The authors acknowledge financial support from the Blazer foundation, the Medical Biotechnology Program at Biomedical Sciences, UIC Rockford, and the Department of Health Science Education, UIC Rockford. The authors thank Ananya Sangineni and James Bradley for their contributions to the project.

1. Preparation of antibodies and buffer
<ol>
	<li>Prepare the primary antibody solution, mouse anti-BrdU at a 1:300 dilution in 5% BSA and rat anti-BrdU at a 1:500 dilution in 5% BSA.</li>
	<li>Prepare the secondary antibody solution, alexafluor 594 rabbit anti-mouse at a 1:300 dilution in 5% BSA and alexafluor 488 chicken anti-rat at a 1:500 dilution in 5% BSA.</li>
	<li>Prepare the tertiary antibody solution, alexafluor 594 goat anti-rabbit at a 1:1,000 dilution in 5% BSA and alexafluor 488 goat anti-chicken at a 1:1,000 dilution in 5% BSA.</li>
	<li>Prepare 10 mL of lysis buffer in ddH2O with 2 mL of 1M Tris (pH 7.4), 1 mL of 0.5 M EDTA, and 0.5 mL of 10% SDS.</li>
</ol>
2. Preparation for the fiber assay
<ol>
	<li>Day 1: Cell culture and nanomaterial treatment for the fiber assay
	<ol>
		<li>Plate RAW 264.5 macrophage cells or the cells of interest, so that the cells are in the log phase of their growth on the day of the experiment. For RAW cells, add 5 x 104 cells/well to 24-well plates for each condition. Maintain the cells in a 37 &#176;C incubator with 5% CO2 and 98% humidity. Table 1 describes the constituents of the media used. Allow the cells to grow to 75%-80% confluency17,37.</li>
		<li>After the cells reach the level of 75%-80%confluency, remove the medium from the plates, take half of the medium and reserve it for the CldU pulse (CldU reserve), and add 5 &#181;L IdU to the other half at a final concentration of 50 &#181;M. Mix and add the IdU-containing medium back to the plates. Place the cells with IdU in the incubator for 20 min.</li>
		<li>Place the CldU reserve medium at 37 &#176;C to keep it pre-warmed. Add CldU to this medium just before the CldU pulse.</li>
		<li>After 20 min, aspirate the IdU-medium mixture, and gently wash the cells with 500 &#181;L of phosphate-buffered saline (PBS, pH 7.4) by gently rotating the plate. Discard the PBS.</li>
		<li>Treat the cells with various concentrations of nanoparticles in 500 &#181;L of the medium, and incubate for another 30 min. For this study, use treated graphene oxide nanoparticles at a 25 &#181;g/mL concentration.</li>
		<li>Aspirate the treatment-medium mixture, and gently wash the cells with 500 &#181;L of PBS by gently rotating the plate. Discard the PBS.Add 5 &#181;L of CldU (100 &#181;M final</li>
		<li>concentration) to the CldU reserve medium, and cover the cells with the CldU reserve medium. Place cells in the incubator for the duration of the CldU pulse for 20 min.</li>
		<li>Wash the cells with PBS, scrape off or trypsinize for 3-4 min, and then add 5 mL of medium to the plate, and collect the cells.</li>
		<li>Spin at 264 x g for 4 min. Remove the medium, and resuspend the cells in 1 mL of ice-cold PBS. Determine the cell numbers using a trypan blue assay (hemocytometer counting), and then dilute the cells to ~200-400 cells/&#181;L. Keep the cell suspension on ice during the cell counting. 
		NOTE: If possible, limit the time the cells sit on the ice. This may help obtain a bead of cell solution along the top of the slide. If they are kept on ice, keep them on for less than 30 min.</li>
	</ol>
	</li>
	<li>Day 1: Preparation of the DNA fibers
	<ol>
		<li>Using a pencil, label the slides (glass slide, 75 mm x 25 mm) with the experimental condition and date.</li>
		<li>Take 2 &#181;L of cells in a pipette, hold the pipette at a ~45&#176; angle, place the pipette about 1 cm below the slide label, and move the pipette horizontally to the slide label. As the pipette moves across the slide, release a little of the cell solution at a time. Make multiple lines of cells on each slide (critical step). 
		NOTE: There should be a horizontal line of cell suspension across the slide. If the cell suspension beads up, then add 5-10 &#181;L of cell medium to the container that has the cell suspension, as this may help the solution to adhere to the slide when being applied again.</li>
		<li>Let the solution with the cells evaporate until the solution looks sticky and tacky. At this point, some liquid is associated with the cells, but not much. This step takes from 8-20 min and varies depending on how much humidity is in the air. 
		NOTE: Ensure that all the solution has not evaporated, since this makes it very hard to obtain straight and aligned DNA fibers as most, if not all, of the DNA will be stuck together and unable to separate.</li>
		<li>When the solution is tacky, overlay the cells with 15 &#181;L of spreading (lysis) buffer per each line of cells, taking care not to let the pipette tip touch the slide. Wait for 10 min. Tilt the slide at a 25&#176; angle, and place the label of the slide horizontally against the edge of a tube rack (the bottom part of the label should line up with the edge of the tube rack).</li>
		<li>Allow the DNA spreads to air dry for a minimum of 4 h from the time the last slides were made. 
		NOTE: A period of 4 h to 12 h is good for drying. However, do not let them dry overnight. If allowed to dry overnight, there will be a lot of relaxed DNA but very few straight and aligned DNA fibers.</li>
	</ol>
	</li>
	<li>Day 1: Fixing the DNA fibers and freezing
	<ol>
		<li>Once the slides have dried, fix the slides with methanol:acetic acid (3:1) by immersing the slides for 2 min. Perform this step under a hood, and let the slides dry overnight in an area with limited light exposure, or cover the slides with a tin foil tent.</li>
	</ol>
	</li>
	<li>Day 2
	<ol>
		<li>Place the slides in a glass slide carrier. Place the slides at &#8722;20 &#176;C for a minimum of 24 h. This step improves the resolution or crispness of the image.</li>
	</ol>
	</li>
</ol>
3. Performing the DNA fiber assay
<ol>
	<li>Denaturation of the DNA
	<ol>
		<li>Prepare a humidified chamber using small pipette tip boxes with lids.&#160;Half-fill the containers with water, and place them in a 37 &#176;C water bath for at least 1 h.</li>
		<li>Take the slides out from &#8722;20 &#176;C, and defrost them for a few seconds. Place the slides carefully into a Coplin jar containing deionized (DI) water so that the coated surfaces do not touch the walls of the jar. Incubate the slide for 20 s, and then carefully pour out the water.</li>
		<li>Add 2.5 M HCL to the Coplin jar such that it covers all the slides. Wait for 80 min. This is a critical step in the protocol. A change in the time can change the outcome of the image.</li>
		<li>Wash the slides 1x with PBS plus Tween (PBS + 0.1% Tween [final concentration]) and then 2x with PBS for 3 min each at room temperature (RT).</li>
	</ol>
	</li>
	<li>Immunostaining
	<ol>
		<li>Keep the slides in the humidified chamber for the following steps. Block the slides in 5% BSA in PBS for 30 min at RT, and cover them with coverslips, coverslips made from a Western blot bag, or transparent plastic sheets.</li>
		<li>After blocking, carefully remove the coverslip, remove the blocking solution, and blot the slide on a paper towel (tap the slide on the towel) to remove the excess blocking solution.</li>
		<li>Add 100 &#181;L of the primary antibody solution along the length of the slide. Add a new plastic coverslip, and wait for 2 h. When adding the coverslip, ensure that no bubbles form.</li>
		<li>After 2 h, knock off the antibody solution, and rinse the slides in a PBS-filled Coplin jar 2x at RT. Use fresh PBS each time to wash the slides.</li>
		<li>Block the slides again by adding 200 &#181;L (three to four drops) of 5% BSA along each slide, and add a plastic coverslip. Wait for 15 min.</li>
		<li>Take off the coverslip, knock off the excess BSA on a paper towel, and add 100 &#181;L of the secondary antibody solution along the length of the slide. Add a new plastic coverslip, and wait for 1 h.</li>
		<li>Take off the coverslip, knock off the excess BSA on a paper towel, and wash the slides 2x in PBS at RT.</li>
		<li>Block the slides again by adding 200 &#181;L (three to four drops) of 5% BSA along each slide, and add a plastic coverslip. Wait for 15 min.</li>
		<li>If needed, remove the coverslip, remove the excess BSA on a paper towel, and add 100 &#181;L of tertiary antibody solution along the length of the slide. Add new plastic coverslips, and wait for 30 min.</li>
		<li>Wash the slides with PBS 3x. Remove the excess PBS, and allow them to dry off completely.</li>
		<li>Once dry, add the mounting medium, and carefully place glass coverslips (24 mm x 60 mm) over the mounting medium so that bubbles are avoided. Re-label the slides, and leave them in the dark overnight.</li>
	</ol>
	</li>
</ol>
4. Image acquisition
<ol>
	<li>Visualize the immunostained DNA fibers using an immunofluorescent microscope equipped with a camera, a 60x objective, and appropriate filter sets to detect alexafluor 488 and 594 dyes.</li>
	<li>Take images for the analysis in regions where the fibers are well separated and not entangled. It is important to capture images in different areas along the slide, as taking images in only one part of the slide may not provide representative data regarding all the replication intermediates found on the slide.</li>
	<li>If possible, obtain fiber images from 20 different regions on a slide. Use only one channel to select the areas for taking images to avoid bias.</li>
</ol>

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J., Tirman, S., Quinet, A., Menck, C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+post-replicative+gaps+accumulation+and+repair+in+human+cells+using+the+DNA+fiber+assay.">Detection of post-replicative gaps accumulation and repair in human cells using the DNA fiber assay.</a> Journal of Visualized Experiments. (180), e63448 (2022).</li></ol>

The authors have no conflicts of interest to disclose.

We discuss here a method to assist in the analysis of the variation in DNA replication dynamics in the presence of nanoparticles through the DNA fiber assay. Major critical steps involved in the standard assay are described in the protocol (step 2.2.2 and step 3.1.3). It is always recommended to use an area with limited overhead light exposure and constant airflow to prevent light-induced DNA breaks in the slides as well as to enhance reproducibility. Careful attention is required on the time duration for the drying of t...

During each cell cycle, DNA replication ensures accurate genome duplication1. Eukaryotic chromosomal replication essentially depends on three factors: the timing of the firing of multiple replication origins, the speed of the forks that emerge from the fired origins, and the termination of the replication process when two replication forks from adjacent origins meet2.&#160;For the high-fidelity transmission of genetic information to daughter cells, as well as the preservation of genetic integrity, accurate DNA replication is crucial. Agents that develop from regular metabolism or are due to artificial or natural environmental materials are constantly attacking the genome. These endogenous and exogenous agents cause replication forks to slow down or stop due to encountering DNA damage caused by these agents, and the forks temporarily slowing down or stopping in response to these difficulties is termed replication stress3. In response to replication stress, cells have developed several molecular pathways that maintain the stability of the disturbed replication forks and allow them to restart4. In terms of genetic stability, cell survival, and human disease, these replication stress response mechanisms have emerged as the key factors for maintaining a healthy genome, ensuring cell survival, and decreasing the likelihood of disease formation5.
One of the exogenous agents capable of producing replication stress is nanoparticles. Nanoparticles are particles that range in size from 1 nm to 100 nm6. Due to their high surface areas, distinctive shapes, and unique chemical properties, nanoparticles are utilized in various medical, pharmaceutical, environmental, and industrial applications7,8. While nanoparticles have a lot of potential benefits, some of them (due to their inherited nature or longevity) can become toxic. Nanoparticles can also form due to the natural wear and tear of medical implants and be released into the peri-prosthetic region9,10.
Due to the exposure of humans to a myriad of nanoparticles produced for various applications, research in the field of nanoparticle toxicity has increased tremendously over the past 10 years11. While these research efforts have revealed information in abundance about the potential threat that nanoparticles pose to human health, knowledge about the potential for nanoparticles to cause genotoxicity is still limited. What has been discovered so far is that these nanoparticles can physically interact with the DNA, promote DNA damage, and damage or interfere with the proteins responsible for repairing or replicating DNA12. To detect how they interfere with DNA replication, DNA fiber combing, radioresistant DNA synthesis (RDS), and DNA fiber analysis are typically used13,14,15,16.
The DNA fiber combing method is flexible and gives information about replication fork dynamics at the single-molecule level17. In essence, a salinized coverslip is gently withdrawn from the DNA solution once the DNA ends bind to it. The DNA molecules are straightened and aligned by the solution&#39;s meniscus. The homogeneity, spacing, and alignment of the DNA fibers support accurate and dependable fiber tract length measurements. By adjusting the length and sequence of the treatments and the drugs used to cause stress or damage, many aspects of fork advancement can be monitored using this application. In this method, a dual labeling system is used, through which the speed and progression of the replication fork are assessed17,18. On the other hand, 2D gel electrophoresis takes advantage of the fact that, in agarose gel electrophoresis, branching DNA structures travel more slowly than linear DNA molecules of the same mass, allowing for the clean separation of the two in a 2D run. In fact, this method is investigated to segregate DNA molecules based on their mass in the first run and based on their shape in the second orthogonal run. After genomic DNA fragmentation, the uncommon replication and recombination intermediates develop a branching form, and they may be distinguished from the more common linear molecules in the 2D gel19.
The RDS method is used for determining how global DNA synthesis is impacted. In this method, the degree of inhibition of global replication is determined by comparing the amount of incorporated radioactively labeled nucleotides, such as [14C] thymidine, in untreated versus treated cells14,20. The percentage difference in radiolabeling between the untreated and treated cells represents the degree to which the DNA-damaging agent impacts DNA synthesis. Similar to this, another method uses the ability of cells to integrate nucleotide analogs like BrdU (5-bromo-2&#8242;-deoxyuridine) for flow cytometry to measure the overall rates of DNA synthesis21,22. While these methods demonstrate how DNA-damaging agents impact global DNA synthesis, they do not show how individual replicons are affected. Indeed, replicon-level assays are imperative to better understand the initiation and extent of genomic instability in the event of toxic particle (nanomaterial) exposure. DNA fiber autoradiography and electron microscopy are some methods used to determine this23,24,25,26.
The concepts of replication bubbles and bidirectional replication from unevenly spaced sources were first developed using single-molecule tests like electron microscopy and DNA fiber autoradiography27,28. The direct observation of branching replication intermediates on specific molecules dispersed across a carbon-coated grid is greatly facilitated by electron microscopy. This method, which is still in use today to track pathological shifts at replication forks, was utilized to locate the first eukaryotic origin of DNA replication28. Fiber autoradiography is centered around the concept of the autoradiographic identification of newly replicated areas and the pulse tagging of chromosomes with tritiated thymidine. The first quantitative evaluation of origin densities and replication fork rates in metazoan genomic sequences was made possible by DNA fiber autoradiography29.
Currently, fiber fluorography methods have taken the place of autoradiography, mainly because fiber fluorography is much faster than autoradiography. In fiber fluorography, two halogenated nucleotide derivatives, such as bromo- (Br), chloro- (Cl), or iododeoxyuridine (IdU), are sequentially incorporated into freshly replicated DNA and then identified by indirect immunofluorescence using antibodies30. Microscopic viewing of the nascent DNA that has incorporated one or both analogs is made possible by immunostaining one of the analogs in one color and the other analog in a different color (e.g., immunostaining nascent DNA with incorporated IdU red and incorporated CldU green) (Figure 1)21. Many different types of replication intermediates can be identified by DNA fiber analysis. The most commonly studied are individual elongating forks, initiations, and terminations. Individual elongating forks have a replication pattern of red followed by green (red-green; Figure 2A).13 The lengths of these intermediates are frequently used to gauge the fork speed (i.e., fork length/pulse time) or the exonucleolytic degradation of nascent DNA through track shortening (Figure 2E)30,31,32. In a study by Mimitou et al., it was found that upon long-term exposure to hydroxyurea, a replication poison that causes double-strand breaks in the DNA, RE11 was recruited33. MRE11 is an exonuclease known for its 3&#39;-5&#39; exonuclease activity, and it is capable of cutting the ends of DNA for repair. Therefore, when exposed to toxic agents, one may observe exonucleolytic degradation of nascent DNA, which is the shortening of the DNA strand due to exposure to a DNA-damaging agent34.
Replication fork breakages brought on by physical obstructions (DNA-protein complexes or DNA lesions), chemical impediments, or mutations may stop replication and necessitate homologous recombination to restart it. This is known as impaired fork progression. Numerous in vitro and in vivo investigations have indicated that transcription may, on occasion, prevent replication fork advancement in this manner35.
Initiations are replication origins that initiate and fire during the first or second pulse. Origins that fire during the first pulse and have replication forks that continue to be active have a green-red-green pattern (Figure 2B, lower). Origins that initiate during the second pulse have a green-only pattern (Figure 2B, upper) and are sometimes called newly initiated origins, so those origins can be differentiated from those that initiate during the first pulse. The comparison of the relative percentages of newly fired origins between two experimental conditions allows one to understand how a cell responds to a DNA-damaging agent or the presence or absence of a protein. Terminations are created when two replication forks from adjacent replicons merge, and they have a red-green-red pattern (Figure 2D)30.
Based on the facts described above, DNA fiber analysis is currently considered a preferred method to study the variation in DNA replication dynamics caused by toxic agents such as nanomaterials. Researchers now have a good understanding and knowledge of the dynamics of genome-wide DNA replication in eukaryotes, both quantitatively and qualitatively, owing to the discovery of this technology36. Based on the outcome variables, several methodologies can be adopted. Some examples of methods to study the variations in DNA damage induced by external agents/nanoparticles are shown in Figure 3. The overall goal of the DNA fiber analysis method described in this study is to determine how nanoparticles impact the replication process in vitro and how they differentially affect various tissues.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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		<tr>
			<td>24 well plate</td>
			<td>Fisher brand</td>
			<td>FB012929</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetic Acid&#160;</td>
			<td>Sigma Aldrich</td>
			<td>695092</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa flour 594 goat anti-rabbit&#160;</td>
			<td>Invitrogen</td>
			<td>A11037</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa fluor 488 chicken anti-rat&#160;&#160;</td>
			<td>Invitrogen</td>
			<td>A21470</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa fluor 488 goat anti-chicken&#160;</td>
			<td>Invitrogen</td>
			<td>A11039</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa fluor 594 rabbit anti-mouse&#160;</td>
			<td>Invitrogen</td>
			<td>A11062</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma Aldrich</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>CldU</td>
			<td>Sigma Aldrich</td>
			<td>50-90-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips (22 x 50 mm)</td>
			<td>Fisher brand</td>
			<td>12-545-EP</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Fisher Scientific</td>
			<td>15575020</td>
			<td></td>
		</tr>
		<tr>
			<td>Frosted Microscope Slides</td>
			<td>Fisher brand</td>
			<td>12-550-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric Acid</td>
			<td>Sigma Aldrich</td>
			<td>320331</td>
			<td></td>
		</tr>
		<tr>
			<td>IdU&#160;</td>
			<td>Sigma Aldrich</td>
			<td>54-42-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Fisher Scientific</td>
			<td>A454-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse Anti-BrdU&#160;</td>
			<td>BD Biosciences</td>
			<td>347580</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffer Saline</td>
			<td>Gibco</td>
			<td>10010072</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-BrdU&#160;</td>
			<td>Abcam</td>
			<td>BU1--75(ICR1)</td>
			<td></td>
		</tr>
		<tr>
			<td>Raw 264.5 macrophage cells&#160;</td>
			<td>ATCC</td>
			<td>TIB-71</td>
			<td></td>
		</tr>
		<tr>
			<td>SDS</td>
			<td>Sigma Aldrich</td>
			<td>L3771</td>
			<td></td>
		</tr>
		<tr>
			<td>Silane-Prep slides</td>
			<td>Sigma Aldrich</td>
			<td>S4651-72EA&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost gold plus slides</td>
			<td>Fischer scientific</td>
			<td>22-035813</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris pH 7.4</td>
			<td>Sigma Aldrich</td>
			<td>77861</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma Aldrich</td>
			<td>P9416</td>
			<td></td>
		</tr>
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demonstration of the dna fiber assay for investigating dna damage and repair dynamics induced by nanoparticles

Nanomaterial exposure can cause replication stress and genomic instability in cells. The degree of instability depends on the chemistry, size, and concentration of the nanomaterials, the time of exposure, and the exposed cell type. Several established methods have been used to elucidate how endogenous/exogenous agents impact global replication. However, replicon-level assays, such as the DNA fiber assay, are imperative to understand how these agents influence replication initiation, terminations, and replication fork progression. Knowing this allows one to understand better how nanomaterials increase the chances of mutation fixation and genomic instability. We used RAW 264.7 macrophages as model cells to study the replication dynamics under graphene oxide nanoparticle exposure. Here, we demonstrate the basic protocol for the DNA fiber assay, which includes pulse labeling with nucleotide analogs, cell lysis, spreading the pulse-labeled DNA fibers onto slides, fluorescent immunostaining of the nucleotide analogs within the DNA fibers, imaging of the replication intermediates within the DNA fibers using confocal microscopy, and replication intermediate analysis utilizing a computer-assisted scoring and analysis (CASA) software.

After obtaining enough images (from 20-100 images per condition), the replication intermediates need to be identified, measured, and counted. Whether analyzing the fibers manually or automatically via a program38, it is necessary to clearly define what characteristics a fiber must have for it to be counted or scored (or not counted or measured)39. For instance, the following questions can be considered. (1) Should one measure and count only fibers with 100% immunof...

Watch this Scientific Journal Video about Demonstration of the DNA Fiber Assay for Investigating DNA Damage and Repair Dynamics Induced by Nanoparticles at JoVE.com

Demonstration of the DNA Fiber Assay for Investigating DNA Damage and Repair Dynamics Induced by Nanoparticles

The methodology assists in the analysis of variation in DNA replication dynamics in the presence of nanoparticles. Different methodologies can be adopted based on the cytotoxicity level of the material of interest. In addition, a description of the image analysis is provided to help in DNA fiber analysis.

Nanomaterial exposure can cause replication stress and genomic instability in cells. The degree of instability depends on the chemistry, size, and ...

demonstration-dna-fiber-assay-for-investigating-dna-damage-repair

Research

JoVE Journal

Biology

2.7K Views.  University of Illinois College of Medicine Rockford. The methodology assists in the analysis of variation in DNA replication dynamics in the presence of nanoparticles. Different methodologies can be adopted based on the cytotoxicity level of the material of interest. In addition, a description of the image analysis is provided to help in DNA fiber analysis.

Video: Demonstration of the DNA Fiber Assay for Investigating DNA Damage and Repair Dynamics Induced by Nanoparticles

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

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

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

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

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

Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not only ensure faithful genome duplication in the face of endogenous and exogenous DNA damage but also prevent genomic instability, a recognized causative factor in tumor development.
Here, we describe a simple and cost-effective fluorescence microscopy-based method to visualize DNA replication in the avian B-cell line DT40. This cell line provides a powerful tool to investigate protein function in vivo by reverse genetics in vertebrate cells1. DNA fiber fluorography in DT40 cells lacking a specific gene allows one to elucidate the function of this gene product in DNA replication and genome stability. Traditional methods to analyze replication fork dynamics in vertebrate cells rely on measuring the overall rate of DNA synthesis in a population of pulse-labeled cells. This is a quantitative approach and does not allow for qualitative analysis of parameters that influence DNA synthesis. In contrast, the rate of movement of active forks can be followed directly when using the DNA fiber technique2-4. In this approach, nascent DNA is labeled in vivo by incorporation of halogenated nucleotides (Fig 1A). Subsequently, individual fibers are stretched onto a microscope slide, and the labeled DNA replication tracts are stained with specific antibodies and visualized by fluorescence microscopy (Fig 1B). Initiation of replication as well as fork directionality is determined by the consecutive use of two differently modified analogues. Furthermore, the dual-labeling approach allows for quantitative analysis of parameters that influence DNA synthesis during the S-phase, i.e. replication structures such as ongoing and stalled forks, replication origin density as well as fork terminations. Finally, the experimental procedure can be accomplished within a day, and requires only general laboratory equipment and a fluorescence microscope.

DT40, a model vertebrate genetic system, provides a powerful tool to analyze protein function. Here we describe a simple method that allows qualitative analysis of parameters that influence DNA synthesis during the S-phase in DT40 cells at the single molecule level.

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not ...

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

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

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

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

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

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

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

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

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

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

gDNA Enrichment by a Transposase-based Technology for NGS Analysis of the Whole Sequence of BRCA1, BRCA2, and 9 Genes Involved in DNA Damage Repair

The widespread use of Next Generation Sequencing has opened up new avenues for cancer research and diagnosis. NGS will bring huge amounts of new data on cancer, and especially cancer genetics. Current knowledge and future discoveries will make it necessary to study a huge number of genes that could be involved in a genetic predisposition to cancer. In this regard, we developed a Nextera design to study 11 complete genes involved in DNA damage repair. This protocol was developed to safely study 11 genes (ATM, BARD1, BRCA1, BRCA2, BRIP1, CHEK2, PALB2, RAD50, RAD51C, RAD80, and TP53) from promoter to 3&#39;-UTR in 24 patients simultaneously. This protocol, based on transposase technology and gDNA enrichment, gives a great advantage in terms of time for the genetic diagnosis thanks to sample multiplexing. This protocol can be safely used with blood gDNA.

gDNA Enrichment by a Transposase-based Technology for NGS Analysis of the Whole Sequence of BRCA1, BRCA2, and 9 Genes Involved in DNA Damage Repair

gDNA enrichment for NGS sequencing is an easy and powerful tool for the study of constitutional mutations. In this article, we present the procedure to analyse simply the complete sequence of 11 genes involved in DNA damage repair.

The widespread use of Next Generation Sequencing has opened up new avenues for cancer research and diagnosis. NGS will bring huge amounts of new data on ...

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

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

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

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

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

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

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

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

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