We thank Marie-Christine Caron for outstanding technical advice. This work is supported by funding from Canadian Institutes of Health Research J.Y.M (CIHR FDN-388879). J.-Y.M. holds a Tier 1 Canada Research Chair in DNA Repair and Cancer Therapeutics. J.O&#39;S is an FRQS PhD student fellow, and S.Y.M is a FRQS postdoctoral fellow.

1. Cell culture, treatments, and coverslip preparation
NOTE: All cell plating, transfections, and treatments, aside from irradiation, should take place under a sterile cell-culture hood.
<ol>
	<li>Day 1
	<ol>
		<li>In a 6-well plate, place a single coverslip in each well for as many conditions as needed. Plate ~150,000 HeLa cells for transfection or drug treatment, as desired. 
		NOTE: If transfecting, it is recommended to do a reverse transfection at the time of plating, or it is possible to do a forward transfection several hours after plating to allow for adherence. A reverse transfection is accomplished by adding the transfection mix to the coverslip before the cells are added; thus, transfection begins before the cells start adhering. A forward transfection, by contrast, is the addition of the transfection mix post-adherence to the surface usually on the following day. However, it is possible to do this on the same day, provided enough time is given for the cells to adhere.</li>
		<li>For this method, use 4 wells: Well 1: siRNA control (termed siCTRL); Well 2: siRNA against PARP-1 (siPARP-1); Well 3: untreated; Well 4: Cells to be treated with 5 &#181;M BMN673 1 h prior to irradiation. 
		NOTE: In this protocol, all the tests were conducted under irradiated conditions (see section 1.3).</li>
		<li>Incubate the cells at 37 &#176;C in a 5% CO2 humidified incubator overnight, although the transfection protocol allows for up to 3 days of incubation. The incubation time prior to BrdU treatment will depend on the siRNA transfection efficiency. If there is no transfection, incubation for 16 h is sufficient to allow for adherence to the coverslip and some cell growth. 
		NOTE: Incubator conditions can be changed according to the cell line used.</li>
	</ol>
	</li>
	<li>Day 2
	<ol>
		<li>Add BrdU at a final concentration of 10 &#181;M in the appropriate culturing media, and incubate for 16 h (one cell cycle). 
		NOTE: The BrdU solution is prepared in dimethylsulfoxide; the stock solution used is 10 mM and is stored in aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Day 3
	<ol>
		<li>Irradiate the plates with a total dose of 5 Gy of X-ray irradiation (vary the dose of irradiation depending on the irradiator type, for example, small animal irradiators vs. benchtop irradiators). See the Table of Materials for the brand and model of irradiator used.</li>
		<li>Return the plates to the incubator, and release the cells for 3 h.</li>
		<li>During the 3 h incubation period, prepare the two buffers, A and B, and 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (PBS). 
		NOTE: Buffers A and B and the sucrose should be prepared fresh the day of fixation, using fresh sucrose solution to prevent contamination.
		<ol>
			<li>Prepare buffer A (Pre-Extraction Buffer), according to the order (30 mL) described in Table 1. 
			NOTE: The 1M sucrose should be prepared the day of use to prevent contamination.</li>
			<li>Prepare buffer B (Cytoskeleton Stripping Buffer) according to the order described in Table 1 (30 mL). 
			&#8203;NOTE: Buffer B must be prepared in the cited order to prevent precipitation of Tween-20 and sodium deoxycholate solution.</li>
			<li>Prepare 4% PFA, 2 mL per condition (10 mL), under a chemical hood (Table 1).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Pre-extraction and fixation
NOTE: All pre-extraction and&#160;fixation are performed with the coverslips remaining in the tissue culture plate on ice or at 4 &#176;C; the coverslip is only lifted in the last step of mounting (see discussion).
<ol>
	<li>Pre-extraction
	<ol>
		<li>Aspirate the medium, carefully wash the cells twice with 1x PBS, and remove the PBS. Add 2 mL of Pre-extraction Buffer A and immediately incubate at 4 &#176;C for 10 min. 
		NOTE: It is important that incubation time in buffer A is not extended. Extended incubation in the pre-extraction buffers will result in an increased number of detached cells from the coverslip. Alternatively, instead of incubation at 4&#176;C, incubation can be done on ice.</li>
		<li>Remove Buffer A through aspiration. 
		NOTE: Do not wash the coverslips after removing buffer A. Proceed to the next step.</li>
		<li>Add 2 mL of Cytoskeleton Stripping Buffer B and immediately incubate at 4 &#176;C for 10 min. Carefully aspirate Buffer B, and carefully wash the cells once with 1x PBS. Carefully aspirate the 1x PBS.</li>
	</ol>
	</li>
	<li>Fixation
	<ol>
		<li>Fix the cells by adding 2 mL of 4% PFA under the chemical hood. 
		NOTE: PFA is toxic and must be manipulated under a chemical hood.</li>
		<li>Incubate the cells at room temperature for 20 min. Wash the coverslips twice with 1x PBS, and aspirate the excess.</li>
		<li>Cover the coverslips with 100% cold methanol, and incubate the coverslips at -20 &#176;C for 5 min. Wash twice with 1x PBS. 
		&#8203;NOTE: The protocol can be paused here. The coverslips can be stored at 4 &#176;C in 1x PBS and the plate wrapped in aluminum foil if necessary. The cells should not be kept more than 5 days before continuing the IF protocol.</li>
	</ol>
	</li>
</ol>
3. Permeabilization
<ol>
	<li>Incubate the cells with 2 mL of 1x PBS containing 0.5% Triton X-100 at room temperature for 15 min. Wash the coverslips three times with 2 mL of 1x PBS.</li>
</ol>
4. Immunostaining
<ol>
	<li>Blocking step
	<ol>
		<li>Prepare enough fresh blocking buffer (3% BSA in 1x PBS) to use 2 mL per coverslip. 
		&#8203;NOTE: Prepare an extra 5mL of blocking buffer to make the antibody solutions.</li>
		<li>Add 2 mL of blocking buffer to each well and incubate at room temperature for 1 h.</li>
	</ol>
	</li>
	<li>Primary antibody incubation
	<ol>
		<li>Prepare the primary antibody solution in fresh blocking buffer: BrdU RPN202 1:1000 and proliferating cell nuclear antigen (PCNA) 1:500, 100 &#181;L per coverslip. 
		NOTE: To preserve the antibody, a smaller volume can be used, 75-100 &#181;L for a 22 x 22 mm coverslip, 50-75 &#181;L for an 18 x 18 mm coverslip.</li>
		<li>On each coverslip, add 100 &#181;L of primary antibody solution. Cover the coverslips with a square of parafilm using tweezers, and carefully position the parafilm to not create bubbles.</li>
		<li>Cover the plate in aluminum foil, and incubate the primary antibody overnight at 4 &#176;C in a humidified chamber.</li>
		<li>Remove the parafilm squares, and wash the coverslips three times with 2 mL of sterile 1x PBS.</li>
	</ol>
	</li>
	<li>Secondary antibody incubation
	<ol>
		<li>Prepare enough secondary antibody solution in fresh blocking buffer: anti-mouse 488 fluorescent secondary A11011 (for BrdU) dilution 1:800 and anti-rabbit 568 fluorescent secondary A11011 (for PCNA) dilution 1:800. 
		NOTE: The colors used can be changed to suit the experimental needs. The specific brand used can be found in the Table of Materials.</li>
		<li>Add on each coverslip 100 &#181;L of secondary antibody solution, cover the coverslips with a square of parafilm using tweezers, and carefully position the parafilm without bubbles.</li>
		<li>Incubate the secondary antibody at room temperature for 1 h with the plate covered in aluminum foil.</li>
		<li>Wash the coverslips three times with 2 mL of 1x PBS.</li>
	</ol>
	</li>
	<li>Nuclear staining
	<ol>
		<li>Prepare a volume of 2 mL per coverslip of 1x PBS containing 4&#39;,6-diamidino-2-phenylindole (DAPI) at a final concentration of 1 &#181;g/mL (1:1000).</li>
		<li>Add on each coverslip 2 mL of the DAPI solution. Incubate the coverslips at room temperature for 10 min with the plate covered in aluminum foil. Wash the coverslips twice with 1x PBS.</li>
		<li>Cover the coverslips with 1x PBS, and use either a needle or fine tweezers to lift the coverslip from the bottom of the well.Carefully blot the off the excess liquid by tapping one edge gently on a paper towel. 
		NOTE: Be careful not to drop the slide or allow the flat surface with the adherent cells to touch the paper.</li>
		<li>Mount the coverslips on slides using 10-20 &#181;L of IF-specific mounting media.</li>
	</ol>
	</li>
</ol>
5. Image acquisition and analysis
NOTE: Image acquisition can be done on several types of fluorescent microscopes. An epifluorescence microscope with a 63x oil objective was used; see the Table of Materials for the brand and model. Z-stacks are not required, although they may be of use depending on the cell line and level of mitochondrial staining.
<ol>
	<li>Image analysis
	<ol>
		<li>For each image, create multiple tiff files; merge all planes, but keep the color channels separate. Import these files into Cell Profiler (The Broad Institute https://cellprofiler.org/home) and analyze using the Speckle Counting pipeline (Supplemental Video 1).</li>
		<li>Upload the images (Supplemental Figure S1A).
		<ol>
			<li>To begin creating the project, load the speckle counting pipeline into the program and the images to be analyzed in the provided area.</li>
			<li>Use the NamesAndTypes module to assign a meaningful name to each image by which other modules will refer to it-C01: Representing BrdU Channel; C02: Representing PCNA Channel; C03: Representing DAPI Channel. 
			NOTE: These identifiers will depend on the microscope; there should be an identifier in the file name to distinguish between the channels.</li>
		</ol>
		</li>
		<li>Identify which file will be used to identify the nuclei and name it (change this name as required) (see Supplemental Figure S1B and Supplemental Figure S1C).
		<ol>
			<li>Select the diameter of the object between 50 and 300 pixel units, which is the acceptable size range for nuclei, but change the value to fit the nuclei in the image. 
			NOTE: It may be that 50 is too large a value and prevents smaller nuclei from being identified; likewise, 300 may allow for large groupings of nuclei to be counted as 1.</li>
			<li>Discard objects outside of the diameter range to ensure only those that match the criteria will be counted.</li>
			<li>Discard objects touching borders to remove cells that may only be partially in the field.</li>
			<li>Apply the threshold to fit the specific images. Note the following settings as an example. Change the thresholding correction factor based on how stringent the thresholding strategy will be. Other settings include threshold strategy: Global; thresholding method: Otsu; two class or three class thresholding: Two class; threshold smoothing scale: 1; threshold smoothing factor: 1; lower and upper bounds on threshold: 0.0 and 1.0; method to distinguish clumped objects: Shape; and method to draw dividing lines between clumped objects: Propagate. 
			NOTE: For each parameter, cell profiler provides complementary definitions and available possibility for the variable setting.</li>
		</ol>
		</li>
		<li>Identify the primary object to identify the foci (Supplemental Figure S2A).
		<ol>
			<li>Use the following settings: select the input image: Maskedgreen; named the primary object to be identified: BrdUFoci; typical diameter of the object: 1-15; discard the object outside the diameter range: Yes; discard the object touching the border of the image : No; threshold strategy: Adaptative; thresholding method: Otsu; two class or three class thresholding: Three class; threshold smoothing scale: 1; threshold smoothing factor: 3; and size of smoothing filter: 4.</li>
		</ol>
		</li>
		<li>Measure intensity (Supplemental Figure S2B).
		<ol>
			<li>Select the image to be measured: OrigGreen.</li>
			<li>Click on Add another image. Select the image to be measured: PCNA. Select objects to measure: Nuclei. 
			NOTE: This will be used to measure the BrdU and PCNA intensity-the final data that will be graphed.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This paper describes a method that makes use of IF staining to measure variations in DNA resection in cellulo. The current standard for observing an effect on DNA resection is through RPA staining; however, this is an indirect method that may be influenced by DNA replication. Previously, another BrdU incorporation-based DNA resection IF technique has been described for classifying the resulting intensities in BrdU-positive and BrdU-negative cells. This method allowed for cells that are not undergoing HR to be counted as positive due to background or mitochondrial staining resulting in a high BrdU intensity14,15,16. The primary novelty of the method described in this paper is the addition of PCNA staining, which allows for selectivity for S and G2 phases of the cell cycle, ensuring that the resulting BrdU signal is due to resection and thus, to homology-directed repair.
The critical step in this protocol is the pre-extraction; without this, the BrdU foci will not be visible, and if done incorrectly, the cells will detach completely. Incorrectly prepared buffers will result in incomplete pre-extraction, such as partial cytoskeletal removal or increased background signal. It is very important to respect the duration of incubation with the pre-extraction buffers, increased duration in the buffers can result in the cells detaching from the coverslip. Importantly, if this protocol is performed with poorly adherent cells, pre-treat the coverslips with polylysine to reduce cell detachment during pre-extraction. Subsequently, without methanol fixation, the PCNA signal will not be visible.
Another important variable in this protocol is the blocking buffer: incompatible blocking buffers will result in loss of BrdU signal. For example, 10% fetal bovine serum in 1x PBS blocking buffer will function when not combined with methanol fixation; however, when combined with methanol fixation as required for the PCNA staining, the BrdU signal is considerably reduced. It is possible other blocking agents and methods will work with this protocol; 3% BSA in 1x PBS provided the best results. A final essential component to this technique is the use of a DNA damage source; without this, there will be little to no nuclear BrdU signal (Supplemental Figure S5). Experience has shown that irradiation is an excellent source of damage for this method. However, if there is no access to an irradiator, radiomimetic drugs such as neocarzinostatin can be used instead.
Cell profiler is a useful and versatile tool that allows easy and consistent analysis of the images produced in this technique. The settings can be customized for both the size of the objects identified and the threshold at which it does so (Supplemental Video 1). As the nuclear intensity of the BrdU staining is the desired readout, the key customization is in the identification of the nuclei from the DAPI files. The analysis of the images from each condition must be done separately as the results are obtained in the form of a single csv file containing cell and image numbers, but not condition names. To determine if the analysis conditions are appropriate, use the Start test mode feature, and make sure the eye icon is open in the desired steps. Close this icon when running the actual analysis as it will result in pop-up windows and slow down the program. In the Start test mode, users can go back and forth, altering the settings as needed to properly identify first the nuclei and then the BrdU foci.
Once satisfied with the resulting identification, use the same settings for each condition within the same experiment; this may require the acquisition of a test image for each condition prior to analyzing the whole batch to ensure that the settings comply with each condition. It is important to note that, depending on image acquisition, there may be variability in the Cell profiler settings between users and between software versions. Hence, it is essential to go through the testing stage to determine the ideal parameters for specific experiments. When preparing to graph the results, the PCNA (S phase)-positive cells must be identified. To do this, the PCNA intensity is measured, in the same fashion as the BrdU intensity, and the resulting values plotted as a scatter plot to best visualize the cut-off intensity value for PCNA-positive cells. The PCNA-negative results will then be removed from the data set to allow for the BrdU intensity graphing. The provided supplementary figures show screenshots from the program highlighting the most optimized settings (Supplemental Figure S1, Supplemental Figure S2, and Supplemental Figure S3).
A possible modification to the protocol is the PCNA antibody used; two different PCNA antibodies were successfully tested, the one listed here and another which is no longer in production-a rat monoclonal antibody (Bulldog Bio PCA16D10). Finally, users should cover the coverslips with parafilm during antibody incubation. It is not strictly necessary, although a significantly larger volume of antibody solution would be required to cover the coverslip entirely without this. Another method is to place a sheet of parafilm surrounding a glass plate, place drops of the antibody solution onto the parafilm, and carefully place the coverslip cell-side down onto the drop of antibody solution. This glass plate can then be placed in a humid chamber at 4 &#176;C overnight for primary incubation and in the dark at room temperature for secondary incubation. This method is completely functional although it does significantly increase the time of handling the coverslips and as a result, increases the probability of dropping or breaking the coverslip.
An alternative method to measure resection is through the use of the DNA-combing SMART method; while this technique provides very clear results, it is much more complicated, requires more time, and is more expensive17. The SMART method requires the extraction of the DNA without damaging it, followed by stretching this DNA onto coverslips to be followed by an IF. The BrdU IF method, by comparison, is both simple and cost-effective, not requiring a greater investment than the cost of the antibody. This method provides an initial way to measure resection that is more accurate than simply measuring RPA foci formation, which can be related to its effects on many cellular functions. However, RPA is phosphorylated on specific residues during resection as part of the DNA damage response.
Single-molecule imaging recently revealed phosphorylated RPA (pRPA) as a negative resection regulator18. Hence, in the long term, this method could be made even more precise by coupling PCNA/phosphorylated RPA staining with BrdU imaging with the use of appropriate antibodies and optimized staining conditions. However, the method presented provides a representation of resection that is observed independently from the proteins involved in the resection process, thus negating bias that can occur from changes in their signal in response to the treatment. This technique provides not only a method to determine if a protein is involved in resection, but also to determine if cell death resulting from drug treatment is related to hypo/hyper-resection. This information can be beneficial in understanding not only the mechanistic aspects of DNA repair, but also the biological mechanism underlying the drug-induced cell death.

Modulation of DNA repair factors is an ever-evolving method for cancer therapy, particularly in DNA DSB repair-deficient tumor environments. The inhibition of specific repair factors is one of the ingenious strategies used to sensitize cancer cells to DNA-damaging agents. Decades of research led to the identification of various mutations of DNA repair genes as biomarkers for therapeutic strategy choices1. Consequently, the DNA repair field has become a hub for drug development to ensure a wide range of treatments, empowering the personalized medicine concept. 
DSBs are repaired by two main pathways: NHEJ and HR2. The NHEJ pathway is error-prone, rapidly ligating the two DNA ends with little to no DNA end-processing and involving the protein kinase (DNA-PKcs), the Ku70/80 complex, 53BP1, and RIF1 proteins3. In contrast, HR is a faithful mechanism initiated by BRCA14. An essential step in HR repair is the DNA-end resection process, which is the degradation of the broken ends leading to single-stranded (ss) DNA with 3'-OH ends. BRCA1 facilitates the recruitment of the downstream proteins that form the resectosome MRN/RPA/BLM/DNA2/EXO1, which are involved in the 5' to 3' DNA resection5. 
The initial end-resection is accomplished through the endonuclease activity of MRE11, allowing for further processing by the DNA2 and EXO1 nucleases. The generated ssDNA overhangs are quickly coated by Replication Protein A (RPA) to protect them from further processing. Subsequently, BRCA2, PALB2, and BRCA1 engage to mediate the displacement of RPA and the assembly of the RAD51 nucleofilament required for homology-directed repair mechanism. A fine balance between the usage of NHEJ and HR is necessary for the optimal maintenance of genomic integrity. The pathway choice depends on the cell cycle phase. HR is preferentially used during the S to G2 phases wherein DNA resection is at the highest level, and the sister chromatids are available to ensure proper repair.
Poly (ADP-ribose) polymerase 1 (PARP-1) is one of the earliest proteins recruited to the DSB. It regulates both resection activity and the assembly of downstream effectors involved in the NHEJ5,6. PARP-1 is also required for DNA single-stranded break repair during replication7,8. Due to its important role in DNA repair, PARP inhibitors (PARPi) are used as cancer therapies. In several HR-deficient cancers, PARPi treatment leads to a synthetic lethal response due to the incapacity of HR-deficient cells to repair the accumulated damage via an alternative pathway9,10. There are currently four FDA approved PARPi: Olaparib, Rucaparib, Niriparib, and Talazoparib (also called BMN 673), which are used for various breast and ovarian cancer treatments11. However, PARPi resistance is common, and one potential cause arises through the reacquisition of HR proficiency12. Loss or inhibition of PARP-1 in the presence of irradiation dysregulates the resectosome machinery, leading to the accumulation of longer ssDNA tracts13. Therefore, an in-depth study of DNA resection in vivo is critical for a clearer understanding of the DNA repair pathways and the subsequent development of new strategies to treat cancer and to overcome PARPi resistance. 
There have been several methods employed to detect DNA resection events5. One such method is the classical IF-based technique allowing for indirect staining and visualization of the resected DNA after stress-induced DSB by using an anti-RPA antibody. Labelling genomic DNA with 5-bromo-2&#8242;-deoxyuridine (BrdU) and detecting only ssDNA is a direct measurement of DNA resection events. It circumvents the monitoring of RPA, which is involved in multiple cellular processes such as DNA replication. In the method described here, cells incubated with BrdU for a single cell cycle allow BrdU to be incorporated into one strand of the replicating cellular DNA. Following resection, IF staining is performed under conditions allowing wherein BrdU detection only in the ssDNA form, with the use of an anti-BrdU antibody. This antibody can only access exposed BrdU nucleotides and will not detect those integrated into double-stranded DNA. Using fluorescence microscopy, the resected DNA can be visualized in the form of punctate BrdU/ssDNA foci. The nuclear intensity of these foci can be used as a readout to quantify resection following DNA damage. This paper describes step-by-step the processes of this method, which can be applied to most mammalian cell lines. This method should be of broad utility as a simple way of monitoring DNA end resection in cellulo, as a proof of concept.

<table><tbody><tr><td>Alexa 568 goat anti-rabbit</td><td>Molecular probes</td><td>A11011</td><td>1:800</td></tr><tr><td>Alexa Fluor 488 goat anti-mouse</td><td>Molecular probes</td><td>A11001</td><td>1:800</td></tr><tr><td>Anti PARP1 (F1-23)</td><td>Homemade</td><td></td><td>1:2500</td></tr><tr><td>Anti PCNA (SY12-07)</td><td>Novus</td><td>NBP2-67390</td><td>1:500</td></tr><tr><td>Anti-Alpha tubulin (DM1A)</td><td>Abcam</td><td>Ab7291</td><td>1:100000</td></tr><tr><td>anti-BrdU</td><td>GE Healthcare</td><td>RPN202</td><td>1:1000</td></tr><tr><td>Benchtop X-ray Irradiator</td><td>Cell Rad</td><td></td><td></td></tr><tr><td>BMN673</td><td>MedChem Express</td><td>HY-16106</td><td></td></tr><tr><td>Bromodeoxyuridine (BrdU)</td><td>Sigma</td><td>B5002</td><td></td></tr><tr><td>BSA</td><td>Sigma</td><td>A7906</td><td></td></tr><tr><td>Cell profiler</td><td>Broad Institute</td><td>V 3.19</td><td>https://cellprofiler.org/</td></tr><tr><td>Curwood Parafilm M Laboratory Wrapping Film 4in / 250 ft</td><td>Fisher scientific</td><td>13-374-12</td><td></td></tr><tr><td>DAPI (4&#039;,6-Diamidino-2-Phenylindole, Dihydrochloride)</td><td>Invitrogen Life Technology</td><td>D1306</td><td></td></tr><tr><td>DMEM high glucose</td><td>Fisher scientific</td><td>10063542</td><td></td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E3889</td><td></td></tr><tr><td>Fetal Bovine serum</td><td>Gibco</td><td>12483-020</td><td></td></tr><tr><td>Fisherbrand Cover Glasses: Squares 22 x 22</td><td>Fisher scientific</td><td>12 541B</td><td></td></tr><tr><td>Fluorescent microscope</td><td>Leica</td><td>DMI6000B</td><td>63x immersion objective</td></tr><tr><td>HeLa</td><td>ATCC</td><td>CCL-2</td><td></td></tr><tr><td>HERACELL 160I CO&lt;sub&gt;2&lt;/sub&gt; INCUBATOR CU 1-21 TC 120V</td><td>VWR</td><td>51030408</td><td>37% CO&lt;sub&gt;2&lt;/sub&gt;</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>BioShop Canada</td><td>MAG520.500</td><td></td></tr><tr><td>NaCl</td><td>BioShop Canada</td><td>SOD002.10</td><td></td></tr><tr><td>Needle</td><td></td><td></td><td></td></tr><tr><td>PBS 1x</td><td>Wisent Bio Products</td><td>311-010-CS</td><td></td></tr><tr><td>PFA 16%</td><td>Cedarlane Labs</td><td>15710-S(EM)</td><td></td></tr><tr><td>PIPES</td><td>Sigma-Aldrich</td><td>P6757-100G</td><td></td></tr><tr><td>ProLong Gold Antifade Mountant</td><td>Invitrogen Life Technology</td><td>P-36930</td><td></td></tr><tr><td>RNAiMAX</td><td>Invitrogen</td><td>13778-075</td><td></td></tr><tr><td>siPARPi</td><td>Dharmacon</td><td></td><td>AAG AUA GAG CGU GAA GGC GAA dTdT</td></tr><tr><td>siRNA control</td><td>Dharmacon</td><td></td><td>UUCGAACGUGUCACGUCAA</td></tr><tr><td>Sodium Deoxycholcate</td><td>Sigma-Aldrich</td><td>D6750-100G</td><td></td></tr><tr><td>Sucrose</td><td>BioShop Canada</td><td>SUC507.5</td><td></td></tr><tr><td>Tris-base</td><td>BioShop Canada</td><td>TRS001.5</td><td></td></tr><tr><td>Trition X-100</td><td>Millipore Sigma</td><td>T8787-250ML</td><td></td></tr><tr><td>Tween20</td><td>Fisher scientific</td><td>BP337500</td><td></td></tr><tr><td>Tweezers</td><td></td><td></td><td></td></tr></tbody></table>

assessment of global dna double-strand end resection using brdu-dna labeling coupled with cell cycle discrimination imaging

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs for cancer treatment. These targets are poly(ADP-ribose) polymerases (PARPs), which initiate various forms of DNA repair. Inhibiting these enzymes using PARP inhibitors (PARPi) achieves synthetic lethality by conferring a therapeutic vulnerability in homologous recombination (HR)-deficient cells due to mutations in breast cancer type 1 (BRCA1), BRCA2, or partner and localizer of BRCA2 (PALB2). 
Cells treated with PARPi accumulate DNA double-strand breaks (DSBs). These breaks are processed by the DNA end resection machinery, leading to the formation of single-stranded (ss) DNA and subsequent DNA repair. In a BRCA1-deficient context, reinvigorating DNA resection through mutations in DNA resection inhibitors, such as 53BP1 and DYNLL1, causes PARPi resistance. Therefore, being able to monitor DNA resection in cellulo is critical for a clearer understanding of the DNA repair pathways and the development of new strategies to overcome PARPi resistance. Immunofluorescence (IF)-based techniques allow for monitoring of global DNA resection after DNA damage. This strategy requires long-pulse genomic DNA labeling with 5-bromo-2&#8242;-deoxyuridine (BrdU). Following DNA damage and DNA end resection, the resulting single-stranded DNA is specifically detected by an anti-BrdU antibody under native conditions. Moreover, DNA resection can also be studied using cell cycle markers to differentiate between various phases of the cell cycle. Cells in the S/G2 phase allow the study of end resection within HR, whereas G1 cells can be used to study non-homologous end joining (NHEJ). A detailed protocol for this IF method coupled to cell cycle discrimination is described in this paper.

In this protocol, the bromodeoxyuridine (BrdU)-based assay was used to quantitatively measure the resection response of HeLa cells to irradiation-induced damage. The generated ssDNA tracks are visualized as distinct foci after immunofluorescence staining (Figure 1A). The identified foci were then quantified and expressed as the total integrated intensity of the BrdU staining in the nuclei (Figure 1B, Supplemental Figure S1, Supplemental Figure S2, and Supplemental Figure S3). It is possible to measure the foci number or mean foci intensity, although this can be less reliable than the total nuclear intensity, largely in part to the variable size of the BrdU foci.
To differentiate between the short-range resection as a result of NHEJ and the long-range resection of HR, co-staining was performed using an anti-PCNA antibody to identify cells going through S-phase (Figure 1 and Supplemental Figure S4). PCNA constitutes the DNA clamp that acts as a processivity factor for DNA polymerase and is essential for replication. PCNA is prominent in the nucleus and reaches maximal expression during the S-phase of the cell cycle. Hence, in early S-phase, the PCNA signal is low and has a granular distribution. In contrast, in late S-phase, the PCNA staining is quite strong (Figure 1A). When first analyzing the PCNA (S-phase) signal, the nuclei must be identified and the PCNA intensity measured.
The resulting values are then plotted as a scatter plot to best determine the cut-off intensity to discriminate between the PCNA-positive and PCNA-negative nuclei (Supplemental Figure S4A). The PCNA-negative results will then be removed from the data set to allow for the BrdU intensity-based analysis because of the low BrdU signal regardless of the condition (Supplemental Figure S4B). In these experimental conditions, the PCNA-negative nuclei harbor a basal integrated intensity of BrdU foci of 900 arbitrary units (A.U.). However, this value reaches 1800 A.U. in PCNA-positive nuclei (Figure 1B). This represents a 100% increase in the amount of the integrated signal of the BrdU foci. Increase in BrdU intensity can be then correlated to an increase in DNA resection, which is more pronounced and efficient when cells go through S and G2 phases and are irradiated.
In this protocol, to demonstrate an increase in DNA resection after irradiation (5 Gy), PARP-1 activity was modulated either through the loss of PARP-1 using an siRNA knockdown condition or after potent inhibition of PARP-1 using Talazoparib (BMN673) (Figure 2 and Supplemental Figure S5). The amounts of resected DNA in each condition were then quantified after IF (Figure 2B). In unperturbed condition (siCTRL), the integrated intensity of BrdU foci per nucleus is approximately 1500 A.U. in the PCNA-positive nuclei (Figure 2A,B). After an efficient knockdown of PARP-1 using an siRNA (Figure 2C), a significant increase in the intensity of the BrdU foci to approximately 1900 A.U. was observed, which corresponds to 26% increase in intensity (p ˂ 0.0001). Similarly, after inhibition of PARP-1 using BMN673 (5 µM final concentration), the intensity of foci increased from approximately 1750 to 2500 A.U. corresponding to 43% increase of the foci intensity (p ˂ 0.0001). Thus, loss of PARP-1 dysregulates DNA resection as reported previously13.
It is important to remember that BMN-673 both inhibits the activity of PARP-1 and traps it in DNA, and the effect of this treatment is much more detrimental to the cell, resulting in the greater increase of BrdU intensity than in siRNA-treated cells. The slight difference in untreated and siCTRL intensities demonstrates how any treatment, such as siRNA transfection, may affect the response and output of an assay. It can further demonstrate the importance of using the proper controls for each condition.
<img alt="Cell imaging of DAPI, PCNA, BrdU foci post-IR 5 Gy; fluorescence microscopy, intensity graph, analysis." class="xfigimg" src="/files/ftp_upload/62553/62553fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: BrdU foci formation is more prone to occur in PCNA-positive cells than in replicating cells. (A) Representative images of BrdU foci formation and PCNA staining following 5 Gy irradiation and 3 h release. A zoomed-in square showing marked BrdU foci is present in the corner of each BrdU image. Scale bars = 5 µm. (B) Quantification of the BrdU nuclear intensity in untreated (without BMN-673) HeLa cells. The data show the mean ± s.e.m (Mann-Whitney U-test). Abbreviations: BrdU = 5-bromo-2′-deoxyuridine; PCNA = proliferating cell nuclear antigen; IR = irradiation; DAPI = 4',6-diamidino-2-phenylindole; A.U. = arbitrary units; s.e.m. = standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/62553fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cellular response analysis; DAPI, PCNA, BrdU staining; immunofluorescence microscopy; chart comparison." class="xfigimg" src="/files/ftp_upload/62553/62553fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Poly(ADP-ribose) polymerase-1 knockdown or inhibition results in increased BrdU foci formation in replicating cells. (A) Representative images of BrdU foci formation and PCNA staining in untreated HeLa cells, treated with 5 µM BMN-673, siCTRL, and siPARP-1, followed by 5 Gy irradiation and 3 h release. A zoomed-in square showing marked BrdU foci is present in the corner of each BrdU image. Scale bars = 5 µm. (B) Quantification of the BrdU nuclear intensity in untreated HeLa cells, treated with 5 µM BMN-673, siCTRL, and siPARP-1 followed by 5 Gy irradiation, the data show the mean ± s.e.m (Mann-Whitney U-test). (C) Western blot to validate the siRNA knockdown of PARP-1. The F1-23 PARP-1 antibody recognizes automodified PARP-1 resulting in the smeared appearance of the siCTRL band and the solid appearance of the catalytically inhibited PARP-1 in BMN-673-treated samples. Abbreviations: PARP-1 = poly(ADP-ribose) polymerase 1; BrdU = 5-bromo-2′-deoxyuridine; PCNA = proliferating cell nuclear antigen; IR = irradiation; DAPI = 4',6-diamidino-2-phenylindole; A.U. = arbitrary units; siCTRL = siRNA control; siPARP-1 = siRNA to knock down PARP-1; s.e.m. = standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/62553fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">Buffer A (Pre-Extraction Buffer) 30 mL</td>
		</tr>
		<tr>
			<td>Reagent</td>
			<td>Volume</td>
			<td>Final Concentration</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>18 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>PIPES (pH 7, 500 mM)</td>
			<td>600 µL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>NaCl (4 M)</td>
			<td>750 µL</td>
			<td>100 mM</td>
		</tr>
		<tr>
			<td>MgCl2 (1 M)</td>
			<td>90 µL</td>
			<td>3 mM</td>
		</tr>
		<tr>
			<td>EGTA (500 mM)</td>
			<td>60 µL</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>Trition X-100 (10%)</td>
			<td>1.5 mL</td>
			<td>0.50%</td>
		</tr>
		<tr>
			<td>Sucrose (1 M)</td>
			<td>9 mL</td>
			<td>300 mM</td>
		</tr>
	</tbody>
</table>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">Buffer B (Cytoskeleton Stripping Buffer) 30 mL</td>
		</tr>
		<tr>
			<td>Reagent</td>
			<td>Volume</td>
			<td>Final Concentration</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>25.035 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris pH 7.5 (1 M)</td>
			<td>300 µL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td> NaCl (4 M)</td>
			<td>75 µL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td> MgCl2 (1 M)</td>
			<td>90 µL</td>
			<td>3 mM</td>
		</tr>
		<tr>
			<td> Tween20 (10%)</td>
			<td>3 mL</td>
			<td>1%</td>
		</tr>
		<tr>
			<td>10% Sodium Deoxycholcate</td>
			<td>1.5 mL</td>
			<td>0.50%</td>
		</tr>
	</tbody>
</table>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>16% PFA</td>
			<td>2.5 mL</td>
		</tr>
		<tr>
			<td> PBS 1x</td>
			<td>7.5 mL</td>
		</tr>
	</tbody>
</table>
Table 1.
Supplemental Figure S1: Visual representation of the Cell Profiler Software and Speckle counting pipeline part 1. (A) Screenshot shows the window used to identify the different channel files of the images to be analyzed. These shots show the cell profiler interface. (B) Screenshot of the Identify Primary Objects menu to identify the Nuclei. The highlighted values are the commonly altered values to best identify nuclei. Abbreviations: BrdU = 5-bromo-2′-deoxyuridine; DAPI = 4',6-diamidino-2-phenylindole. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/supplementalfig01.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Visual representation of the Cell Profiler Software and Speckle counting pipeline part 2. (A) Screenshot of the Identify Primary Objects menu to identify the BrdU foci within the nuclei. (B) Screenshot of Measure Object Intensity menu to measure the BrdU and PCNA nuclear intensity. Abbreviations: BrdU = 5-bromo-2′-deoxyuridine; PCNA = proliferating cell nuclear antigen. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/supplementalfig02.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S3: Visual representation of the Cell Profiler Software and Speckle counting pipeline part 3. (A) Representation of the nuclei identification by cell profiler following optimization. (B) Representation of the BrdU foci identification. (C) Screenshot is a zoom-in on one cell within the initial window. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/supplementalfig03.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S4: Scatter plot of PCNA-positive and PCNA-negative cells. (A) An example of a PCNA scatter plot used to distinguish PCNA-positive from PCNA-negative cells. The PCNA intensity from a single condition was plotted and the distinction between the two populations identified. The green highlights the PCNA-positive cells, the red represents the PCNA-negative cells. This is done for each individual experiment and condition. (B) Quantification of the BrdU nuclear intensity in PCNA-negative untreated HeLa cells, treated with 5 µM BMN-673, siCTRL, and siPARP-1; the data show the mean ± s.e.m (Mann-Whitney U-test). Abbreviations: PARP-1 = poly(ADP-ribose) polymerase 1; BrdU = 5-bromo-2′-deoxyuridine; IR = irradiation; PCNA = proliferating cell nuclear antigen; A.U. = arbitrary units; siCTRL = siRNA control; siPARP-1 = siRNA to knock down PARP-1; s.e.m. = standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/supplementalfig04.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S5: BrdU signal without irradiation. (A) Representative images of BrdU foci formation and PCNA staining without irradiation. Scale bars = 5 µm. A zoomed-in square showing marked BrdU foci is present in the corner of each BrdU image. (B) Quantification of the BrdU nuclear intensity in PCNA-positive untreated Hela cells, treated with 5 µM BMN-673, siCTRL, and siPARP-1; the data show the mean ± s.e.m (Mann-Whitney U-test). (C) Quantification of the BrdU nuclear intensity in PCNA-negative untreated HeLa cells, treated with 5 µM BMN-673, siCTRL, and siPARP-1; the data show the mean ± s.e.m (Mann-Whitney U-test). Abbreviations: PARP-1 = poly(ADP-ribose) polymerase 1; BrdU = 5-bromo-2′-deoxyuridine; PCNA = proliferating cell nuclear antigen; IR = irradiation; DAPI = 4',6-diamidino-2-phenylindole; A.U. = arbitrary units; siCTRL = siRNA control; siPARP-1 = siRNA to knock down PARP-1; s.e.m. = standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/supplementalfig05.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Video 1: Use of Cell Profiler and the Speckle Counting pipeline to analyze BrdU staining for the measurement of DNA resection. This video demonstrates the use of Cell Profiler and the Speckle counting pipeline. It shows how to import and analyze images with this tool specifically for this protocol.Abbreviation: BrdU =5-bromo-2′-deoxyuridine. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62553/62553_R0.mp4" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging at JoVE.com

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

In the present protocol, we demonstrate how to visualize DNA double-strand end resection during S/G2 phase of the cell cycle using an immunofluorescence-based method.

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs ...

assessment-global-dna-double-strand-end-resection-using-brdu-dna

Nanyang Technological University

Research

JoVE Journal

Biology

3.9K Views.  Genome Stability Laboratory, CHU de Québec Research Center, HDQ Pavilion. Dit protocol maakt cellulokwantificering van DNA en resectie na DNA-schade mogelijk. Het belangrijkste voordeel van deze techniek is de celcyclusmarkering die de cellen beperkt tot de S- en G2-fase, waardoor het bromodeoxyuridinesignaal het resultaat is van DNA en resectie. Jean-Yves Masson, een hoofdonderzoeker, en Sofiane Y.Mersaoui, een postdoctoraal onderzoeker, demonstreren de procedure.Werk onder een steriele kweekkap en plaats een enkele dekplaat in elke put van een plaat met ze...

Video: Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

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 ...

Cancer Research

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

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

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

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

Simultaneous Imaging and Flow-Cytometry-based Detection of Multiple Fluorescent Senescence Markers in Therapy-Induced Senescent Cancer Cells

Chemotherapeutic drugs can induce irreparable DNA damage in cancer cells, leading to apoptosis or premature senescence. Unlike apoptotic cell death, senescence is a fundamentally different machinery restraining&#160;propagation of cancer cells. Decades of scientific studies have revealed the complex pathological effects of senescent cancer cells in tumors and microenvironments that modulate cancer cells and stromal cells. New evidence suggests that senescence is a potent prognostic factor during cancer treatment, and therefore rapid and accurate detection of senescent cells in cancer samples is essential. This paper presents a method to visualize and detect therapy-induced senescence (TIS) in cancer cells. Diffuse large B-cell lymphoma (DLBCL) cell lines were treated with mafosfamide (MAF) or daunorubicin (DN) and examined for the senescence marker, senescence-associated &#946;-galactosidase (SA-&#946;-gal), the DNA synthesis marker 5-ethynyl-2&#8242;-deoxyuridine (EdU), and the DNA damage marker gamma-H2AX (&#947;H2AX). Flow cytometer imaging can help generate high-resolution single-cell images in a short period of time to simultaneously visualize and quantify the three markers in cancer cells.

Here we present a flow cytometry-based method for visualization and quantification of multiple senescence-associated markers in single cells.

Chemotherapeutic drugs can induce irreparable DNA damage in cancer cells, leading to apoptosis or premature senescence. Unlike apoptotic cell death, ...

Assessing Cell Cycle Progression of Neural Stem and Progenitor Cells in the Mouse Developing Brain after Genotoxic Stress

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a pseudostratified epithelium lining the lateral ventricles of the embryonic brain. Genotoxic stresses, such as ionizing radiation, have highly deleterious effects on the developing brain related to the high sensitivity of NSPC. Elucidation of the cellular and molecular mechanisms involved depends on the characterization of the DNA damage response of these particular types of cells, which requires an accurate method to determine NSPC progression through the cell cycle in the damaged tissue. Here is shown a method based on successive intraperitoneal injections of EdU and BrdU in pregnant mice and further detection of these two thymidine analogues in coronal sections of the embryonic brain. EdU and BrdU are both incorporated in DNA of replicating cells during S phase and are detected by two different techniques (azide or a specific antibody, respectively), which facilitate their simultaneous detection. EdU and BrdU staining are then determined for each NSPC nucleus in function of its distance from the ventricular margin in a standard region of the dorsal telencephalon. Thus this dual labeling technique allows distinguishing cells that progressed through the cell cycle from those that have activated a cell cycle checkpoint leading to cell cycle arrest in response to DNA damage.
An example of experiment is presented, in which EdU was injected before irradiation and BrdU immediately after and analyzes performed within the 4 hr&#160;following irradiation. This protocol provides an accurate analysis of the acute DNA damage response of NSPC in function of the phase of the cell cycle at which they have been irradiated. This method is easily transposable to many other systems in order to determine the impact of a particular treatment on cell cycle progression in living tissues.

Administration of two analogs of thymidine, EdU and BrdU, in pregnant mice allows the analysis of cell cycle progression in neural and progenitor cells in the embryonic mouse brain. This method is useful to determine the effects of genotoxic stress, including ionizing radiation, during brain development.

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a ...

Neuroscience

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death. Cells repair DSBs using two major pathways: nonhomologous end joining (NHEJ) and homologous recombination (HR). Perturbations of NHEJ and HR are often associated with premature aging and tumorigenesis, hence it is important to have a quantitative way of measuring each DSB repair pathway. Our laboratory has developed fluorescent reporter constructs that allow sensitive and quantitative measurement of NHEJ and HR. The constructs are based on an engineered GFP gene containing recognition sites for a rare-cutting I-SceI endonuclease for induction of DSBs. The starting constructs are GFP negative as the GFP gene is inactivated by an additional exon, or by mutations. Successful repair of the I-SceI-induced breaks by NHEJ or HR restores the functional GFP gene. The number of GFP positive cells counted by flow cytometry provides quantitative measure of NHEJ or HR efficiency.

Analysis of DNA Double-strand Break DSB Repair in Mammalian Cells

This article describes GFP-based fluorescence in vivo assays that separately quantify homologous recombination and nonhomologous end joining in mammalian cells.

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death.  Cells repair DSBs using ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell synchronization can be difficult for many cell systems. Standard practice is to block cell cycle progression at a specific stage and then release the accumulated cells producing a wave of cells progressing through the cycle in unison. However, some cell types find this block toxic resulting in abnormal cell cycling, or even mass death. Bromodeoxyuridine (BrdU) uptake can be used to track the cell cycle stage of individual cells. Cells incorporate this synthetic thymidine analog, while synthesizing new DNA during S phase. By providing BrdU for a brief period it is possible to mark a pool of cells that were in S phase while the BrdU was present. These cells can then be tracked through the remainder of the cell cycle and into the next round of replication, permitting the duration of the cell cycle phases to be determined without the need to induce a potentially toxic cell cycle block. It is also possible to determine and correlate the expression of both internal and external proteins during subsequent stages of the cell cycle. These can be used to further refine the assignment of cell cycle stage or assess effects on other cellular functions such as checkpoint activation or cell death.

This protocol describes the use of bromodeoxyuridine (BrdU) uptake to permit the temporal tracking of cells that were in S phase at a specific point in time. Addition of DNA dyes and antibody labeling facilitates detailed analysis of the fate of the S phase cells at later times.

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell ...

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) has been utilized to investigate replicative/S-phase progression, whereas antibodies against phospho-histone H3 have been utilized to mark mitotic nuclei and cells. A combination of both labels would enable the classification of G0/G1 (Gap phase), S (replicative), and M (mitotic) phases and serve as an important tool to evaluate the effects of mitotic gene knockdowns or null mutants on cell cycle progression. However, the reagents used to mark EdU-labelled cells are incompatible with several secondary antibody-fluorescent tags. This complicates immunostaining, where primary and tagged secondary antibodies are used to mark pH3-positive mitotic cells. This paper describes a step-by-step protocol for the dual-labeling of EdU and pH3 in Drosophila larval neural stem cells, a system utilized extensively to study mitotic factors. Additionally, a protocol is provided for image analysis and quantification to allocate labeled cells in 3 distinct categories, G0/G1, S, S&gt;G2/M (progression from S to G2/M), and M phases.

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Cell cycle analysis with 5-ethynyl-2'-deoxyuridine (EdU) and phospho-histone H3 (pH3) labeling is a multi-step procedure that may require extensive optimization. Here, we present a detailed protocol that describes all steps for this procedure including image analysis and quantification to distinguish cells in different cell cycle phases.

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) ...

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 ...

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

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