Role of Chromatin Damage and Chromatin Trapping of FACT in Mediating the Anticancer Cytotoxicity of DNA-Binding Small-Molecule Drugs

DNA-targeting small molecules have been widely used for cancer treatment for many years. This broad group includes chemicals with different mechanisms of action, but their toxicity was mostly explained by their ability to cause DNA damage (1). Many of these molecules are used for cancer treatment, because tumor cells are more vulnerable to DNA damage due to their high proliferation rate and frequently nonfunctional DNA repair (2, 3). Compounds target DNA via different mechanisms. Some form chemical (covalent) bonds with DNA (e.g., cross-linking agents). Others bind DNA noncovalently via either intercalation between base pairs or accommodation in DNA grooves (1). Some compounds do not stably bind DNA, but their complex with DNA is stabilized by proteins, such as topoisomerases (4, 5). Finally, some compounds do not bind DNA but inhibit enzymes using DNA as a substrate, such as DNA polymerases or topoisomerases (6, 7).

Eukaryotic DNA is packed into chromatin, which is a highly-ordered complex of DNA and histone proteins. The basic unit of chromatin, nucleosome, consists of a core, a complex of four pairs of histones: central H3/H4 tetramer with two H2A/H2B dimers outside, wrapped with DNA. Some nucleosomes are locked by binding the linker histone H1, which forms contacts with entering and exiting strings of DNA and the core histones (8).

The DNA-damaging effect of small molecules depends significantly on chromatin organization, e.g., some agents have a preference for linker versus nucleosomal DNA (9, 10). On the other hand, there are reports that DNA-targeting small molecules perturb chromatin structure (11–14). However, how exactly they affect the chromatin and what impact chromatin alterations have on their biological activity are less studied. One of the reasons of this deficit was difficulty in separation of DNA damage from “chromatin damage” in cells.

We have previously identified small molecule, curaxin CBL0137, which has broad anticancer activity, and binds DNA without detectable DNA damage in mammalian cells (15). Although curaxin does not chemically modify DNA, it changes the shape of the DNA helix, which increases the inter–base-pair distance, unwinds DNA, and leads to the unwrapping of DNA from the histone octamer and to nucleosome disassembly in vitro and in cells (14).

Nucleosome disassembly induced by CBL0137 is “sensed” by the histone chaperone FACT (facilitates chromatin transcription; ref. 14), whose normal function is to control nucleosome stability during replication, transcription, and DNA repair (16). FACT consists of two subunits, Suppressor of Ty 16 (SPT16) and Structure Specific Recognition Protein 1 (SSRP1). It interacts with the nucleosome via several dynamic contacts with histone oligomers and DNA (17). Mammalian FACT binds poorly to the intact nucleosome (18, 19). The weakening of DNA/histone binding provides FACT access to several binding sites hidden inside the nucleosome (18). At lower CBL0137 concentrations (1 molecule per >10–100 bp), DNA is partially unwrapped from the octamer, leading to the dissociation of the H2A/H2B dimers and exposure of the surface of the H3/H4 tetramer (14). FACT binds the H3/H4 surface via its SPT16 subunit (14, 18). At higher CBL0137 concentrations (1 molecule per 1–10 bp), DNA is completely unwrapped from the nucleosome, what culminates in the disassembly of the histone octamer and the appearance of histones in the nucleoplasm (14). Unwrapped DNA undergoes significant negative supercoiling, which results in base unpairing and transition from the normal B-shape helix to alternative DNA structures (ADS). In cells treated with CBL0137, we detected the appearance of left-handed Z-DNA. The SSRP1 subunit binds DNA prone to the Z-DNA transition through its c-terminal intrinsically disordered domain (14). We named the massive binding of FACT to different components of disassembled chromatin in curaxin-treated cells “chromatin trapping” or c-trapping (14).

This study was based on the hypothesis that curaxins may not be unique in their effect on chromatin and, therefore, in the ability to cause c-trapping. We proposed that other small molecules targeting DNA may disturb chromatin, and this could be detected using c-trapping assays. We selected a set of DNA-targeting agents, representing different mechanisms of action, and tested their ability to cause c-trapping and destabilize chromatin in vitro and in vivo. We tried to uncover the reasons why some compounds have stronger chromatin-destabilizing effects than others and how it correlates with their cytotoxicity.

Chemicals, plasmids, antibodies, and cells used in the study are described in the Supplementary Materials and Methods and Supplementary Table S1.

HeLa, HT1080, MCF7, and HCT116 cells were purchased from the ATCC, and HL60/VCR were provided by R. Glazer (Georgetown University Medical Center). MCF7 cells were authenticated using short tandem repeat profiling and found to be a 100% match to original ATCC cells. Other cells were not authenticated. All cells are tested for mycoplasma contamination at least once a month.

HeLa, MCF7, HCT116, and HT1080 cells (3 × 10³ cells/well) were seeded in 96-well plates for overnight adhesion. HL60/VCR suspension cells (5 × 10³ cells/well) were seeded in 96-well plates and treated the same day. Note that 50 μmol/L of 9-Aminoacridine was used as a positive control for complete cell death. Cells were treated for 48 hours with a range of drug concentrations. Cell viability was determined with resazurin saline solution. Fluorescence was measured at 560_Ex/590_Em using Tecan Infinite 200 PRO reader.

Nascent RNA-sequencing was done using the Click-iT Nascent RNA Capture Kit (Invitrogen; cat# C10365). RNA labeling with EU was done in HT1080 cells for 15 minutes in duplicates. Isolation of labeled RNA was done according to the manufacturer's instruction. Library preparation and sequencing were done in Genomics Core of RPCI using Illumina protocols and equipment.

Chromatin immunoprecipitation (ChIP) sequencing was performed as previously described (20). HT1080 cells were treated for 1.5 hours with 3 μmol/L of CBL0137, aclacinomycin A, or 0.3 μmol/L of CBL0100. Experiment was repeated 2 times. The ChIP libraries prepared at UB Genomics and Bioinformatics Core using Illumina ChIP-seq kits were pair-end sequenced on an Illumina HiSeq2000 with 100 bp reads. Raw reads, which passed quality filter from Illumina RTA, were preprocessed by using FASTQC for sequencing base quality control and mapped to human reference genome (hg19) using BWA (21). MACS 2.0 (22) with default parameters for pair-end BAM files is used to identify peaks. Heatmaps and profiles under all conditions were generated using deeptools on RPKM-normalized coverage data using merged bam files of biological replicates. Tag densities for RefGenes body with 3 kb up- and downstream are plotted. The heatmaps for all RefGenes are clustered into four groups using k-mean, within each cluster, and the genes are ordered by mean tag density. Cluster 1 shows the greatest change of binding profiles. The different binding site status is identified by DiffBind R package (23). Heatmaps of the unexpressed and expressed genes were prepared using nascent RNA-Seq data for control-untreated HT1080 cells.

Recombinant human mononucleosomes, purchased from EpiCypher (cat# 16-0009) at 0.125 μmol/L, were incubated with compounds for 20 minutes at room temperature. Following the incubation, samples were run in 5% native PAGE at 120v at 4°C. Gels were stained with ethidium bromide, and nucleosomal DNAs were quantitated with the GelDoc-It TS imaging system equipped with a Series 6100 Camera (UVP). Experiments were repeated at least twice.

Comet assay was done using kit from Cell Biolabs, Inc. (cat# STA-355) in alkali conditions according to the manufacturer's protocol.

Cells were lysed on ice using 1x Cell Culture Lysis Reagent (Promega; cat# E1531) containing protease inhibitors (1:25, Roche; cat# 1838145). Extracts were centrifuged at 10,000 rpm for 10 minutes at 4°C to obtain the soluble fraction. The chromatin-bound proteins were obtained by resuspension of the remaining pellets in 1x Cell Culture Lysis Reagent followed by sonication 3 times for 30 seconds each using the Bioruptor UCD-200, Diagenode. Protein concentrations were measured using Quick Start Bradford 1x Dye Reagent (Bio-Rad; cat# 500-0205). Equal amounts of protein were run on gradient 4%–20% precast gels (Invitrogen) and transferred to Immobilon-P membrane (Millipore). Western blot analysis is described the Supplementary Materials and Methods.

Immunofluorescent staining is described in the Supplementary Materials and Methods. Fluorescent images of live and fixed cells and compound auto-fluorescence were obtained with a Zeiss Axio Observer A1 inverted microscope with N-Achroplan 100×/1.25 oil lens, Zeiss MRC5 camera, and AxioVision Rel.4.8 software.

Sequencing data in the form of bed files are available at GEO: GSE107595, GSE107633.

If change of DNA helical shape is responsible for nucleosome destabilization, then drugs can destabilize nucleosomes either via direct binding to DNA or via inhibition of topoisomerases, enzymes that prevent the accumulation of supercoiling in cells (reviewed in 24). Drugs can either inhibit DNA cleavage by topoisomerases (supercoiling accumulates) or religation (supercoiling is released due to DNA breaks). Reports from Henikoff and Neefjes groups showed that anthracyclines, which bind DNA and inhibit topoisomerases II, caused histone eviction from chromatin in cells and nucleosome disassembly under cell-free conditions (25, 26). However, whether they induced c-trapping was unknown.

To better understand what causes c-trapping, we used a set of known compounds with different modes of DNA binding and inhibition of topoisomerases (Table 1): (i) direct DNA binders that inhibit primarily the cleavage activity of topoisomerases—curaxin CBL0100, aclacinomycin A, Hoechst 33342; (ii) direct DNA binders that inhibit religation of topoisomerases II—doxorubicin and mitoxantrone; (iii) compounds that do not bind DNA but form a cleavable complex with DNA and topoisomerases I—SN38, or topoisomerases II—etoposide; and (iv) catalytic inhibitors of topoisomerases II that do not bind DNA—merbarone and ICRF-193. We also included an inhibitor of DNA synthesis (gemcitabine), which is unable to bind DNA or inhibit topoisomerases (negative control), and CBL0137 as a positive control for c-trapping (14, 15).

Because c-trapping and cytotoxicity of CBL0137 occur at the same concentrations (although with a time delay: c-trapping—within a minute, cell death—around 48 hours after the start of treatment; refs. 14, 15), we identified cytotoxic concentrations of the compounds in several human cell lines (HeLa, HT1080, HCT116, and MCF7; Supplementary Figs. S1–S4, Table 1) and used the range of doses (from nontoxic to maximal toxicity) to detect c-trapping. Using Western blotting, we first screened all compounds for the presence of c-trapping activity at 24 hours after start of treatment in case some compounds might work slowly. FACT subunits disappeared from the soluble and accumulated in the chromatin fractions of all cell types treated with DNA-binding compounds, CBL0137, CBL0100, doxorubicin, aclacinomycin A, mitoxantrone, and Hoechst 33342, whereas there was no change in the distribution of FACT in any cells treated with compounds unable to bind DNA directly (Fig. 1A; Supplementary Figs. S5–S7; Table 1). These data were confirmed via fluorescent imaging of live cells (HeLa or HT1080) expressing GFP-tagged SSRP1. Previously, we have demonstrated using Western blotting and immunofluorescent staining that tagged SSRP1 undergoes c-trapping similarly to endogenous FACT (14). In untreated cells, SSRP1 is diffusely distributed in nucleoplasm and enriched in nucleoli. All compounds directly binding DNA caused significant change in the visual pattern of SSRP1 distribution in nuclei. Neither of non–DNA-binding compounds had this effect (Fig. 1B; Supplementary Figs. S8 and S9). Thus, c-trapping is induced by the binding of small molecules to DNA rather than by inhibition of topoisomerases.

Live cell imaging revealed that DNA-binding compounds induced c-trapping with different kinetics, some of them within minutes, and others required 1 to 2 hours (Fig. 1B). Different kinetic was confirmed using Western blotting (Supplementary Fig. S10). This kinetic correlated with the compound accumulation in cell nuclei, monitored by autofluorescence of the drugs (Supplementary Fig. S11) and with whether compounds are substrate of multidrug transporters or not (Supplementary Fig. S12). Using these observations, we selected optimal time for measurement of c-trapping, caused by six active compound to identify dose-causing redistribution of 50% of SSRP1 and SPT16 from nucleoplasm to chromatin in two cell lines, HeLa and HCT116 (Fig. 1C and Supplementary Fig. S13). This dose positively correlated with concentration of each drug, which kills 90% of cells (IC_90%) 48 hours later (Fig. 1D and E), suggesting that the mechanism underlying c-trapping contributes to the death of tumor cells.

Previously, we established that c-trapping occurs in response to chromatin disassembly in cells (14). To test effect of the compounds on nucleosome stability, we incubated recombinant mononucleosomes with different concentrations of the drugs. All compounds able to induce c-trapping, except Hoechst 33342, caused complete disassembly of the mononucleosome. Hoechst 33342 destabilize but did not disassemble nucleosome (Fig. 2A and B).

To test the effect of the compounds on nucleosome stability within chromatin, we incubated HeLa cell nuclei with the drugs, followed by assessment of chromatin sensitivity to digestion with micrococcal nuclease (MNase), preferentially digesting protein-free DNA. All DNA-binding drugs, except Hoechst 33342, caused loss of nucleosome ladder. Hoechst 33342 inhibits nuclease activity (27) to the extent that its effect cannot be assessed. Curaxins were stronger than anthraquinones, although doxorubicin caused effects similar to curaxins (Fig. 2C and D).

To assess chromatin integrity in cells, we monitored appearance of chromatin-free histones in extracts of cells incubated with the compounds, able to cause c-trapping. A minimal amount of canonical core histones is detected in the soluble fraction of untreated cells (Fig. 2E). With increasing concentrations of the compounds, histone H3 appeared in extracts of cells treated with the curaxins and aclacinomycin A. A weak, but reproducible, increase in histone H3 was observed upon treatment with 10 μmol/L doxorubicin. No free H3 was detected in the lysates of cells treated with mitoxantrone or Hoechst 33342 (Fig. 2E and F). In addition, we monitored the distribution of the outer H2B and inner H4 histones fused with fluorescent tags in cell nuclei via live cell imaging. Excess of free histones, which are not incorporated into chromatin, is accumulated in the nucleoli, first concentrated around and later inside nucleoli (Fig. 2G; refs. 14, 28). Consistent with Western blotting, curaxins and aclacinomycin A caused eviction of both outer and inner histones (Fig. 2H and Supplementary Fig. S14). Doxorubicin caused visible nucleoli accumulation of histones only at high concentrations (≥10 μmol/L), whereas no significant difference from untreated cells was seen in case of mitoxantrone and Hoechst 33342 in line with cell fractionation data (Fig. 2H and Supplementary Fig. S14). Similar response of endogenous H3 was observed via immunofluorescent staining (Supplementary Fig. S15).

The unexpected finding was absence of nucleosome disassembly in cells treated with agents that clearly cause c-trapping, mitoxantrone, and Hoechst 33342. In cells, multiple nucleosomes are stabilized in a “closed” position by the histone H1 (8). Importantly, H1 has a very fast exchange rate (8). We proposed that some of the compounds may have limited access to DNA within a closed nucleosome, but gain access to it if the nucleosomes lack H1. In this case, H1 may not be able to reattach to the nucleosome. We tested this hypothesis by studying the dose- and time-dependent effect of the drugs on the nuclear distribution of the human histone H1 tagged with mCherry. All drugs, inducing c-trapping, caused H1 accumulation in nucleoli (Fig. 3A and B). The dose and time for histone H1 eviction positively correlated with c-trapping, suggesting that c-trapping may occur due to partial opening of the nucleosome without loss of the core histones.

So far we observed that curaxins were the strongest in all cell-based assays, whereas their effect on nucleosome in cell-free conditions is weaker than anthraquinones. We proposed that among tested compounds, curaxins have the highest capacity to bind chromatinized DNA in cells. To test this, we incubated cells with doxorubixin for 3 hours to allow the drug to bind nuclear DNA and then added the curaxins at the same concentration. Because doxorubicin and curaxins have different fluorescence spectra, we were able to monitor each drug separately. The addition of either curaxin quickly displaced doxorubicin from the cell nuclei (Fig. 3C and D). A similar experiment cannot be done with the other compounds because aclacinomycin A and mitoxantrone do not have nuclear fluorescence and Hoechst 33342 has a similar fluorescence spectrum to the curaxins.

In cells treated with CBL0137, FACT is trapped in chromatin by two distinct mechanisms: (i) binding of the SPT16 subunit to the surface of the H3/H4 tetramer, which is exposed by the detachment of the H2A/H2B dimer (“n-trapping”) and (ii) binding of the SSRP1 subunit to ADSs (e.g., Z-DNA; “z-trapping”; ref. 14). SSRP1 can also bind different ADS, such as bent or cruciform DNA, via the HMG domain (29–32). We tested whether other compounds, similarly to CBL0137, induced “n-trapping” and “z-trapping.” Separating these two processes in cells is difficult because SSRP1 and SPT16 are always in a complex, and dissociation of the dimer makes both subunits unstable (33). Previously, we used SSRP1 deletion mutants to distinguish these two phenomena. SSRP1 lacking the c-terminus DNA-binding domains (HMG and CID) can only undergo c-trapping when bound to SPT16 (n-trapping), whereas SSRP1 deletion mutants consisting of only the c-terminal domains bind DNA, either in the Z-form (CID domain, z-trapping) or as cruciform/bent DNA (HMG domain; ref. 14). CBL0137 and CBL0100 caused both n- and z-trapping (Fig. 4A and B). Surprisingly, none of other compounds changed the distribution of either HMG or CID constructs, suggesting that none of them caused z-trapping. However, all of them caused “n-trapping” (Fig. 4A and B). The only exception was that aclacinomycin A did not have an effect on SSRP1 that lacked the HMG domain, suggesting that this domain may be involved in c-trapping by aclacinomycin A.

Thus, although c-trapping looks very similar via immunoblotting, experiments with FACT deletion mutants demonstrated that the compounds induce c-trapping via different FACT subunits. Moreover, they induce different microscopic patterns of FACT distribution in cell nuclei (Fig. 4C). Taking into account difference in the shapes and sizes of nuclei, these patterns were characteristic for all cells in populations (Supplementary Figs. S8 and S9). Curaxins pattern was the most contrasting (Fig. 4C). Anthraquinones caused more subtle changes, and Hoechst 33342, the mildest, transitions between bright and dim zones without clear boundaries. Because the differences were observed even at concentrations of the compounds that caused complete transition of FACT from soluble to chromatin-bound fractions (10 μmol/L, Supplementary Fig. S16), they could not be explained by different amounts of bound and free FACT. We hypothesized that these patterns reflect binding of FACT to different chromatin regions. To test this, we compared genome-wide FACT-chromatin binding in cells treated with 3 μmol/L CBL0137, 0.3 μmol/L CBL0100 (the same visual pattern as CBL0137), and 3 μmol/L aclacinomycin A (different pattern) using ChIP with SSRP1 antibody followed by deep sequencing (seq).

In control cells, SSRP1 peaks coincide with coding regions of transcribed genes (Fig. 5A). Both curaxins induced appearance of multiple new sharp peaks genome-wide, absent in control cells (Fig. 5A). Curaxin-induced peaks were located at regions, which were previously observed in CBL0137-treated cells and identified as mini-satellites, tandem dinucleotide purine/pyrimidine repeats, prone to Z-DNA transition (Fig. 5B and Supplementary Fig. S17; ref. 14). Much more rarely new peaks appeared in aclacinomycin A samples. In most cases, they overlap curaxin-induced peaks (Fig. 5A); however, aclacinomycin-specific peaks appeared at regions annotated as G-quadruplex by Non-B DNA database (Fig. 5B; ref. 34). Although anthracyclines were reported to bind G-quadruplex DNA in cells and cell-free conditions (35, 36), there are no data of FACT binding to this type of DNA. FACT peaks in curaxins treated cells, in contrast to aclacinomycin A, demonstrated absence of colocalization with G-quadruplex regions (Supplementary Fig. S17).

To assess quantitatively FACT redistribution genome wide, we aimed to find (i) where FACT was relocated upon treatments and (ii) what proportion of FACT molecules underwent relocalization. For the first task, we identified peaks in each of the conditions and compared them with control and between-treatment groups (Fig. 5C). In line with visual impression, peaks induced by CBL0100 almost completely overlap with peaks induced by CBL0137, i.e., both curaxins induced similar redistribution of FACT from coding regions to tandem dinucleotide purine/pyrimidine repeats. In contrast, regions bound by FACT in aclacinomycin A–treated cells were not significantly different from FACT-enriched regions in control cells. Novel sites of FACT binding appeared in aclacinomycin A–treated cells were mostly shared with curaxins (Fig. 5C). From the comparison of regions of FACT enrichment, we concluded that curaxins impose FACT to leave regions where it was present in control cells and to bind new regions, whereas what happened with FACT in aclacinomycin A–treated cells was unclear.

Quantitating average FACT enrichment at coding regions of all genes, where FACT presence was changed upon any of treatment, we have found that curaxins significantly reduced FACT presence over coding regions, whereas aclacinomycin A increased (Fig. 5A and D). Proportion of FACT in aclacinomycin A–treated cells was also significantly increased at promoter regions (Fig. 5E). Similar patterns were clearly seen when we did this analysis for all genes, long noncoding RNAs, and miRNAs (Supplementary Figs. S18–S20). Clustering of the genes, based on the degree of changes, up or down, demonstrated that the strongest change was observed for the genes with the highest SSRP1 enrichment in untreated cells (Fig. 5D and F). Because FACT is expected to be enriched at coding regions of transcribed genes, we used data of nascent RNA-sequencing from the same cells to compare SSRP1 distribution at coding regions of expressed versus unexpressed genes (Fig. 5G and H). Curaxins treatment led to the reduction of FACT presence at coding regions and promoter of transcribed genes, but in opposite to the increase of FACT presence at nontranscribed genes, whereas aclacinomycin A treatment has opposite effects (Fig. 5G and H).

In line with different microscopic pattern of c-trapping, curaxins cause redistribution of FACT from transcribed regions to minisatellites, whereas aclacinomycin A induced further accumulation of FACT at coding regions of transcribed genes. These observations suggested that curaxins induce appearance of de novo sites for FACT binding and thus lead to redistribution of FACT, whereas aclacinomycin A makes existing sites more attractive for FACT binding.

To understand the role of “chromatin damage” and c-trapping in the cytotoxic activity of the tested compounds, we quantitated their effects on FACT and chromatin in cells and cell-free systems and ranked them based on these activities to run correlation analysis. Because DNA damage was traditionally seen as a reason of cell death upon treatment with some of these compounds, we also measured their DNA-damaging activity using γH2AX staining and comet assay in the same cells (HeLa and HT1080) and at the same concentration range at which c-trapping was observed (Supplementary Fig. S21). This analysis showed that c-trapping positively correlated with cytotoxicity as well as with eviction of core histones, and positive correlation with the eviction of H1 was also observed, but not significant. Most importantly, cytotoxicity within this group of compounds significantly correlated with c-trapping, but not with DNA-damaging activity of the compounds (Fig. 6 and Supplementary Fig. S22). This observation suggests that ability of compounds directly binding DNA in cells, to cause “chromatin damage,” has much stronger input in their cytotoxicity than their ability to cause DNA damage. Another important observation is that there is a poor correlation between compound effects on chromatin in cells (c-trapping and histone eviction) and their effects on nucleosome in cell-free conditions (Fig. 6 and Supplementary Fig. S22), most probably due to their different abilities to reach and bind genomic DNA in cells.

Our study suggests that many known and some novel molecules exert their biological activity via destabilization of chromatin, i.e., “chromatin damage.” Effects of different small molecules on chromatin in cells were previously noticed in multiple studies (9, 25, 26). We made an attempt to understand why some compounds do this and some not, even if they belong to the similar functional groups, e.g., TOPO inhibitors. We cannot extrapolate our finding to all small molecules, but our data suggest that compounds able to bind genomic DNA in cells cause “chromatin damage.” Importantly, ability of a compound to bind naked or nucleosomal DNA in cell-free conditions is not directly translated into their ability to bind genomic DNA in cells. Many direct DNA binders do not reach nuclear DNA in live cells, e.g., propidium iodide. Some are distributed between nuclei and other organelles (e.g., lysosomes), or emitted from cells by multidrug transporters, and therefore, amount of a drug bound to DNA varies. Unfortunately, there are limited possibilities to measure binding of small molecule to DNA in cells. If on a larger cohort of compounds with well-established ability to bind nuclear DNA, it will be confirmed that they cause c-trapping and histone eviction, then these relatively simple assays may serve as a surrogate markers of compound binding to genomic DNA in cells.

We propose that the binding of almost any small molecules to DNA interferes with nucleosome stability in several ways. Intercalators change the shape of the DNA helix, make DNA less flexible, and disrupt the contacts of amino acids and nucleotides important for nucleosome stability. In addition, many of these compounds are positively charged and therefore their binding neutralizes negative charge of DNA, which plays important role in nucleosome stability. Situation with minor groove binder is less clear: although Hoechst 33342 caused c-trapping in cells, it disassembled nucleosomes neither in vitro nor in cells. It is known from crystal structure of nucleosome that the strongest contacts are formed between the octamer and minor groove of DNA at several regions of nucleosome (37). We can speculate that although Hoechst 33342 may have difficulties in getting to the minor groove at these regions, however if it gets there, for example, as a result of nucleosome breathing, the strong contact of core with DNA will not be restored. Such nucleosome may not be fully disassembled, because other histone/DNA contacts are preserved, but it should have part of DNA uncoiled, what facilitates access of FACT to nucleosome and obstructs histone H1 binding. These phenomena we observed in cells treated with Hoechst 33342.

However, some of these compounds are less capable of binding DNA in cells in the context of chromatin for reasons currently unknown. Because the structure of mammalian chromatin is very complex and cannot be easily visualized at a molecular level, some intermediate system, such as topologically constrained nucleosomal arrays in vitro, may be required to gain an understanding of this phenomenon.

Curaxins happened to be the strongest inducers of “chromatin damage” in cells among tested compounds most probably due to the combination of structural and biological characteristics. First, they are extremely efficient in reaching nuclear DNA because they are not substrate of multidrug transporters. They bind DNA with high affinity (14, 15), yet being relatively small comparing with anthraquinones. The small size may facilitate binding to bent nucleosomal DNA. However, too small size may make intercalation unstable, because compound may “enter” and “exit” DNA too easily without prolonged residence. That is why many planar carbazole compounds are weak DNA binders, i.e., form complex with DNA only at high concentrations. Presence of “ears” in curaxins, two symmetrical carbazole side chains protruding into major groove, is critical for biological activity of curaxins most probably due to anchoring of them between bases. Finally, relatively small and flexible “tail”—side chain filling minor groove with positively charged nitrogen—adds to the stability of binding.

Although all these speculations are based on computer modeling and structural studies of different compounds binding to naked and nucleosomal DNA are needed, we believe that the difference between compounds in their potency to cause chromatin damage and c-trapping may be explained by their binding to different genomic regions in cells. Therefore, we propose the following model. Curaxins can bind both naked and nucleosomal DNAs, which leads to destabilization of chromatin genome wide and occurrence of opened accessible to FACT nucleosomes at multitude of locations, as well as regions of DNA without nucleosomes, undergoing transition to Z-DNA. The latter may be facilitated by underlying sequence of DNA or chromatin structure. Alternatively, regions of nucleosome loss may occur randomly, and just those that are composed of tandem dinucleotide purine/pyrimidine repeats have higher probability of transition to Z-DNA. It is likely that FACT binding to the new regions is stronger than to the regions that it binds in untreated cells, which leads to redistribution of FACT. Alternative explanation is that FACT enrichment at transcribed regions is reduced because curaxins create many more sites for FACT binding, and this leads to the “dilution” of FACT at transcribed regions. However, previously we showed that much higher salt concentrations are needed to extract FACT from chromatin in curaxin-treated cells than in control cells (15), suggesting that affinity of binding is increased upon curaxin treatment.

Doxorubicin, mitoxantrone, and aclacinomycin A share many properties and induce similar pattern of FACT redistribution, which suggests that they have similar effect on chromatin in cells. We propose that these compounds have more difficulties getting to nuclear DNA than curaxins, and therefore much higher concentrations of these compounds are needed to cause effects similar to curaxins in cells, although in cell-free conditions, they cause stronger nucleosome destabilization. Based on literature, they also prefer binding linker versus nucleosomal DNA (9). This should lead to the preferential binding to promoters or coding regions of highly transcribed genes. It was shown by Yang and colleagues that doxorubicin causes the strongest histone eviction from promoter regions, which are also depleted in nucleosomes (25). Even at these regions, they likely do not cause complete nucleosome disassembly, because we do not observe significant eviction of core histones, and experiments with FACT mutants demonstrated that anthraquinones induce only n-trapping, or binding of FACT to destabilized nucleosomes, but no z-trapping or FACT binding to naked DNA.

Based on this, we propose the following model for these molecules. It is known that in untreated cells, passage of RNA polymerase partially uncoils nucleosomal DNA, making core available for FACT binding, whereas recoiling of DNA releases FACT from chromatin. Therefore, in untreated cells, FACT binding to these regions is temporal. Anthraquinones via binding to DNA stabilize uncoiled state of nucleosome, creating permanent sites for FACT binding and therefore increase total amount of binding sites and residence time of FACT at these sites, leading to FACT enrichment.

In conclusion, our findings confirm the previously observed effects of anthracyclines on chromatin and expand these observations toward the proposition that any DNA-binding compound should destabilize nucleosomes in vitro and in cells if they can bind DNA in the context of chromatin and accumulate in the nucleus. FACT is sensitive to a plethora of perturbations of the nucleosomes induced by the binding of the small molecule to DNA, including displacement of the linker histone, eviction of the core histones, and accumulation of negative supercoiling. Most importantly, we suspect that the “chromatin-damaging” effect of small molecules plays a more important role in the toxicity of DNA-binding molecules than their DNA-damaging activity. We do not question the role of DNA damage in the cytotoxic activity of many other molecules; however, “chromatin damage” caused by direct DNA binders may outweigh DNA damage in its influence on cell functioning and viability. The exact mechanism of cell death upon “chromatin damage” still needs to be established.

K.V. Gurova is a consultant/advisory board member for Incuron. No potential conflicts of interest were disclosed by the other authors.

Conception and design: E. Nesher, A. Safina, I. Koman, K.V. Gurova

Development of methodology: A. Safina, K.V. Gurova

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): E. Nesher, A. Safina, E.S. Wang, K.V. Gurova

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): E. Nesher, A. Safina, E.S. Wang, J. Wang, K.V. Gurova

Writing, review, and/or revision of the manuscript: E. Nesher, A. Safina, E.S. Wang, I. Koman, J. Wang, K.V. Gurova

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): A. Safina, I. Aljahdali, S. Portwood, I. Koman, J. Wang, K.V. Gurova

Study supervision: I. Koman, K.V. Gurova

We would like to thank Bruce Specht for the administrative help and Catherine Burkhart from Burkhart Document Solutions for critical review and editing of the article. This work was supported by Incuron LLC (to K.V. Gurova); by National Cancer Institute (R01CA197967 to K.V. Gurova and P30CA016056 to Roswell Park Cancer Center); and by Komen Foundation award (CCR13264604 to K.V. Gurova).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.