We thank Joel Sanneman and Dr. Philine Wangemann of the Confocal Microscopy Core, funded by the Kansas State University College of Veterinary Medicine, for their support of efforts to develop this technique. pCBASceI was a gift from Maria Jasin (Addgene plasmid # 26477)30. U2OS DR-GFP cells were a kind gift from Maria Jasin18.

Please note, this protocol is written for U2OS cells containing an I-SceI recognition site18. The cells need not be U2OS but must contain the I-SceI site. The protocol may need to be adjusted (e.g., number of cells seeded and incubation times) depending on the type of cells used.
1. Defining the Kinetics of DSB Repair Complex Formation
<ol>
	<li>Grow U20S-DRGFP cells on a 10 cm tissue culture plate until 85&#8211;90% confluent.</li>
	<li>Remove the media and incubate at room temperature (RT) in 3 mL of EDTA for 2 min.</li>
	<li>Replace EDTA with 1 mL of trypsin and incubate at 37 &#176;C for 5 min. Ensure that the cells are detached from the surface of the plate by viewing under a microscope. Neutralize trypsin with 5 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin.</li>
	<li>Determine the concentration of cells in this suspension using a hemocytometer and the trypan blue exclusion method.</li>
	<li>Seed 6 x 103 cells/well in 200 &#181;L of a glass-bottom 96-well plate. Alternatively, seed 4 x 106 cells in 8 mL on 12 mm coverslips placed in a 10 cm plate. 
	NOTE: Each well or coverslip represents a single time point, including the control.</li>
	<li>Grow the cells overnight at 37 &#176;C and 5% CO2.</li>
	<li>Inducing DSBs
	<ol>
		<li>Replace the media from the 96-well plate/10 cm dish with H2O2 diluted to a concentration of 25 &#181;M with DMEM + 10% FBS and incubate at 37 &#176;C for 1 h.</li>
		<li>Wash the cells 3x with 1x PBS and replace with fresh DMEM supplemented with 10% FBS and 1% penicillin/streptomycin. Incubate the cells at 37 &#176;C for 0, 1, 4, 8, 24, and 48 h. Ensure that fresh dilutions of H2O2 are used for each experiment. 
		NOTE: To avoid killing cells at the H2O2 concentration used here, ensure that the dose of damage is low enough such that the bulk of cells will avoid apoptosis and senescence. It is critical to observe the recovery process. This is best achieved by measuring sensitivity to a range of doses. Perform an MTT assay or other cell viability assay to determine the minimal concentration of hydrogen peroxide that can be used.</li>
	</ol>
	</li>
	<li>Fixing and permeabilizing cells
	<ol>
		<li>Wash the cells 3 times with 1x PBS. Remove a coverslip from the 10 cm plate for each time point and place it in a single well of a 24-well plate for fixing. Use the needle and forceps to carefully remove the coverslip and avoiding scratching cells off the coverslip surface.</li>
		<li>Fix the cells by incubating for 15 min with 4% paraformaldehyde (PFA) at designated times after 1 h of H2O2 exposure and fresh DMEM replacement (0, 1, 4, 8, 24, and 48 h).</li>
		<li>Wash the fixed coverslips 3 times with PBS. Permeabilize the cells with 0.5% non-ionic detergent diluted in 1x PBS. Incubate for 15 min at RT, before washing 3x with PBS. 
		NOTE: After permeabilization, storage of coverslips is possible in PBS at 4 &#176;C for at least a week.</li>
	</ol>
	</li>
	<li>Repair complex visualization
	<ol>
		<li>Block the 96-well plate/coverslips in a solution of 3% bovine serum albumin (BSA) and 0.1% non-ionic detergent diluted in 1x PBS (3% BSA solution) for 1 h at RT.</li>
		<li>Incubate with primary antibodies that target the repair protein of interest, diluted according to the manufacturer&#39;s specifications in 3% BSA solution for 1 h at RT, before washing 3 times with 1x PBS.</li>
		<li>Incubate the 96-well plate/coverslips with secondary antibody diluted according to the manufacturer&#8217;s protocol in 3% BSA solution. Confirm that the secondary fluorophores do not have overlapping excitation or emission spectra.</li>
		<li>After washing the cells with PBS 3 times add DAPI, diluted according to the manufacturer&#8217;s specifications in 1x PBS and incubate at RT for 5 min.</li>
		<li>Mount the coverslips onto slides with a drop of mounting agent, making sure the cell-side faces down. Press the coverslips carefully and soak up any excess mounting agent with a wipe.</li>
		<li>Seal the coverslips with fast-drying transparent nail polish and ensure a complete seal of the coverslip with the slide.</li>
		<li>Take the confocal microscope images by focusing on the DAPI channel, before other channels (with signals from DNA repair gene of interest) to avoid introducing bias into the experiment.</li>
		<li>Alternatively, images can be acquired by taking Z-sections of the desired region and condensing the image into a single plane to visualize all detectable foci. 
		NOTE: If DSB resolution or protein of interest is not observed at time points chosen, determine when DSBs resolve by selecting a wide range of time points initially (0&#8211;48 h post DSB induction). The time points of interest will vary by the method of DSB induction and the repair protein of interest.</li>
	</ol>
	</li>
	<li>Use free ImageJ software to define the nuclei in the immunofluorescence microscopy images. 
	NOTE: To open microscope images, the Bio-Formats plugin must be installed in ImageJ.
	<ol>
		<li>Open the confocal images (.czi or .tif) by dragging the image file into the ImageJ window, or by selecting &#34;Bio-Formats Importer&#34; from the &#34;Plugin&#34; toolbar in ImageJ. 
		NOTE: The &#34;Bio-Formats Import Option&#34; window will appear.</li>
		<li>Select &#34;Split Channels&#34; to split the image into constitutive DAPI and fluorescent images within the &#34;Bio-Formats Import Option&#34; window.</li>
		<li>Select &#34;Colorized&#34; in &#34;Color options&#34; from the drop-down menu. Select &#34;OK&#34; to open the DAPI and Histone H2AX (H2AX) images as separate windows and choose the DAPI image.</li>
		<li>Select the &#34;Image&#34; toolbar and &#34;Adjust Threshold&#34; option to select nuclei. 
		NOTE: A &#34;Threshold&#34; window will appear.</li>
		<li>Adjust the image threshold by sliding the bottom bar of the &#34;Threshold&#34; window completely to the right. Adjust the top bar until the nuclei appear completely red with distinct outlines, and the background black. DO NOT select apply. Close the &#34;Adjust Threshold&#34; window.</li>
		<li>Select the &#34;Analyze&#34; toolbar and &#34;Analyze Particles&#34; option. See an &#34;Analyze Particles&#34; window appearing on the screen.</li>
		<li>Input the minimum size of a nucleus. 
		NOTE: Particles below the size inputted will not be counted as nuclei.</li>
		<li>From the &#34;Show&#34; dropdown menu, select &#34;Outlines&#34;. Select the &#34;Add to Manager&#34; option. Click &#34;OK&#34; to open an &#34;ROI Manager&#34; window. 
		NOTE: Outlines of nuclei will appear in a separate window.</li>
		<li>Assign numbers to the nuclei that can be selected from the &#34;ROI Manager&#34; window.</li>
	</ol>
	</li>
	<li>Use ImageJ to quantify H2AX foci in immunofluorescence microscopy images.
	<ol>
		<li>Select the &#34;H2AX&#34; foci window (stained with fluorescently conjugated secondary antibody). Then, select the &#34;Process&#34; toolbar and choose &#34;Find Maxima&#34; to find the foci within the nuclei. 
		NOTE: A &#34;Find Maxima&#34; window will open.</li>
		<li>Choose the &#34;Single Points&#34; option from the &#34;Output Type&#34; dropdown menu and select &#34;Preview point selection&#34; to visualize the maxima/foci. Input a &#34;Noise Tolerance&#34; value depending on the fluorescence intensity of the image (Figure 1). Check for the appearance of a white window with small black dots. 
		NOTE: These dots represent the H2AX foci/maxima that have been detected. The noise tolerance value must be maintained constant for all images being analyzed. A low/high noise tolerance will depict too many/few maxima (foci) within the nucleus and will not be an accurate representation of the foci within the nucleus.</li>
		<li>Select the nuclei for which H2AX foci must be quantified for from the &#34;ROI Manager&#34; window. Select all the nuclei by checking the &#34;show all&#34; option.</li>
		<li>Select &#34;Measure&#34; to measure the number of maxima/foci within the selected nuclei. 
		NOTE: The number of maxima/foci appear as multiples of 255, i.e., one maximum has a &#34;RawIntDen&#34; (Integrated density) reading of 255.</li>
		<li>Copy the &#34;RawIntDen&#34; values and paste them into software for data analysis.</li>
	</ol>
	</li>
</ol>
2. Analyzing Localization to a Long-lasting Double-strand DNA Break
<ol>
	<li>Seed cells on 96-well plates/coverslips as described in sections 1.1 and 1.2.</li>
	<li>Induction of double-strand DNA breaks
	<ol>
		<li>Transfect cells with the I-SceI expression vector using a lipid-based transfection reagent according to the manufacturer&#39;s specifications. Wait 24&#8211;72 h after transfection to maximize the endonuclease expression. Determine if the transfection was successful by locating cells with a singular large nuclear &#947;-H2AX focus.</li>
	</ol>
	</li>
	<li>Acquisition of images
	<ol>
		<li>Fix, permeabilize, block, and wash the 96-well plate/coverslips as described in section 1.8.</li>
		<li>Co-incubate cells with an antibody against &#947;-H2AX and the repair protein of interest (here, a primary antibody against RAD51 was used) diluted according to the manufacturer&#39;s specifications in 3% BSA solution.</li>
		<li>Wash the 96-well plate/coverslips 3 times with 1x PBS. Incubate the 96-well plate/coverslips with secondary antibody diluted according to the manufacturer&#39;s protocol in 3% BSA solution. Confirm that the secondary fluorophores do not have overlapping excitation or emission spectra.</li>
		<li>Identify cells that have a large nuclear &#947;-H2AX focus. Only take pictures of these cells to avoid bias.</li>
	</ol>
	</li>
	<li>Analysis of long-lasting I-SceI induced foci
	<ol>
		<li>Analyze the long-lasting foci manually, by counting both the number of cells that have the large &#947;-H2AX foci and co-localized foci of the repair protein of interest.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The analysis of DNA damage repair in general and the repair of double-stranded DNA breaks specifically is an active area of research because its consequences span tumorigenesis to basic biology6,20. This manuscript details an approach that accurately dissects the contribution of RAD51 and &#947;-H2AX proteins to the resolution of DSBs through HR. Looking forward, this method can be used to elucidate additional functions of repair proteins at DSBs<sup class="xref"...

Each day, every cell in the human body is bombarded with an estimated 10,000 DNA lesions1. This existential threat puts us at risk for mutations, oncogenesis as well as cell death. To protect genome fidelity, mammalian cells have evolved to respond to DNA damage with a complex series of protein associations and modifications. This response is organized into multiple pathways, collectively known as the DNA damage response (DDR)2,3. The DDR consists of the accumulation of DNA repair proteins at DNA lesions, coordinated both temporally and spatially. DDR frequently induces cell cycle arrest to avoid the propagation or intensification of damage that can occur during the replication of damaged DNA2,4,5. In turn, it is also necessary for the cellular viability to turn off cell cycle arrest by disassociating repair complexes after repair has been completed.
Among the various types of DNA damage, DSBs are the most deleterious. Failure to repair DSBs can result in chromosome rearrangements or large-scale deletions such as the loss of entire chromosome arms. The repair of DSBs is divided into two pathways6,7,8. Homologous recombination (HR) requires a sister chromosome to use as a DNA template and thus is limited to late S and G2/M phases of the cell cycle9,10. Non-homologous end joining (NHEJ) does not have these restrictions but can cause small deletions when repairing DSBs11,12.
DSB repair specifically and the DDR in general are active areas of investigation. Despite being organized into conveniently separated pathways, there is a great deal of redundancy. Indeed, many proteins (BRCA1, BRCA2, and the RPA complex for example) are involved in multiple pathways13,14,15,16. The repair of a lesion by one pathway, can lead to a damage intermediate that must be repaired by another pathway14. The intertwining of these pathways, combined with their complex task of recruiting the right proteins to the correct place for the precise amount of time necessary, requires a multi-tiered regulatory process.
A recent report highlights the intricacies of DDR by demonstrating that repair complex formation, resolution, and localization can each separately be impaired17. The overall goal of the following protocol is to definitively dissect the ability of cells to repair DSB. Using immunofluorescence microscopy, accumulation of repair proteins at the sites of damage can be visualized at the representative time points following the induction of DSBs.
This technique has several advantages to commonly used approaches. Frequently, repair is investigated at single time points and incapable of representing the dynamic process of assembly and dissociation of repair complexes. Observing the full range of repair from the initial activation to the full resolution ensures that a delay in repair is not misidentified as complete inhibition. Conversely, it assures that the induction of a repair response that is unable to inactivate said response is not misidentified as the normal or the excessive activation.
Delayed protein complex formation and the mis-localization of repair proteins, however, may not be unambiguously distinguished with this approach. To determine if repair proteins are mis-localized versus delayed in their localization, a &#34;long-lasting&#34; DSB can be introduced through enzymatic cleavage of cellular DNA. The resulting lesion is recut each time it is repaired, resulting in a distinct large nuclear repair focus and removing the temporal restriction from recruitment. This can be achieved by modifying the existing approach with the use of a rare cutting endonuclease (e.g., I-SceI) to induce a long-lasting DSB. The longevity of DSBs enables the visualization of elusive repair proteins by immunofluorescence microscopy. The enhanced abundance could also improve the visualization when detection is hindered by limitation in the antibody quality, a feature that could be useful when lesser studied proteins are identified as having an impact on DNA repair.
Notably, we provide explicit instructions for a free image processing and analysis software (e.g., ImageJ). This removes a major financial barrier in image analysis, opening the interrogation of DNA damage repair to a wider audience.

<table><tbody><tr><td>12 mm Coverslips</td><td>VWR</td><td>89015-725</td><td></td></tr><tr><td>16% Paraformaldehyde (PFA)</td><td>ThermoFisher Scientific</td><td>28908</td><td></td></tr><tr><td>24-well plate</td><td>VWR</td><td>82050-892</td><td></td></tr><tr><td>96-well glass bottom plate</td><td>Cellvis</td><td>P96-1.5H-N</td><td></td></tr><tr><td>Anti-H2AX Alexa488</td><td>EMD Millipore</td><td>05-636-AF488</td><td></td></tr><tr><td>Anti-Rad51 (D4B10)&amp;nbsp;</td><td>Cell Signaling Technology</td><td>8875S</td><td></td></tr><tr><td>Bio-formats plugin for ImageJ</td><td>National Institute of Health (NIH)&amp;nbsp;</td><td></td><td>https://imagej.nih.gov/ij/plugins/index.html</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>VWR</td><td>97061-416</td><td></td></tr><tr><td>DAPI</td><td>ThermoFisher Scientific</td><td>D1306</td><td></td></tr><tr><td>DMEM, High Glucose</td><td>ThermoFisher Scientific</td><td>12100046</td><td></td></tr><tr><td>EDTA</td><td>Invitrogen</td><td>15576-028</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>VWR</td><td>89510-194</td><td></td></tr><tr><td>Goat Anti-Rabbit IgG Alexa594</td><td>ThermoFisher Scientific</td><td>&amp;nbsp;A-11012&amp;nbsp;</td><td></td></tr><tr><td>Hydrogen Peroxide</td><td>sigma-Aldrich</td><td>216763-100ML</td><td></td></tr><tr><td>ImageJ Software</td><td>National Institute of Health (NIH)&amp;nbsp;</td><td></td><td>https://imagej.nih.gov/ij/</td></tr><tr><td>I-SceI Expression Vector</td><td>Addgene</td><td>26477</td><td></td></tr><tr><td>Nail Polish- Insta Dri</td><td>Sally and Hansen</td><td>Clearly Quick (103)</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Bio Basic</td><td>PD8117</td><td></td></tr><tr><td>ProLong Gold Antifade Reagent</td><td>Life Technologies</td><td>P36930</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100-100ML</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Sigma-Aldrich</td><td>T4049-500ML</td><td></td></tr><tr><td>TurboFect Transfection Reagent</td><td>ThermoFisher Scientific</td><td>R0531</td><td></td></tr><tr><td>Tween-20</td><td>Fisher Scientific</td><td>BP337-500</td><td></td></tr></tbody></table>

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.

Figure 1 depicts the selection of the correct noise discrimination for maxima/foci quantification using ImageJ. The merged images of DAPI and the repair protein of interest are on the left panel. Figure 1A shows a noise discrimination of 90 and marks the correct number of foci. Nuclei on the edge (depicted with a pink arrow) and foci outside the nuclei (depicted with a yellow arrow) are not counted during the quantification. <str...

Watch this Scientific Journal Video about Characterizing DNA Repair Processes at Transient and Long-lasting Double-strand DNA Breaks by Immunofluorescence Microscopy at JoVE.com

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

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 ...

characterizing-dna-repair-processes-at-transient-long-lasting-double

Universidad de Cartagena UDEC

Research

JoVE Journal

Cancer Research

9.0K Views.  Kansas State University. 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.

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

Application of Laser Micro-irradiation for Examination of Single and Double Strand Break Repair in Mammalian Cells

Highly coordinated DNA repair pathways exist to detect, excise and replace damaged DNA bases, and coordinate repair of DNA strand breaks. While molecular biology techniques have clarified structure, enzymatic functions, and kinetics of repair proteins, there is still a need to understand how repair is coordinated within the nucleus. Laser micro-irradiation offers a powerful tool for inducing DNA damage and monitoring the recruitment of repair proteins. Induction of DNA damage by laser micro-irradiation can occur with a range of wavelengths, and users can reliably induce single strand breaks, base lesions and double strand breaks with a range of doses. Here, laser micro-irradiation is used to examine repair of single and double strand breaks induced by two common confocal laser wavelengths, 355 nm and 405 nm. Further, proper characterization of the applied laser dose for inducing specific damage mixtures is described, so users can reproducibly perform laser micro-irradiation data acquisition and analysis.

Confocal fluorescence microscopy and laser micro-irradiation offer tools for inducing DNA damage and monitoring the response of DNA repair proteins in selected sub-nuclear areas. This technique has significantly advanced our knowledge of damage detection, signaling, and recruitment. This manuscript demonstrates these technologies to examine single and double strand break repair.

Highly coordinated DNA repair pathways exist to detect, excise and replace damaged DNA bases, and coordinate repair of DNA strand breaks. While molecular ...

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of programmed DNA breaks in lymphocytes. The Ataxia-Telangiectasia Mutated kinase (ATM), ATM- and Rad3-Related kinase (ATR) and the catalytic subunit of DNA-dependent Protein Kinase (DNA-PKcs) are among the first that are activated upon induction of DNA damage, and are central regulators of a network that controls DNA repair, apoptosis and cell survival. As part of a tumor-suppressive pathway, ATM and ATR activate p53 through phosphorylation, thereby regulating the transcriptional activity of p53. DNA damage also results in the formation of so-called ionizing radiation-induced foci (IRIF) that represent complexes of DNA damage sensor and repair proteins that accumulate at the sites of DNA damage, which are visualized by fluorescence microscopy. Co-localization of proteins in IRIFs, however, does not necessarily imply direct protein-protein interactions, as the resolution of fluorescence microscopy is limited. 
In situ Proximity Ligation Assay (PLA) is a novel technique that allows the direct visualization of protein-protein interactions in cells and tissues with unprecedented specificity and sensitivity. This technique is based on the spatial proximity of specific antibodies binding to the proteins of interest. When the interrogated proteins are within ~40 nm an amplification reaction is triggered by oligonucleotides that are conjugated to the antibodies, and the amplification product is visualized by fluorescent labeling, yielding a signal that corresponds to the subcellular location of the interacting proteins. Using the established functional interaction between ATM and p53 as an example, it is demonstrated here how PLA can be used in suspension cell cultures to study the direct interactions between proteins that are integral parts of the DNA damage response.

Here, it is demonstrated how the in situ Proximity Ligation Assay (PLA) can be used to detect and visualize the direct protein-protein interactions between ATM and p53 in suspension cell cultures exposed to genotoxic stress.

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of ...

Biochemistry

Immunofluorescence Microscopy of &#947;H2AX and 53BP1 for Analyzing the Formation and Repair of DNA Double-strand Breaks

DNA double-strand breaks (DSB) are serious DNA lesions. Analysis of the formation and repair of DSB is relevant in a broad spectrum of research areas including genome integrity, genotoxicity, radiation biology, aging, cancer, and drug development. In response to DSB, the histone H2AX is phosphorylated at Serine 139 in a region of several megabase pairs forming discrete nuclear foci detectable by immunofluorescence microscopy. In addition, 53BP1 (p53 binding protein 1) is another important DSB-responsive protein promoting repair of DSB by nonhomologous end-joining while preventing homologous recombination. According to the specific functions of &#947;H2AX and 53BP1, the combined analysis of &#947;H2AX and 53BP1 by immunofluorescence microscopy may be a reasonable approach for a detailed analysis of DSB. This manuscript provides a step-by-step protocol supplemented with methodical notes for performing the technique. Specifically, the influence of the cell cycle on &#947;H2AX foci patterns is demonstrated in normal fibroblasts of the cell line NHDF. Further, the value of the &#947;H2AX foci as a biomarker is depicted in x-ray irradiated lymphocytes of a healthy individual. Finally, genetic instability is investigated in CD34+ cells of a patient with acute myeloid leukemia by immunofluorescence microscopy of &#947;H2AX and 53BP1.

This manuscript provides a protocol for the analysis of DNA double-strand breaks by immunofluorescence microscopy of &#947;H2AX and 53BP1.

DNA double-strand breaks (DSB) are serious DNA lesions. Analysis of the formation and repair of DSB is relevant in a broad spectrum of research areas ...

Immunofluorescence Imaging of DNA Damage and Repair Foci in Human Colon Cancer Cells

The purpose of the manuscript is to provide a step-by-step protocol for performing immunofluorescence microscopy to study the radiation-induced DNA damage response induced by neutron-gamma mixed-beam used in boron neutron capture therapy (BNCT). Specifically, the proposed methodology is applied for the detection of repair proteins activation which can be visualized as foci using antibodies specific to DNA double-strand breaks (DNA-DSBs). DNA repair foci were assessed by immunofluorescence in colon cancer cells (HCT-116) after irradiation with the neutron-mixed beam. DNA-DSBs are the most genotoxic lesions and are repaired in mammalian cells by two major pathways: non-homologous end-joining pathway (NHEJ) and homologous recombination repair (HRR). The frequencies of foci, immunochemically stained, for commonly used markers in radiobiology like &#947;-H2AX, 53BP1 are associated with DNA-DSB number and are considered as efficient and sensitive markers for monitoring the induction and repair of DNA-DSBs. It was established that &#947;-H2AX foci attract repair proteins, leading to a higher concentration of repair factors near a DSB. To monitor DNA damage at the cellular level, immunofluorescence analysis for the presence of DNA-PKcs representative repair protein foci from the NHEJ pathway and Rad52 from the HRR pathway was planned. We have developed and introduced a reliable immunofluorescence staining protocol for the detection of radiation-induced DNA damage response with antibodies specific for repair factors from NHEJ and HRR pathways and observed radiation-induced foci (RIF). The proposed methodology can be used for investigating repair protein that is highly activated in the case of neutron-mixed beam radiation, thereby indicating the dominance of the repair pathway.

Radiation-induced DNA damage response activated by neutron mixed-beam used in boron neutron capture therapy (BNCT) has not been fully established. This protocol provides a step-by-step procedure to detect radiation-induced foci (RIF) of repair proteins by immunofluorescence staining in human colon cancer cell lines after irradiation with the neutron-mixed beam.

The purpose of the manuscript is to provide a step-by-step protocol for performing immunofluorescence microscopy to study the radiation-induced DNA damage ...

Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging (ESI)

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in cell biology. Many phenomena cannot be explained by in vitro analysis in simplified systems and need additional structural information in situ, particularly in the range between 1 nm and 0.1 &#181;m, in order to be fully understood. Here, electron spectroscopic imaging, a transmission electron microscopy technique that allows simultaneous mapping of the distribution of proteins and nucleic acids, and an expression tag, miniSOG, are combined to study the structure and organization of DNA double-strand break repair foci.

Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging ESI

Electron spectroscopic imaging can image and distinguish nucleic acid from protein at nanometer resolution. It can be combined with the miniSOG system, which is able to specifically label tagged proteins in transmission electron microscopy samples. We illustrate the use of these technologies using double-strand break repair foci as an example.

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in ...

Biology

Advanced Confocal Microscopy Techniques to Study Protein-protein Interactions and Kinetics at DNA Lesions

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological approach to analyzing protein kinetics at DNA lesions over time or protein-protein interactions on locally microirradiated chromatin. We also show how to recognize individual phases of the cell cycle using the Fucci cellular system to study cell-cycle-dependent protein kinetics at DNA lesions. A methodological description of the use of two UV lasers (355 nm and 405 nm) to induce different types of DNA damage is also presented. Only the cells microirradiated by the 405-nm diode laser proceeded through mitosis normally and were devoid of cyclobutane pyrimidine dimers (CPDs). We also show how microirradiated cells can be fixed at a given time point to perform immunodetection of the endogenous proteins of interest. For the DNA repair studies, we additionally describe the use of biophysical methods including FRAP (Fluorescence Recovery After Photobleaching) and FLIM (Fluorescence Lifetime Imaging Microscopy) in cells with spontaneously occurring DNA damage foci. We also show an application of FLIM-FRET (Fluorescence Resonance Energy Transfer) in experimental studies of protein-protein interactions.

Laser microirradiation is a useful tool for studies of DNA repair in living cells. A methodological approach for the use of UVA lasers to induce various DNA lesions is shown. We have optimized a method for local microirradiation that maintains the normal cell cycle; thus, irradiated cells proceed through mitosis.

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological ...

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 ...

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

Cells are continuously exposed to various DNA damaging agents, inducing different cellular responses. Applying biochemical and genetic approaches is essential in revealing cellular events associated with the recruitment and assembly of DNA repair complexes at the site of DNA damage. In the last few years, several powerful tools have been developed to induce site-specific DNA damage. Moreover, novel seminal techniques allow us to study these processes at the single-cell resolution level using both fixed and living cells. Although these techniques have been used to study various biological processes, herein we present the most widely used protocols in the field of DNA repair, Fluorescence Immunostaining (IF) and Chromatin Immunoprecipitation (ChIP), which in combination with endonuclease-based site-specific DNA damage make it possible to visualize and quantify the genomic occupancy of DNA repair factors in a directed and regulated fashion, respectively. These techniques provide powerful tools for the researchers to identify novel proteins bound to the damaged genomic locus as well as their post-translational modifications necessary for their fine-tune regulation during DNA repair.

This article introduces essential steps of immunostaining and chromatin immunoprecipitation. These protocols are commonly used to study DNA damage-related cellular processes and to visualize and quantify the recruitment of proteins implicated in DNA repair.

Cells are continuously exposed to various DNA damaging agents, inducing different cellular responses. Applying biochemical and genetic approaches is ...

Detection of G1 Phase Single-Stranded DNA Foci via RPA2 Immunofluorescent Staining

Visualizing Single-Stranded DNA Foci in the G1 Phase of the Cell Cycle

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair necessitates collaboration between numerous proteins, responsible for covering a broad range of tasks from recognizing and signaling the presence of a DNA lesion to physically repairing it. During this process, tracks of single-stranded DNA (ssDNA) are often created, which are eventually filled by DNA polymerases. The nature of these ssDNA tracks (in terms of both length and number), along with the polymerase recruited to fill these gaps, are repair pathway-specific. The visualization of these ssDNA tracks can help us understand the complicated dynamics of DNA repair mechanisms.
This protocol provides a detailed method for the preparation of G1 synchronized cells to measure ssDNA foci formation upon genotoxic stress. Using an easy-to-utilize immunofluorescence approach, we visualize ssDNA by staining for RPA2, a component of the heterotrimeric replication protein A complex (RPA). RPA2 binds to and stabilizes ssDNA intermediates that arise upon genotoxic stress or replication to control DNA repair and DNA damage checkpoint activation. 5-Ethynyl-2'-deoxyuridine (EdU) staining is used to visualize DNA replication to exclude any S phase cells. This protocol provides an alternative approach to the conventional, non-denaturing 5-bromo-2'-deoxyuridine (BrdU)-based assays and is better suited for the detection of ssDNA foci outside the S phase.

Author Spotlight: Visualizing Single-Stranded DNA During DNA Repair for Therapeutic Insights 

The following protocol presents the detection of single-stranded DNA foci in the G1 phase of the cell cycle utilizing cell cycle synchronization followed by RPA2 immunofluorescent staining.

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair ...

Immunofluorescence-Based Visualization of DNA Repair Protein Interactions

Source: de la Pe&#241;a Avalos, B., et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/61447">Visualization of DNA Repair Proteins Interaction by Immunofluorescence.</a> J. Vis. Exp. (2020).
In this video, we describe the immunofluorescence method to visualize the localization of the DNA repair proteins &#947;H2AX and 53BP1 at the sites of double-strand DNA breaks in irradiated cell nuclei.

Source: de la Pe&#241;a Avalos, B., et al., Visualization of DNA Repair Proteins Interaction by Immunofluorescence. J. Vis. Exp. (2020).
In this video, we ...