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The linker histone chaperone Prothymosin α (PTMA) is essential for efficient DNA damage repair and the recruitment of PARP1

Epigenetics & Chromatin volume 18, Article number: 32 (2025) Cite this article

Abstract

Background

Mammalian cells have numerous DNA repair pathways to repair lesions generated by replication errors, metabolism, and exogenous agents. Cells can sense and respond to DNA damage within seconds, suggesting that there is a highly effective sensor of lesions although the mechanistic details are unclear. The DNA damage response in mammalian cells results in a localized transient de-condensation of chromatin, loss of linker histones and the recruitment of DNA repair proteins such as PARP1 and chromatin remodelers.

Results

Here we investigated the interactions between poly(ADP-ribose) polymerase-1 (PARP1), the linker histone H1.0 and linker histone chaperone Prothymosin α (PTMA). Using H1.0 tagged with a photoconvertible fluorescent protein, we observed a significant increase in the initial rate of exit of H1.0 from regions of chromatin containing microirradiation-induced DNA lesions. Surprisingly, this was also seen in Parp1−/− cells but not in stable cell lines with homozygous null mutations in the PTMA gene (Ptma−/−). The recruitment of PARP1 to damaged DNA was inhibited by overexpression of a mutant of H1.0 with a tighter chromatin-binding affinity or by reduced expression of PTMA. Relative to the wild type, Ptma−/− cell lines displayed increased sensitivity to DNA-damaging agents.

Conclusion

We suggest that DNA damage alters the interaction of H1.0 with the nucleosome to allow the chaperone PTMA to bind and promote release of linker histones thereby initiating the local chromatin de-condensation necessary for the efficient recruitment of repair proteins such as PARP1. In this context linker histones may serve as in situ “sensors” of DNA damage.

Background

An efficient DNA damage response (DDR) is essential to mammalian cell growth, proliferation, and survival [1]. Each day, more than 104 -105 DNA lesions per cell are generated due to replication errors, metabolism, and UV exposure. To combat such high levels of DNA damage, the cell has numerous evolutionarily conserved pathways in place [2]. Cells can sense and respond to DNA damage within seconds, suggesting that there is a highly effective mechanistic sensor of lesions. A major unresolved question in the DNA repair field is how the DDR specifically recognizes damaged DNA within the 3 × 109 base pairs of the human genome, the vast majority of which exists in a compacted chromatin state [3,4,5]. One of the earliest detectable events following DNA damage in mammalian cells is a localized transient de-condensation of chromatin in the vicinity of the lesion [6, 7]. This is presumed to be necessary to facilitate the access of repair proteins to the underlying damaged DNA [8,9,10]. The cellular mechanism that senses DNA damage and triggers the initial chromatin de-condensation is not well understood. This “sensor” function is often assigned to the proteins that are most rapidly recruited to damage sites.

The nucleosome is the fundamental repeating unit of eukaryotic chromatin [11, 12]. The nucleosome core consists of an octamer of two molecules each of the four core histones around which is wrapped 147 bp of DNA [13]. In eukaryotes, one molecule of the linker or H1 class of histone is bound to nucleosomal DNA and also associates with the linker DNA between adjacent nucleosomes [14,15,16,17]. Photobleaching techniques demonstrated that linker histones interact dynamically with chromatin in living cells [18,19,20]. Most H1 molecules are continuously exchanged between chromatin binding sites with a mean residency time of approximately one minute. As H1 drives the formation and stabilization of the compacted form of chromatin associated with most of the DNA of interphase cells [21,22,23], the dissociation of H1 results in a localized transient chromatin de-condensation and provides a window of opportunity for other DNA-binding factors to access the DNA [24, 25]. We, and others have observed that linker histones are depleted from chromatin in the vicinity of damaged DNA [26,27,28,29].

Poly(ADP-ribose) polymerase-1 (PARP1) has been proposed to be a major sensor of DNA damage due to its abundance, involvement in multiple DNA repair pathways, rapid recruitment to damaged DNA, and ability to bind to the ends of damaged DNA [9, 10, 30,31,32,33,34,35]. In response to DNA damage, PARP1 and the associated co-factor Histone Parylation Factor 1 (HPF1) catalyze addition of polyADP-ribose to serine residues of chromatin proteins, especially histones [36,37,38]. This is thought to create a scaffold for the recruitment of additional chromatin remodelers and repair factors to establish an effective repair complex [10, 39,40,41,42,43,44]. Although PARP1 is an abundant protein, it is not clear how, even acting as a diffusion limited free moving protein in the nucleus, it would be able to scan the entire genome within the time frame of the initial response prompting the proposal that it searches DNA via intersegment transfer or ‘monkey bar’ mechanism [45, 46].

Interestingly, in undamaged cells, PARP1 and H1 bind to overlapping sites on the nucleosome dyad in a mutually exclusive manner, suggesting that they compete for binding sites [47]. An enrichment of PARP1 is associated with active transcription and H1 with repression [48]. Although interactions between H1 and PARP1 in modulating chromatin structure and transcriptional outcomes are independent of PARP1 catalytic activity, linker histones are robustly ADP-ribosylated in response to DNA damage. As the recruitment of PARP1 and the depletion of H1 occur on similar time scales, it has been proposed that PARP1, either through direct competition and/or via ADP-ribosylation promotes the depletion of H1 to facilitate chromatin de-condensation upon DNA damage [28, 49].

Prothymosin α (PTMA) is a small (12.5 kd), unstructured, highly acidic (pI = 3.5) protein ubiquitously expressed in most mammalian tissues [50]. PTMA has been reported to contribute to an astonishing number of normal and aberrant cellular processes including apoptosis [51], the immune response [52], cardiac regeneration [53] and restriction of infectious HIV-1 production [54]. Elevated levels of PTMA correlate with resistance to chemotherapy and poor clinical outcomes in many types of cancer [55,56,57,58]. We previously presented evidence that PTMA functions as a linker histone chaperone to facilitate the release and/or deposition of H1 in chromatin [59].

Here we explore the relationship between PARP1, H1, and PTMA in the early events of DNA damage repair. We have focused on the H1.0 variant in part because of its reported roles in cell proliferation, stem cell maintenance and tumor progression [60,61,62,63]. Surprisingly, we find that the initial depletion of H1.0 from chromatin upon DNA damage induced by microirradiation is mediated by a process that is PTMA-dependent but PARP1-independent. We suggest that H1.0 and perhaps other linker histones may act as a local in situ sensors, facilitating identification of damaged DNA by PARP1.

Results

Depletion of H1 from chromatin containing damaged DNA is PARP1-independent

In wild type mouse fibroblasts, DNA repair proteins, such as PARP1 and XRCC1 are rapidly recruited to sites of DNA damage induced by laser microirradiation with a 405-nm laser (Fig. 1A). We utilized CRISPR/Cas9 technology to generate a cell line containing homozygous null mutations in the Parp1 gene (Fig. 1B, Supplemental Fig. S1) and confirmed that linker histones are robustly ADP-ribosylated by PARP1 in response to DNA damage (Fig. 1C). We then stably transfected a plasmid expressing GFP-tagged H1.0 into these and wild type cells. As has been reported by others [27,28,29], we observed that, following microirradiation of wild type cells, linker histones are excluded from entering regions of chromatin containing lesions in both wild type and the Parp1−/− null cell lines (Fig. 1D). As the GFP chromophore is photobleached by the 405-nm laser, from the results shown in Fig. 1D, we can only conclude that unbleached H1.0 from distal chromatin cannot enter the region of damaged DNA. One explanation for this observation is that the recruitment of PARP1 to damaged DNA (Fig. 1A) physically prevents H1 from returning as PARP1 and H1 have been shown to compete for binding to nucleosomal DNA [47] This exclusion was also observed in the absence of PARP1 although other proteins, such as XRCC1, are also recruited under these conditions (Fig. 1A).

Fig. 1
figure 1

Exclusion of H1.0 from chromatin containing damaged DNA is PARP1-independent. (A) Wild type cells expressing GFP-tagged PARP1 or XRCC1 were microirradiated with a 405-nm laser. (B) Western blot of whole cell extracts from wild type (WT) and Parp1−/− cells with α-PARP1 antibody and α-myc (loading control). (C) Where indicated cells were treated with 1 mM H2O2 for 30 min prior to isolation of total histones. Linker histones were separated on 12% polyacrylamide gels and stained with Coomassie (left) or subjected to Western blotting with mADP-ribose antibody (right). (D) Wild type or Parp1−/− cells expressing GFP-tagged H1.0 were microirradiated in two separate regions with either the 405-nm or the 488-nm laser

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A further limitation imposed by the photobleaching of GFP is that it precludes determining if H1.0 is more rapidly exiting chromatin undergoing damage repair or is simply prevented from returning as part of the normal exchange process inherent to linker histones [20]. To address this, we used the photoconvertible pSMOrange protein [64]. The native form of this protein fluoresces in the orange region (Em λ = 565-nm). Upon brief exposure to 488-nm light, the chromophore undergoes a Stokes shift and fluoresces in the far-red region (Em λ = 662 nm). Importantly, the photoconverted form is not photobleached by 405-nm light. This provides two advantages over conventional photobleaching assays. It allows us to image in the far-red channel of our confocal system and specifically measure the kinetic behavior of the photoconverted species. In addition, we can obtain a better estimate of the initial rate of exit, expressed here as t25, the time for loss of 25% of the protein from the irradiated region. We first generated a wild type cell line stably transfected with two plasmids, one expressing GFP-tagged PARP1 (PARP1-GFP) and another expressing H1.0 tagged with pSMOrange (Or-H1.0). Two adjacent cells, were microirradiated with either 488-nm light to photoconvert the Or-H1.0 or sequentially with 488-nm and 405 nm light to photoconvert Or-H1.0 and damage DNA (Fig. 2A). Imaging of PARP1-GFP (Fig. 2A, upper panel) shows that PARP1 is only recruited to the region microirradiated with the 405-nm laser. Imaging of Or-H1.0 (Fig. 2A, lower panel) revealed that the initial rate of exit of H1.0 from damaged DNA is significantly faster than that from undamaged DNA (Fig. 2C and D). We then stably transfected Or-H1.0 into the Parp1−/− null cell line. Interestingly, the rate of exit of H1 from damaged DNA is significantly faster than that from undamaged DNA in the Parp1−/− line as well (Fig. 2B-D). We also created a Hpf1−/− cell line and observed that the exit of H1.0 from damaged DNA was also accelerated in these cells (Supplemental Fig. S2). These observations suggest that although PARP1 recruitment and H1.0 depletion occur with similar time scales, the processes may not necessarily be mechanistically linked, i.e. due to direct competition between PARP1 and H1.0 for binding to the nucleosome or due to serine ADP-ribosylation by PARP1/HPF1.

Fig. 2
figure 2

Release of H1.0 from chromatin containing damaged DNA is accelerated in a PARP1-independent manner. (A) Wild type cells were stably transfected with two plasmids, one expressing GFP-tagged PARP1 and another expressing H1.0 tagged with pSMOrange (Or-H1.0). Two adjacent cells were microirradiated with either the 488-nm laser to photoconvert the Or-H1.0 or sequentially with the 488-nm and 405-nm lasers to photoconvert Or-H1.0 and damage DNA. Recruitment of PARP1-GFP (upper) and release of photoconverted Or-H1.0 (lower) were simultaneously imaged in the FITC or far-red channels, respectively. (B) Parp1−/− cells were stably transfected with the plasmid expressing Or-H1.0 and microirradiated as described above. (C) Time course of release of Or-H1.0. (D) Quantitation of the time for loss of 25% of the initial fluorescence (t25) from undamaged (U) and damaged (D) regions (✱✱✱✱, unpaired student’s t-test, p-value < 0.0001, ns, not significant, N = 12)

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Expression of H1.0 with enhanced chromatin affinity slows the recruitment of PARP1 to damaged DNA

We then asked the converse question: is H1 depletion necessary for efficient PARP1 recruitment? The basic structure of H1 linker histones is conserved across species and variants, consisting of a short flexible N-terminal tail, a globular domain with a winged-helix motif, and a long, basic, lysine rich C-terminal [16, 65]. The globular domain binds to DNA within or near the nucleosome core to seal two full turns of DNA around the core and to stabilize the chromatosome [21, 66]. The C-terminal domain binds to linker DNA between adjacent nucleosomes and promotes the condensation of chromatin into high order structures [22, 67].

The C-terminal domain of H1.0 consists of four interchangeable subdomains of 20–25 amino acids [68]. We generated a mutant construct (H1.0Cdup) containing a duplication of the two distal subdomains (Fig. 3A, Supplemental Table S1). We introduced a plasmid expressing a pSMOrange-tagged version of H1.0Cdup into mouse 3T3 fibroblasts and measured the exit rate from damaged and undamaged DNA (Fig. 3B). As expected, the H1.0Cdup construct was released from undamaged DNA significantly more slowly than wild type H1.0 (compare to Fig. 2C) indicative of a tighter binding affinity. Release of H1.0Cdup was accelerated upon DNA damage but the exit rate was about three-fold slower than that of wild-type H1.0.

We previously developed a method to express exogenously introduced H1 isotypes by placing them under transcriptional control of the Zn-inducible metallothionein promoter and removing the 3’ UTR sequences that confer S-phase-specific mRNA stability [69]. We generated stable cell lines expressing un-tagged versions of H1.0 and H1.0Cdup. Upon ZnCl2 treatment, these lines expressed significant amounts of the exogenous protein (Fig. 3C). As we previously noted [69] overexpression of individual H1 variants results in a compensatory reduction in the expression of other variants such that the total amount of H1 relative to core histones is not significantly altered (Supplemental Fig. S3). We then transiently transfected a plasmid expressing GFP-tagged human PARP1 into these cells and wild type controls. We observed that recruitment of PARP1 to DNA damage following 405-nm laser microirradiation was significantly impaired in the cell line overexpressing H1.0Cdup (Fig. 3D-F) but not in the cell line overexpressing H1.0. We interpret this to indicate that release of H1 can be rate-limiting for the recruitment of repair proteins in response to DNA damage.

Fig. 3
figure 3

Expression of H1.0 with enhanced chromatin affinity slows the recruitment of PARP1 to damaged DNA. (A) Schematic of the domain structures of wild type H1.0 and H1.0Cdup. The latter contains a duplication of the C-terminal 48 amino acids encompassing the two distal subdomains of the C-terminal domain. (B) Wild type cells were stably transfected with a plasmid expressing Or-tagged H1.0Cdup. Two adjacent cells were micro-irradiated with either the 488-nm laser to photo-convert the H1.0Cdup or simultaneously with the 488-nm and 405-nm lasers to photoconvert H1.0Cdup and damage DNA. The release of H1.0Cdup was monitored as described in Fig. 2. (C) Wild type cells and cell lines expressing an untagged version of either H1.0 or H1.0Cdup were treated with 75 µM ZnCl2 for 48 h prior to isolation of total histones and separation by gel electrophoresis. (D) Wild type, H1.0-overexpressing, and H1.0Cdup-overexpressing cells were transfected with a plasmid expressing PARP1-GFP and treated with ZnCl2 as above. Cells were microirradiated with the 405-nm laser. (E) Quantitation of PARP1-GFP recruitment expressed as the fold-change in fluorescence within the microirradiated region relative to pre-irradiation values (N = 12). Values less than one are due to photobleaching. (F) Parp1-GFP recruitment at 60 s post-irradiation (✱✱✱✱, unpaired student’s t-test, p-value < 0.0001, N = 12)

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Prothymosin α (PTMA), a linker histone chaperone is required for the accelerated release of H1 from chromatin containing damaged DNA under repair

In earlier studies we used siRNA to lower the amounts of PTMA mRNA and protein and demonstrate a role for this protein as a linker histone chaperone [59]. However, this approach has limitations, especially when employing single cell assays as the treated cells are a mixed population with varying amounts of expressed PTMA. Here we used CRISPR/Cas9 technology to generate stable cell lines with null mutations in the endogenous Ptma genes (Fig. 4A, Supplemental Fig. S4). We also created stable “rescued” cell lines in which we introduced a plasmid expressing a myc-tagged version of either wild type or a deletion mutant of PTMA under transcriptional control of the metallothionein promoter (Supplemental Table S1). The mutant construct contains a deletion of sequences encoding amino acids 3–14 of PTMA and was previously shown to be defective in linker histone chaperone functions [59]. Neither the deletion nor the myc tag significantly changes the size or pI of the protein relative to wild type. By treating these cultures with the inducer ZnCl2 we were able to obtain expression of the exogenous PTMA to physiological levels (Fig. 4A, lanes 3–6).

We then transiently transfected a plasmid expressing Or-H1.0 into the wild type, Ptma−/− and rescued cell lines (Ptma−/− ResWT and Ptma−/− ResMut). These cell lines were subjected to microirradiation with the 488-nm or the 488-nm and the 405-nm lasers (Fig. 4B, C). Unlike wild type cells, Ptma−/− cells did not display accelerated loss of H1.0 in response to DNA damage. The reintroduction of wild type but not mutant PTMA restored accelerated linker histone eviction.

Fig. 4
figure 4

PTMA is required for the accelerated release of H1.0 from chromatin containing damaged DNA. (A) Western blot analysis of the expression of PTMA (upper panel) and actin (lower panel) in wild type, Ptma−/−, and rescued cell lines. Samples in lanes 4 and 6 were treated with 75 µM ZnCl2 for 48 h prior to preparing lysates. (B) A plasmid expressing H1.0 tagged with pSMOrange (Or-H1.0) was stably transfected into wild type, Ptma−/−, and the rescued cell lines. Cells were treated with 75 µM ZnCl2 for 48 h prior to microirradiation with either the 488-nm laser alone to photoconvert pSMOrange or sequentially with the 488-nm and 405-nm lasers to photoconvert pSMOrange and damage DNA. Eviction of photoconverted Or-H1.0 was monitored in the far-red region. (C) Quantitation of the time for loss of 25% of the initial fluorescence (t25) from undamaged (U) and damaged (D) regions (✱✱✱✱, unpaired student’s t-test, p-value < 0.0001, ns, not significant, N = 12)

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Ptma −/− cells display reduced recruitment of PARP1 to damaged DNA

We transiently transfected a plasmid expressing GFP-tagged human PARP1 into the wild type, Ptma−/−, and the rescued cell lines (Ptma−/− ResWT and Ptma−/− ResMut) and subjected them to microirradiation with the 405-nm laser (Fig. 5). Ptma−/− cells displayed a dramatically reduced recruitment of PARP1-GFP. Recruitment was partially restored by the reintroduction of wild type but not mutant PTMA (Fig. 5B, C).

Fig. 5
figure 5

Recruitment of PARP1 to damaged DNA is compromised in Ptma−/− cell lines. A plasmid expressing GFP-tagged human PARP1 was stably transfected into wild type, Ptma−/−, and rescued cell lines. Cells were treated with 75 µM ZnCl2 for 48 h prior to microirradiation with the 405-nm laser. A. Gallery of representative experiments. B. Time course of PARP1-GFP recruitment. C. Quantitation of individual measurements. Values are the maximum fold enrichment of fluorescence relative to pre-damage. (✱✱✱✱, unpaired student’s t-test, p-value < 0.0001, ns, not significant, N = 19)

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Chromatin expansion upon induction of DNA damage by irradiation with 405-nm light is dependent on PTMA

Utilizing H2B tagged with photoactivatable GFP, it was previously reported that wild type cells display a localized chromatin relaxation in response to microirradiation-induced DNA damage and that this expansion is dependent on both PARP1 and HPF1 [10]. Here we asked whether this expansion is dependent on PTMA as well (Fig. 6). The indicated cell lines were transfected with CMVOr-H2Bpur. Cells were microirradiated with either the 488-nm laser alone to photoconvert pSMOrange or sequentially with the 488-nm and 405-nm lasers to photoconvert pSMOrange and damage DNA. Photoconverted Or-H2B was monitored in the far-red region immediately after irradiation and after two minutes. The diameter of the region containing photo-converted pSMOrange was measured at t = 0 and t = 2 min. We observed a significant expansion of chromatin in response to microirradiation of wild type cells similar to that previously reported [10]. This expansion was not observed in cells depleted of PARP1, HPF1 or PTMA.

Fig. 6
figure 6

Chromatin expansion upon induction of DNA damage by irradiation with 405-nm light is dependent on PTMA. The indicated cell lines were transfected with CMVOr-H2Bpur. (A) Cells were microirradiated with either the 488-nm laser alone to photoconvert pSMOrange or sequentially with the 488-nm and 405-nm lasers to photoconvert pSMOrange and damage DNA. Photoconverted Or-H2B was monitored in the far-red region immediately after irradiation or after two minutes. (B) The diameter of the region containing photo-converted pSMOrange was measured at t = 0 and t = 2 min. Data are from three independent experiments (✱✱✱✱, unpaired student’s t-test, p-value < 0.0001, ns, not significant, N = 12)

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Ptma −/− cell lines express increased sensitivity to treatment with H202 or ionizing radiation

To further assess the effect of PTMA ablation on DNA damage repair, we performed colony survival assays after treatment with DNA damaging agents (Fig. 7). Compared to the wild type, Ptma−/− cells were significantly more sensitive to treatment with H2O2 or exposure to ionizing radiation, but not to UV irradiation

Fig. 7
figure 7

Ptma−/− cells are more sensitive to treatment with H2O2 or ionizing radiation than wild type cells. Cells were plated on 35 mm dishes and allowed to attach overnight. Following treatments, cells were allowed to grow for 14 days and then stained with Crystal Violet. Colonies larger than 1.5 mm were counted. Plots on the left represent data from three independent experiments. One representative experiment is shown on the right. (A) Cells were treated with the indicated concentration of H2O2 for 15 min then washed and fed with fresh medium. (B) Cells were exposed to the indicated amount of γ-irradiation from a 136Cs source. (C) Cells were exposed to the indicated amount of 265-nm UV irradiation

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Discussion

It was previously reported that H1 is depleted from chromatin containing damaged DNA under repair [27, 28] which we confirmed in our studies (Fig. 1). Using H1.0 tagged with a photoconvertible fluorescent protein, we observed a significant increase in the initial rate of exit from regions of chromatin containing damaged DNA versus untreated regions. This was also observed in Parp1−/− and Hpf1−/− cells suggesting that neither competition for binding between H1.0 and PARP1 nor HPF1-dependent ADP-ribosylation of protein serine residues are involved. The accelerated exit of H1.0 from sites of DNA damage was abrogated by homozygous null mutations in the endogenous genes encoding the linker histone chaperone PTMA. The recruitment of PARP1 to damaged DNA was also compromised by reduced expression of PTMA or overexpression of a mutant of H1.0 with a tighter chromatin-binding affinity. We interpret these results to indicate that depletion of H1.0 or other linker histones can be rate-limiting for the recruitment of repair proteins in response to DNA damage.

Several recent biophysical studies have investigated the interaction of H1.0 and PTMA as a model system for binding of intrinsically disordered proteins [70,71,72]. In our studies, the effects of PTMA depletion on PARP1 recruitment and H1.0 exchange were rescued by reintroduction of expression of wild type PTMA but not a mutant form bearing a small deletion near the amino terminus. This deletion does not significantly change the size or pI of the mutant protein and would not be expected to confer major changes in biophysical properties measured by in vitro assays with purified components. The observation that expression of the mutant form of PTMA does not rescue the biological processes measured here suggests that the mechanism of action of PTMA in vivo might be more specific. From a clinical perspective, PTMA levels are elevated in a number of cancers and associated with poor prognoses and outcomes [55, 56]. Development of cancer lines with increased resistance to treatment with radiation or chemotherapeutic drugs was shown to be associated with a further increase in PTMA levels indicating a possible involvement in the DNA damage response [57, 58]. Here we present evidence suggesting that PTMA, functioning as a linker histone chaperone is essential for an effective DDR.

Collectively these observations lead us to propose the following scenario (Fig. 8). DNA damage such as single- or double-stranded breaks might alter the nucleosome binding properties of H1 without directly promoting release. We, and others have shown that binding of H1 to the nucleosome involves a highly specific orientation of both H1 and the DNA strands entering and exiting the chromatosome [15, 17, 66, 73]. We have also shown that both the globular domain and the highly basic carboxy terminal tail contribute to tight binding of H1 and that metastable intermediates are formed during the exchange process [74]. We consider the possibility that DNA damage might compromise the binding of H1.0 to chromatin and allow the chaperone PTMA to bind and promote release of linker histones thereby initiating the local chromatin de-condensation necessary for the efficient recruitment of repair proteins such as PARP1. In this context linker histones serve as in situ “sensors” of DNA damage. This is not meant to imply that other processes such as competition between H1.0 and repair factors, PARP-dependent ADP-ribosylation of chromatin proteins, or downstream recruitment of chromatin remodelers do not also contribute to chromatin de-condensation. We do suggest the presence of a PTMA-dependent initial chromatin modulation that precedes and is necessary for subsequent.

Fig. 8
figure 8

Model for the role of PTMA in the DNA damage response. (A) Prior to DNA damage, chromatin is mostly in a condensed state due, in part, to H1 binding. (B) Upon DNA damage, a localized change in DNA conformation results in altered H1 binding providing access to PTMA. (C) PTMA promotes the release of H1 allowing PARP1 to bind. (D) PARP1 catalyzes the ADP-ribosylation of chromatin proteins including core histones. Chromatin remodelers are recruited through binding to ADP-ribose. F. Further chromatin remodeling promotes the recruitment of DNA repair proteins and PARP1 is displaced. Created with BioRender.com

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repair factor recruitment. This suggestion is supported by the experiments displayed in Fig. 6.

We were somewhat surprised to observe that Ptma−/− cells do not display accelerated loss of H1.0 in the absence of DNA damage which seems contradictory to the proposed role of PTMA as a linker histone chaperone. It should be noted that in our previous study we focused more on the role of PTMA in promoting H1 incorporation into chromatin previously depleted of linker histones [59]. We consider it possible that PTMA is not rate-limiting for the normal exchange of H1 between regions of undamaged chromatin but accelerates the exit of H1 following altered binding to damaged DNA as proposed in our model. It is also possible that PTMA also affects chromatin expansion via mechanisms independent of H1 binding. Regardless of the mechanism, the observation that Ptma−/− cells display increased sensitivity to DNA damage suggests an important role for PTMA in the DDR.

In this study, we have focused on the H1.0 variant. Our preliminary studies suggest that other variants, notably H1.2 behave in a qualitative if not quantitative manner to PTMA ablation (Supplemental Fig. S5). Interestingly, mouse embryonic stem cells depleted of H1.2, H1.4 and H1.5 were shown to have a significant reduction in the overall stoichiometry of H1 per nucleosome and to display hyper-resistance to DNA damaging agents [75]. This observation is consistent with our proposed model of the role of PTMA in DNA damage repair.

The involvement of an in situ sensor is attractive as it might partially resolve the question of how repair factors are recruited so rapidly. Rather than scan the entire genome, these factors would have preferentially access to the DNA within locally de-condensed chromatin. In the context of the ’monkey bar’ mechanism for PARP1 sensing DNA damages, these locally de-condensed regions may act as the rungs on the monkey bar [45, 46].

Methods

Cell culture

Mouse BALB/c 3T3 fibroblasts (ATCC) were maintained in DMEM-low glucose (Gibco, 11-885-092) supplemented with 10% heat inactivated bovine serum (Gibco, 26170-043) at 37oC in the presence of 5% CO2 and routinely screened for mycoplasma presence. Plasmids expressing exogenous proteins were transfected using Lipofectamine 3000 (Invitrogen). Stable cell lines were established by selection with puromycin (2 µg/ml, Gibco, A11138-03) and/or hygromycin B (200 µg/ml (Invitrogen, 10687010). Transient transfections were conducted 48–72 h prior to experimental protocols. Expression of fluorescently-tagged exogenous proteins does not significantly alter the total amount of the specific protein (18, Supplemental Fig. S6). Furthermore, we image cells expressing the lowest amount of tagged protein that results in an acceptable signal-to-noise ratio.

Plasmid constructs

Plasmid pPSmOrange was a gift from Vladislav Verkhusha (Addgene plasmid # 31898; RRID:31898). Plasmid SuperNova/pRSETB was a gift from Takeharu Nagai (Addgene plasmid # 53234; RRID: 53234).Plasmids were constructed by standard subcloning procedures and verified by DNA sequencing (Eurofins Genomics). Relevant details of the plasmids used in this study are presented in Supplemental Table S1. Plasmid MTH1.0Cduphyg was constructed by mutagenesis of MTsH1.0hyg [69] using the NEBuilderR HiFi DNA cloning kit (NEB). DNA oligonucleotides for guide RNAs, PCR and sequencing (Supplemental Table S2) were purchased from IDT. For CRISPR/Cas9-mediated generation of knockout cell lines, oligonucleotides were annealed and inserted into the vector from the GeneArt®CRISPR Nuclease CD4 Enrichment kit (Invitrogen) following the manufacturer’s instructions. These plasmids express both the sgRNA and Cas9.

Generation of Parp1 −/− and Hpf1 −/− cell lines

Plasmids Parp1CrX1-1, Parp1CrX1-2 and Hpf1CrX2-1 (Supplemental Table S1) were transfected into wild type 3T3 cells using Lipofectamine 3000 (Invitrogen) and plated at low density without selection. Approximately 50 independent colonies were subcloned and screened by sequencing of PCR fragments generated from genomic DNA and Western blotting (Supplemental Figs. S1 and S2).

Generation of Ptma −/− cell lines

As PTMA is essential for mouse embryogenesis and partial depletion of PTMA appears to slow cell proliferation [59, 76] we were concerned that isolation of a cell line completely devoid of PTMA might be problematic. Therefore, we first stably transfected wild type cells with a plasmid expressing low levels of PTMA fused to a fluorescent protein (CMVPTMASNhyg, Supplemental Table S1). Western blot analysis revealed that the tagged protein was expressed at < 5% the level of the endogenous protein (Supplemental Fig. S3A, lane 1). We then designed guide RNAs to the intron/exon junctions of exon 2 of the Ptma gene. Plasmids expressing these guide RNAs and Cas9 were then transfected into the cells described above. We were able to isolate stable cell lines of mouse 3T3 fibroblasts with homozygous frameshift mutations in exon 2 of the Ptma gene which contains codons for amino acids 15–39 of PTMA. We extensively characterized one isolate which bears a single nucleotide insertion in codon 17 of both alleles. Western blot analysis of PTMA levels revealed no detectable signal for endogenous PTMA (Supplemental Fig. S3A, lane 2, Fig. 4A). We refer to this cell line as Ptma−/− with the caveat that it may contain a very small amount of PTMA sequence in the form of a fusion protein.

Live cell imaging

Imaging was performed with a Nikon Eclipse C2 laser scanning confocal system mounted on a Ti-E motorized inverted microscope. Excitation/stimulation was performed with a solid state 405/488/561/640 laser unit using a CFI Apo 60X oil immersion objective, NA 1.40. Images were collected with a high sensitivity C2-DU3 Detection Unit with 435/34, 525/50, 600/50, a660LP filters and NIS-Elements C Imaging software. Cells were plated onto 35 mm glass bottom dishes (MatTek) and allowed to attach overnight. Prior to imaging, cells were sensitized by incubation in medium containing 1 µg/ml Hoechst 33,342 (Thermo Scientific) for 30 min. Cells were washed 3X with PBS followed by the addition of 2 ml of Fluorobrite DMEM imaging medium (Gibco) containing 10% bovine serum. To induce DNA damage a 2 μm diameter circular region of interest (ROI) was microirradiated with a single iteration of the 405-nm laser set to 10% power. SmOrange was photoconverted with a single iteration of the 488-nm laser set to 12% power. For photoconversion and damage, the ROI was sequentially microirradiated with the 488-nm laser followed by the 405-nm laser. We observed no detectable photobleaching of the converted SmOrange by the 405-nm laser. For some of the displayed panels, the pseudo-color was uniformly enhanced to visualize the low responders. For quantitation, grayscale images were imported into ImageJ [77]. Data is only included for individual measurements that remained within the linear range of detection throughout the duration of the experiment and were normalized for loss of signal due to total fluorophore bleaching during subsequent imaging [78]. For PARP1 recruitment assays, data is displayed as the fold-change relative to the pre-irradiation value (set to 1) For H1.0 eviction assays, data is displayed as the change in signal within the ROI relative to the first scan post-photoconversion.

Western blotting

Antibodies used and dilutions: α-PARP1 (abcam ab32138, 1:1,000); α-β-actin (Sigma-Aldrich A5441, 1:5,000); α-PTMA (Fisher Sc. PIPA575828, 1:500); α-mADPribose (Bio-Rad HCA355, 1:500); Goat α-rabbit IgG HRP (abcam ab97051, 1:10,000); Goat α-mouse IgG HRP (abcam ab6789, 1:10,000).

Equivalent numbers of cells from individual cell lines and/or treatments were scraped into PBS, pelleted and extracted with standard RIPA buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 1% SDS, protease inhibitor cocktail (HALT, Fisher, Sci.). Except as described below, samples were electrophoresed on 8–16% Tris-Glycine polyacrylamide gels with Precision Plus markers (Biorad) then transferred to nitrocellulose in Tris/Glycine buffer using the standard semi-dry setting on the Biorad turboblot transfer system. Membranes were blocked with 4.5% non-fat milk. Primary antibody incubations were conducted overnight at 4° C, and secondary antibody incubations were conducted for 2 h at room temperature. Bound antibody was detected with SuperSignal West Pic Plus chemiluminescent substrate kit (Thermo Sci.). Blots were exposed and quantified utilizing the QuantityOne 4.6.1 software on a Biorad chemidoc imager. To conduct western blots using anti-PTMA, the transfer was completed in 20 mM sodium acetate at pH 5.5 using the standard semi-dry setting on a Biorad turboblot transfer system. The membrane was crosslinked with 0.5% glutaraldehyde and the reaction stopped with 50 mM glycine. The antibody incubations were conducted as above but in 5% BSA as opposed to milk.

Colony survival assays

Approximately 50–100 cells were plated on 35 mm dishes and allowed to attach overnight. Following treatments as described in the figure legend, cells were allowed to grow for 14 days and then stained with Crystal Violet. Colonies larger than 1.5 mm were counted.

Statistics

Data were imported into Prism9.5 and analyzed using the statistical tests indicated in the figure legends.

Data availability

No datasets were generated or analysed during the current study.

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Acknowledgements

The authors thank Blaise Seale for generating Fig. 8, Yann Gibert for critical reading of the manuscript, and Michael Garrett, Chair, Department of Cell and Molecular Biology for additional support.

Funding

Research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under Award Number P20GM121334. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

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  1. Department of Cell and Molecular Biology, University of Mississippi School of Medicine, Jackson, MS, 39216, USA

    Ciara A. McKnight, Mary E. Graichen & David T. Brown

  2. Department of Physiology and Biophysics, University of Mississippi School of Medicine, Jackson, MS, 39216, USA

    Eric M. George

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D.T.B conceived and instructed the experimental work. C.A.M. and M.E.G conducted the experiments with input from D.T.B. and E.M.G. C.A.M., E.M.G. and D.T.B. analyzed and interpreted the data. D.T.B. wrote the original draft, which was reviewed and edited with input from all authors.

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Correspondence to David T. Brown.

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McKnight, C.A., Graichen, M.E., George, E.M. et al. The linker histone chaperone Prothymosin α (PTMA) is essential for efficient DNA damage repair and the recruitment of PARP1. Epigenetics & Chromatin 18, 32 (2025). https://doi.org/10.1186/s13072-025-00599-1

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Epigenetics & Chromatin

ISSN: 1756-8935