Name: real-time observation of the dna strand exchange reaction mediated by rad51
Uploaded: 2019-02-13T14:00:00.000+00:00
Duration: 6 min 24 s
Description: Watch this Scientific Journal Video about Real-time Observation of the DNA Strand Exchange Reaction Mediated by Rad51 at JoVE.com

This work was funded by Grants-in-Aid for Scientific Research (A) (18H03985) and on Innovative Areas (15H05974) to HI, for Young Scientists (B) (17K15061) to BA, and for Scientific Research (B) (18H02371) to HT from the Japan Society for the Promotion of Science (JSPS).

1. Preparation of Proteins and DNA Substrates
<ol>
	<li>Purify S. pombe Rad51 and Swi5-Sfr1 proteins to homogeneity (judged by Coomassie staining), as previously reported13,21.</li>
	<li>Prepare oligonucleotide DNA substrates listed in Table 118. 
	NOTE: The oligonucleotides were purchased (see Table of Materials) and synthesized at HPLC grade. For the DNA strand pairing reaction, oligonucleotides 16FA(-), 16A(-)_40bp and 16AR(+)_40bp are required. For the DNA displacement assay, oligonucleotides 16A(-), 16FA(-)_40bp and 16AR(+)_40bp are required (Figure 1 and Table 1). All DNA concentrations in this protocol refer to fragment concentrations as opposed to nucleotide concentrations.</li>
	<li>To form donor dsDNA, mix equimolar amounts of complementary strands in a thin-walled PCR tube with annealing buffer (10 mM Tris-HCl pH 7.5, 100 mM NaCl, 10 mM MgCl2), ensuring a total volume greater than 20 &#181;L. Perform this mixing on a prechilled metal rack on ice (between 2 &#176;C and 4 &#176;C). 
	NOTE: The combination of oligonucleotides for the pairing assay are 16A(-)_40bp and 16AR(+)_40bp. For the displacement assay, anneal oligonucleotides 16FA(-)_40bp and 16AR(+)_40bp.</li>
	<li>Heat the annealing mixture at 90 &#176;C for 5 min and cool down over 3 h to 30 &#176;C using a PCR machine. Store the annealed DNA at -20 &#176;C.</li>
</ol>
2. DNA Strand Pairing and Displacement Assays
<ol>
	<li>Perform DNA strand pairing assay.
	<ol>
		<li>Prepare 1.6 mL of reaction buffer A (30 mM HEPES-KOH pH 7.5, 1 mM dithiothreitol [DTT], 15 mM MgCl2, 0.25 mM ATP, 0.1 mg/mL bovine serum albumin [BSA], and 0.0075% polyoxyethylenesorbitan monolaurate) containing 36 nM 16FA(-) in a 2.0 mL micro-centrifuge plastic tube (polypropylene) and pre-incubate it at 37 &#176;C for 5 min.</li>
		<li>To form Rad51-ssDNA filaments, add Rad51 protein to a final concentration of 1.5 &#181;M in the pre-incubated reaction buffer and incubate it at 37 &#176;C for 5 min.</li>
		<li>Add Swi5-Sfr1 protein to the mixture to a final concentration of 0.15 &#181;M and incubate it at 37 &#176;C for a further 5 min.</li>
		<li>Take 1.5 mL of the mixture and transfer it into a 1.0 x 1.0 cm quartz cuvette containing a magnetic stirrer and set the cuvette into a spectrofluorometer. Configure the peltier temperature controller of the spectrophotometer to 37 &#176;C and set the magnetic stirrer to 450 rpm to ensure rapid mixing of the injected sample.</li>
		<li>Start measuring the fluorescence emission of FAM at 525 nm (bandwidth: 20 nm) upon excitation at 493 nm (bandwidth: 1 nm). Collect data every second.</li>
		<li>After starting the measurement for 100 s, inject ROX-labeled donor dsDNA to a final concentration of 36 nM into the mixture using a syringe and measure the change in emission at 1 s intervals for a further 30 min.</li>
	</ol>
	</li>
	<li>Perform DNA strand displacement assay.
	<ol>
		<li>Prepare 1.6 mL of reaction buffer A containing 36 nM 16A(-) in a 2.0 mL micro-centrifuge plastic tube and pre-incubate it at 37 &#176;C for 5 min.</li>
		<li>Form Rad51-ssDNA filaments in the presence of Swi5-Sfr1 at 37 &#176;C, as described in steps 2.1.2. and 2.1.3.</li>
		<li>Take 1.5 mL of the mixture and transfer it into the quartz cuvette containing a magnetic stirrer and set the cuvette into the spectrophotometer, as described in step 2.1.4.</li>
		<li>Start measuring the fluorescence emission and after 100 s, inject FAM- and ROX-labeled donor dsDNA as described in step 2.1.6. Measure the change in fluorescence emission at 1 s intervals for a further 30 min.</li>
	</ol>
	</li>
</ol>
3. Analyzing Experimental Data from the Pairing and Displacement Assays
<ol>
	<li>Estimate maximum FRET efficiency.
	<ol>
		<li>Prepare 16FA(-) annealed with 16AR(+)_40bp and 16FA(-)_40bp annealed with 16AR(+)_40bp using the same procedure described in steps 1.3 and 1.4.</li>
		<li>Prepare 130 &#956;L of reaction buffer A containing 36 nM of either 16FA(-), 16FA(-) annealed with 16AR(+)_40bp, 16FA(-)_40bp annealed with 16AR(+)_40bp or 16FA(-)_40bp in a 0.2 x 1.0 cm quartz cuvette.</li>
		<li>Set the cuvette into the spectrofluorometer and incubate at 37 &#176;C for 5 min.</li>
		<li>Measure fluorescence spectra from 500 to 600 nm upon excitation at 493 nm.</li>
		<li>To test the effect of Rad51 on the emission of FAM and quenching of FAM by ROX, add Rad51 to a final concentration of 1.5 &#956;M to the mixture and incubate at 37 &#176;C for 5 min.</li>
		<li>Measure fluorescence spectra from 500 to 600 nm upon excitation at 493 nm.</li>
		<li>Calculate maximum FRET efficiency (Emaximum) using the equation described below: 
		Emaximum = (Fluorescence intensity at 525 nm of dsDNA labeled with both FAM and ROX) / (Fluorescence intensity at 525 nm of FAM labeled ssDNA)</li>
	</ol>
	</li>
	<li>Analyze experimental data from the displacement assay.
	<ol>
		<li>To convert the change in fluorescence observed in the displacement assay to the change in the amount of final product, normalize raw experimental data obtained from this assay using the equation described below, where Fraw is fluorescence intensity from raw data and Fnormalized is the change in fluorescence calculated by the following equation. 
		Fnormalized = ([Fraw at time x] - [Fraw at time 0]) / (([Fraw at time 0] / Emaximum) - [Fraw at time 0]) 
		&#8203;Fraw at time 0 is the average fluorescence monitored for the first 5 s after the dead time (i.e., the time required for mixing after an injectant is introduced into the cuvette).</li>
		<li>To exclude the effects of photobleaching and spontaneous displacement, subtract Fnormalized without protein from Fnormalized of the sample to obtain FD, which is the change in the amount of final product in this assay. 
		FD = [Fnormalized of sample] - [Fnormalized without protein]</li>
	</ol>
	</li>
	<li>Analyze experimental data from the pairing assay.
	<ol>
		<li>Normalize raw experimental data obtained from the pairing assay using the equation described below, where Fraw is fluorescence intensity from raw data and Fnormalized is the change in fluorescence calculated by the following equation. 
		Fnormalized = (Fraw at time x) / (Fraw at time 0) 
		&#8203;Fraw at time 0 is the average fluorescence monitored for the last 20 s before initiating the reaction by injecting the dsDNA substrate.</li>
		<li>To convert change in fluorescence into change in amount of substrate and exclude the effects of photobleaching and spontaneous pairing, normalize Fnormalized of sample using the equation described below, where FP is the change in amount of substrate in this assay. 
		&#8203;FP = 1 - (([Fnormalized without protein] - [Fnormalized of sample]) / [1-Emaximum])</li>
		<li>To examine the reaction kinetics of DNA strand exchange, perform non-linear last-square regression analysis of the pairing reaction using the analysis program23 (see Table of Materials).
		<ol>
			<li>Prepare a file in .txt format containing time course data of FP.</li>
			<li>Start the program and paste the script in the Supplemental Code File into a window of the program:</li>
			<li>Start non-linear last-square regression analysis. The results of this analysis will be displayed in the same window.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here, we have described a detailed protocol that utilizes FRET to measure Rad51-driven DNA strand exchange in real time. Importantly, these measurements allow for the determination of reaction kinetics. While the descriptions provided above are sufficient to reproduce our published results, there are several critical points that will be described in this section. Furthermore, the advantages and disadvantages of FRET-based methodologies for studying DNA strand exchange will be discussed, along with the application of such techniques to study other aspects of DNA metabolism.
As with all biochemical reconstitutions, ensuring that all reaction substrates are of high purity is essential. It is negligent to assume the absence of contaminating activities based solely on the apparent purity of a protein preparation judged by Coomassie staining. In particular, the presence of trace amounts of nucleases or helicases can drastically affect the results of the pairing and displacement assays. Thus, we strongly encourage testing for such activities each time a new batch of protein is purified. Furthermore, it is prudent to check the purity of synthesized DNA substrates by native polyacrylamide gel electrophoresis. Despite many companies guaranteeing the purity of oligonucleotides, we have often found through our own testing that purity of the synthesized DNA can vary between batches.
It is important to consider the following two points when conducting experiments with quartz cuvettes. Firstly, some proteins are prone to bind quartz cuvettes nonspecifically. To counter this, BSA and polyoxyethylenesorbitan monolaurate are included in the reaction buffers. Secondly, temperature has a drastic effect on the reaction speed and fluorescence intensity. To minimize this effect, the quartz cuvette should be pre-incubated at 37 &#176;C prior to use.
Although conventional biochemical assays have been hugely useful in studying DNA strand exchange, they have several drawbacks. In a typical time-course experiment, a reaction is incubated at a certain temperature and aliquots are withdrawn at desired timepoints and deproteinized by treatment with detergent and protease to terminate the reaction. Upon completion of the time-course, samples are then subjected to electrophoresis to separate DNA substrates from products. The major advantage of the method described here is that it allows for the real-time observation of the reaction without any disturbance. Any timepoint during the reaction can be inspected without disruption to the reaction itself and there is no need to deproteinize samples or subject them to the potentially disruptive forces of electrophoresis. This is especially relevant when monitoring labile DNA structures.
Despite these strengths over conventional assays, the method described here does have some disadvantages. While the use of oligonucleotide DNA substrates for strand exchange simplifies interpretation of the results, it is important to remember that such substrates do not resemble the DNA substrates involved in HR in the cell. Some conventional assays utilize plasmid-sized DNA substrates, which are more likely to reflect the number of base-pairs that are exchanged in vivo. Furthermore, the use of topologically constrained circular dsDNA substrates in a subset of conventional assays can at least partially recreate the tensions in physiological DNA.
The application of the method described here has started to unravel the molecular mechanisms of Rad51-driven DNA strand exchange. Nonetheless, there are many interesting questions that remain to be answered. There is clear evidence that HR during meiosis requires both Rad51 and Dmc1, the meiosis-specific RecA-type recombinase in eukaryotes24. However, the lack of major biochemical differences between these two recombinases has baffled researchers in the field for years. Moreover, the roles of numerous distinct groups of recombination accessory factors has been a focal topic of research in the field of HR. In addition to elucidating the biochemical differences between Rad51 and Dmc1, we aim to investigate and compare the effects of different recombination accessory factors on the kinetics of DNA strand exchange in the immediate future. Finally, it is important to stress that the FRET-based methodology described here is not limited to the study of DNA strand exchange. With relatively minor modifications, we envision many kinds of applications for this technique in investigating functionally diverse proteins involved in DNA metabolism25,26,27,28. We hope the developments described here will provide further options to researchers belonging to many different disciplines.

Homologous recombination (HR) facilitates the shuffling of genetic information between two different DNA molecules. HR is essential for two fundamental biological phenomena: the generation of genetic diversity during gametogenesis1 and the repair of DNA double-strand breaks (DSBs)2 during mitosis. DSBs are the most severe form of DNA damage and constitute a breakage in the chromosome. Incorrect repair of DSBs can cause extensive chromosomal rearrangements and genomic instability, which are both hallmarks of cancer3.
The DNA strand exchange reaction is the central phase of HR. The Rad51 protein, which is a member of the highly conserved RecA-type family of recombinases, is the key protein that catalyzes this reaction in eukaryotes4,5. In this reaction, Rad51 binds to single-stranded DNA (ssDNA) generated by nucleolytic processing of the DSB end and forms a helical nucleoprotein complex termed the presynaptic filament. This filament catches intact double-stranded DNA (dsDNA) nonspecifically to search for a homologous sequence. When the filament finds a homologous sequence, a reaction intermediate containing three-stranded DNA is formed and the Rad51 filament mediates strand exchange within this structure6,7,8. To accomplish this reaction efficiently, Rad51 requires several kinds of accessory proteins such as BRCA1 and BRCA2, products of breast cancer susceptibility genes9,10.
Understanding how accessory factors regulate Rad51 is an integral step in uncovering the causes of genomic instability during tumorigenesis. Although much research is concerned with the effects of these factors on presynaptic filament formation and stability11,12,13,14,15,16, the contribution of these factors to formation of the three-strand intermediate and its processing into the final product is still unclear. Observing these reaction steps through conventional biochemical experiments is very difficult because the three-strand intermediate is unstable and prone to collapse by common experimental manipulations such as deproteinization of samples or electrophoresis.
To overcome this problem, we adapted two previously developed real-time observation systems of the DNA strand exchange reaction using fluorescence resonance energy transfer (FRET): the DNA strand pairing and DNA strand displacement assays17,18 (Figure 1). In the DNA strand pairing assay, Rad51 forms a presynaptic filament with fluorescein amidite (FAM)-labeled ssDNA and then homologous carboxy-x-rhodamine (ROX)-labeled dsDNA is added to initiate the strand exchange reaction. When the filament catches the ROX-labeled dsDNA and forms the three-strand intermediate, the two fluorophores come into close proximity and fluorescence emission of FAM is quenched by ROX (Figure 1A). In the DNA strand displacement assay, a presynaptic filament formed on unlabeled ssDNA is incubated with FAM and ROX double-labeled dsDNA. When strand exchange is completed and the FAM labeled ssDNA is released from the three-strand intermediate, the emission of FAM increases because FAM is no longer in close proximity to ROX (Figure 1B). These assays enable us to observe the formation of three-strand intermediates and their processing into final products in real-time without any disturbances to the reaction.
Using this real-time observation system, we found that the DNA strand exchange reaction mediated by Rad51 proceeds in three-steps including the formation of the first reaction intermediate (C1), transitioning of the first intermediate into a second intermediate (C2), and release of ssDNA from C219. We also found that fission yeast (S. pombe) Swi5-Sfr1, which is an evolutionarily conserved Rad51 accessory protein complex13,16,20,21,22, stimulates the C1-C2 transitioning and release of ssDNA from C2 in a manner that is dependent on ATP-hydrolysis by Rad5119.
Whether these findings are evolutionarily conserved remains unknown. This protocol is provided with the hope that researchers in the field of HR, especially those working with proteins from organisms other than S. pombe, may apply these techniques to determine the extent to which the molecular mechanism of Rad51-driven strand exchange is conserved. Furthermore, these techniques have proven highly successful in determining the role of S. pombe Swi5-Sfr1. Thus, it is a rational prediction that these techniques will be invaluable in uncovering the precise roles of other HR accessory factors.

<table><tbody><tr><td>0.2 x 1.0 cm quartz cuvette</td><td>Hellma Analytics</td><td>105-250-15-40</td><td></td></tr><tr><td>1.0 x 1.0 cm quartz cuvette</td><td>Hellma Analytics</td><td>101-10-40</td><td></td></tr><tr><td>adenosine triphosphate (ATP)</td><td>Sigma</td><td>A2383</td><td></td></tr><tr><td>DynaFit</td><td>BioKin, Ldt.</td><td></td><td>DynaFit is a program to analyze kinetics of biochemical reactions.</td></tr><tr><td>Fluorescent labeled and non-labeled oligonucleotides</td><td>Eurofins Genomics</td><td></td><td>The sequences of oligos are listed in Table. 1.</td></tr><tr><td>Magnetic stirrer</td><td>Aisis (Japan)</td><td>CM1609</td><td></td></tr><tr><td>PCR machine</td><td>TAKARA (Japan)</td><td>TP600</td><td>TAKARA PCR Thermal Cycler Dice</td></tr><tr><td>Spectrofluorometer</td><td>JASCO</td><td>FP8300</td><td>Contains a peltier temperature controller and magnetic stirrer system</td></tr><tr><td>Syringe</td><td>HAMILTON</td><td>1702RN 25ul SYR (22s/2&quot;/2)</td><td></td></tr></tbody></table>

real-time observation of the dna strand exchange reaction mediated by rad51

The DNA strand exchange reaction mediated by Rad51 is a critical step of homologous recombination. In this reaction, Rad51 forms a nucleoprotein filament on single-stranded DNA (ssDNA) and captures double-stranded DNA (dsDNA) non-specifically to interrogate it for a homologous sequence. After encountering homology, Rad51 catalyzes DNA strand exchange to mediate pairing of the ssDNA with the complementary strand of the dsDNA. This reaction is highly regulated by numerous accessary proteins in vivo. Although conventional biochemical assays have been successfully employed to examine the role of such accessory protein in vitro, kinetic analysis of intermediate formation and its progression into a final product has proven challenging due to the unstable and transient nature of the reaction intermediates. To observe these reaction steps directly in solution, fluorescence resonance energy transfer (FRET)-based real-time observation systems of this reaction were established. Kinetic analysis of real-time observations shows that the DNA strand exchange reaction mediated by Rad51 obeys a three-step reaction model involving the formation of a three-strand DNA intermediate, maturation of this intermediate, and the release of ssDNA from the mature intermediate. The Swi5-Sfr1 complex, an accessary protein conserved in eukaryotes, strongly enhances the second and third steps of this reaction. The FRET-based assays presented here enable us to uncover the molecular mechanisms through which recombination accessary proteins stimulate the DNA strand exchange activity of Rad51. The primary goal of this protocol is to enhance the repertoire of techniques available to researchers in the field of homologous recombination, particularly those working with proteins from species other than Schizosaccharomyces pombe, so that the evolutionary conservation of the findings presented herein can be determined.

In order to effectively analyze the experimental data from the pairing and displacement assays, it is necessary to define how a change in fluorescence emission of FAM corresponds to a conversion of DNA substrates into products. To achieve this, the relevant range of fluorescence intensity must be determined. For the pairing assay, the fluorescence emission of 16FA(-), which corresponds to the ssDNA substrate, is compared with the emission of 16FA(-) annealed with 16AR(+)_40bp, which corresponds to the final products of this reaction (Figure 2A). This equates to the maximum FRET efficiency and hence the maximum reduction in fluorescence intensity that would be expected if all ssDNA substrate was converted into the dsDNA product. For the displacement assay, the emission of 16FA(-)_40bp annealed with 16AR(+)_40bp, which corresponds to the substrate, is compared with the emission of 16FA(-)_40bp, which corresponds to the final product (Figure 2B). In this case, the maximum increase in fluorescence intensity of FAM conveys a scenario in which all of the dsDNA substrate is converted into ssDNA product. S. pombe Rad51 did not affect fluorescence emission of FAM or quenching efficiency of FAM by ROX in both assays (Figure 2). The maximum FRET efficiency should be re-measured with each new preparation of oligonucleotides as it is dependent on the labeling efficiency of oligonucleotides.
Representative data of DNA strand pairing and displacement reactions are shown in Figure 3. The effects of spontaneous reactions between substrate DNAs and photobleaching were small in both assays, as revealed by the negligible changes seen in the emission of FAM without Rad51 compared to the substantial changes seen with Rad51 (Figure 3A and Figure 3B). Based on the data shown in Figure 3A or Figure 3B, the change in fluorescence was converted into the change in amount of substrate (FP) or final product (FD), respectively, using the equations described in steps 3.2.2 or 3.3.2 (Figure 3C and Figure 3D).
The pairing reaction was simulated using a sequential three-step reaction model, consisting of the formation of the first three-strand intermediate (C1), transitioning of the first intermediate into the second intermediate (C1 to C2), and the release of ssDNA from the second intermediate to form the two products (D + E) (Figure 3E). To test whether the simulation using a sequential three-step reaction model fit the experimental data, residuals between experimental data of the DNA strand pairing assay and a theoretical curve obtained by the simulation were calculated (Figure 3F). In addition, residuals between the pairing reaction and a theoretical curve generated using a sequential two-step reaction model were also calculated (Figure 3G). The residuals for the pairing reaction and the two-step model show a systematic deviation in the early stage, whereas the residuals for the pairing reaction and the three-step model do not show such a deviation. This indicates that the three-step model is a better fit than the two-step model for simulating the pairing reaction.
To test whether the three-step model is consistent with the displacement reaction that detects the late step of DNA strand exchange, we generated a theoretical curve of the displacement reaction using the kinetic parameters obtained from simulation of the pairing reaction shown in Figure 3C and compared it with the experimental data of the displacement reaction shown in Figure 3D (Figure 3H). The theoretical curve fit the experimental data of the displacement assay. From these results, we conclude that simulation using the three-step model is able to reasonably evaluate the DNA strand exchange reaction mediated by Rad51.
Representative data of the DNA strand pairing reaction containing Rad51 and the Swi5-Sfr1 complex, an accessary protein of Rad51, are shown in Figure 4A. The Swi5-Sfr1 complex strongly stimulated the pairing activity of Rad51. As was seen in the absence of Swi5-Sfr1, the pairing reaction better fit the three-step model than the two-step model in the presence of Swi5-Sfr1 (Figure 4B). Through simulation of the reaction using the three-step model, reaction equilibrium constants of each reaction step with or without Swi5-Sfr1 were calculated. The reaction equilibrium constants indicated that the Swi5-Sfr1 complex does not stimulate the first reaction step (Figure 4C, panel a), in which a three-strand intermediate is formed, but strongly stimulates C1-C2 transitioning (Figure 4C, panel b) and the release of ssDNA from the C2 intermediate (Figure 4C, panel c).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59073/59073fig1.jpg" /> 
Figure 1: Experimental design of DNA strand pairing and displacement assays. Schematics of DNA strand pairing (A) and displacement (B) assays. Yellow circles represent Rad51 monomers. Green circles containing &#34;F&#34; and red circles containing &#34;R&#34; represent fluorescein amidite (FAM) and carboxy-X-rhodamine (ROX), respectively. Black DNA strands are identical in sequence and complementary to blue DNA strands. Thin black lines with arrowheads point to the name of each oligonucleotide, as depicted in Table 1. This figure has been adapted from Ito et al.19 and modified. <a href="https://www.jove.com/files/ftp_upload/59073/59073fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59073/59073fig2.jpg" /> 
Figure 2: Measurements of the maximum FRET efficiency of the pairing and displacement assays. (A) Comparison of fluorescence spectra between the ssDNA substrate, 16FA(-), and the dsDNA product, 16FA(-) paired with 16AR(+)_40bp, of the pairing assay. Blue and red lines represent the fluorescence spectra of the substrate without and with Rad51, respectively. Green and purple lines show the fluorescence spectra of the final product without and with Rad51, respectively. (B) Comparison of fluorescence spectra between the dsDNA substrate, 16FA(-)_40bp paired with 16AR(+)_40bp, and ssDNA product, 16FA(-)_40bp, of the displacement assay. Blue and red lines represent the fluorescence spectra of the final product without and with Rad51, respectively. Green and purple lines show the fluorescence spectra of the substrate without and with Rad51, respectively. This figure is adapted from Ito et al.19 and modified. <a href="https://www.jove.com/files/ftp_upload/59073/59073fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59073/59073fig3.jpg" /> 
Figure 3: DNA strand pairing and displacement reactions mediated by Rad51. (A) Time course of the normalized fluorescence of the pairing reaction with or without Rad51. (B) Time course of the normalized fluorescence of the displacement reaction with or without Rad51. (C)Time course of the change in the amount of substrate in the pairing reaction with Rad51. (D) Time course of the change in the amount of substrate in the displacement reaction with Rad51. (E) A schematic of the sequential three-step reaction model. A and B correspond to the presynaptic filament and donor dsDNA. C1 corresponds to the first (immature) three-strand intermediate. C2 corresponds to the second (mature) three-strand intermediate. D and E correspond to a heteroduplex and ssDNA released from C2. (F and G) Residuals between experimental data of the DNA strand pairing assay and a theoretical curve obtained by simulation using either the three-step (F) or two-step (G) model. (H) Red dots indicate experimental data from the displacement reaction with Rad51 shown in panel D. Blue line indicates the theoretical curve of the final products. The theoretical curve was generated by simulation using the reaction rate constants obtained from the pairing assay shown in panel C. This figure is adapted from Ito et al.19 and modified. <a href="https://www.jove.com/files/ftp_upload/59073/59073fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59073/59073fig4.jpg" /> 
Figure 4: Swi5-Sfr1 stimulates the second and third steps of the DNA strand exchange reaction. (A) Time course of the change in the amount of substrate in the pairing reaction with or without Swi5-Sfr1 (S5S1). (B)&#160;Residuals between experimental data of the DNA strand pairing assay with Swi5-Sfr1 and a theoretical curve obtained by simulation using the two-step (blue line) or three-step (red line) model. (C)&#160;The pairing reaction shown in Figure 4A was simulated by the three-step model using the analysis program23 (see Table of Materials). The reaction equilibrium constants of each reaction step, K1 (a), K2 (b), and K3 (c), were obtained from the simulation. This figure is adapted from Ito et al.19 and modified. <a href="https://www.jove.com/files/ftp_upload/59073/59073fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="5">Oligonucleotides for DNA strand pairing assay</td>
		</tr>
		<tr>
			<td>16FA(-)</td>
			<td colspan="4">5&#8217;-[FAM]-AAATGAACATAAAGTAAATAAGTATAAGGATAATACA 
			AAATAAGTAAATGAATAAACATAGAAAATAAAGTAAAGGATAT AAA -3&#8217;</td>
		</tr>
		<tr>
			<td>16A(-)_40bp</td>
			<td colspan="4">5&#8217;-AAATGAACATAAAGTAAATAAGTATAAGGATAATACAAAA-3&#8217;</td>
		</tr>
		<tr>
			<td>16AR(+)_40bp</td>
			<td colspan="4">5&#8217;-TTTTGTATTATCCTTATACTTATTTACTTTATGTTCATTT-[ROX]-3&#8217;</td>
		</tr>
		<tr>
			<td colspan="5">Oligonucleotides for DNA strand displacement assay</td>
		</tr>
		<tr>
			<td>16A(-)</td>
			<td colspan="4">5&#8217;-AAATGAACATAAAGTAAATAAGTATAAGGATAATACAAAATA 
			AGTAAATGAATAAACATAGAAAATAAAGTAAAGGATAT AAA -3&#8217;</td>
		</tr>
		<tr>
			<td>16FA(-)_40bp</td>
			<td colspan="4">5&#8217;-[FAM]-AAATGAACATAAAGTAAATAAGTATAAGGATAATACAAAA-3&#8217;</td>
		</tr>
		<tr>
			<td>16AR(+)_40bp</td>
			<td colspan="4">5&#8217;-TTTTGTATTATCCTTATACTTATTTACTTTATGTTCATTT-[ROX]-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 1: A list of oligonucleotides used in the DNA strand pairing and displacement assays. Where applicable, the positions of fluorophores (fluorescein amidite, FAM; carboxy-x-rhodamine, ROX) are indicated in square parentheses.

Watch this Scientific Journal Video about Real-time Observation of the DNA Strand Exchange Reaction Mediated by Rad51 at JoVE.com

Real-time Observation of the DNA Strand Exchange Reaction Mediated by Rad51

Fluorescence resonance energy transfer-based real-time observation systems of the DNA strand exchange reaction mediated by Rad51 were developed. Using the protocols presented here, we are able to detect the formation of reaction intermediates and their conversion into products, while also analyzing the enzymatic kinetics of the reaction.

The DNA strand exchange reaction mediated by Rad51 is a critical step of homologous recombination. In this reaction, Rad51 forms a nucleoprotein filament ...

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DNA Repair and Recombination

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7.9K Views.  Tokyo Institute of Technology. Fluorescence resonance energy transfer-based real-time observation systems of the DNA strand exchange reaction mediated by Rad51 were developed. Using the protocols presented here, we are able to detect the formation of reaction intermediates and their conversion into products, while also analyzing the enzymatic kinetics of the reaction.

Video: Real-time Observation of the DNA Strand Exchange Reaction Mediated by Rad51

Characterizing DNA Repair Processes at Transient and Long-lasting Double-strand DNA Breaks by Immunofluorescence Microscopy

The repair of double-stranded breaks (DSBs) in DNA is a highly coordinated process, necessitating the formation and resolution of multi-protein repair complexes. This process is regulated by a myriad of proteins that promote the association and disassociation of proteins to these lesions. Thanks in large part to the ability to perform functional screens of a vast library of proteins, there is a greater appreciation of the genes necessary for the double-strand DNA break repair. Often knockout or chemical inhibitor screens identify proteins involved in repair processes by using increased toxicity as a marker for a protein that is required for DSB repair. Although useful for identifying novel cellular proteins involved in maintaining genome fidelity, functional analysis requires the determination of whether the protein of interest promotes localization, formation, or resolution of repair complexes. 
The accumulation of repair proteins can be readily detected as distinct nuclear foci by immunofluorescence microscopy. Thus, association and disassociation of these proteins at sites of DNA damage can be accessed by observing these nuclear foci at representative intervals after the induction of double-strand DNA breaks. This approach can also identify mis-localized repair factor proteins, if repair defects do not simultaneously occur with incomplete delays in repair. In this scenario, long-lasting double-strand DNA breaks can be engineered by expressing a rare cutting endonuclease (e.g., I-SceI) in cells where the recognition site for the said enzyme has been integrated into the cellular genome. The resulting lesion is particularly hard to resolve as faithful repair will reintroduce the enzyme's recognition site, prompting another round of cleavage. As a result, differences in the kinetics of repair are eliminated. If repair complexes are not formed, localization has been impeded. This protocol describes the methodology necessary to identify changes in repair kinetics as well as repair protein localization.

Repair of double-strand DNA breaks is a dynamic process, requiring not only formation of repair complexes at the breaks, but also their resolution after the lesion is addressed. Here, we use immunofluorescence microscopy for transient and long-lasting double-stranded breaks as a tool to dissect this genome maintenance mechanism.

The repair of double-stranded breaks (DSBs) in DNA is a highly coordinated process, necessitating the formation and resolution of multi-protein repair ...

Cancer Research

DNA Curtains Shed Light on Complex Molecular Systems During Homologous Recombination

Homologous recombination (HR) is important for the repair of double-stranded DNA breaks (DSBs) and stalled replication forks in all organisms. Defects in HR are closely associated with a loss of genome integrity and oncogenic transformation in human cells. HR involves coordinated actions of a complex set of proteins, many of which remain poorly understood. The key aspect of the research described here is a technology called "DNA curtains", a technique which allows for the assembly of aligned DNA molecules on the surface of a microfluidic sample chamber. They can then be visualized by total internal reflection fluorescence microscopy (TIRFM). DNA curtains was pioneered by our laboratory and allows for direct access to spatiotemporal information at millisecond time scales and nanometer scale resolution, which cannot be easily revealed through other methodologies. A major advantage of DNA curtains is that it simplifies the collection of statistically relevant data from single molecule experiments. This research continues to yield new insights into how cells regulate and preserve genome integrity.

DNA curtains present a novel method for visualizing hundreds or even thousands of DNA-binding proteins in real-time as they interact with DNA molecules aligned on the surface of a microfluidic sample chamber.

Homologous recombination (HR) is important for the repair of double-stranded DNA breaks (DSBs) and stalled replication forks in all organisms. Defects in ...

Genetics

Detection of Homologous Recombination Intermediates via Proximity Ligation and Quantitative PCR in Saccharomyces cerevisiae

DNA damage, including DNA double-stranded breaks and inter-strand cross-links, incurred during the S and G2 phases of the cell cycle can be repaired by homologous recombination (HR). In addition, HR represents an important mechanism of replication fork rescue following stalling or collapse. The regulation of the many reversible and irreversible steps of this complex pathway promotes its fidelity. The physical analysis of the recombination intermediates formed during HR enables the characterization of these controls by various nucleoprotein factors and their interactors. Though there are well-established methods to assay specific events and intermediates in the recombination pathway, the detection of D-loop formation and extension, two critical steps in this pathway, has proved challenging until recently. Here, efficient methods for detecting key events in the HR pathway, namely DNA double-stranded break formation, D-loop formation, D-loop extension, and the formation of products via break-induced replication (BIR) in Saccharomyces cerevisiae are described. These assays detect their relevant recombination intermediates and products with high sensitivity and are independent of cellular viability. The detection of D-loops, D-loop extension, and the BIR product is based on proximity ligation. Together, these assays allow for the study of the kinetics of HR at the population level to finely address the functions of HR proteins and regulators at significant steps in the pathway.

Detection of Homologous Recombination Intermediates via Proximity Ligation and Quantitative PCR in Saccharomyces cerevisiae

The D-loop capture (DLC) and D-loop extension (DLE) assays utilize the principle of proximity ligation together with quantitative PCR to quantify D-loop formation, D-loop extension, and product formation at the site of an inducible double-stranded break in Saccharomyces cerevisiae.

DNA damage, including DNA double-stranded breaks and inter-strand cross-links, incurred during the S and G2 phases of the cell cycle can be repaired by ...

Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for hundreds of individual restriction endonuclease (REase) molecules in one experiment. This assay does not require protein labeling and can be carried out with a single imaging channel. In addition, the results of multiple individual experiments can be pooled to generate well-populated dwell-time distributions. Analysis of the resulting dwell-time distributions can help elucidate the DNA cleavage mechanism by revealing the presence of kinetic steps that cannot be directly observed. Example data collected using this assay with the well-studied REase, EcoRV - a dimeric Type IIP restriction endonuclease that cleaves the palindromic sequence GAT&#8595;ATC (where &#8595; is the cut site) - are in agreement with prior studies. These results suggest that there are at least three steps in the pathway to DNA cleavage that is initiated by introducing magnesium after EcoRV binds DNA in its absence, with an average rate of 0.17 s-1 for each step.

Using quantum-dot-labeled DNA and total internal reflection fluorescence microscopy, we can investigate the reaction mechanism of restriction endonucleases while using unlabeled protein. This single-molecule technique allows for massively multiplexed observation of individual protein-DNA interactions, and data can be pooled to generate well-populated dwell-time distributions.

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for ...

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. Linearized &lambda; DNA is modified to have a biotin on the end of one strand, and a digoxigenin moiety on the other end of the same strand. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead. The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied to exert a drag force on the bead. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate (Figure 1). The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork. Measuring the position of the bead allows precise determination of the rates and processivities of DNA unwinding and polymerization (Figure 2).

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system.

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. ...

Biology

Measuring Ubiquitylated, SUMOylated, and ADP-Ribosylated DNA-Protein Crosslinks

Quantitative Detection of DNA-Protein Crosslinks and Their Post-Translational Modifications

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, methyltransferases, etc.) malfunctioning, or exogenous agents such as chemotherapeutics and crosslinking agents. Once DPCs are induced, several types of post-translational modifications (PTMs) are promptly conjugated to them as early response mechanisms. It has been shown that DPCs can be modified by ubiquitin, small ubiquitin-like modifier (SUMO), and poly-ADP-ribose, which prime the substrates to signal their respective designated repair enzymes and, in some cases, coordinate the repair in sequential manners. As PTMs transpire quickly and are highly reversible, it has been challenging to isolate and detect PTM-conjugated DPCs that usually remain at low levels. Presented here is an immunoassay to purify and quantitatively detect ubiquitylated, SUMOylated, and ADP-ribosylated DPCs (drug-induced topoisomerase DPCs and aldehyde-induced non-specific DPCs) in vivo. This assay is derived from the RADAR (rapid approach to DNA adduct recovery) assay that is used for the isolation of genomic DNA containing DPCs by ethanol precipitation. Following normalization and nuclease digestion, PTMs of DPCs, including ubiquitylation, SUMOylation, and ADP-ribosylation, are detected by immunoblotting using their corresponding antibodies. This robust assay can be utilized to identify and characterize novel molecular mechanisms that repair enzymatic and non-enzymatic DPCs&#160;and has the potential to discover small molecule inhibitors targeting specific factors that regulate PTMs to repair DPCs.

Author Spotlight: Quantitative Detection of DNA Protein Crosslinks and Their Post-Translational Modifications  

The present protocol highlights a modified method to detect and quantify DNA-protein crosslinks (DPCs) and their post-translational modifications (PTMs), including ubiquitylation, SUMOylation, and ADP-ribosylation induced by topoisomerase inhibitors and by formaldehyde, thereby allowing the study of the formation and repair of DPCs and their PTMs.

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, ...

CMG-Mediated DNA Unwinding Observed by ssDNA-RPA Tract Growth

Single-Molecule Real-Time Visualization of DNA Unwinding by CMG Helicase

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a dynamic complex of proteins termed the replisome. At the heart of the replisome is the&#160;CDC45-MCM2-7-GINS (CMG) helicase, which separates the two strands of the DNA double helix such that DNA polymerases can copy each strand. During genome duplication, replisomes must overcome a plethora of obstacles and challenges. Each of these threatens genome stability, as failure to replicate DNA completely and accurately can lead to mutations, diseases, or cell death. Therefore, it is of great interest to understand how CMG functions in the replisome during both normal replication and replication stress. Here, we describe a total internal reflection fluorescence (TIRF) microscopy assay using recombinant purified proteins, which allows for real-time visualization of surface-tethered stretched DNA molecules by individual CMG complexes. This assay provides a powerful platform to investigate CMG behavior at the single-molecule level, allowing helicase dynamics to be directly observed with real-time control over reaction conditions.

Author Spotlight: Unraveling the Dynamics of Eukaryotic DNA Replication Through Single-Molecule Visualization

This protocol demonstrates performing a single-molecule assay for live visualization of DNA unwinding by CMG helicase. It describes (1) preparing a DNA substrate, (2) purifying fluorescently labeled Drosophila melanogaster CMG helicase, (3) preparing a microfluidic flow cell for total internal reflection fluorescence (TIRF) microscopy, and (4) the single-molecule DNA unwinding assay.

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a ...

Investigating the Extent of DNA Breaks Occurring During DNA Replication 

Detection of DNA Breaks in Dividing Human Cells by Neutral Comet Assay

DNA replication is constantly challenged by a wide variety of endogenous and exogenous stressors that can damage DNA. Such lesions encountered during genome duplication can stall replisomes and convert replication forks into double-strand breaks. If left unrepaired, these toxic DNA breaks can trigger chromosomal rearrangements, leading to heightened genome instability and an increased likelihood of cellular transformation. Additionally, cancer cells exhibit persistent replication stress, making the targeting of replication fork vulnerabilities in tumor cells an attractive strategy for chemotherapy. A highly versatile and powerful technique to study DNA breaks during replication is the comet assay. This gel electrophoresis technique reliably detects the induction and repair of DNA breaks at the single-cell level. Herein, a protocol is outlined that allows investigators to measure the extent of DNA damage in mitotically dividing human cells using fork-stalling agents across multiple cell types. Coupling this with automated comet scoring facilitates rapid analysis and enhances the reliability in studying induction of DNA breaks.

Author Spotlight: Unveiling the Role of SNF2L in Replication Fork Stability and Genome Duplication

This protocol is designed to investigate the extent of DNA damage occurring during DNA replication. Under neutral conditions, the induction of DNA breaks can be readily assessed within a short time frame. Additionally, the protocol is adaptable to other cell types and various replication stress reagents.

DNA replication is constantly challenged by a wide variety of endogenous and exogenous stressors that can damage DNA. Such lesions encountered during ...

Bioengineering

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA insults. Genotoxic agents can damage the DNA backbone, break it, or modify the chemical nature of individual bases. Following DNA insult, DNA damage response (DDR) pathways are activated and proteins involved in the repair are recruited. A plethora of factors are involved in sensing the type of damage and activating the appropriate repair response. Failure to correctly activate and recruit DDR factors can lead to genomic instability, which underlies many human pathologies including cancer. Studies of DDR proteins can provide insights into genotoxic drug response and cellular mechanisms of drug resistance.
There are two major ways of visualizing&#160;proteins in vivo: direct observation, by tagging the protein of interest with a fluorescent protein and following it by live imaging, or indirect immunofluorescence on fixed samples. While visualization of fluorescently tagged proteins allows precise monitoring over time, direct tagging in N- or C-terminus can interfere with the protein localization or function. Observation of proteins in their unmodified, endogenous version is preferred. When DNA repair proteins are recruited to the DNA insult, their concentration increases locally and they form groups, or &#8220;foci&#8221;, that can be visualized by indirect immunofluorescence using specific antibodies.
Although detection of protein foci does not provide a definitive proof of direct interaction, co-localization of proteins in cells indicates that they regroup to the site of damage and can inform of the sequence of events required for complex formation. Careful analysis of foci spatial overlap in cells expressing wild type or mutant versions of a protein can provide precious clues on functional domains important for DNA repair function. Last, co-localization of proteins indicates possible direct interactions that can be verified by co-immunoprecipitation in cells, or direct pulldown using purified proteins.

Following DNA damage, human cells activate essential repair pathways to restore the integrity of their genome. Here, we describe the method of indirect immunofluorescence as a means to detect DNA repair proteins, analyze their spatial and temporal recruitment, and help interrogate protein-protein interaction at the sites of DNA damage.

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA ...

Homologous Recombination

The basic reaction of homologous recombination (HR) involves two chromatids that contain DNA sequences sharing a significant stretch of identity. One of these sequences uses a strand from another as a template to synthesize DNA in an enzyme-catalyzed reaction. The final product is a novel amalgamation of the two substrates. To ensure an accurate recombination of sequences, HR is restricted to the S and G2 phases of the cell cycle. At these stages, the DNA has been replicated already and the likelihood of an identical or similar DNA sequence on a sister chromatid is high. Thus, the timing of repair prevents recombination between non-identical sequences. This is a critical feature of HR, particularly during the recombination of parental DNA sequences in an offspring, where faulty HR can lead to loss of the entire gene and the surrounding chromosomal region.
The accurate repair ensured by HR has been applied in gene-editing techniques. HR is the earliest method that has been used to edit genomes in living cells. The CRISPR-Cas9 system is used to create targeted double-strand breaks to correct disease-causing mutations in the genome. The isolated fragments are taken up by cells, where they can recombine with cellular DNA and replace the targeted region of the genome. HR mechanisms govern the repair of the breaks and their accurate recombination with the cellular genome. To help the HR proteins localize precisely at double-strand breaks, Cas9 proteins are fused with HR effector proteins that can recruit repair proteins at the damaged sites. Studies have shown that fusing Cas9 with proteins such as CtIP, Rad52, and Mre11 can increase HR events in the cell by two-folds while discouraging Non-homologous end joining. Such applications of HR in genome editing can revolutionize gene therapy and provide treatment for genetic diseases that are currently considered incurable.

Homologous Recombination and Strand Invasion

Rekombinacja homologiczna

The basic reaction of homologous recombination (HR) involves two chromatids that contain DNA sequences sharing a significant stretch of identity. One of ...

Real-time Observation of the DNA Strand Exchange Reaction Mediated by Rad51

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Characterizing DNA Repair Processes at Transient and Long-lasting Double-strand DNA Breaks by Immunofluorescence Microscopy

DNA Curtains Shed Light on Complex Molecular Systems During Homologous Recombination

Detection of Homologous Recombination Intermediates via Proximity Ligation and Quantitative PCR in Saccharomyces cerevisiae

Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

Quantitative Detection of DNA-Protein Crosslinks and Their Post-Translational Modifications

Single-Molecule Real-Time Visualization of DNA Unwinding by CMG Helicase

Detection of DNA Breaks in Dividing Human Cells by Neutral Comet Assay

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Rekombinacja homologiczna