- Email: [email protected]
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
CHAPTER EIGHTEEN
Single-Molecule Studies of ssDNA-Binding Proteins Exchange Olivia Yang*, Taekjip Ha*,†,‡,1 *Johns Hopkins School of Medicine, Baltimore, MD, United States † Howard Hughes Medical Institute, Baltimore, MD, United States ‡ Johns Hopkins University, Baltimore, MD, United States 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Single-Molecule FRET Experimental Strategies 2.1 Preparation and Assembly of Polyethylene Glycol Slides 2.2 DNA Substrate Preparation 2.3 Single-Molecule FRET With TIR Microscopy 2.4 SSB Replacement Assay 2.5 SSB Transfer Assay 3. Notes 4. Summary Acknowledgments References
464 465 465 466 468 469 472 474 476 477 477
Abstract Single-stranded DNA-binding protein (SSB) is important not only for the protection of single-stranded DNA (ssDNA) but also for the recruitment of other proteins for DNA replication, recombination, and repair. The interaction of SSB with ssDNA is highly dynamic as it exists as an intermediate during cellular processes that unwind dsDNA. It has been proposed that SSB redistributes itself among multiple ssDNA segments, but transient intermediates are difficult to observe in bulk experiments. We can use single-molecule FRET microscopy to observe intermediates of the transfer of a single Escherichia coli SSB from one ssDNA strand to another or exchange of one SSB for another on a single ssDNA in real time. This single-molecule approach can be further applicable to understand relative binding affinities and competitive dynamics for other SSBs and variants across various systems.
Methods in Enzymology, Volume 600 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2017.11.017
#
2018 Elsevier Inc. All rights reserved.
463
464
Olivia Yang and Taekjip Ha
1. INTRODUCTION In order to access and use genomic DNA for processes such as replication, recombination, and repair, single-stranded DNA (ssDNA) is generated as an intermediate. In this state, DNA is more susceptible to breakage, chemical mutagens, or nuclease digestion. ssDNA-binding protein (SSB) not only exists in most biological systems to prevent these damages but also plays an important role in recruitment of relevant proteins to regions of ssDNA to carry out their own functions. In order to efficiently distribute SSB among exposed ssDNA, it is likely that SSB is recycled and redistributed. SSB transfer between adjacent and distant DNA segments can allow rapid recycling and redistribution. Exchange of a bound SSB with another may make it possible to replace a set of proteins bound to the incumbent SSB with another set bound to the incoming SSB. Therefore, a detailed investigation of SSB transfer and exchange kinetics is expected to reveal functionally important properties. While a mechanism of interstrand exchange has been proposed, it is difficult to observe transient intermediates in most ensemble experiments (Kozlov & Lohman, 2002; Lee et al., 2014). To focus on the transfer of a single SSB to another DNA, or the exchange of one SSB for another at the single molecule level, we can use surface-tethered DNA substrates. We have previously described detailed single molecule methods to observe transitions different binding modes of SSB and SSB diffusion along short ssDNA (Roy, Kozlov, Lohman, & Ha, 2007; Zhou & Ha, 2012). For Escherichia coli SSB, there are two major binding modes, (SSB)65 and (SSB)35, which have well-defined ssDNA wrapping lengths. The numbers 65 and 35 refer to the number of nucleotides contacted by the SSB homotetramer in each mode (Roy et al., 2007). Here, we describe an assay of using a mutant SSB with a distinct binding behavior as compared to wild-type SSB (wtSSB) to observe exchange of a DNA–bound SSB for wtSSB in solution. We can also observe an exchange of a labeled SSB for an unlabeled SSB. Exchange kinetics has been measured in ensemble, and it was proposed that there must exist an intermediate of multiple SSB bound to a single ssDNA region (Kunzelmann, Morris, Chavda, Eccleston, & Webb, 2010). Using single-molecule fluorescence resonance energy transfer (smFRET), we can directly observe the real-time formation of a reaction intermediate with more than one SSB bound to the same DNA. We can also use the FRET difference between a DNA fully wrapped around SSB
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
465
and a free DNA to observe the transfer of SSB to a separate DNA strand. We can use these observations to dissect the exchange kinetics and mechanisms. In this chapter, we describe protocols for studying replacement (bound SSB being exchanged for SSB in solution) and transfer (bound SSB transferred to free ssDNA in solution) mechanisms using E. coli SSB as a model.
2. SINGLE-MOLECULE FRET EXPERIMENTAL STRATEGIES 2.1 Preparation and Assembly of Polyethylene Glycol Slides In order to perform single-molecule microscopy experiments using surfacetethered molecules, a clean and passivated surface is required. This minimizes the loss of sample protein and DNA through nonspecific adsorption, as well as potential artifacts caused by interaction with the surface. Generally, polyethylene glycol (PEG)-coated quartz slides and glass coverslips are used for single-molecule total internal reflection fluorescence (TIRF) experiments. PEG slides and coverslips for TIRF microscopy purposes were obtained from Microscope Slides Core facility in Johns Hopkins University School of Medicine. Each slide has multiple channels and is suitable for multiple experiments. 2.1.1 Materials • PEGylated quartz slide and coverslip • Double sided tape • 5-min epoxy • Tubing: WEICO ETT-28 (inner diameter ¼ 0.01500 , outer wall ¼ 0.01600 ) • Needle: BD (26 gauge, 3/800 ) • 1-mL syringe • 20–200-μL pipette tips • Vacuum sealable food saver bags 2.1.2 Procedure 1. Slides are assembled with PEG surfaces of the slide and coverslip facing each other to create passivated channels. PEGylated surfaces can be distinguished from untreated surfaces by dropping water and observing the hydrophobicity of the surface; a treated surface will be more hydrophobic.
466
Olivia Yang and Taekjip Ha
2. After testing, surfaces can be dried with house air or N2(g); PEGylated surfaces should not be physically touched or the passivation will be disrupted. 3. Double-sided tape (100 μm thick) is used as a spacer between the slide and coverslip, and to create the channels for multiple experiments (see Fig. 1). Tape is cut into thin strips on a clean surface with a clean razor blade. Do not move tape once it has been placed. 4. Place the coverslip on top of the slide and tape, PEG side down, and press with pipette tip to secure the adhesion and prevent leaks between channels. 5. Ends of the channel are sealed with 5-min epoxy as marked in yellow (see Fig. 1). Excessive epoxy used in this step may block the inlet holes drilled in the slide, so use sparingly. 6. For real-time flow experiments, additional setup is required (see Fig. 3; Section 3) 7. Slides are typically stored at 20°C in 50-mL tubes, into vacuum sealed bags to avoid moisture. Slides with unused channels can be similarly stored for later use.
2.2 DNA Substrate Preparation A DNA substrate similar to previously used for SSB-binding mode and dynamics studies is tethered to the surface for single-molecule experiments
Fig. 1 PEG slide assembly. Slides are assembled with PEG surfaces facing inward. Double-sided tape is placed as marked in gray, between holes drilled. The long edge of the coverslip and slide is sealed with 5-min epoxy (yellow region). One channel is the space between two tapes.
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
467
(Roy et al., 2007; Zhou et al., 2011). The partial duplex substrate is made of an 18 bp duplex with a polydeoxythymidine (poly-dT) overhang of varying lengths. The 18 bp duplex is used as a spacer to minimize potential surface effects. Poly dT is used for its lack of secondary structure and high binding affinity to SSB, although other sequences may also work (Lohman & Overman, 1985). The length of the ssDNA used is approximately 70 nt, enough to allow for a full wrapping of SSB ((SSB)65 mode) or two partially wrapped SSB ((SSB)35 mode). The end of the duplex without the ssDNA overhang is biotinylated to specifically tether the DNA to the surface, while the other end of the duplex is Cy5 labeled. For smFRET experiments, cyanine dyes Cy3 and Cy5 are commonly used for donor and acceptor molecules, respectively, on DNA due to their high photostability under deoxygenated conditions and high brightness. A detailed labeling procedure is previously published (Joo & Ha, 2012). For experiments with labeled SSB, the substrate is labeled with Cy5 only. For unlabeled SSB, the far end of the ssDNA is additionally labeled in order to observe FRET changes upon SSB binding. SSB is labeled with an average of one Alexa 555, a small organic dye with excitation and emission spectra similar to Cy3, per SSB tetramer as previously described (Lee et al., 2014; Roy, Kozlov, Lohman, & Ha, 2009; Zhou et al., 2011). The DNA oligonucleotides (oligos) used are listed below: 1. 50 -/Cy5/ GCC TCG CTG CCG TCG CCA -/biotin/-30 . 2. 50 -TGG CGA CGG CAG CGA GGC (T)m-/Cy3/-30 (m ¼ 72). 3. 50 -TGG CGA CGG CAG CGA GGC (T)m-30 (m ¼ 70). 4. 50 -(T)m-30 (m ¼ 60). The DNA substrate for labeled SSB experiments was annealed by mixing oligos 1 and 3 at 10 μM and 20 μM, respectively, in 10 mM Tris–HCl (pH 8) and 50 mM NaCl, then slow cooling from 95°C to 4°C over 4 h. The cooling can be done in a thermocycler at a rate of 1°C/min, or with a heat block taken off heat and allowed to cool to RT then stored at 20°C. The substrate for unlabeled SSB experiments was prepared similarly, but using oligos 1 and 2. Typical annealing uses an excess (1.2 to 2) of nonbiotinylated strands in order to ensure that surface-bound DNA contain both strands. DNA substrates are stored at 20°C, covered in foil to prevent photobleaching. For single-molecule experiments, a substock of 5–10 nM DNA is prepared. At lower concentrations, DNA can be lost due to nonspecific adsorption to tube surfaces. 1% v/v BSA (20 mg/mL stock) in solution can act to passivate surfaces and prevent some loss.
468
Olivia Yang and Taekjip Ha
2.3 Single-Molecule FRET With TIR Microscopy A detailed explanation of the prism-type TIRF microscope setup used in these protocols can be found as previously published (Joo & Ha, 2012). Video recordings were processed to extract single molecule fluorescence intensities at each frame, and custom written scripts were used to calculate FRET efficiencies. Data acquisition and analysis software can be downloaded from https://cplc.illinois.edu/software/. Alternating laser excitation (ALEX) is used to filter out molecules with donor only or acceptor only (Kapanidis et al., 2005). We require a minimum intensity cutoff during donor (green) or acceptor (red) excitation. If a fluorescence spot has a combined intensity from donor and acceptor channels less than the cutoff, then it is not counted. Stoichiometry parameter S can also be used to identify the donor and acceptor only spots. The stoichiometry S of each molecule is calculated as S¼
IAG + IDG IAG + IDG + IAR
G Where IG A is the acceptor intensity during green excitation, ID is the donor intensity during green excitation, and IR A is the acceptor intensity during red excitation. Molecules with low S values (0–0.2) correspond to acceptor only molecules, while molecules with high S values (0.8–1) correspond to donor only molecules. Molecules with both donor and acceptor have stoichiometry S 0.5. Our typical data acquisition sequence is an initial set of 10 frames of red laser excitation, followed by green laser excitation. FRET efficiency is calculated from green excitation periods. Long movies had an additional 10 frames of red excitation at the end to test if the acceptor has photobleached during data acquisition. A “snapshot” is defined as 20 frames taken at a single imaging area. “Long movies” are defined as enough frames to photobleach 80% of the molecules within the imaging area, usually around 2000–3000 frames. Leakage of donor signal to the acceptor channel and background signals of both channels were corrected similar to as previously described (Roy, Hohng, & Ha, 2008). Apparent FRET efficiency is calculated with the equation
EFRET ¼
IA leakage ID IA leakage ID + ID
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
469
where IA is the acceptor intensity, leakage is the leakage factor from the donor to the acceptor detection channel, and ID is the donor intensity. Leakage factor will vary depending on the microscope setup used. Background for long movies is corrected to make the intensities after donor photobleaching equal to zero. Histograms were made from at least 10 individual snapshots of a single sample, with approximately 200–300 molecules imaged per area for a total of 2000–3000 molecules used per histogram. The FRET efficiency of each molecule was calculated based on the average intensity of donor and acceptor over eight frames, avoiding the frames around when excitation is changing.
2.4 SSB Replacement Assay In this chapter, we describe the use of E. coli SSB’s well-characterized and controlled binding modes (Roy et al., 2007). The method of purification and labeling of SSB can be found as previously published (Lohman, Green, & Beyer, 1986; Roy et al., 2009). Because we want to observe primarily exchange of a single SSB ((SSB)65), we use NaCl concentrations greater than or equal to 100 mM; any lower NaCl concentration and there would be a significant population of two SSB molecules bound to 70-mer ssDNA ((SSB)35 mode). Similarly, SSB concentration should be kept relatively low (<10 nM in 100 mM NaCl) to minimize binding in the (SSB)35 mode. For an SSB variant with lower binding affinity, lower concentrations of wtSSB will be required for displacement. With only free DNA, 1 nM SSB is expected to bind almost fully within 1 min in high salt conditions. 2.4.1 Materials • T50 buffer: 10 mM Tris–HCl (pH 8.0), 50 mM NaCl in ddH2O, filtered through 0.22-μm membrane • SSB imaging buffer (SSB IB 100 mM NaCl): 100 mM NaCl, 10 mM Tris–HCl (pH 8.0), 0.1 mg/mL BSA, 0.8% (w/v) dextrose, 165 U/mL glucose oxidase, 2170 U/mL catalase, 2-3 mM Trolox, and indicated amounts of SSB. See Section 3. • Neutravidin: 0.2 mg/mL neutravidin diluted in T50 buffer • Assembled PEG slide (see above) • Prism-type TIRF microscope • Labeled DNA substrate (oligomers 1 + 3, see above) • Unlabeled DNA substrate (oligomers 1 + 2, see above) • SSB stock: to be kept at 20°C, secondary stock can be made in SSB IB and kept on ice during the course of one set of experiments.
470
Olivia Yang and Taekjip Ha
2.4.2 Procedure 2.4.2.1 Experiment Procedure
1. Confirm quality of slide (see Section 3). 2. Flow in 20 μL T50 buffer to each channel expected to be used for one experiment. The approximate channel volume is about 20 μL. For thorough flushing or between different buffer conditions, use at least 30 μL of solution. 3. Flow in 20 μL neutravidin to each channel. Incubate 1–2 min, then flush with T50. 4. Prepare 10–30 pM DNA substrate in T50 (“labeled” for labeled SSB displacement, “unlabeled” for mutant SSB displacement) and add to channel. Incubate for 1–2 min, then flush with SSB IB + “gloxy” to check the spot density. For “labeled” substrate, there should only be Cy5 signal, while “unlabeled”substrate should have both Cy3 and Cy5 signal. There should be 200–300 spots of a single dye within one imaging area. If the number of spots is too little, dilute more DNA and add to channel until desired density is reached. 5. Flush with SSB IB + gloxy. Take snapshots to generate DNA only histogram. 6. Add 40 μL of 1 nM SSB (labeled or mutant) in SSB IB 100 mM NaCl and incubate for at least 5 min (Fig. 2A). 7. Remove unbound SSB by flushing with SSB IB 100 mM NaCl + gloxy (Fig. 2B). Take snapshots to get initial time point. 8. Add wtSSB at desired concentration. To observe real-time exchange, a flow cell assembly must be used (see Fig. 3; Section 3). 9. Take at least 10 snapshots at each given time point (0–30 min).
Fig. 2 Scheme of SSB replacement assay. (A) Labeled SSB is added to ssDNA tethered to the PEG slide. (B) Unbound labeled SSB is flushed from sample chamber. (C) wtSSB (dashed outline) is added to sample chamber. (D) Labeled SSB is displaced from tethered DNA.
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
471
Fig. 3 Real-time flow experiment setup—cross section of single channel. Pipette tip segments are stuck to the quartz slide with epoxy. “Sample well” holds the sample to flow in. Teflon tubing is stuck into pipette tip which has been adhered to hole with epoxy. Tubing is attached to a 1-mL syringe with 26-gauge needle.
2.4.2.2 Data Analysis
For labeled SSB, replacement can be observed as the disappearance of fluorescent spots over time. Because the DNA substrate used is labeled with Cy5, we can use Alexa 555 as a label on SSB to distinguish it from unlabeled SSB and to observe FRET upon binding. We include in our analysis only those spots with both the acceptor (Cy5 on DNA) and donor (Alexa 555 labeled SSB) signals using ALEX. The two color colocalization allows us to reject SSB bound to the surface nonspecifically from analysis, and FRET provides an additionally stringent criterion for specific binding. Fluorescence spots with excessively high or low intensity are also rejected because they typically are not due to the molecules of interest. Generally, spots will also disappear due to photobleaching, so it is important to use the condition with no wtSSB added to determine a photobleaching rate. It is important to keep the laser intensity constant to avoid changing this photobleaching rate too much between different experiment repeats. The number of spots is normalized to the initial time point, since for a given experiment the actual spot density at the beginning of experiment may vary. The displacement rate can be determined from a single exponential decay fit, then subtracting the photobleaching rate determined separately. Plotting the rate of spot disappearance against concentration of wtSSB added shows a linear relation.
472
Olivia Yang and Taekjip Ha
For a mutant SSB with less than full wrapping properties, the exchange is observed through a shift in the FRET efficiency. As shown in Fig. 4, the mutant has a FRET efficiency centered around 0.5 (Fig. 4C, top panel), while wtSSB has a FRET efficiency about 0.7 (Fig. 4C, bottom panel). In order to confirm what FRET efficiencies each species corresponds with, controls should be done to see binding in the same buffer conditions at low [SSB]. The simplest way of determining exchange is imposing a FRET cutoff (0.6) to distinguish the mutant-bound DNA population vs the wtSSBbound population. For variants with more overlapping FRET efficiencies, it would be more reasonable to fit the histogram with a Gaussian function for each population and use the area to calculate the population ratio. Using this ratio, we can plot the binding of wtSSB over time as it replaces the mutant. Similar to the labeled SSB assay, we can use this plot to obtain exchange rates at various concentrations of wtSSB. Movies taken during real-time replacement assays will result in FRET traces as sketched in Fig. 4B. Three species are expected to exist in this experiment: the initial, mutant-bound SSB (0.5 FRET); the transient, 2 SSB-bound state with one mutant and one wild type (0.2 FRET, (SSB)35); and the final, wtSSB-bound species (0.7 FRET) (see Fig. 4A) (Roy et al., 2007). Through measuring dwell times of each FRET state on a single molecule trace, the rates of exchange between these three species can be determined. In this case, since we flood the system with wtSSB in a high salt buffer, we expect the reaction to be pushed toward the wtSSBbound state. The EFRET observed in the single molecule traces (Fig. 4B) should correspond with the centers of peaks as observed in histograms (Fig. 4C). Because the (SSB)35 mode is not favored in these salt conditions, it does not exist long enough to be captured in histograms. However, it can be seen as an intermediate during real-time experiments.
2.5 SSB Transfer Assay For this experiment, we use (dT)60 because the expected binding site size for a single SSB is approximately 65 nt. Varying lengths or sequences of DNA oligomers can be tested for transfer activity with the same assay, though may require different concentration ranges. Sequences with secondary structures may also be inefficient in competing with the poly-dT substrate used. 2.5.1 Materials The same materials as listed in the SSB displacement assay are used. Additional/alternative materials required are listed below.
Fig. 4 Results from mutant to wtSSB displacement (A) Scheme of displacement mechanism. (B) Ideal FRET trace of mutant to wtSSB in a “two-step” transition. (C) FRET histograms of populations observed during experiment.
474
• •
Olivia Yang and Taekjip Ha
dT60 (oligomer 4): stock diluted in T50, stored at 20°C SSB imaging buffer (SSB IB 500 mM NaCl): 500 mM NaCl, 10 mM Tris–HCl (pH 8.0), 0.1 mg/mL BSA, 0.8% (w/v) dextrose, 165 U/mL glucose oxidase, 2170 U/mL catalase, 2-3 mM Trolox, and indicated amounts of SSB.
2.5.2 Procedure 2.5.2.1 Experiment Procedure
The same procedure for SSB replacement is followed for SSB transfer, but rather than SSB added at step 6, dT60 at varying concentrations is used in a SSB IB 500 mM NaCl buffer, and throughout the experiment SSB IB 500 mM NaCl is used in place of SSB IB 100 mM NaCl. 2.5.2.2 Data Analysis
The initial state of the tethered substrate is bound by a single SSB, so shows a EFRET 0.7. The EFRET of unbound ssDNA of a dT70 construct in 500 mM NaCl is 0.15. The exact value will vary slightly based on NaCl or other salt concentrations. Since the difference in EFRET is large, a cutoff can be used to determine the bound and unbound DNA substrate populations.
3. NOTES Imaging buffer is generally prepared as follows: 1. Trolox + dextrose solution: 10 mg Trolox (()-6-Hydroxy-2,5,7,8tetramethylchromane-2-carboxylic acid, Sigma-Aldrich) and 80 mg dextrose (Sigma-Aldrich) is dissolved in 10 mL ddH2O and 10 μL 5 M NaOH. Mixture is rotated for at least 4 h, RT, covered in foil. Solution is filtered through 0.22-μm membrane and stored at 4°C. This solution can be used for 2–3 weeks when stored at 4°C, or longer if stored frozen. 2. SSB IB: NaCl, Tris–HCl (pH 8.0), and BSA are added to Trolox + dextrose solution to the final concentrations listed above to make a flush buffer for removing free protein from channel. NaCl, Tris– HCl, and BSA are added with SSB to Trolox + dextrose solution to final concentrations as needed for steps adding SSB to channel. 3. “Gloxy”: 10 mg glucose oxidase (G2133, Sigma-Aldrich) and 20 μL bovine liver catalase (C30, Sigma-Aldrich) is dissolved in 100 μL T50 buffer. Mixture is spun down for 2 min at 10,000 g. The top 100 μL of the solution is transferred and stored at 4°C, and can be used for
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
475
2–4 weeks. This solution is 100 concentrated, and is typically not added until right before samples are added to the slide. Upon mixing with IB, the solution will continually become more acidic upon reacting with atmospheric oxygen (Joo & Ha, 2012). Slides should always be checked for cleanliness and passivation before use, as follows: 1. Cleanliness: With just T50 in the channel, there should be fewer than 10 observable fluorescent spots per imaging area. 2. Passivation: After flowing in 1 nM labeled DNA or protein, there should be less than 30 observable spots stuck to the surface. After flushing with T50, there should be significantly less spots compared to earlier. Protein is generally stickier than DNA and is expected to have slightly more nonspecific binding. 3. Only after the channel has passed these checks should the user proceed with the experiment. Real-time flow experiments have other considerations: 1. 20–200-μL pipette tips are used in this setup. The sample well is made from the wider portion of the tip, while the outlet side is the sharper portion of the tip. Both are adhered to the slide with epoxy. Fig. 3 shows the cross section of a channel assembled for a real-time flow experiment. 2. It is not necessary to epoxy the tubing to the tip, though it helps make the assembly more secure. Care should be taken to avoid clogging the drilled holes with epoxy. 3. A minimum of 40 μL of sample must be added to the sample well to avoid drawing bubbles through the channel. 4. Sample is drawn through by pulling with a syringe. This can be done manually, or in a more controlled way with a syringe pump. With a syringe pump, the exact flow time and rate can also be controlled. At no point should the sample be pushed back through the channel. 5. All solutions added to this channel will be drawn through by adding the solution to the sample well and pulling through with the syringe. 6. Because a sample is being pulled through during the recording of a movie, care must be taken that the pulling action does not perturb the sample stage or any part of the microscope. Slight motions of the slide or sample will affect the focus and the quality of the movie.
476
Olivia Yang and Taekjip Ha
7. After the initial movie is taken, further long movies or snapshots can be taken of the same channel. 8. Tubing and syringes can be reused, but fresh pipette tips should be used for adhering to the slide. Other general notes: 1. During imaging, general exposure time used is 50–100 ms. Highest time resolution for the camera used in these experiments (Andor DU-897E-CSO-#BV) is 30 ms for 512 512 pixels, though the imaging area can be decreased to increase time resolution. To avoid photobleaching, a longer exposure time should be used to minimize laser intensity. To observe faster dynamics, shorter exposure times should be used, but higher laser intensities will be required to increase signal to noise ratios. 2. Slides can be reused. By soaking in methanol, epoxy is weakened and the coverslip and double sided tape can also be removed and disposed of. Then, slides should be boiled in water and any visible junk can be washed off before proceeding with usual treatment (Roy et al., 2008). 3. All concentrations of SSB in this text refer to concentration of SSB tetramer. 4. DNA substrate concentration refers to the concentration of the biotinylated strand. Nonbiotinylated strand is used in excess in order to maximize the amount of annealed DNA that gets specifically tethered to the slide. Nonbiotinylated strand will get flushed from the channel and should not interfere with experiments. 5. High concentrations of DTT cannot be used in imaging buffers for single-molecule experiments, as blinking will be observed.
4. SUMMARY smFRET and single molecule experiments are especially useful for understanding more detailed dynamics of SSB, especially on a small length scale. These details may be helpful to explain steps in protein recruitment processes during replication, recombination, and repair related to ssDNA processing. Though E. coli SSB was used throughout these experiments, similar single molecule studies regarding transfer and replacement mechanisms can be done with the same method for other relevant SSB variants to further understand the differences between SSBs from different organisms. However, the binding of each new variant needs to be well
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
477
characterized for a given substrate before observing exchange in order to properly attribute a given FRET efficiency to a specific variant. Using single-molecule methods can be useful for observing and confirming intermediates as proposed by other ensemble scale SSB experiments.
ACKNOWLEDGMENTS The authors would like to thank Dr. Jichuan Zhang for his guidance and advice throughout this project, and Prof. I-Ren Lee for his helpful ideas. Research in this chapter was funded by NIH Grant R35 GM 122569.
REFERENCES Joo, C., & Ha, T. (2012). Single-molecule FRET with total internal reflection microscopy. Cold Spring Harbor Protocols, 2012(12), 1223–1237. https://doi.org/10.1101/pdb. top072058. pdb.top072058. Kapanidis, A. N., Laurence, T. A., Nam, K. L., Margeat, E., Kong, X., & Weiss, S. (2005). Alternating-laser excitation of single molecules. Accounts of Chemical Research, 38(7), 523–533. https://doi.org/10.1021/ar0401348. Kozlov, A. G., & Lohman, T. M. (2002). Kinetic mechanism of direct transfer of Escherichia coli SSB tetramers between single-stranded DNA molecules. Biochemistry, 41(39), 11611–11627. https://doi.org/10.1021/BI020361M. Kunzelmann, S., Morris, C., Chavda, A. P., Eccleston, J. F., & Webb, M. R. (2010). Mechanism of interaction between single-stranded DNA binding protein and DNA. Biochemistry, 49, 843–852. https://doi.org/10.1021/bi901743k. Lee, K. S., Marciel, A. B., Kozlov, A. G., Schroeder, C. M., Lohman, T. M., & Ha, T. (2014). Ultrafast redistribution of E. coli SSB along long single-stranded DNA via intersegment transfer. Journal of Molecular Biology, 426(13), 2413–2421. https://doi.org/ 10.1016/j.jmb.2014.04.023. Lohman, T. M., Green, J. M., & Beyer, R. S. (1986). Large-scale overproduction and rapid purification of the Escherichia coli ssb gene product. Expression of the ssb gene under lambda PL control. Biochemistry, 25(1), 21–25. https://doi.org/10.1021/ bi00349a004. Lohman, T. M., & Overman, L. B. (1985). Two binding modes in Escherichia coli single strand binding protein-single stranded DNA complexes: Modulation by NaCl concentration. The Journal of Biological Chemistry, 260(6), 3594–3603. Retrieved from http:// www.jbc.org/content/260/6/3594.full.pdf. Roy, R., Hohng, S., & Ha, T. (2008). A practical guide to single-molecule FRET. Nature Methods, 5(6), 507–516. https://doi.org/10.1038/nmeth.1208. Roy, R., Kozlov, A. G., Lohman, T. M., & Ha, T. (2007). Dynamic structural rearrangements between DNA binding modes of E. coli SSB protein. Journal of Molecular Biology, 369(5), 1244–1257. https://doi.org/10.1016/j.jmb.2007.03.079. Roy, R., Kozlov, A. G., Lohman, T. M., & Ha, T. (2009). SSB protein diffusion on singlestranded DNA stimulates RecA filament formation. Nature, 461, 1092–1097. https:// doi.org/10.1038/nature08442. Zhou, R., & Ha, T. (2012). Single-molecule analysis of SSB dynamics on single-stranded DNA. In Single-stranded DNA binding proteins (pp. 85–100). Totowa, NJ: Humana Press. https://doi.org/10.1007/978-1-62703-032-8_5. Zhou, R., Kozlov, A. G., Roy, R., Zhang, J., Korolev, S., Lohman, T. M., et al. (2011). SSB functions as a sliding platform that migrates on DNA via reptation. Cell, 146(2), 222–232. https://doi.org/10.1016/j.cell.2011.06.036.
Single-Molecule Studies of ssDNA-Binding Proteins Exchange Olivia Yang*, Taekjip Ha*,†,‡,1 *Johns Hopkins School of Medicine, Baltimore, MD, United States † Howard Hughes Medical Institute, Baltimore, MD, United States ‡ Johns Hopkins University, Baltimore, MD, United States 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Single-Molecule FRET Experimental Strategies 2.1 Preparation and Assembly of Polyethylene Glycol Slides 2.2 DNA Substrate Preparation 2.3 Single-Molecule FRET With TIR Microscopy 2.4 SSB Replacement Assay 2.5 SSB Transfer Assay 3. Notes 4. Summary Acknowledgments References
464 465 465 466 468 469 472 474 476 477 477
Abstract Single-stranded DNA-binding protein (SSB) is important not only for the protection of single-stranded DNA (ssDNA) but also for the recruitment of other proteins for DNA replication, recombination, and repair. The interaction of SSB with ssDNA is highly dynamic as it exists as an intermediate during cellular processes that unwind dsDNA. It has been proposed that SSB redistributes itself among multiple ssDNA segments, but transient intermediates are difficult to observe in bulk experiments. We can use single-molecule FRET microscopy to observe intermediates of the transfer of a single Escherichia coli SSB from one ssDNA strand to another or exchange of one SSB for another on a single ssDNA in real time. This single-molecule approach can be further applicable to understand relative binding affinities and competitive dynamics for other SSBs and variants across various systems.
Methods in Enzymology, Volume 600 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2017.11.017
#
2018 Elsevier Inc. All rights reserved.
463
464
Olivia Yang and Taekjip Ha
1. INTRODUCTION In order to access and use genomic DNA for processes such as replication, recombination, and repair, single-stranded DNA (ssDNA) is generated as an intermediate. In this state, DNA is more susceptible to breakage, chemical mutagens, or nuclease digestion. ssDNA-binding protein (SSB) not only exists in most biological systems to prevent these damages but also plays an important role in recruitment of relevant proteins to regions of ssDNA to carry out their own functions. In order to efficiently distribute SSB among exposed ssDNA, it is likely that SSB is recycled and redistributed. SSB transfer between adjacent and distant DNA segments can allow rapid recycling and redistribution. Exchange of a bound SSB with another may make it possible to replace a set of proteins bound to the incumbent SSB with another set bound to the incoming SSB. Therefore, a detailed investigation of SSB transfer and exchange kinetics is expected to reveal functionally important properties. While a mechanism of interstrand exchange has been proposed, it is difficult to observe transient intermediates in most ensemble experiments (Kozlov & Lohman, 2002; Lee et al., 2014). To focus on the transfer of a single SSB to another DNA, or the exchange of one SSB for another at the single molecule level, we can use surface-tethered DNA substrates. We have previously described detailed single molecule methods to observe transitions different binding modes of SSB and SSB diffusion along short ssDNA (Roy, Kozlov, Lohman, & Ha, 2007; Zhou & Ha, 2012). For Escherichia coli SSB, there are two major binding modes, (SSB)65 and (SSB)35, which have well-defined ssDNA wrapping lengths. The numbers 65 and 35 refer to the number of nucleotides contacted by the SSB homotetramer in each mode (Roy et al., 2007). Here, we describe an assay of using a mutant SSB with a distinct binding behavior as compared to wild-type SSB (wtSSB) to observe exchange of a DNA–bound SSB for wtSSB in solution. We can also observe an exchange of a labeled SSB for an unlabeled SSB. Exchange kinetics has been measured in ensemble, and it was proposed that there must exist an intermediate of multiple SSB bound to a single ssDNA region (Kunzelmann, Morris, Chavda, Eccleston, & Webb, 2010). Using single-molecule fluorescence resonance energy transfer (smFRET), we can directly observe the real-time formation of a reaction intermediate with more than one SSB bound to the same DNA. We can also use the FRET difference between a DNA fully wrapped around SSB
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
465
and a free DNA to observe the transfer of SSB to a separate DNA strand. We can use these observations to dissect the exchange kinetics and mechanisms. In this chapter, we describe protocols for studying replacement (bound SSB being exchanged for SSB in solution) and transfer (bound SSB transferred to free ssDNA in solution) mechanisms using E. coli SSB as a model.
2. SINGLE-MOLECULE FRET EXPERIMENTAL STRATEGIES 2.1 Preparation and Assembly of Polyethylene Glycol Slides In order to perform single-molecule microscopy experiments using surfacetethered molecules, a clean and passivated surface is required. This minimizes the loss of sample protein and DNA through nonspecific adsorption, as well as potential artifacts caused by interaction with the surface. Generally, polyethylene glycol (PEG)-coated quartz slides and glass coverslips are used for single-molecule total internal reflection fluorescence (TIRF) experiments. PEG slides and coverslips for TIRF microscopy purposes were obtained from Microscope Slides Core facility in Johns Hopkins University School of Medicine. Each slide has multiple channels and is suitable for multiple experiments. 2.1.1 Materials • PEGylated quartz slide and coverslip • Double sided tape • 5-min epoxy • Tubing: WEICO ETT-28 (inner diameter ¼ 0.01500 , outer wall ¼ 0.01600 ) • Needle: BD (26 gauge, 3/800 ) • 1-mL syringe • 20–200-μL pipette tips • Vacuum sealable food saver bags 2.1.2 Procedure 1. Slides are assembled with PEG surfaces of the slide and coverslip facing each other to create passivated channels. PEGylated surfaces can be distinguished from untreated surfaces by dropping water and observing the hydrophobicity of the surface; a treated surface will be more hydrophobic.
466
Olivia Yang and Taekjip Ha
2. After testing, surfaces can be dried with house air or N2(g); PEGylated surfaces should not be physically touched or the passivation will be disrupted. 3. Double-sided tape (100 μm thick) is used as a spacer between the slide and coverslip, and to create the channels for multiple experiments (see Fig. 1). Tape is cut into thin strips on a clean surface with a clean razor blade. Do not move tape once it has been placed. 4. Place the coverslip on top of the slide and tape, PEG side down, and press with pipette tip to secure the adhesion and prevent leaks between channels. 5. Ends of the channel are sealed with 5-min epoxy as marked in yellow (see Fig. 1). Excessive epoxy used in this step may block the inlet holes drilled in the slide, so use sparingly. 6. For real-time flow experiments, additional setup is required (see Fig. 3; Section 3) 7. Slides are typically stored at 20°C in 50-mL tubes, into vacuum sealed bags to avoid moisture. Slides with unused channels can be similarly stored for later use.
2.2 DNA Substrate Preparation A DNA substrate similar to previously used for SSB-binding mode and dynamics studies is tethered to the surface for single-molecule experiments
Fig. 1 PEG slide assembly. Slides are assembled with PEG surfaces facing inward. Double-sided tape is placed as marked in gray, between holes drilled. The long edge of the coverslip and slide is sealed with 5-min epoxy (yellow region). One channel is the space between two tapes.
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
467
(Roy et al., 2007; Zhou et al., 2011). The partial duplex substrate is made of an 18 bp duplex with a polydeoxythymidine (poly-dT) overhang of varying lengths. The 18 bp duplex is used as a spacer to minimize potential surface effects. Poly dT is used for its lack of secondary structure and high binding affinity to SSB, although other sequences may also work (Lohman & Overman, 1985). The length of the ssDNA used is approximately 70 nt, enough to allow for a full wrapping of SSB ((SSB)65 mode) or two partially wrapped SSB ((SSB)35 mode). The end of the duplex without the ssDNA overhang is biotinylated to specifically tether the DNA to the surface, while the other end of the duplex is Cy5 labeled. For smFRET experiments, cyanine dyes Cy3 and Cy5 are commonly used for donor and acceptor molecules, respectively, on DNA due to their high photostability under deoxygenated conditions and high brightness. A detailed labeling procedure is previously published (Joo & Ha, 2012). For experiments with labeled SSB, the substrate is labeled with Cy5 only. For unlabeled SSB, the far end of the ssDNA is additionally labeled in order to observe FRET changes upon SSB binding. SSB is labeled with an average of one Alexa 555, a small organic dye with excitation and emission spectra similar to Cy3, per SSB tetramer as previously described (Lee et al., 2014; Roy, Kozlov, Lohman, & Ha, 2009; Zhou et al., 2011). The DNA oligonucleotides (oligos) used are listed below: 1. 50 -/Cy5/ GCC TCG CTG CCG TCG CCA -/biotin/-30 . 2. 50 -TGG CGA CGG CAG CGA GGC (T)m-/Cy3/-30 (m ¼ 72). 3. 50 -TGG CGA CGG CAG CGA GGC (T)m-30 (m ¼ 70). 4. 50 -(T)m-30 (m ¼ 60). The DNA substrate for labeled SSB experiments was annealed by mixing oligos 1 and 3 at 10 μM and 20 μM, respectively, in 10 mM Tris–HCl (pH 8) and 50 mM NaCl, then slow cooling from 95°C to 4°C over 4 h. The cooling can be done in a thermocycler at a rate of 1°C/min, or with a heat block taken off heat and allowed to cool to RT then stored at 20°C. The substrate for unlabeled SSB experiments was prepared similarly, but using oligos 1 and 2. Typical annealing uses an excess (1.2 to 2) of nonbiotinylated strands in order to ensure that surface-bound DNA contain both strands. DNA substrates are stored at 20°C, covered in foil to prevent photobleaching. For single-molecule experiments, a substock of 5–10 nM DNA is prepared. At lower concentrations, DNA can be lost due to nonspecific adsorption to tube surfaces. 1% v/v BSA (20 mg/mL stock) in solution can act to passivate surfaces and prevent some loss.
468
Olivia Yang and Taekjip Ha
2.3 Single-Molecule FRET With TIR Microscopy A detailed explanation of the prism-type TIRF microscope setup used in these protocols can be found as previously published (Joo & Ha, 2012). Video recordings were processed to extract single molecule fluorescence intensities at each frame, and custom written scripts were used to calculate FRET efficiencies. Data acquisition and analysis software can be downloaded from https://cplc.illinois.edu/software/. Alternating laser excitation (ALEX) is used to filter out molecules with donor only or acceptor only (Kapanidis et al., 2005). We require a minimum intensity cutoff during donor (green) or acceptor (red) excitation. If a fluorescence spot has a combined intensity from donor and acceptor channels less than the cutoff, then it is not counted. Stoichiometry parameter S can also be used to identify the donor and acceptor only spots. The stoichiometry S of each molecule is calculated as S¼
IAG + IDG IAG + IDG + IAR
G Where IG A is the acceptor intensity during green excitation, ID is the donor intensity during green excitation, and IR A is the acceptor intensity during red excitation. Molecules with low S values (0–0.2) correspond to acceptor only molecules, while molecules with high S values (0.8–1) correspond to donor only molecules. Molecules with both donor and acceptor have stoichiometry S 0.5. Our typical data acquisition sequence is an initial set of 10 frames of red laser excitation, followed by green laser excitation. FRET efficiency is calculated from green excitation periods. Long movies had an additional 10 frames of red excitation at the end to test if the acceptor has photobleached during data acquisition. A “snapshot” is defined as 20 frames taken at a single imaging area. “Long movies” are defined as enough frames to photobleach 80% of the molecules within the imaging area, usually around 2000–3000 frames. Leakage of donor signal to the acceptor channel and background signals of both channels were corrected similar to as previously described (Roy, Hohng, & Ha, 2008). Apparent FRET efficiency is calculated with the equation
EFRET ¼
IA leakage ID IA leakage ID + ID
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
469
where IA is the acceptor intensity, leakage is the leakage factor from the donor to the acceptor detection channel, and ID is the donor intensity. Leakage factor will vary depending on the microscope setup used. Background for long movies is corrected to make the intensities after donor photobleaching equal to zero. Histograms were made from at least 10 individual snapshots of a single sample, with approximately 200–300 molecules imaged per area for a total of 2000–3000 molecules used per histogram. The FRET efficiency of each molecule was calculated based on the average intensity of donor and acceptor over eight frames, avoiding the frames around when excitation is changing.
2.4 SSB Replacement Assay In this chapter, we describe the use of E. coli SSB’s well-characterized and controlled binding modes (Roy et al., 2007). The method of purification and labeling of SSB can be found as previously published (Lohman, Green, & Beyer, 1986; Roy et al., 2009). Because we want to observe primarily exchange of a single SSB ((SSB)65), we use NaCl concentrations greater than or equal to 100 mM; any lower NaCl concentration and there would be a significant population of two SSB molecules bound to 70-mer ssDNA ((SSB)35 mode). Similarly, SSB concentration should be kept relatively low (<10 nM in 100 mM NaCl) to minimize binding in the (SSB)35 mode. For an SSB variant with lower binding affinity, lower concentrations of wtSSB will be required for displacement. With only free DNA, 1 nM SSB is expected to bind almost fully within 1 min in high salt conditions. 2.4.1 Materials • T50 buffer: 10 mM Tris–HCl (pH 8.0), 50 mM NaCl in ddH2O, filtered through 0.22-μm membrane • SSB imaging buffer (SSB IB 100 mM NaCl): 100 mM NaCl, 10 mM Tris–HCl (pH 8.0), 0.1 mg/mL BSA, 0.8% (w/v) dextrose, 165 U/mL glucose oxidase, 2170 U/mL catalase, 2-3 mM Trolox, and indicated amounts of SSB. See Section 3. • Neutravidin: 0.2 mg/mL neutravidin diluted in T50 buffer • Assembled PEG slide (see above) • Prism-type TIRF microscope • Labeled DNA substrate (oligomers 1 + 3, see above) • Unlabeled DNA substrate (oligomers 1 + 2, see above) • SSB stock: to be kept at 20°C, secondary stock can be made in SSB IB and kept on ice during the course of one set of experiments.
470
Olivia Yang and Taekjip Ha
2.4.2 Procedure 2.4.2.1 Experiment Procedure
1. Confirm quality of slide (see Section 3). 2. Flow in 20 μL T50 buffer to each channel expected to be used for one experiment. The approximate channel volume is about 20 μL. For thorough flushing or between different buffer conditions, use at least 30 μL of solution. 3. Flow in 20 μL neutravidin to each channel. Incubate 1–2 min, then flush with T50. 4. Prepare 10–30 pM DNA substrate in T50 (“labeled” for labeled SSB displacement, “unlabeled” for mutant SSB displacement) and add to channel. Incubate for 1–2 min, then flush with SSB IB + “gloxy” to check the spot density. For “labeled” substrate, there should only be Cy5 signal, while “unlabeled”substrate should have both Cy3 and Cy5 signal. There should be 200–300 spots of a single dye within one imaging area. If the number of spots is too little, dilute more DNA and add to channel until desired density is reached. 5. Flush with SSB IB + gloxy. Take snapshots to generate DNA only histogram. 6. Add 40 μL of 1 nM SSB (labeled or mutant) in SSB IB 100 mM NaCl and incubate for at least 5 min (Fig. 2A). 7. Remove unbound SSB by flushing with SSB IB 100 mM NaCl + gloxy (Fig. 2B). Take snapshots to get initial time point. 8. Add wtSSB at desired concentration. To observe real-time exchange, a flow cell assembly must be used (see Fig. 3; Section 3). 9. Take at least 10 snapshots at each given time point (0–30 min).
Fig. 2 Scheme of SSB replacement assay. (A) Labeled SSB is added to ssDNA tethered to the PEG slide. (B) Unbound labeled SSB is flushed from sample chamber. (C) wtSSB (dashed outline) is added to sample chamber. (D) Labeled SSB is displaced from tethered DNA.
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
471
Fig. 3 Real-time flow experiment setup—cross section of single channel. Pipette tip segments are stuck to the quartz slide with epoxy. “Sample well” holds the sample to flow in. Teflon tubing is stuck into pipette tip which has been adhered to hole with epoxy. Tubing is attached to a 1-mL syringe with 26-gauge needle.
2.4.2.2 Data Analysis
For labeled SSB, replacement can be observed as the disappearance of fluorescent spots over time. Because the DNA substrate used is labeled with Cy5, we can use Alexa 555 as a label on SSB to distinguish it from unlabeled SSB and to observe FRET upon binding. We include in our analysis only those spots with both the acceptor (Cy5 on DNA) and donor (Alexa 555 labeled SSB) signals using ALEX. The two color colocalization allows us to reject SSB bound to the surface nonspecifically from analysis, and FRET provides an additionally stringent criterion for specific binding. Fluorescence spots with excessively high or low intensity are also rejected because they typically are not due to the molecules of interest. Generally, spots will also disappear due to photobleaching, so it is important to use the condition with no wtSSB added to determine a photobleaching rate. It is important to keep the laser intensity constant to avoid changing this photobleaching rate too much between different experiment repeats. The number of spots is normalized to the initial time point, since for a given experiment the actual spot density at the beginning of experiment may vary. The displacement rate can be determined from a single exponential decay fit, then subtracting the photobleaching rate determined separately. Plotting the rate of spot disappearance against concentration of wtSSB added shows a linear relation.
472
Olivia Yang and Taekjip Ha
For a mutant SSB with less than full wrapping properties, the exchange is observed through a shift in the FRET efficiency. As shown in Fig. 4, the mutant has a FRET efficiency centered around 0.5 (Fig. 4C, top panel), while wtSSB has a FRET efficiency about 0.7 (Fig. 4C, bottom panel). In order to confirm what FRET efficiencies each species corresponds with, controls should be done to see binding in the same buffer conditions at low [SSB]. The simplest way of determining exchange is imposing a FRET cutoff (0.6) to distinguish the mutant-bound DNA population vs the wtSSBbound population. For variants with more overlapping FRET efficiencies, it would be more reasonable to fit the histogram with a Gaussian function for each population and use the area to calculate the population ratio. Using this ratio, we can plot the binding of wtSSB over time as it replaces the mutant. Similar to the labeled SSB assay, we can use this plot to obtain exchange rates at various concentrations of wtSSB. Movies taken during real-time replacement assays will result in FRET traces as sketched in Fig. 4B. Three species are expected to exist in this experiment: the initial, mutant-bound SSB (0.5 FRET); the transient, 2 SSB-bound state with one mutant and one wild type (0.2 FRET, (SSB)35); and the final, wtSSB-bound species (0.7 FRET) (see Fig. 4A) (Roy et al., 2007). Through measuring dwell times of each FRET state on a single molecule trace, the rates of exchange between these three species can be determined. In this case, since we flood the system with wtSSB in a high salt buffer, we expect the reaction to be pushed toward the wtSSBbound state. The EFRET observed in the single molecule traces (Fig. 4B) should correspond with the centers of peaks as observed in histograms (Fig. 4C). Because the (SSB)35 mode is not favored in these salt conditions, it does not exist long enough to be captured in histograms. However, it can be seen as an intermediate during real-time experiments.
2.5 SSB Transfer Assay For this experiment, we use (dT)60 because the expected binding site size for a single SSB is approximately 65 nt. Varying lengths or sequences of DNA oligomers can be tested for transfer activity with the same assay, though may require different concentration ranges. Sequences with secondary structures may also be inefficient in competing with the poly-dT substrate used. 2.5.1 Materials The same materials as listed in the SSB displacement assay are used. Additional/alternative materials required are listed below.
Fig. 4 Results from mutant to wtSSB displacement (A) Scheme of displacement mechanism. (B) Ideal FRET trace of mutant to wtSSB in a “two-step” transition. (C) FRET histograms of populations observed during experiment.
474
• •
Olivia Yang and Taekjip Ha
dT60 (oligomer 4): stock diluted in T50, stored at 20°C SSB imaging buffer (SSB IB 500 mM NaCl): 500 mM NaCl, 10 mM Tris–HCl (pH 8.0), 0.1 mg/mL BSA, 0.8% (w/v) dextrose, 165 U/mL glucose oxidase, 2170 U/mL catalase, 2-3 mM Trolox, and indicated amounts of SSB.
2.5.2 Procedure 2.5.2.1 Experiment Procedure
The same procedure for SSB replacement is followed for SSB transfer, but rather than SSB added at step 6, dT60 at varying concentrations is used in a SSB IB 500 mM NaCl buffer, and throughout the experiment SSB IB 500 mM NaCl is used in place of SSB IB 100 mM NaCl. 2.5.2.2 Data Analysis
The initial state of the tethered substrate is bound by a single SSB, so shows a EFRET 0.7. The EFRET of unbound ssDNA of a dT70 construct in 500 mM NaCl is 0.15. The exact value will vary slightly based on NaCl or other salt concentrations. Since the difference in EFRET is large, a cutoff can be used to determine the bound and unbound DNA substrate populations.
3. NOTES Imaging buffer is generally prepared as follows: 1. Trolox + dextrose solution: 10 mg Trolox (()-6-Hydroxy-2,5,7,8tetramethylchromane-2-carboxylic acid, Sigma-Aldrich) and 80 mg dextrose (Sigma-Aldrich) is dissolved in 10 mL ddH2O and 10 μL 5 M NaOH. Mixture is rotated for at least 4 h, RT, covered in foil. Solution is filtered through 0.22-μm membrane and stored at 4°C. This solution can be used for 2–3 weeks when stored at 4°C, or longer if stored frozen. 2. SSB IB: NaCl, Tris–HCl (pH 8.0), and BSA are added to Trolox + dextrose solution to the final concentrations listed above to make a flush buffer for removing free protein from channel. NaCl, Tris– HCl, and BSA are added with SSB to Trolox + dextrose solution to final concentrations as needed for steps adding SSB to channel. 3. “Gloxy”: 10 mg glucose oxidase (G2133, Sigma-Aldrich) and 20 μL bovine liver catalase (C30, Sigma-Aldrich) is dissolved in 100 μL T50 buffer. Mixture is spun down for 2 min at 10,000 g. The top 100 μL of the solution is transferred and stored at 4°C, and can be used for
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
475
2–4 weeks. This solution is 100 concentrated, and is typically not added until right before samples are added to the slide. Upon mixing with IB, the solution will continually become more acidic upon reacting with atmospheric oxygen (Joo & Ha, 2012). Slides should always be checked for cleanliness and passivation before use, as follows: 1. Cleanliness: With just T50 in the channel, there should be fewer than 10 observable fluorescent spots per imaging area. 2. Passivation: After flowing in 1 nM labeled DNA or protein, there should be less than 30 observable spots stuck to the surface. After flushing with T50, there should be significantly less spots compared to earlier. Protein is generally stickier than DNA and is expected to have slightly more nonspecific binding. 3. Only after the channel has passed these checks should the user proceed with the experiment. Real-time flow experiments have other considerations: 1. 20–200-μL pipette tips are used in this setup. The sample well is made from the wider portion of the tip, while the outlet side is the sharper portion of the tip. Both are adhered to the slide with epoxy. Fig. 3 shows the cross section of a channel assembled for a real-time flow experiment. 2. It is not necessary to epoxy the tubing to the tip, though it helps make the assembly more secure. Care should be taken to avoid clogging the drilled holes with epoxy. 3. A minimum of 40 μL of sample must be added to the sample well to avoid drawing bubbles through the channel. 4. Sample is drawn through by pulling with a syringe. This can be done manually, or in a more controlled way with a syringe pump. With a syringe pump, the exact flow time and rate can also be controlled. At no point should the sample be pushed back through the channel. 5. All solutions added to this channel will be drawn through by adding the solution to the sample well and pulling through with the syringe. 6. Because a sample is being pulled through during the recording of a movie, care must be taken that the pulling action does not perturb the sample stage or any part of the microscope. Slight motions of the slide or sample will affect the focus and the quality of the movie.
476
Olivia Yang and Taekjip Ha
7. After the initial movie is taken, further long movies or snapshots can be taken of the same channel. 8. Tubing and syringes can be reused, but fresh pipette tips should be used for adhering to the slide. Other general notes: 1. During imaging, general exposure time used is 50–100 ms. Highest time resolution for the camera used in these experiments (Andor DU-897E-CSO-#BV) is 30 ms for 512 512 pixels, though the imaging area can be decreased to increase time resolution. To avoid photobleaching, a longer exposure time should be used to minimize laser intensity. To observe faster dynamics, shorter exposure times should be used, but higher laser intensities will be required to increase signal to noise ratios. 2. Slides can be reused. By soaking in methanol, epoxy is weakened and the coverslip and double sided tape can also be removed and disposed of. Then, slides should be boiled in water and any visible junk can be washed off before proceeding with usual treatment (Roy et al., 2008). 3. All concentrations of SSB in this text refer to concentration of SSB tetramer. 4. DNA substrate concentration refers to the concentration of the biotinylated strand. Nonbiotinylated strand is used in excess in order to maximize the amount of annealed DNA that gets specifically tethered to the slide. Nonbiotinylated strand will get flushed from the channel and should not interfere with experiments. 5. High concentrations of DTT cannot be used in imaging buffers for single-molecule experiments, as blinking will be observed.
4. SUMMARY smFRET and single molecule experiments are especially useful for understanding more detailed dynamics of SSB, especially on a small length scale. These details may be helpful to explain steps in protein recruitment processes during replication, recombination, and repair related to ssDNA processing. Though E. coli SSB was used throughout these experiments, similar single molecule studies regarding transfer and replacement mechanisms can be done with the same method for other relevant SSB variants to further understand the differences between SSBs from different organisms. However, the binding of each new variant needs to be well
Single-Molecule Studies of ssDNA-Binding Proteins Exchange
477
characterized for a given substrate before observing exchange in order to properly attribute a given FRET efficiency to a specific variant. Using single-molecule methods can be useful for observing and confirming intermediates as proposed by other ensemble scale SSB experiments.
ACKNOWLEDGMENTS The authors would like to thank Dr. Jichuan Zhang for his guidance and advice throughout this project, and Prof. I-Ren Lee for his helpful ideas. Research in this chapter was funded by NIH Grant R35 GM 122569.
REFERENCES Joo, C., & Ha, T. (2012). Single-molecule FRET with total internal reflection microscopy. Cold Spring Harbor Protocols, 2012(12), 1223–1237. https://doi.org/10.1101/pdb. top072058. pdb.top072058. Kapanidis, A. N., Laurence, T. A., Nam, K. L., Margeat, E., Kong, X., & Weiss, S. (2005). Alternating-laser excitation of single molecules. Accounts of Chemical Research, 38(7), 523–533. https://doi.org/10.1021/ar0401348. Kozlov, A. G., & Lohman, T. M. (2002). Kinetic mechanism of direct transfer of Escherichia coli SSB tetramers between single-stranded DNA molecules. Biochemistry, 41(39), 11611–11627. https://doi.org/10.1021/BI020361M. Kunzelmann, S., Morris, C., Chavda, A. P., Eccleston, J. F., & Webb, M. R. (2010). Mechanism of interaction between single-stranded DNA binding protein and DNA. Biochemistry, 49, 843–852. https://doi.org/10.1021/bi901743k. Lee, K. S., Marciel, A. B., Kozlov, A. G., Schroeder, C. M., Lohman, T. M., & Ha, T. (2014). Ultrafast redistribution of E. coli SSB along long single-stranded DNA via intersegment transfer. Journal of Molecular Biology, 426(13), 2413–2421. https://doi.org/ 10.1016/j.jmb.2014.04.023. Lohman, T. M., Green, J. M., & Beyer, R. S. (1986). Large-scale overproduction and rapid purification of the Escherichia coli ssb gene product. Expression of the ssb gene under lambda PL control. Biochemistry, 25(1), 21–25. https://doi.org/10.1021/ bi00349a004. Lohman, T. M., & Overman, L. B. (1985). Two binding modes in Escherichia coli single strand binding protein-single stranded DNA complexes: Modulation by NaCl concentration. The Journal of Biological Chemistry, 260(6), 3594–3603. Retrieved from http:// www.jbc.org/content/260/6/3594.full.pdf. Roy, R., Hohng, S., & Ha, T. (2008). A practical guide to single-molecule FRET. Nature Methods, 5(6), 507–516. https://doi.org/10.1038/nmeth.1208. Roy, R., Kozlov, A. G., Lohman, T. M., & Ha, T. (2007). Dynamic structural rearrangements between DNA binding modes of E. coli SSB protein. Journal of Molecular Biology, 369(5), 1244–1257. https://doi.org/10.1016/j.jmb.2007.03.079. Roy, R., Kozlov, A. G., Lohman, T. M., & Ha, T. (2009). SSB protein diffusion on singlestranded DNA stimulates RecA filament formation. Nature, 461, 1092–1097. https:// doi.org/10.1038/nature08442. Zhou, R., & Ha, T. (2012). Single-molecule analysis of SSB dynamics on single-stranded DNA. In Single-stranded DNA binding proteins (pp. 85–100). Totowa, NJ: Humana Press. https://doi.org/10.1007/978-1-62703-032-8_5. Zhou, R., Kozlov, A. G., Roy, R., Zhang, J., Korolev, S., Lohman, T. M., et al. (2011). SSB functions as a sliding platform that migrates on DNA via reptation. Cell, 146(2), 222–232. https://doi.org/10.1016/j.cell.2011.06.036.