Doxorubicin impacts chromatin binding of HMGB1, Histone H1 and retinoic acid receptor

Doxorubicin (Dox), a widely used anticancer DNA-binding drug, affects chromatin in multiple ways, and these effects contribute to both its efficacy and its dose-limiting side effects, especially cardiotoxicity. Here, we studied the effects of Dox on the chromatin binding of the architectural proteins high mobility group B1 (HMGB1) and the linker histone H1, and the transcription factor retinoic acid receptor (RARα) by fluorescence recovery after photobleaching (FRAP) and fluorescence correlation spectroscopy (FCS) in live cells. At lower doses, Dox increased the binding of HMGB1 to DNA while decreasing the binding of the linker histone H1. At higher doses that correspond to the peak plasma concentrations achieved during chemotherapy, Dox reduced the binding of HMGB1 as well. This biphasic effect is interpreted in terms of a hierarchy of competition between the ligands involved and Dox-induced local conformational changes of nucleosome-free DNA. Combined, FRAP and FCS mobility data suggest that Dox decreases the overall binding of RARα to DNA, an effect that was only partially overcome by agonist binding. The intertwined interactions described are likely to contribute to both the effects and side effects of Dox.


Results
The anticancer drug Doxorubicin impacts HMGB1 distribution and mobility in the nuclei of live cells. Treatment of U2OS 2FP cells stably expressing GFP-tagged HMGB1 and RFP-tagged H2B with Dox caused a loss of GFP-HMGB1 from nucleoli while its distribution at the chromatin became more structured (Fig. 1a). However, at a Dox concentration of > 9 µM, the structured distribution in the chromatin was lost and the localization of GFP-HMGB1 became more diffuse. This biphasic effect on chromatin distribution was also reflected by the mobility of GFP-HMGB1 measured by point FRAP. The recovery time increased with increasing Dox concentration peaking at about 50 ms for samples treated with 4.5 µM and then declined at higher Dox concentrations ( Fig. 1b and Fig. S1). Similar observations were made following treatment with another intercalating drug, ethidium bromide (EBr) (Fig. S2).
Drug intercalation to DNA increases the base pair rise while reducing the helix twist by an angle dependent on the intercalator molecule. This decrease in helix twist translates into an overall reduction in DNA twist, which is compensated by an increase in writhe within the chromatin loops 23 . To elucidate the possible contribution of superhelicity to the recovery profiles generated above ( Fig. 1b and Fig. S1), we induced topological relaxation by nicking agents.
DNA nicking had no effect on HMGB1 binding in live cells. In view of the well-known winding effect of intercalators 26,27 (and refs therein), we tested the possible role of internucleosomal superhelicity in determining HMGB1 binding in vivo. DNA supercoiling was relaxed by treating live U2OS 2FP cells with hydrogen peroxide (H 2 O 2 ), bleomycin or X-ray irradiation, and GFP-HMGB1 binding to DNA was evaluated by point FRAP. Given the short time interval between nicking and measurement, we expected that the breaks would still be unrepaired, or even if they were, the original levels of internucleosomal superhelicity would not have been re-established. Thus, if HMGB1 binding were sensitive to supercoiling, relaxation would affect its binding. Strik-Scientific Reports | (2022) 12:8087 | https://doi.org/10.1038/s41598-022-11994-z www.nature.com/scientificreports/ ingly, none of these agents caused a change in FRAP recovery rate (Fig. 2), suggesting that intercalators influence HMGB1 binding directly rather than indirectly by changing twist-writhe partitioning.
The lack of effect of DNA supercoiling on HMGB1 binding in live cells was surprising since there was a slight but clear difference between the topological forms of plasmid DNA in their ability to bind recombinant HMGB1 (rHMGB1), when all the forms were simultaneously present (Fig. S3). As seen on the gel image, the migration of supercoiled plasmid DNA was retarded by as low as 60:1 rHMGB1-to-plasmid molar ratio and this retardation became more pronounced with increasing amounts of rHMGB1. Migration of linear DNA was not affected by up to a somewhat higher (100:1) protein-to-plasmid ratio, indicating a slightly lower rHMGB1-binding affinity for linear as compared to supercoiled DNA.
Dox decreases the binding of HMGB1 to supercoiled plasmid DNA. The diffusion constant (D) of rHMGB1 in solution was 73 µm 2 /s as determined by FCS. In the presence of native plasmid DNA, two diffusing components were observed: a fast component corresponding to freely diffusing rHMGB1 with D ≈ 76 µm 2 /s and a slow component with D ≈ 4 µm 2 /s, which was interpreted to be DNA-bound HMGB1 based on the fact that the diffusion constant of Sybr Gold stained native plasmid DNA was ≈ 3 µm 2 /s (Fig. 3a). Addition of Dox The graph shows one representative experiment of three repeats. In this as well as in the other figures, box-and-whiskers plots represent 10th, 25th, 50th, 75th, and 90th percentiles; + , mean value. One-way ANOVA with post hoc Dunnett's test was used to calculate significance compared to 0 Dox (**p < 0.01, ****p < 0.0001). Data were analyzed using GraphPad Prism 8.01.  . 4a). This effect was accompanied by an increased mobility as measured by strip FRAP (Fig. 4b). After 2 h of Dox treatment, the nucleolar component disappeared and there was a generalized loss of H1c-GFP from the cell (Fig. 4a). Similarly, EBr caused displacement of H1c-GFP from chromatin (Fig. S4). The displacement of histone H1 from DNA likely increases the number of available HMGB1 binding sites in the genomic DNA, thus explaining its recruitment to chromatin and the slower recovery rates in the first phase of the bimodal response ( Fig. 1b and Fig. S2).

Scientific
Dox affects the binding of RARα to DNA. To assess if intercalators affect the DNA binding of TFs, we extended our studies to the nuclear receptor RARα. RARα binds to DNA in a sequence-specific manner via its zinc-finger motif not involving intercalating amino acids. Strip-FRAP was used to analyse the average mobility of RARα over distances of several micrometers. Our FRAP data could be fitted to a biexponential model function (see Eqs. 2-5; for FRAP curves see Fig. S5). Figure 5 shows the diffusion behaviour in terms of the weighted average of the recovery times, τ average ; the detailed behaviour of the individual components is shown in Fig. S6. With increasing doses of Dox, the overall mobility of RARα increased; i.e., its DNA-binding decreased as suggested by the reduced τ average . At 1.125 µM Dox concentration it was only slightly reduced, whereas at 4.5 µM it decreased from 1.43 to 0.94 s. A further reduction in RARα binding was observed at 18 µM concentration, to 0.67 s. The mobile pool (Fig. S6d) decreased slightly, from 98% for the control to ~ 93% at 18 µM Dox. Treatment with the specific RARα agonist AM580 (used at 0.1 µM concentration) ameliorated the reduction in RAR binding up to 4.5 µM Dox concentration, while τ average was reduced to 0.69 s upon treatment with 18 µM (Fig. 5, right). For further details see Fig. S6.
To assess if the observed increase in RAR mobility could be due to a change of microviscosity in the nucleus, we applied FRAP to the EGFP dimer, an inert molecule not binding to DNA. Its recovery time was not affected by Dox treatment (see Fig. S7) indicating that microviscosity was not significantly altered. Thus, the FRAP-derived increase of mobility of EGFP-RARα is indeed due to its decreased chromatin binding.
To reveal the subcellular distribution of the chromatin and EGFP-RARα, confocal images were taken. The distribution of EGFP-RARα exhibited only mild spatial variations, whereas that of Dox was more heterogeneous displaying very bright and dark areas, irrespective of treatment with an agonist ligand (0.1 µM AM580) ( Fig. 6a-d).
The uneven distribution of Dox prompted us to probe the local mobility of EGFP-RARα with FCS, an approach capable of distinguishing ligand-specific RARα binding through the fraction of the slow component of the autocorrelation curves 18 . The slow, DNA-bound fraction of RARα in control and in ligand and/or Dox-treated

Discussion
Here we report that Dox intercalation into DNA in vivo affects HMGB1 binding to DNA in a bimodal fashion, i.e. increasing binding of the protein at lower concentrations, and causing a drastic reduction at concentrations higher than 9 µM (Fig. 1). The phase of increased HMGB1 binding is well explained by histone H1 displacement from DNA in view of the facts that both H1 and HMGB1 compete for the linker DNA near the nucleosome dyad 5 .
In line with this notion, histone H1 exhibits higher sensitivity to intercalator binding than HMGB1, i.e. H1 is readily displaced from chromatin by both Dox and EBr at drug concentrations where HMGB1 is still bound to DNA (compare Fig. 4 with Figs. 1 and Fig. S2). The reduction in HMGB1 binding to DNA at high Dox concentrations ( Fig. 1b) could be due to intercalator induced DNA distortion, or alternatively, to competition between Dox and HMGB1 for binding sites on DNA. Dox carries two DNA binding moieties, the anthraquinone moiety which intercalates between adjacent G-C base pair steps, and the amino sugar which is positioned in the DNA minor groove 32 . Intercalation of the anthraquinone moiety increases the base pair rise to 5.2 Å and reduces the helical twist at the site of intercalation. In vivo, unwinding of the DNA due to intercalation would be compensated for by positive writhing within the closed chromatin loops without change in the linking number, in line with the equation ΔLk = ΔTw + ΔWr. The level of the compensatory positive torsion would increase with the number of intercalated molecules. This intercalatorinduced change in torsion together with that induced by ongoing replication and transcription would trigger topoisomerase activity to resolve it, as reviewed in 33,34 . However, at high concentrations (> 9 µM), Dox would inhibit the binding of the topoisomerase to DNA (as reviewed in 35 ), thus leading to accumulation of positive writhe. Although some of the positive torsion may be annihilated by the negative torsion stored in nucleosomebound DNA following destabilization of nucleosomes 36,37 , this may reduce but not completely abrogate positive torsion. The positive writhe would thus hinder the binding of HMGB1, which involves intercalation 38 . This argument would be in line with the monotonous reduction in HMGB1 binding to covalently closed plasmid DNA in the presence of Dox (Fig. 3b). However, the lack of any detectable change in HMGB1 binding upon nicking treatments suggests that the contribution of the above mechanism to the decreased binding may not be  www.nature.com/scientificreports/ significant. Thus, local distortion of DNA conformation or competition of the two ligands for DNA are suggested to account for the results of Fig. 3b.
Regarding the latter type of interaction, the binding of Dox and HMGB1 to DNA overlap in two aspects: both involve intercalation into DNA as well as minor groove binding. HMGB1 possesses two DNA binding domains, box A and B, which are connected by a short linker to an intrinsically disordered, acidic C-terminal tail 39 . The A and B domains form an "L"-shape with a concave DNA binding surface. The binding of HMGB1 to DNA occurs through the DNA minor groove and induces a bend towards the major groove. Additionally, hydrophobic amino acids phenylalanine (Phe38) on box A, phenylalanine (Phe1103) and isoleucine (Ile122) on box B partially intercalate between base pairs following DNA binding, and unwind the DNA 38 . This overlap in the Dox and HMGB1 binding to DNA is bound to create grounds for competition.
Even at the highest concentration of EBr used (100 µM), HMGB1 binding to DNA exceeded that of the control, as seen in the microscopy images and also through the point FRAP profiles (Fig. S2). This is in sharp contrast to Dox, which at drug concentrations > 9 µM reduces HMGB1 binding to DNA. This effect may be attributed to the lower uptake of EBr by live cells due to its positive charge, which means that the amount of dye intercalated in DNA is much lower than in the case of Dox. Doxorubicin is known to accumulate in cells attaining a higher intracellular concentration as compared to the extracellular milieu. Another factor in the higher efficiency of Dox to suppress HMGB1 binding may be the presence of the amino sugar positioned in the DNA minor groove.
Relaxing DNA writhe in chromatin loops by nicking via X-ray irradiation, treatment with H 2 O 2 or bleomycin did not yield any observable effect on the binding of HMGB1 in vivo (Fig. 3). This is in contrast with the well documented preference of HMGB1 binding to supercoiled over relaxed DNA in vitro ( Fig. S3; see also 38 ). At the dosage used in these experiments, X-ray irradiation is estimated to cause 6000-50,000, while H 2 O 2 provokes 15,000-36,000 single strand breaks per nucleus 40,41 (see also Fig. S11). Even if a fraction of single strand breaks may be repaired within minutes 42 , many of the breaks would remain unrepaired given the short time interval between nicking and measurement. Moreover, it appears unlikely that the initial level of supercoiling could be restored by RNA and DNA polymerization according to the twin supercoil model 43 in these circumstances. The fact that HMGB1 binding in vivo was not affected by DNA nicking implies that the protein binds similarly to relaxed and supercoiled DNA when it is the only conformation available. Furthermore, the overall negative supercoiling of internucleosomal DNA may be much smaller than that of the supercoiled plasmid. Therefore, the topological preference of the protein emphasized earlier based on in vitro data 38 is probably irrelevant in the cellular context.
Other groups have observed nuclear to cytoplasmic translocation or extracellular secretion of HMGB1 from immune cells following ionizing radiation or H 2 O 2 treatment, at doses lower than those used here 44,45 . This, however, occurred 3-24 h after exposure, i.e. after most of the single strand breaks would have been repaired and thus cannot be directly related to changes in superhelicity or H1 binding.
The evidence presented here suggests that supercoiling may not affect HMGB1 binding in vivo, or any effect of supercoiling may be overshadowed by interactions involving the chromatin environment, as follows. In the context of the general rules governing ligand binding to chromatin, although the nucleosome-constrained topology and the interrelated structural features of the DNA pose a powerful obstacle of ligand access to DNA 23 , binding to the flexible internucleosomal DNA regions is, in general, not determined by DNA topology; here competition among the ligands or distortion of the DNA structure around the bound anthracyclin seem the dominant controlling factors. The concentration dependent influence of Dox on HMGB1 binding has not been recognized before and may contribute to the effects and side-effects of this medically relevant anthracycline.
We also analyzed how Dox treatment affects the DNA binding of a sequence-specific TF, RARα. This TF is unlikely to have an effect on the balance of H1 and HMGB1 genome-wide, i.e. its effects must be restricted to the regulatory regions of RAR responsive genes. On the other hand, upon ligand dependent RAR binding, HMGB1 may be recruited at these sites while H1 is displaced 46 . This exchange would be facilitated by Dox at low drug concentrations where it evicts H1 without affecting HMGB1 binding, leading perhaps to augmented RAR binding. However, such an effect, if it occurred, was apparently overcome by the decrease in DNA binding, seen also in the absence of the ligand (Fig. 5).
We measured the mobility of EGFP-RARα on a distance scale of a few micrometers, averaged for larger areas, with FRAP (Fig. 5, Fig. S6), and on a shorter distance scale of a few hundred nanometers, locally, with FCS ( Fig. 6e, Fig. S8). The FRAP recovery curves likely reflect the extent of overall DNA binding, while being also influenced by the diffusional constraints. The latter may be altered upon Dox treatment that is known to elicit gross morphological changes in the nucleus due to histone eviction and consequential aggregation 36 . With both methods we could detect a fast and a slow component, and with FRAP an additional fraction appearing as "immobile" over the distance scale of FRAP. The FRAP-detected fast and slow components refer to molecules that leave the few-micrometer wide ROI during the time course of the measurement; thus, both are diffusible and even the slow component can only be transiently bound to DNA. In view of our earlier FCS studies in this system [17][18][19][20]47 , the fast FCS component may likely be identified with the fast component of FRAP, which probably represents the freely diffusing RARα species or what is bound to DNA for very short dwell times, ≈ 1 ms according to our FCS fits (Fig. S10). The slow FCS component likely corresponds to transiently DNA-bound RARα (dwell time in the order of several hundred milliseconds) as well as stably bound RARα moving slowly together with the chromatin. The slow FCS component is likely a mixture of the slow and "immobile" populations detected by FRAP. Similar slow diffusion coefficients were detected by FCS for other DNA-binding proteins as well [47][48][49] .
Our most noteworthy observation via FRAP was that Dox increased the average mobility of RARα (Fig. 5). This increase was confirmed to be due to shorter dwell times spent DNA-bound rather than to a change of microviscosity, since the diffusion coefficient of the EGFP dimer was not affected (Fig. S7). On the other hand, Dox slightly enhanced the immobile fraction (decreased the mobile fraction) in a dose-dependent manner (Fig. S6d). This is in line with the decrease of the FCS-determined slow diffusion coefficient attributed to DNA-bound  (Fig. S8b). Thus, Dox seems to have a dual effect on the DNA-binding of RARα. First, there is a prominent Dox-induced decrease of overall RARα binding to the DNA, and second, a much smaller fraction of RARα displays decreased mobility. This latter minor component may be due to a stronger DNA-binding of the TF to the dehistonized DNA visible in the Dox-reorganized nuclei 36 . Previously, we have shown by FCS, FCCS and FRET that the RAR agonist AM580 augmented heterodimerization of RARα with RXRα, increased the proportion of the slow fraction and enhanced its DNA-binding [18][19][20] . Here, treatment with saturating concentrations (0.1 μM) of AM580 partly counteracted the effect of Dox on the average mobility of RARα; i.e., the reduction of the average FRAP recovery time was mitigated in the presence of the ligand at a Dox concentration of up to 4.5 µM, which is within the range of the therapeutic doses 50 . Our FCS measurements reveal that Dox, used at a concentration easily reached in the tissues during chemotherapy 37 , overcomes the ligand-induced increment of the slow fraction, while not affecting it in the absence of the ligand (Fig. 6e). Thus, Dox decreases ligand-independent as well as ligand-dependent RARα binding, based on FRAP and FCS data, respectively.
The reduction in binding of RARα following Dox treatment may suggest yet another mechanism by which Dox may exert its cytotoxicity. As the binding of RARα to DNA does not involve intercalation, and there was no effect of nicking on RARα binding (Fig. S9), the reduction may be explained by either direct competition between Dox and RAR for the binding sites on the DNA or the drug's effect on its structural features. For example, Dox intercalation increases the stiffness of the DNA 51 and hence may decrease DNA deformability that is required for optimal TF binding 15 .
Despite the high efficacy of Dox in treating malignancies, its therapeutical use is limited by cardiotoxicity 52 . Several studies have linked Dox induced cardiotoxicity (DIC) to topoisomerase poisoning, while others emphasize the role of ROS production 24 . RARγ was recently shown to bind to the Top2β gene at a retinoic acid response element (RARE) positioned downstream of the transcription start site, thereby repressing gene transcription in the presence of ATRA. However, in cells expressing the nonsynonymous SNP variant RARγ-S427L, this transrepression is compromised, thus aggravating DIC 21,25 . Indeed, correction of this RARγ-S427L SNP by genetic engineering reduced the cytotoxicity of Dox 53 . Based on these findings, we propose that Dox inhibition of RAR binding to chromatin may contribute to cardiotoxicity. We speculate that a separate, rather than combined, administration of ATRA and Dox could mitigate DIC, while not affecting Dox cytotoxicity in cancer cells where the role of RAR signalling in Top2β control may not be relevant or tight.
In summary, Dox, at ≤ 4.5 µM concentration, augments HMGB1 binding by causing H1 eviction, while strongly inhibiting HMGB1 binding at higher doses also reached in blood plasma levels during chemotherapy. In addition, Dox reduces RARα binding, thus affecting a TF that contributes to the pathogenesis of DIC.

Materials and methods
Cell culture. HeLa cells stably expressing GFP-tagged histone H1 (HeLa H1c-GFP ) and U2OS cells stably expressing GFP-tagged HMGB1 and RFP-tagged H2B (U2OS 2FP ), (from the labs of Profs. Hiroshi Kimura, Tokyo and Guido Kroemer, Paris, respectively) were maintained in DMEM medium (Gibco) supplemented with 10% FBS 1 × GlutaMAX, 100 U/ml penicillin, 100 µl/ml streptomycin and phenol red. In addition, media for U2OS 2FP cells was supplemented with G418 0.5 mg/mL for continuous selection of transformed cells. HeLa cells and HeLa cells stably expressing EGFP-RARα, (HeLa EGFP-RARα ) were maintained in RPMI supplemented with phenol red, 10% fetal calf serum (Sigma-Aldrich, Saint Louis, MO), 1 × GlutaMAX (Fisher Scientific, Tokyo, Japan), and 50 mg/l gentamycin (KARA, Novo Mesto, Slovenia). All cells were cultured at 37 °C in a humidified, 5% CO 2 incubator and passaged every two days. The expression of the tagged proteins in the transfected cell lines was ~ 20% of the levels of the respective native proteins in the case of H1c-GFP and HMGB1-GFP (data not shown), while the expression level of transfected EGFP-RARα was double of the endogenous level of RARα as determined by RT-QPCR 18 .
For microscopy experiments, HeLa or HeLa EGFP-RARα cells were seeded in 8-well chambered cover slips (ibidi GmbH, Gräfelfing, Germany) 48 h before the measurement and maintained in phenol red-free RPMI supplemented with 10% charcoal-stripped fetal calf serum (PAN-Biotech, Aidenbach, Germany). EBr for 1 h, bleomycin for 2 h, Hoechst 33,342 (10 µM) for 30 min or H 2 O 2 for 20 min at 37 °C in phenol redfree medium, in a volume of 300 µl/well. Irradiation with X-rays was done at room temperature at a distance of 57.6 cm from the window of a 6 MeV linear accelerator (Radiotherapy Department, University of Debrecen) while the cells were kept in phenol red-free cell culture medium (300 µl/well). AM580 was applied to the cells at 0.1 μM final concentration for 30 min at 37 °C before imaging.

Point FRAP.
When it was made possible by the kinetics of recovery, point FRAP was used to better detect possible spatial heterogeneities. For measurement of GFP-HMGB1 mobility, point FRAP measurements were performed on an Olympus FluoView 1000 confocal microscope based on an inverted IX-81 stand with an UPla-nAPo 60 × NA 1.2 oil immersion objective. EGFP was excited by the 488 nm line of an Ar-ion laser, and emission was detected through a 500-520 nm band-pass filter by a PMT. The measurement for point FRAP data acquisition started with a confocal image of a cell (512 × 512-pixel, pixel size: 0.103 µm), followed by the selection of a laser spot at which the laser beam was focused. Before bleaching, 5120 pre-bleach pixels were collected with a Scientific Reports | (2022) 12:8087 | https://doi.org/10.1038/s41598-022-11994-z www.nature.com/scientificreports/ pixel dwell time of 10 μs (51.2 ms) followed by bleach period for 51.2 ms with 100% laser power 55.6 µW and then collecting 40,000 post-bleach pixels from the same spot for a total time of 297.59 ms. In order to change the laser power of the Ar-ion laser shorter than the pixel dwell time, a dedicated Lab-VIEW program was developed. The analogue output of a NI 7833 field programmable gate array (FPGA) card (National Instruments, Austin, TX) was fed into the laser power controller input pin of the acousto-optic tunable filter (AOTF, AA Opto electronic MOD.NC) of the laser combiner unit, allowing fast (~ 1 μs) voltage driven laser power switching by an FPGA card. The operating of the FPGA card was initiated by a TTL output of the trigger port of the FV1000 system, thus controlling the laser power and collecting the fluorescence data were synchronized.
The model function fitting of the recovery post-bleach data was performed by a custom-written Matlab (The Math Works, Natick, MA) program. Data were fitted assuming a one-component exponential recovery with the following equation: where I (t) is intensity at a given time point; t, time; I 0 , amplitude of the intensity; τ, recovery time; I bg , background intensity.
Strip FRAP. When point FRAP recoveries were too fast to provide accurate mobility data, a strip FRAP protocol was followed. These FRAP measurements for histone H1-GFP were carried out on an Olympus FluoView 1000 confocal microscope as described previously 17 with a minor modification; 10 images (512 × 512 pixels, 4 µs/ pixel), were collected before bleaching, followed by 15 bleaching images in a ROI (25 × 12 pixels, 20 µs/pixel, 56 µW) and 80 post-bleach images.
FRAP measurements for EGFP-RARα and EGFP dimers were also carried out in a similar setting with some modifications. In brief, measurements were performed on a Zeiss LSM 880 (Carl Zeiss, Jena, Germany) confocal microscope using a 40 ×, 1.2 NA water immersion objective. The 488 nm laser line was used to excite EGFP with a laser power of 2 μW at the objective (10%), and emission was detected through a 493 to 529 nm band pass filter. For quantitative analysis, a 256 × 256-pixel area was selected and scanned with an open pinhole (5.56 Airy units) and 10 × zoom (pixel size: 0.08 µm), with a pixel dwell time of 1.33 μs. A 405-nm laser was used to bleach the EGFP molecules at a selected strip-shaped region of interest (FRAP ROI) having an area of 140 × 10 pixels, a laser power of 20 μW at the objective, and a pixel dwell time of 8.24 μs. Before bleaching, 10 images were collected at a repetition rate of 204.8 ms/frame followed by one bleach period at the FRAP ROI, and then collecting 189 post-bleach images for a total time of 42 s. To standardize the geometry of the measurement, the scanned field was rotated to make the long axis of the selected nucleus vertically oriented in the image, and the strip shaped FRAP ROI (bleached area) was positioned horizontally at one third of the vertical extension of the nucleus, avoiding nucleoli (Fig. S12).
Images were processed using the open-source FIJI distribution of ImageJ (version 2.0.0-rc-69/1.52i) to acquire the fluorescence intensity recovery curves required for FRAP analysis. The width of the FRAP ROI was cropped to match the width of the nucleus. Another ROI contouring the whole nucleus but excluding the nucleoli was made to calculate the total fluorescence intensity of the nucleus. A third ROI outside the cell was drawn to calculate the intensity of the background. Following the double normalization method 54 the intensity in the FRAP ROI during the recovery period was normalized to its pre-bleach value I ROI (0), and corrected for acquisition bleaching of the whole nucleus using the following equation: where I ROI (t) is intensity of the FRAP ROI at a given time during the recovery, I ROI (0) is average intensity of the ROI before bleaching, I total (t) is the intensity of the whole nucleus at a given time during recovery, I total (0) is the average intensity of the whole nucleus before bleaching over ten frames, and I B is average intensity of the background. Using the Prism version 8.4.0 software, a two-component exponential curve (Eq. 3) was fitted to the normalized recovery curve of the EGFP-RAR while a one-component exponential to the EGFP dimer: The fit yielded the τ fast and τ slow recovery times of the fast and the slow components, their fractions r fast and r slow adding up to 1, the plateau I ∞ at infinite time and the fitted intensity value right after bleaching, I min . The mobile fraction was determined as: The average recovery time was calculated as a weighted average of the fast and slow components:  where N is the average number of fluorescent molecules in the detection volume, T is the fraction of molecules in the triplet state, τ tr is the triplet correlation time. The rate of diffusion is characterized by the diffusion time, τ d , which is the average time that a molecule spends in the illuminated volume. τ D1 and τ D2 are the diffusion times of the fast and slow components, ρ is the fraction of the first component and 1−ρ is the fraction of the second component. Diffusion coefficients (D) of the fast and slow components were determined from the following equation: where ω xy is the lateral e −2 radius of the detection volume. Expression and purification of rHMGB1. The pET19b expression vector carrying the full length human HMGB1 gene with C23S, C45S, C106S and E204C mutations (from Jennifer Kugel's lab, University of Colorado Boulder, USA) was transformed into Rossetta (DE3)pLysS and grown overnight at 37 °C incubator on LB agar plates containing 100 µg/ml ampicillin and 37.4 µg/ml chloramphenicol. A single transformed colony from the plate was used to inoculate 5 ml LB with antibiotic. The small culture was incubated overnight at 37 °C with shaking and was used to inoculate 100 ml of LB containing D glucose (100 ml LB, 2.4 mM NaOH, 0.2 g D-glucose) and antibiotic in a 500 ml culture flask. The culture was grown at 37 °C with shaking until an OD600 of 0.5 was attained upon which expression was induced using 0.5 mM IPTG and the cells were grown for a further 3 h. The culture was transferred into 50 ml centrifuge tubes and pelleted at 5000 RPM for 15 min. The supernatant was discarded leaving a small volume to resuspend the pellet in. The cells were then transferred into a 1.5 ml Eppendorf tube and pelleted again. The supernatant was discarded, the pellet flash-frozen and stored at − 80 °C.
To extract the protein, 2 ml of lysis buffer (20 mM Tris pH 7.9, 500 ml NaCl, 10% glycerol, 5 mM imidazole, 5 mM beta mercaptoethanol (BME) and 0.2 mM PMSF) was added to a thawed pellet from 100 ml culture, the pellet was resuspended then the cells sonicated 5 × on ice (30 s on, 30 s off). The lysate was then centrifuged for 15 min at 4 °C at 15,000 RPM. The supernatant containing the protein was transferred into a packed Nickel column and then kept at 4 °C for 1 h with rocking motion. The supernatant was allowed to flow through, then the resin washed with 1 ml of lysis buffer followed by 3 ml of lysis buffer containing 50 mM imidazole. The protein was eluted using lysis buffer containing 250 mM imidazole. The column was stripped with lysis buffer supplemented with 1 M imidazole.
Following elution from the nickel column, the buffer was exchanged using a 10 K MWCO spin column and 50 ml degassed dialysis buffer (20 mM Tris pH 7.9, 50 mM KCL, 10% glycerol, 100 mM PMSF, 100 mM MgCl 2 and 1 mM DTT). After buffer exchange the protein sample in about 250 µl was centrifuged at 18,000 RPM for 30 min at 4 °C and the supernatant transferred to a dsDNA cellulose column and incubated for 1 h at 4 °C with rocking motion. The supernatant was allowed to flow through, the resin washed 3 × with 3 ml wash buffer (20 mM Tris pH 7.9, 50 mM NaCl, 10% glycerol, 5 mM MgCl 2 and 1 mM DTT) and the protein eluted using wash buffer with 500 mM NaCl. The column was stripped with wash buffer containing 1 M KCl. The protein was then desalted using 10 K MWCO spin column and degassed dialysis buffer as above without DTT. Purification was confirmed using a 12% SDS PAGE gel electrophoresed for 1 h at 150 V (Fig. S3a). Samples were centrifuged at 18,000 RPM for 30 min at 4 °C, aliquoted, flash frozen and stored at − 80 °C. Quantification was carried out using Lowry assay and known amounts of BSA controls. Measurements were also made on the Nanodrop.
Immunofluorescence labelling of rHMGB1. Labelling of rHMGB1 with Alexa 647 C2 maleimide (Thermo scientific) was done following the manufacturer's instructions. To reduce the disulphide bonds in the protein, 750 µg of rHMGB1 in degassed buffer A (100 mM Tris pH 7.1, 50 mM KCl, 10% glycerol, 0.2 mM PMSF and 5 mM MgCl 2 ) was mixed with DTT to a final concentration of 10 mM and incubated at 4 °C for 2 h. DTT was removed using a spin column and 20 ml buffer A. 1.5 µl of 8 mM Alexa 647 C2 maleimide was added to the sample and incubated in the dark at 4 °C overnight. Unbound dye was removed using 10 K MWCO microfuge www.nature.com/scientificreports/ spin column and sample washed with column wash buffer (20 mM Tris pH 7.9, 50 mM KCl and 5 mM MgCl 2 ). Samples were quantified by measuring absorbance at 280 nm and compared to a standard curve prepared from known amounts of BSA. Glycerol was added to a final concentration of 10% and samples stored at either − 20 °C for short term or − 80 °C for long-term storage.

Electrophoretic mobility shift assay (EMSA).
To determine the binding affinity of the labelled rHMGB1 to various topological forms of plasmid DNA, gel electrophoresis was carried out. Native pEGFP C3 plasmid DNA was either nicked or linearized using Nb.Mva1269I and ECoRI (Thermo scientific), respectively, using the manufacturer's instructions. The DNA samples were then cleaned using gel cleaning kit (QIAGEN) following the manufacturer's instructions. DNA was quantified by measuring the absorbance at 260 nm on the Nanodrop. An equal amount (0.5 µg) of linear, nicked and native plasmid DNA was mixed in 20 µl of protein binding buffer (50 mM NaCl, 20 mM Tris HCl pH7.5 and 0.2 mM EDTA). To this DNA mixture, varying amounts of rHMGB1 was added per sample as indicated on the gel (Fig. S3) and incubated on ice for 40 min on ice. The samples were then loaded to a 1% agarose gel in 0.5 × TBE and run for at 36 V for 15 h at 4 °C. The gel was stained with 0.5 µg/ml EBr and then imaged. The image was processed using Fiji ImageJ.
Statistical analysis. For creating plots and for statistical comparisons GraphPad Prism 8.01 was used. Boxand-whiskers plots represent 10th, 25th, 50th, 75th, and 90th percentiles; + , mean value. To compare averages of multiple data sets, ANOVA with Tukey's multiple comparison test was used.