We thank Funda&#231;&#227;o de Amparo a Pesquisa do Estado de S&#227;o Paulo (FAPESP, through Grant Tem&#225;tico 2022/15126-9 to JK and fellowship 21/09439-1 to LARM) and the Conselho Nacional de Desenvolvimento Cientifico e Tecnol&#243;gico (CNPq) for funding this research.

1. Cell plating
NOTE: Cells can be seeded in microscopic fluorescence slides, chambers, or plates. The use of small coverslips or 96/ 384 well plates is recommended for testing multiple conditions and spare reagents. The use of coverslips is advisable for cell lines that detach easily, like HEK293 cells. The coverslips can be previously coated with a poly-L-lysine solution to improve the attachment.
<ol>
	<li>Plate cells considering a vehicle, technical controls, and biological controls. Use technical replicates at two or three wells per condition. 
	NOTE: The use of technical control (both secondaries and one primary only) is mandatory. If possible, biological control should also be included (e.g., overexpression or knockdown of one of the target proteins or a condition that is known to change the interaction, for example).
	<ol>
		<li>Culture U2OS cells in Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% of Fetal Bovine Serum (FBS), 4.5 g/L Glucose, and 4 mM of L-Glutamine on tissue culture-treated plates at 37 &#176;C in 5% CO2 and a humidified atmosphere containing 90% air. Depending on the number of conditions to be tested and if using a 384 or coverslips at a 60 mm plate, start from a 60- or 100-mm plate, respectively.</li>
		<li>When the cells reach 60%-70% confluency, split cells using 0.25% trypsin-EDTA 14. Count cells using a Neubauer chamber or an automatic counter14. Plate 4 x 105 cells in 60 mm plate containing multiple 13 mm round coverslips or 4 x 105 cells per well in 384-well plate(s) and incubate for 12 - 24 hours at 37 &#176;C in a humidified, 5% CO2 incubator.</li>
	</ol>
	</li>
</ol>
2. Preparation of solutions
NOTE: The concentration and times of the treatment were chosen based on literature results, which point out the activation of DNA damage repair pathways15,16.
<ol>
	<li>Prepare the stock solution of Etoposide at 50 mM concentration in DMSO. The stock solution can be kept at -20 &#176;C for several months. Dilute the etoposide solution for use in the complete culture medium at 25 &#181;M final concentration. Use DMSO at 0.05% diluted in the complete culture medium as vehicle control.</li>
	<li>Prepare 20x Saline Sodium Citrate (SSC; 3.0 M NaCl, 0.30 M trisodium citrate pH 7.0). This solution can be kept for several months at 4 &#176;C.</li>
</ol>
3. Cell treatment and fixation
NOTE: Do not allow cells to dry at any moment; try to tap off the previous solutions as much as possible. The appropriate method for cell fixation (ice-cold methanol or 4% paraformaldehyde (PFA)) must be determined previously, as well as the antibody dilution, using a conventional immunofluorescence experiment.
CAUTION: Methanol must be disposed of properly.
<ol>
	<li>Remove the culture medium from the wells and add a culture medium containing etoposide or DMSO at 25 &#181;M or 0.05%, respectively. Incubate cells at 37 &#176;C in 5% CO2 and a humidified atmosphere containing 90% air with etoposide or DMSO for 3 different periods of time: 20 min, 1 h, and 3 h. When using 384 plates, all conditions must be fixed simultaneously.
	<ol>
		<li>For this, start by treating the condition with the longest time of treatment, such as 3 h. After 2 h, treat the 1 h treatment condition, and when there is 20 min left for fixation, treat the 20 min condition. Following this, all conditions can be fixed at the same time.</li>
	</ol>
	</li>
	<li>Remove the medium from the wells and wash the cells with PBS 3x.</li>
	<li>Fix the cells by adding ice-cold methanol with enough volume to cover the well surface. Incubate for 10 min at -20 &#176;C. Wash the wells with PBS 3x. 
	NOTE: At this point, the protocol can be paused. The plates can be stored at 4 &#176;C for 1 week.</li>
</ol>
4. Permeabilization and blocking
NOTE: If using the PFA fixation method, a step of blocking with glycine must be performed before permeabilization. The humidity chamber can be prepared by putting a water pan inside the incubator, resulting in a humidity of around 95%. The 384 plate or slides can be incubated in a closed plastic container above filter paper or moistened sponge and transparent film.
<ol>
	<li>Tap off the PBS from the samples.</li>
	<li>Add 30-40 &#181;L of permeabilization buffer (0.2% Triton X-100 in PBS) and incubate for 20 min at room temperature. Wash 3x with PBS</li>
	<li>Tap off the PBS from the samples and add 30-40 &#181;L of 1x blocking solution provided in the PLA kit (alternatively, 3% of Bovine Serum Albumin and Triton X-100 0.1% in PBS solution can be used) to each well/coverslip.</li>
	<li>Incubate the plate in a pre-heated humidity chamber at 37 &#176;C for 1 h.</li>
</ol>
5. Primary antibody incubation (PLA)
NOTE: If using coverslips, it is important to ensure that the antibody solution covers all coverslip surfaces. Special attention must be paid to antibody selection. The antibodies must be from different animal species, and the secondary antibody used for &#947;-H2AX must not be against the species host of the PLA probes production. For example, the primary antibodies used for PLA were raised in mice and goat animals. The PLA probes, anti-mouse, and anti-goat were produced in the donkey. The antibody against &#947;-H2AX was produced in rabbits, and the secondary utilized is anti-rabbit produced in donkeys. In this way, the secondary will not recognize the PLA probes.
<ol>
	<li>Dilute the primary antibodies (the mouse anti-Ku70 1:100 and goat anti-Nek4 1:50; rabbit anti-Nek5 1:25 and mouse anti-TOPII&#946; 1:25) in antibody diluent provided in the kit (alternatively, 3% of Bovine Serum Albumin and Triton X-100 0.1% in PBS solution can be used). Prepare 30 &#181;L of antibody solution per condition if using coverslips or 20 &#181;L per well if using a 384-well plate.</li>
	<li>Tap off the blocking solution. Add the primary antibody solution to each well. Incubate at 4 &#176;C overnight in a humidity chamber.</li>
</ol>
6. PLA probe incubation
NOTE: Choose the probes, taking into consideration not only the primary antibodies but also those intended for use in the immunofluorescence steps. When using 384-well plates for long 37 &#176;C incubation, slow rotation can be used to improve the distribution of the antibody solution.
<ol>
	<li>Dilute the minus and plus probes 1:5 in the antibody diluent. Prepare 10 &#181;L per well if using 384-well plates or 20 &#181;L per sample if using coverslips. Utilize the following combinations: Donkey anti-mouse minus with Donkey anti-goat plus; Donkey anti-mouse minus with Donkey anti-rabbit plus.</li>
	<li>Tap off the primary antibody solution. Wash 1x with TBS-T (50 mM Tris pH 7.5, 150 mM NaCl, 0.05% of Tween-20). Wash 3x for 5 min each with TBS-T at room temperature. Perform an additional&#160;wash with TBS-T.</li>
	<li>Tap off and add the PLA probe solution with 10 &#181;L for the plates and 20 &#181;L for the coverslips. Incubate in the pre-heated humidity chamber at 37 &#176;C for 1 h.</li>
</ol>
7. Ligation
NOTE: Wait to add Ligase until immediately before adding it to the samples. Ligase must be kept in a freezer block (-20 &#176;C). Remove completely all the wash solution before adding the ligase solution. Avoid leaving the cells without a solution for long periods of time. When using 384-well plates for the 37 &#176;C incubation, slow rotation can be used to improve the distribution of the antibody solution.
<ol>
	<li>Dilute the 5x ligation buffer in high-purity water. Prepare 10 &#181;L per well if using 384-well plates or 20 &#181;L if using coverslips.</li>
	<li>Tap off the PLA probe solution. Wash 1x with TBS-T. Follow with three additional washes, each lasting 5 min, using TBS-T. Wash a final time and tap off the TBS-T before the next step.</li>
	<li>Add Ligase to the 1x Ligation solution at 1:40, immediately before applying it to the samples. Incubate in the pre-heated humidity chamber at 37 &#176;C for&#160;30 min to 1 h.</li>
</ol>
8. Amplification and washing
NOTE: Wait to add the polymerase until immediately prior to addition to the sample. The amplification buffer is light sensitive; protect all solutions containing the buffer from light. Polymerase must be kept in a freezer block (-20 &#176;C). Avoid leaving the cells without a solution for a long period of time. When using 384-well plates for 37 &#176;C incubation, slow rotation can improve the distribution of the antibody solution.
<ol>
	<li>Remove completely all the wash solution before adding the polymerase solution. Avoid leaving the cells without a solution for a long period of time. Dilute the 5x amplification buffer in high-purity water. Prepare 10 &#181;L per well if using 384-well plates or 20 &#181;L if using coverslips.</li>
	<li>Tap off the ligation solution. Wash 1x with TBS-T. Follow with three additional washes, each lasting 5 min, using TBS-T. Wash a final time and tap off the TBS-T before the next step.</li>
	<li>Add the polymerase to the 1x amplification solution at 1:80 dilution, immediately before applying it to the samples. Incubate in the pre-heated humidity chamber at 37 &#176;C for 100 min. 
	NOTE: From this step on, avoid direct light exposition</li>
	<li>Tap off the amplification buffer. Wash 2x with 1x SSC buffer for 10 min each. Wash 2x with 0.01x SSC buffer for 2 min each.</li>
</ol>
9. &#947;-H2AX staining (Immunofluorescence - IF)
NOTE: The labeling of &#947;-H2AX is performed after the PLA is finished.
<ol>
	<li>Primary antibody incubation (IF)
	<ol>
		<li>Dilute the primary antibodies (Rabbit anti yH2AX at 1:100) in blocking solution (alternatively, 3% of Bovine Serum Albumin and Triton X-100 0.1% in PBS solution can be used). Prepare 30 &#181;L of antibody solution per well if using coverslips or 20 &#181;L if using a 384-well plate.</li>
		<li>Tap off the 0.01x SSC solution. Wash 2x with PBS.</li>
		<li>Add the primary antibody solution to each well. Incubate at room temperature for 1 h.</li>
	</ol>
	</li>
	<li>Secondary antibody incubation (IF) 
	NOTE: Make sure the chosen secondary antibodies do not recognize the PLA antibodies or probes. Additionally, the wavelength of the secondary antibody must be different from the PLA detection reagent wavelength. For example: our PLA detection reagent contains the fluorophore 647, and the secondary we used for &#947;-H2AX detection is 488.
	<ol>
		<li>Dilute the secondary antibodies (Donkey anti-Rabbit Alexa Fluor 488) in blocking solution (alternatively, 3% of Bovine Serum Albumin and Triton X - 100 0.1% in PBS solution can be used) at 1:300 dilution. Prepare 30 &#181;L of antibody solution per condition if using coverslips or 20 &#181;L per well if using a 384-well plate. Add Hoechst 33342 dye at 0.6 &#181;g/mL final concentration.</li>
		<li>Tap off the primary antibody solution. Wash 3x with TBS-T.</li>
		<li>Add the secondary antibody solution to each well. Incubate at room temperature for 20 min.</li>
		<li>Wash 5x with TBS-T. Wash 2x with 1x SSC buffer. Wash 2x with 0.01x SSC buffer. 
		NOTE: At this point, the plates can be stored for several weeks at 4 &#176;C (in SSC 0.01X) protected from light. If using coverslips, the slides can be mounted, and after 1 day, the slides can be stored at -20 &#176;C.</li>
	</ol>
	</li>
</ol>
10. Image acquisition
<ol>
	<li>Use a conventional fluorescence microscope and the 63x objective. Determine the time of exposition by comparing negative and positive controls to avoid saturation and dots collapsing. 
	NOTE: The use of confocal microscopes enhances the PLA dots detection. The 20x objective also can be used for increasing the number of nuclei detected.</li>
	<li>Once the exposition time is adjusted, perform all acquisitions of PLA dots with the same parameters. Try to acquire images from different wells or coverslips regions to avoid artifacts due to non-homogeneous incubation.</li>
	<li>Save all the channels with the same name pattern. If the microscope takes photos in the channels separately, save the corresponding images with the same name, adding only PLA or Nucleus at the beginning of the file name to differentiate the channel.</li>
</ol>
11. Image analysis
NOTE: A&#160;macro was created for analysis of dots per nucleus in Image J/FIJI17 and to be used with microscopes where the scale information must be added manually. The pixel size for the objective lens used must be known. Place the images for each channel in different folders (Supplementary Coding File 1, Supplementary Figure 1). 
NOTE: The macro considers fluorescence channels obtained separately. In this case, it is important to save all the images with the same name.
<ol>
	<li>The macro allows the selection of a specific region in the field. Use the option Select a ROI when a region in the image shows artifacts that can be detrimental to analysis. Choose ROI at the beginning of the analysis and use it for all the images. 
	NOTE: This is an optional feature. If not selected, the entire image will be considered in the analysis (Supplementary Figure 2).</li>
	<li>The ROI file should be created prior to the macro usage, as described below. For ROI selection, use a rectangle tool, select the region for analysis, and add it to the ROI manager by clicking Edit &#62; Selection&#62; Add to Manager. Save the ROI by clicking More&#62; Save.</li>
	<li>In the ImageJ 17, open images from different conditions to adjust the parameters for analysis. These parameters must be determined by the user and specified, along with the data, in the macro program (Supplementary Coding File 1)
	<ol>
		<li>Background correction: To remove the background click Process&#62; Subtract Background. Test different rolling ball radius using the preview function). We have used rolling 10 for PLA dots and rolling 100 for nucleus.</li>
		<li>Remove noise by clicking Process &#62; Noise &#62; Despeckle. For nucleus, use the function smooth to improve the filling of the nucleus by clicking Process&#62;Smooth.</li>
		<li>Adjusting threshold: Convert to 8-bit image and adjust the threshold by clicking Image &#62; Type &#62; 8 bit and Image &#62; Adjust &#62;Threshold. Adjust the threshold accordingly to visualize all the nucleus or dots present in the image but avoiding over saturation and structures collapsing. Take notes of the values and test in different images to check if the threshold works for different images.</li>
		<li>Convert to mask&#160;by clicking Process &#62; Binary &#62; Convert to mask. In the case of the nucleus, click Process &#62; Binary &#62; Fill holes followed by Process &#62; Binary &#62; Watershed. In the case of the PLA dots, click Process &#62; Binary &#62; Watershed.</li>
		<li>For counting nuclei, measure the area or length of the different nuclei to estimate the average size. Use an approximate value to analyze particles. Click Analyze &#62; Analyze particles, then select Set size: 15-infinity (these values depend on the size of the nuclei). Check the following options: Show: Mask; Display results; Clear results; Summarize; Add to Manager; Exclude on edges.</li>
		<li>For counting PLA dots, measure the area or the radius of the dots to estimate the average size.&#160;Select Apply Nucleus Mask (apply mask is a function on GDSC plugin, check if it is installed) followed by Analyze &#62; Analyze particles&#160;and set. 0.02-3 (these values depend on the size of the nuclei). Check the following options: Show: Mask; Display results; Clear results; Summarize; Add to Manager; Exclude on edges.</li>
	</ol>
	</li>
	<li>The results show the number of dots per nucleus in each nucleus present in the image. Save this data.</li>
</ol>
12. Visualizing the merged images
<ol>
	<li>Open the channels corresponding to PLA, nucleus, and &#947;-H2AX. Subtract the background and remove noise, as mentioned in steps 11.4.1 and 11.4.2 for all images.</li>
	<li>Adjust brightness and contrast automatically or manually using the same value for all conditions from the same channel.&#160;Select Image &#62; Adjust &#62; Bright/contrast.</li>
	<li>Combine images (merging)by selecting Image &#62; Color &#62; Merge channels. Add scale bar by selecting Analyze &#62; Tools &#62; Scale bar. Save as.TIFF file.</li>
</ol>

Assessing Protein Interaction and Localization in DNA Damage

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The authors declare that they have no competing financial interests.

The data show that the use of PLA concomitantly to a DNA-damaged marker can provide the most information in a DNA damage response profiling, showing the spatial and temporal behavior of the interaction after the insult. PLA is a versatile method that has been used for dimerization identification, organelles contact determination, protein-nucleic acid interaction, and mainly protein-protein interaction10,11,19,<sup cla...

Cellular DNA damage occurs daily because of the spontaneous chemical reactions and is also increased by exogenous factors such as genotoxic agents (radiation and chemicals such as etoposide) and oxidative stress1,2,3. The cells have a complex machinery that corrects a myriad of different types of DNA damage, from removal of bases to replicative fork torsion or interruption to the most deleterious lesion: the DNA double-strand break4,5.
Several proteins that participate in the DDR have already been characterized, and excellent revisions explore the pathways of non-homologous end join (NHEJ) and homologous recombination (HR), the main pathways implied in double-strand breaks (DSB) repair5,6. The phosphorylation of the histone H2A variant, H2AX, at serine 139 (thereafter named y-H2AX) has been used for many years as a sensitive and reliable marker of the double-strand break. The phosphorylation of H2AX is observed in the first minutes following the damage and can persist for hours7.
The advantage of using the immunofluorescence detection of &#947;-H2AX foci is the characterization of the temporal and spatial distribution of the foci, as well as its co-staining with other proteins. Moreover, immunofluorescence does not require lysis, which preserves cellular context information and makes it more informative. Besides, as &#947;-H2AX is formed de novo, it is not abundantly present in the absence of DNA damage, making it a specific marker7.
H2AX is mostly phosphorylated by protein kinase ataxia-telangiectasia mutated (ATM) and DNA-dependent protein kinase (DNA-PK), the sensors of DSBs. Ku70/80 (XRCC6 X-ray repair cross-complementing 6/ XRCC5 X-ray repair cross-complementing 5) and DNA-protein kinase catalytic subunit (DNA-PKcs) are responsible for DSB identification and the protection of DNA ends, whereas Artemis carries out end-processing to facilitate ligation by the XRCC4-Ligase IV complex. The phosphorylation of H2AX at sites flanking DSBs acts as a signal for the recruitment of several repair proteins and signaling for cell cycle arrest, death, or survival8.
The analysis of changes in y-H2AX foci is the most common assay for studying if the protein of interest is implicated in DNA damage response. Whether a direct role in the DNA damage site is expected, fluorescence microscopy is used to verify the co-localization of the protein of interest with y-H2AX foci. However, except for the new super-resolution fluorescence methods, concluding the local interaction with DNA damage sites can be tricky. Furthermore, immunofluorescence detection usually depends on many protein molecules at the DNA damage site, making it difficult to identify scarce proteins9.
Here, we show an assay to evaluate the localization of proteins involved in the DDR pathway using y-H2AX as a marker of the damage site. This assay can be used to characterize temporal localization under different insults that cause DNA damage.
The Proximity Ligand Assay (PLA) emerged as a tool for estimating interaction between proteins as well as spatial proximity among organelles or cellular structures10,11and allows the temporal localization and co-localization analysis under stress conditions, for instance. The method is simple because it is like conventional immunofluorescence and allows the simultaneous staining of organelle or cellular structure-specific markers, such as mitochondria, endoplasmic reticulum, PML bodies, or DNA double-strand marker, yH2AX. The use of PLA allows the identification of protein interaction during DNA damage in a sensitive, specific, and reliable manner that can be used to monitor the interaction during the cell cycle, in response to different stimuli, and at different times after genotoxic stress.
We show here the use of PLA concomitantly to conventional &#947;-H2AX immunofluorescence to localize Nek4-Ku70 interaction after etoposide-induced DNA damage. Nek4, a Nek (NIMA-related kinases) family member, has no clear role in DNA Damage response. In 2012, Nguyen and colleagues showed that Nek4 interacts with Ku70/Ku80 proteins, and in the Nek4 absence, these proteins are not recruited to the DNA damage site, and H2AX phosphorylation is impaired12. The cellular localization of this interaction is unknown so far. We have observed a reduction in Ku70 phosphorylation in cells expressing Nek4 kinase-dead mutant13but whether Nek4 phosphorylates directly Ku70 remains to be elucidated. Considering that Nek4 interaction with Ku70 is increased after etoposide treatment according to immunoprecipitation studies12, we attempt to localize this interaction using PLA and the &#947;-H2AX marker.
Usually, the interaction studies are based on immunoprecipitation assays, but these are in vitro and do not provide information about the location of the interaction. When possible, an immunoprecipitation of fractioned cells (nucleus, mitochondrial, cellular membrane, or cytosol) can be performed, but performing a time course of the interaction is costly and laborious. Immunofluorescence can give information about the cellular localization of the proteins, but proving the interaction can be difficult and depends on the resolution of the microscopes. Here, we show the advantage of using the Proximity Ligand Assay to monitor the interaction of two proteins in a DNA damage context.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Black 384-well plates</td>
			<td>Perkin Elmer Cell carrier plates</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey&#160; anti- Rabbit Alexa Fluor 488</td>
			<td>Invitrogen</td>
			<td>A21206</td>
			<td>1:300</td>
		</tr>
		<tr>
			<td>Duo link Donkey anti Mouse Minus</td>
			<td>Sigma</td>
			<td>DUO92004</td>
			<td></td>
		</tr>
		<tr>
			<td>Duolink antibody diluent</td>
			<td>Sigma</td>
			<td>DUO82008</td>
			<td></td>
		</tr>
		<tr>
			<td>Duolink blocking solution 1X</td>
			<td>Sigma</td>
			<td>DUO82007</td>
			<td></td>
		</tr>
		<tr>
			<td>Duolink Detection reagent Far red</td>
			<td>Sigma</td>
			<td>DUO92013</td>
			<td></td>
		</tr>
		<tr>
			<td>Duolink Donkey anti goat plus</td>
			<td>Sigma</td>
			<td>DUO92003</td>
			<td></td>
		</tr>
		<tr>
			<td>Duolink Donkey anti rabbit plus</td>
			<td>Sigma</td>
			<td>DUO92002</td>
			<td></td>
		</tr>
		<tr>
			<td>Etoposide</td>
			<td>Sigma</td>
			<td>E1383</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti Nek4</td>
			<td>Santa Cruz Biotechnology</td>
			<td>SC-5517</td>
			<td>goat anti Nek4 was used at 1:50 dilution</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Thermo</td>
			<td>H1399</td>
			<td>0.6 &#181;g/mL</td>
		</tr>
		<tr>
			<td>Leica DMI microscope</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti Ku70</td>
			<td>Thermo</td>
			<td>MA5-13110</td>
			<td>mouse anti Ku70 was used at 1:100 dilution</td>
		</tr>
		<tr>
			<td>Mouse anti TOPII&#946;</td>
			<td>Santa Cruz Biotechnology</td>
			<td>SC-365071</td>
			<td>1:25</td>
		</tr>
		<tr>
			<td>Rabbit anti Nek5</td>
			<td>Santa Cruz Biotechnology</td>
			<td>SC-84527</td>
			<td>1:25</td>
		</tr>
		<tr>
			<td>Rabbit anti Y H2AX</td>
			<td>Cell Signalling</td>
			<td>9718S</td>
			<td>1:100 dilution</td>
		</tr>
		<tr>
			<td>U2OS cell line</td>
			<td>ATCC</td>
			<td>HTB-96&#160;</td>
			<td></td>
		</tr>
	</tbody>
</table>

proximity ligand assay to localize proteins in dna damage sites

The DNA damage response is a genetic information safeguard that protects cells from perpetuating damaged DNA. The characterization of the proteins that cooperate in this process allows the identification of alternative targets for therapeutic intervention in several diseases, such as cancer, aging-related diseases, and chronic inflammation. The Proximity Ligand Assay (PLA) emerged as a tool for estimating interaction between proteins as well as spatial proximity among organelles or cellular structures and allows the temporal localization and co-localization analysis under stress conditions, for instance. The method is simple because it is similar to conventional immunofluorescence and allows the staining of an organelle, cellular structure, or a specific marker such as mitochondria, endoplasmic reticulum, PML bodies, or DNA double-strand marker, yH2AX simultaneously. The phosphorylation of the S139 at Histone 2A variant, H2AX, then referred to as yH2AX, is widely used as a very sensitive and specific marker of DNA double-strand breaks. Each focus of yH2AX staining corresponds to one break in DNA that occurs a few minutes after the damage. The analysis of changes in yH2AX foci is the most common assay for studying if the protein of interest is implicated in DNA damage response (DDR). Whether a direct role in the DNA damage site is expected, fluorescence microscopy is used to verify the colocalization of the protein of interest with yH2AX foci. However, except for the new super-resolution fluorescence methods, to conclude, the local interaction with DNA damage sites can be a little subjective. Here, we show an assay to evaluate the localization of proteins in the DDR pathway using yH2AX as a marker of the damage site. This assay can be used to characterize the temporal localization under different insults that cause DNA damage.

We have observed Nek4-Ku70 interaction in the absence of etoposide treatment. However, this interaction can occur outside of the nucleus (Figure 1A). The Nek4-Ku70 interaction increases after DNA damage and is concentrated in the nucleus (Figure 1A). In the case of Nek5-Topoisomerase II &#946; (TOPII&#946;) interaction, used in this assay as a positive control based on literature results18, the interaction is greatly increased by etoposid...

Author Spotlight: Combining Proximity Ligand Assay with Gamma-H2AX Staining to Characterize Protein Interactions in DNA Damage Response

Here, we present a simple and rapid protocol for the detection of protein interaction at DNA damage sites.

Proximity Ligand Assay to Localize Proteins in DNA Damage Sites

The DNA damage response is a genetic information safeguard that protects cells from perpetuating damaged DNA. The characterization of the proteins that ...

proximity-ligand-assay-to-localize-proteins-in-dna-damage-sites

Research

JoVE Journal

Biology

353 Views.  University of Campinas. Here, we present a simple and rapid protocol for the detection of protein interaction at DNA damage sites.

Video: Author Spotlight: Combining Proximity Ligand Assay with Gamma-H2AX Staining to Characterize Protein Interactions in DNA Damage Response

Proteomics to Identify Proteins Interacting with P2X2 Ligand-Gated Cation Channels

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys loop, the glutamate-gated and the P2X receptor channels2. In each case binding of transmitter leads to the opening of a pore through which ions flow down their electrochemical gradients. Many ligand-gated channels are also permeable to calcium ions3, 4, which have downstream signaling roles5 (e.g. gene regulation) that may exceed the duration of channel opening. Thus ligand-gated channels can signal over broad time scales ranging from a few milliseconds to days. Given these important roles it is necessary to understand how ligand-gated ion channels themselves are regulated by proteins, and how these proteins may tune signaling. Recent studies suggest that many, if not all, channels may be part of protein signaling complexes6. In this article we explain how to identify the proteins that bind to the C-terminal aspects of the P2X2 receptor cytosolic domain.
P2X receptors are ATP-gated cation channels and consist of seven subunits (P2X1-P2X7). P2X receptors are widely expressed in the brain, where they mediate excitatory synaptic transmission and presynaptic facilitation of neurotransmitter release7. P2X receptors are found in excitable and non-excitable cells and mediate key roles in neuronal signaling, inflammation and cardiovascular function8. P2X2 receptors are abundant in the nervous system9 and are the focus of this study. Each P2X subunit is thought to possess two membrane spanning segments (TM1 &amp; TM2) separated by an extracellular region7 and intracellular N and C termini (Fig 1a)7. P2X subunits10 (P2X1-P2X7) show 30-50% sequence homology at the amino acid level11. P2X receptors contain only three subunits, which is the simplest stoichiometry among ionotropic receptors. The P2X2 C-terminus consists of 120 amino acids (Fig 1b) and contains several protein docking consensus sites, supporting the hypothesis that P2X2 receptor may be part of signaling complexes. However, although several functions have been attributed to the C-terminus of P2X2 receptors9 no study has described the molecular partners that couple to the intracellular side of this protein via the full length C-terminus. In this methods paper we describe a proteomic approach to identify the proteins which interact with the full length C-terminus of P2X2 receptors.

We describe a simple protocol to identify brain proteins that bind to the full length C terminus of ATP-gated P2X2 receptors. The extension and systematic application of this approach to all P2X receptors is expected to lead to a better understanding of P2X receptor signaling.

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys ...

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution of existing proteins, and the assembly of new protein complexes can be stimulated by a variety of DNA lesions and mismatched DNA base pairs. Fluorescence microscopy has been used as a powerful experimental tool for visualizing and quantifying these and other responses to DNA lesions and to monitor DNA replication status within the complex subcellular architecture of a living cell. Translational fusions between fluorescent reporter proteins and components of the DNA replication and repair machinery have been used to determine the cues that target DNA repair proteins to their cognate lesions in vivo and to understand how these proteins are organized within bacterial cells. In addition, transcriptional and translational fusions linked to DNA damage inducible promoters have revealed which cells within a population have activated genotoxic stress responses. In this review, we provide a detailed protocol for using fluorescence microscopy to image the assembly of DNA repair and DNA replication complexes in single bacterial cells. In particular, this work focuses on imaging mismatch repair proteins, homologous recombination, DNA replication and an SOS-inducible protein in Bacillus subtilis. All of the procedures described here are easily amenable for imaging protein complexes in a variety of bacterial species.

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

A detailed protocol is described for imaging the real time formation of DNA repair complexes in Bacillus subtilis cells.

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution ...

PAR-CliP - A Method to Identify Transcriptome-wide the Binding Sites of RNA Binding Proteins

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ribonucleoprotein complexes (miRNPs) that are often expressed in a cell-type dependently. To understand how the interplay of these RNA-binding factors affects the regulation of individual transcripts, high resolution maps of in vivo protein-RNA interactions are necessary1.
A combination of genetic, biochemical and computational approaches are typically applied to identify RNA-RBP or RNA-RNP interactions. Microarray profiling of RNAs associated with immunopurified RBPs (RIP-Chip)2 defines targets at a transcriptome level, but its application is limited to the characterization of kinetically stable interactions and only in rare cases3,4 allows to identify the RBP recognition element (RRE) within the long target RNA. More direct RBP target site information is obtained by combining in vivo UV crosslinking5,6 with immunoprecipitation7-9 followed by the isolation of crosslinked RNA segments and cDNA sequencing (CLIP)10. CLIP was used to identify targets of a number of RBPs11-17. However, CLIP is limited by the low efficiency of UV 254 nm RNA-protein crosslinking, and the location of the crosslink is not readily identifiable within the sequenced crosslinked fragments, making it difficult to separate UV-crosslinked target RNA segments from background non-crosslinked RNA fragments also present in the sample.
We developed a powerful cell-based crosslinking approach to determine at high resolution and transcriptome-wide the binding sites of cellular RBPs and miRNPs that we term PAR-CliP (Photoactivatable-Ribonucleoside-Enhanced Crosslinking and Immunoprecipitation) (see Fig. 1A for an outline of the method). The method relies on the incorporation of photoreactive ribonucleoside analogs, such as 4-thiouridine (4-SU) and 6-thioguanosine (6-SG) into nascent RNA transcripts by living cells. Irradiation of the cells by UV light of 365 nm induces efficient crosslinking of photoreactive nucleoside-labeled cellular RNAs to interacting RBPs. Immunoprecipitation of the RBP of interest is followed by isolation of the crosslinked and coimmunoprecipitated RNA. The isolated RNA is converted into a cDNA library and deep sequenced using Solexa technology. One characteristic feature of cDNA libraries prepared by PAR-CliP is that the precise position of crosslinking can be identified by mutations residing in the sequenced cDNA. When using 4-SU, crosslinked sequences thymidine to cytidine transition, whereas using 6-SG results in guanosine to adenosine mutations. The presence of the mutations in crosslinked sequences makes it possible to separate them from the background of sequences derived from abundant cellular RNAs.
Application of the method to a number of diverse RNA binding proteins was reported in Hafner et al.18

RNA transcripts are subject to extensive posttranscriptional regulation that is mediated by a multitude of trans-acting RNA-binding proteins (RBPs). Here we present a generalizable method to identify precisely and on a transcriptome-wide scale the RNA binding sites of RBPs.

RNA transcripts are subjected to post-transcriptional gene regulation by interacting with hundreds of RNA-binding proteins (RBPs) and microRNA-containing ...

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the messagefor review see 1. However, it's now widely accepted that synonymous codon usage is the primary cause of non-uniform translational elongation rates1. Synonymous codons are not used with identical frequency. A bias exists in the use of synonymous codons with some codons used more frequently than others2. Codon bias is organism as well as tissue specific2,3. Moreover, frequency of codon usage is directly proportional to the concentrations of cognate tRNAs4. Thus, a frequently used codon will have higher multitude of corresponding tRNAs, which further implies that a frequent codon will be translated faster than an infrequent one. Thus, regions on mRNA enriched in rare codons (potential pause sites) will as a rule slow down ribosome movement on the message and cause accumulation of nascent peptides of the respective sizes5-8. These pause sites can have functional impact on the protein expression, mRNA stability and protein foldingfor review see 9. Indeed, it was shown that alleviation of such pause sites can alter ribosome movement on mRNA and subsequently may affect the efficiency of co-translational (in vivo) protein folding1,7,10,11. To understand the process of protein folding in vivo, in the cell, that is ultimately coupled to the process of protein synthesis it is essential to gain comprehensive insights into the impact of codon usage/tRNA content on the movement of ribosomes along mRNA during translational elongation.
Here we describe a simple technique that can be used to locate major translation pause sites for a given mRNA translated in various cell-free systems6-8. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA. The rationale is that at low-frequency codons, the increase in the residence time of the ribosomes results in increased amounts of nascent peptides of the corresponding sizes. In vitro transcribed mRNA is used for in vitro translational reactions in the presence of radioactively labeled amino acids to allow the detection of the nascent chains. In order to isolate ribosome bound nascent polypeptide complexes the translation reaction is layered on top of 30% glycerol solution followed by centrifugation. Nascent polypeptides in polysomal pellet are further treated with ribonuclease A and resolved by SDS PAGE. This technique can be potentially used for any protein and allows analysis of ribosome movement along mRNA and the detection of the major pause sites. Additionally, this protocol can be adapted to study factors and conditions that can alter ribosome movement and thus potentially can also alter the function/conformation of the protein.

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

A technique to identify translational pause sites on mRNA is described. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA, followed by the size analysis of the nascent chains using a denaturing gel electrophoresis.

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage checkpoint signaling as a surveillance mechanism to sense DNA damage and direct cellular responses to DNA damage. There are several groups of proteins called sensors, transducers and effectors involved in DNA damage checkpoint signaling (Figure 1). In this complex signaling pathway, ATR (ATM and Rad3-related) is one of the major kinases that can respond to DNA damage and replication stress. Activated ATR can phosphorylate its downstream substrates such as Chk1 (Checkpoint kinase 1). Consequently, phosphorylated and activated Chk1 leads to many downstream effects in the DNA damage checkpoint including cell cycle arrest, transcription activation, DNA damage repair, and apoptosis or senescence (Figure 1). When DNA is damaged, failing to activate the DNA damage checkpoint results in unrepaired damage and, subsequently, genomic instability. The study of the DNA damage checkpoint will elucidate how cells maintain genomic integrity and provide a better understanding of how human diseases, such as cancer, develop.
Xenopus laevis egg extracts are emerging as a powerful cell-free extract model system in DNA damage checkpoint research. Low-speed extract (LSE) was initially described by the Masui group1. The addition of demembranated sperm chromatin to LSE results in nuclei formation where DNA is replicated in a semiconservative fashion once per cell cycle.
The ATR/Chk1-mediated checkpoint signaling pathway is triggered by DNA damage or replication stress 2. Two methods are currently used to induce the DNA damage checkpoint: DNA damaging approaches and DNA damage-mimicking structures 3. DNA damage can be induced by ultraviolet (UV) irradiation, &gamma;-irradiation, methyl methanesulfonate (MMS), mitomycin C (MMC), 4-nitroquinoline-1-oxide (4-NQO), or aphidicolin3, 4. MMS is an alkylating agent that inhibits DNA replication and activates the ATR/Chk1-mediated DNA damage checkpoint 4-7. UV irradiation also triggers the ATR/Chk1-dependent DNA damage checkpoint 8. The DNA damage-mimicking structure AT70 is an annealed complex of two oligonucleotides poly-(dA)70 and poly-(dT)70. The AT70 system was developed in Bill Dunphy's laboratory and is widely used to induce ATR/Chk1 checkpoint signaling 9-12.
Here, we describe protocols (1) to prepare cell-free egg extracts (LSE), (2) to treat Xenopus sperm chromatin with two different DNA damaging approaches (MMS and UV), (3) to prepare the DNA damage-mimicking structure AT70, and (4) to trigger the ATR/Chk1-mediated DNA damage checkpoint in LSE with damaged sperm chromatin or a DNA damage-mimicking structure.

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

Xenopus egg extract is a useful model system to investigate the DNA damage checkpoint. This protocol is for the preparation of Xenopus egg extracts and DNA damage checkpoint inducing reagents. These techniques are adaptable to a variety of DNA damaging approaches in the study of the DNA damage checkpoint signaling.

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage ...

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently attach a tagged form of dUTP to 3&#8217; ends of double- and single-stranded DNA breaks in cells. It is a reliable and useful method to detect DNA damage and cell death in situ. This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method of semi-automated TUNEL signal quantitation. Inherent normal tissue features and tissue processing conditions affect the ability of the TdT enzyme to efficiently label DNA. Tissue processing may also add undesirable autofluorescence that will interfere with TUNEL signal detection. Therefore, it is important to empirically determine tissue processing and TUNEL labeling methods that will yield the optimal signal-to-noise ratio for subsequent quantitation. The fluorescence-based assay described here provides a way to exclude autofluorescent signal by digital channel subtraction. The TUNEL assay, used with appropriate tissue processing techniques and controls, is a relatively fast, reproducible, quantitative method for detecting apoptosis in tissue. It can be used to confirm DNA damage and apoptosis as pathological mechanisms, to identify affected cell types, and to assess the efficacy of therapeutic treatments in vivo.

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method for semi-automated analysis of TUNEL labeling.

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently ...

In Vivo Alkaline Comet Assay and Enzyme-modified Alkaline Comet Assay for Measuring DNA Strand Breaks and Oxidative DNA Damage in Rat Liver

Unrepaired DNA damage can lead to genetic instability, which in turn may enhance cancer development. Therefore, identifying potential DNA damaging agents is important for protecting public health. The in vivo alkaline comet assay, which detects DNA damage as strand breaks, is especially relevant for assessing the genotoxic hazards of xenobiotics, as its responses reflect the in vivo absorption, tissue distribution, metabolism and excretion (ADME) of chemicals, as well as DNA repair process. Compared to other in vivo DNA damage assays, the assay is rapid, sensitive, visual and inexpensive, and, by converting oxidative DNA damage into strand breaks using specific repair enzymes, the assay can measure oxidative DNA damage in an efficient and relatively artifact-free manner. Measurement of DNA damage with the comet assay can be performed using both acute and subchronic toxicology study designs, and by integrating the comet assay with other toxicological assessments, the assay addresses animal welfare requirements by making maximum use of animal resources. Another major advantage of the assays is that they only require a small amount of cells, and the cells do not have to be derived from proliferating cell populations. The assays also can be performed with a variety of human samples obtained from clinically or occupationally exposed individuals.

In Vivo Alkaline Comet Assay and Enzyme-modified Alkaline Comet Assay for Measuring DNA Strand Breaks and Oxidative DNA Damage in Rat Liver

The alkaline comet assay measures DNA strand breaks in eukaryotic cells. By adding an Endonuclease III or human 8-oxoguanine-DNA-N-glycosylase digestion step, the assay can efficiently detect oxidative DNA damage. We describe methods for using these assays to detect DNA damage in rat liver.

Unrepaired DNA damage can lead to genetic instability, which in turn may enhance cancer development. Therefore, identifying potential DNA damaging agents ...

Amplification, Next-generation Sequencing, and Genomic DNA Mapping of Retroviral Integration Sites

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of diverse libraries of retroviral integration sites using ligation-mediated PCR (LM-PCR) amplification and next-generation sequencing (NGS), (2) mapping the genomic location of each virus-host junction using BEDTools, and (3) analyzing the data for statistical relevance. Genomic DNA extracted from infected cells is fragmented by digestion with restriction enzymes or by sonication. After suitable DNA end-repair, double-stranded linkers are ligated onto the DNA ends, and semi-nested PCR is conducted using primers complementary to both the long terminal repeat (LTR) end of the virus and the ligated linker DNA. The PCR primers carry sequences required for DNA clustering during NGS, negating the requirement for separate adapter ligation. Quality control (QC) is conducted to assess DNA fragment size distribution and adapter DNA incorporation prior to NGS. Sequence output files are filtered for LTR-containing reads, and the sequences defining the LTR and the linker are cropped away. Trimmed host cell sequences are mapped to a reference genome using BLAT and are filtered for minimally 97% identity to a unique point in the reference genome. Unique integration sites are scrutinized for adjacent nucleotide (nt) sequence and distribution relative to various genomic features. Using this protocol, integration site libraries of high complexity can be constructed from genomic DNA in three days. The entire protocol that encompasses exogenous viral infection of susceptible tissue culture cells to integration site analysis can therefore be conducted in approximately one to two weeks. Recent applications of this technology pertain to longitudinal analysis of integration sites from HIV-infected patients.

We describe a protocol for amplifying retroviral integration sites from the genomic DNA of infected cells, sequencing the amplified virus-host junctions, and then mapping these sequences to a reference genome. We also describe techniques to quantify the distribution of integration sites relative to various genomic annotations using BEDTools.

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of ...

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.