Monitoring helicase-catalyzed unwinding of multiple duplexes simultaneously

Abstract

Helicases catalyze the unwinding of duplex nucleic acids to aid a variety of cellular processes. Although helicases unwind duplex DNA in the same direction that they translocate on single-stranded DNA, forked duplexes provide opportunities to monitor unwinding by helicase monomers bound to each arm of the fork. The activity of the helicase bound to the displaced strand can be discerned alongside the helicase bound to the translocase strand using a forked substrate with accessible duplexes on both strands labelled with different fluorophores. In order to quantify the effect of protein-protein interactions on the activity of multiple monomers of the Bacteroides fragilis Pif1 helicase bound to separate strands of a forked DNA junction, an ensemble gel-based assay for monitoring simultaneous duplex unwinding was developed (Su et al., 2019). Here, the use of that assay is described for measuring the total product formation and rate constants of product formation of multiple duplexes on a single nucleic acid substrate. Use of this assay may aid characterization of protein-protein interactions between multiple helicase monomers at forked nucleic acid junctions and can assist with the characterization of helicase action on the displaced strand of forked duplexes.

1. INTRODUCTION

Helicases catalyze the unwinding of duplex nucleic acids to form single-stranded products for a diverse array of cellular processes (Brosh & Matson, 2020). Duplex DNA unwinding catalyzed by ATP hydrolysis proceeds in the direction of single-stranded DNA (ssDNA) translocation, either 5′-to-3′ or 3′-to-5′, giving helicases a specified direction of unwinding (Dillingham et al., 1999; Morris & Raney, 1999; K. D. Raney & Benkovic, 1995; Saikrishnan et al., 2009). Ensemble unwinding assays are frequently performed using gel electrophoresis to resolve nucleic acid products of helicase-catalyzed unwinding of a single duplex with a radio- or fluorophore-labelled strand (S. W. Matson et al., 1983; Steven W. Matson & Kaiser-Rogers, 1990; Venkatesan et al., 1982) . Ensemble duplex DNA unwinding can also be monitored over time using real-time fluorescence measurement (e.g. stopped-flow fluorescence spectroscopy) with fluorophore-labelled DNA duplexes (Belon & Frick, 2008; Bjomson et al., 1994; Donmez & Patel, 2008; Jeong et al., 2004; Özeş et al., 2014; Kevin D Raney et al., 1994). Additionally, single-molecule Förster resonance energy transfer (smFRET) and magnetic or optical tweezers can be used to monitor unwinding of individual DNA molecules (Amit et al., 2004; Bianco et al., 2001; Dawid et al., 2004; Myong et al., 2007; Rasnik et al., 2008; Yodh et al., 2009). These assays have uncovered novel unwinding events such as strand switching, where a helicase switches strands while unwinding a duplex and consequently reanneals the duplex at a similar rate by translocating down the opposite strand (Byrd et al., 2012; Comstock et al., 2015; Dessinges et al., 2004), and reeling (also called patrolling, shuttling, looping, and repetitive activity), where a helicase remains fixed in place and can pull in and unwind DNA without translocating relative to a fixed point on the substrate (Carney et al., 2021; Lu et al., 2019; Park et al., 2010; R. Zhou et al., 2014).

Robust experimental design in ensemble and single-molecule duplex unwinding assays has enabled detailed mechanistic descriptions of nucleic acid unwinding by helicases such as the bacteriophage T4 superfamily 1B helicase Dda (Aarattuthodiyil et al., 2014; Byrd et al., 2012; Byrd & Raney, 2005; Eoff & Raney, 2006; He et al., 2012; Nanduri et al., 2002), E. coli superfamily 1A helicase UvrD (Ali et al., 1999; Ali & Lohman, 1997; Carney et al., 2021; Comstock et al., 2015; Maluf et al., 2003) and the Hepatitis C Virus superfamily 2 helicase NS3 (Jennings et al., 2009; Levin et al., 2004; Myong et al., 2007; Rajagopal & Patel, 2008; Sikora et al., 2008; T. Zhou et al., 2018). DNA footprinting, protein-DNA crosslinking, and X-ray crystallography have established that helicases such as Dda (Aarattuthodiyil et al., 2014), E. coli PriA (Windgassen et al., 2018; Windgassen & Keck, 2016), and Bacteroides fragilis Pif1 (BaPif1) can bind to the displaced strand of a forked duplex opposite an active helicase on the translocase strand. However, use of forked duplex unwinding substrates with only a single labelled duplex prevented the investigation of simultaneous unwinding of duplexes by helicases monomers translocating on opposite strands of the same forked duplex. The need to assess simultaneous unwinding of multiple duplexes on a forked substrate was established by the solution of a crystal structure with two BaPif1 monomers bound to a forked duplex (Su et al., 2019). Unexpectedly, the structure illustrated that the BaPif1 bound to the 3′-ssDNA arm of the displaced strand was situated closer to the forked junction than the BaPif1 bound to the 5′-ssDNA arm of the translocase strand, so the activity of both helicases on the duplex was assessed to confirm the relevance of the structure for forked duplex unwinding by BaPif1 in vitro (Su et al., 2019). To accomplish this, a traditional duplex unwinding assay was modified to include multiple fluorophore-labelled duplexes on a forked duplex substrate (Su et al., 2019). This enabled simultaneous monitoring of unwinding of the duplex on each strand of the forked duplex by each BaPif1 monomer and demonstrated the existence of forked duplexes with active BaPif1 bound to each strand during in vitro unwinding experiments.

The multiple duplex unwinding assay can further be used to assess features of unwinding mechanisms of helicases on forked duplexes, as mechanisms for the increased unwinding of forked duplexes by helicases such as Saccharomyces Pif1 and human FANCJ remain unclear (Gupta et al., 2005; Lahaye et al., 1993; Ramanagoudr-Bhojappa et al., 2013). For example, an increased rate of unwinding of a forked duplex could be explained by either interactions between a helicase bound to the translocase strand with structural features of a forked duplex such as the 3′-ssDNA arm of the displaced strand or the ssDNA-dsDNA fork junction, as demonstrated for Dda (Aarattuthodiyil et al., 2014), or could be explained by protein-protein interactions between active or inactive helicases bound on both the translocase and displaced strand. To distinguish between these mechanisms, the multiple duplex unwinding assay could be used alongside DNA footprinting to assess whether simultaneous binding and/or unwinding of a displaced strand duplex occurs alongside unwinding of the translocase strand and whether protein-protein interactions between the helicases on each strand influence unwinding, as demonstrated for BaPif1 (Su et al., 2019).

Here, we describe and generalize the ensemble multiple duplex unwinding assay originally developed for use with BaPif1 (Su et al., 2019) for use with other helicases, and we demonstrate its utility for determining both total product formation and rate constants of product formation for multiple duplexes on a single substrate.

2. KEY RESOURCE TABLES

Table 1:

Key oligonucleotide substrates and sequences.

Oligonucleotide	Source	Sequence (5′-3′)
T50	Integrated DNA Technologies	TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TT
Parent16	Integrated DNA Technologies	CCA GGC GAC ATC AGC G
Lead16	Integrated DNA Technologies	GCA GTG ACC AGA CAG G
Parent16_10T_Lead16	Integrated DNA Technologies	CCA GGC GAC ATC AGC GTT TTT TTT TTG CAG TGA CCA GAC AGG
5′F-10TParent16comp	Integrated DNA Technologies	6-FAM-TTT TTT TTT TCG CTG ATG TCG CCT GG
5′Cy5-Lead16comp	Integrated DNA Technologies	Cy5-CCT GTC TGG TCA CTG C

3. SAFETY CONSIDERATIONS AND STANDARDS

Care should be taken when performing this protocol. Gloves, a lab coat, and eye protection should be worn as the transport and mixing of large volumes of liquid is required. A chemical fume hood is also required for the handling of volatile substances.

The relevant safety data sheet (SDS) for each reagent should be reviewed before performing this protocol. Dichlorodimethylsilane is a respiratory hazard, causes eye damage, and is flammable, so it should only be used in a fume hood. Acrylamide is a neurotoxin, carcinogen, and may cause reproductive toxicity, so precautions should be taken to avoid contact with acrylamide. Wash hands well after using either of these reagents.

Relevant synthetic DNA safety standards and regulations should be followed at all times when performing this protocol. Synthetic oligonucleotides as used in this protocol are exempt from the NIH Guidelines for Research Involving Recombinant or Synthetic Nucleic Acid Molecules (see Sections III-F-1, III-F-2, and III-F-8).

4. MATERIALS AND EQUIPMENT

4.1. Equipment and Materials

• Equipment

  • Heat block set at 95°C
  • Heat block set at 25°C (or the appropriate temperature for your enzyme)
  • Fluorescent gel imager such as Amersham™ Typhoon™ RGB Imager
  • High voltage power supply (capable of operating at ~1000 V)
  • Sequencing gel system such as Life Technologies/BRL Sequencing System, Model S2 gel system including glass plates, 0.4 mm spacers, 32-well comb, casting clamp, and gel stand
  • Fume hood
  • Magnetic stir plate
  • Benchtop centrifuge
  • Vortex
• Materials

  • 6 large binder clips
  • Plastic wedge
  • Low autofluorescence plastic wrap such as Anchor wrap All-Purpose Foodservice Film
  • KimWipes
  • 1.5 ml tubes; clear and amber
  • Absorbent bench paper
  • Software for analysis of band intensities such as ImageQuant TL or ImageJ

4.2. Reagents

• Fluorophore-labelled and unlabeled oligonucleotide substrates (see Table 1)

• 10 mM HEPES, pH 7.5, 1 mM EDTA solution

• ddH₂O

• 10% (v/v) Dichlorodimethylsilane in hexanes – Dichlorodimethylsilane is corrosive. Store at 4°C with a tightly closed cap wrapped with parafilm.

• 40% (w/v) 19:1 acrylamide/bis-acrylamide – Store at 4°C.

• 10% (w/v) Ammonium persulfate (APS) in ddH₂O – Prepare fresh immediately before use.

• Tetramethylethylenediamine (TEMED)

• 10X Tris-borate-EDTA (TBE), pH 8.0

• 2X Quenching solution – 200 mM EDTA, 0.2% sodium dodecyl sulfate (SDS), 5% glycerol, 0.1% Orange G in ddH₂O

• 5X helicase assay buffer – We used 50 mM Tris, pH 7.5, 250 mM NaCl, 0.5 mM EDTA, 10 mM DTT, 0.5 mg/ml bovine serum albumin (BSA), 25% glycerol for Pif1 and 125 mM HEPES, pH 7.5, 50 mM potassium acetate (KOAc), 0.5 mM EDTA, 10 mM BME, and 0.5 mg/ml bovine serum albumin (BSA) for Dda.

• 100 mM MgCl₂ or magnesium acetate (MgOAC₂)

• 100 mM ATP – Prepare in ddH₂O and bring pH to 7 by adding 1M NaOH dropwise to reduce spontaneous hydrolysis. Store at −20°C.

• Purified helicase – We used Dda, BaPif1, and human PIF1. Purifications are described elsewhere (Byrd et al., 2021; George et al., 2009; X. Zhou et al., 2016).

Alternatives:

• Depending on the lasers and filters available on the fluorescent gel imager, it may be necessary to use different fluorophores whose excitation and emission spectra overlap the available lasers and filters. Any combination of fluorophores with well separated spectra can be used.

• Any vertical electrophoresis system can be utilized instead of a sequencing gel system. However, the gel will have to be carefully monitored to ensure separation of substrate, intermediates, and products without running the smallest products off the gel.

• The preferred reaction buffer will vary based on the specific helicase being studied. 5X helicase assay buffer should be modified to match the appropriate reaction conditions for the helicase of interest.

5. PREPARATION OF DUAL DUPLEX DNA SUBSTRATES

Substrates should be designed to match the characteristics of the helicase of interest. For example, as illustrated in Figure 1A, the substrates used for Pif1 and Dda contained 10 thymidines 5′ to each duplex to allow for binding and unwinding by these 5′-to-3′ helicases. Additionally a short ssDNA tail can be added to the 3′-end of the leading duplex displaced strand to create a forked leading duplex for enzymes that prefer to unwind a forked duplex such as Saccharomyces cerevisiae Pif1 (Su et al., 2019).

Fig. 1.

Expected separation of reaction products following PAGE. (A) Schematic of the FAM- (blue star) and Cy5-labelled (red star) dual duplex unwinding substrate (S), the unwinding intermediates containing the FAM-labelled parental duplex (I_PD) and the Cy5-labelled leading strand duplex (I_LD), and the trapped unwinding products of FAM-labelled parental duplex (P_PD) and Cy5-labelled leading duplex (P_LD). (B) The fluorophore-labelled duplexes were separated on a 20% acrylamide gel by electrophoresis. The FAM-labelled duplexes appear in blue, Cy5-labelled duplexes in red, and the dual-labelled substrate in magenta.

Each duplex is labeled with a fluorophore (FAM and Cy5, in this example) so that unwinding of each duplex can be monitored simultaneously. Fluorophore pairs other than those utilized here can be utilized, provided that imaging equipment is available, and that spectral overlap and background fluorescence are not encountered.

Before an unwinding reaction can be carried out with dual duplex DNA substrates, the appropriate oligonucleotides must be annealed, and successful annealing must be confirmed with polyacrylamide gel electrophoresis (PAGE). The expected outcome of this step is the verification that the dual-duplex substrate, the intermediates, and the reaction products can be properly separated and identified with PAGE following the completion of an unwinding reaction.

5.1. Procedure

5.1.1. Annealing oligonucleotides to form duplexes

Timing: 2 hours-overnight

1. Lyophilized oligonucleotides were resuspended in 10 mM HEPES/1 mM EDTA. Absorbance was measured at 260 nm, and the concentration of each oligonucleotide was calculated using the extinction coefficient. Oligonucleotides were diluted to 50 μM in 10 mM HEPES, 1 mM EDTA and stored at −20°C until further use (see note 5.1).

2. To prepare substrate, intermediates, and products, dilute appropriate oligonucleotides in 10 mM HEPES, 1 mM EDTA in an amber tube.

   1. Dual duplex substrate (S)

      • Mix equimolar amounts of Parent16_10T_Lead16, 5′ F-10TParent16comp, and 5′ Cy5-Lead16comp. See note 5.2 for considerations on substrate design. See critical note 5.1 for important considerations when annealing the dual duplex substrate.)
   2. Intermediate-Parental Duplex (I_PD)

      • Mix 1 equivalent of 5′F-10TParent16comp with a 1.2 fold excess of Parent16_10T_Lead16. Addition of excess unlabeled strand will ensure complete duplex formation of the FAM-labeled duplex.
   3. Intermediate-Leading Duplex (I_LD)

      • Mix 1 equivalent of 5′Cy5-Lead16comp with a 1.2 fold excess of Parent16_10T_Lead16. Addition of excess unlabeled strand will ensure complete duplex formation of the Cy5-labeled duplex.
   4. Product-Parental Duplex (P_PD)

      • Mix 1 equivalent of 5′F-10TParent16comp with a 1.2 fold excess of Parent16. Addition of excess unlabeled strand will ensure complete duplex formation of the FAM-labeled duplex.
   5. Product-Leading Duplex (P_LD)

      • Mix 1 equivalent of 5′Cy5-Lead16comp with a 1.2 fold excess of Lead16. Addition of excess unlabeled strand will ensure complete duplex formation of the Cy5-labeled duplex.

3. Heat oligonucleotides to 95 °C for 10 minutes followed by slow cooling to room temperature to anneal duplexes.

4. Vortex and briefly centrifuge duplexes in a benchtop centrifuge before using.

Pause Point:

Annealed oligonucleotides can be stored at −20 °C protected from light until further use.

5.1.2. Preparing a non-denaturing 20% polyacrylamide gel

Timing: 1.5 - 2 hours

1. Move glass plates into a chemical fume hood and apply a thin layer 10% dichlorodimethylsilane in hexanes with a KimWipe. Wash excess silane off using ddH₂O and a KimWipe. It is critical that this step be performed in a fume hood as dichlorodimethylsilane is a respiratory hazard. See critical note 5.2 for storage and safety considerations when working with dichlorodimethylsilane.

2. Remove the plates from the chemical fume hood and place the large glass plate on a flat surface covered with bench paper. Place the 0.4 mm spacers along each vertical (long) edge of the large glass plate.

3. Invert the short glass plate onto the spacers and the long glass plate, so that the silanized surfaces face each other. Adjust the spacers so that they run parallel to the vertical edges of both plates.

4. Hold the two plates in position and wrap a rubber casting clamp around the corners and sides of the plates to create a sealed chamber with the closed drainage holes in the casting clamp on the top (closest to the shorter plate).

5. Gently adjust the spacers to ensure that the foam seal of each spacer presses down onto the top edge of the shorter plate. This seals the acrylamide chamber against leakage of electrophoresis buffer while running the gel.

6. For a 20% gel, mix 37.5 ml 20% 19:1 acrylamide:bis-acrylamide, 7.5 ml 10X TBE, 30 ml ddH₂O, 1 ml 10% ammonium persulfate, and 75 μl TEMED in a plastic beaker with stirring. Add ammonium persulfate and TEMED immediately before pouring gel as this will initiate polymerization. Acrylamide is toxic; see critical note 5.3 for safety information.

7. Hold the sealed glass plates at a gentle angle and pour the acrylamide solution onto a corner where the large plate, short plate, and spacer meet. Allow the acrylamide solution to flow between the plates to the bottom of the gel and maintain a pool of acrylamide solution that has not yet flowed between the plates to prevent the introduction of air bubbles into the gel. See critical note 5.3 for safety and disposal considerations when working with acrylamide.

8. Prop the gel at a gentle angle by placing an empty rack for 15 ml tubes or a 200 μl pipette tip box under the top of the gel. Insert a 32 well comb and clamp the glass plates with 6 large binder clips along the top of the gel.

9. Wait until the acrylamide has just polymerized (~20-30 minutes).

10. Remove the binder clips and use a clean razor blade to carefully remove the polymerized acrylamide between the top of the comb and the top of the short plate.

11. Apply ddH₂O to the newly exposed comb surface and use a razor blade to gently lift the comb.

12. Apply ddH₂O under the comb to loosen it from the glass plate.

13. Gently remove the comb from between the plates and ensure that the wells are intact.

14. Remove the casting clamp and place the gel and plates into the gel stand with the longer plate facing out.

15. Secure the gel into the gel stand and ensure that a tight seal is formed in the top buffer reservoir.

16. Add 1 liter of 1X TBE split between the top and bottom buffer reservoirs. Flush any air bubbles from the wells using a 30 ml syringe filled with 1X TBE from the top buffer reservoir.

17. Pre-run the gel at 22 mA for 60 minutes.

18. Flush any air bubbles from the wells using a 30 ml syringe filled with 1X TBE from the top buffer reservoir before loading samples.

5.1.3. Separating samples by electrophoresis

Timing: 3 hours

1. Dilute duplex oligonucleotides to a final concentration of 50 nM with ddH₂O and 1X freshly made quenching solution. Vortex to mix, and spin down oligonucleotides before using. See note 5.3 for considerations about loading dye selection for this procedure.

2. Load 5 μl of each substrate into an individual well in the pre-run gel.

3. Run the gel at 22 mA for 2.5 hours. See critical note 5.4 for safety considerations while running the gel, and see note 5.4 for modifications for duplexes shorter than 14 base pairs.

4. Remove the gel from the gel stand, and gently slide the spacers out from between the glass plates.

5. Use a plastic wedge to gently pry one glass plate off of the other, and ensure that the gel remains attached only to a single plate.

6. Carefully place a sheet of plastic wrap onto the exposed gel surface. Wrinkles and bubbles in the plastic will be visible in the scan so take care not to trap air bubbles or wrinkle the plastic wrap in the area of interest. See note 5.5 for considerations on plastic wrap selection.

7. Wipe the back of the glass plate with 100% ethanol before scanning.

8. Scan the gel, plastic-side down, with the FAM and Cy5 channels of a fluorescent gel imager such as the Amersham™ Typhoon™ RGB Imager (Cytiva) using a 488 nm laser and 525BP20 emission filter for FAM and a 635 nm laser and a 670BP30 filter for Cy5.

9. Open the image file in the software provided with your imager or in ImageJ.

5.2. Expected Outcomes

As illustrated in Figure 1B, the success of oligonucleotide annealing to form the fluorescently-labelled substrate can be assessed after electrophoresis. The dual-duplex substrate for use in unwinding reactions appears as the highest molecular weight band in both FAM and Cy5 detection channels. The expected intermediates and products of a helicase-catalyzed unwinding reaction also separate by molecular weight. The presence of a single band with the appropriate fluorescent label(s) in each lane is expected and confirms that the duplexes were annealed with equivalent quantities of all three strands and were not denatured before or during electrophoresis.

5.3. Optimization and Troubleshooting

Presence of single-fluorophore-labelled species after substrate annealing along with annealed substrate band.

An imbalance of concentrations of fluorophore-labelled oligonucleotides and unlabeled strand can lead to the presence of singly-labelled duplexes that are structurally identical to intermediate (i.e. I_PD or I_LD) bands or the presence of bands at low molecular weight corresponding to unannealed fluorophore-labelled oligonucleotides.

If singly-labelled duplexes labelled with both FAM and Cy5 are present after substrate annealing but no free fluorophore-labelled oligonucleotides are present, too much unlabeled strand was added before annealing.

If singly-labelled duplexes labelled with both FAM and Cy5 are present after substrate annealing and free fluorophore-labelled oligonucleotides are also present, too little unlabeled strand was added before annealing.

If a singly-labelled duplex and unannealed fluorophore-labelled oligonucleotides with the same fluorophore are present, an excess of that oligonucleotide was added before annealing.

Potential Solution

Check the concentration of each oligonucleotide by measuring the absorbance at 260 nm and calculating the concentration using the extinction coefficient. Ensure careful pipetting and proper mixing when creating dilutions and solutions.

Presence of smeared bands starting from the expected size of the appropriate duplex and ending at the expected size of the fluorophore-labelled ssDNA.

Melting of the duplexes while the gel is running will lead to smearing of each band.

Potential Solution

Ensure that the polyacrylamide gel components and run conditions (e.g. gel temperature exceeding T_m of the duplex) do not promote denaturation of the substrate. See note 5.4 for substrate design considerations.

Presence of multiple species of a lower-than-expected molecular weight.

Degradation of any of the oligonucleotides or the presence of partial synthesis products can result in the appearance of a ladder of products on the gel. The presence of extra bands can interfere with proper quantitation of unwinding reactions.

Potential Solution

Gel purify the oligonucleotides, and select the slowest migrating species for each oligonucleotide because the full length synthesis product that has not been degraded will be the largest species. To prevent this problem from occurring in the first place, gel purify oligonucleotides before beginning experimentation and prepare all solutions using reagents that are free from DNase contamination.

5.4. Notes

Note 5.1: Protect FAM- and Cy5-labelled oligonucleotides from light to prevent photobleaching by storing stocks and dilutions in amber tubes.

Note 5.2: The Pif1 and Dda helicases that we used translocate 5′-to-3′ on ssDNA. If the helicase being studied translocates 3′-to-5′ on the ssDNA, the substrate will need to be inverted so that both helicases would translocate 3′-to-5′ towards a duplex.

Note 5.3: Orange G dye is used because, although it is visible in the FAM fluorescence scan at 525 nm, it migrates faster than a 12mer oligonucleotide so it does not interfere with quantitation. Xylene cyanol is also visible in a scan for FAM fluorescence at 525 nm, and bromophenol blue is visible in a scan for Cy5 fluorescence at 670 nm. Both xylene cyanol and bromophenol blue migrate slower than Orange G and have the potential to interfere with imaging of FAM or Cy5 labeled oligonucleotides.

Note 5.4: If either duplex is shorter than 14 base pairs, run the gel at 13 mA for 5 hours to prevent the gel from heating enough that the duplex(es) melt(s) while the gel runs.

Note 5.5: We use Anchor Wrap All-Purpose Foodservice Film because it does not display background fluorescence with FAM or Cy5. Other plastic film options would need to be evaluated for background fluorescence during imaging for optimal quantitation. We have observed high background fluorescence from AEP Industries Inc. Sealwrap brand plastic wrap and do not recommend this brand. The background will be affected by the sensitivity of the imager used and the concentration of fluorescently labeled DNA.

5.5. Critical Notes

Critical Note 5.1: It is essential that equimolar quantities of each of the 3 oligonucleotides are mixed when preparing the substrate. If an excess of any of the oligonucleotides is added, the substrate will contain intermediates or free fluorescently labeled oligonucleotides.

Critical Note 5.2: Dichlorodimethylsilane is corrosive and toxic. Apply silane by briefly inverting bottle over a KimWipe, not by pipetting the dichlorodimethylsilane solution, because it will corrode the metal components of the pipette. Always use dichlorodimethylsilane in a chemical fume hood.

Critical Note 5.3: Acrylamide is a neurotoxin. Disposable gloves, lab coat, and eye protection should be worn. When inserting the comb, the acrylamide solution can spray upwards. Let extra acrylamide solution polymerize in a beaker before disposal as polyacrylamide is non-toxic.

Critical Note 5.4: Ensure that the buffer does not leak from the upper buffer chamber and that the gel does not float up out of the silanized plates. If loss of contact between the gel and the buffer in either chamber occurs, the high voltage across the buffer reservoirs creates an electrical fire hazard.

6. MEASURING HELICASE-CATALYZED UNWINDING OF DUAL-DUPLEX DNA SUBSTRATES

Once the dual-labelled substrate has been prepared, it can be used to measure product formation of both duplexes simultaneously. As illustrated in Figures 3 and 4, helicase-catalyzed unwinding of each duplex creates single-fluorophore-labelled products that can be separated with gel electrophoresis and quantified. This method allows for the measurement of duplex unwinding preference for the helicase of interest and can be used to examine how factors such as protein-protein interactions regulate unwinding of each duplex (Su et al., 2019). The same basic experimental design can be used to determine the quantity of product formed (amplitude) at the completion of a single cycle reaction (Figure 3) or to measure product formation over time (Figure 4). Optional addition of a T₅₀ oligonucleotide to the reaction in 30-fold excess relative to the unwinding substrate will create a competing binding site for helicases that have dissociated from the unwinding substrate, so product formation will assess the result of a single unwinding attempt (i.e. single-turnover conditions) per substrate. Omission of the T₅₀ oligonucleotide from the reaction will allow for rebinding of helicases to the substrate after dissociation, so product formation will assess the result of multiple unwinding attempts (i.e. multi-turnover conditions) per substrate.

Fig. 3.

Simultaneous visualization of multiple duplex unwinding. (A) Schematic of the helicase-catalyzed multiple duplex unwinding reaction. Helicase monomers (green) are bound to the ssDNA regions of the dual duplex unwinding substrate (S). Each helicase monomer will translocate 5′-to-3′ (as indicated by the arrow) and unwind the downstream parental duplex (blue) or the leading duplex (red). After ATP, DNA trapping strands (orange), and protein trap (gray) are added, reaction products (I_PD, I_LD, P_PD, and P_LD) are formed, and the helicase is prevented from rebinding to substrate by the protein trap. (B) Unwinding of the dual duplex substrate was performed in triplicate using Dda, and products were visualized after electrophoresis on a 20% native polyacrylamide gel. Multiple time points were collected to ensure completion of the unwinding reaction. (C) The fraction of product formed (either P_PD or P_LD) by unwinding of the duplex was quantified for each time point of each replicate. The mean fraction product formed was averaged to determine the quantity of product formed for each duplex. Plotted are means with standard deviations; n = 3. (D) Rectangles drawn for quantification of the FAM channel of a single replicate of the reaction in 3C are illustrated. (E) Rectangles drawn for quantification of the Cy5 channel of a single replicate of the reaction in 3C are illustrated.

Fig. 4.

Determining the rate of unwinding for multiple duplexes on the same substrate. (A) A multi-turnover unwinding reaction of a dual duplex substrate by human PIF1 helicase was performed (Su et al., 2019). (B) Product formation was quantified, and data was fit to a linear equation to determine that the rate of unwinding was greater for the parental duplex than the leading duplex (Su et al., 2019). (C) A single-turnover unwinding reaction of a dual duplex substrate by BaPif1 helicase was performed with a rapid chemical quench flow instrument (Su et al., 2019). (D) The fraction product formed for BaPif1-catalyzed unwinding of each duplex quantified (Su et al., 2019). (E) The rate constants of unwinding and dissociation were derived by fitting the data to a 5-step sequential mechanism. The rate constants for unwinding the leading and parental duplexes were the same, but the dissociation rate was higher for the parental duplex, leading to a reduced amplitude of product formation for the parental duplex (Byrd et al., 2018; Su et al., 2019). This figure was adapted from Su et al., 2019 and Byrd, Bell, and Raney, 2018 under CC BY Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/). Portions of two figures were combined with minor changes to the font and title. Additionally, data that was originally plotted in two separate plots is combined in panel B.

6.1. Procedure

6.1.1. Unwinding reaction

Timing: 4-5 hours

1. Prepare a native 20% polyacrylamide gel and pre-run it for 60 minutes at 22 mA (see Preparing a native 20% polyacrylamide gel in section 5.1.2).

2. Prepare and label a 0.6 ml microcentrifuge tube for each condition and replicate of the experiment. These tubes will be referred to as reaction tubes (blue tube in Figure 2). Reactions should be performed in triplicate.

3. For each reaction, label a microcentrifuge tube for each time point, including a zero-second time point and a heated control. Add 10 μl of 2X quenching solution to each tube. These tubes will be referred to as time point tubes (orange tube in Figure 2). See note 6.1 for considerations of the number of time points to collect and for considerations on the duration of each time point depending on the desired experimental outcome.

4. Prepare an adequate amount of 2X ATP/trap solution (1X helicase assay buffer, 8 mM ATP, 10 mM Mg(OAc)₂, 0.8 μM Parent16, 0.8 μM Lead16, and 24 μM T₅₀ in ddH₂O) (purple tube in Figure 2) to have a total volume five times the number of time point tubes plus an additional 10 μl. This will provide sufficient volume to use 5 μl for each time point and accounts for sample loss due to sticking to tubes. See note 6.2 for considerations on inclusion of the T₅₀ protein trapping strand. See note 6.3 for considerations on the Parent16 and Lead16 annealing traps.

5. Prepare an equivalent volume of 2X helicase/substrate solution (1X helicase assay buffer, 40 nM dual-fluorophore-labelled substrate, and 2X helicase in ddH₂O (green tube in Figure 2). The concentration of helicase needed will depend on the particular helicase used. See note 6.4 for details relating to experiments performed in Figures 3 and 4. Prepare the helicase/substrate solution without helicase, chill on ice, and then add helicase 5 minutes before initiating the reaction. This incubation allows the enzyme to bind the substrate before beginning the reaction.

6. For the zero-second time point, mix 5 μl of the 2X ATP/trap solution with the 10 μl of 2X quenching solution already in the zero-second time point tube (orange in Figure 2). Then add 5 μl of the enzyme/substrate solution and mix well.

7. For the heated controls, mix 5 μl of the 2X ATP/trap solution with the 10 μl of 2X quenching solution already in the heat time point tube (orange in Figure 2). Then add 5 μl of the enzyme/substrate solution and heat at 95°C for 10 minutes.

8. To begin an unwinding reaction, transfer 5 μl times the number of time points (excluding time zero and heated sample) plus an additional 5 μl of 2X ATP/trap solution to a reaction tube. To initiate the reaction, add an equivalent volume of 2X enzyme/substrate solution, and simultaneously start a timer counting up. Place the reaction tube at the appropriate temperature for the helicase of interest. See note 6.5 for temperature details relating to experiments performed in Figures 3 and 4. The final reaction volume should be 10 μl per time point plus an additional 10 μl.

9. At the appropriate time, remove 10 μl of the reaction solution, and mix the reaction solution with the 10 μl of 2X quench solution in the appropriate time point tube to quench the unwinding reaction. The reaction solution should be added to the quench solution when the timer reaches the time corresponding to the time point. Pipette the solution up 3 times after adding the reaction to the quench to ensure mixing.

10. Repeat step 9 for additional time points as appropriate.

11. Vortex the quenched reaction solutions in the product tubes and spin them down in a tabletop centrifuge.

12. Separate reaction products by gel electrophoresis as described in section 5.1.3 Separating samples by electrophoresis.

Fig. 2.

Illustration of reaction setup. The reaction is prepared in two halves, one containing 2X helicase and substrate, and the other containing 2X ATP, Mg²⁺, and trapping strands. The reaction is initiated by mixing equivalent quantities of each 2X reaction half. At various times, 10 μl are removed from the reaction tube and mixed with an equal volume of 2X quench solution to stop the reaction.

6.1.2. Quantitation of product formation

1. Open the 2 channel fluorescent overlay image and confirm that the substrate, intermediate, and product migrate as expected based on your control gel from section 5. PREPARATION OF DUAL DUPLEX DNA SUBSTRATES. An example FAM and Cy5 channel fluorescent overlay of a multiple duplex unwinding reaction is in Figure 3B.

2. Open the image from a single fluorescent channel. Analysis is performed on single color images, not the composite image of both fluorophores, so that quantitation of product formation for each duplex is unaffected by the signal from the other duplex. Fluorescence intensity of each duplex unwinding product can be measured using the software provided with your imager or ImageJ.

3. As observed in the gel to confirm substrate formation and migration patterns of the substrates and intermediates, three bands should be present in each fluorescence channel. An example FAM channel image of a single replicate from the gel in Figure 3B is shown in Figure 3D, and an example Cy5 channel image from the same replicate is shown in Figure 3E.

4. Identify the substrate from the zero-second time point. Identify the product from the heated sample. The intermediate should migrate between the substrate and product.

5. Measure the density of the substrate, intermediate, and product in each lane. Many programs have built-in methods to do this automatically; however, manual quantitation is preferred to avoid missing lower intensity bands. To quantify manually, draw a rectangle around the substrate in the first lane and copy and paste for substrate, intermediate, and product in each lane. Copy and paste ensures that all rectangles are the same size, which reduces the effect of background. Example rectangles for quantitation are overlayed in green in Figures 3D and 3E. Three extra rectangles are also pasted in regions with no bands for background correction.

6. Copy the data into a spreadsheet and average the three values for background density. Subtract this average background density from the density of each of the substrate, intermediate, and product band.

7. Calculate the fraction product in each lane by dividing the background corrected density of the product band by the sum of the background corrected densities of all three bands. The intermediate is counted as substrate because, in both cases, formation of the intermediate is due to unwinding of the duplex labeled with the fluorophore that is not scanned. For the FAM-labeled parental duplex, the intermediate I_PD has an unwound Cy5-labeled duplex, but the FAM-labeled duplex is intact. Likewise, for the Cy5-labeled leading duplex, the intermediate I_LD has an unwound FAM-labeled duplex, but the Cy5-labeled duplex is intact, as shown in Figure 3A.

8. The fraction product for each duplex is corrected for any product present at time zero as follows:

   $$\mathrm{F}\mathrm{r}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\mathrm{P}\mathrm{r}\mathrm{o}\mathrm{d}\mathrm{u}\mathrm{c}\mathrm{t}=\frac{\frac{{\mathrm{P}}_{\mathrm{t}}}{{\mathrm{S}}_{\mathrm{t}}+{\mathrm{I}}_{\mathrm{t}}+{\mathrm{P}}_{\mathrm{t}}}-\frac{{\mathrm{P}}_0}{{\mathrm{S}}_0+{\mathrm{I}}_0+{\mathrm{P}}_0}}{1-\frac{{\mathrm{P}}_0}{{\mathrm{S}}_0+{\mathrm{I}}_0+{\mathrm{P}}_0}}$$

   P_t is the background-corrected density of product (P_PD for FAM fluorescence; P_LD for Cy5 fluorescence) at a time point. P₀ is the background-corrected density of product at time zero. S_t is the background-corrected density of substrate at a time point. S₀ is the background-corrected density of substrate at time zero. I_t is the background-corrected density of intermediate with the other duplex unwound (I_PD for FAM fluorescence; I_LD for Cy5 fluorescence) at a time point. I₀ is the background-corrected density of intermediate with the other duplex unwound at time zero.

9. For experiments where the desired output is the quantity of product formed in a single cycle, the fraction product formed for each duplex is then averaged across each time point (except zero) and can be graphed as in Figure 3C.

10. For experiments where a reaction time course has been collected, the shape of the reaction progress curve will determine the appropriate equation to use. For multi-turnover reactions, a linear function can be used to fit reactions that have not reached completion and have a linear appearance, as shown in Figure 4A and 4B. For multi-turnover reactions that reach completion, a single exponential function is often appropriate. For single-turnover reactions, the reaction progress curve may be exponential or may be more complicated like the example in Figure 4C and 4D which exhibits a lag phase followed by a burst phase. This data was fit to a n-step sequential mechanism shown in Figure 4E using KinTek Explorer (Johnson et al., 2009). Alternatively, the data can be fit with an equation describing an n-step mechanism (Ali et al., 1999; Ali & Lohman, 1997). In any case, the goodness of fit will need to be evaluated to determine whether the chosen function is appropriate.

6.2. Expected Outcomes

As illustrated in Figure 3, the fraction product formed from the parental duplex and the leading duplex (either P_PD or P_LD) can be assessed following electrophoresis. The zero-second time point serves as a negative control for substrate unwinding, and the heated sample serves as a positive control for duplex unwinding. As shown in Figure 3C, quantification of the mean fraction product formed for each duplex allows for comparison of the processivity of the helicase for unwinding one duplex relative to the other.

As illustrated in Figure 4, the product formation over time can be measured. For multi-turnover reactions, as in Figure 4A, the rate of product formation for one duplex relative to the other can be determined. The data can fit with a linear function, as shown in Figure 4B, or a single exponential function, depending on the shape of the curve. See section 6.1.2 Quantitation of product formation for more information on data fitting. As shown in Figure 4C, a time course of product formation under single turnover conditions can be measured. This allows information about both the rate and processivity of product formation for each of the duplexes to be compared, as the amplitude of the product formation curve is affected by the processivity of the helicase, and the slope of the burst phase is affected by the unwinding rate of the helicase. Differences in the rate or amplitude of the product formation curves for the leading duplex and the parental duplex can be used to infer information about the helicase mechanism (Su et al., 2019).

6.3. Statistical Approaches

Fraction product measurements of duplex unwinding are assumed to be normally distributed, so a student’s t-test can be used to assess statistical significance between fraction product formation measurements of each duplex of interest. If assumptions of normality are violated, a non-parametric Mann-Whitney U-test can be conducted instead.

6.4. Advantages

Measuring unwinding of two duplexes on the same substrate simultaneously provides an opportunity to measure the activity of enzymes bound to both strands of a forked DNA substrate. Since the activity of a helicase bound to the displaced strand is invisible in a traditional DNA unwinding assay, use of substrates with dual duplexes provides additional information about the unwinding mechanism. The use of fluorescently-labelled oligonucleotides reduces hazards created by use of ³²P-labelled oligonucleotides and allows for the monitoring of unwinding of more than one duplex at a time when each duplex is labelled with a different fluorophore. Additionally, the volume of purified enzyme required is small, and the required equipment is commonly available.

6.5. Limitations

The use of this ensemble assay does not elucidate any reaction intermediates or transient events that could be seen with single-molecule techniques.

6.6. Optimization and Troubleshooting

Intermediates and/or products present in zero-second time point

The zero-second time point serves as a negative control for unwinding, as ATP hydrolysis is inhibited before addition of ATP to the reaction. A high abundance of intermediate and product will prevent accurate quantitation.

Potential Solution

Remake the substrate according to section 5.1.1, ensuring that the concentrations of each component are equal. See also section 5.3, “Presence of single-fluorophore-labelled species after substrate annealing along with annealed substrate band”.

After initial rapid product formation, product formation slowly increases in a single-turnover reaction

If the product formation slowly increases after initially forming at a faster rate, it is likely that the concentration of the protein trap is not sufficiently high to ensure single-turnover conditions.

Potential Solution

Increase the concentration of T₅₀ protein trap to prevent rebinding of the helicase to the substrate to ensure single-turnover reaction conditions.

6.7. Notes

Note 6.1: The number of time points required depends on the desired output. For determining product amplitude, three time points should be collected in addition to time zero to ensure the reaction has reached completion. If only one time point is collected, it is difficult to determine whether changes in the quantity of product formed are due to changes in rate or processivity of product formation. If the quantity of product formed increases over time, the experiment can be repeated with longer time points where the fraction product formed is the same for each time point to determine the total quantity of product formed. Alternatively, the experiment can be performed across a broad time course to determine the rate of product formation. The number of time points required for a full time course depends on the helicase and substrate used. If product formation is linear or single exponential, 3 or 6 time points, respectively, are sufficient to define the curve. For more complex reaction progress curves with lag phases, more time points are required.

Note 6.2: T₅₀ serves as a protein trap to prevent helicase that dissociates from the substrate from rebinding a substrate molecule and re-initiating unwinding (i.e., single-turnover conditions). The protein trap is added in at least 30-fold excess of binding sites relative to the helicase concentration. The protein trap can be omitted for multi-turnover reactions where the helicase dissociates and rebinds substrate throughout the reaction.

Note 6.3: The Parent16 and Lead16 are complementary to the FAM- and Cy5-labelled oligonucleotides. They are used to trap the unwound fluorescently labelled oligonucleotides to prevent reannealing. The annealing traps are added at a 20-30-fold excess, relative to the substrate.

Note 6.4: For the experiment in Figure 3, 400 nM Dda was used in the 2X solution, 2 μM hPIF1 was used in the 2X solution for the experiments in Figure 4A and 4B, and 4 μM BaPif1 was used in the 2X solution for the experiments in Figure 4C and 4D.

Note 6.5: For Dda, hPIF1, and BaPif1, reactions were performed at 25°C.

7. ALTERNATIVE METHODS/PROCEDURES

In addition to this procedure, there are other methods which allow monitoring of the unwinding of multiple duplexes simultaneously. This assay could be adapted for real-time measurement of unwinding of multiple duplexes on a single substrate by monitoring the increase in fluorescence upon unwinding of duplexes, each with a quenched fluorophore. Similar experiments using a plate reader or stopped flow spectrophotometer have been performed to monitor unwinding of a single duplex (Donmez & Patel, 2008; Houston & Kodadek, 1994; Marecki et al., 2019; Özeş et al., 2014).

Single molecule FRET (smFRET) microscopy with alternating laser excitation (ALEX)(Hohlbein et al., 2014) allows simultaneous measurement of unwinding of multiple duplexes in a single molecule (Su et al., 2019). The main advantage of smFRET with ALEX is that by measuring unwinding of single molecules, intermediates in the unwinding process can be observed. In the case of BaPif1 unwinding a dual duplex substrate similar to those described here was monitored by smFRET with ALEX, and a mid-FRET intermediate was observed that was missed in the ensemble experiments (Su et al., 2019). This intermediate was observed for unwinding of both the leading and parental duplex (Su et al., 2019). DNA footprinting and DNA protein-crosslinking can also be used to determine if helicases interact with both the translocase and displaced strand ssDNA arms of a forked duplex (Aarattuthodiyil et al., 2014; Windgassen et al., 2018; Windgassen & Keck, 2016), although these methods do not provide information about the activity of the helicases bound to the two strands.