The authors thank Michele Pagano for his support and helpful insights, Ashley Chui and Sharon Kaisari for critically reading the manuscript, and Jeffrey Estrada and Vilma Diaz for their continuous support. This work was supported by a diversity supplement to the National Institutes of Health grant GM136250.

1. Maintenance of hTERT-immortalized retinal pigment epithelial cells (RPE1)
<ol>
	<li>Maintain RPE1 cell lines in Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (Hi-FBS) and 100 &#181;g/mL Penicillin-Streptomycin (referred to as culturing medium from now on) in a humidified incubator with 5% CO2 at 37 &#176;C. For routine culturing, grow RPE1 cells in a 15 cm tissue culture-treated dish and split when reaching 80-90% confluency (~16-18 &#215; 106 cells per 15 cm dish).</li>
	<li>When splitting, remove the medium and rinse the cells with 10 mL of 1x phosphate-buffered saline (PBS).</li>
	<li>Add 3 mL of 0.05% Trypsin-EDTA to cover the entire surface of the dish. Keep the cells at 37 &#176;C with the trypsin until they detach.</li>
	<li>After trypsinization, resuspend the cells with culturing medium and spin them down at 150 &#215; g for 5 min at room temperature (RT, 22-25 &#176;C). Remove the supernatant and gently resuspend the cells in 10 mL of culturing medium.</li>
	<li>Seed 1.6-1.8 &#215; 106 cells into a new 15 cm dish (~1 mL of the cell suspension). 
	&#8203;NOTE: All tissue culture work should be done under BSL-2 safety levels. The incubation time for trypsinization depends on cell confluency. Usually, the process takes 2-3 min to complete for a 90% confluent plate. Cells should be screened for Mycoplasma contamination on a regular basis with commercially available kits (see examples in the Table of Materials).</li>
</ol>
2. siRNA knocking down of the gene of interest (GOI)
<ol>
	<li>Seed 1.0 &#215; 106 RPE1 cells into a 10 cm tissue culture treated plate with 10 mL of culturing medium on the day before the transfection.</li>
	<li>On the day of the transfection, complex the siRNA. For a 10 cm plate, use a final concentration of 20 nM siRPA2 and 12 &#181;L of lipid-based transfection reagent in 500 &#181;L of low-serum transfection medium. Gently mix all the components by flicking the tube and incubate at RT (22-25 &#176;C) for 5 min.</li>
	<li>Add the complexed siRNA mixture to the cells dropwise and incubate the cells with the siRNA for 48 h.</li>
</ol>
3. Synchronization of RPE1 cells into G0 phase
<ol>
	<li>Trypsinize the RPE1 cells from step 2.3 as outlined in section 1 (~2 &#215; 106 cells).</li>
	<li>Transfer the cell suspension into 15 mL centrifuge tubes and centrifuge it at 150 &#215; g, RT (22-25 &#176;C) for 5 min.</li>
	<li>Remove the supernatant and resuspend the cells in 12 mL of PBS. Centrifuge the cells at 150 &#215; g at RT (22-25 &#176;C) for 5 min. Repeat the removal of the supernatant and centrifugation twice.</li>
	<li>Resuspend the cells in 10 mL of serum-free DMEM supplemented with 100 &#956;g/mL Penicillin-Streptomycin,&#160;1 mM Sodium Pyruvate, 15 mM HEPES, and plate them onto a 10 cm tissue culture dish. 
	NOTE: If the cells tend to clump, resuspend them in just 1 mL of serum-free DMEM and pipette them up and down 5x using a P1000 tip to dislodge the clumps before diluting the suspension up to a final volume of 10 mL.</li>
	<li>After 24 h of serum starvation, introduce the second round of silencing using the same procedure as described in section 2 by adding the complexed siRNA to the serum-starving cells.</li>
	<li>Keep the RPE1 cells in serum-free DMEM for 72 h before proceeding to G1 release.</li>
</ol>
4. Coverslip coating and releasing cells into G1 phase
<ol>
	<li>Sterilize tweezers with 70% ethanol and place a single glass coverslip (12 mm in diameter and #1.5 thickness [0.17 mm]) in a well of a 24-well plate.</li>
	<li>Dilute the vitronectin coating matrix with PBS to obtain a final concentration of 10 &#181;g/mL. Add 500 &#956;L of the vitronectin solution into each well containing the coverslips and incubate for 1 h at RT.</li>
	<li>Remove the coating solution and wash the coverslips with 1 mL of PBS.</li>
	<li>Detach the serum-starved RPE1 cells from the 10 cm tissue cultured treated plate using 1 mL of 0.05% Trypsin after a PBS wash for 1 min at 37 &#176;C. 
	NOTE: Cells detach much faster after serum starvation. Use caution when washing the cells with PBS and use short trypsinization times.</li>
	<li>To inactivate the trypsin, resuspend the RPE1 cells in a total of 6 mL of culturing medium. Remove the inactivated trypsin by spinning down the cells using 150 &#215; g at RT (22-25 &#176;C) for 5 min.</li>
	<li>Resuspend the cells in 1 mL of culturing medium and measure the cell number.</li>
	<li>Seed 4 &#215; 104 RPE1 cells onto the coated coverslip in a total of 500 &#956;L of culturing medium. 
	NOTE: Make sure cell viability is above 90% before proceeding to downstream steps. Cell viability could be quickly assessed by trypan blue staining during the cell counting step.</li>
	<li>After 6 h of plating the cells into the culture medium, G0-released cells will be in early G1 phase. Perform experiments in G1 in this 6-12 h window before the cells start entering S phase.</li>
	<li>Before introducing DNA damage, pulse the cells with 10 &#956;M 5-ethynyl-2&#39;-deoxyuridine (EdU) for 30 min at 37 &#176;C, diluted in culturing medium.</li>
	<li>Remove the medium containing EdU and chase the cells with 10 &#956;M thymidine for 10 min at 37 &#176;C to prevent remaining EdU incorporation during DNA damage induction.</li>
	<li>Remove the medium with thymidine and treat the cells with 250 &#956;M H2O2 for 1 h, diluted in culturing medium.</li>
</ol>
5. Immunofluorescence staining of ssDNA
<ol>
	<li>Wash cells once with 1 mL of RT (22-25 &#176;C) PBS to remove the medium and serum components. 
	NOTE: Be gentle when washing the cells to avoid detachment and drying. Do not process many wells at the same time.</li>
	<li>Pre-extraction: Incubate the washed cells in 1 mL of CSK extraction buffer (Table 1) for 5 min at RT (22-25 &#176;C). 
	NOTE: CSK pre-extraction removes all the non-chromatin-bound proteins, including soluble RPA2. 
	CAUTION: Triton X-100 is harmful if swallowed and may cause skin irritation and eye damage.</li>
	<li>Remove the CSK buffer from the cells and fix them directly by adding 0.5 mL of 3.6% paraformaldehyde solution (in PBS) containing 0.05% Triton X-100 for 10 min at RT (22-25 &#176;C). 
	CAUTION: It is important to prepare 3.6% PFA from 32% PFA stock freshly. Paraformaldehyde may cause serious eye damage, skin irritation, and respiratory irritation.</li>
	<li>Wash the cells once with 1 mL of PBS containing 0.05% Triton X-100 to remove the PFA.</li>
	<li>Further permeabilize cells using 1 mL of PBS containing 0.5% Triton X-100 for 15 min at RT (22-25 &#176;C).</li>
	<li>EdU click-IT reaction to visualize replicating cells (S phase)
	<ol>
		<li>Remove the permeabilization solution and wash the cells 2x using 1 mL of blocking buffer (Table 1). 
		CAUTION: Bovine serum albumin (BSA) may cause respiratory irritation.</li>
		<li>Add 1 mL of blocking buffer (Table 1) and gently rock the coverslip-containing plate for 10 min at RT (22-25 &#176;C).</li>
		<li>Remove the blocking buffer and add 500 &#956;L of click reaction cocktail containing picolyl azide 647 (Table 1). Incubate the coverslips for 30 min at RT (22-25 &#176;C) using gentle rocking and perform downstream incubations in the dark. 
		NOTE: When using BrdU antibodies, use double the amount (1 mL) and time (60 min) for the click-reaction as recommended by the manufacturer to ensure that the reaction is saturated and the incorporated EdU is labeled. This limits the cross-reactivity of the BrdU antibodies27.</li>
	</ol>
	</li>
	<li>Remove the click reaction mixture and wash the cells 2x with PBS with 0.05% Triton X-100 for 10 min at RT (22-25 &#176;C) (Figure 1 and Figure 2).</li>
	<li>Add 1 mL of blocking buffer and incubate at RT (22-25 &#176;C) for 30 min. Alternatively, keep the cells in blocking buffer at 4 &#176;C overnight.</li>
	<li>Apply primary antibody (anti-RPA2 rat, 1:1,000 dilution) for 2 h at RT (22-25 &#176;C) in 250-500 &#956;L of blocking buffer with gentle rocking.</li>
	<li>Wash the cells 2x with PBS containing 0.05% Triton X-100 to quickly remove most of the antibody solution.</li>
	<li>Continue washing the cells for 3 x 10 min with blocking buffer at RT (22-25 &#176;C).</li>
	<li>Apply secondary antibody (anti-rat Alexa-488, 1:1,000 dilution) in 250-500 &#956;L of blocking buffer at RT (22-25 &#176;C) for 2 h with gentle rocking.</li>
	<li>Wash the cells with blocking buffer 2x to quickly remove most of the secondary antibody. Continue washing the cells for 3 x 10 min with PBS containing 0.05% Triton X-100 at RT (22-25 &#176;C).</li>
	<li>To counterstain the nuclei, wash cells once with PBS containing 0.05% Triton X-100 and 1 &#956;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI) for 10 min at RT (22-25 &#176;C). Wash cells once with PBS for 5 min at RT (22-25 &#176;C).</li>
	<li>Mount the cover glass onto microscope slides using 10 &#956;L of mounting medium/coverslip. Dip the coverslips into distilled water before mounting to get rid of any salt crystals. Image the slides on the following day and store them at 4 &#176;C for weeks (Figure 3).</li>
</ol>
6. Image acquisition and quantification
<ol>
	<li>To capture images, use any available epifluorescent microscope equipped with routine filter sets to image DAPI, FITC, and Cy5 channels with at least 60-63x magnification, high numeric aperture, and oil objectives to visualize nuclear foci. 
	NOTE: Optimal DAPI excitation is ~359 nm; Alexa 488 excitation is ~488 nm; while Alexa 647 excitation is ~647 nm.</li>
	<li>For image analysis, open image files in Fiji/ImageJ.
	<ol>
		<li>Make nuclear masks using the DAPI staining (Figure 4A-F and Supplementary Video S1).
		<ol>
			<li>Open the DAPI image.</li>
			<li>Select Process | Enhance Contrast and set saturated pixel to 0.35.</li>
			<li>Click Process | Binary | Convert to Mask. Choose Binary | Fill Holes and click on Analyze | Analyze Particles. Set the size to 10-Infinity.</li>
			<li>In ROI manager, click on Show all.</li>
		</ol>
		</li>
		<li>Finding RPA2 foci in the nucleus (Figure 4G,H and Supplementary Video S1)
		<ol>
			<li>Open the RPA2 image.</li>
			<li>Choose Process | Find Maxima. Set the prominence to a value that highlights the RPA2 foci (between 500 and 750), separating it from the background.</li>
			<li>Finally, click the Measure button in ROI Manager.</li>
			<li>Calculate the total number of nuclear ssDNA foci by dividing the value in the RawinDen column by 255 (the maximum value of pixel intensity in each foci).</li>
		</ol>
		</li>
		<li>Perform statistical analysis using the preferred statistical software tool. 
		NOTE: Exclude all EdU-positive cells and improperly segmented DAPI masks from the analysis.</li>
	</ol>
	</li>
</ol>

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

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I., Haaf, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nuclear+foci+of+mammalian+recombination+proteins+are+located+at+single-stranded+DNA+regions+formed+after+DNA+damage.">Nuclear foci of mammalian recombination proteins are located at single-stranded DNA regions formed after DNA damage.</a> Proc Natl Acad Sci U S A. 96 (5), 1921-1926 (1999).</li><li>Forment, J. V., Walker, R. V., Jackson, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+high-throughput,+flow+cytometry-based+method+to+quantify+DNA-end+resection+in+mammalian+cells.">A high-throughput, flow cytometry-based method to quantify DNA-end resection in mammalian cells.</a> Cytometry A. 81 (10), 922-928 (2012).</li><li>Mistrik, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cells+and+stripes:+A+novel+quantitative+photo-manipulation+technique.">Cells and stripes: A novel quantitative photo-manipulation technique.</a> Sci Rep. 6, 19567 (2016).</li><li>Aten, J. A., Bakker, P. J., Stap, J., Boschman, G. A., Veenhof, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+double+labelling+with+IdUrd+and+CldUrd+for+spatial+and+temporal+analysis+of+cell+proliferation+and+DNA+replication.">DNA double labelling with IdUrd and CldUrd for spatial and temporal analysis of cell proliferation and DNA replication.</a> Histochem J. 24 (5), 251-259 (1992).</li><li>Podgorny, O., Peunova, N., Park, J. H., Enikolopov, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Triple+S-phase+labeling+of+dividing+stem+cells.">Triple S-phase labeling of dividing stem cells.</a> Stem Cell Reports. 10 (2), 615-626 (2018).</li><li>Cappella, P., Gasparri, F., Pulici, M., Moll, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+proliferation+method:+click+chemistry+based+on+BrdU+coupling+for+multiplex+antibody+staining.">Cell proliferation method: click chemistry based on BrdU coupling for multiplex antibody staining.</a> Curr Protoc Cytom. Chapter 7, (2008).</li><li>Ligasova, A., Koberna, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strengths+and+weaknesses+of+cell+synchronization+protocols+based+on+inhibition+of+DNA+synthesis.">Strengths and weaknesses of cell synchronization protocols based on inhibition of DNA synthesis.</a> Int J Mol Sci. 22 (19), 10759 (2021).</li></ol>

The authors have no competing interests to declare.

Maintaining a healthy, mycoplasma-free cell culture is critical for all the experiments described above. RPE1 cells have a strong attachment to tissue culture-treated plasticware under normal culturing media; however, their binding characteristics significantly diminish when kept in serum-free conditions. Additionally, to capture high-resolution images of ssDNA foci under a microscope, the cells need to be plated onto a 0.17 mm thick cover glass, which is not hydrophilic enough to support proper attachment of RPE1 cells. Without properly flattened and evenly distributed cells, it is very challenging to visualize individual ssDNA foci. Therefore, it is critical to choose the proper coating material (e.g., vitronectin) and to leave adequate time (6-12 h) for the cells to spread and attach after releasing them into G1 phase.
A challenging part of the protocol is to obtain homogeneous G1 synchronized RPE1 cells. This requires two critical steps. First, for efficient serum starvation, the cells need to be trypsinized, washed thoroughly with PBS, and directly seeded onto new tissue culture dishes using serum-free media. Washing the cells directly in tissue culture dishes to remove the serum will not yield efficient G0 synchronization. Second, when releasing cells into G1 phase, the cells must be trypsinized again and seeded onto fresh tissue culture plates. Similarly, just changing the medium and adding serum-containing culturing medium to the cells will not result in a synchronous G1 entry. Additionally, for proper G1 entry, the seeding density of the cells on the coated cover glasses must be at certain confluency levels. While perfect cell synchronization is generally unachievable, this synchronization protocol described here gives a ~97% pure G1 population. The recommended seeding density for RPE1 on a 12 mm diameter coverslip is ~4 &#215; 104 to acquire a homogeneous field of view for imaging, with approximately 70% confluency. Higher seeding density causes cells to detach and &#34;peel-off&#34; after CSK extraction and will result in a higher background signal during image acquisition.
To reduce any background signal and achieve a favorable signal-to-noise ratio, thorough washing after primary and secondary antibody incubation is essential. Since numerous washing steps are to be applied, it is also essential to prevent the well from drying out during each washing step. We minimize this artifact by applying a minimum of 0.05% Triton X-100 in all the washing and incubation steps. Once the wells dried out, the cells displayed an altered signal-to-noise ratio; this leads to a mosaic-like pattern under the microscope and could interfere with evaluation. Z-stack image acquisition combined with deconvolution can assist in capturing foci in different focal planes to improve the analysis.
Conventional methods rely on the detection of incorporated BrdU under non-denaturing conditions. These methods, however, depend on the pretreatment of the cells with high dosages of BrdU for at least 1-2 days (or time equivalent to a full cell cycle in the cell line used) to ensure uniform genomic incorporation. Undesirably, extensive BrdU incorporation can cause cell cycle interference36. To address these limitations, this method utilizes endogenous RPA2 to detect ssDNA foci. This approach does not require replication-driven BrdU incorporation, it can also be used in post-mitotic cells. Since extensive BrdU incorporation is not needed, this saves time and reduces experimental complexity. By using RPA2 staining to visualize ssDNA, we can use 2&#8242;-deoxy-5-ethynyluridine (EdU) and click-chemistry to mark DNA replication while avoiding possible crossreactivity of the BrdU antibodies against EdU27,37,38. Special care must be taken to properly mask the incorporated EdU during the click-reaction so that the BrdU antibodies will not cross-react with EdU27,39.
Finally, an important benefit of utilizing RPA2 instead of BrdU is simply having a superior signal-to-noise ratio when compared to BrdU staining outside S phase. We found that the non-denaturing BrdU staining and its ability to visualize ssDNA is restricted to S phase even in replicating cells (Figure 2). BrdU antibody binds only to the sufficiently exposed BrdU in ssDNA stretches. The binding of repair proteins, including RPA2, to the ssDNA stretches may suppress or hamper the sufficient exposure of BrdU in ssDNA. We also found that CSK pre-extraction is necessary for ssDNA visualization using BrdU antibody. This is possible because the ssDNA tracks are not accessible for the antibody without removing lightly bound protein components from them.
Nonetheless, there are some limitations associated with this protocol. A limitation of using RPA2 for ssDNA detection is the need to optimize the CSK pre-extraction step. Unbound, excess RPA2 must be washed away from the DNA before fixing the cells. On the one hand, underextraction leads to a high background due to the RPA2 protein fraction that is not bound to ssDNA. On the other hand, overextraction will lead to signal loss. For BrdU detection, this is not a variable since BrdU is stably incorporated into the DNA and cannot be washed away by pre-extraction. Therefore, the time of the CSK pre-extraction, the amount of Triton X-100 in the buffer, the volume, and the temperature at which the pre-extraction is performed must be carefully considered. CSK pre-extraction also limits the use of nucleus size to discriminate G0/G1 cells from S/G2 cells.
Additionally, we cannot exclude the possibility that some of the signal that comes from RPA2 originates from it being bound to other chromatin-binding protein interactors. One also must consider the species specificity of the RPA2 antibody. The antibody used in this protocol can recognize human, mouse, rat, hamster, and monkey RPA2. Another limitation of this approach is that not all cell lines can be serum-starved for G0 synchronization. Most cancer cell lines can bypass cell cycle checkpoints and proliferate even in serum-deprived media. Though serum starvation is beneficial, since it does not cause DNA damage, one must carefully monitor their cell synchronization efficiency to make sure that proper cell cycle phase enrichment is achieved. For cells that do not respond to serum deprivation, other cell synchronization methods must be considered (e.g., mitotic shake off, CDK1 inhibition for G2 arrest, or non-invasive techniques such as centrifugal elutriation). Another possible method is using high-content imaging to measure EdU and nuclear DNA content for cell cycle profiling of asynchronous cells31. One must consider the implications of utilizing alternate synchronization methods to prevent interference with downstream analysis. For instance, the use of double thymidine block or aphidicolin, often used in literature, will result in replication stress and DNA damage40.
The investigation of DNA repair mechanisms continues to be a focal point of discussion in the fields of cancer and cell biology. The protocol presented here offers a valuable approach for the preparation of cells, enabling the visualization and quantitative analysis of ssDNA upon exposure to DNA-damaging agents. Notably, this protocol highlights the utilization of the ssDNA binding protein, RPA2, demonstrating its high specificity to visualize small amounts of ssDNA foci while avoiding unwanted cross-reactivity in all the cell cycle phases. Using RPA2 confers numerous advantages, particularly for researchers aiming to analyze cells in the G1 phase of the cell cycle. This protocol considers several limitations and addresses concerns related to signal interference, undesired background noise, and cross-reactivity when using RPA2 or BrdU staining to detect ssDNA.

To sustain life, cells constantly survey and repair DNA to maintain their genomic integrity. Cells may accumulate various types of DNA damage due to both endogenous (e.g., oxidation, alkylation, deamination, replication errors) and exogenous (e.g., UV, ionizing irradiation) sources of DNA stressors. Failure to repair these lesions results in either apoptosis, cell cycle arrest, or senescence and can lead to diseases1. DNA lesions could be addressed by any of the following main DNA repair pathways: DR (direct reversal repair), which mainly repairs alkylated bases2; BER (base excision repair), which targets non-bulky DNA bases errors and single-stranded DNA breaks (SSBs)3; NER (nucleotide excision repair) correcting bulky, helix-distorting DNA lesions4; MMR (mismatch repair) mainly targeting DNA mismatches, insertion/deletion loops (IDLs), and certain base damages5; NHEJ (non-homologous end joining) and HRR (homologous recombination repair) that are both active at double-stranded DNA breaks (DSBs)6; and TLS (translesion synthesis), which is a DNA lesion bypass mechanism7. Though these pathways have distinct substrate specificities, there are certain overlaps amongst them to ensure redundancy for efficient repair. Understanding the action of the different DNA repair pathways in various cell cycle phases is crucial as these DNA repair factors could serve as essential targets for therapeutic approaches to treat cancer, aging, and neurological disorders8,9.
Single-stranded DNA (ssDNA) is generated throughout the cell cycle due to the repair of DNA lesions generated by both endogenous and exogenous DNA-damaging agents. Upon genotoxic stress, ssDNA is generated abundantly in the S and G2 phases where HRR and MMR have their highest activity and when the replication machinery stalls or collapses when encountering DNA lesions6,10,11. Other DNA repair pathways (e.g., NHEJ/microhomology-mediated end joining (MMEJ)/single-strand annealing [SSA]) also generate ssDNA during DSB repair12. These ssDNA tracks usually arise from DNA resection, carried out by exonucleases such as EXO1, DNA2, and CtIP during HR and MMR, endonucleases such as XPF and XPG during NER, or through the combined action of POLB and FEN1 during BER4,13,14,15,16,17,18,19. Due to the work of the replication machinery, ssDNA tracks are also generated when the DNA helicases unwind DNA in front of the PCNA-bound replicative polymerases20. In contrast, in the G1 phase, the lack of HRR and DNA replication and the limited activity of MMR reduces the extent of ssDNA tracks generated and are therefore more challenging to detect10,11,21.
Cellular ssDNA tracks are highly sensitive structures that must be protected to avoid the formation of DSBs. This is achieved by coating the ssDNA tracks with RPA. RPA is an abundant heterotrimeric protein complex composed of multiple subunits (RPA1, RPA2, and RPA3, also referred to as RPA70, RPA32, and RPA14, respectively), which are ubiquitously expressed throughout the cell cycle22. Each RPA subunit contains a DNA-binding domain (DBD), capable of interacting with 4-6 nucleotides, and the combined subunits form a stable trimerization core. Altogether, RPA binds to approximately 20-30 nucleotides with sub-nanomolar affinity23,24.
Conventional methods use immunofluorescence (IF) microscopy to visualize ssDNA foci by labeling 5-bromo-2&#39;-deoxyuridine (BrdU) incorporated into genomic DNA using BrdU antibodies25. This approach relies on the fact that BrdU antibodies can only detect BrdU in exposed ssDNA25. Although this approach is straightforward, it also displays certain limitations. For instance, cells are pretreated to incorporate BrdU before the start of the experiment, which is time-consuming and can interfere with downstream effectors. Therefore, BrdU-based ssDNA detection is limited to replicating cells and cannot be used for quiescent cells. This excludes the application of this method to study DNA repair in non-replicating cells despite its importance in several diseases such as cancer and neurodegeneration5,26. Additionally, because the structures of BrdU and EdU are very similar, most BrdU antibodies display crossreactivity towards EdU, which must be considered when aiming for dual labeling experiments27. RPA staining has been previously utilized to show ssDNA foci mainly in S phase cells; however, some papers have also successfully used it outside the S phase28,29,30,31,32,33,34,35. The following protocol efficiently utilizes the properties of RPA, allowing the visualization of ssDNA foci following DNA damage in the G1 phase of the cell cycle (although it can be used in all cell cycle phases).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alpha-tubulin antibody</td>
			<td>Sigma-Aldrich</td>
			<td>T6074</td>
			<td>primary antibody (1:5,000)</td>
		</tr>
		<tr>
			<td>Axio Observer Inverted Microscope</td>
			<td>Zeiss</td>
			<td>na</td>
			<td>microscope</td>
		</tr>
		<tr>
			<td>Bis-Tris Plus Mini Protein Gels, 4-12%</td>
			<td>Invitrogen&#160;</td>
			<td>NW04127BOX</td>
			<td>Western Blot</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin&#160;</td>
			<td>Jackson ImmunoResearch&#160;</td>
			<td>001-000-162</td>
			<td>blocking</td>
		</tr>
		<tr>
			<td>BrdU (5-Bromo-2&#39;-deoxyuridine)&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>B5002-100MG</td>
			<td>nucleotide analogue</td>
		</tr>
		<tr>
			<td>BrdU antibody BU1/75</td>
			<td>Abcam</td>
			<td>ab6326</td>
			<td>primary antibody (1:500)</td>
		</tr>
		<tr>
			<td>CellAdhere Dilution Buffer</td>
			<td>Stemcell Technologies</td>
			<td>07183</td>
			<td>coating reagent</td>
		</tr>
		<tr>
			<td>Click-iT Plus EdU Flow Cytometry Assay Kits&#160;</td>
			<td>Invitrogen&#160;</td>
			<td>C10632</td>
			<td>flow cytomery&#160;</td>
		</tr>
		<tr>
			<td>Click-iT Plus EdU Cell Proliferation Kit for Imaging, Alexa Fluor 647 dye</td>
			<td>Thermo Fisher Scientific</td>
			<td>C10640</td>
			<td>click-reaction kit</td>
		</tr>
		<tr>
			<td>cOmplete ULTRA Protease inhibitor tablets</td>
			<td>Sigma-Aldrich</td>
			<td>5892791001</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Countess 3 Automated cell counter&#160;</td>
			<td>Thermo Scientific</td>
			<td>AMQAX2000</td>
			<td>cell counter</td>
		</tr>
		<tr>
			<td>Coverslip&#160;</td>
			<td>neuVitro</td>
			<td>GG12PRE</td>
			<td>tissue culture</td>
		</tr>
		<tr>
			<td>Cyclin A2 antibody</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-271682</td>
			<td>primary antibody (1:1,000) for IF and WB</td>
		</tr>
		<tr>
			<td>Cyclin B1 antibody</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-245</td>
			<td>primary antibody (1:5,000)</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma-Aldrich</td>
			<td>D2650-100ML</td>
			<td>vehicle control</td>
		</tr>
		<tr>
			<td>DMEM, high glucose, with HEPES</td>
			<td>Gibco</td>
			<td>12430051</td>
			<td>cell culture medium for RPE cells</td>
		</tr>
		<tr>
			<td>DPBS, no calcium, no magnesium&#160;</td>
			<td>Gibco</td>
			<td>14190144</td>
			<td>the PBS used throughout the protocol</td>
		</tr>
		<tr>
			<td>D-Sucrose</td>
			<td>Thermo Fisher Scientific</td>
			<td>bp220-1</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Eclipse Ti2 Series Epifluorescent Microscope&#160;</td>
			<td>Nikon</td>
			<td>na</td>
			<td>microscope</td>
		</tr>
		<tr>
			<td>EdU (5-Ethynyl-2&#39;-deoxyuridine)&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>C10637</td>
			<td>nucleotide analog</td>
		</tr>
		<tr>
			<td>Falcon 24-well plate</td>
			<td>Corning&#160;</td>
			<td>351147</td>
			<td>tissue culture</td>
		</tr>
		<tr>
			<td>Falcon Cell Culture Dishes 100 mm</td>
			<td>Corning&#160;</td>
			<td>353003</td>
			<td>tissue culture</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum, heat inactivated</td>
			<td>Gibco</td>
			<td>16140071</td>
			<td>media supplement</td>
		</tr>
		<tr>
			<td>Fiji (ImageJ)</td>
			<td>NIH</td>
			<td>version 1.54f</td>
			<td>software and algorithms</td>
		</tr>
		<tr>
			<td>FxCycle PI/RNase Staining Solution</td>
			<td>Invitrogen&#160;</td>
			<td>F10797</td>
			<td>PI staining</td>
		</tr>
		<tr>
			<td>Goat anti-mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor Plus 555</td>
			<td>Thermo Fisher Scientific</td>
			<td>A21422</td>
			<td>secondary antibody (1:1,000)</td>
		</tr>
		<tr>
			<td>Goat anti-rat IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor Plus 488</td>
			<td>Thermo Fisher Scientific</td>
			<td>A48262</td>
			<td>secondary antibody (1:1,000)</td>
		</tr>
		<tr>
			<td>Histone H3 antibody</td>
			<td>Abcam</td>
			<td>ab1791</td>
			<td>primary antibody (1:10,000)</td>
		</tr>
		<tr>
			<td>hTERT RPE1</td>
			<td>ATCC</td>
			<td>CRL-3216</td>
			<td>cell line</td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td>Sigma-Aldrich</td>
			<td>H1758-100ML</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide 30% soultion</td>
			<td>Sigma-Aldrich</td>
			<td>H1009-100ML</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Hydroxyurea,98% powder</td>
			<td>Sigma-Aldrich</td>
			<td>H8627-5G</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Invitrogen Ultra Pure 0.5 M EDTA pH 8.0</td>
			<td>Thermo Fisher Scientific</td>
			<td>15-575-020</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Lipfectamine RNAiMAX Transfection Reagent</td>
			<td>Invitrogen&#160;</td>
			<td>13778150</td>
			<td>transfection reagent</td>
		</tr>
		<tr>
			<td>Magnesium chloride solution 1 M</td>
			<td>Sigma-Aldrich</td>
			<td>M1028-100ML</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>MycoFluor</td>
			<td>Thermo Fisher</td>
			<td>M7006</td>
			<td>Mycoplasma Detection Kit</td>
		</tr>
		<tr>
			<td>Neocarzinostatin from Streptomyces carzinostaticus&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>N9162-100UG</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>NuPage MES SDS Running Buffer (20x)</td>
			<td>Invitrogen&#160;</td>
			<td>NP0002</td>
			<td>Western Blot</td>
		</tr>
		<tr>
			<td>onTARGETplus Human RPA2 siRNA</td>
			<td>Dharmacon</td>
			<td>L-017058-01-0005</td>
			<td>siRNA</td>
		</tr>
		<tr>
			<td>p27 antibody</td>
			<td>BD Biosciences&#160;</td>
			<td>610241</td>
			<td>primary antibody (1:1,000)</td>
		</tr>
		<tr>
			<td>Paraformaldehyde aqueous solution (32%)</td>
			<td>Electron Microscopy Sciences</td>
			<td>50-980-494</td>
			<td>fixative</td>
		</tr>
		<tr>
			<td>PARP1 antibody</td>
			<td>Cell Signaling Technology&#160;</td>
			<td>9542S</td>
			<td>primary antibody (1:1,000)</td>
		</tr>
		<tr>
			<td>PCNA antibody</td>
			<td>Cell Signaling Technology&#160;</td>
			<td>13110S</td>
			<td>primary antibody (1:2,000)</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140163</td>
			<td>media supplement</td>
		</tr>
		<tr>
			<td>pH3 antibody</td>
			<td>Cell Signaling Technology</td>
			<td>3377S</td>
			<td>primary antibody (1:2,000)</td>
		</tr>
		<tr>
			<td>PhosSTOP phosphatase inhibitor tablets</td>
			<td>Sigma-Aldrich</td>
			<td>4906837001</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>PIPES Buffer 0.5 M solution, pH 7.0</td>
			<td>Bioworld</td>
			<td>41620034-1</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Precision Plus Protein Kaleidoscope Prestained Protein Standards</td>
			<td>Bio-Rad</td>
			<td>1610395</td>
			<td>Western Blot</td>
		</tr>
		<tr>
			<td>Prism</td>
			<td>GraphPad</td>
			<td>version 10</td>
			<td>statistical analysis and graph</td>
		</tr>
		<tr>
			<td>ProLong Diamond Antifade Mountant</td>
			<td>Thermo Scientific</td>
			<td>P36961</td>
			<td>mounting media</td>
		</tr>
		<tr>
			<td>Reduced serum media (Opti-MEM)</td>
			<td>Gibco</td>
			<td>31985070</td>
			<td>used for transfection</td>
		</tr>
		<tr>
			<td>Rpa32/rpa2 antibody (mouse)</td>
			<td>EMD Millipore</td>
			<td>NA19L</td>
			<td>primary antibody (1:1,000) for WB</td>
		</tr>
		<tr>
			<td>Rpa32/rpa2 antibody (rat)</td>
			<td>Cell Signaling Technology&#160;</td>
			<td>2208S</td>
			<td>primary antibody (1:1,000) for IF</td>
		</tr>
		<tr>
			<td>Sodium Chloride solution (5 M)</td>
			<td>Sigma-Aldrich</td>
			<td>S5150</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Sodium Pyruvate (100 mM)</td>
			<td>Gibco</td>
			<td>11360070</td>
			<td>media supplement</td>
		</tr>
		<tr>
			<td>Sodium tetraborate decahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>B3535-500G</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Thermo Scientific Pierce DAPI Nuclear Counterstain&#160;</td>
			<td>Thermo Scientific</td>
			<td>62248</td>
			<td>nucleic acid stain</td>
		</tr>
		<tr>
			<td>Thymidine,powder</td>
			<td>Sigma-Aldrich</td>
			<td>T1985-1G</td>
			<td>reagent</td>
		</tr>
		<tr>
			<td>Triton X-100 aqueous solution (10%)</td>
			<td>Sigma-Aldrich</td>
			<td>11332481001</td>
			<td>detergent</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.5%), no phenol red</td>
			<td>Gibco</td>
			<td>1540054</td>
			<td>cell dissociation agent</td>
		</tr>
		<tr>
			<td>Vitronectin XF</td>
			<td>Stemcell Technologies</td>
			<td>07180</td>
			<td>coating reagent</td>
		</tr>
		<tr>
			<td>ZE5 Cell Analyzer</td>
			<td>Bio-Rad</td>
			<td>na</td>
			<td>flow cytomery&#160;</td>
		</tr>
	</tbody>
</table>

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.

To overcome the limitations of detecting ssDNA in G1, we utilized RPA2, which enhances both the specificity and intensity of ssDNA foci detection35. To achieve precise cell synchronization, we used RPE1 cells that can be efficiently serum-starved and synchronized into G0 phase. They can then be induced to re-enter the cell cycle by the addition of serum following serum deprivation. To confirm the synchronization efficiency, we labeled the cells with EdU and their DNA content with propidium-iodide. We further gathered qualitative and quantitative results via flow cytometry (Supplementary Figure S1A). The dot-plots show that after 72 h of serum starvation, ~98% of the cells are in G0 phase. Following the addition of serum-containing media for 6 h, cells reenter the cell cycle (as seen by the increase in p27 levels in Figure 1A), having ~97% of cells in G1, while only having &#60;1% cells in S phase, &#60;2% cells in G2 phase (Supplementary Figure S1A). After 20-28 h of addition of serum to the cells, they gradually pass through S phase, as shown by the flow cytometry plots (Supplementary Figure S1A). This cell synchronization protocol gives a ~97% pure G1 population (6 h post serum addition following 72 h of serum starvation). To further validate the synchronization efficiency, we compared the expression of cell cycle markers following serum release using western blotting (Figure 1A and Supplementary Figure S1B) and in parallel, performed an EdU incorporation assay to visualize DNA replication. The EdU staining also highlights synchronization efficiency and the lack of DNA replication in G1 phase (Figure 1B,C).
Conventional methods to detect ssDNA in mammalian cells rely on the detection of BrdU in ssDNA. Figure 2A demonstrates that upon H2O2 and neocarzinostatin (NCS) treatment, the BrdU foci were detectable only in S phase cells, while no ssDNA foci were detectable in non-S phase cells. The BrdU antibody staining also showed a noticeable nucleolar background staining that could be detected in all the nuclei, independent of the cell cycle stage or treatments applied. Using the EdU click protocol described here, we could not detect co-localizing EdU and BrdU foci, which is evident in the untreated samples of Figure 2A. To completely rule out any BrdU signal that arose from cross-reactivity, we avoided EdU labeling and rather used cyclin A2 as an S-G2 marker. However, cyclin A2 staining did not allow CSK pre-extraction, and under this condition, we did not see any BrdU foci, even after genotoxic stress (Supplementary Figure S2A). This highlights the fact that CSK pre-extraction is necessary for anti-BrdU-based ssDNA staining. As a control, we tested BrdU antibody staining under denaturing conditions. This opens the DNA to expose the incorporated BrdU, which reveals that BrdU was uniformly incorporated (Supplementary Figure S2B).
In contrast, RPA2 staining shows NCS- and H2O2-dependent foci formation not only in the S phase but also in other cell cycle phases (Figure 2B). As a control, we also treated the cells with HU, which only causes ssDNA accumulation in cells undergoing replication. As expected, we only detected a signal increase upon HU treatment with the RPA2 antibody in EdU-positive cells, highlighting the specificity of this approach. The RPA2 antibody can also detect naturally occurring ssDNA formation during replication in the absence of exogenous genotoxic stress (Figure 2B). The highly sensitive nature of the RPA2 antibody prompted us to try to utilize it in the G1 phase where conventional BrdU staining failed to detect any signal upon genotoxic stress (Supplementary Figure S2C). Figure 3A shows that the formation of ssDNA foci upon H2O2 treatment was detectable when using an anti-RPA2 antibody, even in G1. There was a significant increase in the number of RPA2 foci in these nuclei upon H2O2 treatment (Figure 3B). These foci were specific to RPA2 as silencing of RPA2 abolished the IF signal (Figure 3A,B). Figure 3C and Supplementary Figure S1C show the efficiency of RPA2 silencing in these cells. Compared to conventional methods, RPA2-based detection of ssDNA is highly sensitive, and its application can therefore be extended to G1 phase cells.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65926/65926fig01.jpg" /> 
Figure 1: Synchronization efficiency of RPE1 cells following serum starvation. (A) Immunoblots show indicated protein levels in asynchronous, G1, and S phase synchronized RPE1 cells. (B) Representative images show asynchronous, G1, and S phase synchronized RPE1 cells that were exposed to 10 &#181;M EdU for 30 min before fixation and visualized by Click-IT reaction. DAPI was used to counterstain nuclear DNA. Scale bars = 50 &#181;m. (C) Graph shows percentage of EdU-positive cells over the total cell population assessed by DAPI. The error bar represents standard error of mean, and the analyzed numbers of nuclei were the following: AS n = 219, G1 n = 630, S n = 437. Abbreviations: RPE1 = hTERT-immortalized retinal pigment epithelial cells; AS = asynchronous; EdU = 5-ethynyl-2&#39;-deoxyuridine; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www.jove.com/files/ftp_upload/65926/65926fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65926/65926fig02.jpg" /> 
Figure 2: ssDNA detection with either BrdU antibody or RPA2 antibody upon DNA damage. (A) Representative images illustrate ssDNA foci using &#945;BrdU (green), S phase cells are highlighted by EdU (purple), and DAPI was used to counterstain nuclear DNA (blue). RPE1 cells were kept in 10 &#181;M BrdU for 48 h prior to any additional treatment. After 48 h, cells were pulsed with 10 &#181;M EdU for 30 min followed by treatment of H2O2 (250 &#956;M) for 1 h or Neocarzinostatin (0.5&#160;&#956;g/mL) for 4 h. Cells were fixed after CSK pre-extraction. A white dashed line denotes the border of each nucleus. Scale bar = 5 &#181;m. Panels on the right are enlarged images of the indicated S phase or non-S phase nuclei. (B) Representative images illustrate ssDNA foci using &#945;RPA2 antibody (green). S phase cells are highlighted by EdU (purple), and DAPI was used to counterstain nuclear DNA (blue). RPE1 cells were pulsed with 10 &#181;M EdU for 30 min followed by either 1 h H2O2 (250 &#956;M), 4 h of Hydroxyurea (2 mM), or 4 h of NCS (0.5 &#160;&#956;g/mL). Cells were fixed after CSK pre-extraction. A white dashed line denotes the border of each nucleus. Scale bar = 10 &#181;m. Panels on the right are enlarged images of the indicated S phase or non-S phase nuclei. Abbreviations: ssDNA = single-stranded DNA; BrdU = 5-bromo-2&#39;-deoxyuridine; DAPI = 4&#39;,6-diamidino-2-phenylindole; RPE1 = hTERT-immortalized retinal pigment epithelial cells; EdU = 5-ethynyl-2&#39;-deoxyuridine; NCS = Neocarzinostatin; HU = hydroxyurea. <a href="https://www.jove.com/files/ftp_upload/65926/65926fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65926/65926fig03.jpg" /> 
Figure 3: Detection of ssDNA foci in G1 phase using RPA2 antibody. (A)&#160;RPE1 cells were transfected with either siRNAs targeting RPA2 or a non-targeting siRNA control, and subsequently synchronized in G1 and pulse-labeled with 10 &#181;M EdU for 30 min before treating them with H2O2 (250 &#956;M) for 1 h where indicated. DAPI was used to counterstain nuclear DNA. Cells were fixed after CSK pre-extraction. A white dashed line denotes the border of each nucleus. Scale bar = 5 &#181;m. (B) The measurements for the number of RPA2 foci/nucleus were carried out from two independent experiments. Only EdU-negative cells were considered during the analysis. Lines represent the mean value on the plots. Non-parametric ANOVA test (Kruskal-Wallis) was performed for statistical analysis. The stars indicate P &#60; 0.0001. The analyzed number of nuclei were the following: siNT no H2O2 n = 513, siNT H2O2 n = 603, siRPA2 no H2O2 n = 266, siRPA2 H2O2 n = 536. (C) The efficiency of the siRNA knockdown is shown in the immunoblotting. Abbreviations: siNT = non-targeting siRNA control; BrdU = 5-bromo-2&#39;-deoxyuridine; DAPI = 4&#39;,6-diamidino-2-phenylindole; RPE1 = hTERT-immortalized retinal pigment epithelial cells; EdU = 5-ethynyl-2&#39;-deoxyuridine. <a href="https://www.jove.com/files/ftp_upload/65926/65926fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65926/65926fig04.jpg" /> 
Figure 4: Quantification of ssDNA foci using Fiji. Detailed steps in Fiji showing how to assess RPA2 foci numbers in the nucleus. (A-E) The creation of a nuclear mask using the DAPI channel. (F-H) Thresholding to identify individual nuclear ssDNA foci from the background signal. Abbreviations: ssDNA = single-stranded DNA; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www.jove.com/files/ftp_upload/65926/65926fig04large.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="2">Cytoskeletal (CSK) buffer</td>
		</tr>
		<tr>
			<td>PIPES pH 7.0&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>NaCl&#160;</td>
			<td>100 mM</td>
		</tr>
		<tr>
			<td>EDTA pH 8&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>MgCl2&#160;</td>
			<td>3 mM</td>
		</tr>
		<tr>
			<td>D-sucrose</td>
			<td>300 mM</td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>0.20%</td>
		</tr>
		<tr>
			<td>Phosphatase inhibitor cocktail</td>
			<td>1 tablet per 10 mL</td>
		</tr>
		<tr>
			<td>Protease inhibitor cocktail</td>
			<td>1 tablet per 10 mL</td>
		</tr>
		<tr>
			<td>diluted in ddH2O</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td colspan="2">Washing buffer</td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>0.05%</td>
		</tr>
		<tr>
			<td>diluted in PBS</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td colspan="2">Permeabilization buffer</td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>0.50%</td>
		</tr>
		<tr>
			<td>diluted in PBS</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td colspan="2">Fixation solution</td>
		</tr>
		<tr>
			<td>Paraformaldehyde&#160;</td>
			<td>3.60%</td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>0.05%</td>
		</tr>
		<tr>
			<td>diluted in PBS</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td colspan="2">Blocking buffer</td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)&#160;</td>
			<td>5%</td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>0.10%</td>
		</tr>
		<tr>
			<td>diluted in PBS</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td colspan="2">Click-iT Plus reaction cocktail</td>
		</tr>
		<tr>
			<td>1x Click-iT reaction buffer&#160;</td>
			<td>435 mL</td>
		</tr>
		<tr>
			<td>Alexa Fluor PCA solution</td>
			<td>5 mL</td>
		</tr>
		<tr>
			<td>CuSO4-copper protectant premix</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>1x Click-iT buffer additive</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>Total volume&#160;</td>
			<td>500 mL</td>
		</tr>
	</tbody>
</table>
Table 1: Composition of the buffers used in this protocol. 
Supplementary Figure S1. (A) RPE1 cells were synchronized to G0 phase using serum starvation for 72 h and subsequently released into different cell cycle phases by reintroducing serum. Dot plots show cells in G0/G1, S, or G2/M phases, where hours indicate the time after the re-addition of serum following serum starvation. Graph on the right shows the percentage of G0/G1, S, and G2/M cells in each condition. FACS analysis was carried out using a commercially available cell proliferation kit using EdU and propidium iodide according to the manufacturer&#39;s recommendations. (B) Uncropped western blot scans for Figure 1. Numbers show molecular weight markers in kDa. PARP1 was used as a loading control and loaded on the gel that was also developed against CCNA2, p27 (further stripped for PCNA), and pH3 (S10) (further stripped for H3) by cutting up the membrane. CCNB1 and RPA2 were loaded onto a separate gel, using the same amount of protein lysate to ensure comparability. (C) Uncropped western blot scans for Figure 3. Numbers show molecular weight markers in kDa. Abbreviation: EdU = 5-ethynyl-2&#39;-deoxyuridine. <a href="https://www.jove.com/files/ftp_upload/65926/Supplementary Figure 1.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Figure S2: (A) Representative images illustrate ssDNA foci using BrdU antibody (green); S phase cells are highlighted by cyclin A2 (red); and DAPI was used to counterstain nuclear DNA (blue). RPE1 cells were kept in 10 &#181;M BrdU for 48 h prior to further treatment. After 48 h, cells were treated with H2O2 (250 &#956;M) for 1 h or Neocarzinostatin (0.5 &#181;g/mL) for 4 h before fixation. A white dashed line denotes the border of each nucleus. Scale bar = 5 &#181;m. (B) BrdU staining of RPE1 cells with and without denaturing condition. Asynchronous RPE1 cells were pre-treated with 10 &#181;M BrdU for 48 h. Scale bar = 10 &#181;m. (C) The measurements for the number of BrdU foci/nucleus were carried out from two independent experiments in G1 synchronized RPE1 cells. Only EdU-negative cells were considered during the analysis. Lines represent the mean value on the plots. Non-parametric ANOVA test (Kruskal-Wallis) was performed for statistical analysis. The &#39;ns&#39; indicates non-significant difference. The analyzed number of nuclei were the following: NT n = 52, NCS n = 105, H2O2 n = 82. Abbreviations: siNT = non-targeting siRNA control; BrdU = 5-bromo-2&#39;-deoxyuridine; DAPI = 4&#39;,6-diamidino-2-phenylindole; RPE1 = hTERT-immortalized retinal pigment epithelial cells; NCS = Neocarzinostatin. <a href="https://www.jove.com/files/ftp_upload/65926/Supplementary Figure 2_revised2.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Video S1: Screen recording of Fiji-based RPA2 foci analysis. <a href="https://www.jove.com/files/ftp_upload/65926/Supplementary Video 1.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Visualizing Single-Stranded DNA During DNA Repair for Therapeutic Insights at JoVE.com

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.

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 ...

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2.1K Views.  NYU Grossman School of Medicine. 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.

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

Extracting DNA from the Gut Microbes of the Termite (Zootermopsis Angusticollis) and Visualizing Gut Microbes

Termites are among the few animals known to have the capacity to subsist solely by consuming wood.  The termite gut tract contains a dense and species-rich microbial population that assists in the degradation of lignocellulose predominantly into acetate, the key nutrient fueling termite metabolism (Odelson &amp; Breznak, 1983).  Within these microbial populations are bacteria, methanogenic archaea and, in some ("lower") termites, eukaryotic protozoa. Thus, termites are excellent research subjects for studying the interactions among microbial species and the numerous biochemical functions they perform to the benefit of their host.  The species composition of microbial populations in termite guts as well as key genes involved in various biochemical processes has been explored using molecular techniques (Kudo et al., 1998; Schmit-Wagner et al., 2003; Salmassi &amp; Leadbetter, 2003). These techniques depend on the extraction and purification of high-quality nucleic acids from the termite gut environment.  The extraction technique described in this video is a modified compilation of protocols developed for extraction and purification of nucleic acids from environmental samples (Mor  et al., 1994; Berthelet et al., 1996; Purdy et al., 1996; Salmassi &amp; Leadbetter, 2003; Ottesen et al. 2006) and it produces DNA from termite hindgut material suitable for use as template for polymerase chain reaction (PCR).

Extracting DNA from the Gut Microbes of the Termite Zootermopsis Angusticollis and Visualizing Gut Microbes

This video illustrates the technique for extracting DNA from the species of microbes resident in the termite hindgut. The preparation of a wet mount slide, which is useful for visualizing the gut microbial community is also illustrated, and a tour through the species-rich gut environment is given.

Termites are among the few animals known to have the capacity to subsist solely by consuming wood.  The termite gut tract contains a dense and ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Quantitation of &gamma;H2AX Foci in Tissue Samples

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) processes or by exogenous factors, particularly ionizing radiation and radiomimetic compounds. Phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX, is an early response to DNA double-strand breaks1. This phosphorylation event is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Overall, DSB induction results in the formation of discrete nuclear &gamma;H2AX foci which can be easily detected and quantitated by immunofluorescence microscopy2. Given the unique specificity and sensitivity of this marker, analysis of &gamma;H2AX foci has led to a wide range of applications in biomedical research, particularly in radiation biology and nuclear medicine. The quantitation of &gamma;H2AX foci has been most widely investigated in cell culture systems in the context of ionizing radiation-induced DSBs. Apart from cellular radiosensitivity, immunofluorescence based assays have also been used to evaluate the efficacy of radiation-modifying compounds. In addition, &gamma;H2AX has been used as a molecular marker to examine the efficacy of various DSB-inducing compounds and is recently being heralded as important marker of ageing and disease, particularly cancer3. Further, immunofluorescence-based methods have been adapted to suit detection and quantitation of &gamma;H2AX foci ex vivo and in vivo4,5. Here, we demonstrate a typical immunofluorescence method for detection and quantitation of &gamma;H2AX foci in mouse tissues.

Quantitation of &#947;H2AX Foci in Tissue Samples

Quantitation of DNA double-strand breaks on the basis of &gamma;H2AX foci has become an invaluable tool, particularly in radiation biology, for the evaluation of tissue radiosensitivity and effects of radiation modifying compounds. Here we demonstrate the use of an immunofluorescence assay for quantitation of &gamma;H2AX foci in tissue samples.

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) ...

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

Agarose Gel Electrophoresis for the Separation of DNA Fragments

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from the seaweed genera Gelidium and Gracilaria, and consists of repeated agarobiose (L- and D-galactose) subunits2. During gelation, agarose polymers associate non-covalently and form a network of bundles whose pore sizes determine a gel&#39;s molecular sieving properties. The use of agarose gel electrophoresis revolutionized the separation of DNA. Prior to the adoption of agarose gels, DNA was primarily separated using sucrose density gradient centrifugation, which only provided an approximation of size. To separate DNA using agarose gel electrophoresis, the DNA is loaded into pre-cast wells in the gel and a current applied. The phosphate backbone of the DNA (and RNA) molecule is negatively charged, therefore when placed in an electric field, DNA fragments will migrate to the positively charged anode. Because DNA has a uniform mass/charge ratio, DNA molecules are separated by size within an agarose gel in a pattern such that the distance traveled is inversely proportional to the log of its molecular weight3. The leading model for DNA movement through an agarose gel is &#34;biased reptation&#34;, whereby the leading edge moves forward and pulls the rest of the molecule along4. The rate of migration of a DNA molecule through a gel is determined by the following: 1) size of DNA molecule; 2) agarose concentration; 3) DNA conformation5; 4) voltage applied, 5) presence of ethidium bromide, 6) type of agarose and 7) electrophoresis buffer. After separation, the DNA molecules can be visualized under uv light after staining with an appropriate dye. By following this protocol, students should be able to: 
 
1. Understand the mechanism by which DNA fragments are separated within a gel matrix 
2. Understand how conformation of the DNA molecule will determine its mobility through a gel matrix 
3. Identify an agarose solution of appropriate concentration for their needs 
4. Prepare an agarose gel for electrophoresis of DNA samples 
5. Set up the gel electrophoresis apparatus and power supply 
6. Select an appropriate voltage for the separation of DNA fragments 
7. Understand the mechanism by which ethidium bromide allows for the visualization of DNA bands 
8. Determine the sizes of separated DNA fragments 
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A basic protocol for the separation of DNA fragments using agarose gel electrophoresis is described.

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Flow cytometry has been widely used to obtain information about DNA content in a population of cells, to infer relative percentages in different cell cycle phases. This technique has been successfully extended to the mitotic tissues of the model organism Drosophila melanogaster for genetic studies of cell cycle regulation in vivo. When coupled with cell-type specific fluorescent protein expression and genetic manipulations, one can obtain detailed information about effects on cell number, cell size and cell cycle phasing in vivo. However this live-cell method has relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye, limiting users to flow cytometers equipped with a UV laser. We have modified this protocol to use a newer live-cell DNA dye, Vybrant DyeCycle Violet, compatible with the more common violet 405nm laser. The protocol presented here allows for efficient cell cycle analysis coupled with cell type, relative cell size and cell number information, in a variety of Drosophila tissues. This protocol extends the useful cell cycle analysis technique for live Drosophila tissues to a small benchtop analyzer, the Attune Acoustic Focusing Cytometer, which can be run and maintained on a single-lab scale.

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Flow cytometry has  been widely used to obtain information about DNA content in a population of  cells, to infer relative percentages in different cell ...

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

The cell cycle is important for growth, genome replication, and development in all cells. In bacteria, studies of the cell cycle have focused largely on unsynchronized cells making it difficult to order the temporal events required for cell cycle progression, genome replication, and division. Caulobacter crescentus provides an excellent model system for the bacterial cell cycle whereby cells can be rapidly synchronized in a G0 state by density centrifugation. Cell cycle synchronization experiments have been used to establish the molecular events governing chromosome replication and segregation, to map a genetic regulatory network controlling cell cycle progression, and to identify the establishment of polar signaling complexes required for asymmetric cell division. Here we provide a detailed protocol for the rapid synchronization of Caulobacter NA1000 cells. Synchronization can be performed in a large-scale format for gene expression profiling and western blot assays, as well as a small-scale format for microscopy or FACS assays. The rapid synchronizability and high cell yields of Caulobacter make this organism a powerful model system for studies of the bacterial cell cycle.

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

Synchronization of bacterial cells is essential for studies of the bacterial cell cycle and development. Caulobacter crescentus is synchronizable through density centrifugation allowing a rapid and powerful tool for studies of the bacterial cell cycle. Here we provide a detailed protocol for the synchronization of Caulobacter cells.

The cell cycle is important for growth, genome replication, and development in all cells.  In bacteria, studies of the cell cycle have focused largely on ...