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><tbody><tr><td>&lt;strong&gt;Reagent&lt;/strong&gt;</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>NaBH&lt;sub&gt;4&lt;/sub&gt;</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&#039;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>&lt;strong&gt;Instruments&lt;/strong&gt;</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>&lt;strong&gt;1GC10H&lt;/strong&gt;</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 45&lt;sup&gt;o&lt;/sup&gt;</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>&lt;strong&gt;Software&lt;/strong&gt;</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

3.7K Views.  Lomonosov Moscow State University. We presenteren een protocol voor de elektronenmicroscopische visualisatie van de replicerende domeinen in de cel door de EdU-etikettering van het nieuw gesynthetiseerde DNA en labeldetectie met zilververbetering van Nanogold-deeltjes en ChromEM-kleuring van het chromatine. Dit protocol geeft de pre-embedding-etikettering met hoog contrast met traditionele glutaaraldehydefixatie die de beste structurele conservering van het chromatine geeft voor de monsterverwerking bij kamertemperatuur. Dit p...

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

Using Tomoauto: A Protocol for High-throughput Automated Cryo-electron Tomography

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native context at a molecular level. The technique involves initially preserving the sample in its native state by rapidly freezing the specimen in vitreous ice, then collecting a series of micrographs from different angles at high magnification, and finally computationally reconstructing a 3-D density map. The frozen-hydrated specimen is extremely sensitive to the electron beam and so micrographs are collected at very low electron doses to limit the radiation damage. As a result, the raw cryo-tomogram has a very low signal to noise ratio characterized by an intrinsically noisy image. To better visualize subjects of interest, conventional imaging analysis and sub-tomogram averaging in which sub-tomograms of the subject are extracted from the initial tomogram and aligned and averaged are utilized to improve both contrast and resolution. Large datasets of tilt-series are essential to understanding and resolving the complexes at different states, conditions, or mutations as well as obtaining a large enough collection of sub-tomograms for averaging and classification. Collecting and processing this data can be a major obstacle preventing further analysis. Here we describe a high-throughput cryo-ET protocol based on a computer-controlled 300kV cryo-electron microscope, a direct detection device (DDD) camera and a highly effective, semi-automated image-processing pipeline software wrapper library tomoauto developed in-house. This protocol has been effectively utilized to visualize the intact type III secretion system (T3SS) in Shigella flexneri minicells. It can be applicable to any project suitable for cryo-ET.

We present a protocol on how to utilize high-throughput cryo-electron tomography to determine high resolution in situ structures of molecular machines. The protocol permits large amounts of data to be processed, avoids common bottlenecks and reduces resource downtime, allowing the user to focus on important biological questions.

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native ...

Bioengineering

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in situ. Owing to the strong interaction of an electron with the matter, high-resolution cryo-ET studies are limited to specimens with a thickness of less than 200 nm, which restricts the applicability of cryo-ET only to the peripheral regions of a cell. A complex workflow that comprises the preparation of thin cellular cross-sections by cryo-focused ion beam micromachining (cryo-FIBM) was introduced during the last decade to enable the acquisition of cryo-ET data from the interior of larger cells. We present a protocol for the preparation of cellular lamellae from a sample vitrified by plunge freezing utilizing Saccharomyces cerevisiae as a prototypical example of a eukaryotic cell with wide utilization in cellular and molecular biology research. We describe protocols for vitrification of S. cerevisiae into isolated patches of a few cells or a continuous monolayer of the cells on a TEM grid and provide a protocol for lamella preparation by cryo-FIB for these two samples.

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

We present a protocol for lamella preparation of plunge frozen biological specimens by cryo-focused ion beam micromachining for high-resolution structural studies of macromolecules in situ with cryo-electron tomography. The presented protocol provides guidelines for the preparation of high-quality lamellae with high reproducibility for structural characterization of macromolecules inside the&#160;Saccharomyces cerevisiae.

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in ...

Biochemistry

Electron Cryotomography of Bacterial Cells

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for instance how they generate and maintain specific cell shapes, establish polarity, segregate their genomes, and divide. In order to understand these phenomena, imaging technologies are needed that bridge the resolution gap between fluorescence light microscopy and higher-resolution methods such as X-ray crystallography and NMR spectroscopy.
Electron cryotomography (ECT) is an emerging technology that does just this, allowing the ultrastructure of cells to be visualized in a near-native state, in three dimensions (3D), with "macromolecular" resolution (~4nm).1, 2 In ECT, cells are imaged in a vitreous, "frozen-hydrated" state in a cryo transmission electron microscope (cryoTEM) at low temperature (&lt; -180&deg;C). For slender cells (up to ~500 nm in thickness3), intact cells are plunge-frozen within media across EM grids in cryogens such as ethane or ethane/propane mixtures. Thicker cells and biofilms can also be imaged in a vitreous state by first "high-pressure freezing" and then, "cryo-sectioning" them. A series of two-dimensional projection images are then collected through the sample as it is incrementally tilted along one or two axes. A three-dimensional reconstruction, or "tomogram" can then be calculated from the images. While ECT requires expensive instrumentation, in recent years, it has been used in a few labs to reveal the structures of various external appendages, the structures of different cell envelopes, the positions and structures of cytoskeletal filaments, and the locations and architectures of large macromolecular assemblies such as flagellar motors, internal compartments and chemoreceptor arrays.1, 2
In this video article we illustrate how to image cells with ECT, including the processes of sample preparation, data collection, tomogram reconstruction, and interpretation of the results through segmentation and in some cases correlation with light microscopy.

We illustrate here how to use electron cryotomography (ECT) to study the ultrastructure of bacterial cells in near-native states, to "macromolecular" (~4 nm) resolution.

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for ...

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 ...

Simplified Volumetric Models as an Effective Strategy for Segmenting Actin Networks in Cryo-Electron Tomograms

Efficient methods for the extraction of features of interest remain one of the biggest challenges for the interpretation of cryo-electron tomograms. Various automated approaches have been proposed, many of which work well for high-contrast datasets where the features of interest can be easily detected and are clearly separated from one another. Our inner ear stereocilia cryo-electron tomographic datasets are characterized by a dense array of hexagonally packed actin filaments that are frequently cross-connected. These features make automated segmentation very challenging, further aggravated by the high-noise environment of cryo-electron tomograms and the high complexity of the densely packed features. Using prior knowledge about the actin bundle organization, we have placed layers of a highly simplified ball-and-stick actin model to first obtain a global fit to the density map, followed by regional and local adjustments of the model. We show that volumetric model building not only allows us to deal with the high complexity, but also provides precise measurements and statistics about the actin bundle. Volumetric models also serve as anchoring points for local segmentation, such as in the case of the actin-actin cross connectors. Volumetric model building, particularly when further augmented by computer-based automated fitting approaches, can be a powerful alternative when conventional automated segmentation approaches are not successful.

Here, we present a protocol to place simplified volumetric models into noisy, complex, tomographic 3D volumes. This allows the fast segmentation of actin filament densities, detection of systematic filament bending and of gaps in hair bundle filaments, as well as convenient quantification of volumetric model properties, such as distances.

Efficient methods for the extraction of features of interest remain one of the biggest challenges for the interpretation of cryo-electron tomograms. ...

Neuroscience

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 ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

Cryo-Electron Tomography Remote Data Collection and Subtomogram Averaging

Cryo-electron tomography (cryo-ET) has been gaining momentum in recent years, especially since the introduction of direct electron detectors, improved automated acquisition strategies, preparative techniques that expand the possibilities of what the electron microscope can image at high-resolution using cryo-ET and new subtomogram averaging software. Additionally, data acquisition has become increasingly streamlined, making it more accessible to many users. The SARS-CoV-2 pandemic has further accelerated remote cryo-electron microscopy (cryo-EM) data collection, especially for single-particle cryo-EM, in many facilities globally, providing uninterrupted user access to state-of-the-art instruments during the pandemic. With the recent advances in Tomo5 (software for 3D electron tomography), remote cryo-ET data collection has become robust and easy to handle from anywhere in the world. This article aims to provide a detailed walk-through, starting from the data collection setup in the tomography software for the process of a (remote) cryo-ET data collection session with detailed troubleshooting. The (remote) data collection protocol is further complemented with the workflow for structure determination at near-atomic resolution by subtomogram averaging with emClarity, using apoferritin as an example.

The present protocol describes high-resolution cryo-electron tomography remote data acquisition using Tomo5 and subsequent data processing and subtomogram averaging using emClarity. Apoferritin is used as an example to illustrate detailed step-by-step processes to achieve a cryo-ET structure at 2.86 &#197; resolution.

Cryo-electron tomography (cryo-ET) has been gaining momentum in recent years, especially since the introduction of direct electron detectors, improved ...

Visualization of Organelles In Situ by Cryo-STEM Tomography

Cryogenic electron microscopy (cryo-EM) relies on the imaging of biological or organic specimens embedded in their native aqueous medium; water is solidified into a glass (i.e., vitrified) without crystallization. The cryo-EM method is widely used to determine the structure of biological macromolecules recently at a near-atomic resolution. The approach has been extended to the study of organelles and cells using tomography, but the conventional mode of wide-field transmission EM imaging suffers a severe limitation in the specimen thickness. This has led to a practice of milling thin lamellae using a focused ion beam; the high resolution is obtained by subtomogram averaging from the reconstructions, but three-dimensional relations outside the remaining layer are lost. The thickness limitation can be circumvented by scanned probe imaging, similar to the scanning EM or the confocal laser scanning microscope. While scanning transmission electron microscopy (STEM) in materials science provides atomic resolution in single images, the sensitivity of cryogenic biological specimens to electron irradiation requires special considerations. This protocol presents a setup for cryo-tomography using STEM. The basic topical configuration of the microscope is described for both two- and three-condenser systems, while automation is provided by the non-commercial SerialEM software. Enhancements for batch acquisition and correlative alignment to previously-acquired fluorescence maps are also described. As an example, we show the reconstruction of a mitochondrion, pointing out the inner and outer membrane and calcium phosphate granules, as well as surrounding microtubules, actin filaments, and ribosomes. Cryo-STEM tomography excels in revealing the theater of organelles in the cytoplasm and, in some cases, even the nuclear periphery of adherent cells in culture.

Visualization of Organelles In Situ by Cryo-STEM Tomography

Cryo-STEM tomography provides a means to visualize organelles of intact cells without embedding, sectioning, or other invasive preparations. The 3D resolution obtained is currently in the range of a few nanometers, with a field of view of several micrometers and an accessible thickness in the order of 1 &#181;m.

Imaging Replicative Domains in Ultrastructurally Preserved Chromatin by Electron Tomography

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