The authors wish to thank Drs. Philina S. Lee and Alan D. Grossman for initially training L.A.S. in fluorescence microscopy. The authors also thank Drs. Melanie Berkmen and Hajime Kobayashi for help and tips for imaging. This work was supported by start-up funds from the College of Literature, Science &amp; Arts and from the Department of Molecular, Cellular, and Developmental Biology at the University of Michigan.

Growing cultures of cells for microscopy
1. One or two days prior to imaging, prepare the B. subtilis strain containing the translational fusion protein you wish to visualize. Trial rounds of steps 1A and 1B will be necessary to determine which growth condition provides the best images of your strain. In most cases, cells should be imaged during exponential growth phase.
<ol class="ualpha">
<li>For some B. subtilis strains (in particular, strains that grow poorly due to the integration of the translational fusion between the gene of interest and GFP at its endogenous locus), initial growth for two days prior to imaging is necessary for high quality images. On the first day, streak the B. subtilis strain to be imaged on LB agar with selective antibiotic(s) and incubate overnight at 30&deg;C. On the second day, inoculate 100 &mu;l of 0.85% saline with a single colony and perform three 10-fold serial dilutions on LB agar plates (with selective antibiotics), plating 100 &mu;L of each dilution, followed by incubation overnight at 30&deg;C. On the third day, select the dilution plate with light confluent growth of cells. Light confluent growth on a plate will provide the growth medium with a healthy, exponentially growing starter inoculum to potentially yield excellent imaging results (see step 4). This step is particularly useful for the highest quality membrane imaging with the stain FM4-64. </li>
<li>Other B. subtilis strains may simply be applied to selective LB agar medium for single colonies with a sterile stick the day prior to imaging and incubate overnight at 30&deg;C. The following day, cells on this plate will be sufficient to use as an inoculum for imaging (see step 4). NOTE: For B. subtilis, overnight cultures in liquid medium are not appropriate due to a combination of spore formation and quorum responses, which will cause an extended period of lag phase growth 1.</li>
</ol>
2. On the morning of imaging, place 10 mL of S750 1-3 medium into a 125 mL sterile Erlenmeyer flask. 
3. Resuspend the cells in up to 2 mL of S750 medium from the LB agar plate with single colonies or the light confluent cells to obtain an inoculum. 
4. Add enough S750 medium-cell inoculum to the Erlenmeyer flask containing 10 mL S750 medium for a culture with an initial optical density measured at 600 nm (OD600) of ~0.08 to 0.1. At this step it is important to have at least a 10-fold air to medium ratio. Thus, we use 10 to 12.5 mL of inoculated medium in a 125 mL Erlenmeyer flask. 
5. Grow each culture at 30&deg;C until the culture reaches an OD600 ~0.5-0.8. Shaking water baths are preferred with revolutions per minute from 150-200. Cultures challenged with DNA damaging or mismatch inducing agents should be at an early exponential growth phase OD600 such that the cells will reach the target OD600 with the additional period of growth that is required for treatment (see Table 1) 
 For example, 2-aminopurine requires one hour of treatment, and therefore a culture treated with 2-aminopurine should be at OD600~0.3-0.4, such that the one hour of treatment/growth results in an OD600~0.5-0.8, whereby cells are ready for imaging. 
Sample Preparation
1. Pipette 300 &mu;l of cultured cells into a 1.5 mL microcentrifuge tube.
2. If imaging of cellular membranes is desired, add 1:1000 dilution of FM4-64 membrane stain (Invitrogen) to each sample from a 1 mg/mL stock solution. A titration of FM4-64 may be required to achieve the desired fluorescence signal. A typical titration range is 1:100, 1:1000, and 1:10,000.
3. Allow the samples to sit at room temperature for up to ten minutes or while the slide is being prepared with 1% agarose in 1X Spizizens medium as described below (see step 1 of "slide preparation").
4. If concentration of cells is required, we prefer filtering cells using a 0.2 &mu;m filter with a vacuum apparatus. Cells can then be gently washed from the surface of the filter with PBS, another appropriate buffer or medium prior to visualization.
Slide Preparation
1. Prepare 1% agarose with 1X Spizizens. Add 5 mL of 10X Spizizens stock to 45 ml ddH2O along with 0.5 g agarose, and microwave to melt. Hot molten agarose is difficult to pipette, and solidified agarose will not enter the pipette tip. The agarose should be equilibrated in a water bath to a temperature of ~55-65&deg;C for it to enter the pipette tip without solidifying within the tip.
2. Draw 20 &mu;l of Spizizens with 1% melted agarose into the pipette tip. Pipette a single drop (approximately 1-2 &mu;L) of the agarose solution into each slide well (we use 15-well slides, see Table 2) to form shallow agarose pads. The agarose will solidify immediately. Slides must also be made immediately prior to application of the cells. If the agarose pads are left at room temperature alone, they can dehydrate within minutes and become unusable. 
Agarose pads should be centered and fill each well but have minimum height to avoid disrupting the coverslip; if droplets are too high (bubble-like) or if the sample spreads beyond the well boundary, simply swipe the well clean with a Kimwipe and apply a new agarose pad. 
3. Apply the entire 300 &mu;l sample (see step 6) to the slide, distributed on the top of each agarose pad. This amount of cells should be sufficient to cover each well on the slide.
4. Allow the slide loaded with cells to sit at room temperature (23&deg;C) for approximately ten minutes. This step will allow for the cells to settle on the agarose pad becoming immobile and ready for imaging. For specialized needs this temperature may need to be changed, for example, when performing experiments that require a temperature shift4.
5. Aspirate away the excess medium from each well without removing the pads themselves.
6. Apply the coverslip to the slide, pressing firmly and evenly, but gently, across the area of the slide. Check to make sure that the cover slip is against the slide and that the cover slip is not raised above the slide.
Visualization of cells
1. Many different fluorescence microscopes will work well. It is critical to have a 100X oil immersion lens. The Simmons Lab uses:
<ul>
 <li>Olympus BX61 microscope equipped with 1,45 NA TIRFM 100X oil immersion objective lens</li>
 <li>Hamamatsu ORCAR2 CCD cooled camera</li>
 <li>Lumen 200 arc metal (Prior) light source. </li>
 <li>For detection of GFP (FITC), filter excitation 460-500 and emission 510-560</li>
 <li>For detection of FM4-64 (TRITC), filter excitation 510-560 and emission 572-648. </li>
 <li>Images were captured using SlideBook 4.2 (Figure 1) </li>
</ul>
2. Bring the cells into focus using white light.
3. Capture the image using appropriate filters for each fluorophore. 
Representative Results
Representative images are shown (Figure 1). GFP foci should be well defined, while FM4-64 staining should be bright and clear 4-7. GFP or membrane images can be pseudo-colored with any color, to allow for the highest quality image to be presented without losing any data.
<img src="/files/ftp_upload/1736/1736fig1.jpg" alt="Figure 1" /> Figure 1: Representative GFP images of B. subtilis translational fusion proteins. GFP proteins are shown in green, while FM4-64 membrane staining is shown in red. The white scale bar indicates 3 &mu;m. (A) MutS-GFP [relevant genotype: mutS-gfp (spc), amyE::PspacmutL (cat)] in the presence of 600 &mu;g/mL 2-aminopurine and 1 mM IPTG. FM4-64 signal is not presented in order to more clearly show the MutS-GFP foci. (B) MutL-GFP [relevant genotype: mutL::mutL-GFP (spc)] in the presence of 2-aminopurine. (C) DnaX-GFP [relevant genotype: dnaX::dnaX-GFP (spc)]. (D) RecA-GFP [relevant genotype: recA::recA-GFP (spc)] in the presence of 20 ng/mL mitomycin C. (E) TagC-GFP [relevant genotype: tagC::tagC-GFP (spc)] in the presence of 1 &mu;g/mL mitomycin C. 

<table border="1" cellspacing="0" cellpadding="0">
<tbody>
 <tr>
 <td>Treatment /concentration or amount</td>
 <td>Function</td>
 <td>Time of incubation</td>
 </tr>
 <tr>
 <td>Mitomycin C (MMC) [20 to 150 ng/mL]</td>
 <td>DNA alkylating agent</td>
 <td>1 hour</td>
 </tr>
 <tr>
 <td>2-aminopurine (2-AP) [600 &mu;g/mL]</td>
 <td>mismatch inducing</td>
 <td>1 hour</td>
 </tr>
 <tr>
 <td>Hydroxyurea (HU)</td>
 <td>depletes dNTPs</td>
 <td>3 hours</td>
 </tr>
 <tr>
 <td>ultraviolet light (UV) 20 J/m2</td>
 <td>Thymine-thymine dimers and 6-4 photoproducts</td>
 <td>varies depending on source, usually 20 seconds</td>
 </tr>
 <tr>
 <td>grays (ionizing radiation) 5 to 100 Gy</td>
 <td>Double strand breaks, single strand breaks and base damage sites</td>
 <td>varies depending on source </td>
 </tr>
</tbody>
</table>
Table 1. Summary of DNA damaging and mismatch inducing treatments

<table border="1" cellspacing="0" cellpadding="0">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Multitest slide, 15-well</td>
 <td>MP Biomedicals</td>
 <td>6041505E</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope Cover Glass</td>
 <td>Fisher</td>
 <td>12-544-B</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Agarose</td>
 <td>Fisher</td>
 <td>BP160-500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Mitomycin C</td>
 <td>Sigma</td>
 <td>M0503-2MG</td>
 <td>mutagen, wear gloves</td>
 </tr>
 <tr>
 <td>2-Aminopurine</td>
 <td>Sigma</td>
 <td>A3509-250MG</td>
 <td>gloves</td>
 </tr>
 <tr>
 <td>Hydroxyurea</td>
 <td>Sigma</td>
 <td>H8627-25G</td>
 <td>gloves</td>
 </tr>
 <tr>
 <td>FM4-64</td>
 <td>Invitrogen</td>
 <td>T13320</td>
 <td>light sensitive</td>
 </tr>
</tbody>
</table>
Table 2. List of specific reagents and equipment:

<ol>
	<li>Hardwood, C. R., Cutting, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Molecular Biological Methods for Bacillus. , (1990).</li><li>Berkmen, M. B., Grossman, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+and+temporal+organization+of+the+Bacillus+subtilis+replication+cycle.">Spatial and temporal organization of the Bacillus subtilis replication cycle.</a> Mol. Microbiol. 62, 57-71 (2006).</li><li>Simmons, L. A., Grossman, A. D., Walker, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clp+and+Lon+proteases+occupy+distinct+subcellular+positions+in+Bacillus+subtilis.">Clp and Lon proteases occupy distinct subcellular positions in Bacillus subtilis.</a> J. Bacteriol. 190, 6758-6768 (2008).</li><li>Simmons, L. A., Grossman, A. D., Walker, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replication+is+required+for+the+RecA+localization+response+to+DNA+damage+in+Bacillus+subtilis.">Replication is required for the RecA localization response to DNA damage in Bacillus subtilis.</a> Proc. Natl. Acad. Sci. U S A. 104, 1360-1365 (2007).</li><li>Simmons, L. A., Davies, B. W., Grossman, A. D., Walker, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=b+Clamp+Directs+Localization+of+Mismatch+Repair+in+Bacillus+subtilis.">b Clamp Directs Localization of Mismatch Repair in Bacillus subtilis.</a> Mol. Cell. 29, 291-301 (2008).</li><li>Simmons, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+responses+to+double-strand+breaks+between+Escherichia+coli+and+Bacillus+subtilis+reveals+different+requirements+for+SOS+induction.">Comparison of responses to double-strand breaks between Escherichia coli and Bacillus subtilis reveals different requirements for SOS induction.</a> J. Bacteriol. 191, 1152-1161 (2009).</li><li>Smith, B. T., Grossman, A. D., Walker, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+mismatch+repair+in+bacterial+cells.">Visualization of mismatch repair in bacterial cells.</a> Mol. Cell. 8, 1197-1206 (2001).</li></ol>

Trial and error are required to find exposure conditions for the highest quality images for each strain; we find that 1 millisecond is appropriate for white light images, while exposures of 100 to 2000 ms are appropriate for GFP (FITC) and FM4-64 (TRITC) images. Exposure time will vary depending on the imaging equipment used. We recommend the use of one strain per 15-well microscope slide for the simplest imaging, as pad quality and diffusion of cells from the pad border could complicate strain differentiation if multipl...

<table><tbody><tr><td colspan="4">Specific solution recipes&lt;sup&gt;1&lt;/sup&gt;:</td></tr><tr><td colspan="4">&lt;strong&gt;10x S7&lt;sub&gt;50&lt;/sub&gt; salts&lt;/strong&gt;&lt;br/&gt; 0.5 M MOPS&lt;br/&gt; 100 mM Ammonium Sulfate&lt;br/&gt; 50 mM Potassium Phosphate Monobasic&lt;br/&gt; Filter sterilize, and wrap S7&lt;sub&gt;50&lt;/sub&gt; media in foil prior to storage</td></tr><tr><td colspan="4">&lt;strong&gt;100x Metals&lt;/strong&gt;&lt;br/&gt; 0.2 M MgCl&lt;sub&gt;2&lt;/sub&gt;&lt;br/&gt; 70 mM CaCl&lt;sub&gt;2&lt;/sub&gt;&lt;br/&gt; 5 mM MnCl&lt;sub&gt;2&lt;/sub&gt;&lt;br/&gt; 0.1 mM ZnCl&lt;sub&gt;2&lt;/sub&gt;&lt;br/&gt; 100 &amp;#x03BC;g/mL Thiamine HCl&lt;br/&gt; 2 mM HCl&lt;br/&gt; 0.5 mM FeCl&lt;sub&gt;3&lt;/sub&gt;*&lt;br/&gt; dH&lt;sub&gt;2&lt;/sub&gt;O to final volume&lt;br/&gt; *FeCl&lt;sub&gt;3&lt;/sub&gt; should be added last, to prevent precipitation.&lt;br/&gt; After filter sterilization, wrap 100x Metal solution in foil.</td></tr><tr><td colspan="4">&lt;strong&gt;S7&lt;sub&gt;50&lt;/sub&gt; media&lt;/strong&gt;&lt;br/&gt; 1x S7&lt;sub&gt;50&lt;/sub&gt; salts&lt;br/&gt; 1x Metal&lt;br/&gt; 1% Glucose&lt;br/&gt; 0.1% Glutamate&lt;br/&gt; 40 &amp;#x03BC;g/mL Tryptophan&lt;br/&gt; 40 &amp;#x03BC;g/mL Phenylalanine&lt;br/&gt; distilled H&lt;sub&gt;2&lt;/sub&gt;O to final volume</td></tr><tr><td colspan="4">&lt;strong&gt;10x Spizizens (grams/L)&lt;/strong&gt;&lt;br/&gt; 151.4 mM Ammonium Sulfate (20g/L)&lt;br/&gt; 803.8 mM Potassium Phosphate Monobasic (140g/L)&lt;br/&gt; 440.9 mM Potassium Phosphate Dibasic (60g/L)&lt;br/&gt; 34.0 mM Sodium Citrate (10g/L)&lt;br/&gt; 16.6 mM MgSO&lt;sub&gt;4&lt;/sub&gt; (2g/L)&lt;br/&gt; dH&lt;sub&gt;2&lt;/sub&gt;O to final volume&lt;br/&gt; Filter sterilize</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis at JoVE.com

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.

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 ...

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10.9K Views.  University of Michigan-Ann Arbor. A detailed protocol is described for imaging the real time formation of DNA repair complexes in Bacillus subtilis cells.

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

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 ...

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 ...

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

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

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

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

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 ...

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types and is widely believed to be directly associated with the development of cancer in humans. DNA damage, such as that induced by UV irradiation, severely impairs the ability of replication to duplicate the genomic template accurately. A number of gene products have been identified that are required when replication encounters DNA lesions in the template. However, a remaining challenge has been to determine how these proteins process lesions during replication in vivo. Using Escherichia coli as a model system, we describe a procedure in which two-dimensional agarose-gel analysis can be used to identify the structural intermediates that arise on replicating plasmids in vivo following UV-induced DNA damage. This procedure has been used to demonstrate that replication forks blocked by UV-induced damage undergo a transient reversal that is stabilized by RecA and several gene products associated with the RecF pathway. The technique demonstrates that these replication intermediates are maintained until a time that correlates with the removal of the lesions by nucleotide excision repair and replication resumes.

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

We present a procedure by which two-dimensional agarose-gel analysis can be used to identify the structure of replication intermediates that occur following UV irradiation.

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types ...

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 ...

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA insults. Genotoxic agents can damage the DNA backbone, break it, or modify the chemical nature of individual bases. Following DNA insult, DNA damage response (DDR) pathways are activated and proteins involved in the repair are recruited. A plethora of factors are involved in sensing the type of damage and activating the appropriate repair response. Failure to correctly activate and recruit DDR factors can lead to genomic instability, which underlies many human pathologies including cancer. Studies of DDR proteins can provide insights into genotoxic drug response and cellular mechanisms of drug resistance.
There are two major ways of visualizing&#160;proteins in vivo: direct observation, by tagging the protein of interest with a fluorescent protein and following it by live imaging, or indirect immunofluorescence on fixed samples. While visualization of fluorescently tagged proteins allows precise monitoring over time, direct tagging in N- or C-terminus can interfere with the protein localization or function. Observation of proteins in their unmodified, endogenous version is preferred. When DNA repair proteins are recruited to the DNA insult, their concentration increases locally and they form groups, or &#8220;foci&#8221;, that can be visualized by indirect immunofluorescence using specific antibodies.
Although detection of protein foci does not provide a definitive proof of direct interaction, co-localization of proteins in cells indicates that they regroup to the site of damage and can inform of the sequence of events required for complex formation. Careful analysis of foci spatial overlap in cells expressing wild type or mutant versions of a protein can provide precious clues on functional domains important for DNA repair function. Last, co-localization of proteins indicates possible direct interactions that can be verified by co-immunoprecipitation in cells, or direct pulldown using purified proteins.

Following DNA damage, human cells activate essential repair pathways to restore the integrity of their genome. Here, we describe the method of indirect immunofluorescence as a means to detect DNA repair proteins, analyze their spatial and temporal recruitment, and help interrogate protein-protein interaction at the sites of DNA damage.

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA ...

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

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

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

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

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

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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

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

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

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

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

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

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage