Chromosome territories reposition during DNA damage-repair response
© Mehta et al.; licensee BioMed Central Ltd. 2013
Received: 30 July 2013
Accepted: 13 December 2013
Published: 13 December 2013
Local higher-order chromatin structure, dynamics and composition of the DNA are known to determine double-strand break frequencies and the efficiency of repair. However, how DNA damage response affects the spatial organization of chromosome territories is still unexplored.
Our report investigates the effect of DNA damage on the spatial organization of chromosome territories within interphase nuclei of human cells. We show that DNA damage induces a large-scale spatial repositioning of chromosome territories that are relatively gene dense. This response is dose dependent, and involves territories moving from the nuclear interior to the periphery and vice versa. Furthermore, we have found that chromosome territory repositioning is contingent upon double-strand break recognition and damage sensing. Importantly, our results suggest that this is a reversible process where, following repair, chromosome territories re-occupy positions similar to those in undamaged control cells.
Thus, our report for the first time highlights DNA damage-dependent spatial reorganization of whole chromosomes, which might be an integral aspect of cellular damage response.
DNA-damaging agents, both endogenous as well as exogenous, constantly pose a threat to the genome. Protecting the genome from damage and an efficient machinery to repair damage are essential for maintaining genomic integrity. A failure to do so can lead to mutations or result in cell death. Since, DNA is non-randomly and spatially distributed within nuclei, it is important to understand the interplay between DNA damage response (DDR) and nuclear or chromosome organization.
Genomes within interphase nuclei occupy discrete, three-dimensional regions known as chromosome territories (CTs) [1, 2]. The non-randomness of CT organization within an interphase nucleus is conserved in organisms throughout evolution from flies to humans [1, 3, 4]. It has been speculated that the conserved arrangement of CTs is important for mediating genome functions [1, 5–10].
Although non-random, changes in CTs have been associated with altered cellular physiology. Interestingly, the organization of interphase CTs has been shown to change during differentiation, quiescence and senescence [1, 11–13] (Mehta IS, et al., unpublished work). Altered chromosome positioning has also been observed in some diseases and cancers [11, 14, 15]. Specifically, recent reports suggest that changes in whole chromosome or chromosomal domain positions are associated with modulations in transcriptional status [1, 16–19] and chromatin architecture .
The link between transcription and CTs has been supported by studies where gene-rich chromosomes are present in the nuclear interior while gene-poor chromosomes are located at the periphery [20, 21]. Further, studies have demonstrated that regions of DNA that are within the nuclear interior show higher transcriptional activity [22–24]. Although chromosome repositioning has been associated with varied cellular outputs, the underlying biological significance and the molecular mechanisms that mediate these relocalizations are poorly understood.
Local chromatin structure affects the damage frequency and repair mechanisms. It is interesting that open chromosomal material or chromatin incurs damage more easily than closed heterochromatin [25–29]. In addition, DNA composition is also known to impinge upon damage and repair mechanisms. DNA associated with features such as being at nuclear center, having higher gene density or GC content, or chromatin enriched with hyperacetylated histones, is more prone to damage [27, 30, 31]. Moreover, gene-rich regions and euchromatin are repaired faster than gene-poor and heterochromatic regions [25–29]. In addition to local changes in chromatin structure and functions [29, 32], global alterations in chromosome architecture and position are likely to play an important role during DDR [33–38]. While studies have demonstrated that chromosome domains have to remain stationary following laser irradiation to assist repair, the studies focused on specific chromosomal loci and domains or only certain chromosomes [35, 39–42]. However, in contrast, recent biophysical modelling studies have indicated that structural reorganization of genome and chromosome domains assists DDR [43, 44]. Specifically, changes in the nuclear matrix attachment of chromosomes and repositioning of chromosome domains [45–49] have been hypothesized to assist cell cycle arrest, alter the transcription profile and the accessibility of damage sites to repair proteins [33, 35, 43]. Very recent reports for yeast have shown that damaged chromosomal domains undergo large-scale nuclear movements and it has been suggested that this phenomenon is vital for homology searches during homologous recombination (HR) [36, 50–52]. Therefore, investigations of alterations in CTs, if any, are important for understanding the global effects of chromatin, chromosome packaging and nuclear architecture vis-à-vis DNA repair.
In this study, we have addressed the effect of DNA damage on CTs in primary human fibroblasts. We find that while most of the chromosomes do not change location, some gene-rich chromosomes, such as chromosomes 17, 19 and 20, reposition themselves after DNA damage in a dose- and time-dependent manner. Our observations also implicate the importance of DNA damage sensing in bringing about such alterations, whereby inhibiting the activity of DNA-dependent protein kinase (DNA-PKcs) and Ataxia Telangiectasia mutated kinase (ATM kinase) perturb this relocalization. The results also show that these spatial changes in CTs are reversible, since the DNA damage is repaired. Our report highlights an intricate relation between CTs and DNA damage response.
Results and discussion
Repositioning of select chromosomes after DNA damage
Positions of all human chromosomes in fibroblasts before and after DNA damage
Number of genes/megabases
Chromosome size (base pairs × 106)
Although, it is still far from clear why only certain chromosomes reposition after DNA damage, previous research by Falk and co-workers [29, 54] clearly indicated an association between gene-rich chromosomes and the high frequency of phosphorylated histone 2A.X (γH2AX) foci. To assess if the frequency of damage affects DNA damage-dependent CT repositioning, we performed combined immuno-FISH experiments to analyze the occurrence of γH2AX foci on chromosomes that show relocalization after DNA damage. In concurrence with the previous research [29, 54], we observed a slight increase in the number of γH2AX foci on gene-rich chromosomes that show relocation after DNA damage vis-à-vis the chromosomes that do not change location after DNA damage (Additional file 4: Panel 2). These results, suggest that chromosome relocalization after DNA damage might be an intrinsic component of DDR.
Chromosome repositioning is DNA damage dose and time dependent
Further, to assess whether chromosome relocation was linked to DSB recognition following breaks, we also analyzed the DSB kinetics (TUNEL assay and γH2AX). We observed that chromosome 19 relocation starts at about 2 hours following cisplatin treatment (Figure 4G) and coincides with an increasing γH2AX response (Additional file 4: Panel 2 and Figure 4K). This relocation is complete after 3 to 4 hours (Figure 4E,F,G,H,I). These findings were interesting because the repositioning was initiated only after DDR was elicited and thus, suggested that damage sensing and recognition preceded territory repositioning.
Repositioning of chromosome territories is dependent upon DNA damage sensing
Since CT repositioning after DNA damage occurs post phosphorylation of H2AX, we hypothesized that factors involved in DSB recognition and downstream signaling might be involved. Specifically, we wanted to investigate the role of DSB sensors Ataxia Telangiectasia mutant (ATM)/Ataxia Telangiectasia and Rad3 related (ATR) and DNA Protein Kinase C (DNA-PKcs), which are involved in eliciting a repair response including cell cycle arrest, phosphorylation of H2AX and altering chromatin structure at the sites of damage [55–59].
Chromosome territories revert back after repair
Reversal of chromosome territories to locations similar to those of control cells post repair may require passage through mitosis
This study reports a hitherto unknown association between DNA damage sensing and chromosome territories (CTs). We demonstrate that CT positions alter after DNA damage and are intrinsically coupled to the ability of cells to elicit a damage response dependent on ATM/ATR and DNA-PKcs activity. Local chromatin structure and DNA damage sensing mechanisms have been known to crosstalk with each other and mediate an efficient repair response [25–29, 33]. Although the type and kinetics of repair are governed by the local chromatin structure (heterochromatin versus euchromatin) [25, 29, 32], the behavior of chromosomes as a whole has not been addressed thus far. In this regard, our results show reorganization of whole CTs in response to DNA damage. While some gene-rich CTs relocate (for example, 12 and 17) and others do not (for example, 11 and 14), a better correlation is observed in the lack of relocation of gene-poor chromosomes. It would be thus interesting to investigate further the properties of gene-rich CTs to determine their propensity to relocate.
Recent studies of yeast have hinted at the possibility of large-scale chromosome movement that ensue because of heightened mobility of broken DNA ends [29, 35, 36, 49, 50]. Chromatin repositioning, including the relocation of single or multiple chromosome loci or whole chromosomes, has been hypothesized to be a normal cellular response to radiation exposure [35–37, 43, 44]. Our findings not only support the hypothesis but also evidence the involvement of whole chromosomal mobility in DDR. These studies have also shown that the mobility of broken DNA ends influences HR efficiency. Although we cannot rule out such a possibility in our system (at least for the chromosomes that move to the interior), we do not hypothesize this to be a determining factor, since HR is not predominant in mammalian cells unlike yeast. Instead, we speculate that a potential bias in transcriptional activity and chromatin structure in these chromosomes might be relevant for such repositioning and needs to be addressed in the future. More importantly, we have addressed the relation between DNA damage response and nuclear organization vis-à-vis chromosomal localization.
Our results show that repositioned CTs revert to distributions similar to their control counterparts in a repair-dependent manner whereby restoration coincides with loss of γH2AX foci. Importantly, these raise fundamental questions about the link between DNA repair proteins and chromosomal relocalization, and suggest that CT reorganization could be an integral component of cellular DNA damage response. In conclusion, our report has investigated a novel aspect of DNA damage response and should lead to further studies aimed at understanding the role of nuclear complexity in maintaining genomic integrity.
Materials and methods
Cell culture and treatments
Early passage primary normal human dermal fibroblasts (NHDFs; Lonza) and normal human lung fibroblasts (Lonza) were cultured in 15% fetal bovine serum (FBS) supplemented DMEM and fibroblast growth medium (FGM) media supplemented with 2% FBS and growth supplements (Lonza). ATM mutant fibroblast lines AT2BE and AT5B1 (kindly provided by Dr Michael Weinfeld) were cultured in 10% FBS supplemented Dulbecco’s modified Eagle’s medium/nutrient mixture F-12 (D-MEM/F-12) (1:1 ratio). Cells were either treated with 1 mM H2O2 or with 25 μM cisplatin (Calbiochem) for 1.5 or 4 hours, respectively, unless mentioned otherwise in figure legends.
Two-dimensional fluorescence in situhybridization
2D-FISH was performed using the standard protocol . Briefly, cells treated with hypotonic solution (0.075 M KCl) were fixed with 3:1 (v/v) methanol:acetic acid using the standard protocol. Cells dropped on slides were taken through an ethanol row followed by denaturation in 70% formamide at 70°C for 2 minutes. The slides were then taken through another ethanol row and air-dried at 37°C. Directly labelled total human chromosome DNA probes (Applied Spectral Imaging), denatured at 80°C for 7 minutes and re-annealed at 37°C for 20 minutes, were then applied to the denatured cells followed by hybridization for 18 hours. Post hybridization stringency washes were performed and then the cells were counter-stained with 4′,6-diamidino-2-phenylindole (DAPI) in Vectashield mounting medium.
Three-dimensional fluorescence in situhybridization
To conserve the three-dimensional structure of fibroblast nuclei, a previously described FISH methodology was used . Cells, grown on slides, were fixed with 4% paraformaldehyde (PFA) and permeabilized using Triton X/saponin solution followed by four or five freeze-thaw cycles and depurination in 0.1 N HCl. Denaturation was performed using 70% and 50% formamide for precisely 3 and 1 minutes, respectively, at 73°C. Hybridization with processed probes (see 2D-FISH section) was carried out for 48 hours at 37°C. Post hybridization washes were carried out similar to 2D-FISH.
Stacks of 0.3-μm optical sections (with an average of eight) were captured using a Zeiss confocal laser-scanning microscope (LSM510). The distances between the geometric center of each chromosome territory and the nuclear center in at least 40 nuclei were measured (Bitplane Imaris software) (Additional file 3F) and frequency distribution curves were plotted. Simple statistical analyses were performed using the two-tailed Student’s t-test.
kinase Ataxia telangiectasia mutated kinase
Ataxia telangiectasia and Rad3 related
DNA damage response
Dulbecco’s modified Eagle’s medium
DNA-dependent protein kinase
Double stranded breaks
Fluorescence associated cell sorting
Fetal bovine serum
Fibroblast growth medium
Normal human dermal fibroblast
Normal human dermal fibroblast
Normal human lung fibroblast
Standard error of the mean
Terminal deoxynucleotidyl transferase dUTP nick end labeling
Phosphorylated histone 2A.X.
Research funding support from TIFR (Department of Atomic Energy, India) is acknowledged. BJR wants to acknowledge JC Bose award fellowship. We are grateful to Dr Michael Weinfeld, (Department of Experimental Oncology, Cross Cancer Institute, 11560 University Avenue, Edmonton, AB, T6G 1Z2, Canada) for providing the ATM mutant fibroblast lines. We are thankful to Amba Priya Dube and Saurabh Kokane for their contributions in the discussions on this work. We thank an anonymous referee for suggesting an enhancement to the IMACULAT methodology for incorporating equal-volume analyses.
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