Our cells constantly withstand damage that can lead to breaks in the strands of our DNA. These cuts need to be fixed for the cell to stay healthy. When a break happens, one of the first responders to the scene is a protein known as PARP1. It binds to the ruptured strand (or strands) and then it recruits other repair agents to that location. But first, PARP1 needs to scan for cuts and notches amongst an overwhelming amount of DNA that is still intact. This is a complicated task, especially since the protein tends to bind both broken and unbroken DNA. How does it not stay ‘stuck’ on an undamaged portion of the genome?

Here, Rudolph et al. use a combination of biochemical techniques and cell biology to show that PARP1 travels through our genome by swinging from one DNA location to another, the way a child grabs onto monkey bars. One of the DNA-binding domains of PARP1, known as the WGR-domain, acts like an arm and initiates the movement by gripping onto a new segment of DNA. In fact, chopping off the WGR-domain or disabling it through mutations makes PARP1 worse at finding DNA breakages in the cell.

Unfixed DNA damage can lead to a cell becoming cancerous; ultimately, if the breakages keep accumulating, the cell does not survive. This makes PARP1 an important target for cancer treatment. Indeed, certain drugs already rely on trapping the protein so that tumor cells die. Understanding how cells cope with DNA damage and exactly how PARP1 works could help in the fight against cancer.

Poly(ADP-ribose) polymerase 1 (PARP1) serves as a first responder to DNA damage and is the founding member and most abundant representative of the large family of diphtheria toxin-like ADP-ribosyltransferases (ARTDs) (Bai, 2015; Bock and Chang, 2016; Beck et al., 2014; Daniels et al., 2015; De Vos et al., 2012; Mashimo et al., 2014; Morales et al., 2014). Binding to either single or double-strand DNA breaks (SSBs or DSBs) enzymatically activates PARP1 to use NAD⁺ in polymerizing long chains of poly(ADP)-ribose (PAR) onto itself and other nuclear acceptor proteins such as histones and DNA repair proteins. These PAR chains then recruit the appropriate DNA repair machinery containing PAR-binding motifs (Karlberg et al., 2013; Teloni and Altmeyer, 2016). PARP1 is of special interest because it is a validated target for cancer therapy (Tangutoori et al., 2015; Liu et al., 2014). Most notably, olaparib and rucaparib are in clinical use for treatment of ovarian and/or breast cancer in BRCA1/2 negative patients, and there are many on-going phase III clinical trials for inhibitors of PARPs either as monotherapy or in combination with chemo- or radiotherapy.

Overall, the domain structures of the 16 members of the ARTD family are quite diverse, but they all share a common catalytic core domain (~40 kDa) (Barkauskaite et al., 2015). Clinically relevant inhibitors of PARP1 bind in the catalytic domain. The N-terminal region of PARP1 contains five additional domains; three Zn-finger domains, an automodification domain that contains a BRCT-fold, and a WGR domain (Figure 1A). Seminal work from the Pascal laboratory has provided a molecular understanding of how Zn1, Zn2, Zn3, and the WGR domain collaborate to recognize DNA strand breaks in a structure-specific and sequence-independent manner, and subsequently activate the catalytic activity of PARP1. Zn1 and Zn2 separately (Langelier et al., 2011a), and together in the context of an SSB (Eustermann et al., 2015), bind one DNA end each using two points of contact, termed the phosphate backbone grip and the base stacking loop. In the context of a DSB, this grip-loop interaction mode is maintained by Zn1, while the Zn3 and WGR domains make additional contacts to the DNA (Figure 1B) (Langelier et al., 2012). Importantly, stepwise assembly of the different domains of PARP1 on DNA leads to the destabilization of the helical subdomain (HD) of the catalytic domain, which results in activation of its ADP-ribosyl transferase activity (Eustermann et al., 2015; Langelier et al., 2018; Dawicki-McKenna et al., 2015).

Figure 1 with 1 supplement see all

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In cells, PARP1 is known to contribute to many types of DNA repair mechanisms, including base excision repair, homologous recombination, nucleotide excision repair, and alternative non-homologous end-joining (de Murcia et al., 1997). In vitro, PARP1 is activated by a wide variety of DNA damage models including nicks, gaps, blunt ends, 5’- or 3’- extensions, all with or without a 5’-phosphate (Langelier et al., 2014). There is now clear evidence from multiple laboratories that PARP1 also binds tightly to undamaged DNA. For example, the Kraus laboratory has shown that PARP1 binds to and condenses intact chromatin, represses Pol II-dependent transcription, and is activated for auto-PARylation (Kim et al., 2004). We have previously shown that PARP1 serves as a chromatin architectural protein and interacts tightly (K_d ~nM) with and is activated by various nucleosome constructs (Clark et al., 2012; Muthurajan et al., 2014). Additionally, atomic force microscopy has shown that PARP1 binds not only to DNA ends or specific nicks, but also has significant affinity for undamaged DNA (Sukhanova et al., 2016). Most recently, single molecule tightrope assays have demonstrated that PARP1 interacts with and moves along undamaged DNA (Liu et al., 2017).

Thus PARP1 faces a similar ‘speed-stability’ paradox (Mirny et al., 2009; Zandarashvili et al., 2015; Halford and Marko, 2004) as transcription factors that need to find their target recognition site in an overwhelming excess of non-specific sites for which they also have significant affinity. PARP1 must rapidly search the genome for damaged DNA, yet it has significant affinity for the billions of base pairs of undamaged DNA that are present at concentrations of ~100 mg/mL in the nucleus (Krebs et al., 2017). In fact, laser micro-irradiation experiments in live cells have shown that PARP1 significantly accumulates at DNA damage sites in less than 10 s (Mortusewicz et al., 2007). The conundrum is that repeated cycles of release of PARP1 from undamaged DNA, random diffusional collisions, and rebinding to a different location may not be fast enough to explain how PARP1 can rapidly localize to sites of DNA damage. Various models have been put forth and tested for explaining how ‘facilitated diffusion’ could accelerate this search process, all of which recognize the importance, as opposed to hindrance, of non-specific binding to DNA for efficient site localization (Halford and Marko, 2004; Berg et al., 1981; Iwahara et al., 2006; Doucleff and Clore, 2008). These models include binding followed by one-dimensional sliding along DNA, hopping to a near-by site in the same chain, and intersegment transfer via an intermediate loop that is formed when the protein binds two different DNA sites at the same time. While some localized sliding along DNA has been reported for PARP1 (Liu et al., 2017), a more thorough kinetic characterization of binding to and dissociation from DNA is needed in order to address how PARP1 can efficiently localize to sites of DNA damage to initiate repair.

PARP1 has been found to associate more tightly with DNA in vivo in the presence of clinically relevant inhibitors. This phenomenon, known as PARP ‘trapping' (Tangutoori et al., 2015; Brown et al., 2016; Shen et al., 2015; Murai et al., 2012; Pommier et al., 2016) is thought to be in part responsible for the clinical effects of PARP inhibitors and has been used to explain the numerous discrepancies between in vitro inhibition of PARP1 vs. potency in preclinical models. For example, talazoparib is 100-fold more potent at trapping PARP1 on DNA and >50 fold more potent at killing cancer cells than rucaparib and olaparib, although the apparent IC-50’s for all three compounds are quite similar (1–5 nM) (Shen et al., 2015). Further complicating matters, extensive biochemical investigations of PARP1 trapping failed to provide evidence for an allosteric interaction between DNA- and inhibitor-binding (Hopkins et al., 2015), suggesting that trapping is due solely to inhibition of catalytic activity (but see [Langelier et al., 2018]). Thus, an evaluation of PARP inhibitors in a quantitative assay that measures DNA binding and release has the potential to shed further light on this controversial issue.

Here, we report on the kinetics of association and dissociation of PARP1 with DNA. We find that association of PARP1 with DNA is extremely fast, and that dissociation depends on the formation of a ternary complex where a second DNA molecule binds before release of the original DNA. We find that the WGR-domain, more specifically the conserved residue Trp589, is essential for triggering DNA-dependent release of DNA from PARP1, and we demonstrate the importance of this mechanism of DNA release for the accumulation of PARP1 at sites of DNA damage in the cell. Finally, we find that clinically relevant inhibitors do not perturb the rate or mechanism of release of DNA from PARP1.

Our results regarding the mechanisms of association and dissociation of PARP1 to and from DNA have important implications for our understanding of how PARP1 can move around the nucleus to scan for DNA damage. In vitro, PARP1 binds to DNA at or above the commonly accepted diffusion-limited rate (Record et al., 1991) of 1–2 nM⁻¹s⁻¹, consistent with its extremely rapid accumulation at sites of DNA damage in vivo (Mortusewicz et al., 2007; Aleksandrov et al., 2018). In fact, compared to other model systems using DNA oligomers, PARP1 – DNA association is among the fastest previously reported. For comparison, the rates of association of eukaryotic mismatch repair complex Msh2-Msh6 (Biro et al., 2010), RNA polymerase (Johnson and Chester, 1998), 8-oxoguanine-DNA-glycosylase (Lukina et al., 2017), and papillomavirus E2 protein (Ferreiro and de Prat-Gay, 2003) with double-stranded DNA are 0.002, 0.004, 0.13, and 0.6–1.4 nM⁻¹s⁻¹, respectively, effectively spanning three orders of magnitude. The fast association of PARP1 with DNA means that when dissociation does occur, re-association is most likely to the same site on the same DNA, as association is faster than diffusion carrying PARP1 away from its original binding site. This observation suggests that PARP1, like other DNA-binding proteins such as transcription factors (Mirny et al., 2009), must have a mechanism for moving around the genome that does not rely on simple dissociation and re-association. Although protein sliding along DNA in one-dimension has previously been invoked as a potential mechanism for accelerating the search for specific binding sites (Berg et al., 1981), more recent publications point out potential difficulties with such a long-distance sliding model (Halford, 2009), which is even more difficult to envision given the organization of DNA into nucleosomes in the eukaryotic genome.

Instead, we have found that PARP1 dissociation from DNA is triggered by binding of an additional DNA oligomer or undamaged plasmid prior to dissociation from the first DNA oligomer. We envision a ‘monkey bar’ model (Vuzman et al., 2010a; Vuzman et al., 2010b), wherein PARP1 moves from one DNA molecule to another DNA molecule, much like a child swings from bar to bar, transferring one hand at a time. This mechanism allows PARP1 to effectively scan the genome, moving to new and different sections of DNA. In the absence of competing DNA, PARP1 would remain effectively stuck at or near one site given its fast rate of association. Interestingly, undamaged DNA appears to be very effective at promoting the monkey-bar mechanism (Table 1), raising the question of how PARP1 remains stationary at sites of DNA damage. We have found that the WGR domain provides the other weaker ‘hand’ to facilitate the movement from one DNA strand to the next. Based on the structure of PARP1 bound to a DSB (Langelier et al., 2012), the WGR domain would need to first dissociate from the bound DNA prior to association with a second different molecule of DNA. Such a model is consistent with an NMR study that provides evidence for the stepwise assembly of PARP1 on a site of DNA damage where the Zn1, Zn2, and Zn3 domains engage a DNA molecule prior to final engagement of the WGR and catalytic domains (Eustermann et al., 2015). Thus, one can readily imagine a partial reversal of this process wherein the WGR domain releases the original DNA to bind the incoming DNA, followed by release of the original DNA and subsequent rearrangement of the domain around the newly bound DNA (Figure 8).

Figure 8

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A recent study of PARP1 using single-molecule DNA tightrope assays provides strong evidence for such a monkey bar mechanism (Liu et al., 2017). It was shown that micro-dissociation of one of PARP1’s multiple DNA-binding domains from DNA allows it to bind to a free 37 bp fragment, thus preventing rebinding of the domain to the tightrope and accelerating overall macro-dissociation. Our mechanism for DNA-dependent DNA dissociation also provides a compelling explanation for the wide diversity and significantly weaker dissociation constants previously reported for PARP1 with DNA (Langelier et al., 2018; Clark et al., 2012; Langelier et al., 2010): the measured apparent K_D depends strongly on the experimental conditions (i.e. [DNA]) under which the experiment is performed, since higher concentrations of DNA promote release of DNA.

Strong in vitro evidence for a monkey bar mechanism, also known as intersegment transfer, exists for various other proteins, all of them transcription factors. Kinetic experiments similar to ours have demonstrated strong dependence of DNA release on additional DNA for the lac repressor (Ruusala and Crothers, 1992), cAMP receptor protein (Fried and Crothers, 1984) and glucocorticoid receptor (Lieberman and Nordeen, 1997). NMR methods have demonstrated that both HoxD9 (Iwahara et al., 2006) and Oct1 (Doucleff and Clore, 2008) can bridge one DNA strand to another. A combination of methods including NMR, rate measurements, and computational modeling have elegantly demonstrated how EgrI uses its three Zn fingers to both bind and scan other DNA fragments as it moves between different recognition sites on different DNA molecules (Zandarashvili et al., 2015). Interestingly, mutagenesis of specific residues in EgrI was used to shift the equilibrium between binding and scanning modes.

One major caveat to any biochemical investigation of the mechanism of DNA – protein interactions is the artificial nature of a DNA oligomer compared to intact chromatin. This limitation affects studies of PARP1 interactions with DNA in particular because it is quite difficult to prepare completely intact DNA without ends or nicks, preferably wrapped around nucleosomes. Thus, it was important to test the significance of the monkey bar mechanism in a more physiologically relevant model. We have demonstrated the validity of interstrand transfer in vivo by demonstrating that the point mutant W589A, which disrupts DNA-dependent release of DNA, accumulates slower and to a lesser amount than wild-type PARP1 at sites of laser microirradiation (Table 2). The slower accumulation of W589A in cells is a particularly powerful demonstration of the importance of the monkey bar mechanism for PARP1 in finding sites of DNA damage for two reasons. First, the rate of dissociation for W589A from DNA is greater than for wild-type (20 s⁻¹ vs. < 10 s⁻¹). Second, the apparent K_D of W589A for DNA is weaker than that of wild-type PARP1 (5 nM vs. < 3 nM, respectively). Simplistically, these two observations might suggest that W589A should arrive at sites of DNA damage more rapidly than wild-type PARP1 since its interaction with DNA is not as tight or as long-lived (i.e. it spends less time occupying irrelevant sites); yet we observe the opposite. The monkey bar mechanism provides the explanation for these results: high concentrations of intranuclear DNA allow PARP1 to explore the nucleus rapidly. A dysfunctional monkey (W589A) surrounded by a lot of DNA does not move as rapidly. Thus, we have quantitatively demonstrated the importance of intersegment transfer in the accumulation of a DNA-binding protein at its target destination in vivo.

Finally, our results also provide further insight into the much-discussed topic of PARP1 ‘trapping’, wherein cells treated with inhibitors of PARP1 exhibit a shift of PARP1 from the soluble fraction to a chromatin-associated insoluble fraction (Murai et al., 2012). Many cell-based studies have since confirmed the phenomenon of PARP-trapping (Pommier et al., 2016). Our data showing no effect of four different tight-binding inhibitors of PARP1 on the release of DNA agree with a previous thorough biochemical analysis that also could not find any effects of inhibitors on DNA binding constants or rates of dissociation (Hopkins et al., 2015). Thus, the mechanistic basis for PARP-trapping is more complex than can be reconstituted in vitro. Our results are in agreement with recent findings that PARP inhibitors lead to defective fork recovery and/or homologous recombination-mediated repair, and thus an increase in DNA damage where PARP1 is bound to DNA, and lacking activity due to inhibitor binding (Maya-Mendoza et al., 2018).

We have provided Excel files for all the figures and tables.

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Author details

1. Johannes Rudolph

   Department of Chemistry and Biochemistry, University of Colorado Boulder, Boulder, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Validation, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0230-3323

2. Jyothi Mahadevan

   Department of Chemistry and Biochemistry, University of Colorado Boulder, Boulder, United States

   Contribution

   Formal analysis, Validation, Investigation, Methodology

   Competing interests

   No competing interests declared

3. Pamela Dyer

   1. Department of Chemistry and Biochemistry, University of Colorado Boulder, Boulder, United States
   2. Howard Hughes Medical Institute, University of Colorado Boulder, Boulder, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-0142-5073

4. Karolin Luger

   1. Department of Chemistry and Biochemistry, University of Colorado Boulder, Boulder, United States
   2. Howard Hughes Medical Institute, University of Colorado Boulder, Boulder, United States

   Contribution

   Conceptualization, Funding acquisition, Project administration, Writing—review and editing

   For correspondence

   karolin.luger@colorado.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5136-5331

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