This work was supported in part by RSF (grant #17-15-01290) and RFBR (grant #19-015-00273). The authors thank Lomonosov Moscow State University development program (PNR 5.13) and Nikon Center of Excellence in correlative imaging at Belozersky Institute of Physico-Chemical Biology for access to imaging instrumentation.

The protocol is optimized for adherent cells and was tested on HeLa, HT1080, and CHO cell lines.
1. Cell labeling and fixation
<ol>
	<li>Plate cells on acid-cleaned coverslips in a 3 cm Petri dish. Grow the cells in the media recommended for the cell line being used to 70% confluency.</li>
	<li>Add EdU (5-ethynyl-2&#39;-deoxyuridine) from 10 mM stock to 10 &#181;M final concentration and place the cells in the incubator for 10 min or longer (depending on the experiment objective). For shorter pulses (up to 2 min), prepare a Petri dish with pre-warmed fresh culture media supplemented with 10 &#181;M EdU, and transfer coverslips in it, rather than adding EdU to the cells directly. 
	NOTE: All subsequent steps are carried out at room temperature if not indicated otherwise.</li>
	<li>Fix the labeled cells with 2.5% glutaraldehyde, freshly made by diluting 25% stock in 100 mM cacodylate buffer, for 1 h. 
	NOTE: The subsequent EdU detection steps are sensitive to fixation timing. Longer fixation times may adversely affect EdU labeling, causing lower signal-to-noise ratio.</li>
	<li>Remove glutaraldehyde by washing the samples with PBS supplemented with 5 mM MgCl2 (PBS* thereafter). Wash three times with a 10 min incubation for each wash.</li>
	<li>Permeabilize plasma membranes with 1% Triton X-100 in PBS* (PBS*T thereafter). Wash twice with a 5 min incubation for each wash.</li>
	<li>Extensively wash the sample with PBS*. Perform five changes and incubate for 5 min in each change.</li>
	<li>Quench the residual-free aldehyde groups with 20 mM glycine in PBS*, two times for 10 min each.</li>
	<li>Block the samples in 1% BSA in PBS* for 30 min.</li>
</ol>
2. Click-reaction
NOTE: This procedure is modified from a previously published protocol19.
<ol>
	<li>Immediately before use, prepare Click-reaction mix for EdU detection: in a microcentrifuge tube, mix 430 &#181;L of 100 mM Tris-HCl (pH = 8.5), 20 &#181;L of 100 mM CuSO4, 1.2 &#181;L of biotin-azide (10 mM in DMSO), and 50 &#181;L of 0.5 M ascorbic acid. For quality control of the replicative labeling and Click-procedure at the level of fluorescent microscopy, biotin-azide can be replaced by AlexaFluor 488-azide. 
	NOTE: The order of addition of components in the above reaction is important.</li>
	<li>Perform click-reaction in a moist chamber to minimize the evaporation and concentration shifts in the reaction cocktail. Prepare the moist chamber by placing a wet sheet of filter paper on the bottom of the Petri dish and covering it with a paraffin film. Place the coverslips on the film&#39;s surface with the cells facing up and layer 50-100 &#181;L of the reaction cocktail onto the coverslip. The reaction takes 30 min at room temperature.</li>
	<li>Stop the reaction by washing the sample in 0.1% Triton X-100 in PBS* (PBS*T), five times for 5 min each.</li>
	<li>Block in 1% BSA in PBS*T for 30 min at room temperature.</li>
	<li>Prepare streptavidin-Nanogold solution with PBS*T containing 1% BSA. Incubate the sample with streptavidin, conjugated with 1.4 nm Nanogold particles (Nanoprobes) in 1% BSA + PBS*T overnight at +4 &#176;C in the newly prepared moist chamber.</li>
	<li>Stop the reaction and wash the sample in PBS*T, five times for 10 min each.</li>
	<li>Stabilize biotin streptavidin complex by postfixing in 1% glutaraldehyde in PBS* for 30 min.</li>
	<li>Remove glutaraldehyde by intense washing with deionized water and wash the sample in PBS*, five times for 20 min each.</li>
	<li>Quench free aldehyde groups with freshly prepared 1 mg/mL NaBH4 in water, two times for 10 min each. During incubation, bubbles of H2 are formed which can lift the coverslips. Carefully push them down with the tweezers.</li>
	<li>Wash with deionized water five times for 5 min each. 
	NOTE: For quality control at the labeling step, it is advisable to perform control labeling with fluorescently labeled streptavidin (Figure 2). This would provide an estimate of labeling intensity and background level before switching to tedious and artifact-prone subsequent steps.</li>
</ol>
3. Ag-amplification
NOTE: This procedure is modified from Gilerovitch et al., 1995 (see 20,21).
<ol>
	<li>Resuspend 50 g of acacia powder in 100 mL of deionized water. Dissolve for 3 days, degas, and filter through four layers of cheesecloth. Store frozen in 5 mL aliquots in 50 mL tubes. Thaw immediately before use.</li>
	<li>Add 2 mL of 1 M MES (pH = 6.1) to 5 mL of thawed aliquot of acacia powder solution to make 7 mL of 0.28 M MES (pH = 6.1) in acacia powder solution. Mix by slowly rocking the tube for 30 min. Wrap the tube with foil to protect from light.</li>
	<li>Simultaneously with step 3.2., wash coverslips with washing buffer (50 mM MES, pH = 5.8, 200 mM sucrose), three times for 10 min each.</li>
	<li>Prepare the fresh solution of N-propyl gallate (NPG). In a 15 mL tube, dissolve 10 mg of NPG in 250 &#181;L of 96% ethanol, and adjust to 5 mL with deionized water.</li>
	<li>Put 36 mg of silver lactate in a foil-wrapped 15 mL tube.</li>
	<li>After step 3.2. is completed, add 1.5 mL of NPG solution to acacia powder mix and rock for another 3 min. 
	NOTE: All the subsequent steps are carried out in a darkroom. Use non-actinic safelight (yellow or red).</li>
	<li>Equilibrate the sample with washing buffer and proceed to the dark room. Simultaneously, add 5 mL of deionized water to silver lactate and dissolve by vigorous shaking for 1-2 min.</li>
	<li>Add 1.5 mL of silver lactate solution to acacia powder mix, rock slowly for 1 min. Avoid bubble formation, as oxygen in the mix will slow down the reaction.</li>
	<li>Drain the solution from the dish with coverslips and pour 3 mL of silver lactate-acacia powder mix on the cells. Rock the dish several times and incubate for 2-5 min. Incubation time depends on acacia powder batch and the temperature. This should be defined experimentally. 
	NOTE: A longer incubation time may result in unspecific silver binding.</li>
	<li>To stop the reaction, drain the reaction mix and wash the dish extensively with three to five changes of deionized water, followed by three more changes for 5 min each.</li>
	<li>Check the staining under the brightfield microscope. Good staining should be light to dark brown with a very faint yellowish background. 
	&#8203;NOTE: If the staining is not developing, it is possible to repeat immediately with the already made mix (the mix is active for about 15 min if kept in the dark). However, the staining in this case might develop too fast.</li>
</ol>
4. Gold toning
NOTE: In order to protect silver nanoparticles from dissolution by OsO4 oxidation in the subsequent procedures, an impregnation with gold is used at this step (Sawada, Esaki, 1994)22.
<ol>
	<li>Rinse coverslips with deionized water.</li>
	<li>In order to protect silver nanoparticles from dissolution by OsO4 oxidation, incubate the samples in 0.05% tetrachloroauric acid for 2 min in the dark.</li>
	<li>Wash thoroughly with deionized water for 10 min. 
	&#8203;NOTE: At this stage additional contrast is added to the DNA containing material. The color of the sample should change from brown to black.</li>
</ol>
5. ChromEM
NOTE: This protocol was modified from Ou et al., 201716.
<ol>
	<li>Incubate the coverslips and saturate the samples in 10 &#181;M DRAQ5 in PBS for 10 min in the dark.</li>
	<li>Prepare 0.2% diaminobenzidine tetrahydrochloride (DAB) solution in 50 mM Tris or PBS: 
	dissolve DAB in 90% of final volume by stirring vigorously in the dark, then adjust the pH to 7.0-7.4 with NaOH. Adjust the volume to 100% and filter through a 0.22 &#181;M filter, store wrapped in foil.</li>
	<li>Add DAB solution to cells (2 mL per 3 cm Petri dish) and incubate for 1 min.</li>
	<li>Move the coverslip into the glass bottom Petri-dish and place it to the stage of an inverted microscope. Irradiate the sample (several fields of view) on an inverted microscope with metal-halide light source and Cy5 filter set (640 nm, 1 W/cm2 at focal plane) through Plan Apo &#955; 40&#1093; (NA = 0.95) lens for 10 min. 
	&#8203;NOTE: An indication of successful DAB photoconversion is complete DRAQ5 photobleaching and dark DAB precipitate formation in irradiated nuclei. However, the latter is not always apparent over intense EdU-silver-gold signal (especially when extended EdU pulses are utilized).</li>
	<li>Wash with deionized water three times for 5 min each.</li>
</ol>
6. Dehydration and epoxy resin embedding
<ol>
	<li>In a microcentrifuge tube, prepare partially reduced osmium tetraoxide solution by mixing 2% aqueous solution of K4Fe(CN)6 with an equal amount of 2% aqueous solution of OsO4, to get 1% final concentration of both components. Incubate coverslips in reduced OsO4 for 1 h and wash in deionized water three times for 5 min each. 
	NOTE: At this stage the reduced osmium compound reacts with DAB depositions and increases the contrast of DNA. 
	CAUTION: Osmium is toxic, work under a fume hood and handle waste according to local safety regulations.</li>
	<li>Dehydrate the samples in a series of graded ethanol solutions in increasing percentages: 50% EtOH - 3 x 10 min; 70% EtOH - 3 x 10 min, 80% EtOH - 3 x 10 min, 96% EtOH - 3 x 10 min, 100% EtOH - 3 x 10 min.</li>
	<li>For infiltration and embedding use the resin formula with the component volumetric ratio as follows: epoxy resin monomer:DDSA:MNA:DMP-30 = 9:6:4:0.23 (w/w). This formula is miscible with ethanol.
	<ol>
		<li>Incubate the coverslip in the ethanol-resin mix (3 parts 100% EtOH:1 part epoxy resin) for 30 min.</li>
		<li>Incubate the coverslip in the ethanol-resin mix (1 part 100% EtOH:1 part epoxy resin) for 2 h.</li>
		<li>Incubate the coverslip in the ethanol-resin mix (1 part 100% EtOH:3 parts epoxy resin) overnight.</li>
		<li>Replace ethanol-resin mix with freshly prepared pure resin. Incubate in pure resin mix for 8 h. Open the dish to let remnants of ethanol to evaporate.</li>
		<li>Transfer coverslips into a new dish with pure resin and incubate for 2 h at 37-42 &#176;C.</li>
		<li>Fill a silicon mold with freshly prepared resin. Place the coverslips with the cells facing down on appropriate silicon mold filled with epoxy resin. Cure the resin for 24 h at 37 &#176;C, then at 60 &#176;C for at least 2 days. 
		CAUTION: Components of epoxy resin mix are potential carcinogens. Wear gloves and work under the fume hood.</li>
	</ol>
	</li>
	<li>Remove the resin slab from the mold, clean the surface of the coverslip with the scalpel or razor blade. To remove the coverslip, drop the coverslip into liquid nitrogen and then transfer the slab into the boiling water. Repeat if necessary.</li>
	<li>Locate the irradiated area under a stereomicroscope and cut it out from the slab with a hacksaw or any other suitable instrument. Pre-warm the slab at 70 &#176;C on a hot plate, if needed.</li>
	<li>Fasten the cut-out into an ultramicrotome sample holder for final block trimming. Using the razor, prepare the pyramid shaped sample. Mount the holder with pyramid onto the ultramicrotome and install the knife.
	<ol>
		<li>Trim the block so that the irradiated cells bearing EdU label are included. Prepare a semi-thin section (250 nm thickness) suitable for electron tomography. 
		NOTE: Section thickness can be adjusted to suit the experimental requirements.</li>
	</ol>
	</li>
	<li>Detach section from the knife&#39;s edge and place them on the single-slot grids. For electron tomography, 250 nm sections are picked onto 1 mm single-slot grids and after drying these sections can be carbon-coated from both sides. No additional contrasting is necessary. 
	NOTE: Since the sections already contain high contrast Au-Ag particles, there is no need to add fiducial gold particles to the surface of the section.</li>
	<li>Examine the grids in a transmission electron microscope at low magnification (about 600x) to locate the cells with appropriate replication patterns. 
	&#8203;NOTE: This protocol generates labeling density high enough for easy detection of replication foci even at low magnification. It is advisable first to create a map of the section for subsequent navigation and angular projections collection for tomography. Switch to higher magnification and take high-resolution images.</li>
</ol>
7. Electron tomography
<ol>
	<li>Insert the grid into the transmission electron microscope with a high-tilt holder. Load the sample into the electron microscope and locate the region of interest.</li>
	<li>Align the microscope in EFTEM mode in near parallel illumination mode. Tune the energy filter with 20 eV energy-selecting slit placed at zero loss peak.</li>
	<li>Pre-irradiate the tomography acquisition area at a dose rate of 40 e/&#197;2/s for at least 1.5-2 min, which corresponds to the total dose at least of 3000 e/&#197;2.</li>
	<li>Adjust eucentric height with automatic task in SerialEM. Check autofocus task to ensure that it is working correctly at zero tilt angle and -0.8 mkm target defocus value.</li>
	<li>Tilt the sample holder to -60&#176; with the Walk-Up task.</li>
	<li>Set up the tomography acquisition from -60 to +60&#176; with step 2.0&#176;. Exposure should be set dependent on the tilt angle to keep the average intensity on the camera. Limit the image shift in case it causes an energy shift and thus slit misalignment. The tomography data should be saved as .mrc file.</li>
	<li>Import the .mrc file into IMOD software for tomography reconstruction. Remove the outlier pixel values due to X-rays.</li>
	<li>Align the tilt series. Perform initial cross-correlation, then manually mark 10-12 gold particles as fiducials. Track the fiducial model and inspect that every fiducial is tracked correctly through the tilt series. Create sample tomograms (slices) and manually mark the thin section borders to prevent the possible tilt of the reconstructed density inside the volume. The mean residual error for fine alignment was less than 1.2 pix.</li>
	<li>Reconstruct the tomogram with a filtered back projection algorithm and then trim the output to fit the region of interest into the final volume.</li>
</ol>

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The authors have nothing to disclose

The method described here has several advantages over previously published protocols. First, the use of Click-chemistry for labeling replicated DNA eliminates the necessity of DNA denaturation prerequisite for BrdU detection with antibodies, thus better preserving chromatin ultrastructure. 
Second, utilization of biotin as a secondary ligand that is generated after glutaraldehyde fixation and proper quenching of unbound aldehyde groups minimizes chemical modification of the target, thus improv...

DNA replication is a basic biological process required for faithful copying and transmission of the genetic information during cell division. In higher eukaryotes, DNA replication is subjected to tight spatio-temporal regulation, which is manifested in sequential activation of replication origins1. Neighboring replication origins firing synchronously form clusters of replicons2. At the level of optical microscopy, sites of ongoing DNA replication are detected as replication foci of various number and size. Replication foci display specific patterns of spatial distribution within the cell nucleus depending on the replication timing of the labeled DNA3,4, which, in turn, is tightly correlated with its gene activity. Thanks to well-defined sequence of DNA replication, strictly ordered in space and time, replicative labeling is a powerful method of precise DNA labeling not only for the study of replication process per se, but also to discriminate a specific DNA sub-fraction with defined transcription activity and compaction level. Visualization of replicating chromatin is usually performed through the detection of major protein components of DNA replication machinery (either by immunostaining or by expression of fluorescent protein tags5,6) or by the incorporation of modified DNA synthesis precursors7,8,9,10. Of these, only methods based on the incorporation of modified nucleotides into newly replicated DNA allow for the capture of conformational changes in chromatin during replication, and trace the behavior of replicative domains after their replication is completed.
In higher eukaryotes, DNA packaging into chromatin adds another level of complexity to the regulation of basic genetic functions (transcription, replication, reparation, etc.). Chromatin folding affects accessibility of DNA to regulatory trans-factors and DNA conformational changes (double helix unwinding) required for the template synthesis. Therefore, it is generally accepted that DNA-dependent synthetic processes in the cell nucleus require a structural transition of chromatin from its condensed, repressive state to a more accessible, open conformation. Cytologically, these two chromatin states are defined as heterochromatin and euchromatin. However, there is still no consensus concerning the mode of DNA folding in the nucleus. The hypotheses range from a &#34;polymer melt&#34; model11, where nucleosomal fiber behaves as a random polymer for which the packing density is controlled by phase separation mechanisms, to hierarchical folding models postulating sequential formation of chromatin fiber-like structures of increasing thickness12,13. Hierarchical folding models recently gained support from molecular approaches based on the analysis of in situ DNA-DNA contacts (chromosome conformation capture, 3C), demonstrating the existence of the hierarchy of chromatin structural domains14. It is important to note that replication units correlate very well to these chromatin domains15. The major criticism of these models is based on potential artificial chromatin aggregation caused by sample preparation procedures, such as permeabilization of cell membranes and removal of non-chromatin components, in order to improve chromatin contrast for ultrastructural studies while improving chromatin accessibility for various probes (e.g., antibodies). Recent technical advances in selective DNA staining for electron microscopy by DNA-binding fluorophore-mediated photo-oxidation of diaminobenzidine (ChromEMT6) has allowed for the elimination of this obstacle. However, the same considerations hold true for electron microscopy visualization of replicating DNA17,18. Here we describe a technique that allows for the simultaneous high-resolution ultrastructural mapping of newly synthesized DNA and total chromatin in intact aldehyde-crosslinked cells. The technique combines detection of EdU-labeled DNA by Click-chemistry with biotinylated probes and streptavidin-Nanogold, and ChromEMT.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5-ethynyl-2`-deoxyuridine (EdU)</td>
			<td>Thermo Fisher</td>
			<td>A10044</td>
			<td></td>
		</tr>
		<tr>
			<td>2-(4-Morpholino)ethane Sulfonic Acid (MES)</td>
			<td>Fisher Scientific</td>
			<td>BP300-100</td>
			<td></td>
		</tr>
		<tr>
			<td>AlexaFluor 555-azide</td>
			<td>Termo Fisher</td>
			<td>A20012</td>
			<td></td>
		</tr>
		<tr>
			<td>biotin-azide</td>
			<td>Lumiprobe</td>
			<td>C3730</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumine</td>
			<td>Boval</td>
			<td>LY-0080</td>
			<td></td>
		</tr>
		<tr>
			<td>DDSA</td>
			<td>SPI-CHEM</td>
			<td>26544-38-7</td>
			<td></td>
		</tr>
		<tr>
			<td>DMP-30</td>
			<td>SPI-CHEM</td>
			<td>90-72-2</td>
			<td></td>
		</tr>
		<tr>
			<td>DRAQ5</td>
			<td>Thermo Scientific</td>
			<td>62251</td>
			<td></td>
		</tr>
		<tr>
			<td>Epoxy resin monomer</td>
			<td>SPI-CHEM</td>
			<td>90529-77-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde (25%, EM Grade)</td>
			<td>TED PELLA, INC</td>
			<td>18426</td>
			<td></td>
		</tr>
		<tr>
			<td>Gum arabic</td>
			<td>ACROS Organics</td>
			<td>258850010</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>Panreac</td>
			<td>141396.1209</td>
			<td></td>
		</tr>
		<tr>
			<td>NaBH4</td>
			<td>SIGMA-ALDRICH</td>
			<td>213462</td>
			<td></td>
		</tr>
		<tr>
			<td>NMA</td>
			<td>SPI-CHEM</td>
			<td>25134-21-8</td>
			<td></td>
		</tr>
		<tr>
			<td>N-propyl gallate</td>
			<td>SIGMA-ALDRICH</td>
			<td>P3130</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>MP Biomedicals</td>
			<td>2810305</td>
			<td></td>
		</tr>
		<tr>
			<td>Silver lactate</td>
			<td>ALDRICH</td>
			<td>359750-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin-AlexaFluor 488 conjugate</td>
			<td>Termo Fisher</td>
			<td>S11223</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin-Nanogold conjugate</td>
			<td>Nanoprobes</td>
			<td>2016</td>
			<td></td>
		</tr>
		<tr>
			<td>tetrachloroauric acid</td>
			<td>SIGMA-ALDRICH</td>
			<td>HT1004</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris(hydroxymethyl)aminomethane (Tris)</td>
			<td>CHEM-IMPEX INT&#39;L</td>
			<td>298</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fluka Chemica</td>
			<td>93420</td>
			<td></td>
		</tr>
		<tr>
			<td>Instruments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Carbon Coater</td>
			<td>Hitachi</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Copper single slot grids</td>
			<td>Ted Pella</td>
			<td>1GC10H</td>
			<td></td>
		</tr>
		<tr>
			<td>Cy5 fluorescence filter set (Ex620/60 DM660 Em700/75)</td>
			<td>Nikon</td>
			<td>Cy5 HQ</td>
			<td>Alternatives: Zeiss, Leica, Olympus</td>
		</tr>
		<tr>
			<td>Diamond knife Ultra Wet 45o</td>
			<td>Diatome</td>
			<td>DU</td>
			<td>Alternatives: Ted Pella</td>
		</tr>
		<tr>
			<td>Fluorescent microscope</td>
			<td>Nikon</td>
			<td>Ti-E</td>
			<td>Alternatives: Zeiss, Leica, Olympus</td>
		</tr>
		<tr>
			<td>High-tilt sample holder</td>
			<td>Jeol</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rotator</td>
			<td>Biosan</td>
			<td>Multi Bio RS-24</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission electron microscope operating at 200 kV in EFTEM mode, with high-tilt goniometer</td>
			<td>Jeol</td>
			<td>JEM-2100</td>
			<td>Alternatives: FEI, Hitachi</td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>Ted Pella</td>
			<td>523</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome</td>
			<td>Leica</td>
			<td>UltraCut-E</td>
			<td>Alternatives: RMC</td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Image acquisition</td>
			<td>Open Source</td>
			<td>SerialEM (https://bio3d.colorado.edu/SerialEM/)</td>
			<td></td>
		</tr>
		<tr>
			<td>Image processing</td>
			<td>Open Source</td>
			<td>IMOD (https://bio3d.colorado.edu/imod/)</td>
			<td></td>
		</tr>
	</tbody>
</table>

imaging replicative domains in ultrastructurally preserved chromatin by electron tomography

Principles of DNA folding in the cell nucleus and its dynamic transformations that occur during the fulfillment of basic genetic functions (transcription, replication, segregation, etc.) remain poorly understood, partially due to the lack of experimental approaches to high-resolution visualization of specific chromatin loci in structurally preserved nuclei. Here we present a protocol for the visualization of replicative domains in monolayer cell culture in situ,&#160;by combining EdU labeling of newly synthesized DNA with subsequent label detection with Ag-amplification of Nanogold particles and ChromEM staining of chromatin. This protocol allows for the high-contrast, high-efficiency pre-embedding labeling, compatible with traditional glutaraldehyde fixation that provides the best structural preservation of chromatin for room-temperature sample processing. Another advantage of pre-embedding labeling is the possibility to pre-select cells of interest for sectioning. This is especially important for the analysis of heterogeneous cell populations, as well as compatibility with electron tomography approaches to high-resolution 3D analysis of chromatin organization at sites of replication, and the analysis of post-replicative chromatin rearrangement and sister chromatid segregation in the interphase.

Replication foci in mammalian cell nuclei display distinct patterns of distribution within the nucleus depending on S-phase progression. These patterns correlate with transcriptional activity of the loci being replicated. Since the method presented here utilizes a rather strongfixation procedure, it is fairly straightforward to use replicative pulse labeling for specific detection of chromatin loci in various transcriptional states, even under conditions offering best structural preservation of chromatin obtained by room...

Watch this Scientific Journal Video about Imaging Replicative Domains in Ultrastructurally Preserved Chromatin by Electron Tomography at JoVE.com

Imaging Replicative Domains in Ultrastructurally Preserved Chromatin by Electron Tomography

This protocol presents a technique for high-resolution mapping of replication sites in structurally preserved chromatin in situ that employs a combination of pre-embedding EdU-streptavidin-Nanogold labeling and ChromEMT.

Principles of DNA folding in the cell nucleus and its dynamic transformations that occur during the fulfillment of basic genetic functions (transcription, ...

imaging-replicative-domains-ultrastructurally-preserved-chromatin

Research

JoVE Journal

Biology

3.6K Views.  Lomonosov Moscow State University. This protocol presents a technique for high-resolution mapping of replication sites in structurally preserved chromatin in situ that employs a combination of pre-embedding EdU-streptavidin-Nanogold labeling and ChromEMT.

Video: Imaging Replicative Domains in Ultrastructurally Preserved Chromatin by Electron Tomography

Chromatin Isolation by RNA Purification (ChIRP)

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental gene expression 1,2,3,4,5,6,7. The recent discovery of thousands of lncRNAs in association with specific chromatin modification complexes, such as Polycomb Repressive Complex 2 (PRC2) that mediates histone H3 lysine 27 trimethylation (H3K27me3), suggests broad roles for numerous lncRNAs in managing chromatin states in a gene-specific fashion 8,9. While some lncRNAs are thought to work in cis on neighboring genes, other lncRNAs work in trans to regulate distantly located genes. For instance, Drosophila lncRNAs roX1 and roX2 bind numerous regions on the X chromosome of male cells, and are critical for dosage compensation 10,11. However, the exact locations of their binding sites are not known at high resolution. Similarly, human lncRNA HOTAIR can affect PRC2 occupancy on hundreds of genes genome-wide 3,12,13, but how specificity is achieved is unclear. LncRNAs can also serve as modular scaffolds to recruit the assembly of multiple protein complexes. The classic trans-acting RNA scaffold is the TERC RNA that serves as the template and scaffold for the telomerase complex 14; HOTAIR can also serve as a scaffold for PRC2 and a H3K4 demethylase complex 13.
Prior studies mapping RNA occupancy at chromatin have revealed substantial insights 15,16, but only at a single gene locus at a time. The occupancy sites of most lncRNAs are not known, and the roles of lncRNAs in chromatin regulation have been mostly inferred from the indirect effects of lncRNA perturbation. Just as chromatin immunoprecipitation followed by microarray or deep sequencing (ChIP-chip or ChIP-seq, respectively) has greatly improved our understanding of protein-DNA interactions on a genomic scale, here we illustrate a recently published strategy to map long RNA occupancy genome-wide at high resolution 17. This method, Chromatin Isolation by RNA Purification (ChIRP) (Figure 1), is based on affinity capture of target lncRNA:chromatin complex by tiling antisense-oligos, which then generates a map of genomic binding sites at a resolution of several hundred bases with high sensitivity and low background. ChIRP is applicable to many lncRNAs because the design of affinity-probes is straightforward given the RNA sequence and requires no knowledge of the RNA's structure or functional domains.

Chromatin Isolation by RNA Purification ChIRP

ChIRP is a novel and rapid technique to map genomic binding sites of long noncoding RNAs (lncRNAs). The method takes advantage of the specificity of anti-sense tiling oligonucleotides to allow the enumeration of lncRNA-bound genomic sites.

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental ...

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 ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Isolation of Specific Genomic Regions and Identification of Associated Molecules by enChIP

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. To enable the non-biased identification of molecules interacting with a specific genomic region of interest, we recently developed the engineered DNA-binding molecule-mediated chromatin immunoprecipitation (enChIP) technique. Here, we describe how to use enChIP to isolate specific genomic regions and identify the associated proteins and RNAs. First, a genomic region of interest is tagged with a transcription activator-like (TAL) protein or a clustered regularly interspaced short palindromic repeats (CRISPR) complex consisting of a catalytically inactive form of Cas9 and a guide RNA. Subsequently, the chromatin is crosslinked and fragmented by sonication. The tagged locus is then immunoprecipitated and the crosslinking is reversed. Finally, the proteins or RNAs that are associated with the isolated chromatin are subjected to mass spectrometric or RNA sequencing analyses, respectively. This approach allows the successful identification of proteins and RNAs associated with a genomic region of interest.

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. This protocol describes procedures to perform engineered DNA-binding molecule-mediated chromatin imunoprecipitation (enChIP) for identification of proteins and RNAs associated with a specific genomic region.

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the ...

Preparation and Observation of Thick Biological Samples by Scanning Transmission Electron Tomography

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It also describes an imaging method for studying the 3D structure of thick biological specimens by scanning transmission electron tomography. The sample preparation protocol is based on conventional methods in which the sample is fixed using chemical agents, treated with a heavy atom salt contrasting agent, dehydrated in a series of ethanol baths, and embedded in resin. The specific imaging conditions for observing thick samples by scanning transmission electron microscopy are then described. Sections of the sample are observed using a through-focus method involving the collection of several images at various focal planes. This enables the recovery of in-focus information at various heights throughout the sample. This particular collection pattern is performed at each tilt angle during tomography data collection. A single image is then generated, merging the in-focus information from all the different focal planes. A classic tilt-series dataset is then generated. The advantage of the method is that the tilt-series alignment and reconstruction can be performed using standard tools. The collection of through-focal images allows the reconstruction of a 3D volume that contains all of the structural details of the sample in focus.

This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique cristae-rich double membrane ultrastructure of this organelle is elegantly arranged to carry out multiple functions by partitioning biomolecules. Mitochondrial ultrastructure is intimately linked with various functions; however, the fine details of these structure-function relationships are only beginning to be described. Here, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle. Serial-section electron tomography may be adapted to study any cellular structure in three-dimensions.

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

In this protocol, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle.

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique ...

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 ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...