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 border="0" cellpadding="0" cellspacing="0">
	<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&#160;</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&#8242;-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 208carbon</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><a href="mailto:tsienlab@yahoo.com">tsienlab@yahoo.com</a></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 &#34;MARK 1&#34;</td>
			<td>&#160;(J.L. Shepherd &#38; 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&#176;</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>lifetechnologies</td>
			<td>16000-044</td>
			<td></td>
		</tr>
		<tr>
			<td>G418</td>
			<td>lifetechnologies</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

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

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System 

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers. Genetic studies in mammalian systems have been limited due to the lack of readily available tools including defined mutant genetic cell lines, regulatory expression systems, and appropriate selectable markers. To circumvent these difficulties, model genetic systems in lower eukaryotes have become an attractive choice for the study of functionally conserved DNA repair proteins and pathways. We have developed a model yeast system to study the poorly defined genetic functions of the Werner syndrome helicase-nuclease (WRN) in nucleic acid metabolism. Cellular phenotypes associated with defined genetic mutant backgrounds can be investigated to clarify the cellular and molecular functions of WRN through its catalytic activities and protein interactions. The human WRN gene and associated variants, cloned into DNA plasmids for expression in yeast, can be placed under the control of a regulatory plasmid element. The expression construct can then be transformed into the appropriate yeast mutant background, and genetic function assayed by a variety of methodologies. Using this approach, we determined that WRN, like its related RecQ family members BLM and Sgs1, operates in a Top3-dependent pathway that is likely to be important for genomic stability. This is described in our recent publication [1] at <a href="http://www.impactaging.com">www.impactaging.com</a>. Detailed methods of specific assays for genetic complementation studies in yeast are provided in this paper.

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Genetic studies in yeast can be employed to investigate the molecular and cellular functions of human genes in cellular DNA metabolism. Methods are described for the genetic characterization of the human WRN gene product defective in the premature aging disorder Werner syndrome in functionally conserved pathways using yeast as a tractable model system.

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers.  Genetic studies in mammalian ...

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

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

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

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

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

Visualization of MG53-mediated Cell Membrane Repair Using in vivo and in vitro Systems

Repair of acute injury to the cell membrane is an elemental process of normal cellular physiology, and defective membrane repair has been linked to many degenerative human diseases. The recent discovery of MG53 as a key component of the membrane resealing machinery allows for a better molecular understanding of the basic biology of tissue repair, as well as for potential translational applications in regenerative medicine. Here we detail the experimental protocols for exploring the in vivo function of MG53 in repair of muscle injury using treadmill exercise protocols on mouse models, for testing the ex vivo membrane repair capacity by measuring dye entry into isolated muscle fibers, and for monitoring the dynamic process of MG53-mediated vesicle trafficking and cell membrane repair in cultured cells using live cell confocal microscopy.

Visualization of MG53-mediated Cell Membrane Repair Using in vivo and in vitro Systems

Described here are protocols used to visualize the dynamic process of MG53-mediated cell membrane repair in whole animals and at the cellular level. These methods can be applied to investigate the cell biology of plasma membrane resealing and regenerative medicine.

Repair of acute injury to the cell membrane is an elemental process of normal cellular physiology, and defective membrane repair has been linked to many ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

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

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

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

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

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

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 ...

Photodiode-Based Optical Imaging for Recording Network Dynamics with Single-Neuron Resolution in Non-Transgenic Invertebrates

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light represents a revolutionary advance for studies of the neural basis of behavior. However, a downside of this development is its tendency to focus investigators on a very small number of &#8220;designer&#8221; organisms (e.g., C. elegans and Drosophila), potentially negatively impacting the pursuit of comparative studies across many species, which is needed for identifying general principles of network function. The present article illustrates how optical recording with voltage-sensitive dyes in the brains of non-transgenic gastropod species can be used to rapidly (i.e., within the time course of single experiments) reveal features of the functional organization of their neural networks with single-cell resolution. We outline in detail the dissection, staining, and recording methods used by our laboratory to obtain action potential traces from dozens to ~150 neurons during behaviorally relevant motor programs in the CNS of multiple gastropod species, including one new to neuroscience &#8211; the nudibranch Berghia stephanieae. Imaging is performed with absorbance voltage-sensitive dyes and a 464-element photodiode array that samples at 1,600 frames/second, fast enough to capture all action potentials generated by the recorded neurons. Multiple several-minute recordings can be obtained per preparation with little to no signal bleaching or phototoxicity. The raw optical data collected through the methods described can subsequently be analyzed through a variety of illustrated methods. Our optical recording approach can be readily used to probe network activity in a variety of non-transgenic species, making it well suited for comparative studies of how brains generate behavior.

This protocol presents a method for imaging neuronal population activity with single-cell resolution in non-transgenic invertebrate species using absorbance voltage-sensitive dyes and a photodiode array. This approach enables a rapid workflow, wherein imaging and analysis can be pursued over the course of a single day.

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light ...