips (CELLocate, Eppendorf)

fixed with freshly prepared 4% paraformaldehyde for

1 h at 4-C in 0.1 M Soerensen phosphate buffer (pH

7.4) were dehydrated in an increasing ethanol series

and finally embedded in LR White resin which was

allowed to polymerize for 48 h at 60-C. The blocks

with embedded cells were separated from the cover-

slips by a short treatment with liquid nitrogen. Ultrathin

sections (~ 80 nm), were obtained using a Leica

Ultracut UCT ultramicrotome, placed on uncoated gold

grids (400 mesh) and contrasted with osmium ammine

staining solution of 0.2% for 1 h after HCl hydrolysis

(5 N HCl, 20 min at RT) to specifically visualize DNA

by the Feulgen-type method (Cogliati & Gautier

1973). Immunocytochemical double-labeling assays

were carried out using anti-DNA and anti-GFP anti-

bodies followed by secondary colloidal gold markers

of 15 and 10 nm respectively, according to a

previously reported protocol (Cmarko et al. 1999).

Grids were observed with Philips CM12 or CM10

electron microscopes (EM) operated at 80 kV, using a

30Y40 2m objective aperture.

Image processing and 3D image analysis

Prior to 3D image reconstructions (generated with

Amira 3.1, TGS) and a quantitative 3D image anal-

ysis, 3D data stacks of light optical sections were

deconvolved with the Huygens (SVI) maximum-


likelihood-estimation algorithm (quality factor 0.1;

20Y40 iterations; signal-to-noise ratio was set to 30)

using measured point-spread functions. To evaluate

the 3D location of nuclear components in nuclei with

HCC, we applied the ADS program, developed by us,

which allowed absolute 3D distance measurements

from a given signal to the nearest surface. Following

a user-set threshold a surface was generated around

HCC bundles. To support the choice of threshold by

an objective criterion we compared EM sections with

corresponding, deconvolved confocal sections. These

comparisons allowed us to determine thresholds for

confocal sections, which yielded a rather perfect

overlay with the EM section (Figure S13*). There-

after, for each voxel attributed to a segmented signal

(e.g. a PML body) the smallest 3D distance was

measured to this chromatin surface. The plotted

occurrence frequencies of the measured and addi-

tionally intensity-weighted distances give a descrip-

tion of the spatial distribution of the signals with

respect to the nearest chromatin aggregates. The

frequencies were plotted in classes with 200 nm

width. Positive values reflect signals located in the IC

interior, negative values reflect signals in the interior

of HCC bundles. We applied the U-test to compare

the obtained distributions with each other or with a

control distribution. For the control distribution we

performed ADS measurements for voxels filling

uniformly the complete 3D nuclear volume.

To quantify the similarity and dissimilarity,

respectively, between 3D images of cell nuclei

recorded during repeated NCCYHCCYNCC cycles

we developed a histogram-based approach. Given

two 3D images we first segmented the nuclei using a

global thresholding scheme after applying an aniso-

tropic diffusion filter to reduce the noise; Then we

computed the histograms of the two 3D images. Note

that the histograms of the intensities were computed

only for the segmented parts of the images. Therefore,

the intensities of the background were not included

and did not cause a deterioration in the result. In

addition, we normalized the histograms with respect

to the volumes of the nuclei to be invariant to object

scaling. To this end we determined the volume of each

nucleus by counting the number of voxels and

dividing the histogram values accordingly. Conse-

quently, the results did not depend on the size of the

nuclei. Moreover, in order to make our analysis

independent of a global change of the intensities we

normalized the histograms with respect to the mean

intensities. Finally, we computed the sum-of-

squared-differences between the two histograms and

used the mean-squared error (MSE) between the

histograms as similarity/dissimilarity measure for

cell nuclei. For better readability each value was

multiplied by 107.

Results

Giant chromatin loops are presentas finger-like protrusions

For a complete visualization of the gene-dense region

of 11p15.5, which was reported to expand into a giant

loop away from the human CT 11 (Mahy et al. 2002),

we performed 3D FISH with a BAC contig of

2.35 Mb (Figure 1A). When tested individually each

BAC yielded a dot-like FISH signal in the nucleus. In

agreement with the model proposed by Bickmore and

co-workers the signals were typically, though not

consistently, observed at the border or outside of the

painted HSA 11 CTs (Figure 1BYD). In a multicolor

3D FISH experiment we painted the HSA 11 CT and

visualized the two terminal BACs as well as two

BACs located in the middle of the contig as dot-like

signals (Figure 1B). The BAC at the telomeric end of

the contig contains 11 genes, many of them highly

transcribed in human fibroblasts, while the BAC at

the centromeric end contains only one gene with low

transcriptional activity (data not shown). The telo-

meric BAC was typically located further away from

the HSA 11 CT body than the centromeric BAC.

Notably, both BACs revealed dot-like chromatin

structures and we were not able to detect a clear size

difference. The two BACs chosen from the middle of

the contig always revealed a single compact signal

(Figure 1B), although a region of about 350 kb

between these BACs was not covered. According to

the model proposed by Bickmore and colleagues we

would expect that 3D FISH experiments with the

entire BAC contig should result in a dispersed pattern

of BAC signals in the vicinity of the HSA 11 CT core.

To test this prediction we modified this experiment so

*Electronic Supplementary Material

Supplementary material is available in the online version

of this article at http://dx.doi.org/10.1007/s10577-006-

1086-x and is accessible for authorized users.


that the two BACs representing the telomeric and

centromeric ends of the contig were visualized in red

and green as before, while all other BACs in between

were visualized in yellow (Figure 1C). In this exper-

iment we typically observed finger-like chromatin

protrusions expanding from the CT periphery and

having a maximal length of up to 3 2m and a width of

typically several hundred nanometers. Occasionally a

thicker part of the protrusion was connected by a much

thinner fiber segment (Figure 1C, arrow). In other

cases the contig probes revealed rather compact struc-

tures at the CT surface (Figure 1D). In any case, the

compaction rates of ~ 1:300, estimated for this 2.3 Mb

segment, argue against a giant 30 nm fiber (compac-

tion ~ 1:30 to 1:40) meandering in the neighborhood

of the corresponding CT.

Figure 1. Multicolor 3D FISH on a human fibroblast of a 2.34 Mb region on HSA 11p15.5: A: Schematic draft of the 15 BACs used for the

contiguous delineation of this region (with one interruption of 350 kb in the middle). The most telomeric (red), the most centromeric (green)

clone and the intermediate clones (yellow) are labeled in different colors. BYD: Maximum Z projections after 3D-FISH of the CT 11 (blue)

and the 11p15.5 clones as indicated in the schematic draft. B: In addition to the two BACs marked green and red, two BACs marked by an

asterisk in A were visualized together with the CT 11. C,D: Here, the most telomeric (red) and centromeric (green) BAC were visualized

together with all other BACs (yellow). B and C: The stained region forms a finger-like chromatin protrusion with a compaction factor of

~ 1:300 expanding from CT 11. The inset in C outlines the contiguous structure of the full length Fcontig_, delineated by all BACs. The arrow

points to a much thinner fiber segment connecting the thicker parts of the protrusion. D: Here, the stained region presents itself as an even

more condensed structure.


Chromosome territories built up from chromatinwith higher-order compaction

To test whether this conclusion could be extended to

the entire genome, we generated living HeLa cells

harboring a few CTs partially labeled by fluorescent

~ 1 Mb chromatin domains, while the majority of

CTs were unlabeled (see Methods). Figure 2 shows a

typical nucleus with three distinct labeled clusters of

~ 1 Mb chromatin domains. Each area represents one or

possibly several closely adjacent in-vivo-labeled CTs.

Chromatin was simultaneously visualized by expres-

sion of histone H2B-GFP (Figure 2A1). Nuclear

regions between the three intensely labeled areas

revealed evenly distributed signals similar to back-

ground fluorescence outside the nucleus after applica-

tion of a low threshold (Figure 2B1). A slight

accumulation of weak fluorescence signals is seen

in the close vicinity of intensively labeled clusters

of ~ 1 Mb chromatin domains (Figure 2B1), which

disappeared after application of a higher threshold

(Figure 2B2). We also searched in deconvolved light

optical sections for labeled chromatin fibers expand-

ing from fluorescent ~ 1 Mb chromatin domains

within a given CT. After deconvolution, measured

fluorescence intensities in non-labeled areas within

labeled CTs were not higher compared to nuclear

zones occupied by non-labeled CTs (Figure 2C1).

This finding conforms to the assumption that most, if

not all, fluorescence observed in the vicinity of ~ 1 Mb

chromatin domains resulted from out-of-focus light,

and not from looping-out of labeled chromatin fibers.

The maximum intensity projection of all deconvolved

light-optical sections, which comprise all labeled foci

present in this nucleus, reveals a few ~ 1 Mb chro-

matin domains located between the three intensely

labeled nuclear areas (Figure 2C2, arrow).These

domains may be part of higher-order chromatin pro-

trusions expanding from a labeled CT body as des-

cribed above (Figure 1). These findings are typical for

numerous nuclei harboring fluorescently labeled CTs

observed in this and previous studies (Walter et al.2003). Furthermore, we followed living HeLa cells

with a few labeled CTs from one cell cycle to the next

(Figure S2) (Walter et al. 2003). In case of a sizeable

fraction of labeled giant loops expanding from a given

CT, we would expect that the retraction of such loops

during the formation of the corresponding prometa-

phase chromosome would result in a much smaller

and accordingly more intensely labeled halo around

the condensing CT. The reverse event should take

place, when mitotic chromosomes form CTs during

the telophase/early G1 transition. However, besides a

modest shrinkage (Figure S2C: G2Yprometaphase)

and swelling (Figure S2C: prometaphaseYG1) of the

observed CTs this expectation could be ruled out.

In summary, present evidence argues against the

hypothesis that extended 30 nm giant chromatin loops

represent a major part of the DNA of a given CT.

Evidence for the interchromatin compartment

The existence of an interchromatin space was already

evident by the observation of DNA-free zones in light

optical confocal sections (Figure S1). EM images of

corresponding osmium ammine-stained thin sections

of HeLa cell nuclei confirmed the existence of this

DNA-free space with variable width between domains

of compact chromatin (Figure S1). Although the

osmium ammine staining technique is highly specific

for DNA molecules (Cogliati & Gautier 1973), some

background cannot be avoided, still giving room to

speculations of occasional giant loops expanding

within the IC. For this reason we performed

additional immunogold labeling for DNA and H2B-

GFP tagged chromatin (Figure 3) and found super-

position of these signals with DNA-containing

domains revealed by osmium ammine staining, but

no labeling in the unstained regions in between.

Images at higher resolution (Figure 3A2 and B2)

reveal a chromatin substructure at a scale much

smaller than the average diameter of about ~ 500 nm

measured by us (Figure 2 and Figure S7) and others

in light optical sections for replication foci and

persistent chromatin domains with a DNA content

in the order of 1 Mb (Ma et al. 1998, Zink et al.1998; for review see Berezney et al. 2000). An IC

space with variable width and smallest visible

channels G 100 nm can be distinguished between

these substructures. Taken together with the compel-

ling evidence for ~ 1 Mb chromatin domains at the

light microscopic level, the fact that we cannot

distinguish morphologically distinct ~ 1 Mb chroma-

tin domains in EM images rules out the assumption

that these domains consist of a homogeneous mass of

intermingling chromatin fibers clearly separated at all

sites by an interchromatin domain. Instead, taken

together, light and electron microscopic evidence

supports the idea that ~ 1 Mb chromatin domains are

interconnected with each other into a higher-order


chromatin network and possess a substructure with

smaller chromatin domains and smallest branches of

the IC expanding into their interior.

Experimentally induced transient chromatinhypercondensation and concomitantexpansion of the IC

To study dynamic features of higher-order chromatin

architecture and the IC we applied an approach first

described by Bank 1939 and by Kuwada & Sakamura

1927. It allows the transient change of chromatin

compaction in living cells from normally condensed

chromatin (NCC) to hypercondensed chromatin

(HCC) by an increase of the osmolarity of the culture

medium (Figure 4A). HCC formation occurred within

1 min after the increase of osmolarity (Figure 4A) and

at all stages of the cell cycle (Figure S3 and Video S1).

Time-lapse images recorded during the first 90 s after

the replacement of physiological medium (290

mOsm) with hyper-osmolar medium (570 mOsm)

demonstrate that HCC formation was accompanied

with an enlargement of IC lacunas already detected in

the untreated nuclei (Figure 4A).

Alternatively, we increased the osmolarity of the

medium stepwise from 290 to 750 mOsm. This

procedure resulted in a stepwise increase of chromatin

condensation together with a stepwise widening of the

pre-existent IC-lacunas space (Figure 4B, arrowheads).

Additional IC channels became apparent, which

were not visible in untreated nuclei (Figure 4B2,

arrows). Due to the limited resolution of light

microscopy we were not able to determine whether

such channels were already present in untreated

nuclei or were too narrow to be detected. In spite of

this limitation our data strongly argue that the

majority of the DNA-free space observed between

HCC bundles reflects the widening of a pre-existing

IC space.

When cells with HCC were reincubated in medium

with physiological osmolarity (290 mOsm) the state

of NCC was fully recovered (Figure 5) with a similar

time-scale as for HCC formation (data not shown).

We call this experimentally induced, transient

change of chromatin condensation a NCCYHCCYNCC cycle (Figure 5A). As previously described,

cells retained their proliferative potential (Robbins

et al. 1970). Hyper-osmolar medium prepared by the

addition of different salts such as NaCl, MgCl2,

CaCl2, KCl or NaAc, as well as of saccharose, was

similarly efficient (data not shown). The effect was

visible in all studied cell types including HeLa cells,

human neuroblastoma cell lines Kelly and LAN-5,

human breast cancer cell line MCF-7 and Chinese

hamster ovary cells, as well as normal diploid human

fibroblasts, lymphocytes and amniotic fluid cells

(data not shown). An intact semi-permeable cell

membrane was obligatory for HCC formation medi-

ated by an increase in the osmolarity of the medium.

In cells with membranes permeabilized by a mild

digitonin treatment, HCC formation could no longer

be induced by an increase in the concentration of

monovalent cations but could still be induced by an

increase in that of divalent cations (Mg2+ and Ca2+)

(Figure S4). We conclude that an increase in

osmolarity of the medium likely had an indirect

effect on intact cells. The induction of a loss of

water from such cells leads to a decrease of the

nuclear volume (Figure 5B) and an increase in the

concentration of divalent cations and possibly of

other unknown factors involved in HCC forma-

tion. ATP depletion (see Methods) on itself led to

condensation of chromatin in cells kept in phys-

iological medium (Gorisch et al. 2004, Shav-Tal

Figure 2. HeLa cell nucleus with a few in-vivo-labeled CTs. A: Midsection of a fixed nucleus reveals GFP tagged histone H2B (A1) and

three clusters of densely located ~ 1 Mb chromatin domains (A2) representing (at least) three fluorescently labeled CTs. B: Pixels with gray

values above the assigned thresholds (B1: TH = 4; B2: TH =16; eight-bit format) are highlighted in red. Most labeled ~ 1 Mb chromatin

domains appear clearly separated from each other in the image segmented with the higher threshold. The low threshold image reveals a

Fcloud_ of fluorescence in the immediate vicinity of CTs, as well as between labeled domains. Note that nuclear regions between the CTs do

not reveal signals above background fluorescence intensity outside the nucleus for both thresholds. C1: Same mid-section as shown in B after

deconvolution of the original 3D image stack (12-bit format) to remove out-of-focus light and conversion to the eight-bit format. Pixels with

intensities above zero are highlighted in red. Note that detectable fluorescent signal is now restricted to the compact, labeled ~ 1 Mb

chromatin domains. C2: Maximum Z projection of the deconvolved image stack reveals clusters with all labeled ~ 1 Mb chromatin domains

representing CTs. Large zones between these labeled clusters do not contain detectable signal, clearly contradicting a major contribution of

intermingling, labeled chromatin loops to CT architecture. A few labeled ~ 1 Mb chromatin domains found apart from the CT (arrow) may be

part of chromatin protrusion as described in Figure 1.

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et al. 2004). Incubation of such cells in hyper-

osmolar medium had a slight additional, but clearly

not major, effect on further chromatin compaction

(Figure S5).

In agreement with light optical sections of nuclei

with HCC (Figure 5A2) and 3D reconstructions from

light optical image stacks (Figure 5B2), transmission

electron microscopic (TEM) images (Figure 5D)

confirmed the existence of a wide DNA-free space

between the HCC bundles. The ultrastructure of

nuclei fixed after restoration of NCC (Figure 5E)

was the same as in control nuclei (Figure 5C). The

relative arrangements of CTs were largely main-

tained over a full NCCYHCCYNCC cycle as detected

in nuclei with fluorescently labeled CTs (Figure S6).

To test whether the compaction of labeled ~ 1 Mb

chromatin domains changes as a result of HCC

formation or after incubation of cells in hypo-osmolar

medium (140 mOsm) (Figure S7), we measured the

diameters of these domains. Stacks of light optical

serial sections were deconvolved and the diameters of

clearly separated ~ 1 Mb chromatin domains were

Figure 3. Transmission electron microscopy provides evidence for interchromatin compartment (IC). Immunoelectron microscopic

visualization of DNA and GFP tagged histone H2B on HeLa cell nucleus after double-labeling with specific anti-DNA and anti-GFP

antibodies and colloidal gold markers of 15 nm and 10 nm, respectively. The ultrathin section was contrasted by a Feulgen-type staining

specific for DNA. A1: Raw electron micrograph; A2: Enlargement of box indicated in A1. B1: Same image as A1 with DNA contrast

enhanced by red color, 15 nm gold particles (anti-DNA) colored in green and 10 nm particles (anti-GFP) colored in blue. B2: Enlargement of

box indicated in B1. IC channels (arrows) expand between chromatin regions marked by the red DNA and the pseudocolored gold particles.


measured for both treated and untreated cells after

application of the lowest possible threshold level

(Figure S7A3YC3). These measurements revealed

only few, if any, changes. A minor, yet not significant

shift to smaller sizes was observed in nuclei with

HCC compared to control nuclei and nuclei of cells

exposed to hypo-osmolar medium (Figure S7D). The

latter result can be accounted for by the model of

Munkel et al. (1999), which argues that ~ 1 Mb

chromatin domains are firmly connected by chromatin

linkers. In a case where ~ 1 Mb chromatin domains were

formed as local assemblies of 30 nm chromatin fibers

kept together by weak interactions we expected that

hypo-osmolar treatment should strongly disrupt these

domains.

Chromatin bundles and interchromatin channelsform two separate, contiguous 3D networks

3D image reconstructions of nuclei in living HeLa

cells exposed to hyper-osmolar medium revealed a 3D

Figure 4. Formation of hypercondensed chromatin (HCC) and concomitant enlargement of the IC space. A: Time-lapse recording of confocal

midsections from the nucleus of a living HeLa cell (H2B-GFP) before the treatment (0 s) and subsequently at 30, 50, 60 and 90 s after

subjection to medium with 570 mOsm. HCC formation implied an expansion of IC lacunas already visible under physiological conditions

(A2, arrowheads). Maximum condensation was obtained after about 60 s. A similar time-scale was found for the decondensation process (data

not shown). B: Exemplary confocal midsections from the nucleus of a living HeLa cell with H2B-GFP tagged chromatin in physiological

medium (290 mOsm) and after incubation for 5 min each in medium with increasing osmolarity (340 mOsm; 425 mOsm; 570 mOsm; 750

mOsm). Chromatin hypercondensation increases from 340 to 570 mOsm. No further increase is apparent at 750 mOsm. Corresponding close-

ups (B2) indicate that the space between HCC bundles was generated at last to a large extent by the expansion of the pre-existing IC

compartment (arrowheads) and additional IC channels (arrows) not visible in untreated nuclei.


contiguous network of HCC bundles and a second

contiguous network of concomitantly expanded IC

lacunas and channels (Figure 6A; compare Video S2).

The relative volume occupied by the condensed chro-

matin network was in the order of 40%. The remaining

volume of about 60% was occupied by the expanded

IC space and the nucleoli. 3D-FISH with paint probes

for human chromosomes 7 and 8 in nuclei of human

diploid fibroblasts with HCC revealed spatially dis-

crete, differentially painted CTs (Figure 6B). In nuclei

Figure 5. HCC formation and restoration on normally condensed chromatin (NCC) in HeLa cell nuclei. A: Confocal midsections of a nucleus

(H2B-GFP) of a living HeLa cell in physiological medium (290 mOsm) (A1), after formation of HCC in medium with 520 mOsm (A2) and

after restoration of NCC (A3). B: 3D reconstructions (top view) of a nucleus with NCC (B1), HCC (B2) and restored NCC (B3) reveal a

pattern of interconnected H2B-GFP tagged HCC bundles together with a largely increased interchromatin space (B2). Side views (insets)

show shrinkage in the z-extension after HCC formation of ~20%. CYE: EM midsections of HeLa cell nuclei fixed in physiological medium

(C), during HCC state (D) and after restoration of NCC (E) stained by the osmium ammine technique. Note the complete restoration of the

normal chromatin architecture (C) after NCC recovery (E).


in which the two CTs were accidentally located side

by side, we observed persistent connections of HCC

bundles from one CT to the next (Figure 6B).

Stability of higher-order chromatin patterns duringrepeated NCCYHCCYNCC cycles

When the formation of HCC bundles occurred by

random clustering of chromatin domains, we expected

that the exposure of living cells to repeated NCCYHCCYNCC cycles should lead to highly variable

chromatin and IC patterns. We followed individual

HeLa cells with H2B-GFP tagged chromatin through

three NCCYHCCYNCC cycles of 5 min in hyper-

osmolar medium (570 mOsm) followed by 5 min

in physiological medium (290 mOsm). During each

cycle confocal serial sections were recorded from

nuclei with HCC and NCC (Figure 7A1Y6). Best-

fit overlays from maximum-intensity projections

recorded at subsequent cycles were generated for

nuclei with NCC (not shown) and nuclei with HCC,

respectively (Figure 7A7Y9). In addition, 3D recon-

structions from the same nuclei were generated for

comparison (Figure 7A10Y12). The apparent repro-

ducibility of the observed higher-order chromatin

patterns during repeated NCCYHCCYNCC cycles sup-

ports the hypothesis that structural patterns observed

in nuclei with HCC reflect a higher-order chromatin

and interchromatin domain topology that already

exists in nuclei with NCC. The observed similarity

was confirmed by measuring the differences between

3D data sets with a histogram-based approach, which

provides the mean-square error MSE (see Methods).

MSE values are small for a comparison of similar

input data (MSE = 0 for identical data sets), while a

comparison of increasingly dissimilar data sets yields

increasingly larger MSE values. The comparison of

repeated NCC and HCC states of the same nucleus

yielded MSE values of 7.7 T 1.5 and 8.8 T 1.9 respect-

ively (n = 15), while the pairwise comparison of MSE

values for 54 different nuclei yielded significantly

higher MSE values of 28 T 2.9 for NCC and 25 T 2.7

for HCC (p G 0.001, U-test). To further test the

reproducibility of higher-order chromatin arrange-

ments during repeated NCCYHCCYNCC cycles, we

bleached stripes into HeLa cell nuclei with H2B-

GFP tagged chromatin using an intense laser beam

(Figure 7B). The stripes were fully maintained

during repeated cycles both in the nuclear periphery

and in the nuclear interior.

RNA and DNA synthesis is inhibited in nuclei withHCC but rapidly resumed after restoration of NCC

Neither DNA nor RNA synthesis was detected, when

living HeLa cells were pulse-labeled for 10 min with

Cy3-dUTP and BrUTP after HCC formation

(Figure S8). When cells with NCC were first loaded

with BrUTP immediately before induction of HCC

formation, RNA synthesis was absent in the nucleo-

plasm, but still detected in nucleoli (Figure S9). These

findings indicate that the transcription of genes by

RNA polymerase II was quickly stalled after the

beginning of HCC formation, whereas RNA poly-

merase I was still active during early stages of HCC

formation in accordance with previous observations

(Pederson & Robbins 1970). When RNA polymerase

II transcription was selectively inhibited in control

cells by !-amanitin (2 2g/ml), incorporation of

BrUTP was also exclusively obtained into nascent

ribosomal RNA (Figure S9). Transcription and DNA

replication, however, were observed in cells that were

pulse-labeled immediately after recovery of the NCC

state (Figure S8), giving rise to patterns undistinguish-

able from those found in control cells (Figure S8). The

rapid restoration (G 10 min) of transcription and DNA

replication was unexpected considering the drastic

difference of higher-order chromatin compaction

between nuclei with HCC and NCC.

HeLa cells kept in hyper-osmolar medium for

10 min and cultured thereafter in normal medium did

not show a noticeable difference in their growth

behavior when compared to untreated controls (data

not shown). Normal cell proliferation was even

observed after the HCC state had been maintained

for 60 min (Video S1) or after cells had been

subjected to successive NCCYHCCYNCC cycles

(Video S3). With increasing duration of the HCC

state ( 91 h), however, we noted an increasing

fraction of cells that died during the procedure or

after restoration of the NCC state (data not shown).

Topography of nascent RNA, RNA polymerase IIand newly replicated DNA in nuclei with HCC

Immunofluorescence staining was used to further

investigate the 3D localization of RNA polymerase

II, nascent RNA and newly replicated DNA in HeLa

cell nuclei which were induced to form HCC 10 min

after scratch-loading with BrUTP or Cy3-dUTP

(Figure 8AYC). In all three cases signals appeared


preferentially at the surface of the HCC bundles. To

quantify this observation we measured absolute 3D

distances from all voxels contributing to nascent

RNA, newly replicated DNA or RNA polymerase II

to the closest voxel representing the surface of HCC

(Figure 9A). Indeed we found that the majority of

nascent RNA and newly replicated DNA was located

in narrow shells expanding T200 nm on either side of

HCC bundle surfaces. A similar distribution was

found for RNA polymerase II. As a control distribu-

tion we performed ADS measurements for voxels

filling uniformly the complete 3D nuclear volume

(Figure 9B, yellow bars). All evaluated distributions

differed highly significantly from this control distri-

bution (p G 0.001; U-test). As a control for the

accessibility of antibodies to the interior of HCC

bundles we labeled DNA in S-phase nuclei with

BrdU for several hours followed by the formation of

HCC (Figure 8D). As expected, absolute 3D distance

measurements showed BrdU signals within the

boundaries of H2B-GFP tagged HCC bundles

(Figure 9B, orange bars).

The choice of threshold level used to delimit HCC

bundles was guided by the patterns usually seen in

EM sections of nuclei with HCC (see Methods). We

further tested a range of threshold values above and

below the chosen level (Figure S10). This evaluation

demonstrated a highly significant difference (p G0.001; U-test) between the observed signal distribu-

tions and the control distribution for the entire range

of tested threshold levels.

Topography of nuclear speckles and bodiesin nuclei with HCC

For the quantitative evaluation of the localization of

nuclear speckles, PML bodies, and Rad51 foci with

respect to the surface of HCC bundles, we recorded the

3D location of the respective signals (Figure 8E,F). 3D

distance-to-surface measurements (see above) revealed

that the large majority of voxels belonging to nuclear

speckles and PML bodies (Figure 9C) was located in

the interior of the expanded IC, highly significant

different ( p G 0.001, U-test) to the control distribution.

In the present experiments speckles were visualized

by antibodies against SC-35, a protein that is not an

exclusive marker of nuclear speckles, but is also

present in perichromatin fibrils (Spector et al. 1991).

SC-35 labeling of the latter, however, gives a more

diffuse signal, which can be distinguished from nuclear

speckles by proper thresholding. Our finding that

speckles were preferentially located in the IC interior

is consistent with EM evidence for the topography of

interchromatin granules (the equivalent of nuclear

speckles (Spector et al. 1991). The distribution of

Rad51 foci, however, (Figure 9C) was not different

from the control distribution ( p = 0.756, U-test). It

seems possible that a fraction of Rad51 foci was

bound to chromatin and engaged in repair processes,

while Rad51 foci located in the interior of the IC

may serve for storage of repair factors.

Discussion

In contrast to the general acceptance of CT, the

architecture of CT, their interaction with neighboring

CT and the question of whether an interchromatin

compartment (IC) exists as a distinct nuclear domain

have remained a matter of controversy (for review see

Cremer et al. 2006). Figure 10 provides an updated

version of the CT-IC model, which summarizes our

present views of the global nuclear architecture. The

following discussion provides a critical evaluation of

compatible and conflicting results and interpretations

from us and others. Data and models described by

different groups including ours remind us of the

elephant studied by blind people from different sites.

Present models reflect rather the nuclear sites and scales

of resolution where their studies were undertaken than a

deep understanding of the global architecture and

functional implications of the elephant.

Figure 6. 3D networks of HCC bundles and interchromatin channels. A1: 3D reconstruction of a HeLa cell nucleus (H2B-GFP, green) after

HCC formation. The space between HCC bundles (red) represents both the expanded IC space and the nucleoli. Removal of a nuclear segment

allows a view into the nuclear interior. The hypercondensed chromatin (A2) and expanded IC space (A3) form two contiguous networks. B1:

Confocal midsection of a human fibroblast nucleus after HCC formation with painted CT 7 (red) and 8 (blue) and TO-PRO-3 stained

chromatin (green) indicates a complex folding of CTs with IC channels expanding from the territory periphery to the interior. Close-up views

of HCC bundles with and without chromosome painting (B2 and B3) suggest a direct connection between the chromatin of two adjacent CTs

further emphasized by 3D reconstructions (B4).

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Figure 7. A: Repeated NCCYHCCYNCC cycles in a living HeLa cell reveal reproducible chromatin and IC patterns. 1Y9: Nucleus of a living

HeLa cell (H2B-GFP, confocal maximum Z projections) during repeated cycles of HCC formation (2, 4, 6) and release (1, 3, 5). Overlays of

pseudocolored projections (2,red, 4,green, 6,blue) revealed merged colors expected in case of a high reproducibility of the patterns of HCC

bundles, i.e. yellow for red/green (7), pink for red/blue (8) and turquoise for green/blue (9) image overlays. 10Y12: 3D reconstructions of the

image stacks recorded from nuclei with HCC (compare 2, 4 and 6) further demonstrate the reproducibility of the HCC networks, suggesting

that these were formed on the basis of a structure pre-existing in nuclei with NCC. B: Crosswise stripes of bleached chromatin are maintained

during repeated NCCYHCCYNCC cycles. Crosswise stripes of chromatin were bleached in the nucleus (H2B-GFP) of a living HeLa cell by an

intense laser beam (1). Confocal midsections of the nucleus were obtained during repeated HCC formation (2, 4, 6) and release (3, 5). The

first image (1) was obtained 5 min after bleaching, the others (2Y6) thereafter at time intervals of 10 min. The bleached cross remained visible

during all cycles.


Chromatin hypercondensation reveals a 3Dchromatin network

In the present study we observed a contiguous 3D

network of chromatin bundles with a width of several

hundred nanometers after the formation of HCC. Little

is known about the mechanism of HCC formation, but

an increase in the concentration of divalent cations and

the resulting decrease in the negative charge of the

DNA backbone may play an essential role (Hansen

2002, Horn & Peterson 2002). The observed inhibi-

tion of transcription and replication in nuclei with

HCC possibly resulted from an inhibitory effect of

increased concentrations of divalent cations on the

activity of RNA polymerase II and DNA polymerase.

In addition transcription and replication require

strand separation, which may be hindered in nuclei

with HCC. The rapid resumption of RNA synthesis

and DNA replication, and the viability of cells kept for

up to 1 h in the HCC state or subjected to repeated

NCCYHCCYNCC cycles suggest that the functional

topography of transcription and replication was not

severely disrupted during HCC formation.

Our observation that the global architecture of the

3D networks of chromatin bundles was largely

maintained during repeated NCCYHCCYNCC cycles

suggests permanent connections between CT and

chromatin domains in both nuclei with HCC and

NCC. In nuclei with NCC a structural continuity

between labeled and unlabeled chromatin domains

with little interpenetration was previously demon-

strated at the EM level (Visser et al. 2000). The

concept of a global 3D chromatin network, estab-

lished in early G1, is consistent with the reported

overall stability of CT and chromatin domains from

mid-G1 to late G2 despite evidence for locally

constrained chromatin movements (Gerlich et al.2003, Walter et al. 2003). Specific chromatin con-

nections to the nuclear lamina play an important

role in the maintenance of higher-order chromatin

arrangements, and disruptions of these connections

result in severe diseases called laminopathies

(Gruenbaum et al. 2005). In spite of the important

role of the nuclear lamina as a structure for the

attachment of gene-poor, midYlate-replicating chro-

matin, we think it unlikely that specific chromatinYnuclear envelope connections fully explain the

apparent stability of higher-order chromatin arrange-

ments during repeated NCCYHCCYNCC cycles. In

addition to nuclear crowding effects (Hancock 2004)

non-covalent or possibly also covalent proteinYprotein, DNAYprotein, DNAYDNA and DNAYRNA

interactions may play a role as interchromatin

Flinkers_ (for a recent review see Adkins et al.2004). Such linkers could also explain the mainte-

nance of higher-order chromatin arrangements from

one cell generation to the next (Gerlich et al. 2003).

Specific and stable chromatin linker patterns estab-

lished during terminal cell differentiation may ensure

the long-term stability of cell-type-specific chromatin

arrangements. Evidence for cell-type-specific changes

of higher-order chromatin arrangements during post-

mitotic differentiation (Manuelidis 1990, Martou &

De Boni 2000, Moen et al. 2004, Solovei et al. 2004,

Su et al. 2004) raises questions on the mechanism(s)

responsible for this plasticity of nuclear architecture.

Attempts to identify the molecular nature, plasticity

and possible cell-type-specific diversity of interchro-

matin linkers will likely become a major focus of

future analysis.

Our observations open a new avenue to explore the

long-disputed question of whether a nuclear matrix is

involved in higher chromatin organization and

nuclear functions (Pederson 2000, Nickerson 2001).

Beyond all pro and con discussions the introduction

of the nuclear matrix concept (Berezney et al. 1995,

for review see Pederson 1998) argued for a higher-

order organization with distinct nuclear compart-

ments at a time when the nucleus was typically

considered as a bag of complex biochemistry

performed on intermingling chromatin fibers. Bio-

chemical nuclear matrix preparations contain pro-

teins involved in the assembly of nuclear speckles,

bodies and functional machineries (Berezney et al.1995), which are part of the IC and perichromatin

region, respectively. We have suggested that bio-

chemical matrix preparations obtained by high salt

and DNase treatment may yield an enrichment of the

content of the IC space, independent of whether the

various components are associated with a contiguous

nuclear matrix in vivo or whether such a matrix does

in fact not exist (Cremer et al. 1995). If it exists, this

nuclear matrix may provide the contiguous scaffold

structure to which small-scale chromatin loops

retract during HCC formation. Accordingly, the IC

and the perichromatin region may still be considered

as candidate compartments, where a nuclear matrix

involved in nuclear biochemistry may form tran-

siently in vivo, but present evidence does not

strengthen this assumption. Alternatively, it may be


argued that branched 10 nm core filaments, which

have been described as the hallmark of the nuclear

matrix in situ (Nickerson 2001), are located in the

interior of HCC bundles. In the absence of more

decisive evidence for a contiguous 3D nuclear matrix

within or at the surface of HCC bundles, we argue

that the role which has been contributed to the

nuclear matrix in the organization of higher-order

chromatin arrangements can be fulfilled by a 3D

chromatin network per se. CT modeling showed that

the assumption of local proteinYprotein and proteinYDNA interactions is sufficient to model CT (Munkel

et al. 1999). It is a question of terminology, if one

chooses to summarize under the generic term nuclear

matrix all inter- and intra-CT linker proteins involved

in the formation of a contiguous 3D chromatin

network.

The chromatin compaction and interminglingcontroversy

It has been reported that both relatively small

chromatin loops (in the range of about 50Y200 kb)

and giant loops (in the Mb range) exist in the cell

nucleus (for review see Kosak & Groudine 2004).

The question which has fueled controversial discus-

sions, concerns the relative proportions of small- and

large-scale loops, their compaction ratios, and their

relevance for the global nuclear architecture (for

review see Cremer et al. 2006). The data discussed

below, as well as data from computer modeling

(Dietzel et al. 1998, Munkel et al. 1999), argue

against the random walk/giant loop model of CT

architecture (Sachs et al. 1995), as well as against the

claim that a large fraction of extended chromatin

fibers expand from CT and meander in their

neighborhood (see Introduction).

By 3D-FISH and confocal microscopy we analyzed

the nuclear topography of the particularly gene-dense,

transcriptionally active chromatin region on HSA

11p15.5. previously analyzed by the Bickmore group

(Mahy et al. 2002, Chubb & Bickmore 2003); see

Introduction). We demonstrate that the complete

region frequently expanded from the HSA 11 CT as

a finger-like protrusion with a degree of compaction

at least 10-fold higher than the compaction of an

extended 30 nm chromatin fiber. While it is not

certain whether the higher-order compaction mea-

sured for this region is typical for all other gene-

dense regions, our postulate that most chromatin has

a degree of compaction much higher than extended

30 nm fibers was substantiated by our analysis of

clusters of fluorescently labeled ~ 1 Mb chromatin

domains representing parts of chromosome territo-

ries. The present understanding of DNA replication

maintains that the whole genome is replicated in

so-called replication foci in a precise spatial and

temporal order during S-phase (Sporbert et al. 2002).

Since these foci persist as ~ 1 Mb chromatin domains

outside of S-phase, we argue that the possible range

of compaction of such domains represents the global

compaction level of chromatin in the cell nucleus.

We measured an average diameter of 500 nm for

these domains on deconvolved images. Considering

that their DNA content ranges between a few

hundred kb and several Mb (Jackson & Pombo

1998, Ma et al. 1998, Berezney et al. 2000, Koberna

et al. 2005), we can estimate compaction levels

between 1:200 (for 300 kb) and 1:660 (for 1 Mb). We

consider these numbers as conservative estimates

given the fact that our measurements of foci

diameters should rather be considered as overesti-

mates. In addition, most recent data indicate that the

diameters of replication foci are typically smaller

than 500 nm (Koberna et al. 2005). Considering the

still-unexplored range of variability with respect to

chromatin domain size and DNA content, we do not

presently exclude the possibility that segments

G 200 kb as detected by FISH experiments with

typical BAC probes may have compaction levels close

to 30 nm fibers. Despite this uncertainty present evi-

dence strongly argues against a major contribution of

extended 30 nm giant loops to the global chromatin

architecture.

While our imaging system was not sensitive

enough to detect individual 30 nm giant loops, we

should have detected fields of intermingling labeled

30 nm chromatin fibers expanding from labeled CT

in our present experiments. Accordingly, we consider

the fact that we could not demonstrate specific

Figure 8. Visualization of nascent RNA, DNA, RNA polymerase II, SC-35 speckles, and nuclear bodies in HeLa cell nuclei with HCC. In

nuclei with HCC bundles (H2B-GFP, red) the following components (green) were visualized directly or by immunocytochemistry (see

Methods): A: nascent transcripts after 10 min incorporation of BrUTP, B: nascent DNA after 10 min pulse-labeling with Cy3-dUTP, C: RNA

polymerase II, D: DNA after 7 h BrdU labeling, E: SC-35 speckles, F: Rad51-foci and G: PML-bodies.

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fluorescent label between the labeled CT as strong

support that such fields do not exist. In this context it

is interesting to note that in-vivo swelling of nuclei

by incubation of cells in hypotonic medium did not

reveal major changes in the measured diameters of

~ 1 Mb chromatin domains compared with nuclei of

cells kept in physiological or hypertonic medium (see

also below). This finding argues for linkers, keeping

these chromatin foci together even under conditions

which should facilitate the spreading of unlinked

convolutes of 30 nm fibers.

The detailed structure of ~ 1 Mb chromatin

domains is still a matter of speculation. While it is

possible that many short segments of 10 and 30 nm

chromatin fibers may exist within the boundaries of

such domains, estimates of average compaction

levels of DNA in these domains suggest a chromatin

compaction significantly above the level of extended

30 nm chromatin fibers. A deep gap of understanding

and interpretation still exists between light micro-

scopic and EM studies (exemplified by Figures 2

and 3). Whereas evidence for CT and discrete ~ 1 Mb

chromatin domains is compelling at the light micro-

scopic level, the interconnection of CT into a 3D

chromatin network, and the lack of possibilities to

color CT differentially in EM studies, provide two

decisive reasons why electron microscopists failed to

discern individual CT despite the much higher

resolution they could use. The same reasons likely

account for the failure to clearly distinguish discrete

~ 1 Mb chromatin domains at the EM level. These

domains are arguably not simple chromatin clumps

but have a still-unidentified higher-order spaceYtime

structure (for models see Cremer et al. 2000). Given

the lack of a quantitative assessment of 10 and 30 nm

fiber segments present within such domains, as well

as the lack of robust data on the variability of domain

sizes and DNA content, our present estimates

indicating an average compaction of DNA in chro-

matin domains considerably higher than expected for

30 nm chromatin fibers are suggestive but not

compelling. The stability of these domains in our

view, however, strongly argues against a structure

simply built up from randomly intermingling 30

nm fibers. An average diameter of about 500 nm

for ~ 1 Mb chromatin domains sets a limit to the

maximum length of an extended 30 nm fiber within

such a domain. While our experiments leave open

the possibility that a substantial fraction of the

genome is decondensed at any given time to the 30 nm

fiber level within chromatin domains, the idea that a

large fraction of 30 nm fibers meanders between

chromatin domains is clearly not supported by our

data. Whatever the true contribution of short seg-

ments of 10 and 30 nm chromatin fibers to higher-

order chromatin organization may be, present data

strongly argue against the view that 30 nm chromatin

giant loops expanding several microns into the

neighborhood of chromosome territories are essen-

tially involved in regulatory events requiring the spatial

co-localization of gene sequences located on different

chromosomes (Osborne et al. 2004, Spilianakis et al.2005, Bacher et al. 2006, Ling et al. 2006, Xu et al.2006).

Experimental work recently reported by Branco

and Pombo in support of their interchromosomal

network (ICN) model requires special attention

(Branco & Pombo 2006). The authors painted CT

on ultrathin cryosections (~ 150 nm) of human

lymphocyte nuclei with different fluorochromes,

and analyzed regions of overlapping colors between

neighboring CT, which they considered as regions

with significant intermingling of chromatin fibers

from both CT. Overlap measurements were based on

the subjective setting of gray value thresholds and

the extent of potential intermingling critically

depends on this choice. Intermingling chromatin

fibers at sites where chromatin domains form direct

contacts is consistent with our results, which argue

for direct contacts between CT at many sites

resulting in a contiguous 3D chromatin network, but

do not exclude the presence of an interchromatin

Figure 9. Quantitative analysis of the topography of nascent RNA, DNA, RNA polymerase II, nuclear speckles and bodies in HeLa cell

nuclei. A: 3D distance values for signal voxels attributed to nascent DNA (bright green bars), nascent transcripts (dark green bars) and RNA

polymerase II (olive bars) demonstrate the location of most of these signals close to the surface of HCC bundles (compare scheme in the

inset). B: H2B-GFP signals (red bars), as expected, were located exclusively inside of HCC bundles. BrdU signals (orange bars) recorded

after 7 h BrdU labeling were also mainly located within HCC bundles (comparable to a schematic distribution drawn in the inset). Yellow

bars show a control distribution (see Methods). C: More than 90% of signal voxels from PML bodies (bright blue) and SC-35 speckles (dark

blue) located within the expanded IC space (compare scheme in the inset). In contrast, only about 70% of signal voxels attributed to Rad51

foci (turquoise) located outside of HCC bundles, the remaining fraction suggests a more intimate connection with HCC bundles.

R


Figure 10. Update of the chromosome territoryYinterchromatin compartment (CT-IC) model. A: Cartoon of a partial interphase nucleus with

differentially colored higher-order chromatin domains (red and green) from neighboring CTs separated by the IC (white). This model

postulates that the nucleus and each CT is built up from two structurally distinct compartments: a 3D network of chromatin domains with

compaction levels much higher (10 times and more) than the compaction level of an extended 30 nm fiber (for details see text) and an

integrated IC channel network with nuclear speckles and bodies (blue), which expands between these domains, independently of whether they

belong to the same or different CTs. The width of the IC varies from the micrometer scale, e.g. IC lacunas containing large nuclear speckles,

to nanometer scales (see B). Intrachromosomal, respectively interchromosomal, rearrangements can occur when double-strand breaks are

induced in neighboring chromatin domains of the same respectively different CTs. Opportunities for rearrangements are increased, when

constrained Brownian movements of neighboring chromatin domains result in a transient decrease of the width of small IC channels. The

perichromatin region (gray) is located at the periphery of chromatin domains and forms a functionally important border zone (100Y200 nm)

with certain genes or segments thereof poised for, or in the process of, transcription. Although the CT-IC model postulates that permanently

silenced genes are hidden in the interior of compact chromatin domains, the possibility that most or all genes are located at chromatin domain

borders has not been excluded. BYD: Enlargements of nuclear sites indicated in A show ~1 Mb chromatins domains (red and green) and the

interchromatin space (white) with nuclear speckles, bodies (blue), as well as preformed modules of the transcription and splicing machineries

(pink). Diffusion of individual proteins into the interior of compact chromatin domains is likely not prevented. Several ~1 Mb chromatin

domains may form still larger domains seen in EM images as chromatin clumps. The finest branches of the IC with a width G100 nm may

penetrate into the interior of ~1 Mb chromatin domains and end between ~100 kb loop domains (not shown). B: The red ~1 Mb chromatin

domain denotes the end of a higher-order chromatin protrusion, which expands from the respective red CT into the interior of the green CT

(compare A). We assume that the expansion of these higher-order protrusions is guided by the IC. Locally decondensed chromatin loops

contribute to the perichromatin region (gray). Note that the narrow IC channel allows for direct contact of loops from neighboring ~1 Mb

chromatin domains (arrow). C: This enlargement shows somewhat wider IC channels compared to B. Note one larger decondensed loop

(arrow) expanding along the perichromatin region. D: Direct contact between chromatin domains from neighboring CTs (arrow). The possible

extent of intermingling of chromatin fibers at such connections is not known.


compartment. Zones of color overlap can also result,

when distinct chromatin domains locate in close

vicinity or on top of each other (Figure S11).

Images presented by Branco and Pombo (e.g. their

Figure 1AYF) suggest the presence of chromatin foci

with sizes typical for ~ 1 Mb chromatin domains.

Branco and Pombo also carried out transmission EM

of ~ 150 nm sections, in which chromatin of

neighboring CT was differentially labeled with 5

and 10 nm gold particles, respectively. The spatial

distribution of these gold grains provides an exper-

imentally testable criterion to distinguish between

the ICN and CT-IC models. In case of true

intermingling of chromatin fibers one would expect

to see 5 and 10 nm gold particles evenly distributed

in close neighborhood throughout the section. In

case of the CT-IC model, however, we would expect

that distinct clusters of 5 and 10 nm gold particles in

close proximity represent distinct ~ 1 Mb or (given

the resolution of the EM) ~ 100 kb chromatin

domains.

We do not rule out that more sensitive approaches

will reveal a small fraction of extended 30 nm fibers/

loops, which escaped our notice, or that zones of

intermingling 30 nm giant loops can form in other

cell types or under conditions not yet tested. Yet the

weight of our present data, as well as previously

published data (Jackson & Pombo 1998, Munkel

et al. 1999, Sadoni et al. 1999, Berezney et al. 2000)

supports the conclusion that the large majority of the

genome is present at any given time in the form of

higher-order chromatin domains with compaction

levels more than 10 times higher than the compaction

of extended 30 nm chromatin fibers. This idea

becomes more plausible if one considers (1) that only

a small fraction of the genome is actually transcribed

at any given time and (2) that it may suffice that at

any given time point of transcription only a small

segment of the entire gene is actually exposed to the

transcription or replication machinery located in the

perichromatin region (Figure 10) (Cremer et al.2006).

The interchromatin compartment controversy

A structurally and functionally defined interchroma-

tin compartment (IC) is a cornerstone of the CT-IC

model (Figure 10), but the existence and functional

importance of an IC has been disputed by other

models (see Introduction). While a field of intermin-

gling chromatin fibers expanding between CT pro-

vides sufficient space for nuclear speckles and

bodies, the functional implications of such an

interchromatin space differ from the structurally and

functionally defined interchromatin compartment

(IC) postulated by the CT-IC model (Figure 10).

Branco & Pombo (2006) have argued against the

existence of an interchromosomal domain (ICD),

which would isolate CT from each other (Zirbel et al.1993). Consistent with their model, the visualization of

all heterologous CT in human fibroblast nuclei by

multicolor 3D FISH experiments revealed a topogra-

phy in which CT are close neighbors, clearly not

isolated from each other by a wide, light microscop-

ically distinguishable ICD (Bolzer et al. 2005). In

contrast with the ICN model, however, our study

demonstrates a 3D network of chromatin bundles in

nuclei with HCC. A demarcation of individual CT was

possible only, when neighboring CT were differen-

tially colored. In the light of evidence provided by us

and others, CT may be compared with sponges, which

consist of a network of interconnected higher-order

chromatin domains/fibers and an integrated, internal

IC channel network (Cremer et al. 2006).

EM investigations have provided evidence that the

IC comprises about half of the nuclear space and that

its interior is mostly DNA-free (Lopez-Velazquez

et al. 1996, Esquivel et al. 1989, Visser et al. 2000).

A border zone of decondensed chromatin, called the

perichromatin region, separates the interior of IC

lacunas and channels from the compact interior of

higher-order chromatin domains (for review see

Cremer et al. 2004, Fakan 2004a). The perichromatin

region contains perichromatin fibrils (Monneron &

Bernhard 1969), which represent the in-situ forms of

hnRNP transcripts (Nash et al. 1975, Fakan et al.1976, for review see Fakan 1994). In addition to the

synthesis and co-transcriptional splicing of RNA

(Fakan & Bernhard 1971; Fakan et al. 1984), the

perichromatin region serves as a nuclear site for

DNA replication (Jaunin et al. 2000).

Further strong evidence for an IC as a structurally

defined compartment stems from the observation that

certain proteins are able to form artificial 3D net-

works of filamentous protein structures (Bridger

et al. 1998, Gorisch et al. 2005, Richter et al.2005). The formation of closed networks could be

explained by the guidance of filament growth within a

pre-existent IC, although this prediction has not yet

been supported by direct observations. According to


this view the complete 3D network provides a struc-

tural marker for the IC, or at least the part of the IC,

which is available for the growing filamentous

structures. In case of the ICN model filamentous fibers

are considered to grow anywhere between intermin-

gling chromatin fibers and an explanation must be

provided as to why these fibers do not end anywhere in

the nucleoplasm without making contacts.

The present study for the first time presents direct

evidence for the widening of a sizeable part of pre-

existing IC channels/lacunas upon induction of HCC

formation. Notably, we did not observe a shrinkage

of ~ 1 Mb chromatin domains in nuclei with HCC

compared to nuclei with NCC. Therefore, we argue

that the observed widening of the IC at least in part

results from a contraction of chromatin linkers,

which connect neighboring chromatin domains

(Figure S12). This contraction results in the observed

3D network of chromatin bundles with diameters of

several hundred nanometers. We observed nuclear

speckles and PML bodies within this expanded IC

space and were able to locate most nascent DNA,

nascent RNA and RNA polymerase II within a region

of about T200 nm from the surface of HCC bundles.

Within the limits of light microscopic resolution

these results are consistent with ultrastructural obser-

vations from RNA labeling experiments that con-

firmed the location of hnRNA transcription sites

within the perichromatin region and support the

functional role of this border zone (Fakan &

Bernhard 1971, Spector et al. 1991, Cmarko et al.1999, Verschure et al. 1999). We assume that this

topography is largely maintained in nuclei with

HCC. Accordingly, rapid functional recovery after

HCCYNCC transition should require movements of

DNA and functional machineries only in the nano-

meter scale, not large-scale movements.

Modeling experiments demonstrate that translo-

cations frequencies measured for pairs of CT after

ex-vivo exposure to ionizing radiation can be

explained by computer simulations based on the CT-

IC model (Kreth et al. 1998, 2004). These simulations

assume that a close proximity of two double-strand

breaks is required to allow the erroneous rejoining of

broken ends by the same repair machine. The CT-IC

model provides many opportunities for such events

not only at sites of direct contacts between chromatin

domains, but at any site where small-scale loops

from different chromatin domains come into contact

within narrow interchromatin channels (Figure 10

and Figure S11). These opportunities become even

more frequent when we consider constrained Brow-

nian movements of chromatin (Marshall et al. 1997,

Bornfleth et al. 1999, Chubb et al. 2002, Gasser

2002) paralleled by a continuous widening and

narrowing of IC channels. Furthermore, the CT-IC

model can explain the experimental observation that

intrachromosomal rearrangements occur in large

excess compared to interchromosomal rearrange-

ments (Cornforth et al. 2002), which would not be

expected in a case of intense CT intermingling.

These studies emphasize the importance of nuclear

architecture for the formation of chromosome aber-

rations versus the maintenance of genome integrity.

Finally, we wish to emphasize that none of the

present models can claim to provide final answers.

The question of basic principles and cell-type-

specific modifications of nuclear architecture, and

how this architecture is involved in major nuclear

processes, has not been solved. Understanding nuclear

architecture remains a future goal, yet the develop-

ment of new approaches during the past few years has

made this goal accessible, and the present discussion

of controversial findings and models demonstrates

that this exciting field of research is coming of age.

We hope that the formulation of a generally agreed

model will become possible in the future, but a much

broader and more secure basis of data is required to

accomplish this goal. Accordingly, we are aware that

the CT-IC model, like other present models, does not

provide answers which we would consider as written

in stone, but rather an incentive for further studies.

However, despite the deficiencies of all present

models, including the CT-IC model, we conclude that

two features have now been firmly established,

namely (1) that nuclei show higher-order chromatin

arrangements with CT and chromatin domains, which

form a global chromatin network with compaction

levels above 30 nm chromatin fibers, and (2) that a

structurally defined interchromatin compartment

exists. Accordingly, we argue that the perichromatin

region, which supposedly provides the structural and

functional link between these two networks, should

become a central focus for future research.

Acknowledgements

We thank Christian Lanctot (University of Munich)

for helpful comments on the manuscript, and K.


Sullivan (Scripps Research Institute, La Jolla, USA)

for the gift of a HeLa cell line expressing histone

H2B-GFP. We are grateful to Elisabeth Kremmer

(GSF) for raising the anti-RNA polymerase II anti-

bodies, and to Jitka Fakan and Francine Voinesco for

excellent technical assistance with electron micros-

copy. This work was supported by grants to T.

Cremer (Deutsche Forschungsgemeinschaft CR59/

22, Wilhelm-Sanderstiftung 2001.079.2 and from

the Bundesministerium fur Bildung und Forschung

NGFN II- EP (0313377A)) and to S. Fakan (Swiss

National Science Foundation 3100-064977.01).

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