This research was funded by the National Research, Development and Innovation Office grant GINOP-2.3.2-15-2016-00020, GINOP-2.3.2-15-2016-00036, GINOP-2.2.1-15-2017-00052, EFOP 3.6.3-VEKOP-16-2017-00009, NKFI-FK 132080, the J&#225;nos Bolyai Research Scholarship of the Hungarian Academy of Sciences BO/27/20, &#218;NKP-20-5-SZTE-265, EMBO short-term fellowship 8513, and the Tempus Foundation.

1. Immunodetection of specific proteins
<ol>
	<li>Preparation of cell culture and experimental setup
	<ol>
		<li>Maintain U2OS cells in monolayers in DMEM culture medium supplemented with 10% fetal calf serum, 2 mM glutamine, and 1% antibiotic-antimycotic solution. 
		NOTE: For endonuclease-based DNA damage induction, use charcoal-treated or steroid-free medium to avoid system leakiness.</li>
		<li>Grow cells in a humidified 5% CO2 environment at 37 &#176;C until 80% confluency, renewing medium every 2-3 days.</li>
		<li>Aspirate the medium and wash the cells with 1x PBS. Detach cells with Trypsin-EDTA solution. When the cells detach, stop the trypsin activity by adding culture medium to the cells, yielding a cell suspension.</li>
		<li>Count the cells using a cell counting chamber. Plate 2 x 104 cells/mL/well on a 24-well plate, with sterile 12 mm round coverslips in each well.</li>
		<li>Incubate cells for 24 h at 37 &#176;C in a humidified 5% CO2 atmosphere to allow attachment onto the coverslips.</li>
		<li>Treat the cells with 10 ng/mL of neocarzinostatin (NCS) by directly pipetting the damaging agent to the cultured medium. Incubate the cells with the NCS-containing medium for 15 min, then wash them with 1x PBS and add fresh, supplemented culture medium to the cells. Otherwise, use appropriate agent (i.e., 4-OHT) to induce DSBs via endonuclease-based systems without refreshing the medium22. 
		NOTE: Alternatively, use irradiation to induce DNA damage, ranging from 30 min up to 8 h of recovery time by using neutron flux between 2-20 Gy23.</li>
		<li>Incubate cells for 1-8 h at 37 &#176;C in a humidified 5% CO2 atmosphere to follow the kinetics of DNA repair.</li>
	</ol>
	</li>
	<li>Fixation of cells 
	&#8203;NOTE: 300-500 &#181;L of solutions/well should be used in the following steps (steps 1.2-1.5) to cover all cells adequately. Each incubation and washing step (except the antibody incubation) should be performed on an orbital shaker with gentle agitation.
	<ol>
		<li>Following DSB induction and incubation of cells, remove the medium from the attached cells and wash the cells once with 1x PBS.</li>
		<li>Fix cells with 4% formaldehyde-PBS solution for 20 min at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Permeabilization of cells
	<ol>
		<li>Remove the fixing solution and wash cells three times with 1x PBS for 5 min each.</li>
		<li>Remove the PBS and add 0.2% Triton X-100 dissolved in PBS. Incubate the samples for 20 min.</li>
	</ol>
	</li>
	<li>Blocking of non-specific binding sites
	<ol>
		<li>Wash the cells three times with 1x PBS.</li>
		<li>Block non-specific binding sites with 5% BSA (Bovine Serum Fraction V albumin) diluted in PBST (1x PBS supplemented with 0.1% Tween-20), and incubate the permeabilized samples for at least 20 min.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining
	<ol>
		<li>Add the proper amount of primary antibody (i.e., anti-&#947;H2AX, anti-DNA-PKcs) diluted in 1% BSA-PBST solution. Place each coverslip upside down onto a paraffin film over a 10 &#181;L droplet of the diluted anti-&#947;H2AX antibody. 
		NOTE: In case of co-immunostaining dilute appropriately both antibodies in the same 1% BSA-PBST solution.</li>
		<li>Incubate the samples in a humidity chamber for 1.5 h at 4 &#176;C. 
		NOTE: Incubation can also be performed at 4 &#176;C overnight.</li>
		<li>Place the coverslips back being side up into the 24-well plate and wash three times for 5 min with 1x PBS.</li>
		<li>Add the proper amount of secondary antibody diluted in 1% BSA-PBST. Place each coverslip upside down onto a paraffin film over a 10 &#181;L droplet of the diluted antibody.</li>
		<li>Incubate the samples in a humidity chamber at 4 &#176;C for 1 h.</li>
		<li>Place the coverslips back being side up into the 24-well plate and wash three times for 5 min with 1x PBS.</li>
		<li>Before removing the last PBS washing solution, gently take out the coverslips using a tweezer and needle and then place them upside down onto glass slides with droplets of mounting medium (supplemented with DAPI). 
		&#8203;NOTE: Avoid the formation of air bubbles. When the mounting medium dries, it is recommended to seal the edges of the coverslips with nail polish to prevent shriveling of the samples.</li>
	</ol>
	</li>
</ol>
2. Chromatin immunoprecipitation
<ol>
	<li>Cell collection, crosslinking, cell and nuclear lysis, and DNA fragmentation
	<ol>
		<li>Culture approximately 5&#160;x 106 cells/mL in a 150 mm dish for each sample.</li>
		<li>Remove the culture medium and wash the cells twice with ice-cold 1x PBS.</li>
		<li>Fix the cells with 1% formaldehyde-PBS solution, place the plates on an orbital shaker, and agitate gently for 20 min. 
		NOTE: Formaldehyde is volatile; always prepare a fresh working solution. In some cases, the formaldehyde solution contains methanol to stabilize it, but it is better to use a methanol-free solution to avoid interference with downstream reactions.</li>
		<li>Stop the fixation with 125 mM glycine and incubate on an orbital shaker with gentle agitation for 5 min at 25 &#176;C.</li>
		<li>Place the plates on ice and wash twice with ice-cold 1x PBS.</li>
		<li>Scrape the cells in ice-cold 1x PBS and transfer them into 15 mL conical tubes.</li>
		<li>Centrifuge the cells at 2,500 x g for 5 min at 4 &#176;C.</li>
		<li>Carefully aspirate the supernatant and resuspend the pellet in 2 mL of cell lysis buffer [5 mM PIPES pH 8.0, 85 mM KCl, 0.5% NP-40, 1x PIC (protease inhibitor cocktail)] and incubate on ice for 10 min.</li>
		<li>Centrifuge the cell suspension at 2,500 x g for 5 min at 4 &#176;C.</li>
		<li>Carefully discard the supernatant and resuspend the pellet in 500-1,500 &#181;L of nuclear lysis buffer (50 mM Tris-HCl pH 8.0, 10 mM EDTA pH 8.0, 0.8% SDS, 1x PIC) and incubate on ice for 30-60 min. Transfer the lysate into a polystyrene conical tube suitable for sonication. 
		NOTE: Since the nuclear lysis buffer contains SDS, it will precipitate on ice, and the solution will turn white. The solution should turn transparent following sonication.</li>
		<li>Sonicate the lysate to shear DNA to an average fragment size of 300-1,000 bp. 
		NOTE: The appropriate sonication cycles and conditions should be set according to the cell type and the sonication equipment. Fragments smaller than 200 bp are not suitable for ChIP, because the nucleosome-DNA interactions can be disrupted.</li>
	</ol>
	</li>
	<li>Reversal of crosslinking, determination of sonicated fragment sizes
	<ol>
		<li>Take out 100 &#181;L of the sonicated sample to verify the fragment size of the sonicated chromatin. The remaining chromatin should be stored at &#8722;80 &#176;C.</li>
		<li>Add 0.5 mg/mL RNase A to each 100 &#181;L of the sample and incubate them at 37 &#176;C for 20 min to activate the RNase.</li>
		<li>Incubate the samples at 65 &#176;C overnight.</li>
		<li>The next day, add 500 &#181;g/mL of Proteinase K and 0.5% SDS, and incubate the samples at 50 &#176;C for 3 h.</li>
		<li>Add 0.5 volume of phenol and 0.5 volume chloroform-isoamyl alcohol mix (24:1) to each sample.</li>
		<li>Vortex for 1 min.</li>
		<li>Centrifuge at 13,000 x g for 10 min.</li>
		<li>Transfer the upper aqueous phase to a new microcentrifuge tube.</li>
		<li>Add 1 volume chloroform-isoamyl alcohol mix (24:1) to each sample.</li>
		<li>Vortex for 1 min.</li>
		<li>Centrifuge at 13,000 x g for 10 min.</li>
		<li>Transfer the upper aqueous phase to a new microcentrifuge tube.</li>
		<li>Add 2.5 volumes of 96% ethanol and 0.1 volume of 3 M Na-Acetate pH 5.2.</li>
		<li>Incubate for at least 20 min at &#8722;80 &#176;C.</li>
		<li>Centrifuge the samples at 13,000 x g for 10 min at 4 &#176;C.</li>
		<li>Remove the ethanol and wash the pellet with 400 &#181;L of 70% ethanol.</li>
		<li>Centrifuge the samples at 13,000 x g for 10 min at 4 &#176;C.</li>
		<li>Remove the ethanol and air dry the pellet.</li>
		<li>Resuspend the pellet in 10 &#181;L of TE.</li>
		<li>Run the samples on a 0.8% agarose gel. The sonicated chromatin size should be around 500 bp. 
		NOTE: Use bromophenol blue-free loading buffer because the size of this dye is approximately 500 bp, which can disturb the proper detection of chromatin fragments. Instead, it is recommended to use loading buffer complemented with xylene-cyanole, which is approximately 3,000 bp.</li>
		<li>If the chromatin size is acceptable, dilute the frozen chromatin samples from step 2.1. in 3 volumes of dilution buffer (10 mM Tris-HCl pH 8.0, 0.5 mM EGTA pH 8.0, 1% Triton X-100, 140 mM NaCl, 1x PIC) and mix the samples via rotation for 10 min at 4 &#176;C. 
		&#8203;NOTE: This step is necessary to dilute the SDS present in the nuclear lysis buffer to avoid interference with downstream reactions, including the measurement of chromatin concentration.</li>
		<li>Measure the DNA concentration of the chromatin samples at 260/280 nm using a spectrophotometer.</li>
	</ol>
	</li>
	<li>Preparation of beads, pre-clearing, and immunoprecipitation
	<ol>
		<li>Prepare beads (sheep anti-rabbit or mouse IgG) for the pre-clearing and immunoprecipitation steps. Wash beads twice for 10 min at 4 &#176;C with RIPA buffer (50 mM Tris-HCl pH 8.0, 1 mM EDTA pH 8.0, 1% Triton X-100, 0.1% Na-DOC, 0.1% SDS, 150 mM NaCl and 1X PIC).</li>
		<li>Resuspend the beads in the same volume of RIPA buffer as in step 2.3.1.</li>
		<li>Pre-clear 25-30 &#181;g chromatin of each sample with 4 &#181;L of the beads via rotation for 1-2 h at 4 &#176;C. 
		NOTE: Add RIPA buffer to each chromatin sample up to 500 &#181;L of final volume to let the samples mix properly under rotation. Do not forget to take out chromatin for NAC (No Antibody Control) and TIC (Total Input Control) in the case of each sample set. TICs only require a final volume of up to 200 &#181;L.</li>
		<li>Precipitate the beads with a magnet and transfer the supernatant to a new microcentrifuge tube.</li>
		<li>Add the appropriate amount of antibody to each chromatin sample (except NAC and TIC) and rotate overnight at 4 &#176;C.</li>
		<li>Next day, add 40 &#181;L of washed beads to each sample (except TIC) and incubate them overnight, rotating at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Washing
	<ol>
		<li>Wash once with 300 &#181;L of Low Salt buffer (20 mM Tris-HCl pH 8.0, 150 mM NaCl, 2 mM EDTA pH 8.0, 1% Triton X-100, 0.1% SDS, 1x PIC) for 10 min via rotation at 4 &#176;C.</li>
		<li>Wash once with 300 &#181;L of High Salt buffer (20 mM Tris-HCl pH 8.0, 300 mM NaCl, 2 mM EDTA pH 8.0, 1% Triton X-100, 0.1% SDS, 1x PIC) for 10 min via rotation at 4 &#176;C.</li>
		<li>Wash once with 300 &#181;L of LiCl buffer (250 mM LiCl, 1% NP-40, 1% Na-DOC, 1 mM EDTA pH 8.0, 10 mM Tris-HCl pH 8.0, 1x PIC) for 10 min via rotation at 4 &#176;C.</li>
		<li>Wash twice with 300 &#181;L of TE (10 mM Tris-HCl pH 8.0, 1 mM EDTA pH 8.0) for 10 min via rotation, for the first wash at 4 &#176;C, and the second wash at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Elution
	<ol>
		<li>Add 200 &#181;L of the elution buffer (1% SDS and 100 mM NaHCO3) to the beads and incubate at 65 &#176;C in a thermo-shaker for 15 min with continuous shaking (approx. 400 RPM). Transfer the supernatant to a new tube and elute beads again in 200 &#181;L of Elution buffer. Combine eluates (400 &#181;L final volume).</li>
		<li>Add NaCl to a final concentration of 200 mM in each sample. Supplement the TIC samples with 200 &#181;L of Elution buffer and add NaCl as well. 
		&#8203;NOTE: From this step, TIC should be handled under the same conditions as the other samples.</li>
		<li>Incubate the samples at 65 &#176;C (without shaking) for at least 6 h.</li>
		<li>Add 1 mL of cold 100% ethanol to each sample, rotate the tubes twice to mix, and precipitate DNA overnight at &#8722;80 &#176;C.</li>
		<li>The next day, centrifuge for 30 min at 13,000 x g at 4 &#176;C.</li>
		<li>Discard the supernatant and wash the pellet with 70% EtOH.</li>
		<li>Centrifuge the samples at 13,000 x g for 10 min at 4 &#176;C.</li>
		<li>Discard the supernatant and air dry the pellets.</li>
		<li>Resuspend the pellets in 100 &#181;L of TE and add 0.5 mg/mL of RNase A to each sample. Incubate at 37 &#176;C for 20 min to activate the RNase.</li>
	</ol>
	</li>
	<li>Reversal of crosslinking
	<ol>
		<li>Add 500 &#181;g/mL of Proteinase K and 0.5% SDS then incubate the samples at 50 &#176;C for 2 h. 
		NOTE: If proceeding to ChIP-seq, avoid phenol-chloroform extraction, as it inhibits the downstream NGS process. Instead, it is recommended to use a commercially available kit (see Table of Materials).</li>
		<li>Add 0.5 volume of phenol and 0.5 volume of chloroform-isoamyl alcohol mix (24:1) to each sample.</li>
		<li>Vortex for 1 min.</li>
		<li>Centrifuge at 13,000 x g for 10 min.</li>
		<li>Transfer the upper aqueous phase to a new microcentrifuge tube.</li>
		<li>Add 1 volume of chloroform-isoamyl alcohol mix (24:1) to each sample.</li>
		<li>Vortex for 1 min.</li>
		<li>Centrifuge at 13,000 x g for 10 min.</li>
		<li>Transfer the upper aqueous phase to a new microcentrifuge tube.</li>
	</ol>
	</li>
	<li>DNA extraction
	<ol>
		<li>Add 2.5 volumes of 96% ethanol and 0.1 volume of 3 M Na-Acetate pH 5.2.</li>
		<li>Incubate for at least 20 min at &#8722;80 &#176;C.</li>
		<li>Centrifuge the samples at 13,000 x g for 10 min at 4 &#176;C.</li>
		<li>Remove the ethanol and wash the pellet with 400 &#181;L of 70% ethanol.</li>
		<li>Centrifuge the samples at 13,000 x g for 10 min at 4 &#176;C.</li>
		<li>Remove the ethanol and air dry the pellet.</li>
		<li>Resuspend the pellet in 50 &#181;L of TE.</li>
	</ol>
	</li>
</ol>

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A., Kruhlak, M., Dotiwala, F., Nussenzweig, A., Haber, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterochromatin+is+refractory+to+gamma-H2AX+modification+in+yeast+and+mammals.">Heterochromatin is refractory to gamma-H2AX modification in yeast and mammals.</a> Journal of Cell Biology. 178 (2), 209-218 (2007).</li><li>Daniels, D. L., Urh, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+intracellular+protein--DNA+complexes+using+HaloCHIP,+an+antibody-free+alternative+to+chromatin+immunoprecipitation.">Isolation of intracellular protein--DNA complexes using HaloCHIP, an antibody-free alternative to chromatin immunoprecipitation.</a> Methods in Molecular Biology. 977, 111-124 (2013).</li></ol>

Although DNA repair is a relatively recent research field, our knowledge is rapidly expanding with the help of various biochemical and microscopic methods. Preserving genetic information is crucial for cells since mutations occurring in genes involved in repair processes are among the leading causes of tumorigenesis and therefore elucidating the key steps of DNA repair pathways is essential.
Biochemical techniques (i.e., western blot, immunoprecipitation, mass-spectrometry, etc.) require large number of cells and the studied repair processes represent a snapshot of the desired cell population. Performing ChIP experiments is laboring, troublesome and many considerations have to be taken into account when designing specific experiment to study the process of DSB repair. The following steps are some examples: (I) cells should be properly lysed; it is highly recommended to apply a two-step lysation method by using separate cell- and nuclear lysis buffer to ensure higher accessibility to the chromatin fraction (II) chromatin should not be over- or undersonicated; the appropriate conditions of sonication should be optimized to each cell type beforehand (III) the appropriate amount of purified antibody should be determined since antibodies against the same protein from different companies exhibit non-identical characteristics (IV) for the efficient immunoprecipitation, 25-30 ug initial chromatin should be used in each condition (V) the appropriate time of fixation should be optimized, as overfixation can result in false positive results by cross-linking the distant protein complexes and underfixation can prevent the proper cross-linking between DNA and the desired protein (VI) based on the applied antibody, the type of the beads (protein A or G) should be carefully determined (VII) during washing steps, the order of the washing buffers should be thoroughly kept to avoid the release of the antibody from the beads (VIII) during DNA extraction, phenol traces should be properly eliminated to avoid decreasing the efficiency of the further downstream reactions. Since this method reduces the yield of recovered DNA, our personal advice is to rather use specific DNA-purification kit. When all critical points have been properly addressed, ChIP can provide valuable data for the occupancy of the desired protein at different genomic loci and can unravel critical steps in DNA repair processes.
However, ChIP combined with qPCR is an indirect approach to study the protein distribution at selected genomic regions and does not allow to specifically recognize the DNA binding site or directly examine the function of the protein. Mono- or polyclonal antibodies used to capture the protein-DNA complexes can also cross-react with other proteins leading to false-positive data and therefore, the antibodies used in this technique should be ChIP-grade and highly specific against the protein of interest. However, ChIP is a widely used technique and further approaches based on this have been developed, such as ChIP-on-chip and ChIP-seq. The former one relies on hybridizing the immunoprecipitated and purified DNA fragments on a microarray with a large variety of small random DNA sequences which further amplifies the annealed sequences providing valuable information about protein binding sites. However, ChIP-seq has emerged as an attractive alternative approach since it provides genome-wide mapping of protein-DNA complexes with higher resolution than ChIP-on-chip and high-throughput genome sequencing. ChIP-seq has revolutionized the field of DNA repair by disclosing DNA binding sites of various transcription factors providing insights into the gene regulation and unravelling chromatin landscapes in a genome-wide scale25. According to this, the field of DNA repair has tremendously profited from ChIP-seq since these data play a crucial role in various diseases and biological pathways, such as cancer progression. Nonetheless, various modifications of the ChIP method have been developed such as HaloChIP, which does not require specific antibody against the protein of interest but rather uses sequences encoding DNA-binding proteins fused with HaloTag, which are transfected to cells and subsequently to crosslinking, the desired protein-DNA complexes can be captured by using HaloLink Resin. However, this technique relies on overexpression and not the endogenous level of the desired proteins which can result in misinterpreted data26.
Furthermore, microscopic techniques provide valuable information about the spatiotemporal tracking of DNA damage repair, even in a single-cell level. The rapid improvement of antibodies raised against specific repair proteins has led to a deeper understanding of the mechanism of NER and DSBR sub-pathways, as well as their post-translational regulation. The microscopic field has been revolutionized by high-resolution techniques, such as super-resolution microscopy, which allows the visualization of DNA damage-induced cellular processes at the nucleosomal level, as well as ensuring the accurate mapping of protein co-localization24. However, it should be noted that the variance in cell lineages must be considered during the experiment as the rate of the repair can diverge, which can make the results difficult to interpret. Considering the rapid evolution of fluorescence imaging methodology and the deliberate design of the experimental setup, a precious opportunity to investigate the DNA damage-induced cellular and molecular responses at a single protein resolution in a single-cell level is on its way to perfection.
In conclusion, the combination of super-resolution microscopy and single-cell sequencing methodology can significantly improve our understanding of the DNA repair field.

Our genome is constantly being challenged by various DNA damaging agents. These assaults can derive from environmental sources, such as UV light or irradiation, as well as from endogenous sources, such as metabolic by-products caused by oxidative stress or replication errors1,2. These lesions can affect the integrity of either one or both DNA strands, and if the generated errors become persistent, it frequently leads to translocations and genome instability, which may result in tumorigenesis3,4. To maintain genome integrity, multiple repair systems have been developed during evolution. According to the chemical and physical properties of specific types of DNA damage, multiple repair mechanisms can be activated. Mismatches, abasic sites, single-strand breaks, and 8-oxoguanine (8-oxoG) can be removed either by mismatch repair or base-excision repair pathway5,6. Lesions caused by UV-induced photoproducts and bulky adducts can be repaired either by nucleotide-excision repair (NER) or DNA double-strand break repair (DSBR) process7,8. NER consists of two main sub-pathways: transcription-coupled NER (TC-NER) and global genomic NER (GG-NER). Regarding the cell cycle phase, following DNA double-strand break induction, two sub-pathways can be activated: non-homologous end joining (NHEJ) and homologous recombination (HR)1,9. NHEJ, which is the dominant pathway in resting cells, can be activated in all cell cycle phases, representing a faster but error-prone pathway10. On the other hand, HR is an error-free pathway, in which the DSBs are repaired based on sequence-homology search of the sister chromatids, therefore it is mainly present in S and G2 cell cycle phases11. Furthermore, microhomology-mediated end joining (MMEJ) is another DSB repair mechanism, distinct from the aforementioned ones, based on a KU70/80- and RAD51-independent way of re-ligation of previously resected microhomologous sequences flanking the broken DNA ends. Therefore, MMEJ is considered to be error-prone and highly mutagenic12. During DNA repair, DSBs can induce the DNA damage response (DDR), which results in the activation of checkpoint kinases that halt the cell cycle during repair13,14,15. The DDR is activated as a response to the recruitment and extensive spreading of initiator key players of the repair process around the lesions, contributing to the formation of a repair focus. In this early signaling cascade, the ATM (Ataxia Telangiectasia Mutated) kinase plays a pivotal role by catalyzing the phosphorylation of the histone variant H2AX at Ser139 (referred to as &#947;H2AX) around the lesion16. This early event is responsible for the recruitment of additional repair factors and the initiation of downstream repair processes. Although the exact function of the recruited proteins at the repair focus has not yet been fully characterized, the formation and the dynamics of repair foci have been investigated by several laboratories. These markers are extensively used to follow the repair kinetics, but their precise role during the repair process remains elusive. Due to the great importance yet poor understanding of DNA repair-related cellular processes, several methods have been developed so far to induce and visualize the DDR.
Various methods and systems have been established to induce the desired type of DNA damage. For instance, some agents [such as neocarzinostatin (NCS), phleomycin, bleomycin, &#947;-irradiation, UV] can induce large numbers of random DNA breaks at non-predictive genomic positions, while others (endonucleases, such as AsiSI, I-PpoI or I-SceI, as well as laser striping) can induce DNA breaks at known genomic loci17,18,19,20,21. Here, we focus on the endonuclease-based techniques currently used to study the DDR in mammalian and yeast cells. Aside from highlighting the principles of these techniques, we emphasize both their advantages and disadvantages.

<table><tbody><tr><td>4-OHT</td><td>Sigma Aldrich</td><td>H7904</td><td></td></tr><tr><td>Agarose</td><td>Lonza</td><td>50004</td><td></td></tr><tr><td>Antibiotic-Antimycotic Solution (100&amp;times;), Stabilized</td><td>Sigma Aldrich</td><td>A5955</td><td></td></tr><tr><td>Anti-gamma H2A.X (phospho S139) antibody</td><td>Abcam</td><td>ab26350</td><td></td></tr><tr><td>Bovine Serum Fraction V albumin</td><td>Biosera</td><td>PM-T1725</td><td></td></tr><tr><td>TrackIt&amp;trade; Cyan/Yellow Loading Buffer</td><td>Thermo Fisher Scientific</td><td>10482035</td><td></td></tr><tr><td>DMEM with 1.0 g/L Glucose, without L-Glutamine</td><td>Lonza</td><td>12-707F</td><td></td></tr><tr><td>Doxycycline</td><td>Sigma Aldrich</td><td>D9891</td><td></td></tr><tr><td>Dynabeads&amp;trade; M-280 Sheep Anti-Mouse IgG</td><td>Invitrogen</td><td>11202D</td><td></td></tr><tr><td>Dynabeads&amp;trade; M-280 Sheep Anti-Rabbit IgG</td><td>Invitrogen</td><td>11204D</td><td></td></tr><tr><td>EDTA</td><td>Sigma Aldrich</td><td>E6758</td><td></td></tr><tr><td>EGTA</td><td>Sigma Aldrich</td><td>E3889</td><td></td></tr><tr><td>Ethanol</td><td>Molar Chemicals</td><td>02910-101-340</td><td></td></tr><tr><td>Fetal Bovine Serum (South America Origin), EU-approved</td><td>Gibco</td><td>ECS0180L</td><td></td></tr><tr><td>Formaldehyde 37% solution free from acid</td><td>Sigma Aldrich</td><td>1.03999</td><td></td></tr><tr><td>GlutaMAX&amp;trade; Supplement</td><td>Thermo Fisher Scientific</td><td>35050038</td><td></td></tr><tr><td>Glycine</td><td>Sigma Aldrich</td><td>50046</td><td></td></tr><tr><td>IPure kit v2</td><td>Diagenode</td><td>C03010015</td><td></td></tr><tr><td>Isoamyl alcohol</td><td>Sigma Aldrich</td><td>W205702</td><td></td></tr><tr><td>LiCl</td><td>Sigma Aldrich</td><td>L9650</td><td></td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S5886</td><td></td></tr><tr><td>Na-DOC</td><td>Sigma Aldrich</td><td>D6750</td><td></td></tr><tr><td>NaHCO3</td><td>Sigma Aldrich</td><td>S5761</td><td></td></tr><tr><td>Neocarzinostatin from Streptomyces carzinostaticus</td><td>Sigma Aldrich</td><td>N9162</td><td></td></tr><tr><td>NP-40</td><td>Sigma Aldrich</td><td>I8896</td><td></td></tr><tr><td>PBS Powder without Ca2+, Mg2+</td><td>Sigma Aldrich</td><td>L182-50-BC</td><td></td></tr><tr><td>Phenol</td><td>Sigma Aldrich</td><td>P4557</td><td></td></tr><tr><td>PIPES</td><td>Sigma Aldrich</td><td>P1851</td><td></td></tr><tr><td>Polysorbate 20 (Tween 20)</td><td>Molar Chemicals</td><td>09400-203-190</td><td></td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P5405</td><td></td></tr><tr><td>ProLong&amp;trade; Gold Antifade Mountant with DAPI</td><td>Thermo Fisher Scientific</td><td>P36935</td><td></td></tr><tr><td>Protease Inhibitor Cocktail Set I</td><td>Roche</td><td>11873580001</td><td></td></tr><tr><td>Proteinase K</td><td>Sigma Aldrich</td><td>P2308</td><td></td></tr><tr><td>P-S2056 DNAPKcs antibody</td><td>Abcam</td><td>ab18192</td><td></td></tr><tr><td>RNase A</td><td>Roche</td><td>10109169001</td><td></td></tr><tr><td>CH&lt;sub&gt;3&lt;/sub&gt;COONa</td><td>Sigma Aldrich</td><td>S2889</td><td></td></tr><tr><td>SDS</td><td>Sigma Aldrich</td><td>L3771</td><td></td></tr><tr><td>Tris Acetate-EDTA buffer</td><td>Sigma Aldrich</td><td>T6025</td><td></td></tr><tr><td>Tris-HCl</td><td>Sigma Aldrich</td><td>91228</td><td></td></tr><tr><td>TRITON X-100</td><td>Molar Chemicals</td><td>09370-006-340</td><td></td></tr><tr><td>Trypsin from porcine pancreas</td><td>Sigma Aldrich</td><td>T4799</td><td></td></tr><tr><td>Trypsin-EDTA (0.5%), no phenol red</td><td>Gibco</td><td>15400054</td><td></td></tr></tbody></table>

visualizing and quantifying endonuclease-based site-specific dna damage

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

Studying site-directed DSB-induced repair processes in cells can be achieved&#160;via either stable or transient transfection. However, it should be noted that stable transfection ensures a homogenous cell population, which gives a unified and thus more reliable cellular response. In the case of transient transfection, only a small proportion of the cell population takes up and maintains the plasmid, which introduces diversity into the experiment. Establishing ER-I-PpoI or ER-AsiSI endonuclease-based cell systems require a 50% confluent cell population, which is more effectively transfected with plasmids encoding the endonuclease. For transfection, commercially available transfection reagents or viral infection-based methods can also be used. If a microscopic visualization technique is to be applied and transient transfection is required, the directed DSBs can be induced by 4-OHT addition 24 h after transfection, which binds to the ER-fused endonucleases and allows the nuclear translocation and DSB induction. To determine the most appropriate time-points, immunofluorescence-based microscopy and western blot detection of &#947;H2AX at different time-points following 4-OHT treatment can be performed. Under physiological conditions, a maximum of 10-15 &#947;H2AX foci per cell can be detected, and the formation of strong repair foci can be triggered by endonucleases (or various other techniques not discussed here e.g., laser microirradiation). A typical I-PpoI endonuclease leads to the formation of elevated &#947;H2AX signals around the nucleolus by inducing DSBs at ribosomal DNA (rDNA). If the breaks are repaired by NHEJ or HR, the number of repair foci decreases over time. For this reason, representative time-points at 0 h, 30 min, 1, 2, 4, and 8 h following 4-OHT treatments are recommended. To track DNA repair processes, the most commonly used cell line is U2OS, as all known repair pathways are fully functional in these cells. When investigating several proteins in the same cells, co-localization can be studied by combining antibodies conjugated with different fluorophores with different emission wavelengths raised in different animal species as shown in Figure 1. Therein, the induction of DSBs via an inducible stable cell line is represented which is based on the ER-AsiSI restriction endonuclease fused with hemagglutinin tag (HA). Doxycycline can induce the expression and sequestration of HA-ER-AsiSI in the cytoplasm which can be tracked using an antibody against HA (Figure 1. third column, second raw). Incubation for 4 h with 4-OHT, 24 h after doxycycline addition, can induce high number of DSBs since the endonuclease has been translocated to the nucleus (Figure 1. third column, third raw and second column, third raw). DSBs can be visualized by using an antibody recognizing &#947;H2AX.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62175/62175fig01.jpg" /> 
Figure 1:&#160;Immunofluorescence microscopy is used to detect &#947;H2AX in cultured cells expressing HA-ER-AsiSI. DOX (doxycycline) addition activates the cytoplasmic expression of HA-ER-AsiSI and 4-OHT (4-hydroxytamoxifen) (4 h) induces the nuclear translocation of the fusion protein, which leads to the induction of DSBs at known genomic positions. HA (hemagglutinin) staining (anti-HA antibody) represents the HA-ER-AsiSI fusion protein in green, and the induction of breaks is verified by &#947;H2AX staining (anti-&#947;H2AX antibody) in red. Scale bars represent 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62175/62175fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Co-localization of repair proteins at the damage site indicates that they are recruited to the same DNA lesion site, but they do not necessarily interact with each other. The resolution of the confocal microscopy is approximately 300 nm; to determine the binding pattern of specific repair proteins at the break site, super-resolution microscopy (STORM) is instead recommended 24. However, this method requires expensive microscopic equipment and an expert researcher. Alternatively, the binding pattern of the repair proteins can be examined by chromatin immunoprecipitation using DIvA or U2OS-pEP15 stable cell lines, which can express AsiSI and I-PpoI endonucleases in a regulated fashion, respectively17,21. Upon 4-OHT addition, both endonucleases can cut the DNA in a sequence specific manner which provides us the opportunity to design locus specific primers to the expected break sites and their surrounding genomic regions. By applying &#947;H2AX antibody in the immunoprecipitation part of the ChIP, we can temporally follow the DNA repair kinetics upon different conditions (such as silencing or inhibition of certain repair factors of interest, i.e., DNA-PKcs). A typical experimental result obtained using ChIP-qPCR is represented in Figure 2. Therein, the temporal enrichment of &#947;H2AX is demonstrated as a response to I-PpoI-induced DNA damage. On the left part of the image, the timely detected &#947;H2AX signal is shown at the break site while on the right part, the &#947;H2AX distribution is represented at a control gene region at which DSBs have not been induced.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62175/62175fig02.jpg" /> 
Figure 2: Temporal enrichment of &#947;H2AX as determined by chromatin immunoprecipitation in response to I-PpoI-induced DNA damage. Left, &#947;H2AX signal at the break site; right, &#947;H2AX distribution at a control region where DSBs were not induced (anti-&#947;H2AX antibody). The represented results are derived from one biological experiment, and error bars indicate the variations of the corresponding sample replicates. N=3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62175/62175fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage at JoVE.com

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

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

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

visualizing-quantifying-endonuclease-based-site-specific-dna

Nanyang Technological University

Research

JoVE Journal

Biology

3.5K Views.  University of Szeged. This article introduces essential steps of immunostaining and chromatin immunoprecipitation. These protocols are commonly used to study DNA damage-related cellular processes and to visualize and quantify the recruitment of proteins implicated in DNA repair. 

Video: Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

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

Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage

DNA damage induces specific signaling and repair responses in the cell, which is critical for protection of genome integrity. Laser microirradiation became a valuable experimental tool to investigate the DNA damage response (DDR) in vivo. It allows real-time high-resolution single-cell analysis of macromolecular dynamics in response to laser-induced damage confined to a submicrometer region in the cell nucleus. However, various laser conditions have been used without appreciation of differences in the types of damage induced. As a result, the nature of the damage is often not well characterized or controlled, causing apparent inconsistencies in the recruitment or modification profiles. We demonstrated that different irradiation conditions (i.e., different wavelengths as well as different input powers (irradiances) of a femtosecond (fs) near-infrared (NIR) laser) induced distinct DDR and repair protein assemblies. This reflects the type of DNA damage produced. This protocol describes how titration of laser input power allows induction of different amounts and complexities of DNA damage, which can easily be monitored by detection of base and crosslinking damages, differential poly (ADP-ribose) (PAR) signaling, and pathway-specific repair factor assemblies at damage sites. Once the damage conditions are determined, it is possible to investigate the effects of different damage complexity and differential damage signaling as well as depletion of upstream factor(s) on any factor of interest.

Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage

The goal of this protocol is to describe how to use laser microirradiation to induce different types of DNA damage, including relatively simple strand breaks and complex damage, to study DNA damage signaling and repair factor assembly at damage sites in vivo.

DNA damage induces specific signaling and repair responses in the cell, which is critical for protection of genome integrity. Laser microirradiation ...

Genetics

Investigation of Protein Recruitment to DNA Lesions Using 405 Nm Laser Micro-irradiation

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand genome maintenance mechanisms. Since recruitment and exchange of proteins at lesions are highly dynamic, their study requires the ability to generate DNA damage in a rapid and spatially-delimited manner. Here, we describe procedures to locally induce DNA damage in human cells using a commonly available laser-scanning confocal microscope equipped with a 405 nm laser line. Accumulation of genome maintenance factors at laser stripes can be assessed by immunofluorescence (IF) or in real-time using proteins tagged with fluorescent reporters. Using phosphorylated histone H2A.X (&#947;-H2A.X) and Replication Protein A (RPA) as markers, the method provides sufficient resolution to discriminate locally-recruited factors from those that spread on adjacent chromatin. We further provide ImageJ-based scripts to efficiently monitor the kinetics of protein relocalization at DNA damage sites. These refinements greatly simplify the study of the DDR dynamics.

Studying DNA damage repair kinetics requires a system to induce lesions at defined sub-nuclear regions. We describe a method to create localized double-stranded breaks using a laser-scanning confocal microscope equipped with a 405 nm laser and provide automated procedures to quantify the dynamics of repair factors at these lesions.

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand ...

A Simple, Rapid, and Quantitative Assay to Measure Repair of DNA-protein Crosslinks on Plasmids Transfected into Mammalian Cells

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in mammalian cell lines. Rather than globally assaying removal of xenobiotic-induced or spontaneous chromosomal DPC removal, this assay examines the repair of a homogeneous, chemically defined lesion specifically introduced at one site within a plasmid DNA substrate. Importantly, this approach avoids the use of radioactive materials and is not dependent on expensive or highly-specialized technology. Instead, it relies on standard recombinant DNA procedures and widely available real-time, quantitative polymerase chain reaction (qPCR) instrumentation. Given the inherent flexibility of the strategy utilized, the size of the crosslinked protein, as well as the nature of the chemical linkage and the precise DNA sequence context of the attachment site can be varied to address the respective contributions of these parameters to the overall efficiency of DPC repair. Using this method, plasmids containing a site-specific DPC were transfected into cells and low molecular weight DNA recovered at various times post-transfection. Recovered DNA is then subjected to strand-specific primer extension (SSPE) using a primer complementary to the damaged strand of the plasmid. Since the DPC lesion blocks Taq DNA polymerase, the ratio of repaired to un-repaired DNA can be quantitatively assessed using qPCR. Cycle threshold (CT) values are used to calculate percent repair at various time points in the respective cell lines. This SSPE-qPCR method can also be used to quantitatively assess the repair kinetics of any DNA adduct that blocks Taq polymerase.

The goal of this protocol is to quantify the repair of defined DNA-protein crosslinks on plasmid DNA. Lesioned plasmids are transfected into recipient mammalian cell lines and low-molecular weight harvested at multiple time points post-transfection. DNA repair kinetics are quantified using strand-specific primer extension followed by qPCR.

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in ...

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 ...

Using Next Generation Sequencing to Identify Mutations Associated with Repair of a CAS9-induced Double Strand Break Near the CD4 Promoter

Double strand breaks (DSBs) in DNA are the most cytotoxic type of DNA damage. Because a myriad of insults can result in these lesions (e.g., replication stress, ionizing radiation, unrepaired UV damage), DSBs occur in most cells each day. In addition to cell death, unrepaired DSBs reduce genome integrity and the resulting mutations can drive tumorigenesis. These risks and the prevalence of DSBs motivate investigations into the mechanisms by which cells repair these lesions. Next generation sequencing can be paired with the induction of DSBs by ionizing radiation to provide a powerful tool to precisely define the mutations associated with DSB repair defects. However, this approach requires computationally challenging and cost prohibitive whole genome sequencing to detect the repair of the randomly occurring DSBs associated with ionizing radiation. Rare cutting endonucleases, such as I-Sce1, provide the ability to generate a single DSB, but their recognition sites must be inserted into the genome of interest. As a result, the site of repair is inherently artificial. Recent advances allow guide RNA (sgRNA) to direct a Cas9 endonuclease to any genome locus of interest. This could be applied to the study of DSB repair making next generation sequencing more cost effective by allowing it to be focused on the DNA flanking the Cas9-induced DSB. The goal of the manuscript is to demonstrate the feasibility of this approach by presenting a protocol that can define mutations that stem from the repair of a DSB upstream of the CD4 gene. The protocol can be adapted to determine changes in the mutagenic potential of DSB associated with exogenous factors, such as repair inhibitors, viral protein expression, mutations, and environmental exposures with relatively limited computation requirements. Once an organism's genome has been sequenced, this method can be theoretically employed at any genomic locus and in any cell culture model of that organism that can be transfected. Similar adaptations of the approach could allow comparisons of repair fidelity between different loci in the same genetic background.

Presented here is sgRNA/CAS9 endonuclease and next-generation sequencing protocol that can be used to identify the mutations associated with double strand break repair near the CD4 promoter.

Double strand breaks (DSBs) in DNA are the most cytotoxic type of DNA damage. Because a myriad of insults can result in these lesions (e.g., replication ...

Quantifying DNA Double Strand Break Repair with Nanoluciferase Reporter Assays

Assessment of DNA Double Strand Break Repair Activity Using High-throughput and Quantitative Luminescence-Based Reporter Assays

The repair of DNA double strand breaks (DSBs) is crucial for the maintenance of genome stability and cell viability. DSB repair (DSBR) in cells is mediated through several mechanisms: homologous recombination (HR), non-homologous end joining (NHEJ), microhomology-mediated end joining (MMEJ), and single strand annealing (SSA). Cellular assays are essential to measure the proficiency and modulation of these pathways in response to various stimuli.
Here, we present a suite of extrachromosomal reporter assays that each measure the reconstitution of a nanoluciferase reporter gene by one of the four major DSBR pathways in cells. Upon transient transfection into cells of interest, repair of pathway-specific reporter substrates can be measured in under 24 h by the detection of Nanoluciferase (NanoLuc) luminescence.
These robust assays are quantitative, sensitive, titratable, and amenable to a high-throughput screening format. These properties provide broad applications in DNA repair research and drug discovery, complementing the currently available toolkit of cellular DSBR assays.

Author Spotlight: Developing Novel Anticancer Therapeutics Targeting the DNA Damage Response

We present protocols for the assessment of double strand break repair pathway proficiency in cells using a suite of luminescence-based extrachromosomal reporter substrates.

The repair of DNA double strand breaks (DSBs) is crucial for the maintenance of genome stability and cell viability. DSB repair (DSBR) in cells is ...

Quantitative, Real-time Analysis of Base Excision Repair Activity in Cell Lysates Utilizing Lesion-specific Molecular Beacons

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates using base excision repair (BER) molecular beacons. The substrate (beacon) is comprised of a deoxyoligonucleotide containing a single base lesion with a 6-Carboxyfluorescein (6-FAM) moiety conjugated to the 5'end and a Dabcyl moiety conjugated to the 3' end of the oligonucleotide. The BER molecular beacon is 43 bases in length and the sequence is designed to promote the formation of a stem-loop structure with 13 nucleotides in the loop and 15 base pairs in the stem1,2. When folded in this configuration the 6-FAM moiety is quenched by Dabcyl in a non-fluorescent manner via F&ouml;rster Resonance Energy Transfer (FRET)3,4. The lesion is positioned such that following base lesion removal and strand scission the remaining 5 base oligonucleotide containing the 6-FAM moiety is released from the stem. Release and detachment from the quencher (Dabcyl) results in an increase of fluorescence that is proportionate to the level of DNA repair. By collecting multiple reads of the fluorescence values, real-time assessment of BER activity is possible. The use of standard quantitative real-time PCR instruments allows the simultaneous analysis of numerous samples. The design of these BER molecular beacons, with a single base lesion, is amenable to kinetic analyses, BER quantification and inhibitor validation and is adaptable for quantification of DNA Repair activity in tissue and tumor cell lysates or with purified proteins. The analysis of BER activity in tumor lysates or tissue aspirates using these molecular beacons may be applicable to functional biomarker measurements. Further, the analysis of BER activity with purified proteins using this quantitative assay provides a rapid, high-throughput method for the discovery and validation of BER inhibitors.

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates. The assay yields rates of DNA Repair activity amenable to kinetic analysis and is adaptable for quantification of DNA Repair activity in tissue and tumor lysates or with purified proteins.

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates using base ...

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

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

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

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

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

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

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

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

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

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

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

Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage

Investigation of Protein Recruitment to DNA Lesions Using 405 Nm Laser Micro-irradiation

A Simple, Rapid, and Quantitative Assay to Measure Repair of DNA-protein Crosslinks on Plasmids Transfected into Mammalian Cells

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

Using Next Generation Sequencing to Identify Mutations Associated with Repair of a CAS9-induced Double Strand Break Near the CD4 Promoter

Assessment of DNA Double Strand Break Repair Activity Using High-throughput and Quantitative Luminescence-Based Reporter Assays

Quantitative, Real-time Analysis of Base Excision Repair Activity in Cell Lysates Utilizing Lesion-specific Molecular Beacons

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

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