We thank Dr. Roger Tsien for providing us with the miniSOG constructs. Dr. Xuejun Sun for help with the TEM. Lisa Lem, and Peter Shipple from the Cross Cancer Institute for supplying oxygen. Hilmar Strickfaden holds a postdoctoral fellowship by the Alberta Cancer foundation and was supported by the Bayrische Forschungsallianz. This work was supported by grants from the Canadian Institutes of Health Research and Alberta Cancer Foundation.

1. Generation of miniSOG Cell-lines
<ol>
	<li>Grow U2OS (human osteosarcoma) cells in a 35 mm dish containing 2 ml&#160;of low glucose Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) that is supplemented with 10% foetal bovine serum (FBS) at 37 &#176;C in a humidified incubator with 5% CO2 atmosphere.</li>
	<li>Transfect the cells when they are 80% confluent by lipofection with purified transfection quality (A269/280 ratio 1.8-2.0) plasmid constructs containing the sequences for miniSOG and mCherry tagged repair proteins according to the protocol of the manufacturer of the transfection reagent19. The miniSOG expression plasmids can be ordered from the Tsien-Lab:&#160;<a href="http://www.tsienlab.ucsd.edu/Samples.htm">http://www.tsienlab.ucsd.edu/Samples.htm</a></li>
	<li>Sort the cells expressing the recombinant protein with the miniSOG and mCherry tag by fluorescence activated cell sorting two days post transfection. Collect cells with intermediate fluorescence intensity in order to avoid cells that are overexpressing20.</li>
	<li>Culture the positive cells on a 10 cm dish in 10 ml&#160;of DMEM medium that is supplemented with 10 % FBS and 0.6 &#181;g/ml&#160;of the selection agent G418 (Geneticin).</li>
	<li>After visible colonies have emerged on the plates, screen for mCherry positive colonies using an inverted fluorescence microscope and draw circles around them (onto the bottom of the plate) with a permanent marker. Use a low magnification air objective lens (e.g., 10x) and pick clones using sterile technique by carefully scratching the colonies off and slowly sucking them into a sterile 1,000 &#181;l&#160;pipette tip.</li>
	<li>Expand the selected colonies and freeze parts of them down using the growth medium described in step 1.1 supplemented with 10% dimethyl sulphoxide. Test the cells for DRF formation by growing them on sterile 18 x 18 mm2 cover slips and exposing them to 2 Gy of radiation. The irradiated cells stably expressing a miniSOG mCherry tagged repair protein must show the typical focal pattern characteristic of DSBs when visualized with a fluorescence microscope using a filter-set for imaging mCherry.</li>
</ol>
2. Cell Culture
<ol>
	<li>Culture U2OS cells stably expressing the miniSOG and mCherry tagged repair proteins under the conditions outlined in step 1.1. Grow cells to 80% confluency in a 35 mm diameter glass bottom dish containing 2 ml&#160;of DMEM. Make sure that the glue that attaches the cover slip to the plastic and the used plastic is resistant to aqueous and alcoholic solutions and resistant to the resin used!</li>
	<li>Before conducting the experiment, outline a small area of approximately 1 mm2 that contains cells expressing the ectopic proteins by scratching with a sterile diamond pen using a fluorescence microscope to identify the region of interest. Alternatively use a glass bottom dish that contains a coverslip with a pre-etched grid system built into the coverslip. 
	NOTE: If using high quality correlative microscopy combining ESI and fluorescence microscopic pictures21, make sure that the thickness of the coverslip is compatible with oil immersion objectives. It should have the thickness No. 1.5 (0.16-0.18 mm). Choose an area &#160;close to the center of the dish for the region to be examined in the TEM in order to avoid problems with polymerization of the resin that may occur near the edges (see point 4.10-4.13).</li>
</ol>
3. DNA Damage Induction
<ol>
	<li>Irradiate the cells with gamma radiation, use a radiomimetic drug, or induce the damage by laser micro irradiation on a confocal microscope (e.g., as described in18,22 or&#160;23).</li>
	<li>In this example, irradiate the cells with 2 and 6 Gy of gamma irradiation in a cesium-137 radiation source or laser microirradiate using the 405 nm solid state laser of a confocal microscope after sensitizing the cells for 20 min with 0.5 &#181;g/ml&#160;Hoechst.</li>
</ol>
4. Sample Preparation
<ol>
	<li>Fix the cells with 1 ml&#160;of 4% paraformaldehyde CAUTION in 0.1 M sodium cacodylate buffer or in 0.1 M phosphate buffer pH 7.4 for 30 min at RT in the dark. 
	CAUTION: Paraformaldehyde and sodium cacodylate are toxic! Wear gloves and dispose the toxic substances adequately.</li>
	<li>Wash the cells twice with 4 ml&#160;of 0.1 M sodium cacodylate buffer pH 7.4 or phosphate buffer pH 7.4.</li>
	<li>Treat the fixed cells with 2 ml&#160;of 50 mM glycine, 5 mM aminotriazole and 10 mM potassium cyanide CAUTION in 0.1 M sodium cacodylate buffer or 0.1 M phosphate buffer (pH 7.4) (check the pH!) for 30 min in order to block unreacted aldehyde groups and to suppress the reactive oxygen species generated by heme groups, which can increase the background. 
	CAUTION: Potassium cyanide is extremely toxic! Wear gloves, work under a hood and dispose the toxic substances adequately. The reaction is very sensitive to pH so it is important that the pH verified and accurate.</li>
	<li>Record the area that was selected in step 2.2 in 2D or 3D on an inverted fluorescence microscope, if correlative microscopy is desired.</li>
	<li>Prepare a 1 mg/ml&#160;diaminobenzidine hydrochloride CAUTION solution by placing a 10 mg diaminobenzidine hydrochloride tablet into a 2 ml&#160;microcentrifuge tube. Add 980 &#181;l&#160;of distilled H2O and 20 &#181;l of concentrated hydrochloric acid (11.65 M) and mix until the solution is light brown and translucent. Dilute this solution in 0.1 M phosphate or 0.1 M sodium cacodylate buffer to a final concentration of 1 mg/ml&#160;while adjusting the pH with sodium hydroxide to a pH between 7.0 and 7.6 (pH 7.4 is recommended). 
	CAUTION: Diaminobenzidine is toxic! Wear gloves and dispose the toxic substances adequately.</li>
	<li>For photooxidation, replace the buffer with 2 ml&#160;of a 1 mg/ml&#160;diaminobenzidine hydrochloride solution in 0.1 M sodium cacodylate buffer or phosphate buffer. Protect this solution from light and cool it down on ice to 4 oC in order to increase the solubility of oxygen. Saturate the solution with oxygen by bubbling oxygen through the solution (use a hose that is inserted into the solution and connected to an oxygen bottle). Check the pH again and adjust if necessary. 
	NOTE: It is crucial for the photooxidation reaction that the pH is between pH 7.0 and pH 7.6.</li>
	<li>Mount the dish with the fixed cells carefully onto an inverted fluorescence microscope equipped with a 40x oil immersion objective lens and move it to the area of interest. Excite the miniSOG tag with blue light by using filter cubes for GFP or CFP. MiniSOG has an excitation maximum at 448 nm with a shoulder at 473 nm10.</li>
	<li>Continue with the illumination even after the green miniSOG fluorescence has completely disappeared and until the brown photo-oxidation product of the polymerized diaminobenzidine emerges in the transmitted light channel. When the oxidation product is visible in most of the cells, turn off the fluorescence illumination to stop the photo-oxidation. 
	NOTE: Photo-oxidation can take up to several minutes depending on the objective lens, filter transmission wavelengths and efficiency, and the illumination source.</li>
	<li>Postfix the cells with 1 ml&#160;of 2% glutaraldehyde in 0.1 M sodium cacodylate pH 7.4 buffer or phosphate buffer pH 7.4 for 30 min.</li>
	<li>Fix the membranes of the cells with 0.1%-0.5% osmium tetroxide for 20 min in 0.1 M sodium cacodylate buffer pH 7.4 or in 0.1 M phosphate buffer pH 7.4. 
	NOTE: It is recommended to use the lowest possible concentration of osmium tetroxide required to stabilize the membranes. Since the polymerized DAB precipitates have a high density of nitrogen, which can be detected directly by ESI, a strong osmium staining is not necessary to identify the miniSOG-tagged protein and might be detrimental to the ESI. Osmium tetroxide is very toxic! Work in a fume hood and dispose the toxic waste adequately.</li>
	<li>Dehydrate, the cells through an ethanol series using 30%, 50%, 70%, 90%, 98% 100% ethanol steps (each 5 min). Subsequently, incubate the cells in a 1:1 mix of 100% ethanol and acrylic resin (LR White) and put the dish onto a shaker for 4 hr to facilitate infiltration into the cells before incubating the cells for at least another 4 hr in 100% acrylic resin (LR White).</li>
	<li>In order to polymerize the resin, cut off the lid of a labeled 2 ml&#160;microcentrifuge tube with a razor blade and coat the rim with acrylic resin accelerator. Make sure that the accelerator only covers the rim and does not flow into the tube. Subsequently, carefully fill the tube approximately two-thirds with LR White resin using a Pasteur pipette. Take care that the resin does not get into contact with the accelerator on the rim!</li>
	<li>Remove the resin that infiltrated the cells in the dish and place the dish upside down onto the upright standing microcentrifuge tube so that the width of the tube almost completely fills up the glass covered observation window of the dish.</li>
	<li>Wait 1 to 2 min&#160;for the accelerator to seal the tube and the coverslip, then invert the tube so that the resin in the tube now covers the cells.</li>
	<li>Place the dish with the tube into an oven at 60 &#176;C and cure it for 12 hr.</li>
	<li>When the block is cured, remove the dish with the attached resin-filled microcentrifuge tube from the oven and separate the microcentrifuge tube from the dish. Facilitate separation by freeze-thaw cycles in liquid nitrogen and hot water.</li>
	<li>Discard the glass bottom dish and carefully cut open the microcentrifuge tube with a razor blade in order to remove the block.</li>
	<li>Label the block with a permanent marker.</li>
	<li>Use a razor blade to trim the block so that nothing but the 1 mm2 region that contains the area previously marked with the diamond or tungsten pen (or contains the area with the cells of interest) remains.</li>
	<li>Mount the block in an ultramicrotome, trim the block with a trimming knife and cut ultra-thin sections of approximately 50 nm using a diamond knife.</li>
	<li>Pick up the sections on high-transmission 300 mesh grids. These grids have very small grid bars and thus cover fewer cells in the area of interest.</li>
	<li>Coat the sections on the grids with about 0.2-0.4 nm of carbon using a carbon coater to help stabilize them under the electron beam of the TEM.</li>
</ol>
5. Electron Microscopy
<ol>
	<li>Load the grids into a TEM that is equipped with an energy filter. After tuning the microscope and the energy filter according to the manufacturer&#8217;s instructions, inspect the cells on the sections in the low magnification mode (normal transmission) and compare them to fluorescence data when appropriate (obtained in step 4.4).</li>
	<li>Once a nucleus with interesting features (DNA damage track or DNA repair foci) is found, switch to the energy filtering mode and record a thickness map. 
	NOTE: This procedure includes the recording of a zero-loss and an unfiltered image. Thicknesses up to 0.3 mean free path (30% of the electrons of the incident beam gets scattered by the sample) are thin enough to generate good elemental maps.</li>
	<li>Record ratio maps of the elements phosphorus and nitrogen using the software that controls the energy filter of the microscope used. Record the phosphorus map post edge images at 175 eV energy loss with a slit width of 20 eV and pre-edge images at 120 eV energy loss, also with a slit width of 20 eV. For the nitrogen maps, record post edge images at 447 eV with a slit width of 35 eV and pre-edge images at 358 eV, also with a slit width of 35 eV. 
	NOTE: Since the content of the imaged elements (phosphorus and nitrogen) is relatively low (approx. 1%), choose &#8220;Ratio Image&#8221; (qualitative elemental map) over the &#8220;3 window method&#8221; (quantitate elemental map) in order to generate images with a better signal to noise ratio.</li>
</ol>
6. Image Processing
<ol>
	<li>Open the elemental ratio maps in an image processing software that can handle the file format of the acquired pictures (e.g., Digital Micrograph). Copy the nitrogen map into the red channel and the phosphorus map into the green channel of an RGB image, then superimpose and align the maps.</li>
	<li>Export the composite image as a tagged image file format (TIFF) file. Open the image in an image processing software that can use layers (e.g., Photoshop).</li>
	<li>Adjust the dynamic range of each channel by rescaling the minimum and maximum values in the image to an 8-bit data set (0 to 255).</li>
	<li>Subtract the phosphorus content from the nitrogen map (using the &#8220;Layers&#8221; window) in order to generate a qualitative map that shows the distribution of protein.</li>
	<li>Convert the maps of the nucleic acid (phosphorus) and the protein (nitrogen)&#160;created in the previous step into &#8220;indexed color&#8221; and create a yellow lookup table in the CMYK color space for the map showing the nucleic acid distribution and a cyan lookup table to show the non-nucleoprotein map. The colors are chosen to generate maximum contrast between the two elemental maps.</li>
	<li>Create a new image and import the maps showing the distributions of phosphorus and protein as different layers. Use the &#8220;screen&#8221; transparency mode in order to see both layers superimposed on each other.</li>
	<li>Open the zero-loss image acquired in step 5.2. This image represents an image of the recorded area that looks like a conventional TEM image but only contains the electrons that have passed through the specimen without colliding with the specimen. Export this image as a TIFF image.</li>
	<li>Open the exported zero loss image in a photo editor and copy it into the uppermost layer of the image showing the elemental map. Align the image using the &#8220;screen&#8221; transparency mode.</li>
	<li>Optionally threshold the zero-loss image to segment the signal that originates from the miniSOG. Add the resulting image as a separate layer as described above.</li>
</ol>

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The authors have nothing to disclose.

ESI can serve as an excellent tool for examining different states of chromatin and ribonucleoproteins in the nucleus because it is able to specifically map areas that are rich in phosphorus. It profoundly augments the amount of detail that can be obtained by an electron microscope and is not dependent on nonspecific contrasting methods. One disadvantage of ESI is that it requires thinner sections than conventional TEM at the same accelerating voltage. This can be overcome by working with serial sections or using tomograp...

Despite the significant advancements in light microscopy over recent years1, cell-biologists still suffer from a gap in resolution. This limits understanding of the structure-function relationships in fundamental cellular processes that involve the coordinated interplay between macromolecular complexes (e.g., in chromatin remodelling, DNA repair, RNA transcription and DNA replication). Although transmission electron microscopes (TEM)s provide the required resolution, it has been challenging to define these processes structurally because of the inability to label specific proteins while also being able to determine the biochemical composition of the visualized structures. In the absence of internal membranes to help differentiate nuclear structures, the nucleus has been particularly challenging. Electron spectroscopic imaging (ESI) solves some of these limitations by allowing the simultaneous detection and differentiation of DNA, RNA, and protein-based nuclear structures2-5.
Electron spectroscopic imaging:
In order to map elemental distributions at high sensitivity and resolution in the electron microscope, one can use an imaging spectrometer that selects electrons that have been inelastically scattered through interactions with inner shell electrons of an element in the specimen6. Because element-specific amounts of energy are lost as a consequence of ionization of atoms in the specimen, these electrons can be separated and visualized using a spectrometer that is attached to the electron microscope. Thus, analysis of the spectrum of the electrons that have interacted with the specimen reveals qualitative and quantitative information about the elemental composition of the sample7. Electrons that do not lose energy when passing through the specimen are found in the &#8220;zero-loss peak&#8221; of the electron energy loss spectrum. The abundance of these electrons is related to the mass, density and thickness of the specimen and is comprised of electrons that pass through the specimen without colliding with the specimen or losing energy during passage through the specimen. This information can be useful for absolute quantification of the numbers of atoms of a specific element present in the specimen8.
Since biological samples consist mostly of light elements that poorly deflect the electrons in the incident beam of the TEM, staining methods using heavy metal salts have to be applied in order to generate contrast in the sample. The lack of specificity of most of these contrasting agents and the inability to visualize more than one stain where specificity is possible has limited the value of conventional electron microscopy in the study of the nucleus. ESI has significant advantages over conventional TEM, particularly for the study of structures of the cell nucleus. It is possible to exploit the phosphorus-rich nature of DNA- and RNA-containing macromolecular complexes to distinguish nucleoprotein complexes from protein complexes and to resolve different nucleoprotein complexes based on their density of nucleic acids. The remaining biological material can be imaged based on its abundance of nitrogen. Mapping just these two elements and analysis of their distribution and relative abundance within different anatomical structures provides us with a lot of information about the nucleus. For example, it is easy to identify chromatin and the ribosomes in the map representing phosphorus abundance. The interchromatin space, nuclear pore complexes, and nuclear bodies, on the other hand, can be easily detected in the nitrogen map image.
Mini singlet oxygen generator system (miniSOG)
While ESI represents a powerful technique to study in situ chromatin structure because it takes advantage of the characteristic ratios in elemental composition between phosphorus and nitrogen, the elemental composition cannot normally be used to discriminate between different populations of protein complexes. Antibodies labeled with small gold particles in the nanometer range have been widely used to map the location of individual molecules. Since the gold particle is usually attached to a secondary antibody, it will appear within a circumference of approximately 20 nm around the epitope detected by the primary antibody. In post-embedding samples, antibodies can only detect the epitopes that are exposed at the surface of the sections. While it is possible to prove the presence of an antigen and relate it to a certain anatomical structure of the cell, the information that is obtained is incomplete, since most of the epitopes are obscured by resin. Pre-embedding techniques, which use similar protocols to those used for fluorescence microscopy, allow access to antigens throughout the entire depth of the sample but the permeabilization steps that are required to allow the antibody to penetrate the cell typically require removal of lipid membranes and remove components that cannot be fixed by aldehydes. Moreover, the preferred aldehyde fixative for ultrastructure preservation, glutaraldehyde, commonly destroys epitopes and, consequently, paraformaldehyde is typically required. This is less effective at protein-protein crosslinking. Another disadvantage of antibodies that are labeled with a gold nanoparticle is that gold is a very electron-dense material that creates a strong contrast that can obscure interesting structural details of the sample, which has weaker contrast.
The emergence of the green fluorescent protein (GFP) as an expressed protein tag has transformed the use of fluorescence microscopy to answer questions in cell biology. Tagging a protein with a small fluorescent domain allows mapping of its distribution in vivo without the need for permeabilization procedures that can alter the structure. In order to translate the elegant principle of using protein tags to map protein distributions in a cell to electron microscopy, a system was needed that is able to produce a signal close to the structure of interest and generate contrast in the TEM. Polymerized diaminobenzidine is a stain that is often used in histology to detect antibody binding. This stain is usually deposited using horseradish peroxidase (HRP) conjugated to a secondary antibody. Although the reaction produces a reliable result, HRP is not catalytically active in the cytosol of cells9. The reaction products of HRP can also diffuse away from the site of generation so that their resolution is worse than the nanogold method10. In order to bypass these problems, the FlAsH/ReAsH system was developed11. It consists of recombinant fusion proteins that possess a tetracysteine motif. This motif allows binding of a biarsenic fluorophore. When excited, the bound FlAsH or ReAsH is able to generate the highly reactive singlet oxygen and thus photoconvert diaminobenzidine (DAB) into a polymer that precipitates immediately at the site of the tagged proteins. The DAB polymer can be stained with osmium tetroxide, which is electron dense and thus can be used to map the distribution of the recombinant fusion protein in the TEM.
In 2011, Shu et al.10 presented the miniSOG system, which consists of a small 106 amino acids fusion tag derived from a flavoprotein of Arabidopsis that is fluorescent and able to create many singlet oxygen radicals when excited with 448 nm blue light. Those singlet oxygen radicals can be used to photo-oxidize diaminobenzidine to form polymers on and near the surface of the tagged protein, which is considerably closer than immunogold labeled antibodies11. While the FlAsH/ReAsH system requires bringing the fluorochrome and the diaminobenzidine into the cells before doing the photoconversion, the miniSOG system only requires diaminobenzidine and light and in addition is about twice as effective at polymerizing diaminobenzidine. Here, miniSOG is employed in combination with ESI in order to map the ultrastructure of DNA repair foci.
DNA repair foci (DRF)
Unrepaired DNA double strand breaks pose a serious threat to the cell since they can lead to translocations and loss of genetic information. In turn, this can lead to senescence, cancer, and cell death. Many proteins that are involved in DNA double strand break repair accumulate in foci that assemble around a DNA double strand break12-14. Although their function is not known, they represent the site in the nucleus that contains the DNA double strand break and is the site of DNA double strand break repair.
DNA repair foci (DRF) have been characterized by fluorescence microscopy and they serve as biomarkers for DNA damage12,15. They are massive relative to the size of the double-strand break and were considered relatively homogeneous until recent super-resolution microscopy studies revealed some evidence of sub-compartmentalization of molecules within each focus16. In order to understand how these sites are organized, it is necessary to visualize all of the underlying biological structures relative to each other. This cannot be achieved by fluorescence microscopy but is possible through electron microscopy17,18. Here, a method is described that combines electron spectroscopic imaging with the miniSOG method to illustrate the potential of this combined approach to explore the ultrastructure of DNA double-strand break repair.

<table><tbody><tr><td>35 mm glass bottom dishes</td><td>MatTek</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Gridded 35 mm glass bottom dishes</td><td>MatTek</td><td>P35G-2-14-CGRD&amp;nbsp;</td><td>not made for immersion objectives</td></tr><tr><td>LR White: acryl resin</td><td>emsdiasum</td><td>14381</td><td></td></tr><tr><td>LR White accelerator</td><td>emsdiasum</td><td>14385</td><td></td></tr><tr><td>3,3&amp;prime;-Diaminobenzidine tetrahydrochloride 10 mg tablets</td><td>Sigma</td><td>D5905</td><td></td></tr><tr><td>Slim Bar Grids 300 Mesh</td><td>SPI</td><td>1161123</td><td></td></tr><tr><td>Tungsten-Point Lab Pen</td><td>emsdiasum</td><td>41148</td><td></td></tr><tr><td>Osmium Tetroxide</td><td>emsdiasum</td><td>19100</td><td></td></tr><tr><td>Carbon Coater 208 carbon</td><td>Cressington</td><td></td><td></td></tr><tr><td>Ultra microtome Leica EM UC6</td><td>Leica</td><td></td><td></td></tr><tr><td>Photoshop CS5</td><td>Adobe</td><td></td><td>might even work with older versions</td></tr><tr><td>Digital Micrograph V.2.30</td><td>Gatan</td><td></td><td>might even work with older versions</td></tr><tr><td>Hoechst 33342</td><td>Sigma</td><td>H-1399</td><td></td></tr><tr><td>Effectene</td><td>Quiagen</td><td></td><td></td></tr><tr><td>MiniSOG-Constructs</td><td>Tsien-Lab</td><td></td><td>&lt;a href=&quot;mailto:tsienlab@yahoo.com&quot;&gt;tsienlab@yahoo.com&lt;/a&gt;</td></tr><tr><td>MDC1 miniSOG mCherry</td><td></td><td></td><td></td></tr><tr><td>53BP1 miniSOG mCherry</td><td></td><td></td><td></td></tr><tr><td>Rad52 miniSOG mCherry</td><td></td><td></td><td></td></tr><tr><td>cesium 137 radiation source &quot;MARK 1&quot;</td><td>&amp;nbsp;(J.L. Shepherd &amp;amp; Associated)</td><td></td><td></td></tr><tr><td>ImageJ/FiJi</td><td>open source</td><td></td><td>http://fiji.sc/Fiji</td></tr><tr><td>2 ml Eppendorf tubes</td><td>Fisherbrand</td><td>05-408-146</td><td></td></tr><tr><td>Diamond Knife ultra 35&amp;deg;</td><td>Diatome</td><td></td><td></td></tr><tr><td>Trimming Knife ultratrim</td><td>Diatome</td><td></td><td></td></tr><tr><td>Sodium cacodylate trihydrate</td><td>emsdiasum</td><td>12300</td><td>Caution Toxic!</td></tr><tr><td>Glutaraldehyde EM Grade 8%</td><td>emsdiasum</td><td>16020</td><td>Caution Toxic!</td></tr><tr><td>Sodium phosphate dibasic</td><td>emsdiasum</td><td>21180</td><td></td></tr><tr><td>Sodium phosphate monobasic</td><td>emsdiasum</td><td>21190</td><td></td></tr><tr><td>Paraformaldehyde</td><td>emsdiasum</td><td>19202</td><td></td></tr><tr><td>Osmium tetroxide 4% solution</td><td>emsdiasum</td><td>19150</td><td></td></tr><tr><td>Inverted Fluorescence Micoscope Axiovert 200M</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Hydrochloric Acid</td><td>Fisherbrand</td><td>A142-212</td><td></td></tr><tr><td>Sodium Hydroxide Solution 10M</td><td>Fluka</td><td>72068</td><td></td></tr><tr><td>Oxygen</td><td>Medigas</td><td></td><td></td></tr><tr><td>3-Amino-1,2,4-triazole</td><td>Sigma</td><td>A8056</td><td></td></tr><tr><td>Potassium cyanide</td><td>Sigma</td><td>207810</td><td>Caution Toxic!</td></tr><tr><td>Ethanol</td><td>emsdiasum</td><td>15058</td><td>Caution Toxic!</td></tr><tr><td>Razor blade Single Edge Carbon Steel</td><td>emsdiasum</td><td>71960</td><td>Caution Sharp!</td></tr><tr><td>DMEM</td><td>Sigma</td><td>D 5546</td><td></td></tr><tr><td>FBS</td><td>Life Technologies</td><td>16000-044</td><td></td></tr><tr><td>G418</td><td>Life Technologies</td><td>11811-023</td><td></td></tr><tr><td>DMSO</td><td>Sigma</td><td>D2650</td><td></td></tr><tr><td>Transmission Electron Microscope 200 kV</td><td>JEOL</td><td>2100</td><td></td></tr><tr><td>GIF Tridiem 863 Energy filter</td><td>Gatan</td><td></td><td></td></tr></tbody></table>

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.

ESI
Comparing the ESI images of the nucleus (Figure 1) with the conventional TEM images (e.g., Figure 7-1) reveals a dramatic increase in anatomical structures that can be easily distinguished. The chromatin ridges appear in yellow and it is quite easy to identify the chromocenters in mouse cells. Nucleoli can easily be distinguished from&#160;chromatin by their round structure and different color, since the pre-assembled ribosomes al...

Watch this Scientific Journal Video about Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging ESI at JoVE.com

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.

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 ...

visualization-minisog-tagged-dna-repair-proteins-combination-with

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

10.0K Views.  University of Alberta. 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.

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

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of programmed DNA breaks in lymphocytes. The Ataxia-Telangiectasia Mutated kinase (ATM), ATM- and Rad3-Related kinase (ATR) and the catalytic subunit of DNA-dependent Protein Kinase (DNA-PKcs) are among the first that are activated upon induction of DNA damage, and are central regulators of a network that controls DNA repair, apoptosis and cell survival. As part of a tumor-suppressive pathway, ATM and ATR activate p53 through phosphorylation, thereby regulating the transcriptional activity of p53. DNA damage also results in the formation of so-called ionizing radiation-induced foci (IRIF) that represent complexes of DNA damage sensor and repair proteins that accumulate at the sites of DNA damage, which are visualized by fluorescence microscopy. Co-localization of proteins in IRIFs, however, does not necessarily imply direct protein-protein interactions, as the resolution of fluorescence microscopy is limited. 
In situ Proximity Ligation Assay (PLA) is a novel technique that allows the direct visualization of protein-protein interactions in cells and tissues with unprecedented specificity and sensitivity. This technique is based on the spatial proximity of specific antibodies binding to the proteins of interest. When the interrogated proteins are within ~40 nm an amplification reaction is triggered by oligonucleotides that are conjugated to the antibodies, and the amplification product is visualized by fluorescent labeling, yielding a signal that corresponds to the subcellular location of the interacting proteins. Using the established functional interaction between ATM and p53 as an example, it is demonstrated here how PLA can be used in suspension cell cultures to study the direct interactions between proteins that are integral parts of the DNA damage response.

Here, it is demonstrated how the in situ Proximity Ligation Assay (PLA) can be used to detect and visualize the direct protein-protein interactions between ATM and p53 in suspension cell cultures exposed to genotoxic stress.

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of ...

Biochemistry

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 ...

Genetics

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 ...

Cancer Research

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome ...

Immunology and Infection

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

Immunofluorescence is one of the most widely used techniques to visualize target antigens with high sensitivity and specificity, allowing for the accurate identification and localization of proteins, glycans, and small molecules. While this technique is well-established in two-dimensional (2D) cell culture, less is known about its use in three-dimensional (3D) cell models. Ovarian cancer organoids are 3D tumor models that recapitulate tumor cell clonal heterogeneity, the tumor microenvironment, and cell-cell and cell-matrix interactions. Thus, they are superior to cell lines for the evaluation of drug sensitivity and functional biomarkers. Therefore, the ability to utilize immunofluorescence on primary ovarian cancer organoids is extremely beneficial in understanding the biology of this cancer. The current study describes the technique of immunofluorescence to detect DNA damage repair proteins in high-grade serous patient-derived ovarian cancer organoids (PDOs). After exposing the PDOs to ionizing radiation, immunofluorescence is performed on intact organoids to evaluate nuclear proteins as foci. Images are collected using z-stack imaging on confocal microscopy and analyzed using automated foci counting software. The described methods allow for the analysis of temporal and special recruitment of DNA damage repair proteins and colocalization of these proteins with cell-cycle markers.

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

The present protocol describes methods for evaluating DNA damage repair proteins in patient-derived ovarian cancer organoids. Included here are comprehensive plating and staining methods, as well as detailed, objective quantification procedures.

Immunofluorescence is one of the most widely used techniques to visualize target antigens with high sensitivity and specificity, allowing for the accurate ...

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 ...

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 ...

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 ...

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

Visualizing Single-Stranded DNA Foci in the G1 Phase of the Cell Cycle

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair necessitates collaboration between numerous proteins, responsible for covering a broad range of tasks from recognizing and signaling the presence of a DNA lesion to physically repairing it. During this process, tracks of single-stranded DNA (ssDNA) are often created, which are eventually filled by DNA polymerases. The nature of these ssDNA tracks (in terms of both length and number), along with the polymerase recruited to fill these gaps, are repair pathway-specific. The visualization of these ssDNA tracks can help us understand the complicated dynamics of DNA repair mechanisms.
This protocol provides a detailed method for the preparation of G1 synchronized cells to measure ssDNA foci formation upon genotoxic stress. Using an easy-to-utilize immunofluorescence approach, we visualize ssDNA by staining for RPA2, a component of the heterotrimeric replication protein A complex (RPA). RPA2 binds to and stabilizes ssDNA intermediates that arise upon genotoxic stress or replication to control DNA repair and DNA damage checkpoint activation. 5-Ethynyl-2'-deoxyuridine (EdU) staining is used to visualize DNA replication to exclude any S phase cells. This protocol provides an alternative approach to the conventional, non-denaturing 5-bromo-2'-deoxyuridine (BrdU)-based assays and is better suited for the detection of ssDNA foci outside the S phase.

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

The following protocol presents the detection of single-stranded DNA foci in the G1 phase of the cell cycle utilizing cell cycle synchronization followed by RPA2 immunofluorescent staining.

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair ...

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

Summary

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Chapters in this video

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

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

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

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

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

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Visualizing and Quantifying Endonuclease-Based Site-Specific DNA Damage

Visualizing Single-Stranded DNA Foci in the G1 Phase of the Cell Cycle