The excluded DNA strand is SEW important for hexameric helicase unwinding

Methods xxx (2016) xxx–xxx

Contents lists available at ScienceDirect

Methods journal homepage: www.elsevier.com/locate/ymeth

The excluded DNA strand is SEW important for hexameric helicase unwinding Sean M. Carney a, Michael A. Trakselis a,b,⇑ a b

Molecular Biophysics and Structural Biology Program, University of Pittsburgh, Pittsburgh, PA 15260, United States Department of Chemistry and Biochemistry, Baylor University Waco, TX 76798, United States

a r t i c l e

i n f o

Article history: Received 1 March 2016 Received in revised form 7 April 2016 Accepted 7 April 2016 Available online xxxx Keywords: Helicase Unwinding Steric exclusion Hydrogen-deuterium mass spectrometry Single molecule FRET DNA damage sensing

a b s t r a c t Helicases are proposed to unwind dsDNA primarily by translocating on one strand to sterically exclude and separate the two strands. Hexameric helicases in particular have been shown to encircle one strand while physically excluding the other strand. In this article, we will detail experimental methods used to validate specific interactions with the excluded strand on the exterior surface of hexameric helicases. Both qualitative and quantitative methods are described to identify an excluded strand interaction, determine the exterior interacting residues, and measure the dynamics of binding. The implications of exterior interactions with the nontranslocating strand are discussed and include forward unwinding stabilization, regulation of the unwinding rate, and DNA damage sensing. Ó 2016 Elsevier Inc. All rights reserved.

Contents 1. 2.

3.

4. 5. 6.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Experimental methods for detecting excluded strand contacts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Specific excluded DNA strand footprinting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Nontranslocating strand-protein crosslinking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Hydrogen deuterium exchange mass spectrometry (HDX-MS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Single molecule fluorescence energy transfer (smFRET). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Explicit probability and rate transition (ExPRT) plots. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Impact of the excluded strand in unwinding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Excluded strand as a ‘molecular ratchet’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Excluded strand as a ‘molecular brake’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Nonhexameric helicases that engage both strands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sensing of DNA damage. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Role of the excluded strand in the unwindosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Final thoughts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

00 00 00 00 00 00 00 00 00 00 00 00 00 00 00 00

Abbreviations: AAA+, ATPases associated with various activities; ARDD, average relative deuterium uptake; BER, base excision repair; BrdU, bromodeoxyuridine; CHiP, chromatin immunoprecipitation; CMG, Cdc45/MCM2-7/GINS; CPD, cyclobutane pyrimidine dimer; DNA, deoxyribonucleic acid; dsDNA, double stranded DNA; Ec, Escherichia coli; FA, Fanconi anemia; FRET, fluorescence resonance energy transfer; FT, Fourier transform; HDX, hydrogen-deuterium exchange; HR, homologous recombination; IdU, iododeoxyuridine; MCM, minichromosomal maintenance; MMR, mismatch repair; MS, mass spectrometry; MS-MS, tandem mass spectrometry; NER, nucleotide excision repair; NTP, nucleotide triphosphate; SE, steric exclusion; SEW, steric exclusion and wrapping; SF, superfamily; smFRET, single molecule FRET; ssDNA, single stranded DNA; Sso, Sulfolobus solfataricus; Ta, Thermoplasma acidophilum; TIRF, total internal reflectance fluorescence; UV, ultraviolet light. ⇑ Corresponding author at: One Bear Place #97365, Waco, TX 76798, United States. E-mail address: [email protected] (M.A. Trakselis). http://dx.doi.org/10.1016/j.ymeth.2016.04.008 1046-2023/Ó 2016 Elsevier Inc. All rights reserved.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

2

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

1. Introduction The loading, activation, and action of hexameric DNA helicases are tightly regulated to occur during the initiation and elongation phases of DNA replication. Hexameric helicases have generally evolved a toroidal geometry and are structurally classified based on having either RecA folds or within the broader ATPases associated with a variety of cellular activities (AAA+) clade [1–3]. RecA hexameric helicases are within the superfamily (SF) 4 and are of some of the most well studied including: T7 gp4, T4 gp41, bacterial DnaB, and mitochondrial Twinkle. Hexameric AAA+ SF3 helicases have conserved Walker A and B motifs, an arginine finger coming from an adjacent subunit to make up the ATPase site, and a unique motif C and are from DNA viruses that include E1 and SV40 Large T-antigen. The archaeal and eukaryotic SF6 hexameric AAA+ MCM helicases have similar ATPase sites comprised from adjacent subunits but also include additional sensor-1 and sensor-2 motifs that function in trans to one another to control ATP hydrolysis [4]. Although both structural and functional motifs differ across the three hexameric DNA helicase SFs [3,4,6], the overall three dimensional structure has evolved to encircle and separate DNA strands processively [5]. The prevailing view is that most hexameric helicases encircle one DNA strand (ssDNA) within their central channel, while physically excluding the complementary strand to the exterior [6]. Although it has been shown that hexameric helicases can accommodate double stranded DNA (dsDNA) within the central channel, dsDNA translocation does not appear to contribute to effective DNA unwinding [7]. The exception may be SV40 Large-T-antigen which is known to encircle dsDNA, however, the added ability to bind and melt origin DNA of this SF3 helicase may have captured this specific conformation prior to conversion of encircling only a single-strand [8,9]. Nevertheless, in all cases, hexameric helicases utilize the energy from nucleoside triphosphate (NTP) hydrolysis to propagate the destabilization of hydrogen bonding within the duplex [10]. Hydrolysis at the conserved Walker A and B motifs combined with the other cis and trans acting elements engage contacts with the translocating strand and propel the helicase forward in a series of steps. NTP hydrolysis is proposed to occur in a sequential mechanism around the hexamer [11–13]. The conformation of the hexamer has been shown to exist in multiple states including a flat washer, a cracked-ring, or a split spiral [12,14–18]. The degree of out-of-plane spiraling may correlate with the NTPase associated step size, such that additional contacts of the translocating strand within a spiral ring contribute to greater step sizes [19,20]. Whether the global conformation of the hexamer changes or remains fixed during the course of unwinding or for every step is fascinating aspect of the helicase mechanism that is not yet solved. Once loaded onto ssDNA, hexameric helicases have specific unwinding polarities, where SF4 helicases translocate 50 –30 and SF3 (except for E1) and SF6 helicases translocate 30 –50 . Strand separation is stimulated by the presence of a steric block, such as the nontranslocating strand. The steric exclusion (SE) model (Fig. 1) of unwinding has been accepted for decades to explain the unwinding action of not only hexameric helicases, but also monomeric and dimeric helicases. One limitation of the SE model is that it generally ignores any contribution of the excluded strand in the unwinding mechanism. Interactions with the excluded strand have been shown previously with hexameric helicases [21], but it was only recently that their role in the DNA unwinding mechanism has been revealed [22]. Although, it has been reported previously, that the ssDNA is bound in the central channel and not wrapped around the DnaB hexamer [23,24], we would hypothesize that external surface binding of ssDNA is not thermodynamically stable

Fig. 1. Structural models for hexameric helicase DNA unwinding include steric exclusion (SE) or steric exclusion and wrapping (SEW) where the nontranslocating strand makes contact with the exterior surface (orange).

when the primary central binding site is available. Only after encircling the 50 -strand of fork DNA would the excluded 30 -strand be conformationally favored for exterior surface binding. In support, binding of a second ssDNA strand to T7 gp4 and EcDnaB helicases has been measured but with lower affinity [25,26]. Therefore, we have expanded the SE model to include favorable interactions with not only the translocating strand but also the excluded strand and termed this the steric exclusion and wrapping (SEW) model for unwinding (Fig. 1). In this methods review, we will detail experimental techniques used to determine the importance and influence of the excluded strand in the DNA unwinding mechanism. 2. Experimental methods for detecting excluded strand contacts To determine whether exterior interactions of helicases with the nontranslocating DNA strand exist, a variety of qualitative and quantitative biochemical and biophysical experiments can be performed. Both stable and dynamic binding of the nontranslocating strand to the exterior surface may aid in DNA unwinding, and assays are needed to differentiate strands and quantitate specificities. Precise detection of nontranslocating strand binding coupled with mutagenesis can unequivocally determine whether the excluded strand plays any role in DNA unwinding and stabilization for DNA helicases (Table 1).

Table 1 Comparison of methods used to validate the Hexameric Helicase SEW Model. Method

Experimental advantages

DNA footprinting

Identify specific regions and lengths of each strand of DNA protected upon binding to the helicase Captures both transient and stable covalent protein-DNA complexes for analyses of strand specificities and amino acid identification Global unbiased measurement of DNA binding to the helicase in solution without perturbations Determines populations of distance-based DNA conformations and their changes upon wild-type or mutant helicase binding Allows for easy visualization of conformational transitions, binding dynamics, and transition rates of fluorescently labeled DNA strands between two or more experimental conditions

DNA crosslinking

HDX-MS smFRET

ExPRT analyses

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

3

with different translocation polarities. In all cases, protection from digestion when helicases are added can confirm SEW interactions. It is essential that the helicase and DNA concentrations are stoichiometric and above the binding Kd in these experiments. If helicase is in excess, then encircling of both ssDNA tails is possible providing spurious protection. Trace amounts of 32P or fluorescently labeled DNA can be added to higher concentrations of unlabeled DNA as a marker for digestion. Both the reaction time and the concentration of the chosen nuclease should be systematically tested to achieve optimal footprinting and prevent over-digestion. We have previously used single-strand specific mung bean nuclease footprinting to show that the excluded 50 -tail, even up to 80 bases, is protected from degradation when archaeal SF6 SsoMCM is added (Fig. 2B) [22]. 2.2. Nontranslocating strand-protein crosslinking

Fig. 2. A) Excluded strand specific nuclease footprinting experiments used to test the SEW model. B) Fork DNA substrates with variable 20–80 base excluded strands in the absence () or presence (+) of stoichiometric concentrations of SsoMCM digested with mung bean nuclease for 30 min. DNA was labeled with 32P at the 30 duplex end. Schematics of the fork substrates are above. Samples were resolved on denaturing acrylamide gels and the fraction protected quantified using phosphorimaging. (Figure modified from Ref. [22].)

2.1. Specific excluded DNA strand footprinting A variety of DNA footprinting approaches have been used ubiquitously for decades to map DNA/RNA binding sites and substrate specificities of protein binding [27]. Generally, DNA footprinting can be chemical (i.e. OH radical [28] or OP-Cu [29]) or enzymatic (DNAseI [30] or other nucleases) but care needs to be taken as to whether the method digests ssDNA, dsDNA or both. Many excellent review and methods articles exist on the specificities, intricacies, and limitations of DNA footprinting [28,31–33]. Instead, we will focus on the footprinting applications that probe the binding of only the nontranslocating strand while leaving the encircled translocating strand intact. To specifically detect binding of only the excluded strand, single strand specific nucleases are particularly useful. Hydroxyl radical or OP-Cu footprinting could also be performed but would digest unbound single or doubled stranded DNA creating a complete pattern of binding. Single strand specific DNA endonucleases including S1 and mung bean nucleases and can digest unbound DNA tails or show the length dependence of excluded stand binding (Fig. 2A). Promiscuous ds/ssDNA nucleases such as DNaseI or micrococcal nuclease could also be used, however, the digestion pattern will be more complicated and include digestion of the duplex region similar to chemical methods. Alternatively, ssDNA exonucleases with specific polarities (i.e. RecJf, 50 –30 or ExoI, 30 –50 ) could be used to digest longer tailed substrates down towards the duplex and test excluded strand binding for helicases

In order to more thoroughly characterize exterior binding path residues, a crosslinking protocol can be designed to map specific helicase binding sites of the nontranslocating strand (Fig. 3A). Biotinylated-DNA can be used either for detection with antibodies or enrichment of peptides later using streptavidin beads. Crosslinking can be initiated either through chemical agents (i.e. glutaraldehyde) or at specific modified DNA bases (i.e. BrdU or IdU) using ultraviolet (UV) light. Again, it is essential that these experiments be performed with stoichiometric concentrations to prevent loading of multiple hexamers. Glutaraldehyde has been used for crosslinking because of its ease of use and lack of specificity in reacting any two amino groups in close proximity through Schiff base formation [34–39]. Generally, glutaraldehyde crosslinking is used in chromatin immunoprecipitation (ChIP) experiments to identify the DNA sequence for protein binding [40] by targeting free amino or imino groups on proteins and nucleic acids [41,42]. We have used this protocol instead to identify the exterior protein region of helicases that bind the nontranslocating strand. Although crosslinking can easily occur at the imino nitrogen of T, faster crosslinking can occur with exocyclic amines of A, G, and C and can be incorporated site specifically into the sequence. Addition of 0.001–0.01% glutaraldehyde to the bound helicase-DNA complex for 1–10 min is usually sufficient to induce crosslinking. The glutaraldehyde concentration and time should be systematically titrated for optimal crosslinking yield. Glutaraldehyde crosslinking can be seen in Coomassie stained and phosphorimaged 32P labeled DNA within the same gel (Fig. 3B). Trypsin digestion of the crosslinked DNA-SsoMCM followed by biotin/streptavidin bead selection can be used to isolate the crosslinked peptide fragment (Fig. 3A). The degree of crosslinking will affect the efficiency of the trypsin digestion. Therefore crosslinking conditions should be optimized to minimize spurious crosslinks. At this stage, reversal of the glutaraldehyde crosslinks can simplify downstream mass spectrometry (MS), however, glutaraldehyde crosslinks are very stable, and if lysines are targeted, they cannot be easily regenerated even after 24 h at 110 °C in 6 N HCl [43]. In our experience, heating the sample to 95 °C in Tris buffer for up to one hour, releases less than 10% of the glutaraldehyde crosslinks which may complicate downstream analyses. In order to more specifically target crosslinking to just the excluded strand site, bromodeoxyuridine (BrdU) or iodouracil (IdU) can be incorporated into the excluded single strand at different positions which primarily targets tyrosines, histidines and tryptophans for crosslinking [44–46]. BrdU or IdU can be synthesized directly into oligos by a number of DNA synthesis companies. Briefly, reactions are placed under the UV transilluminator and exposed to UV light (254 nm) for crosslinking at a distance of 5–10 cm. Increases in molecular weight can be visualized after phosphorimaging 32P-labeled DNA at a yield of roughly 15-20%

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

4

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

Fig. 3. A) Experimental flow for crosslinking of the excluded strand and identification of exterior helicase binding sites using tandem mass spectrometry (MS-MS). B) Global glutaraldehyde crosslinking of the SsoMCM/fork DNA complex or C) more specific UV induced BrdU crosslinking of the excluded strand can be resolved on a SDS-PAGE gel and phosphorimaged to visualize the DNA followed by Coomassie staining to visualize the protein and determine the degree of crosslinking.

(Fig. 3C). Reversal of UV induced BrdU crosslinks can also be performed at 95 °C for 5 min and typically releases 50% of these crosslinks. Following crosslinking, trypsin digestion, and then reversal, isolated peptides can be analyzed by tandem mass spectrometry for identification of the external binding residues. The advantage of specific BrdU crosslinking compared to more general crosslinking methods is the reduced background leading to less complex mixtures and reduced sample noise. Although we will not focus on the specific mass spectrometry techniques, high resolution MS-MS is recommended to counter low sample concentrations, yields, and the diversity of the possible crosslink chemistries. Flexibility in controlling the detection mass algorithms are essential for identifying the crosslinked site because of the conjugated ssDNA fragment [47]. The MS1 spectrum can be used to detect crosslinked peptides compared to an uncrosslinked control. The MS2 spectrum can be quite complex because of fragmentation of the DNA in addition to the peptide. In most cases, analysis is performed manually because of the complexity and diversity of crosslinking approaches and the lack of appropriate software. This basic approach has been used to characterized and map a variety of DNA-protein and RNA-protein interactions [48–56].

2.3. Hydrogen deuterium exchange mass spectrometry (HDX-MS) Hydrogen deuterium exchange mass spectrometry (HDX-MS) detects amide proton exchange with solution and can identify protein regions that become protected from exchange upon ligand binding [57–59]. A major benefit of HDX-MS is that native conformations of complexes can be explored without any perturbations typically needed for analytical quantifications [60,61]. Differences in HDX when ligands (including DNA) are bound can be mapped back onto the protein surface to determine the interactions surfaces [60,62]. Although difficult because of competing symmetry in the hexamer blurring specific ssDNA binding to one or two subunits, high resolution HDX-MS can provide an unbiased global representation of the excluded strand DNA binding path. In order to compare the DNA binding paths on helicases using HDX-MS, experiments should be performed with different DNA substrates where the nontranslocating strand is either present or absent (Fig. 4A). After stoichiometric binding, samples are diluted 10-fold into D2O to initiate exchange before quenching in 8 M urea, 1.0% formic acid, and protease type XIII (Sigma-Aldrich). Protease type XIII has nonspecificity of cleavage resulting in a beneficial

Fig. 4. A) Experimental flow of the Fourier transform hydrogen-deuterium exchange tandem mass spectrometry (FT-HDX-MS). B) Example of the deuterium uptake kinetics for a single peptide compared between two experiments (i.e. with and without a 50 -tail). The D-uptake rate is then globally compared across the entire peptide space using the ARDD equation to identify differences in HDX.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

5

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

library of overlapping peptides used in the analysis. Peptides are rapidly separated using in-line liquid chromatography and electrosprayed on a FT-ICR-MS at the National High Magnetic Field Laboratory (Tallahassee, FL) [63], although other high resolution instrumentation facilities could perform similar analyses. The measured deuterium uptake (D) is quantified for exponential rates and standard deviations from multiple experiments (Fig. 4B) [64]. From the rates, the average relative deuterium uptake differences (ARDD) between free and DNA-bound helicase can be calculated giving a fractional exchange difference from 1 to 1. These values are then color coded and mapped back onto the three-dimensional structure of the helicase to identify DNA binding sites that perturb HDX exchange (Fig. 4C). One caveat is that for symmetrical homoligomeric helicases, unbound subunits contribute neutrally to the ARDD intensity increasing the noise and obfuscating detection of DNA bound to one or two specific subunits. However, using this global approach, we have not only identified exterior DNA binding paths but can also show how electrostatic mutations that reduce DNA unwinding also alter the binding path on the exterior of SsoMCM validating the SEW model [65]. In addition to helicases, this high resolution HDX-MS method has identified RNA binding sites on phage packaging motors [60] and on RIG-I that sense viral RNA infection for vertebrate immunity [66].

2.4. Single molecule fluorescence energy transfer (smFRET) Even though techniques like footprinting and crosslinking can be used to identify interactions between helicases and DNA, they cannot inform on the dynamics and heterogeneity of those interactions. Single-molecule FRET (smFRET) is a technique that can yield detailed information concerning the dynamics of an interaction, and it has been used in several studies to characterize the interactions between hexameric helicases and the excluded nontranslocating strand during unwinding [22,67–69]. Generally, smFRET utilizes a flow cell where the inside surfaces have been pegylated to reduce non-specific binding to the surface. A small fraction of these PEG molecules are biotinylated so that biotin-streptavidin interactions can immobilize the protein, RNA, or DNA of interest (Fig. 5B). The flow cell is imaged using a total internal reflection fluorescence (TIRF) microscope, and a diode laser is used to excite the donor dye (Fig. 5A). Subsequent donor and acceptor fluorescent signals are imaged with dual channel emission optics (Fig. 5C). This allows for the quantification of corresponding donor and acceptor dye signals from each single molecule imaged over time (Fig. 5D). Quantification of the donor and acceptor signals allow the FRET efficiency (E) to be calculated using:

E¼

IA ID þ I A

ð1Þ

Fig. 5. smFRET experimental setup. A) TIRF microscopy. A 532 nm diode laser is focused and directed through a prism sitting on top of the flow cell. Emitted light is captured by the objective, and separated based on wavelength before entering the CCD camera. B) smFRET flow cell design. The flow cell is coated in polyethylene glycol (PEG) to prevent non-specific binding. A small fraction of these immobilized PEG molecules are biotinylated. Streptavidin is used to tether biotinylated DNA to the biotinylated PEG surface. The DNA substrates are model fork structures where a duplex region branches into two poly-deoxythymidylates. The 30 and 50 ends of the DNA fork are labeled with Cy3 and Cy5 respectively. C) CCD imaging. The separated donor and acceptor signals are detected by the CCD camera. Two resulting channels show superimposable images of the Cy3 signal in the donor channel and the Cy5 signal in the acceptor channel. D) Single-molecule time trace. Top Panel: Corresponding donor and acceptor intensity can be followed over time as shown by the outlined circles in C). Each single pair (e.g. arrows) can be plotted over time to produce a single-molecule fluorescence intensity time trace. Bottom Panel: The FRET efficiency is calculated and plotted over time (blue) and fit to ideal states (red) to aid in downstream analysis.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

6

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

where IA and ID are the intensity values of the donor and acceptor respectively. The relation between FRET efficiency and the distance between the dyes can be described by:

E¼

1þ

1  6

ð2Þ

r R0

where r is the distance between the dyes and R0 is the Förster distance for the FRET pair, or distance between the dyes where half of the energy is transferred. Details concerning specific protocols for performing smFRET can be found in reviews dedicated to the practical and theoretical considerations of smFRET experiments [70–72]. The DNA substrates used in the helicase-interaction assays are designed as model forks, where a duplex region is immobilized via biotin-streptavidin interactions to the pegylated flow cell surface and two single-strand arms of the fork project outwards away from the surface (Fig. 5B). To avoid spurious base pairing of the single-strand fork arms, the arms for the fork can consist solely of oligo-deoxythymidylates. The termini of the fork arms are

labeled with the Cy3-Cy5 FRET pair. The energy transferred from the Cy3 donor dye to the Cy5 acceptor dye is a function of the distance between the dyes as shown in Eq. (2). In the absence of any protein, the FRET signal is low due to the ends of the fork arms not being within close proximity as can be seen in the histogram corresponding to the DNA3050 fork substrate alone (Fig. 6A). When introducing a hexameric helicase that encircles one of the fork arms in its central channel as it loads onto the fork substrate, one might expect to see a further decrease in the FRET signal due to the hexamer acting as a wedge that would drive the two arms of the fork further apart as in the SE model (Fig. 1). However, the first time this assay was implemented to study the interaction with the SsoMCM helicase, an increase in FRET was observed upon helicase binding [67]. Performing a similar experiment, we see consistent results (Fig. 6A), where the wild-type (WT) SsoMCM helicase induces a high FRET state on the DNA3050 fork. An increase in FRET efficiency is consistent with the excluded strand physically interacting with the outer surface of the hexamer so that the Cy3 – Cy5 pair are brought within close proximity. Surface mutations along the proposed path of interaction with the

Fig. 6. smFRET analysis of WT and mutant SsoMCM bound to DNA. A) A histogram of all single-molecule FRET data collected for the DNA3050 substrate alone, with WT SsoMCM, and SsoMCM (K323D/R440D). B) and C) show the ExPRT plots for WT SsoMCM and SsoMCM (K323D/R440D), respectively. D) The legend for the ExPRT plots: Transition Probability and Dwell Times. smFRET time traces for E) WT and F) K323D/R440D showing Cy3 (green) and Cy5 (red) intensities and FRET efficiencies (blue). The red line is a fit to ideal FRET states.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

excluded-strand were made (SsoMCM K323D/R440D) and the resulting histogram profile shows a broad distribution of FRET signals, suggesting a disruption of the wrapping interaction seen for the wild type. These residues have been previously suggested to mediate the excluded strand wrapping interaction in the case of SsoMCM [22] and point to an electrostatic-based wrapping interaction consistent with the SEW model (Fig. 1). We have used similar smFRET assays to further characterize the SF6 SsoMCM helicase as well as the SF4 EcDnaB helicase and the SF4 human Twinkle helicase. For each of these helicases, the length of the excluded arm of the fork (50 arm for SsoMCM and 30 arm for EcDnaB and Twinkle) was varied. The length of each arm of the model fork can be altered to examine how the excluded strand interaction changes with the length of the single strand available for binding. In all cases, the helicase induces a significant shift from low FRET for the naked DNA forks to high FRET when bound by the helicase [22,68,69]. Surface mutations of SsoMCM and EcDnaB have been created that disrupt the excluded strand interaction as monitored by smFRET [22,68]. Although a histogram of the FRET signal can show differences in the FRET state(s) being sampled, it does not characterize or quantify the dynamics of the interaction. 2.5. Explicit probability and rate transition (ExPRT) plots smFRET allows for the monitoring the distances between single Cy3 – Cy5 pairs on single molecules in real time. Therefore, single events such as binding interactions, conformational changes, and protein translocation can be observed explicitly in many singlemolecules. Analyses that quantify the probabilities and rates of these events can be used to provide a detailed characterization of the dynamics being observed. There are several programs available to perform analyses on smFRET data [73–76]. These programs rely on using Hidden Markov modeling programs, such as HaMMY and vbFRET to fit data to ideal states [73,77]. The analysis programs then use the ideal state fits to the raw data to generate transition plots, where the initial FRET value (the FRET state preceding the transition) is on the x-axis, and the final FRET value (the FRET state transitioned into) is on the y-axis. For example, the Transition Density Plot (TDP) program created by the Ha laboratory uses ideal state fits to each individual trace and visualizes all transitions from all traces as heat map. This allows heterogeneity within the transitions to be visualized and for a qualitative comparison of transition probability to be visualized, but lacks visualization features that provide explicit quantification of the data directly on the plot [73,74]. Another analysis program is the Probability and Kinetically Indexed Transition (POKIT) Plot program developed by the Walter laboratory [75,76]. This program relies on fitting all traces as one stitched trace, which allows for the analysis of the transition rate and probability between the number of ideal states fitted to the stitched trace. The POKIT program utilizes color and concentric markers to plot transitions into bins of probabilities and rates. Each single marker and its corresponding x,y coordinate together represent a unique transition. However, plotting the data into bins diminishes the power of single-molecule methods to determine the explicit parameters such as probability and rates based on single-molecule events. There are limitations and shortcomings for all analysis programs. The currently available analyses either cannot visualize both probability and kinetic information on a single plot or rely on binning data into ranges of values in order to do so. We sought to overcome some of these limitations by creating our own smFRET analysis program termed Explicit Probability and Rate Transition (ExPRT) plots. The ExPRT analysis program is a Matlab executable program that produces ExPRT plots from raw smFRET values. The ExPRT program follows an initial data processing and analysis approach

7

similar to the POKIT program, where the raw data that is collected must be stitched together, and fit to ideal states using software such as vbFRET [77]. That ideal stitched trace is then unstitched to avoid analyzing ‘false transitions’ and to generate single-trace statistics. The unstitched traces can then be fed into the ExPRT program. Fig. 6E-F show example unstitched traces for WT SsoMCM and SsoMCM (K323D/R440D) on the DNA3050 fork. The upper panel shows the raw Cy3 donor and Cy5 acceptor intensities, and the lower panel shows the FRET efficiency calculated from the raw data. The overlaid red line is the ideal state fit to calculated FRET values. The program goes through each trace and collects all transitions that occur in addition to the dwell time of each state between transitions. Once all traces have been analyzed, the ExPRT program generates the transition plots (Fig. 6B-C). Each transition is plotted as a marker based on its initial FRET state on the x-axis and final FRET state on the y-axis. The size of the marker corresponds to the probability of that transition occurring within a measured single trace. We define probability as the fraction of analyzed traces that exhibit the given transition at least once. The color of the marker corresponds to the time spent in the initial state before transitioning (Fig. 6D). Although the ExPRT plots resemble POKIT plots in that they both use circular markers to represent transitions, ExPRT plots use and visualize the explicit probabilities and dwell time values rather than displaying the data in bins. In addition, the ExPRT program also plots the dwell times of each transition in the form of a survival curve, and fits the subsequent curve to single and double exponential decay functions. The user is then able to determine which fit is appropriate based on the differences in R2 values between fits. If a double exponential decay is selected, then the resulting marker on the ExPRT plot will be concentric, containing two colors where each color corresponds to the dwell time based on the rates given by the exponential fit. For SsoMCM (K323D/R440D), there is an increase in transition probabilities and a decrease in the observed dwell times in each state compared with WT (Fig. 6B-C). In all, this mutation is responsible for a large increase in excluded strand binding dynamics. The ExPRT program illustrates all observed dynamics on a single plot, which allows the reader to more easily gain greater insight into the measured interactions. The ExPRT plots have been used to characterize the dynamics of the excluded strand interactions with SsoMCM, EcDnaB, and Twinkle [22,68,69]. It captures changes in FRET states, transitions, and kinetics when altering the length of the excluded strand or when studying mutants designed to disrupt the interaction. The ExPRT analysis is the only single-molecule FRET analysis program that produces transition plots that simultaneously visualize explicit transitions, probability of transitions, and dwell times on a single plot. Moreover, the ExPRT program tests for and can visualize multiple rates that govern a transition. For these reasons, we believe that the ExPRT program may be a preferable method of analyzing and visualizing data compared to other smFRET analysis programs [73–76]. 3. Impact of the excluded strand in unwinding 3.1. Excluded strand as a ‘molecular ratchet’ Now that we and others have found that the excluded (nontranslocating) strand interacts with the exterior surface of a number of helicases, the question is why? How does the excluded strand contribute to the dsDNA unwinding mechanism? Generally, we have found that surface exposed and conserved positively charged residues define a path on the exterior surface [22,65,68]. Mutation of individual residues alter the DNA binding path and perturb DNA unwinding. Specifically for SsoMCM, a 30 –50 helicase,

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

8

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

Fig. 7. SEW models showing the impact on the excluded strand on unwinding. A) The ratchet is used by SsoMCM to stabilize unwinding in a forward direction. B) The regulator is used by SF4 helicases to control the unwinding rate. When engaged with the external surface, the unwinding rate is slowed. C) Various DNA helicase will recognize a variety of DNA damages and stall, directing downstream DNA repair processes.

mutation of positively charged residues along the proposed binding path reduce the unwinding efficiency 2–10 fold. For this helicase, loss of contact with the excluded strand makes dsDNA unwinding more futile. We hypothesize that maintaining a grip on both ssDNA strands allows for better destabilization of the hydrogen bonding within the duplex for efficient unwinding. Specific contacts within the central channel will propel the helicase forward along DNA, coupled with ATP hydrolysis steps [78–81]. However when contact with the excluded strand is lost, the helicase can slip backwards after an ATP hydrolysis cycle, effectively rezipping the duplex region, and ineffectively unwinding DNA. Therefore, the excluded strand interaction acts as a ‘molecular ratchet’ to promote SsoMCM unwinding in a forward direction and prevent slippage backwards (Fig. 7A). 3.2. Excluded strand as a ‘molecular brake’ Similarly, exterior excluded strand contracts are present for the SF4 50 –3 DNA helicase, EcDnaB [68], however, the effect on DNA unwinding may be opposite to that of SsoMCM. Mutation of exterior positively charged residues on EcDnaB resulted in increases in dsDNA unwinding rates. It is intriguing that the excluded strand contact may provide a ‘molecular brake’ to control the unwinding rate. It is also possible that a greater force is applied to the encircled strand by the EcDnaB motor domain to destabilize the duplex and that the excluded strand modulates that force. A greater force may be necessary thermodynamically at mesophilic temperatures for EcDnaB compared to thermophilic temperatures for SsoMCM where higher temperature can aid in destabilization of the duplex. It is clear for the exterior mutants of EcDnaB that show faster and more efficient unwinding, that loss of contact with the excluded strand released the imposed molecular restrictions rendering the helicase ungoverned with regards to unwinding (Fig. 7B). In addition to EcDnaB, other SF4 helicases, T7 gp4 and T4 gp41, have also been proposed to have an excluded-strand interaction based on unwinding rates that varied with the length of excluded-strand [82,83]. The T7 gp4-excluded-strand interaction was disrupted using hybrid DNA:morpholino substrates. Morpholinos carry no charge on their backbone, and would neutralize any electrostatic interaction a protein has with the phosphate sugar backbone. Unwinding studies showed that the T7 gp4 unwinding rate and amplitude were greatly enhanced when the morpholino was on the lagging strand [84]. This result is consistent with what has been observed for DnaB specifically that disrupting the electrostatic-based excluded-strand interaction greatly stimulates unwinding activity. It seems that for these SF4 helicase, the excluded-strand interaction acts to regulate the DNA unwinding rate, opposite to that for SsoMCM, despite the similarities of the interactions.

3.3. Nonhexameric helicases that engage both strands Even nonhexameric helicases that exist primarily as monomers or dimers have been shown to make contact with both DNA strands. SF1 helicases are comprised of at least two RecA-like domains, and are characterized by a conserved arrangement of several core motifs that are involved in ATP hydrolysis, DNA-binding, and ssDNA translocation. SF1A helicases translocate from 30 – 50 , and SF1B helicases translocate from 50 – 30 [3,85]. The best known example of SF1 helicases engaging both DNA strands is the bacterial recombination dependent helicase/nuclease RecBCD [86–88]. The RecB subunit is a SF1A helicase, and the RecD subunit is a SF1B helicase allowing unidirectional translocation along dsDNA with opposing polarities acting on each strand. The combination of domain motors allows engagement of both DNA strands to efficiently unwind the duplex in search of a Chi recognition site for recombination. Other model SF1A helicases include Rep, PcrA, and UvrD and are involved in rolling circle replication, replication fork progression, and repair of various DNA damage events [89–91]. These homologous helicases unwind DNA through an open to closed transition griping the DNA in an inchworm stepping mechanism [92–96]. The ‘closed’ form of the helicase corresponds to robust helicase activity, while the ‘open’ conformation is associated with a strand-switching event and translocation on the opposite strand effectively reannealing the duplex. This observed strand switching suggests that interactions between the helicase and DNA go beyond solely interacting with the translocating strand. The helicase must also interact with the duplex region or the nontranslocating strand in order to anchor the helicase through the flexible 2B domain to switch and prevent dissociation. The eukaryotic SF1B Pif1 helicase has been implicated in a number of activities including Okazaki fragment processing [97,98], repair of double-stranded DNA breaks [99–101], and telomere maintenance [102–104]. Pif1 has a preference for unwinding DNA:RNA over DNA:DNA duplexes due to enhanced processivity [105]. Additionally, measured dissociation rates for singlestranded tailed duplex substrates were 2–4 fold greater than those for measured for forked duplexes suggesting that an interaction with the nontranslocating strand was responsible. The increased processivity on forked DNA:RNA substrates could have implications for Pif1’s role inhibiting telomerase by removing the telomerase complex. Reduced processivity on dsDNA tailed substrates may be important for Pif1 binding to double-strand breaks and subsequently recruiting other repair proteins. Similarly, the monomeric SF1B T4 bacteriophage Dda helicase involved in DNA replication initiation during T4 phage infection [85,106,107] has been shown to interact with both the translocating and nontranslocating strand to enhance unwinding of DNA fork substrates [108,109].

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

Like SF1 helicases, SF2 helicases are characterized by a distinctive organization of ATPase motifs, however, SF2 helicases generally interact with their nucleic acid substrates mostly through the phosphodiester backbone, while SF1 helicases mainly utilize base stacking to mediate interactions [85,110,111]. The BLM helicase belongs to the RecQ group of SF2 helicases and mutations in this protein result in genomic instability, sunlight sensitivity, and the early onset of cancers. BLM is thought to function in homologous recombination, and its loss increases sister chromatid exchange [111]. Interestingly, human BLM also exhibits strandswitching. Single-molecule magnetic tweezing experiments show that BLM unwinding is interrupted by short rezipping events [112]. BLM unwinds 30 – 50 , but intermittently the two motor domains pause and loosen their interaction with the translocating strand. An accessory RQC domain maintains contact with the nontranslocating strand while BLM ‘slips’ backwards as the DNA fork rezips. A study of RecQ2 and RecQ3 from Arabidopsis thaliana also showed that these RecQ helicases also exhibit rewinding events by interacting with both the translocating and nontranslocating strands simultaneously [113]. The hepatitis virus C (HCV) NS3 (SF2) helicase processes the viral ssRNA genome, but it has also been proposed to interact with the host’s DNA genome [114,115]. NS3 directionally unwinds substrates with a 30 -overhang in the absence of ATP, but not 50 -tailed or blunt substrates [116]. It is proposed that NS3 diffuses on the 30 -tail until it encounters and engages the duplex junction where local basepair melting is trapped through an interaction with the translocating strand as well as interactions with the newly displaced 50 -nontranslocating strand. These additional interactions would provide specificity for forward unwinding and allow for additional NS3 molecules to load behind the leading and engaged helicase for more efficient strand separation. The dual interactions with both DNA strands measured for various SF1 and SF2 helicases may be required for the strand switching and unwinding activities of these helicases similar to hexameric helicases. 4. Sensing of DNA damage In addition to the apparent effects that interactions with the displaced strand have on functions such as strand switching, unwinding, and rewinding, interactions with the nontranslocating strand have been shown to detect DNA damage directly. Helicases perform important functions in a variety of DNA repair processes including nucleotide excision repair (NER), base excision repair (BER), homologous recombination (HR), and mismatch repair (MMR). Deficiencies or mutations in these helicases are clinically associated with genome instabilities in the forms of heightened predisposition to cancers and age-related genetic conditions such as Fanconi anemia, Werner syndrome, Bloom’s syndrome, and Xeroderma pigmentosum among others [117–120]. Recently, there have been multiple reports identifying an interaction between a DNA repair helicase and the nontranslocating strand which may have strong implications in the sensing of DNA damage (Fig. 7C). FANCJ is one of the 15 or so genes associated directly with Fanconi anemia (FA) that is primarily involved in recognizing and repairing interstrand DNA crosslinks that impede DNA replication processes. FANCJ is also known to directly interact with BRCA1 to resolve G-quadruplex structures [120–124]. A recent study has shown that the unwinding activity of FANCJ is completely disrupted when a single polyglycol linker is included in the sugar phosphate backbone of either the translocating or nontranslocating strand [125]. This suggests that FANCJ can sense the DNA backbone chemistry of both strands which is crucial for unwinding. However, when three adjacent abasic sites were included in either strand, unwinding was only partially inhibited when the abasic

9

sites were in the translocating strand. This result is consistent with a later study where FANCJ unwinding was only sensitive to a cyclopurine lesion in the translocating strand, and not in the nontranslocating strand [126]. Interestingly, FANCJ unwinding is also hindered by a thymine glycol site on either strand, but it is not sensitive to 8-oxoguanine on either strand [127]. Collectively, these data suggest a lesion-specific and strand-specific damage recognition mechanism (including the nontranslocating strand) by FANCJ that could direct various post-recognition processing pathways (Fig. 7C). Similar specific damage and strand recognition properties also exist for the XPD (SF2) helicase involved in eukaryotic NER [118]. The structure and unwinding kinetics of the archaeal homologue, XPD from Thermoplasma acidophilum (TaXPD) have been widely studied [128–132]. TaXPD to unwinds DNA substrates containing either a fluorescein-modified thymine or a cyclobutane pyrimidine dimer (CPD). Although XPD stalls when the fluorescein adduct is located on the translocating strand, it recognizes and stalls at CPD sites only when they are located on the nontranslocating strand [133]. This suggests that the helicase must interact with both strands, and monitors ssDNA for specific DNA damage on each strand. These distinct modes of DNA damage searching and recognition may play a role in mediating distinct responses from downstream repair machinery. The ability of SF4 hexameric replication helicases to unwind over damaged DNA has also been tested. EcDnaB showed no decrease in unwinding when a cyclopurine or thymine glycol was located on either the translocating or nontranslocating strand [126,127]. However, the related SF4 mitochondrial Twinkle helicase showed a decrease in unwinding when acting upon DNA substrates with a thymine glycol site in the translocating strand. This effect was not seen when the thymine glycol was present only on the nontranslocating strand. DNA substrates with a cyclopurine either in the translocating strand or nontranslocating strand were also tested. In these cases, Twinkle was able to unwind the substrate with cyclopurine in the translocating strand, but not when the damage was on the displaced strand [69]. This is another example where different mechanisms of damage recognition may lead to distinct downstream processing (Fig. 7C). 5. Role of the excluded strand in the unwindosome Interactions with the nontranslocating strand have been shown for replicative helicases from all three domains of life, SsoMCM, EcDnaB, and human Twinkle. The replication of genetic material is a complex and dynamic process, involving many protein components and interactions with various roles. Until recently, the displaced strand had only been considered a passive component at the replication fork. However, it seems that the excluded-strand serves a variety of functions that differ depending on the helicase being studied. At the fork, immediate strand separation will be maintained though interactions both within the central channel and the exterior surface. However, the extent of the SEW interaction of the excluded strand with the exterior of the helicase may be modulated by other interacting proteins within the replisome. In eukaryotes, the MCM2-7 helicase is in contact with additional subunits, Cdc45 and GINS, to form the CMG complex or unwindosome [134–138]. Although direct interactions of the excluded strand with the exterior surface of MCM2-7 have not been tested, an interaction of the excluded strand with Cdc45 have been visualized using EM [139,140]. Interestingly, CMG unwinding was stalled when molecular roadblocks were placed on the excluded lagging strand [141]. The authors note that the presence of a bulky lesion on the excluded-strand could cause stalling if the outer surface of CMG interacts closely with the lagging strand. A

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

10

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

recently reported secondary channel or groove has been characterized when the MCM2-MCM5 ‘gate’ is closed that could potentially accommodate the excluded-strand [140]. When closed, this creates a second channel bound by the outer surface of MCM2-7 and the inner surfaces of GINS and Cdc45. It is possible that during active unwinding by CMG, the leading strand is restricted to the interior channel of MCM2-7, while the lagging strand is separated and fed into this secondary exterior channel. This would provide for strand sequestration and prevent reannealing. It may also provide a mechanism to hand-off the lagging strand to the DNA primase for priming. In archaea and SV40, and similar to the bacterial systems, the DNA primase has been shown to directly interact with the helicase to mediate hand-off the displaced lagging strand for efficient priming [142–144]. Interestingly, an EM structure of the minimal eukaryotic replisome, places Pol e in front of the CMG complex contacting the leading strand, while the lagging strand traverses the length of the MCM2-7 complex to contact the Pola-primase complex afterwards for priming [145]. The architecture of the eukaryotic replisome is still being revealed, and the preliminary studies are pointing towards replication fork organization and dynamics that are more complex than bacterial and viral model systems [145–148]. However, based on the various roles and interactions between the nontranslocating strand and the CMG helicase, it would not be surprising to find that such an interaction is important in coordinating function at the replication fork. Clearly, higher resolution data is required to determine the precise paths of both strands through the MCM2-7 helicase and within the eukaryotic replisome for better understanding. It has been reported that dsDNA unwinding by T7 gp4 helicase is greatly stimulated by the presence of the DNA polymerase placed very close to the duplex junction so that little to no ssDNA is exposed [19,149]. However, in the absence of the polymerase or when the DNA polymerase stalls at a lesion, T7 gp4 can continue unwinding at a slower rate possibly engaging the excluded strand as a molecular brake [150]. During rapid synthesis, SEW interactions with the external surface are prevented by the polymerase thus stimulating coupled unwinding and replication in T7. A similar model has been proposed for the coupling of T4 gp41 helicase and gp43 polymerase and Escherichia coli DnaB and Pol III at the replication fork [151,152]. For these 50 –30 lagging strand translocating helicases, the excluded leading strand would be disengaged from the helicase in favor of promoting synthesis within the leading strand DNA polymerase. The binding and unbinding of the excluded strand to the exterior of the helicase may regulate the speed of unwinding and replication by coupling helicase activity to the polymerase (Fig. 7B). As an example, incorporation of abasic DNA lesions on either the leading or lagging strand causes the uncoupling of leading and lagging strand synthesis in E. coli. Leading strand synthesis is halted, while lagging strand synthesis continues in 2/3 of cases for up to 1 kb past the point of uncoupling at a slower rate [153]. Studies of the E. coli replisome have also shown that the speed of replication fork migration and helicase unwinding is regulated by the other components of the replisome such as primases and polymerases. For example, inclusion of the DnaG primase reduces the processivity of leading strand synthesis upon interaction with DnaB preventing uncoupling of leading and lagging strand synthesis when priming is rate limiting [154]. Furthermore, inclusion of translesion polymerases, Pol II and Pol IV, slow the DnaB helicase compared to solely the leading strand polymerase, Pol III [155]. It may be that more contact with the excluded strand on the exterior of DnaB slows the replisome to allow time for DNA polymerase switching and translesion synthesis. It is possible that the excluded-strand interaction takes place after the leading strand polymerase becomes uncoupled, and acts to slow the rest of the replisome to a stop by acting as an electrostatic brake to

unwinding (Fig. 7B). This would reduce the amount of vulnerable ssDNA produced by helicase unwinding, stimulating proper repair before continuing with normal replication and releasing the brake. The excluded-strand interaction may also provide a platform for the accessory helicase, Rep, to bind and interact to restart stalled replication forks. Rep unwinds 30 – 50 , opposite to the polarity of DnaB helicase, translocating on the leading strand instead. Rep and DnaB physically interact and unwind forks cooperatively [156]. An important finding was that there must be ssDNA available on the leading strand for Rep to interact and subsequently unwind cooperatively with DnaB. This ssDNA may be interacting with DnaB’s exterior surface until assembly of Rep on the leading strand stimulates fork restart. These studies suggest that the unwinding activity of DnaB is a highly regulated component of the replisome and that the excluded-strand interaction can control the speed of the replisome to direct dynamic processes at the fork.

6. Final thoughts At the heart of the replisome is the hexameric DNA helicase. Assembly of the hexameric helicase at origins controls the initiation of replication, and the rate of unwinding controls elongation, both proposed to require interactions with the excluded strand. Independent of unwinding polarity, dynamic interactions with the excluded nontranslocating strand have now been shown to control the DNA unwinding rate, sensing of DNA damage, and modulating interactions with other proteins within the replisome (Fig. 7). Higher resolution studies are required to continue mapping ssDNA interactions on both the interior and exterior of DNA helicases. As the excluded strand has now been revealed to have an almost equal or greater importance than the translocating strand, future experiments will need to test contributions of both strands within the replisome to determine and differentiate interactions directing DNA unwinding and sensing. Acknowledgements The publication of this work was supported by American Cancer Society (RSG-11-049-01-DMC) and Baylor University. I am thankful to Brian Graham, Grant Schauer, Sanford Leuba, and Alan Marshall whose work and collaboration has provided for some of the data and conclusions presented in this article. References [1] J.P. Erzberger, J.M. Berger, Evolutionary relationships and structural mechanisms of AAA+ proteins, Annu. Rev. Biophys. Biomol. Struct. 35 (2006) 93–114. [2] A.F. Neuwald, L. Aravind, J.L. Spouge, E.V. Koonin, AAA+: A class of chaperonelike ATPases associated with the assembly, operation, and disassembly of protein complexes, Genome Res. 9 (1999) 27–43. [3] M.R. Singleton, M.S. Dillingham, D.B. Wigley, Structure and mechanism of helicases and nucleic acid translocases, Annu. Rev. Biochem. 76 (2007) 23–50. [4] M.L. Bochman, A. Schwacha, The Mcm complex: unwinding the mechanism of a replicative helicase, Microbiol. Mol. Biol. Rev. 73 (2009) 652–683. [5] M.A. Trakselis, Structural mechanisms of hexameric helicase loading, assembly, and unwinding, F1000Res. (2016) 5. [6] S.S. Patel, K.M. Picha, Structure and function of hexameric helicases, Annu. Rev. Biochem. 69 (2000) 651–697. [7] D.L. Kaplan, M.J. Davey, M. O’Donnell, Mcm 4,6,7 uses a ‘‘pump in ring” mechanism to unwind DNA by steric exclusion and actively translocate along a duplex, J. Biol. Chem. 278 (2003) 49171–49182. [8] I. Cuesta, R. Nunez-Ramirez, S.H. Scheres, D. Gai, X.S. Chen, E. Fanning, J.M. Carazo, Conformational rearrangements of SV40 large T antigen during early replication events, J. Mol. Biol. 397 (2010) 1276–1286. [9] D. Gai, R. Zhao, D. Li, C.V. Finkielstein, X.S. Chen, Mechanisms of conformational change for a replicative hexameric helicase of SV40 large tumor antigen, Cell 119 (2004) 47–60. [10] A.E. Gorbalenya, E.V. Koonin, Viral proteins containing the purine NTPbinding sequence pattern, Nucleic Acids Res. 17 (1989) 8413–8440.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx [11] M.R. Singleton, M.R. Sawaya, T. Ellenberger, D.B. Wigley, Crystal structure of T7 gene 4 ring helicase indicates a mechanism for sequential hydrolysis of nucleotides, Cell 101 (2000) 589–600. [12] N.D. Thomsen, J.M. Berger, Running in reverse: the structural basis for translocation polarity in hexameric helicases, Cell 139 (2009) 523–534. [13] J.C. Liao, Y.J. Jeong, D.E. Kim, S.S. Patel, G. Oster, Mechanochemistry of T7 DNA helicase, J. Mol. Biol. 350 (2005) 452–475. [14] J.M. Miller, B.T. Arachea, L.B. Epling, E.J. Enemark, Analysis of the crystal structure of an active MCM hexamer, Elife 3 (2014) e03433. [15] E.J. Enemark, L. Joshua-Tor, Mechanism of DNA translocation in a replicative hexameric helicase, Nature 442 (2006) 270–275. [16] A.S. Brewster, G. Wang, X. Yu, W.B. Greenleaf, J.M. Carazo, M. Tjajadia, M.G. Klein, X.S. Chen, Crystal structure of a near-full-length archaeal MCM: functional insights for an AAA+ hexameric helicase, Proc. Natl. Acad. Sci. U.S.A. 105 (2008) 20191–20196. [17] O. Itsathitphaisarn, R.A. Wing, W.K. Eliason, J. Wang, T.A. Steitz, The hexameric helicase DnaB adopts a nonplanar conformation during translocation, Cell 151 (2012) 267–277. [18] A. Costa, L. Renault, P. Swuec, T. Petojevic, J.J. Pesavento, I. Ilves, K. MacLellanGibson, R.A. Fleck, M.R. Botchan, J.M. Berger, DNA binding polarity, dimerization, and ATPase ring remodeling in the CMG helicase of the eukaryotic replisome, Elife 3 (2014) e03273. [19] M. Pandey, S.S. Patel, Helicase and polymerase move together close to the fork junction and copy DNA in one-nucleotide steps, Cell Rep. 6 (2014) 1129– 1138. [20] C.A. Froelich, S. Kang, L.B. Epling, S.P. Bell, E.J. Enemark, A conserved MCM single-stranded DNA binding element is essential for replication initiation, Elife 3 (2014) e01993. [21] R. Galletto, M.J. Jezewska, W. Bujalowski, Unzipping mechanism of the double-stranded DNA unwinding by a hexameric helicase: the effect of the 30 arm and the stability of the dsDNA on the unwinding activity of the Escherichia coli DnaB helicase, J. Mol. Biol. 343 (2004) 101–114. [22] B.W. Graham, G.D. Schauer, S.H. Leuba, M.A. Trakselis, Steric exclusion and wrapping of the excluded DNA strand occurs along discrete external binding paths during MCM helicase unwinding, Nucleic Acids Res. 39 (2011) 6585– 6595. [23] M.J. Jezewska, S. Rajendran, D. Bujalowska, W. Bujalowski, Does singlestranded DNA pass through the inner channel of the protein hexamer in the complex with the Escherichia coli DnaB Helicase? Fluorescence energy transfer studies, J. Biol. Chem. 273 (1998) 10515–10529. [24] W. Bujalowski, M.J. Jezewska, Interactions of Escherichia coli primary replicative helicase DnaB protein with single-stranded DNA. The nucleic acid does not wrap around the protein hexamer, Biochemistry 34 (1995) 8513–8519. [25] M.M. Hingorani, S.S. Patel, Interactions of bacteriophage T7 DNA primase/ helicase protein with single-stranded and double-stranded DNAs, Biochemistry 32 (1993) 12478–12487. [26] M.J. Jezewska, U.S. Kim, W. Bujalowski, Interactions of Escherichia coli primary replicative helicase DnaB protein with nucleotide cofactors, Biophys. J. 71 (1996) 2075–2086. [27] T.D. Tullius, Physical studies of protein-DNA complexes by footprinting, Annu. Rev. Biophys. Biophys. Chem. 18 (1989) 213–237. [28] T.D. Tullius, B.A. Dombroski, M.E. Churchill, L. Kam, Hydroxyl radical footprinting: a high-resolution method for mapping protein-DNA contacts, Methods Enzymol. 155 (1987) 537–558. [29] D.S. Sigman, D.R. Graham, V. D’Aurora, A.M. Stern, Oxygen-dependent cleavage of DNA by the 1,10-phenanthroline cuprous complex. Inhibition of Escherichia coli DNA polymerase I, J. Biol. Chem. 254 (1979) 12269–12272. [30] D.J. Galas, A. Schmitz, DNAse footprinting: a simple method for the detection of protein-DNA binding specificity, Nucleic Acids Res. 5 (1978) 3157–3170. [31] A.J. Hampshire, D.A. Rusling, V.J. Broughton-Head, K.R. Fox, Footprinting: a method for determining the sequence selectivity, affinity and kinetics of DNA-binding ligands, Methods 42 (2007) 128–140. [32] S.S. Jain, T.D. Tullius, Footprinting protein-DNA complexes using the hydroxyl radical, Nat. Protoc. 3 (2008) 1092–1100. [33] T. Moss, B.t. Leblanc, DNA-Protein Interactions: Principles and Protocols, 3rd ed., Humana Press, New York, 2009. [34] V.E. Fadouloglou, M. Kokkinidis, N.M. Glykos, Determination of protein oligomerization state: two approaches based on glutaraldehyde crosslinking, Anal. Biochem. 373 (2008) 404–406. [35] P. Slusarewicz, K. Zhu, T. Hedman, Kinetic characterization and comparison of various protein crosslinking reagents for matrix modification, J. Mater. Sci. – Mater. Med. 21 (2010) 1175–1181. [36] J.R. Kuykendall, M.S. Bogdanffy, Efficiency of DNA-histone crosslinking induced by saturated and unsaturated aldehydes in vitro, Mutat. Res. 283 (1992) 131–136. [37] R.C. Adami, K.G. Rice, Metabolic stability of glutaraldehyde cross-linked peptide DNA condensates, J. Pharm. Sci. 88 (1999) 739–746. [38] I. Migneault, C. Dartiguenave, M.J. Bertrand, K.C. Waldron, Glutaraldehyde: behavior in aqueous solution, reaction with proteins, and application to enzyme crosslinking, Biotechniques 37 (790–796) (2004) 798–802. [39] Y. Wine, N. Cohen-Hadar, A. Freeman, F. Frolow, Elucidation of the mechanism and end products of glutaraldehyde crosslinking reaction by X-ray structure analysis, Biotechnol. Bioeng. 98 (2007) 711–718.

11

[40] O. Aparicio, J.V. Geisberg, K. Struhl, Chromatin Immunoprecipitation for Determining the Association of Proteins with Specific Genomic Sequences in Vivo, John Wiley & Sons Inc., 2001. [41] J.D. McGhee, P.H. von Hippel, Formaldehyde as a probe of DNA structure. II. Reaction with endocyclic imino groups of DNA bases, Biochemistry 14 (1975) 1297–1303. [42] J.D. McGhee, P.H. von Hippel, Formaldehyde as a probe of DNA structure. I. Reaction with exocyclic amino groups of DNA bases, Biochemistry 14 (1975) 1281–1296. [43] K. Peters, F.M. Richards, Chemical cross-linking: reagents and problems in studies of membrane structure, Annu. Rev. Biochem. 46 (1977) 523–551. [44] T.M. Dietz, T.H. Koch, Photochemical coupling of 5-bromouracil to tryptophan, tyrosine and histidine, peptide-like derivatives in aqueous fluid solution, Photochem. Photobiol. 46 (1987) 971–978. [45] Q. Dong, E.E. Blatter, Y.W. Ebright, K. Bister, R.H. Ebright, Identification of amino acid-base contacts in the Myc-DNA complex by site-specific bromouracil mediated photocrosslinking, EMBO J. 13 (1994) 200–204. [46] L.A. Chodosh, UV crosslinking of proteins to nucleic acids, Curr. Protoc. Mol. Biol. (2001). Chapter 12, Unit 12 15. [47] B. Schilling, R.H. Row, B.W. Gibson, X. Guo, M.M. Young, MS2Assign, automated assignment and nomenclature of tandem mass spectra of chemically crosslinked peptides, J. Am. Soc. Mass Spectrom. 14 (2003) 834– 850. [48] C.M. Santiveri, Y. Mirassou, P. Rico-Lastres, S. Martinez-Lumbreras, J.M. PerezCanadillas, Pub1p C-terminal RRM domain interacts with Tif4631p through a conserved region neighbouring the Pab1p binding site, PLoS ONE 6 (2011) e24481. [49] M.C. Golden, K.A. Resing, B.D. Collins, M.C. Willis, T.H. Koch, Mass spectral characterization of a protein-nucleic acid photocrosslink, Protein Sci. 8 (1999) 2806–2812. [50] R.A. Rieger, M.M. McTigue, J.H. Kycia, S.E. Gerchman, A.P. Grollman, C.R. Iden, Characterization of a cross-linked DNA-endonuclease VIII repair complex by electrospray ionization mass spectrometry, J. Am. Soc. Mass Spectrom. 11 (2000) 505–515. [51] H. Steen, O.N. Jensen, Analysis of protein-nucleic acid interactions by photochemical cross-linking and mass spectrometry, Mass Spectrom. Rev. 21 (2002) 163–182. [52] C.E. Doneanu, P.R. Gafken, S.E. Bennett, D.F. Barofsky, Mass spectrometry of UV-cross-linked protein-nucleic acid complexes: identification of amino acid residues in the single-stranded DNA-binding domain of human replication protein A, Anal. Chem. 76 (2004) 5667–5676. [53] P.R. Gafken, C.E. Doneanu, S.E. Bennett, D.F. Barofsky, Comparison of ESI-MS interfaces for the analysis of UV-crosslinked peptide-nucleic acid complexes, J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 860 (2007) 145–152. [54] K. Kramer, T. Sachsenberg, B.M. Beckmann, S. Qamar, K.L. Boon, M.W. Hentze, O. Kohlbacher, H. Urlaub, Photo-cross-linking and high-resolution mass spectrometry for assignment of RNA-binding sites in RNA-binding proteins, Nat. Methods 11 (2014) 1064–1070. [55] A. Leitner, T. Walzthoeni, A. Kahraman, F. Herzog, O. Rinner, M. Beck, R. Aebersold, Probing native protein structures by chemical cross-linking, mass spectrometry, and bioinformatics, Mol. Cell. Proteomics 9 (2010) 1634–1649. [56] A. Maiolica, D. Cittaro, D. Borsotti, L. Sennels, C. Ciferri, C. Tarricone, A. Musacchio, J. Rappsilber, Structural analysis of multiprotein complexes by cross-linking, mass spectrometry, and database searching, Mol. Cell. Proteomics 6 (2007) 2200–2211. [57] Z. Zhang, D.L. Smith, Determination of amide hydrogen exchange by mass spectrometry: a new tool for protein structure elucidation, Protein Sci. 2 (1993) 522–531. [58] G.F. Pirrone, R.E. Iacob, J.R. Engen, Applications of hydrogen/deuterium exchange MS from 2012 to 2014, Anal. Chem. 87 (2015) 99–118. [59] S.W. Englander, Hydrogen exchange and mass spectrometry: a historical perspective, J. Am. Soc. Mass Spectrom. 17 (2006) 1481–1489. [60] J. Lisal, D.E. Kainov, T.T. Lam, M.R. Emmett, H. Wei, P. Gottlieb, A.G. Marshall, R. Tuma, Interaction of packaging motor with the polymerase complex of dsRNA bacteriophage, Virology 351 (2006) 73–79. [61] P. Nevin, J.R. Engen, P.J. Beuning, Steric gate residues of Y-family DNA polymerases DinB and pol kappa are crucial for dNTP-induced conformational change, DNA Repair (Amst) 29 (2015) 65–73. [62] M.J. Chalmers, S.A. Busby, B.D. Pascal, G.M. West, P.R. Griffin, Differential hydrogen/deuterium exchange mass spectrometry analysis of protein-ligand interactions, Expert Rev. Proteomics 8 (2011) 43–59. [63] T.M. Schaub, C.L. Hendrickson, S. Horning, J.P. Quinn, M.W. Senko, A.G. Marshall, High-performance mass spectrometry: Fourier transform ion cyclotron resonance at 14.5 Tesla, Anal. Chem. 80 (2008) 3985–3990. [64] G.T. Blakney, C.L. Hendrickson, A.G. Marshall, Predator data station: a fast data acquisition system for advanced FT-ICR MS experiments, Int. J. Mass Spectrom. 306 (2011) 246–252. [65] B.W. Graham, Y. Tao, K.L. Dodge, C.T. Thaxton, D. Olaso, N.L. Young, A.G. Marshall, M.A. Trakselis, DNA interactions probed by H/D exchange FT-ICR mass spectrometry confirm external binding sites on the MCM helicase. J. Biol. Chem., (Epub ahead of print) http://dx.doi.org/10.1074/jbc.M116. 719591, in press. [66] J. Zheng, H.Y. Yong, N. Panutdaporn, C. Liu, K. Tang, D. Luo, High-resolution HDX-MS reveals distinct mechanisms of RNA recognition and activation by RIG-I and MDA5, Nucleic Acids Res. 43 (2015) 1216–1230.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

12

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx

[67] E. Rothenberg, M.A. Trakselis, S.D. Bell, T. Ha, MCM forked substrate specificity involves dynamic interaction with the 50 -tail, J. Biol. Chem. 282 (2007) 34229–34234. [68] S.M. Carney, H.N. McFarland, S.H. Leuba, M.A. Trakselis, Exterior excluded strand interactions regulate DNA unwinding by replicative hexameric helicases. Nucleic Acids Res., in revision. [69] I. Khan, J.D. Crouch, S.K. Bharti, J.A. Sommers, S.M. Carney, E. Yakubovskaya, M. Garcia-Diaz, M.A. Trakselis, R.M. Brosh, Biochemical characterization of the human mitochondrial replicative Twinkle helicase: substrate specificity, DNA branch-migration, and ability to overcome blocades to DNA unwinding, J. Biol Chem., accepted. [70] R. Roy, S. Hohng, T. Ha, A practical guide to single-molecule FRET, Nat. Methods 5 (2008) 507–516. [71] M.V. Fagerburg, S.H. Leuba, Optimal practices for surface-tethered single molecule total internal reflection fluorescence resonance energy transfer analysis, in: G. Zuccheri, B. Samori (Eds.), DNA Nanotechnology, Methods and Protocols, 1st ed., Humana Press, New York, NY, 2011, p. 361. [72] C. Joo, T. Ha, Single-molecule FRET with total internal reflection microscopy, Cold Spring Harbor Protoc. (2012). [73] S.A. McKinney, C. Joo, T. Ha, Analysis of single-molecule FRET trajectories using hidden Markov modeling, Biophys. J. 91 (2006) 1941–1951. [74] C. Joo, S.A. McKinney, M. Nakamura, I. Rasnik, S. Myong, T. Ha, Real-time observation of RecA filament dynamics with single monomer resolution, Cell 126 (2006) 515–527. [75] J. Abelson, M. Blanco, M.A. Ditzler, F. Fuller, P. Aravamudhan, M. Wood, T. Villa, D.E. Ryan, J.A. Pleiss, C. Maeder, C. Guthrie, N.G. Walter, Conformational dynamics of single pre-mRNA molecules during in vitro splicing, Nat. Struct. Mol. Biol. 17 (2010) 504–512. [76] M. Blanco, N.G. Walter, Analysis of complex single-molecule FRET time trajectories, Methods Enzymol. 472 (2010) 153–178. [77] J.E. Bronson, J. Fei, J.M. Hofman, R.L. Gonzalez Jr., C.H. Wiggins, Learning rates and states from biophysical time series: a Bayesian approach to model selection and single-molecule FRET data, Biophys. J. 97 (2009) 3196–3205. [78] E.R. Barry, J.E. Lovett, A. Costa, S.M. Lea, S.D. Bell, Intersubunit allosteric communication mediated by a conserved loop in the MCM helicase, Proc. Natl. Acad. Sci. U.S.A. 106 (2009) 1051–1056. [79] E.R. Barry, A.T. McGeoch, Z. Kelman, S.D. Bell, Archaeal MCM has separable processivity, substrate choice and helicase domains, Nucleic Acids Res. 35 (2007) 988–998. [80] M.J. Moreau, A.T. McGeoch, A.R. Lowe, L.S. Itzhaki, S.D. Bell, ATPase site architecture and helicase mechanism of an archaeal MCM, Mol. Cell 28 (2007) 304–314. [81] A.T. McGeoch, M.A. Trakselis, R.A. Laskey, S.D. Bell, Organization of the archaeal MCM complex on DNA and implications for the helicase mechanism, Nat. Struct. Mol. Biol. 12 (2005) 756–762. [82] P. Ahnert, S.S. Patel, Asymmetric interactions of hexameric bacteriophage T7 DNA helicase with the 50 - and 30 -tails of the forked DNA substrate, J. Biol. Chem. 272 (1997) 32267–32273. [83] R.W. Richardson, N.G. Nossal, Characterization of the bacteriophage T4 gene 41 DNA helicase, J. Biol. Chem. 264 (1989) 4725–4731. [84] Y.J. Jeong, V. Rajagopal, S.S. Patel, Switching from single-stranded to doublestranded DNA limits the unwinding processivity of ring-shaped T7 DNA helicase, Nucleic Acids Res. 41 (2013) 4219–4229. [85] M.E. Fairman-Williams, U.P. Guenther, E. Jankowsky, SF1 and SF2 helicases: family matters, Curr. Opin. Struct. Biol. 20 (2010) 313–324. [86] M. Spies, I. Amitani, R.J. Baskin, S.C. Kowalczykowski, RecBCD enzyme switches lead motor subunits in response to chi recognition, Cell 131 (2007) 694–705. [87] M.R. Singleton, M.S. Dillingham, M. Gaudier, S.C. Kowalczykowski, D.B. Wigley, Crystal structure of RecBCD enzyme reveals a machine for processing DNA breaks, Nature 432 (2004) 187–193. [88] M.S. Dillingham, M. Spies, S.C. Kowalczykowski, RecBCD enzyme is a bipolar DNA helicase, Nature 423 (2003) 893–897. [89] I. Husain, B. Van Houten, D.C. Thomas, M. Abdel-Monem, A. Sancar, Effect of DNA polymerase I and DNA helicase II on the turnover rate of UvrABC excision nuclease, Proc. Natl. Acad. Sci. U.S.A. 82 (1985) 6774–6778. [90] J.E. Yancey-Wrona, S.W. Matson, Bound Lac repressor protein differentially inhibits the unwinding reactions catalyzed by DNA helicases, Nucleic Acids Res. 20 (1992) 6713–6721. [91] S. Iordanescu, J. Bargonetti, Staphylococcus aureus chromosomal mutations that decrease efficiency of Rep utilization in replication of pT181 and related plasmids, J. Bacteriol. 171 (1989) 4501–4503. [92] S.S. Velankar, P. Soultanas, M.S. Dillingham, H.S. Subramanya, D.B. Wigley, Crystal structures of complexes of PcrA DNA helicase with a DNA substrate indicate an inchworm mechanism, Cell 97 (1999) 75–84. [93] E.J. Tomko, C.J. Fischer, A. Niedziela-Majka, T.M. Lohman, A nonuniform stepping mechanism for E. coli UvrD monomer translocation along singlestranded DNA, Mol. Cell 26 (2007) 335–347. [94] S. Arslan, R. Khafizov, C.D. Thomas, Y.R. Chemla, T. Ha, Protein structure. Engineering of a superhelicase through conformational control, Science 348 (2015) 344–347. [95] M.J. Comstock, K.D. Whitley, H. Jia, J. Sokoloski, T.M. Lohman, T. Ha, Y.R. Chemla, Protein structure. Direct observation of structure-function relationship in a nucleic acid-processing enzyme, Science 348 (2015) 352– 354.

[96] M.N. Dessinges, T. Lionnet, X.G. Xi, D. Bensimon, V. Croquette, Singlemolecule assay reveals strand switching and enhanced processivity of UvrD, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 6439–6444. [97] M.L. Rossi, J.E. Pike, W. Wang, P.M. Burgers, J.L. Campbell, R.A. Bambara, Pif1 helicase directs eukaryotic Okazaki fragments toward the two-nuclease cleavage pathway for primer removal, J. Biol. Chem. 283 (2008) 27483– 27493. [98] M.E. Budd, C.C. Reis, S. Smith, K. Myung, J.L. Campbell, Evidence suggesting that Pif1 helicase functions in DNA replication with the Dna2 helicase/ nuclease and DNA polymerase delta, Mol. Cell. Biol. 26 (2006) 2490–2500. [99] N. Saini, S. Ramakrishnan, R. Elango, S. Ayyar, Y. Zhang, A. Deem, G. Ira, J.E. Haber, K.S. Lobachev, A. Malkova, Migrating bubble during break-induced replication drives conservative DNA synthesis, Nature 502 (2013) 389–392. [100] K. Myung, C. Chen, R.D. Kolodner, Multiple pathways cooperate in the suppression of genome instability in Saccharomyces cerevisiae, Nature 411 (2001) 1073–1076. [101] M.A. Wilson, Y. Kwon, Y. Xu, W.H. Chung, P. Chi, H. Niu, R. Mayle, X. Chen, A. Malkova, P. Sung, G. Ira, Pif1 helicase and Poldelta promote recombinationcoupled DNA synthesis via bubble migration, Nature 502 (2013) 393–396. [102] J. Zhou, E.K. Monson, S.C. Teng, V.P. Schulz, V.A. Zakian, Pif1p helicase, a catalytic inhibitor of telomerase in yeast, Science 289 (2000) 771–774. [103] J.B. Boule, L.R. Vega, V.A. Zakian, The yeast Pif1p helicase removes telomerase from telomeric DNA, Nature 438 (2005) 57–61. [104] V.P. Schulz, V.A. Zakian, The saccharomyces PIF1 DNA helicase inhibits telomere elongation and de novo telomere formation, Cell 76 (1994) 145– 155. [105] S. Chib, A.K. Byrd, K.D. Raney, Yeast helicase Pif1 unwinds RNA:DNA hybrids with higher processivity than DNA:DNA duplexes, J. Biol. Chem. 291 (2016) 5889–5901. [106] P. Gauss, K. Park, T.E. Spencer, K.J. Hacker, DNA helicase requirements for DNA replication during bacteriophage T4 infection, J. Bacteriol. 176 (1994) 1667–1672. [107] J. Barry, B. Alberts, A role for two DNA helicases in the replication of T4 bacteriophage DNA, J. Biol. Chem. 269 (1994) 33063–33068. [108] S. Aarattuthodiyil, A.K. Byrd, K.D. Raney, Simultaneous binding to the tracking strand, displaced strand and the duplex of a DNA fork enhances unwinding by Dda helicase, Nucleic Acids Res. 42 (2015) 11707–11720. [109] R.L. Eoff, K.D. Raney, Intermediates revealed in the kinetic mechanism for DNA unwinding by a monomeric helicase, Nat. Struct. Mol. Biol. 13 (2006) 242–249. [110] K.D. Raney, A.K. Byrd, S. Aarattuthodiyil, Structure and mechanisms of SF1 DNA helicases, Adv. Exp. Med. Biol. 973 (2013) E1. [111] D.C. Beyer, M.K. Ghoneim, M. Spies, Structure and mechanisms of SF2 DNA helicases, Adv. Exp. Med. Biol. 767 (2013) 47–73. [112] S. Wang, W. Qin, J.H. Li, Y. Lu, K.Y. Lu, D.G. Nong, S.X. Dou, C.H. Xu, X.G. Xi, M. Li, Unwinding forward and sliding back: an intermittent unwinding mode of the BLM helicase, Nucleic Acids Res. 43 (2015) 3736–3746. [113] D. Klaue, D. Kobbe, F. Kemmerich, A. Kozikowska, H. Puchta, R. Seidel, Fork sensing and strand switching control antagonistic activities of RecQ helicases, Nat. Commun. 4 (2013) 2024. [114] R.K. Beran, B.D. Lindenbach, A.M. Pyle, The NS4A protein of hepatitis C virus promotes RNA-coupled ATP hydrolysis by the NS3 helicase, J. Virol. 83 (2009) 3268–3275. [115] A.M. Lam, D.N. Frick, Hepatitis C virus subgenomic replicon requires an active NS3 RNA helicase, J. Virol. 80 (2006) 404–411. [116] K.A. Reynolds, C.E. Cameron, K.D. Raney, Melting of duplex DNA in the absence of ATP by the NS3 helicase domain through specific interaction with a single-strand/double-strand junction, Biochemistry 54 (2015) 4248–4258. [117] N.B. Larsen, I.D. Hickson, RecQ helicases: conserved guardians of genomic integrity, Adv. Exp. Med. Biol. 767 (2013) 161–184. [118] J. Kuper, C. Kisker, DNA helicases in NER, BER, and MMR, Adv. Exp. Med. Biol. 767 (2013) 203–224. [119] J.M. Daley, H. Niu, P. Sung, Roles of DNA helicases in the mediation and regulation of homologous recombination, Adv. Exp. Med. Biol. 767 (2013) 185–202. [120] A.N. Suhasini, R.M. Brosh Jr., DNA helicases associated with genetic instability, cancer, and aging, Adv. Exp. Med. Biol. 767 (2013) 123–144. [121] Y. Wu, A.N. Suhasini, R.M. Brosh Jr., Welcome the family of FANCJ-like helicases to the block of genome stability maintenance proteins, Cell. Mol. Life Sci.: CMLS 66 (2009) 1209–1222. [122] S.B. Cantor, D.W. Bell, S. Ganesan, E.M. Kass, R. Drapkin, S. Grossman, D.C. Wahrer, D.C. Sgroi, W.S. Lane, D.A. Haber, D.M. Livingston, BACH1, a novel helicase-like protein, interacts directly with BRCA1 and contributes to its DNA repair function, Cell 105 (2001) 149–160. [123] T.B. London, L.J. Barber, G. Mosedale, G.P. Kelly, S. Balasubramanian, I.D. Hickson, S.J. Boulton, K. Hiom, FANCJ is a structure-specific DNA helicase associated with the maintenance of genomic G/C tracts, J. Biol. Chem. 283 (2008) 36132–36139. [124] Y. Wu, K. Shin-ya, R.M. Brosh Jr., FANCJ helicase defective in Fanconia anemia and breast cancer unwinds G-quadruplex DNA to defend genomic stability, Mol. Cell. Biol. 28 (2008) 4116–4128. [125] R. Gupta, S. Sharma, K.M. Doherty, J.A. Sommers, S.B. Cantor, R.M. Brosh Jr., Inhibition of BACH1 (FANCJ) helicase by backbone discontinuity is overcome by increased motor ATPase or length of loading strand, Nucleic Acids Res. 34 (2006) 6673–6683.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008

S.M. Carney, M.A. Trakselis / Methods xxx (2016) xxx–xxx [126] I. Khan, A.N. Suhasini, T. Banerjee, J.A. Sommers, D.L. Kaplan, J. Kuper, C. Kisker, R.M. Brosh Jr., Impact of age-associated cyclopurine lesions on DNA repair helicases, PLoS ONE 9 (2014) e113293. [127] A.N. Suhasini, J.A. Sommers, A.C. Mason, O.N. Voloshin, R.D. Camerini-Otero, M.S. Wold, R.M. Brosh Jr., FANCJ helicase uniquely senses oxidative base damage in either strand of duplex DNA and is stimulated by replication protein A to unwind the damaged DNA substrate in a strand-specific manner, J. Biol. Chem. 284 (2009) 18458–18470. [128] S.C. Wolski, J. Kuper, P. Hanzelmann, J.J. Truglio, D.L. Croteau, B. Van Houten, C. Kisker, Crystal structure of the FeS cluster-containing nucleotide excision repair helicase XPD, PLoS Biol. 6 (2008) e149. [129] H. Liu, J. Rudolf, K.A. Johnson, S.A. McMahon, M. Oke, L. Carter, A.M. McRobbie, S.E. Brown, J.H. Naismith, M.F. White, Structure of the DNA repair helicase XPD, Cell 133 (2008) 801–812. [130] L. Fan, J.O. Fuss, Q.J. Cheng, A.S. Arvai, M. Hammel, V.A. Roberts, P.K. Cooper, J. A. Tainer, XPD helicase structures and activities: insights into the cancer and aging phenotypes from XPD mutations, Cell 133 (2008) 789–800. [131] Z. Qi, R.A. Pugh, M. Spies, Y.R. Chemla, Sequence-dependent base pair stepping dynamics in XPD helicase unwinding, Elife 2 (2013) e00334. [132] R.A. Pugh, Y. Lin, C. Eller, H. Leesley, I.K. Cann, M. Spies, Ferroplasma acidarmanus RPA2 facilitates efficient unwinding of forked DNA substrates by monomers of FacXPD helicase, J. Mol. Biol. 383 (2008) 982–998. [133] C.N. Buechner, K. Heil, G. Michels, T. Carell, C. Kisker, I. Tessmer, Strandspecific recognition of DNA damages by XPD provides insights into nucleotide excision repair substrate versatility, J. Biol. Chem. 289 (2014) 3613–3624. [134] S.E. Moyer, P.W. Lewis, M.R. Botchan, Isolation of the Cdc45/Mcm2-7/GINS (CMG) complex, a candidate for the eukaryotic DNA replication fork helicase, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 10236–10241. [135] T. Aparicio, E. Guillou, J. Coloma, G. Montoya, J. Mendez, The human GINS complex associates with Cdc45 and MCM and is essential for DNA replication, Nucleic Acids Res. 37 (2009) 2087–2095. [136] S.P. Bell, J.M. Kaguni, Helicase loading at chromosomal origins of replication, Cold Spring Harbor Perspect. Biol. 5 (2013) a010124. [137] S.D. Bell, M.R. Botchan, The minichromosome maintenance replicative helicase, Cold Spring Harbor Perspect. Biol. 5 (2013) a012807. [138] I. Ilves, T. Petojevic, J.J. Pesavento, M.R. Botchan, Activation of the MCM2-7 helicase by association with Cdc45 and GINS proteins, Mol. Cell 37 (2010) 247–258. [139] T. Petojevic, J.J. Pesavento, A. Costa, J. Liang, Z. Wang, J.M. Berger, M.R. Botchan, Cdc45 (cell division cycle protein 45) guards the gate of the eukaryote replisome helicase stabilizing leading strand engagement, Proc. Natl. Acad. Sci. U.S.A. 112 (2015) E249–E258. [140] A. Costa, I. Ilves, N. Tamberg, T. Petojevic, E. Nogales, M.R. Botchan, J.M. Berger, The structural basis for MCM2-7 helicase activation by GINS and Cdc45, Nat. Struct. Mol. Biol. 18 (2011) 471–477. [141] Y.V. Fu, H. Yardimci, D.T. Long, T.V. Ho, A. Guainazzi, V.P. Bermudez, J. Hurwitz, A. van Oijen, O.D. Scharer, J.C. Walter, Selective bypass of a lagging

[142]

[143]

[144]

[145]

[146]

[147] [148]

[149]

[150] [151]

[152]

[153]

[154]

[155]

[156]

13

strand roadblock by the eukaryotic replicative DNA helicase, Cell 146 (2011) 931–941. R.J. Bauer, B.W. Graham, M.A. Trakselis, Novel interaction of the bacterial-Like DnaG primase with the MCM helicase in archaea, J. Mol. Biol. 425 (2013) 1259–1273. N. Marinsek, E.R. Barry, K.S. Makarova, I. Dionne, E.V. Koonin, S.D. Bell, GINS, a central nexus in the archaeal DNA replication fork, EMBO Rep. 7 (2006) 539– 545. B. Zhou, D.R. Arnett, X. Yu, A. Brewster, G.A. Sowd, C.L. Xie, S. Vila, D. Gai, E. Fanning, X.S. Chen, Structural basis for the interaction of a hexameric replicative helicase with the regulatory subunit of human DNA polymerase alpha-primase, J. Biol. Chem. 287 (2012) 26854–26866. J. Sun, Y. Shi, R.E. Georgescu, Z. Yuan, B.T. Chait, H. Li, M.E. O’Donnell, The architecture of a eukaryotic replisome, Nat. Struct. Mol. Biol. 22 (2015) 976– 982. R.E. Georgescu, G.D. Schauer, N.Y. Yao, L.D. Langston, O. Yurieva, D. Zhang, J. Finkelstein, M.E. O’Donnell, Reconstitution of a eukaryotic replisome reveals suppression mechanisms that define leading/lagging strand operation, Elife 4 (2015) e04988. I. Kurth, M. O’Donnell, New insights into replisome fluidity during chromosome replication, Trends Biochem. Sci. 38 (2013) 195–203. L.D. Langston, D. Zhang, O. Yurieva, R.E. Georgescu, J. Finkelstein, N.Y. Yao, C. Indiani, M.E. O’Donnell, CMG helicase and DNA polymerase epsilon form a functional 15-subunit holoenzyme for eukaryotic leading-strand DNA replication, Proc. Natl. Acad. Sci. U.S.A. 111 (2014) 15390–15395. D. Nandakumar, M. Pandey, S.S. Patel, Cooperative base pair melting by helicase and polymerase positioned one nucleotide from each other, Elife (2015) 4. B. Sun, M. Pandey, J.T. Inman, Y. Yang, M. Kashlev, S.S. Patel, M.D. Wang, T7 replisome directly overcomes DNA damage, Nat. Commun. 6 (2015) 10260. M. Manosas, M.M. Spiering, F. Ding, V. Croquette, S.J. Benkovic, Collaborative coupling between polymerase and helicase for leading-strand synthesis, Nucleic Acids Res. 40 (2012) 6187–6198. S. Kim, H.G. Dallmann, C.S. McHenry, K.J. Marians, Coupling of a replicative polymerase and helicase: a tau-DnaB interaction mediates rapid replication fork movement, Cell 84 (1996) 643–650. K. Higuchi, T. Katayama, S. Iwai, M. Hidaka, T. Horiuchi, H. Maki, Fate of DNA replication fork encountering a single DNA lesion during oriC plasmid DNA replication in vitro, Genes Cells 8 (2003) 437–449. N.A. Tanner, S.M. Hamdan, S. Jergic, K.V. Loscha, P.M. Schaeffer, N.E. Dixon, A. M. van Oijen, Single-molecule studies of fork dynamics in Escherichia coli DNA replication, Nat. Struct. Mol. Biol. 15 (2008) 998. C. Indiani, L.D. Langston, O. Yurieva, M.F. Goodman, M. O’Donnell, Translesion DNA polymerases remodel the replisome and alter the speed of the replicative helicase, Proc. Natl. Acad. Sci. U.S.A. 106 (2009) 6031–6038. J. Atkinson, M.K. Gupta, P. McGlynn, Interaction of Rep and DnaB on DNA, Nucleic Acids Res. 39 (2011) 1351–1359.

Please cite this article in press as: S.M. Carney, M.A. Trakselis, Methods (2016), http://dx.doi.org/10.1016/j.ymeth.2016.04.008