Watching individual proteins acting on single molecules of DNA

C H A P T E R T H I R T E E N Watching Individual Proteins Acting on Single Molecules of DNA Ichiro Amitani,*,†,1 Bian Liu,*,†,‡,1 Christopher C. Dombrowski,*,† Ronald J. Baskin,† and Stephen C. Kowalczykowski*,†,‡ Contents 1. Introduction 2. Preparation of DNA Substrates 2.1. Preparation of biotinylated l DNA 2.2. Preparation of DNA–bead complexes 2.3. Preparation of DNA–bead complexes end-labeled with Cy3-labeled antibody 3. Preparation of Fluorescent Proteins 3.1. RecBCD labeled with a fluorescent nanoparticle (RecBCD–nanoparticle) 3.2. Rad54/Tid1 labeled with a fluorescent antibody (FITC–Rad54/Tid1) 3.3. Chemically modified fluorescent RecA or Rad51 proteins (RecAFAM/Rad51FAM) 4. Instrument 4.1. Flow cell design 4.2. Flow cell fabrication 4.3. Microscope with laser trap and microfluidic system 4.4. Temperature determination and control 5. Single-Molecule Imaging of Proteins on DNA 5.1. Unwinding of DNA by a single RecBCD enzyme 5.2. Direct observation of RecBCD–nanoparticle translocation 5.3. Rad54/Tid1 translocation 5.4. Real-time Rad51 assembly 5.5. Real-time Rad51 disassembly 5.6. Visualization of RecAFAM/RecA-RFP/Rad51FAM filament formation 262 265 265 265 266 268 268 268 269 270 270 271 273 276 280 281 281 283 284 285 286 * Department of Microbiology, University of California, Davis, California, USA Department of Molecular and Cellular Biology, University of California, Davis, California, USA Biophysics Graduate Group, University of California, Davis, California USA 1 These authors contributed equally to this work. { { Methods in Enzymology, Volume 472 ISSN 0076-6879, DOI: 10.1016/S0076-6879(10)72007-3 # 2010 Elsevier Inc. All rights reserved. 261 262 Ichiro Amitani et al. 6. Data Analysis Methods 6.1. Two-dimensional Gaussian fitting 6.2. Automatic DNA length measurement Acknowledgments References 287 288 288 289 289 Abstract In traditional biochemical experiments, the behavior of individual proteins is obscured by ensemble averaging. To better understand the behavior of proteins that bind to and/or translocate on DNA, we have developed instrumentation that uses optical trapping, microfluidic solution delivery, and fluorescent microscopy to visualize either individual proteins or assemblies of proteins acting on single molecules of DNA. The general experimental design involves attaching a single DNA molecule to a polystyrene microsphere that is then used as a microscopic handle to manipulate individual DNA molecules with a laser trap. Visualization is achieved by fluorescently labeling either the DNA or the protein of interest, followed by direct imaging using high-sensitivity fluorescence microscopy. We describe the sample preparation and instrumentation used to visualize the interaction of individual proteins with single molecules of DNA. As examples, we describe the application of these methods to the study of proteins involved in recombination-mediated DNA repair, a process essential for the maintenance of genomic integrity. 1. Introduction In traditional ensemble experiments, the behavior of individual proteins is averaged by the obligatory need to study a population of molecules. However, it has become increasingly evident that the analysis of single molecules is not only possible, but that it can reveal novel information about the behavior and function of enzymes (see, e.g., Amitani et al., 2006; Bianco et al., 2001; Galletto et al., 2006; Handa et al., 2005; Nimonkar et al., 2007; Spies et al., 2003, 2007). To better understand the molecular behavior of individual proteins, we have used optical trapping to capture and visualize the action of individual proteins on single molecules of DNA (Bianco et al., 2001). The general experimental design involves attaching a single DNA molecule to a polystyrene microsphere. The microsphere is then used as a handle to manipulate the DNA molecule. Visualization is achieved by using a fluorescence microscope to image fluorescently labeled DNA or protein (Amitani et al., 2006; Bianco et al., 2001; Galletto et al., 2006; Handa et al., 2005, 2009; Hilario et al., 2009). To both extend the DNA and exchange solutions rapidly, we designed and fabricated multichannel microfluidic flow cells that provide parallel paths for different solutions that remain separated by laminar flow (Fig. 13.1) (Bianco et al., 2001; Brewer and 263 Watching Individual Proteins Acting on Single Molecules of DNA Divider 100 mm A 750 mm B 200 mm Region inside the mean diffusion boundary 10 mm 4.5 mm Figure 13.1 Illustration of the three-channel flow cells used in the single-molecule experiments. (A) Photograph of a three-channel flow cell. The flow cell is fabricated using the process described in Section 4.2. To demonstrate the flow path, green dye flows through channels I and III, whereas yellow dye flows through channel 2. (B) Schematic (drawing not to scale) of a three-channel flow cell showing typical dimensions; magnification shows the detail at the end of channel divider. The divider is 100 mm wide with a semicircular end of radius of  50 mm. The gray area to the right of the divider illustrates the region inside the mean diffusion length boundary. Experiments are conducted at a point  200 mm downstream of the divider,  750 mm into each channel, and  35 mm from the surface where the effects of diffusion are minimal. Bianco, 2008). These flow paths are used to introduce the optically trapped DNA to solutions that contain the proteins of interest, or that permit the controlled initiation of enzymatic reactions. We have applied these methods to the study of proteins involved in recombinational DNA repair, a conserved biological process responsible for the repair of DNA breaks. A DNA double-strand break (DSB) is a lethal type of DNA damage. These breaks are constantly created by many endogenous and exogenous sources in cells. Because an unrepaired DSB often leads to cell death, all organisms have evolved various methods to repair broken DNA. Among them, homologous recombination (HR) is the most accurate method for DSB repair (Kowalczykowski, 2000). The process of HR consists of three stages. First, the end of a broken double-stranded DNA (dsDNA) molecule is processed by helicase and nuclease to generate a 30 -ended, single-stranded DNA (ssDNA) tail onto which a DNA strand exchange protein self-assembles. Second, this protein– ssDNA complex searches for homology on a donor dsDNA molecule and then catalyzes the pairing and exchange of DNA strands. Finally, the heteroduplex DNA product is resolved (Kowalczykowski, 2000). 264 Ichiro Amitani et al. In Escherichia coli, the RecBCD helicase/nuclease is responsible for the resection of dsDNA ends (Spies and Kowalczykowski, 2005). RecBCD is a bipolar DNA helicase and nuclease (Dillingham and Kowalczykowski, 2008). It unwinds and degrades dsDNA rapidly and processively (Bianco et al., 2001; Roman and Kowalczykowski, 1989). Its biological activities are regulated by an octameric DNA sequence called Chi (Crossover hotspot instigator, Chi: 50 -GCTGGTGG-30 ) (Dillingham and Kowalczykowski, 2008). Single-molecule analysis uniquely revealed that, upon interaction with Chi, the enzyme pauses for a few seconds, and then it translocates at a reduced rate due to a switch in motor usage (Spies et al., 2003, 2007). The interaction also downregulates the nuclease activity (Dixon and Kowalczykowski, 1993) and switches the polarity of DNA degradation (Anderson and Kowalczykowski, 1997a). These alterations of nuclease activity generate a processed dsDNA ending with a 30 -ssDNA tail (Taylor and Smith, 1995) onto which RecA is loaded by RecBCD to form a nucleoprotein filament (Anderson and Kowalczykowski, 1997b). DNA strand exchange is catalyzed by RecA in bacteria and Rad51 in eukaryotes (Bianco et al., 1998). Both RecA and Rad51 form a helical nucleoprotein filament on either ssDNA or dsDNA in the presence of ATP. In the filament, RecA/Rad51 occupies 3 nucleotides or base pairs (depending on whether ssDNA or dsDNA is used), and it stretches DNA to 150% of its B-form DNA length (Benson et al., 1994; Chen et al., 2008; Conway et al., 2004; Ogawa et al., 1993; Stasiak et al., 1981). Although the typical active form of RecA/Rad51 is the ssDNA–RecA/Rad51 complex, when assembled on dsDNA, RecA can also promote DNA pairing with ssDNA (Zaitsev and Kowalczykowski, 2000). However, when RecA/Rad51 forms a complex with dsDNA, DNA strand exchange with a RecA/Rad51– ssDNA complex is impeded, resulting in defective recombination (Campbell and Davis, 1999; Sung and Robberson, 1995). Assembly of RecA/Rad51 nucleoprotein filaments occurs by nucleation and growth, a process that was imaged at the single-molecule level (Galletto et al., 2006; Handa et al., 2009; Hilario et al., 2009; Modesti et al., 2007; Prasad et al., 2006; Robertson et al., 2009; van der Heijden et al., 2007). In eukaryotes, the inhibitory Rad51 bound to chromosomes is removed by Rad54 (Solinger et al., 2002), a chromatin-remodeling protein (Alexeev et al., 2003). Rad54 and Tid1, a Rad54 homolog with an important role in meiosis (Klein, 1997; Shinohara et al., 1997), work together with Rad51 (Mazin et al., 2000, 2003; Petukhova et al., 1998; Solinger and Heyer, 2001; Solinger et al., 2001, 2002) and Dmc1, the meiotic Rad51 homolog (Holzen et al., 2006; Shinohara et al., 2000), respectively. Both Rad54 and Tid1 are dsDNA translocases as defined by the direct visualization of their movements on individual DNA molecules (Amitani et al., 2006; Nimonkar et al., 2007; Prasad et al., 2007). Watching Individual Proteins Acting on Single Molecules of DNA 265 In this chapter, we detail the sample preparation and instrumentation used to visualize the interaction of individual proteins with single molecules of DNA. 2. Preparation of DNA Substrates 2.1. Preparation of biotinylated l DNA Bacteriophage l DNA (New England Biolabs, Ipswich, MA) is biotinylated by ligation to a 30 -biotinylated 12-mer oligonucleotide (50 -GGGCGGCG ACCT-30 or 50 -AGGTCGCCGCCC-30 , Operon Technologies, Huntsville, AL) that is complementary to one of the cohesive ends of l DNA (Bianco et al., 2001). In all the subsequent protocols, the pipetting of solutions containing l DNA should be performed with cut pipette tips to minimize shearing of the DNA. 1. Phosphorylate the oligonucleotide by incubating the oligonucleotide (5 mM) in 50 ml of polynucleotide kinase (PNK) buffer (5 mM dithiothreitol (DTT)), 10 mM MgCl2, 70 mM Tris–HCl (pH 7.6), 1 mM ATP, and 0.2 U/ml PNK at 37  C for 1 h. 2. Stop the reaction by incubation at 75  C for 10 min. 3. Anneal the phosphorylated oligonucleotide and the l DNA by preparing a reaction (90 ml) containing 28 ng/ml of l DNA, 0.56 mM phosphorylated oligonucleotide, and 100 mM NaCl. 4. Incubate the reaction at 75  C for 20 min in a heat block to denature the annealed cohesive ends of the l DNA. 5. Remove the heat block and place it on the bench to slowly cool the reaction to room temperature (2–3 h), and then chill the reaction on ice. 6. Ligate the phosphorylated oligonucleotide to the l DNA by adding 10 ml of 10 T4 DNA ligase buffer (10 mM ATP, 100 mM DTT, 100 mM MgCl2, 500 mM Tris–HCl, pH 7.5) and 1 ml of T4 ligase (400 Units) to the annealing reaction from the previous step. 7. Incubate the reaction at 16  C overnight or at room temperature for 1 h. 8. Inactivate the ligase by incubating at 75  C for 10 min. 9. Remove excess oligonucleotide and ATP by filtration through a spin column (MicroSpin S-400 HR, GE Healthcare, Piscataway, NJ). 2.2. Preparation of DNA–bead complexes DNA–bead complexes are prepared by incubating 1 ml of 35 pM streptavidin-coated polystyrene beads (1.0 mm, Bangs Laboratories, Fishers, IN), 1 ml of 100 mM NaHCO3 (pH 8.3), and 2 ml of 100 pM biotinylated 266 Ichiro Amitani et al. l DNA for 1 h at 37  C. The ratio of beads to DNA may be varied and optimized for different experiments. To fluorescently stain the DNA, add 500 ml of sample buffer (see below for experiment-specific recipes) containing 20 nM YOYO-1 (Invitrogen, Carlsbad, CA) to the DNA–bead complex and stain in the dark at room temperature for at least 1 h. The dye to DNA (in base pairs) ratio can be altered to vary from 1:1 to 1:5. The sample buffer is degassed for at least 1 h to remove oxygen and to reduce oxygen-mediated photobleaching and cleavage of DNA. 2.3. Preparation of DNA–bead complexes end-labeled with Cy3-labeled antibody 2.3.1. Fluorescent secondary antibody To visualize the end of a DNA molecule in order to measure its length without the use of a nonspecifically binding dye such as YOYO-1, we attach a fluorescent tag at the free end of the DNA–bead complex (Hilario et al., 2009). To obtain a strong signal for imaging, we fluorescently label a secondary antibody and bind it to a primary antibody that is bound to the end of DNA, which is labeled with digoxigenin (DIG). 1. Exchange the storage solution of donkey antisheep IgG antibody (50 ml, 2 mg/ml, Millipore, Billerica, MA) to a buffer lacking primary amines by using a P30 spin column (850  g for 4 min; Bio-Rad, Hercules, CA) equilibrated with labeling buffer (50 mM sodium borate (pH 9.3), 140 mM NaCl, and 2.7 mM KCl). 2. Add a 20-fold molar excess of Cy3 succinimidyl ester (Cy3-NHS, GE Healthcare) and incubate at room temperature for 1 h in the dark. 3. Remove the unreacted Cy3-NHS with a P30 spin column equilibrated with phosphate buffered saline (PBS; 10 mM Na2HPO4, 1.8 mM KH2PO4, (pH 7.4), 137 mM NaCl, and 2.7 mM KCl). 4. Determine the Cy3 and antibody concentrations by using the extinction coefficients e552 ¼ 1.5105 M 1cm 1 for Cy3, and e280 ¼ 1.7  105 M 1cm 1 for the antibody. The effect of absorption by Cy3 at 280 nm is corrected by: [antibody] ¼ (A280 – (0.08  A552))/ 1.7  105 (GE Healthcare, Amersham product booklet, ‘‘CyDyeTM monoreactive NHS Esters’’). 5. Determine the degree of labeling by calculating the ratio of Cy3 and antibody concentrations. A typical degree of labeling is 6–8 dyes/ protein. 6. Store Cy3-antisheep antibody at 4  C in the dark and use within a few days. Watching Individual Proteins Acting on Single Molecules of DNA 267 2.3.2. DNA labeled with biotin and digoxigenin (Biotin-l DNA–DIG) Bacteriophage l DNA that is labeled with biotin at one end and DIG at the other end is prepared by attaching a biotin-labeled oligonucleotide and DIG-labeled oligonucleotide to opposite cohesive ends of l DNA in successive steps. 1. Incubate 750 pM of l DNA (molecules) with 375 nM of DIG-labeled oligonucleotide (Operon Technologies) in 90 ml of 100 mM NaCl at 75  C for 15 min in a heat block. 2. Remove the heat block and place it on the bench to slowly cool the reaction mixture to room temperature (2–3 h). 3. Add 10 ml of 10 T4 DNA ligase buffer and T4 DNA ligase to a final concentration of 4 U/ml. 4. Incubate at room temperature for 1 h. 5. Inactivate the DNA ligase at 65  C for 10 min. 6. Remove unreacted DIG-oligonucleotide with an S-400 spin column (850  g for 5 min) equilibrated with TE buffer (10 mM Tris–HCl (pH 7.5), 1 mM EDTA). 7. Add 50-fold molar excess of biotinylated-oligonucleotide and 4 U/ml T4 DNA ligase to DIG-labeled l DNA. 8. Incubate at room temperature for 1 h. 9. Inactivate the DNA ligase at 65  C for 10 min. 2.3.3. Binding Cy3-labeled antibody to the DNA–bead complex DNA–bead complexes that are end-labeled with Cy3-labeled antibody are prepared by binding the sheep anti-DIG antibody to DNA–bead complex, and then binding the Cy3-antisheep secondary antibody to the anti-DIG antibody. 1. Attach the biotin-l DNA–DIG to streptavidin-coated beads as described in Section 2.2. 2. Add bovine serum albumin (BSA; stock 10 mg/ml) and sheep anti-DIG antibody (stock 200 g/ml) to final concentrations of 1 mg/ml and 20 g/ml, respectively. 3. Incubate at room temperature for 2 min. 4. Add Cy3-antisheep IgG antibody to a final concentration of 60 g/ml. The final volume is 7 l. 5. Incubate at room temperature for 2 min. 6. Immediately dilute the Cy3-antibody end-labeled DNA–bead complex into 400 l of single-molecule buffer (SMB; 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose). The final concentration of streptavidin-beads is about 90 fM. 268 Ichiro Amitani et al. 3. Preparation of Fluorescent Proteins 3.1. RecBCD labeled with a fluorescent nanoparticle (RecBCD–nanoparticle) Translocation by individual RecBCD enzyme molecules can be directly visualized by labeling the protein with a fluorescent nanoparticle, which provides a strong and stable fluorescence signal (Handa et al., 2005). Biotinylated RecBCD was purified from an E. coli strain that expresses RecD with an N-terminal hexahistidine tag, followed by an amino-acid sequence that directs the biotinylation in vivo of a single lysine residue (Handa et al., 2005; Schatz, 1993). To attach the fluorescent nanoparticle: 1. Mix 4.8 ml of the biotinylated RecBCD enzyme (1.22 mM in storage buffer: 20 mM Tris–HCl (pH 7.5), 0.1 mM EDTA, 0.1 mM DTT, 100 mM NaCl, and 50% (v/v) glycerol) with 3 ml of a streptavidincoated fluorescent nanoparticle (0.5% solids in 50 mM sodium phosphate (pH 7.5), 50 mM NaCl, and 5 mM sodium azide; 40 nm TransFluoSpheres; excitation 488 nm; emission 645 nm; Molecular Probes, Carlsbad, CA). 2. Incubate for 10 min at 37  C. The RecBCD–nanoparticle is subsequently bound to the DNA–bead complex (see below, Section 5.2). 3.2. Rad54/Tid1 labeled with a fluorescent antibody (FITC–Rad54/Tid1) As an alternative to biotinylation, proteins can be prepared as fusion products; the choice of using a biotinylation tag versus a fusion protein depends on a number of empirical factors, including the efficiency of biotinylation in the organism used for protein expression versus the expression, solubility, and activity of the modified protein. Rather than attaching a streptavidincoated nanoparticle to the biotin, a fluorescent antibody can be used to label the fusion protein. Yeast Rad54 and Tid1 proteins are purified as a GST fusion product. Consequently, one can directly visualize the translocation of Rad54 or Tid1 by binding a fluorescent antibody to the GST moiety of Rad54 or Tid1 (Amitani et al., 2006; Nimonkar et al., 2007). 1. Prepare DNA–bead complexes as described in Section 2.2. 2. Add DNA translocase to a final concentration of 10 nM. 3. Add 670 nM FITC-anti-GST antibody (an average degree of labeling of six fluorophores/antibody; RGST-45F-Z, Immunology Consultants Laboratory, Newberg, OR) in PBS containing 0.2% (w/v) BSA. The final volume is  5 ml. Watching Individual Proteins Acting on Single Molecules of DNA 269 4. Incubate the mixture at room temperature for 10 min. 5. Immediately dilute the complex into 400 l of degassed buffer containing 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose. The final bead concentration is 90 fM. 3.3. Chemically modified fluorescent RecA or Rad51 proteins (RecAFAM/Rad51FAM) RecA and Rad51 can be imaged by the covalent addition of a fluorescent adduct to the N-terminus of the protein (Galletto et al., 2006; Hilario et al., 2009). Chemical modification is performed by coupling 5(6)-carboxyfluorescein succinimidyl ester (FAM-SE, Invitrogen) to the N-terminal amine. Because of the difference in pKa between N-terminal a-amino group (pKa7) and the e-amino group of lysine (pKa  10–11), by a judicious adjustment of dye concentration and incubation time, the N-terminal amine of the protein can be relatively specifically labeled ( 103-fold over other primary amino groups) at near neutral pH to yield products with typically 1 dye/protein monomer. Here, we describe the protocol to label RecA. The reaction conditions need to be optimized for the protein of interest. 1. The protein is first dialyzed extensively against a solution lacking primary amines (50 mM K2HPO4/KH2PO4 (pH 7.0), 1 M NaCl, 0.1 mM DTT, and 10% glycerol). 2. Dissolve the FAM-SE in dry DMSO to a stock concentration of  50– 75 mM. The precise concentration of the stock is determined spectroscopically by making a 1:10,000 dilution into 10 mM Tris–HCl (pH 9.0) and using an extinction coefficient of 7.8  104 M 1 cm 1 at 492 nm. 3. Add a 12-fold molar excess of FAM-SE to typically 80–100 mM RecA (500 ml) and incubate at 4  C for 4 h in the dark. 4. Stop the reaction by adding Tris–HCl (pH 7.5) to a final concentration of 50 mM. 5. Remove unreacted fluorescein (FAM) by using a Bio-Gel P10 (Bio-Rad) column (1 cm  16 cm). 6. Dialyze the sample against storage buffer (20 mM Tris–HCl (pH7.5), 0.1 mM EDTA, 0.5 mM DTT, 10% (v/v) glycerol). 7. Determine the RecA and FAM concentrations by measuring the absorption at 280 and 492 nm using the extinction coefficients of e280 ¼ 2.7  104 M 1 cm 1 for RecA and e492 ¼ 7.8  104 M 1 cm 1 for fluorescein measured at pH 9. 8. Determine the degree of labeling by calculating the ratio of FAM and RecA concentrations. A correction factor (CF ¼ A280/A492) of 0.32 measured for the free dye in the absence of protein is used to account for the absorption of FAM at 280 nm using the following calculation: 270 Ichiro Amitani et al. ARecA ¼ A280 – CF  A492. The correction factor may be sensitive to buffer conditions, specifically pH. It is recommended that the CF provided by the manufacturer is verified experimentally. The chemical modification of human Rad51 (at 50–60 M) is performed in buffer containing 50 mM KH2PO4 (pH 7.1), 200 mM KCl, 0.1 mM DTT, and 25% (v/v) glycerol, using 20-fold molar excess of FAM, reacted for 6 h at 4  C. The labeled protein is stored in storage buffer (50 mM Tris–HOAc (pH 7.5), 200 mM KCl, 1 mM DTT, 0.1 mM EDTA, 50% glycerol) (Hilario et al., 2009). 4. Instrument 4.1. Flow cell design Single-molecule reactions are carried out in multi-channel flow cells. The photograph of a three-channel flow cell is shown in Fig. 13.1A. Flow cells are designed to ensure laminar flow and to minimize mixing of solutions from different channels (Figs. 13.1 and 13.2). Channel dividers are 100 mm in width with an approximately semicircular end of 10–100 mm, depending on the manufacturing process. Each flow channel is 1.5 mm in width and 70 m in depth. Fluid flow in this geometry is at a very low Reynolds number (<1) and laminar. The velocity field is a Poiseuille flow with a parabolic profile; maximum velocity is midway between the top and bottom of the flow cell, and zero velocity at the top and bottom surfaces. To ensure that experiments are conducted in regions where diffusion from adjacent channels is minimal, trapping must occur at a position that is downstream of the confluence and away from the boundary between flow channels. The mean diffusion distance, x, of a solute can be calculated as hx2 i ¼ 2Dt ð13:1Þ kB T 6pa ð13:2Þ D¼ Here, D is the diffusion rate of the solute, kB is the Boltzmann constant, T is the absolute temperature,  is the viscosity of the solution, and a is the radius of the diffusing particle. The mean diffusion distance provides a good approximation of the boundary where little diffusion between the channels has occurred. However, the absolute concentration will contain solute or ligand from adjacent channels some distance past the mean diffusion boundary. Fig. 13.2 shows a simulation of a solute concentration for two typical solutes (Mg2þ and ATP) in our flow cells under nominal conditions. The 271 Watching Individual Proteins Acting on Single Molecules of DNA Mg2+ I -200 Concentration Distance across (mm) 100 100% -400 0 200 Concentration (%) A II 400 100 200 300 400 80 60 40 20 0 -400 0% Distance along flow (mm) B ATP 0 200 II 400 100 200 300 400 Distance along flow (mm) 0% Concentration (%) -200 -200 0 200 Distance across (mm) 400 100 100% I Concentration Distance across (mm) -400 50 mm 150 mm 250 mm 350 mm 450 mm 80 60 40 50 mm 150 mm 250 mm 350 mm 450 mm 20 0 -400 -200 0 200 Distance across (mm) 400 Figure 13.2 The calculated concentrations of Mg2þ, D ¼ 10 5 cm2/s (A, left panel), and ATP, D ¼ 10 6 cm2/s (B, left panel) as a function of position for diffusion from channel I into channel II. The flow cell has the same dimensions as described in Fig. 13.1, and the flow rate, v, is 50 mm/s from left to right; the end of the divider is at the origin of the plot. The cross section of concentration as a function of the distance downstream of the flow cell is also shown (right panels). The solid white line in the left panels indicates the mean diffusion distance of the solute from and into each channel. The calculations were performed in MATLAB (MathWorks). simulations are based on the exact solution of Fick’s equation for one-dimensional diffusion in a pipe. Typically, a flow velocity of 100–200 m/s is used (Fig. 13.1B). Experiments are conducted by optically trapping 200 m downstream of the dividers into the flow cell, halfway between the top and bottom surfaces ( 35 mm), and halfway into the channel ( 750 m). This position ensures that experiments are conducted in a region where the local solution concentration is identical to the bulk concentration within a channel. 4.2. Flow cell fabrication Several methods have been used to make multichannel flow cells (for a review, see Brewer and Bianco, 2008). MMR Technologies employs a dry etching technique to create channels on a glass slide; the coverslip is attached to the slide by melting powdered glass at 660  C (MMR Technologies, 272 Ichiro Amitani et al. Mountain View, CA). Another way to construct flow cells is to use chemically amplified, negative-tone, epoxy-based photoresists. A multichannel flow cell  70 mm deep can be made using the following process (Fig. 13.3). 1. Prepare a mask on a chrome borosilicate photomask using standard lithography techniques. 2. Drill inlet and outlet holes on a glass coverslip or slide for either an upright or an inverted microscope using diamond grinding bit (model 750, Dremel, WI). The glass should be submerged under water during grinding. 3. Clean the coverslip and slide with hot piranha treatment (96% H2SO4:30% H2O2 ¼ 3:1 (v/v)). 4. Spin coat KMPR 1050 photoresist (MicroChem Corp., Newton, MA) onto the slide at 2000 rpm for 30 s. 5. Soft bake at 100  C for 20 min. 6. Expose the coated slide with 365 nm light using the prepared mask on a Karl-Suss MA4 Mask Aligner (Karl Suss America, Inc., Waterbury Center, VT). 7. Bake at 100  C for 4 min. 8. Remove unexposed photoresist in SU-8 developer (MicroChem Corp.) for 4 min with slow shaking. Slide Coverslip Spin-coat photoresist Exposure KMPR 1050 Develop KMPR 1050 Flip and assemble Expose KMPR 1005 Develop KMPR 1005 Figure 13.3 Flow diagram for the microfabrication of a three-channel flow cell (see text for details). Watching Individual Proteins Acting on Single Molecules of DNA 273 9. Spin coat KMPR 1005 photoresist (MicroChem Corp.) onto the coverslip at 1600 rpm for 30 s. 10. Put both the coverslip and the slide on a hotplate at 60  C with the KMPR-coated side facing up. 11. Flip the coverslip onto the slide. Carefully align the coverslip and the slide before dropping the coverslip onto the slide. 12. Increase the hotplate temperature to 90  C. When the temperature reaches 90  C, gently touch the coverslip so that the coverslip and the slide bond together, then decrease the temperature to 60  C. 13. Expose the coverslip and slide assembly to 365 nm light using the mask prepared in step 1 on a Karl-Suss MA4 Mask Aligner. 14. Bake at 100  C on a hotplate for 1 min. 15. Remove unexposed photoresist in SU-8 developer by running the developer through the channels with vacuum. 16. Attach machined connectors (P-770-01, Upchurch Scientific, Oak Harbor, WA) to the coverslip at the holes using epoxy. An alternative way to make flow cells is to use thermobond film or Parafilm: 1. Drill inlet and outlet holes (1 mm diameter) on the coverslip or slide, depending on the type of microscope (upright or inverted) used. 2. Place either thermobond film (Thermobond film 668EG, 2.5 mil (62 m), 3 M, St. Paul, MN) or Parafilm on the slide. Cut the desired pattern using a razor blade. 3. Place a coverslip on top of the spacer. 4. Place the assembly on a heat block at 150  C for 20–30 s. Gently press the coverslip so that the slide and the coverslip bond evenly. 5. Using either a handheld grinding tool or a razor blade, create a V-shape at one end of a short piece (1–2 cm) of PEEK tubing (Upchurch Scientific). 6. Attach the V-shaped end of the PEEK tubing into the holes and glue it using epoxy. Care must be taken not to block the channels. 4.3. Microscope with laser trap and microfluidic system The laser-trap systems are constructed around a Nikon Eclipse E400 or a Nikon TE2000U microscope (Nikon, Tokyo, Japan). Schematics of the instruments are shown in Figs. 13.4 and 13.5. A brief description of the components follows. 4.3.1. Single optical trap imaging system A high-pressure mercury lamp (USHIO America, Inc., Cypress, CA) and Y-FL 4-cube Epi-Fluorescence (Nikon) attachment are used for illumination. Images are captured using a high-sensitivity electron bombardment 274 Ichiro Amitani et al. Monitor Camera Lamp Camera controller Syringe+ pump DM2 L2 S L1 Beam expander IR laser DM1 OBJ Stage PC Figure 13.4 Schematic diagram of the microscope, optical trap, and flow cell. The trapping IR laser initially passes through a 20 beam expander, and is then further collimated and steered by lenses L1 and L2 an electronic shutter (S) is in-between. A high-pass IR dichroic mirror (DM1) directs laser beam into the objective (OBJ). The flow cell is mounted on an x–y translocation stage that is controlled by a computer (PC) solutions are delivered to the flow cell using a multisyringe pump. A high-pressure mercury arc lamp is used for illumination (fluorescence and bright field). A second dichroic mirror (DM2) is used to image the fluorescent protein–DNA–bead complex onto an electron bombardment camera; the real-time image is displayed on a monitor. CCD camera (EB-CCD C7190, Hamamatsu Photonics, Hamamatsu City, Japan), recorded on video tape, and subsequently digitalized using an LG-3 frame grabber at 30 frame/s (Scion Corporation, Frederick, MD). The optical trap is created by focusing a 1064 nm laser (Nd:YVO4, 6 W max, J-series power supply, Spectra Physics, Mountain View, CA) through a high numerical aperture (NA) objective (100/1.3 oil DICH, Nikon). A high NA objective is necessary to create an intensity gradient sufficiently large to form the trap (Neuman and Block, 2004). The laser is expanded with a 20 beam expander (HB-20XAR.33, Newport, Irvine, CA) to fill the back aperture of the objective. The laser is collimated and aligned using two lenses with the same focal length forming a 1 telescope. The laser is reflected along the optical axis of the microscope by means of a low-pass dichroic mirror (DM) placed between the objective and the fluorescence cube. Experiments are carried out in a multichannel microfluidic flow cell held on a computer controlled motorized stage (MS-2000, Applied Scientific Instruments, Eugene, OR) mounted on the microscope. The solutions are introduced into the flow cell by a syringe pump with multiple syringes 275 Watching Individual Proteins Acting on Single Molecules of DNA Xe lamp BS L9 M PC M Stage L5 L4 L3 AOM L8 L7 L6 Objective lens Camera M BS M HWP M ck ba L1 Fe QPD L11 HM L10 L2 ed DM Amplifier+ low-pass filter IR laser PC Figure 13.5 Schematic diagram for a dual laser-trap microscope. Lenses L1 and L2 initially collimate and expand the laser. The first beam path (black line) passes through an AOM which is imaged on the back aperture of the objective lens by lenses L3, L4, L5, and L9. The second beam path (gray line) is reflected off a movable mirror which is imaged onto the back aperture of the objective by lenses L6, L7, L8, and L9. The image from the objective (dashed line) is split between a camera that images the fluorescent protein–DNA–bead complex, and a quadrant photodiode (QPD) for position detection of the bead in the first trap by means of a half-mirror (HM). The signal from the QPD passes through an amplifier and a low-pass filter before being processed by a PC which uses the information to control the AOM, thus providing feedback control on the position of the first trap with nm resolution. Mirrors (M) and beam splitters (BS) serve to direct the beam path. A dichroic mirror (DM) is used to direct the trapping lasers into the objective and to pass light from the Xenon lamp to the camera and QPD; the realtime image is displayed on a monitor. Lenses L10 and L11 image the trapped bead onto the camera and QPD. (KD Scientific, Hollston, MA). PEEK tubing (Upchurch Scientific) is used to connect the syringes to the flow cell. The microfluidic system permits the imaging of protein–DNA complexes on a single molecule of flow-stretched DNA; it also enables the rapid movement of the sample to the different buffers in the channels of the flow cell. The position of the stage and hence the flow cell, is controlled using a custom-built program. Because the translation speed of the motorized sample stage is typically 0.5–1 mm/s, and the distance being moved to the adjacent flow channel is 0.7–1.5 mm, the time required to move between solution channels is 1–2 s. 4.3.2. Dual optical trap imaging system For force measurements, a double laser-trap system is constructed around a TE2000U microscope (Fig. 13.5). An infrared laser (Nd:YVO4, 6 W max, J-series power supply, Spectra Physics) beam is passed through lenses L1 and 276 Ichiro Amitani et al. L2 ( f ¼ 6, 12 mm, respectively) serving as a collimator. The beam is then passed through a half-wave plate (HWP) and a polarizing beam splitter (BS) creating two separate beam paths. The first beam is steered with an acoustic optical modulator (AOM) for force feedback. The first beam path is then magnified with lenses L3, L4, L5, and L9 ( f ¼ 75, 25, 25, 800 mm, respectively) to fill the back aperture of the microscope objective. Similarly, the second path is magnified with lenses L6, L7, L8, and L9 ( f ¼ 75, 25, 25, 800 mm, respectively) to fill the back aperture of the microscope objective. The laser-trap strength can be up to 0.4 pN/nm, but heating at high-power settings can be a problem. A dichroic mirror (DM) is used to direct the trapping lasers into the objective and to pass light from the Xenon lamp to the camera and quadrant photodiode (QPD). The florescent image of the trapped protein–DNA–bead complex is focused onto a CCD camera (iXonþ, Andor) via the objective through another BS. One of the trapped beads is also imaged onto the QPD (S1557-03, Hamamatsu) to provide precise high-bandwidth information about its position. Signals are digitized (PCI6052E, National Instruments) and processed with software written in LabView (LabView 6.1, National Instruments). The position data from the QPD controls the deflection angle of the AOM allowing for feedback between the bead position and the trap position. This arrangement permits the movement of one optical trap relative to the other. The total moveable range of the AOM is 2.4 m, but the linear range is limited to 200 nm. 4.4. Temperature determination and control To achieve reliable trapping in a flow field, laser power in the range of several hundred milliwatts (mW) is used. Water has a measurable absorption at the near-infrared wavelength typically used for an optical trap (l ¼ 1064 nm). Consequently, the effect of local heating on sample temperature is an important consideration. Different ways of estimating the temperature in an optical trap have been reported (Celliers and Conia, 2000; Liu et al., 1995; Peterman et al., 2003). We adapted the methods using fluorescence, and we measured temperature based on the thermal quenching of rhodamine B (RhB) fluorescence (Karstens and Kobs, 1980; Romano et al., 1989). The temperature measurement is carried out using a customized sample chamber (Fig. 13.6). This chamber is constructed as described in Section 4.3. A thermocouple (model CHAL-002, Omega Engineering, Stamford, CT) is placed in the middle of the channel before assembling the coverslip and slide. 4.4.1. Temperature determination The following procedure is used to measure temperature based on the fluorescence intensity measurements of RhB relative to Alexa-488. 277 Watching Individual Proteins Acting on Single Molecules of DNA Thermocouple Slide Parafilm Coverslip Thermistor Brass jacket Water circulation Objective Copper tubing Figure 13.6 Schematic illustration of the components used for temperature measurement and control. Top: the flow cell used for temperature determination is made of a glass slide, a coverslip, and a layer of Parafilm sandwiched in between. A thermocouple is placed inside the channel. Bottom: an illustration of the temperature controller (side view; not to scale). 1. Construct a flow cell as described in Figure 13.6, Section 4.3. Place a thermocouple (model CHAL-002, Omega Engineering, Stamford, CT) in the middle of the channel before assembling the coverslip and slide. 2. Fill the flow cell with 10 dye solution (10 M RhB (Wako Pure Chemical Industries, Ltd.) and 15 M Alexa-488 (Invitrogen) in TE buffer) and incubate at 40  C overnight to coat the flow cell. 3. Replace the 10 dye solution with degassed 1 dye solution. Seal the outlets using Parafilm. 4. Set the flow cell temperature using either a thermoelectric microscope slide temperature controller (BC-100, 20/20 technologies, Wilmington, NC) or an objective jacket (see next Section 4.4.3). Wait for at least 30 min for the system to equilibrate. 5. Select a region (5 mm  10 m) around the trapping position and record the fluorescence images of RhB and Alexa-488 with the appropriate filter sets (Ethidium Bromide set 41006 and Blue set 11001v2; Chroma Technology Corp., Rockingham, VT). 6. Repeat steps 4 and 5 for at least four different temperatures. These measurements are used to relate the fluorescence intensity to temperature (see below). 7. Turn on the IR laser, set the desired power, and wait for about 30 min. 8. Select the same region as in step 4. Record the fluorescence images of RhB and Alexa-488. 9. Change the IR laser power and repeat steps 7 and 8. 10. Replace the dye solution with TE buffer. Record the background fluorescence images of RhB and Alexa-488. 278 Ichiro Amitani et al. 11. Remove the flow cell and the objective lens. Measure the laser power at the back aperture of the objective. Calculate the power delivered to the focus using the infrared transmission coefficient of the objective used (60% for the objective used) (Neuman and Block, 2004). To determine the average temperature around the focus, the fluorescence images are analyzed with ImageJ (NIH; http://rsb.info.nih.gov/ij/). A calibration curve of relative fluorescent intensity versus temperature is generated for the data acquired in the absence of trapping laser power. The temperature at any given laser power setting is then obtained from the calibration curve: 1. Calculate the relative fluorescence intensity, Ir(T), at each temperature, T, measured above in the absence of IR irradiation using Ir ðTÞ ¼ ðIR IR;BG Þ=ðIA IA;BG Þ ð13:3Þ where IR and IA are the average fluorescence intensity of RhB and Alexa-488 at temperature T, respectively; IR,BG and IA,BG are the average background fluorescence intensities using RhB and Alexa-488 filter sets, respectively. 2. Normalize the relative intensity Ir(T) to an arbitrary reference temperature, T0 (e.g., 25  C), using rðT Þ ¼ Ir ðT Þ=Ir ðT0 Þ ð13:4Þ where r(T) is the normalized intensity ratio. 3. Determine the constant, a, in the empirical linear relationship (Kato et al., 1999) (Fig. 13.7A): rðT Þ ¼ 1 T0 Þ aðT ð13:5Þ 4. The temperature at any given IR laser power is given by T¼ r 1 a ð13:6Þ þ T0 where r is the observed intensity ratio normalized to the reference temperature at that given IR laser power. 5. The temperature change for any given IR laser power is DT ¼ T T0 ¼ r 1 a ð13:7Þ For our instrument, the IR laser-induced temperature change is about 1.2  C/100 mW of laser power delivered at the focus (Fig. 13.7B). 279 Watching Individual Proteins Acting on Single Molecules of DNA Normalized fluorescence intensity ratio, r(T) A a = -0.021 ± 0.001 per ⬚C 1.0 0.9 0.8 25 30 Temperature (⬚C) 35 B Temperature increase (⬚C) 20 ΔT = 1.20 ± 0.02 ⬚C/100 mW 15 10 5 0 0 500 1000 Laser power (mW) 1500 Figure 13.7 Measurement of IR laser-induced temperature changes. (A) Relative fluorescence as a function of temperature. The ratio of the background-corrected fluorescence of RhB and Alexa-488 is normalized to that at 25  C. The relative fluorescence intensity decreases 2% per  C increase. (B) Relationship between induced temperature change and the IR laser power delivered at the focus, at a starting temperature of 25  C. 4.4.2. Temperature gradient around the trap center In an optical trap, hundreds of mW of laser power are focused on a micrometer-sized spot. The temperature gradient caused by such localized heating is another concern. To experimentally determine this gradient, the fluorescence images of RhB and Alexa-488 are recorded as described in Section 4.4.1. The images are then analyzed using the same procedure as in Section 4.4.1 except that the images are analyzed on a pixel-by-pixel basis, using the intensity measured at each pixel, instead of the average fluorescence intensity of the region. The constant a (Eq. (13.5)) for each pixel is then calculated from the normalized intensity ratios r(T) (Kato et al., 1999; Romano et al., 1989). The temperature distribution in a selected region of our instrument is shown in Fig. 13.8A. The temperature gradient in the absence of flow is about 0.06  C/mm (Fig. 13.8B). 280 Ichiro Amitani et al. A B 35 36 °C T 31 °C 10 mm Trap position The region used to calculate the temperature gradient shown in Fig. B on the right Temperature (°C) Temperature distribution at an IR power of 587 mW 0.061 ± 0.002 ⬚C/mm 34 33 32 -5 0 5 10 15 Distance from the trap center (mm) 20 Figure 13.8 Thermal gradient in a flow cell due to infrared heating by an optical trap. (A) Example showing the measured temperature distribution in an optical trap with an IR laser power of 587 mW when the temperature is set at a starting temperature of 24  C using only the microscope slide temperature controller. The circle indicates the position of the trap. The rectangle indicates the region where the temperature shown in B is measured. (B) Plot of the temperature distribution in the horizontal direction of the region shown in (A). Black: measured temperature; gray: linear fitting from 0.5 to 14.5 mm away from the trap center. Linear fitting gives a temperature gradient of 0.06  C/mm. 4.4.3. Temperature control Due to the need to use an oil-immersion lens, we discovered that the temperature of the flow cell is largely determined by the heat transfer from the objective to the sample. Although a thermoelectric microscope slide temperature controller (BC-100, 20/20 Technologies, Wilmington, NC) can be used for temperatures within a few degrees of ambient and for low power IR laser settings, a thermostated objective lens is a more effective regulator of sample temperature (Mao et al., 2005). A water circulation system is used to control the temperature of the oil-immersion objective (Fig. 13.6). The temperature control module consists of a brass jacket that fits onto the objective lens and copper tubing that is soldered for several turns around the jacket. The temperature of the objective is controlled by circulating temperature-controlled water from a water bath (Isotemp Refrigerated Circulator, Model 910, Fisher Scientific, Pittsburgh, PA) through the tubing. Water circulation does not perturb the optical trapping. This temperature control module can control the temperature of the sample from 15 to 45  C and reduces laser-induced heating by 60%. 5. Single-Molecule Imaging of Proteins on DNA A general single-molecule experiment includes the following steps: (1) prepare DNA attached to the polystyrene bead, either with or without a bound protein of interest; (2) introduce appropriate solutions into different Watching Individual Proteins Acting on Single Molecules of DNA 281 channels of the multichannel flow cell; (3) capture a DNA–bead complex in the optical trap; and (4) move it into other channels containing ligands or proteins of interest. 5.1. Unwinding of DNA by a single RecBCD enzyme To visualize DNA unwinding by an individual RecBCD enzyme (Bianco et al., 2001; Spies et al., 2003), a complex of RecBCD enzyme bound to YOYO-1-stained DNA is optically trapped in the absence of ATP; the complex is then moved into the reaction channel, which contains ATP, to initiate DNA unwinding. DNA unwinding and RecBCD enzyme translocation are monitored as a shortening of the DNA length. 1. Prepare a sample buffer containing 45 mM NaHCO3 (pH 8.3), 20% sucrose (w/v), and 50 mM DTT; degas for at least 1 h. 2. Wash all syringes, tubing, and flow cell with 500 ml of 0.5% (v/v) of blocking reagent (B-10710, Molecular Probes, Carlsbad, CA) in sample buffer using a flow rate of 800 ml/h. 3. Prepare DNA–bead complexes as described in Section 2.2. 4. Add 500 ml of 20 nM YOYO-1 in sample buffer to the DNA–bead reaction. 5. Incubate in the dark at room temperature for at least 1 h. 6. Add Mg(OAc)2 and RecBCD to the stained DNA–bead complex at final concentrations of 2 mM and 50 nM; immediately transfer to the sample syringe (first channel). 7. Prepare 500 ml reaction solution containing the sample buffer supplemented with 2 mM Mg(OAc)2 and ATP at various concentrations; load the reaction syringe (second channel). 8. Trap a RecBCD–DNA–bead complex in the first channel. 9. Immediately move the trapped complex to the second channel to start the reaction. The unwinding of dsDNA is manifested by the shorting of the YOYO-1-labeled DNA (Fig. 13.9A and B). 5.2. Direct observation of RecBCD–nanoparticle translocation Another way to visualize translocation by individual RecBCD enzyme molecules is to attach a fluorescent nanoparticle to RecBCD (Handa et al., 2005). 1. Prepare a sample buffer containing 45 mM NaHCO3 (pH 8.3), 20% (w/v) sucrose, and 50 mM DTT; degas for at least 1 h. 2. Wash all syringes, tubing, and flow cell with 500 ml of 0.5% (v/v) of blocking reagent (B-10710, Molecular Probes) in sample buffer using a flow rate of 800 ml/h. 282 Ichiro Amitani et al. A B RecBCD DNA length (bp) 60,000 50,000 1300 ± 4 bp/s 40,000 30,000 20,000 10,000 0 0 0⬙ 10 20 Time (s) 40⬙ 30 40 RecBCD 0⬙ 35⬙ Nanoparticle position (bp) D C 60,000 50,000 1199 ± 5 bp/s 40,000 30,000 20,000 10,000 0 0 10 20 Time (s) 30 Figure 13.9 RecBCD translocating through, and unwinding, an individual l DNA molecule. (A) Kymograph showing a YOYO-1 stained l dsDNA molecule being unwound by a RecBCD molecule bound to the free DNA end. The drawing to the left of the kymograph depicts the optically trapped bead–YOYO-1-DNA–RecBCD complex. (B) Plot of DNA length versus time. Black line shows the fit to a straight line. (C) Kymograph showing translocation by a fluorescent nanoparticle-labeled RecBCD molecule on l dsDNA. The drawing on the left side of the kymograph depicts the optically trapped bead–DNA–RecBCD–nanoparticle complex. (D) Plot of the position of the RecBCD molecule, indicated by the nanoparticle, versus time. The black line shows the fit to a straight line. Note that the difference in unwinding rates in (B) and (D) is not due to a difference in the techniques, but rather reflects the intrinsic heterogeneity of individual RecBCD enzyme behavior. 3. Prepare DNA–bead complexes as described in Section 2.2. 4. Label biotinylated RecBCD using a fluorescent nanoparticle as described in Section 3.1. 5. Add the DNA–bead complex and 2 mM Mg(OAc)2 to the biotinylated RecBCD–nanoparticle complex, and incubate the resulting mixture for 2 min. 6. Dilute the nanoparticle–RecBCD–DNA–bead complex with 400 ml of degassed sample buffer supplemented with 2 mM Mg(OAc)2 and 0.5% (v/v) blocking solution (B-10710, Molecular Probes); transfer to the sample syringe (first channel). 283 Watching Individual Proteins Acting on Single Molecules of DNA 7. Load the reaction syringe with the reaction buffer consisting of 1 mM ATP, 2 mM Mg(OAc)2, and 0.5% (v/v) blocking solution in sample buffer (second channel). 8. Trap a nanoparticle–RecBCD–DNA–bead complex in the first channel. 9. Immediately move the trapped complex to the second channel. The fluorescent particle is seen to move toward the trapped bead as RecBCD translocates on the DNA (Fig. 13.9C and D). 5.3. Rad54/Tid1 translocation To observe Rad54/Tid1 translocation (Amitani et al., 2006; Nimonkar et al., 2007), a two-channel flow cell is used. Figure 13.10A shows a schematic illustration of the translocation assay, a kymograph showing A FITC-Rad54 Downstream l DNA Optical trap Upstream Flow B Start End Bead 0⬙ Rad54 position from bead (bp) C 370⬙ 50,000 40,000 30,000 20,000 10,000 0 0 100 200 300 400 Time (s) Figure 13.10 Rad54 translocating on a single dsDNA molecule. (A) Schematic illustration of the optically trapped l DNA–bead complex with a bound FITC–Rad54 complex. (B) Kymographs depicting upstream translocation (in the direction opposite to flow) of Rad54 on the dsDNA. (C) Plot of FITC–Rad54 position relative to the bead versus time. 284 Ichiro Amitani et al. translocation by Rad54 labeled with FITC-antibody (Fig. 13.10B), and a graph of FITC–Rad54 position as a function of time (Fig. 13.10C). 1. To reduce nonspecific binding, the syringe, tubing, and flow cell are incubated with 0.5 mg/ml of BSA in 50 mM Tris–HOAc (pH 7.5) at room temperature for 15 min, then rinsed with a 10-fold volume of 50 mM Tris–HOAc (pH 7.5). 2. Prepare FITC–Rad54/Tid1–DNA–bead complexes as described in Section 3.2. 3. Load 400 ml of the FITC–Rad51/Tid1–DNA–bead complex (typically 90 fM) in 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose into the first channel. 4. Load 400 ml of solution containing 1 mM ATP, 2 mM Mg(OAc)2, 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose into the second channel. 5. Trap a FITC–Rad54/Tid1–DNA–bead complex in the optical trap in the first channel. 6. Move the complex to the second channel containing the ATP to initiate translocation. The fluorescently tagged Rad54 is seen to translocate toward the trapped bead (upstream) (Fig. 13.10B and C) or away from the trapped bead (downstream). 5.4. Real-time Rad51 assembly To detect the assembly of Rad51 in real time, a two-channel flow cell is used (Hilario et al., 2009). Rad51 assembly can be measured by monitoring the increase in the length of fluorescently end-labeled DNA. A kymograph is shown in Fig. 13.11A; Fig. 13.11B shows a graph of DNA length versus time. 1. To reduce nonspecific binding, the syringe, tubing, and flow cell are washed at 800 ml/h with 0.5 mg/ml of BSA and 0.5 mg/ml casein in 50 mM Tris–HOAc (pH 7.5) at room temperature for 1 h, followed by a rinse with sample buffer for 1 h. 2. Prepare Cy3–DNA–bead complexes as described in Section 2.3. 3. Load 400 ml of solution containing Cy3–DNA–bead complexes (about 90 fM), 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose into the first channel. 4. Load 400 ml of solution containing Rad51, 10 mM Mg(OAc)2, 2 mM ATP, 40 mM Tris–HOAc (pH 7.5), 30 mM DTT, and 15% (w/v) sucrose into the second channel. The Rad51 concentration can be varied from 50 nM to 1 mM. 5. Trap a Cy3–DNA–bead complex in the first channel. 6. Move the complex to the second channel to initiate Rad51 assembly. 285 Watching Individual Proteins Acting on Single Molecules of DNA A B C 20 15 10 0 Rad51 nucleoprotein filament disassembly 0⬙ 25 50⬙ 60⬙ 10 D DNA extension (mm) 0⬙ DNA extension (mm) Rad51 nucleoprotein filament assembly 30 20 Time (s) 40 50 25 20 15 10 0 20 40 60 Time (s) 80 Figure 13.11 Rad51 assembling onto, and dissociating from, a single dsDNA molecule. (A) Kymograph of Rad51 assembly on Cy3-end-labeled l DNA. The schematic on the left side of kymograph depicts the optically trapped bead, initial position of Cy3end-label of the DNA (star), and DNA (solid line); the schematic on the right side depicts the extended Rad51 nucleoprotein filament. DNA length is measured from the center of the bead to the Cy3-end-label. (B) Plot of DNA length versus time for the assembly of Rad51 on DNA analyzed using two-dimensional Gaussian fitting of the end-label position. (C) Kymograph of Rad51 disassembly from Cy3-end-labeled l DNA. The schematic on the left side of kymograph depicts the optically trapped bead, initial position of Cy3-end-label of the DNA (star), Rad51 (filled circles), and DNA (solid line). (D) Plot of DNA length versus time for the disassembly of Rad51 from DNA. 5.5. Real-time Rad51 disassembly To visualize the disassembly of a Rad51 nucleoprotein filament in real time, a three-channel flow cell is used (Hilario et al., 2009). Rad51 disassembly can be measured by monitoring the decrease in the length of fluorescently end-labeled DNA onto which Rad51 is assembled. A kymograph of Rad51 disassembly is shown in Fig. 13.11C; Fig. 13.11D is the graph of DNA length versus time. 1. The syringe, tubing, and flow cell are washed at 800 ml/h with 0.5 mg/ml of BSA and 0.5 mg/ml casein in 50 mM Tris–HOAc (pH 7.5) at room temperature for 1 h, followed by a rinse with sample buffer for 1 h. 2. Prepare Cy3–DNA–bead complexes as described in Section 2.3. 286 Ichiro Amitani et al. 3. Load 400 ml of solution containing Cy3–DNA–bead complexes (typically about 90 fM), 40 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 15% (w/v) sucrose into the first channel. 4. Load 400 ml of solution containing 40 mM Tris–HOAc (pH 7.5), 10 mM Mg(OAc)2, 30 mM DTT, and 15% (w/v) sucrose into the second channel; the ATP concentration can be varied from 0 to 2 mM. 5. Load 400 ml of solution containing 1 mM Rad51, 40 mM Tris–HOAc (pH 7.5), 10 mM Mg(OAc)2, 2 mM ATP, 30 mM DTT, and 15% (w/v) sucrose into the third channel. 6. Trap a Cy3–DNA–bead complex in the first channel. 7. Move the complex to the third channel to assemble Rad51 onto the DNA. 8. Move the complex to the second channel to initiate Rad51 disassembly from the DNA. 5.6. Visualization of RecAFAM/RecA-RFP/Rad51FAM filament formation To directly visualize nucleation (cluster formation) by fluorescent RecA or Rad51 (Galletto et al., 2006; Handa et al., 2009; Hilario et al., 2009), a threechannel flow cell is used. To confirm that a single molecule of DNA is present, YOYO-1-stained DNA is used in the initial trapping. YOYO-1 is removed by washing the DNA in a buffer containing 5–10 mM Mg(OAc)2 before starting the assay. A time course of RecA-RFP cluster formation is shown in Fig. 13.12. Here, we describe the protocol for imaging RecARFP cluster formation. 1. The syringe, tubing, and flow cell are incubated with 0.5 mg/ml of BSA in 50 mM Tris–HOAc (pH 7.5) at room temperature for 15 min, then rinsed with a 10-fold volume of 50 mM Tris–HOAc (pH 7.5). 2. Prepare YOYO-1-stained DNA–bead complexes as described in Section 2.2. 3. Load 400 ml of solution containing YOYO-1-stained DNA–bead complex, 20 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 20% (w/v) sucrose into the first channel. 4. Load 400 ml of solution containing 0.5 mM ATPgS, 5 mM Mg(OAc)2, 20 mM Tris–HOAc (pH 8.2), 30 mM DTT, and 20% (w/v) sucrose into the second channel. 5. Load 400 ml of solution containing RecA-RFP, 0.5 mM ATPgS, 1 mM Mg(OAc)2, 20 mM MES (pH 6.2), 30 mM DTT, and 20% (w/v) sucrose into the third channel. The RecA-RFP concentration is varied from 150 to 400 nM. 6. Trap a YOYO-1-stained DNA–bead complex in the first channel. 287 Watching Individual Proteins Acting on Single Molecules of DNA A RecA-RFP cluster Bead l DNA B 110 nM RecA-RFP 140 nM RecA-RFP 170 nM RecA-RFP 200 nM RecA-RFP 10 s 10 s 10 s 5s 20 s 20 s 20 s 10 s 30 s 30 s 30 s 15 s 40 s 40 s 40 s 20 s Bead Figure 13.12 Direct visualization of nucleation and growth of RecA-RFP nucleoprotein filaments on individual molecules of dsDNA. (A) Illustration of the nucleation of RecA-RFP clusters on DNA. (B) Representative video frames showing nucleation at four different RecA-RFP concentrations. Flow is left to right. Each vertical strip represents the same DNA molecule repeatedly dipped into the RecA-RFP protein solution for the incubation times indicated. The trapped bead position is indicated by an arrow; the bead is fluorescent due to the nonspecific binding of the RecA-RFP. 7. Move the complex to the second channel to confirm by visual inspection that a single molecule of intact l DNA is attached to the bead. 8. Incubate the complex for 2–3 min in the second channel to dissociate the YOYO-1. In this step, the shutter for excitation light is closed to avoid the photocleavage of DNA. 9. Move the complex to the third channel containing the RecA-RFP. 10. Incubate the complex for 5–30 s in the third channel. In this step, the shutter for excitation light is closed to protect the CCD camera from the strong signal due to a high concentration of fluorescent protein. 11. Move the complex to the second channel to observe the RecA-RFP clusters. To avoid the photobleaching of RecA-RFP, the illumination time should be minimized (1–2 s). 12. Repeat steps 9 and 10. 6. Data Analysis Methods To improve the efficiency of the single-molecule studies and increase the reliability of the data analysis, we developed programs to analyze the fluorescence images. The software is available upon request. 288 Ichiro Amitani et al. 6.1. Two-dimensional Gaussian fitting To determine the center of a fluorescent intensity distribution (e.g., for Cy3-end-labeled DNA or FITC–Rad54/Tid1), the distribution is fitted to the following two-dimensional Gauss function (Hilario et al., 2009; Nimonkar et al., 2007): " # ðx xc Þ2 ðy yc Þ2 f ðx; yÞ ¼ A  exp þB ð13:8Þ s2x s2y Here, f(x, y) is the point spread function (PSF), A the maximum intensity of PSF, x, y the coordinates of image, xc, yc the center of PSF, sx, sy the width of the PSF, and B the background intensity. To perform nonlinear regression, Eq. (13.8) is expanded to a Taylor series, and the zeroth and first derivative terms are used. The fitting process is iterated until the relative change of fitting parameters falls within a predefined threshold (5%; empirically chosen as a threshold because the fluctuation of fluorescent spot position is greater than the fluorescent spot size). To reduce the calculation time, a small region surrounding the fluorescent spot is selected for fitting. The image in the region of interest is median-filtered and averaged before fitting. The initial parameters for fitting are determined automatically from the fluorescent intensity distribution in the region. For the DNA length measurement, the radius of bead is known and subtracted from the observed length. Figs. 13.10C, 13.11B, and D show the results of a two-dimensional Gaussian fitting. 6.2. Automatic DNA length measurement To measure the length of DNA that is either fluorescently labeled with YOYO-1 or decorated with fluorescent proteins, we developed a plug-in for ImageJ to automate the analysis. Fig. 13.9B shows an example of the analysis. 1. To improve the data quality, fluorescence images are first averaged every 5–10 frames. 2. The position of the trapped bead is initially determined manually, taking advantage of the nonspecific binding of dye to the bead. 3. For each frame, a radial line scan originating from the bead center is determined. The orientation of the DNA is determined as the direction that has the maximum mean gray value. 4. The profile along the orientation of the DNA is calculated. 5. The derivative of the profile is calculated. The position of the maximum (excluding the bead) in the derivative is the position of the DNA end. The length of the DNA is calculated as the distance from the DNA end to the bead center, minus the radius of the bead. Watching Individual Proteins Acting on Single Molecules of DNA 289 ACKNOWLEDGMENTS We wish to thank Jason Bell, Aura Carreira, Petr Cejka, Anthony Forget, Joe Hilario, Taeho Kim, Hsu-Yang Lee, Katsumi Morimatsu, Amitabh Nimonkar, Behzad Rad, and Lisa Vancelette for their comments on this manuscript, and members of the Kowalczykowski lab for their contribution to this research. The research in our lab has been funded by grants from National Institutes of Health T32 CA-108459 to C. C. 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