This project has received funding from the European Commission Horizon 2020 iNEXT-Discovery project. I. M. Dobbie acknowledges funding from the Wellcome Trust (107457/Z/15/Z). This work was carried out with the support of the Diamond Light Source, instrument B24 (proposal BI25512). Our thanks to the staff at Micron and all our excellent users and collaborators for helping us establish the cryoSIM and its correlative potential.

NOTE: This protocol pertains to samples containing cells grown or deposited on transmission electron microscopy (TEM) 3 mm flat gold grids with a holey carbon support film that have been vitrified by plunge freezing or high-pressure freezing. This protocol assumes that samples have already been imaged using a conventional epifluorescence and brightfield microscope to map locations of interest for imaging in cryoSIM. See Figure 1 for an overview of the entire protocol.
1. Preparation of the cryo-stage
<ol>
	<li>Prepare ice-free liquid nitrogen (LN2) by passing LN2 through a funnel lined with any standard scientific dry paper wipes. 
	NOTE: As LN2&#160;causes burns, wear appropriate personal protective equipment, including cryo-protective gloves and goggles, when handling it. Such liquids may displace oxygen and cause rapid suffocation and should be handled in a properly ventilated area.</li>
	<li>Remove surface dust from the cryo-stage with pressurized air. Expel liquid from the pressurized air container before using it.</li>
	<li>Ensure the sample-loading cartridge is in place with the appropriate chambered sample holder (check that there is no grid left in the sample holder from a previous experiment) (Figure 2).</li>
	<li>Remove the lid from the external dewar of the cryo-stage, and pour filtered LN2&#160;until approximately 1/4th full. Wait until initial boiling subsides before pouring more; fill the vessel to about 2/3rd full. Replace the lid carefully, pointing the nozzle away from the handler and the stage/optics while LN2&#160;boils out from the outlet.</li>
	<li>Once LN2&#160;has stopped coming out of the outlet, place the outlet pipe over the stage dewar on the cryo-stage.</li>
	<li>Plug-in the power source of the stage, and connect the USB cable to the cryo-stage. Ensure the heated sample chamber lid is plugged in. Plug in the external dewar to the stage. 
	NOTE: Do not plug in the external dewar until the outlet pipe has been positioned over the stage dewar on the cryo-stage. If LN2 overflows (or to prevent it from overflowing), pull out the USB cable for ~10 s, allow it to equilibrate, and reconnect the USB cable to reactivate the sensor.</li>
	<li>After LN2&#160;has been delivered into the stage dewar, press the release button on the cryo-stage to allow LN2&#160;to enter the sample chamber. 
	NOTE: Do not leave the system unattended while LN2 is filling the chamber.</li>
	<li>Wait for ~30-45 min to allow the system to cool and stabilize before commencing image acquisition. Periodically check the external dewar, and top-up with filtered LN2 if it is less than a quarter full (approximately every hour).</li>
</ol>
2. Transfer of the sample storage box into the cryo-stage
NOTE: Immerse the sample storage box, holder, and the tips of any instruments (e.g., forceps) inside filtered LN2 to cool them before touching any cold surfaces such as the sample or any objects inside the sample chamber. Wear a laboratory coat and gloves when handling biological samples.
<ol>
	<li>Ensure vitrified samples are in a cryo-compatible container, and bring them to the microscope. Press the corresponding button on the cryo-stage to turn on the light in the sample chamber.</li>
	<li>Use the hex key on the cassette tool to open the two plates of the sample transfer cassette. Open the plates wide enough to drop in the grid between the two plates, but avoid opening to the maximum open position to prevent the grid falling through the other side.</li>
	<li>Use long forceps to lift the sample grid box out of the LN2, turn it where the notch aligns with the position of the storage position inside the stage, and place it onto the stage. Use the appropriate device (e.g., a screwdriver) to open the storage box lid to the correct sample position.</li>
	<li>Using inverted forceps (or any fine-tip surgical forceps), remove the TEM grid from the sample holder, immerse it inside the LN2, and drop it into position in the sample transfer cassette, keeping close to the LN2&#160;during the transfer process. Ensure the carbon film side is placed so that it will ultimately be facing the objective on the sample bridge.</li>
	<li>Close the sample cartridge using the hex key on the cassette tool. Close and remove the storage box along with any remaining samples.</li>
	<li>Use the magnet point on the cassette tool to lift and mount the cartridge containing the grid onto the sample bridge. Keep it immersed/close to the LN2&#160;during the transfer process. Ensure the orientation is appropriate for the bridge (the two magnets will make contact with the magnets on the bridge). Place the cassette flat within the positioning pins of the bridge, and gently nudge to ensure it is fixed. 
	&#8203;NOTE: A sample cassette for clipped grids that have been prepared for subsequent focused ion beam milling is also available at the CryoSIM facility at beamline B24.</li>
</ol>
3. Stage docking and focusing
<ol>
	<li>Move the cryo-stage lid opening to the imaging position, and turn off the sample chamber light. Slide the stage towards the optics to align it under the objective lens. Gently drop the objective into position using the lever, ensuring that it rests within the lid of the cryo-stage, but does not touch it. 
	NOTE: Ensure the external Dewar&#39;s outlet pipe does not touch the stage dewar on the cryo-stage at any point during data collection.</li>
	<li>Cover the stage and optics with an opaque black curtain. Start the control software Cockpit on the cryoSIM PC (Figure 3 and Figure 4).</li>
	<li>Click on the readout mode button for each camera, and set it to CONV 3 MHz. Check that the temperature of each camera is -80 &#176;C, and the camera fan is off.</li>
	<li>Turn on the reflected camera. Under Lights, choose ambient (exposure 20 ms), and under linkam, click on condenser. Click on the Video mode button.</li>
	<li>In the Mosaic view window, zoom out (mouse scroll) to see the grid outline. Click on Find stage if it cannot be seen. Center the grid by double-left-clicking in the middle of the circle.</li>
	<li>Focus the sample until the grid support film (or any other relevant sample feature) is in focus, using the up and down keys to move the cryo-stage up and down, and using the 9 and 3 keys on the numerical pad to change the z-step (set it to 20 &#181;m for initial focusing). 
	&#8203;NOTE: If the user runs out of travel in z, the stage can be manually moved up or down using the wheel under the cryo-stage. Turn it by one notch at a time, and check if the sample is within the range of view. Alter the direction again if focusing worsens.</li>
</ol>
4. Brightfield mosaic acquisition
<ol>
	<li>Once the stage is centered, turn off video mode, and collect a visible light mosaic (click on Run mosaic in the Mosaic view to produce tiles of visible light images that spiral outward from the center). If the grid is bent, try different positions on the grid (double left click within Mosaic view), re-focus, and click on Run mosaic again to collect partial mosaics. Alternatively, drop-in a focused image on top of the mosaic (Figure 3).</li>
	<li>Save the view by clicking on Save mosaic. Give it a short filename containing information on the storage box and respective grid number (a timestamp will be appended automatically to the file name).</li>
</ol>
5. Identification of areas of interest
<ol>
	<li>Inspect the brightfield mosaic alongside any previous fluorescence &#34;map&#34; images for where cells or biological features of interest have previously been located. Check if those areas produce suitable fluorescence.
	<ol>
		<li>Turn off the ambient light and condenser as well as the video mode&#160;if active. Turn on the required excitation laser (405, 488, 561, or 647), and choose the corresponding camera and filter, initially at 50 mW, for 50 ms exposure time. 
		NOTE: Increase/decrease these camera and filter settings depending on the fluorescence signal.</li>
		<li>Press 0 to snap an image and * to auto-contrast. Alternatively, manually adjust the contrast by using the slider at the bottom of the image. 
		NOTE: Switch on the Laser on sign for the room.</li>
		<li>Once biologically interesting cells with suitable fluorescence have been found, mark their positions using the Mark site button in Mosaic view. 
		NOTE: These marked sites will appear in the enclosed list and can be accessed by double-clicking on the coordinates. When returning to a marked site, zoom into the area before double-clicking.</li>
		<li>Continue marking all potential sites before commencing image acquisition. Re-save the mosaic with the marked sites; click on Save sites to file.
		<ol>
			<li>To stitch the mosaic images together using the StitchM software (developed inhouse at beamline B24), drag and drop the .txt mosaic file into the StitchM file with extension .bat and save the combined tiff image of the mosaic tiles in the same folder. To save an image with the marked sites, drag and drop the mosaic.txt file and the markers.txt file into the icon at the same time. 
			&#8203;NOTE: Balance data collection against the needs of the project (e.g., if correlative imaging will be done, check how many images can be taken with the partner-imaging modality, and choose the appropriate number of sites for cryoSIM imaging). In addition, if the external dewar requires refilling with LN2 , the cryo-stage system will change position, and the marked sites will likely not return to the same locations; therefore consider this when choosing the number of sites to be imaged.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
6. Data collection strategy
<ol>
	<li>Set the laser exposure time based on the counts in the dynamic range in the fluorescence image (at the bottom of the camera views window). Choose which filter to apply, and optimize these settings for each wavelength of excitation light to be used, turning each laser on separately. 
	&#8203;NOTE: Although 10,000-20,000 counts are optimal, lower counts are acceptable if there is a good contrast. Ideally, check filter settings before each image stack is acquired because cells at different areas of the grid could have variable fluorescence levels.</li>
</ol>
7. Data collection
<ol>
	<li>Click on both cameras to turn them on. Return to one of the marked sites, and focus on the desired depth again. Once in focus in an area of interest, move out of focus upwards (&#8593; key) in&#160;the XY window in the Macro Stage to choose the height of the z stack to acquire, and click on Save top. Move out of focus downwards (&#8595; key), click on Save bottom and then on Go to the centre, and check that the image is still in focus. 
	NOTE: The sample height will be shown in the window.</li>
	<li>Right-click on slm (spatial light modulator) in the Cockpit window, and make sure the angle is set at 0.41. In the Cockpit window, select Single-site experiment. 
	NOTE: Do not click the slm on; Cockpit will automatically turn it on during image acquisition.
	<ol>
		<li>From the dropdown list, select Structured Illumination. Alter the Stack height so that it equals the z-height + 1 &#181;m. 
		NOTE: The addition of 1 &#181;m ensures the capture of the entire sample in z and minimizes reconstruction artefacts.</li>
		<li>Enter the exposure times (ms) for the 405 nm and 488 nm lasers in the upper row (for the reflected camera) and the exposure times for the 561 nm and 647 nm lasers in the lower row (for the transmitted camera). 
		NOTE: The values for exposure time should match the values decided on previously and shown in the main Cockpit window.</li>
		<li>Input a file name (naming convention: box number_grid area_filters_FL) (FL (fluorescence) or BF (brightfield) depending on what type of imaging is being done). Click on Update to produce a new file containing the date and time without overwriting previous files. Then, click on Start.</li>
	</ol>
	</li>
	<li>Check the Camera view while data is being collected. Re-take the images if there is any xy displacement. If the LN2&#160;dewar refills the cryo-stage during image acquisition, abort the process by clicking on the Abort button in the Cockpit software.
	<ol>
		<li>Once the external dewar has finished refilling the stage dewar, repeat the experiment because the refill displaces the sample vertically. Refocus the image to re-center it in z, and repeat the Single-site experiment. Repeat the Single-site experiment for all combinations of lasers and filters needed. 
		NOTE: It takes approximately 30 s-1 min to finish refilling. During the imaging of one grid, refilling will happen ~4-8x.</li>
	</ol>
	</li>
	<li>At each position, collect a z stack using visible light.
	<ol>
		<li>Switch off the lasers, and switch on Ambient light and condenser. Under Single-site experiment, select Z-stack, set the Ambient light to 20 ms exposure, and keep z height as above.</li>
	</ol>
	</li>
	<li>Repeat steps 7.2 to 7.4 for all sites marked on the grid.</li>
	<li>Before moving to another sample, delete the mosaic by clicking on Delete tiles.
	<ol>
		<li>Draw a square around all the tiles to delete them. Delete the markers as well by selecting all of them in the list and then clicking on Delete selected sites.</li>
	</ol>
	</li>
	<li>Turn off the ambient light and condenser before proceeding to change the grid. Follow the steps in section 4 in reverse order to undock the stage from under the objective and change the grid.</li>
</ol>
8. After imaging
<ol>
	<li>After imaging is finished, undock the stage, and remove all the samples. Turn off the sample chamber light. Unplug the external dewar, and decant any remaining LN2&#160;into another cryo-compatible container, allowing the dewar to safely return to a normal temperature.</li>
	<li>Put the lid plug on the cryo-stage. Wait until the option to bake-out the cryo-stage display becomes available after no more LN2&#160;remains in the stage dewar. Press the bake-out button to enter the heating mode, and remove any moisture from the cryo-stage to avoid ice formation.</li>
</ol>
9. Reconstruction
<ol>
	<li>Transfer raw SIM data files to the appropriate workstation for reconstruction. Run processing in batches through the Processing Task builder window using channel-specific optical transfer functions (OTFS) (calculated from multi-fluorescent beads point spread functions) and K0 angles (0.29278, -1.8028, 2.3786) with a constant Wiener filter for all channels of 0.004 and a bias offset of 200.</li>
	<li>In the Additional options panel, ensure that negative intensities are discarded by keeping other options unchecked. Save the SIR images into a folder named &#34;processed&#34;. 
	&#8203;NOTE: Commercial SIM reconstruction software usually produces reconstructed SIM data and retains the file name, but appends SIR.dv at the end. A log file is also created that contains the processing protocol, steps, and statistical information on reconstruction success.</li>
</ol>
10. Chromatic shift correction
<ol>
	<li>Download the software Chromagnon7 to correct for the chromatic shift.</li>
	<li>Use the chromatic shift reference matrix that corresponds to the fluorescence wavelengths of the data collected (provided by the beamline). 
	&#8203;NOTE: The reference file is a &#39;chromagnon.csv&#39; file, which contains the alignment parameters and has been obtained from calibration images using multi-fluorescent nanoparticles. It can be used to batch-process multiple data sets at once.
	<ol>
		<li>Choose the appropriate reference file which matches the laser wavelength and filter used for imaging the sample, and add it to the reference field. Add the reconstructed SIR data to be aligned in the source field, and click on Run.</li>
		<li>Check that the fluorescence signal is now aligned in the images. For batch-processing, place all SIR images to be aligned in the source field and the reference file in the reference field, and press Run all.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Schermelleh, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super-resolution+microscopy+demystified.">Super-resolution microscopy demystified.</a> Nature Cell Biology. 21 (1), 72-84 (2019).</li><li>Ball, G., Demmerle, J., Kaufmann, R., Davis, I., Dobbie, I. M., Schermelleh, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SIMcheck:+A+toolbox+for+successful+super-resolution+structured+illumination+microscopy.">SIMcheck: A toolbox for successful super-resolution structured illumination microscopy.</a> Scientific Reports. 5 (1), 15915 (2015).</li><li>Kounatidis, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+Correlative+cryo-structured+illumination+fluorescence+and+soft+X-ray+microscopy+elucidates+reovirus+intracellular+release+pathway.">3D Correlative cryo-structured illumination fluorescence and soft X-ray microscopy elucidates reovirus intracellular release pathway.</a> Cell. 182 (2), 515-530 (2020).</li><li>Gustafsson, M. G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surpassing+the+lateral+resolution+limit+by+a+factor+of+two+using+structured+illumination+microscopy.">Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy.</a> Journal of Microscopy. 198 (2), 82-87 (2000).</li><li>Rego, E. H., Shao, L., Rego, E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Practical+structured+illumination+microscopy.">Practical structured illumination microscopy.</a> Methods in Molecular Biology. 1251, 1251 (2015).</li><li>Phillips, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CryoSIM:+super-resolution+3D+structured+illumination+cryogenic+fluorescence+microscopy+for+correlated+ultrastructural+imaging.">CryoSIM: super-resolution 3D structured illumination cryogenic fluorescence microscopy for correlated ultrastructural imaging.</a> Optica. 7 (7), 802 (2020).</li><li>Matsuda, A., Schermelleh, L., Hirano, Y., Haraguchi, T., Hiraoka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accurate+and+fiducial-marker-free+correction+for+three-dimensional+chromatic+shift+in+biological+fluorescence+microscopy.">Accurate and fiducial-marker-free correction for three-dimensional chromatic shift in biological fluorescence microscopy.</a> Scientific Reports. 8, 7583 (2018).</li><li>Kaufmann, R., Hagen, C., Gr&#252;newald, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+cryo-microscopy:+current+challenges+and+prospects.">Fluorescence cryo-microscopy: current challenges and prospects.</a> Current Opinion in Chemical Biology. 20, 86-91 (2014).</li><li>Phillips, M. A., Pinto, D. M. S., Dobbie, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SPEKcheck-fluorescence+microscopy+spectral+visualisation+and+optimisation:+a+web+application,+javascript+library,+and+data+resource.">SPEKcheck-fluorescence microscopy spectral visualisation and optimisation: a web application, javascript library, and data resource.</a> Wellcome Open Research. 3, 92 (2018).</li><li>Demmerle, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategic+and+practical+guidelines+for+successful+structured+illumination+microscopy.">Strategic and practical guidelines for successful structured illumination microscopy.</a> Nature Protocols. 12 (5), 988-1010 (2017).</li></ol>

The authors declare no competing financial interests.

3D SIM at cryogenic temperatures has many advantages over other SR imaging techniques for imaging vitrified biological material. It requires significantly fewer images per z slice compared to other SR methods, resulting in less irradiation and a lower chance of ice crystal formation for vitrified samples. It is also able to image whole cells and can be correlated with X-ray tomography to match structure with function. Interestingly, most commercially available fluorophores and fluorescence tags bleach less under cryogenic conditions than at room temperature. However, given the high quantum yield of most common fluorophores at room temperature (more than 80% in some cases), the absolute gain detected in photons is not due to changes in quantum yield, but due to a reduction in the complex bleaching processes. More information on the yield of fluorophores at cryogenic temperatures can be found in 8.
It is critical that samples arriving at the cryoSIM have been premapped using a conventional cryofluorescence microscope with brightfield capability to produce a grid &#34;map&#34; that includes highlighting of all potential AOIs for further imaging (Figure 5). Access time at the cryoSIM is allocated via a competitive process that involves the submission of a proposal, which is subsequently evaluated for technique feasibility and biomedical impact. Time at the equipment, therefore, is always &#34;at a premium&#34;, and premapped grids allow the most efficient use of an allocation. It is also essential that the sample is kept vitrified, especially during sample transfer from the sample holder to the imaging platform, to minimize the formation of ice crystals and subsequent sample damage. The sample should be of good quality to produce the best SIM images. A well-prepared sample will be characterized by the following features: (a) it will have no ice crystal contamination, (b) the grid used will be a finder grid, (c) the carrier will be flat, (d) the grid mesh and substrate surface will not be auto-fluorescent, and (e) there will be no breaks in the support membrane. These prerequisites can be achieved by careful sample handling and ensuring that samples always stay vitrified.
It is important to check in advance whether proposed sample fluorophores will give enough signal in the cryoSIM microscope. Tools such as SPEKCheck9 can aid with choosing the optimal fluorophore and filter combinations. If there are issues with the raw data collection or the reconstruction process, artefacts can appear in the images after reconstruction. Examples of various artefacts have been documented by Demmerle et al.10 The image reconstruction parameters can be reviewed in the SoftWoRx log file if the reconstruction is not optimal by opening the reconstruction summary file. There should be consistent line spacing across angles in a given channel and relatively consistent amplitude. Variation of more than 30% and values significantly above 1 (if bead size compensation is applied) should be more closely investigated and are likely to indicate failed reconstructions. In addition, the SIMcheck2 software in Fiji can also be used to perform various checks on the raw and reconstructed data to diagnose the cause of errors in the imaging or reconstruction parameter settings. SIM-check and its modulation contrast map can also aid in the assessment of the quality of reconstructed data by interpreting which areas of an image are likely to be real structures versus artefacts.
Low modulation contrast (shown by dark color, in Figure 5E) within the nuclear area means that this region is going to be more susceptible to reconstruction artifacts, therefore implying that the hash patterns shown in the nucleus could be classified (Figure 5D) as an artefact. Strong fluorescence signal areas are more likely to accurately reflect native structures in the processed data. In areas of weak signal where fluorophores are distributed over wider areas, such as the total surface of a vesicle, it is likely that real signal coexists with processing artefacts, and care should be taken in the interpretation of that data. After inspection of the full-range reconstructed data to ensure there are no strange artifacts, and that the background is generally Gaussian and centered near zero, the data is generally clipped at zero, or the modal value-the peak of the background signal-should be very near zero. This ensures that the dynamic range of the displayed image is not dominated by negative background artifacts. When a weaker signal is expected, extra care should be taken in analyzing the features and ensuring they are real structures rather than reconstruction artefacts.
There are some limitations of the imaging system. Because the sample stage is flat, samples with variable thickness or grids that are not flat are not ideal subjects for imaging. Additionally, if correlative imaging will be done using soft X-ray tomography, cells near grid boundaries should not be imaged as these will not be visible in the X-ray microscope during tilt series acquisition. Finally, the amount of blotting of the sample before plunge freezing has a significant impact on the imaging quality; too little blotting results in samples that are too thick, giving suboptimal SIM images with high background noise, while too much blotting can cause cells to become misshaped and therefore more susceptible to heat damage from the incident laser beam. In summary, cryoSIM is a powerful fluorescence microscopy tool for imaging biological samples in 3D in a near-native stage and has wide-ranging applications in many areas.

SR imaging techniques have become widely accessible to biologists over the last decade1. They allow high-resolution imaging of fluorescently tagged samples beyond the diffraction limit. However, it has been challenging to adapt SR microscopy methods to work with samples at cryogenic temperatures2. This would be advantageous for correlative imaging in combination with electron or X-ray tomography. Recently, SIM has been adapted for use with cryogenic samples and has successfully been shown to enable correlative studies of biological cells in conjunction with soft X-ray tomography (SXT)3 at the correlative cryo-imaging beamline B24 at the Diamond Light Source Synchrotron (https://www.diamond.ac.uk/Instruments/Biological-Cryo-Imaging/B24.html). SIM can double the resolution of conventional wide-field microscopy by illuminating the sample with striped patterns of light (Moir&#233; fringes) at three angles and in five phases. The interference between these light patterns and the sample fluorescence can be used to computationally uncover extra information about sub-diffraction structures4,5.
There are several advantages of SIM over other SR techniques for cryogenic applications. First, it can work without specially designed blinking fluorophores; conventional fluorophores can be used, giving access to a wider range of potential fluorescent tagging agents6. In addition, it only requires 15 images per z slice (in 3D; 9 images for 2D), whereas other SR methods take approximately 1000 images per slice, increasing the chance of the sample being heated and therefore increasing the risk of ice crystal formation, which can cause artefacts. Finally, this technique can image thicker biological samples of over 10 &#181;m, allowing whole cells to be imaged in their near-native state6. The cryoSIM has been built using standard optical components and with open-access software for imaging, making it easy to document and duplicate if desired6. The cryoSIM has a 100x/0.9 numerical aperture objective (see the Table of Materials); further information on its optical components, design parameters, and performance has been described by Phillips et al.6 Here, this protocol demonstrates how to use the cryoSIM microscope including how to load and unload samples on the cryogenic stage, how to collect data on the microscope, and how to reconstruct the SIM images.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Auto grids</td>
			<td>FEI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Autogrids</td>
			<td>Thermo Fisher Scientific</td>
			<td>1036173</td>
			<td></td>
		</tr>
		<tr>
			<td>BioTracker 488 Green Microtubule Cytoskeleton Dye</td>
			<td>Sigma-Aldrich</td>
			<td>SCT142</td>
			<td></td>
		</tr>
		<tr>
			<td>Cockpit Software</td>
			<td>Oxford University</td>
			<td></td>
			<td>https://github.com/MicronOxford/cockpit</td>
		</tr>
		<tr>
			<td>cryo compatible polyurethane container</td>
			<td>Jena bioscience</td>
			<td>CC-FD-800</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryo TEM grid storage box</td>
			<td>Thermo Fisher Scientific</td>
			<td>Model#AutoGrid</td>
			<td></td>
		</tr>
		<tr>
			<td>cryo-SIM microscope</td>
			<td>Custom made</td>
			<td>N/A</td>
			<td>Custom made, see following reference for the design:&#160; Michael A. Phillips, Maria Harkiolaki, David Miguel Susano Pinto, Richard M. Parton, Ana Palanca, Manuel Garcia-Moreno, Ilias Kounatidis, John W. Sedat, David I. Stuart, Alfredo Castello, Martin J. Booth, Ilan Davis, and Ian M. Dobbie, &#34;CryoSIM: super-resolution 3D structured illumination cryogenic fluorescence microscopy for correlated ultrastructural imaging,&#34; Optica 7, 802-812 (2020). Has a Nikon TU Plan Apo 100x/0.9 NA.&#160;</td>
		</tr>
		<tr>
			<td>Cryostage system</td>
			<td>Linkam Scientific Instruments</td>
			<td>CMS196</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine tip surgical forceps</td>
			<td>Ted Pella</td>
			<td>38125</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoTracker Deep Red FM</td>
			<td>Thermo Fisher Scientific</td>
			<td>M22426</td>
			<td></td>
		</tr>
		<tr>
			<td>Python Microscope Software</td>
			<td>Oxford University</td>
			<td></td>
			<td>https://www.python-microscope.org/</td>
		</tr>
		<tr>
			<td>Scientific dry paper wipes</td>
			<td>Kimberly-Clark 7551</td>
			<td>2</td>
			<td></td>
		</tr>
		<tr>
			<td>SIM Reconstruction Software</td>
			<td>softWoRx, GE Healthcare</td>
			<td>Version&#160; 6.5.2</td>
			<td></td>
		</tr>
		<tr>
			<td>StitchM Software</td>
			<td>Diamond Light Source</td>
			<td></td>
			<td>https://github.com/DiamondLightSource/StitchM</td>
		</tr>
		<tr>
			<td>TEM grids for samples</td>
			<td>Quantifoil Micro Tools</td>
			<td>G200F1</td>
			<td></td>
		</tr>
	</tbody>
</table>

cryo-structured illumination microscopic data collection from cryogenically preserved cells

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than conventional fluorescence microscopy. This super-resolution (SR) technique enables visualization of molecular processes in whole cells and has the potential to be used in conjunction with electron microscopy and X-ray tomography to correlate structural and functional information. A SIM microscope for cryogenically preserved samples (cryoSIM) has recently been commissioned at the correlative cryo-imaging beamline B24 at the UK synchrotron. 
It was designed specifically for 3D imaging of biological samples at cryogenic temperatures in a manner compatible with subsequent imaging of the same samples by X-ray microscopy methods such as cryo-soft X-ray tomography. This video article provides detailed methods and protocols for successful imaging using the cryoSIM. In addition to instructions on the operation of the cryoSIM microscope, recommendations have been included regarding the choice of samples, fluorophores, and parameter settings. The protocol is demonstrated in U2OS cell samples whose mitochondria and tubulin have been fluorescently labelled.

A sample containing U2OS cells was stained with a mixture of green microtubule cytoskeleton dye and red mitochondria dye, resulting in the staining of the microtubule component of the cytoskeleton (green) and the mitochondria (red). Subsequent imaging showed the localization of mitochondria within the cell as well as the arrangement of the microtubules, highlighting the structural framework that they provide to the cell and the assembly of the cytoskeleton around organelles such as the nucleus. The resolution in cryoSIM is significantly higher than that in standard epifluorescence microscopy (Figure 5). Figure 6 demonstrates how the fluorescent &#34;map&#34; from a conventional epifluorescence microscope can be used to locate areas of interest for imaging and the corresponding cryoSIM-reconstructed image from a location on the grid.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62274/62274fig01.jpg" /> 
Figure 1: Flow chart showing the stages of the cryoSIM imaging protocol. Abbreviation: cryoSIM = Cryo-structured illumination microscopy. <a href="https://www.jove.com/files/ftp_upload/62274/62274fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62274/62274fig02.jpg" /> 
Figure 2:&#160;The cryo-stage. (A) The cryo-stage setup. (B) A sample grid shown held by inverted forceps. (C) Components of the cryo-stage. The connection ports are labelled, with the colors corresponding to orange: power supply, yellow: heated stage lid, blue: external dewar, green: connection to PC. Abbreviations: PC = personal computer; LN2 = liquid nitrogen. <a href="https://www.jove.com/files/ftp_upload/62274/62274fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62274/62274fig03.jpg" /> 
Figure 3: Views of the Cockpit software panels. (A) Main panel, (B) macro stage XY, (C) mosaic view, (D) camera views. Abbreviation: SLM = spatial light modulator. <a href="https://www.jove.com/files/ftp_upload/62274/62274fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62274/62274fig04.jpg" /> 
Figure 4: Views of the Cockpit software panels. (A) Z stack single site experiment. (B) SI single site experiment. (C) Keyboard shortcuts for the cockpit software used during image acquisition. (D) The cryoSIM microscope is on-site at beamline B24 at the Diamond Light Source synchrotron. Abbreviation: cryoSIM = Cryo-structured illumination microscopy. <a href="https://www.jove.com/files/ftp_upload/62274/62274fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62274/62274fig05.jpg" /> 
Figure 5: Resolution of cryoSIM. (A) Mosaic view of a grid under examination. (B) Brightfield image of an Area of Interest (AOI). (C) Pseudo-widefield image compared to its (D) SIM image showing the increase in resolution. The white arrow indicates SIM reconstruction artefacts. (E) Modulation contrast map combining the pixel intensity information of the reconstructed image with the color information of the respective modulation contrast-to-noise ratio (MCNR) values of the raw data generated by SIMCheck2. The bright and dark regions show high and low contrast, respectively. Scale bar = 10 &#181;m. CryoSIM imaging settings: excitation/emission wavelengths: 488/525 nm, 50 mW laser power, 50 ms exposure time and 647/655 nm, 20 mW laser power, 5 ms exposure time. Abbreviation: cryoSIM = Cryo-structured illumination microscopy. <a href="https://www.jove.com/files/ftp_upload/62274/62274fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62274/62274fig06.jpg" /> 
Figure 6: Image reconstruction in cryoSIM from a location on the grid in a fluorescence map from a conventional epifluorescence map. (A,B) Overlay of brightfield and fluorescence image maps generated with a conventional epifluorescence microscope. This map is used to locate regions of interest to subsequently image in cryoSIM. (C) The reconstructed cryoSIM image obtained at the location shown in (B). Abbreviation: cryoSIM = Cryo-structured illumination microscopy.&#160;<a href="https://www.jove.com/files/ftp_upload/62274/62274fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells at JoVE.com

Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells

This protocol demonstrates how to image biological cryo-preserved samples using cryo-structured illumination microscopy. We demonstrate the methodology by imaging the cytoskeleton of U2OS cells.

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than ...

cryo-structured-illumination-microscopic-data-collection-from

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JoVE Journal

Biology

4.0K Views.  Beamline B24, Diamond Light Source. This protocol demonstrates how to image biological cryo-preserved samples using cryo-structured illumination microscopy. We demonstrate the methodology by imaging the cytoskeleton of U2OS cells.

Video: Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells

Blood Collection from the American Horseshoe Crab, Limulus Polyphemus

The horseshoe crab has the best-characterized immune system of any long-lived invertebrate. The study of immunity in horseshoe crabs has been facilitated by the ease in collecting large volumes of blood and from the simplicity of the blood. Horseshoe crabs show only a single cell type in the general circulation, the granular amebocyte. The plasma has the salt content of sea water and only three abundant proteins, hemocyanin, the respiratory protein, the C-reactive proteins, which function in the cytolytic destruction of foreign cells, including bacterial cells, and &alpha;2-macroglobulin, which inhibits the proteases of invading pathogens. Blood is collected by direct cardiac puncture under conditions that minimize contamination by lipopolysaccharide (a.k.a., endotoxin, LPS), a product of the Gram-negative bacteria. A large animal can yield 200 - 400 mL of blood. For the study of the plasma, blood cells are immediately removed from the plasma by centrifugation and the plasma can then be fractionated into its constituent proteins. The blood cells are conveniently studied microscopically by collecting small volumes of blood into LPS-free isotonic saline (0.5 M NaCl) under conditions that permit direct microscopic examination by placing one of more LPS-free coverglasses on the culture dish surface, then mounting those coverglasses in simple observation chambers following cell attachment. A second preparation for direct observation is to collect 3 - 5 mL of blood in a LPS-free embryo dish and then explanting fragments of aggregated amebocytes to a chamber that sandwiches the tissue between a slide and a coverglass. In this preparation, the motile amebocytes migrate onto the coverglass surface, where they can readily be observed. The blood clotting system involves aggregation of amebocytes and the formation of an extracellular clot of a protein, coagulin, which is released from the secretory granules of the blood cells. Biochemical analysis of washed blood cells requires that aggregation and degranulation does not occur, which can be accomplished by collecting blood into 0.1 volumes of 2% Tween-20, 0.5 M LPS-free NaCl, followed by centrifugation of the cells and washing with 0.5 M NaCl.

The American horseshoe crab, Limulus polyphemus, is arguably the most convenient source for large quantities of blood of any invertebrate. The blood is simple in composition, with only one cell-type in the general circulation, the granular amebocyte, and only three abundant proteins in the plasma, hemocyanin, the C-reactive proteins, and &alpha;2-macroglobulin. Blood is collected from the heart and the blood cells and plasma are separated by centrifugation.

The horseshoe crab has the best-characterized immune system of any long-lived invertebrate.  The study of immunity in horseshoe crabs has been facilitated ...

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 ...

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

Collection and Identification of Pollen from Honey Bee Colonies

Researchers often collect and analyze corbicular pollen from honey bees to identify the plant sources on which they forage for pollen or to estimate pesticide exposure of bees via pollen. Described herein is an effective pollen-trapping method for collecting corbicular pollen from honey bees returning to their hives. This collection method results in large quantities of corbicular pollen that can be used for research purposes. Honey bees collect pollen from many plant species, but typically visit one species during each collection trip. Therefore, each corbicular pollen pellet predominantly represents one plant species, and each pollen pellet can be described by color. This allows the sorting of samples of corbicular pollen by color to segregate plant sources. Researchers can further classify corbicular pollen by analyzing the morphology of acetolyzed pollen grains for taxonomic identification. These methods are commonly used in studies related to pollinators such as pollination efficiency, pollinator foraging dynamics, diet quality, and diversity. Detailed methodologies are presented for collecting corbicular pollen using pollen traps, sorting pollen by color, and acetolyzing pollen grains. Also presented are results pertaining to the frequency of pellet colors and taxa of corbicular pollen collected from honey bees in five different cropping systems.

We describe methods for collecting corbicular pollen from honey bees as well as protocols for color sorting, acetolysis, and microscope slide preparation of pollen for taxonomic identification. In addition, we present pellet color and taxonomic diversity of corbicular pollen collected from five cropping systems using pollen traps.

Researchers often collect and analyze corbicular pollen from honey bees to identify the plant sources on which they forage for pollen or to estimate ...

Tissue Collection and RNA Extraction from the Human Osteoarthritic Knee Joint

Osteoarthritis (OA) is a chronic and degenerative joint disease most often affecting the knee. As there is currently no cure, total knee arthroplasty (TKA) is a common surgical intervention. Experiments using primary human OA tissues obtained from TKA provide the capability to investigate disease mechanisms ex vivo. While OA was previously thought to impact mainly the cartilage, it is now known to impact multiple tissues in the joint. This protocol describes patient selection, sample processing, tissue homogenization, RNA extraction, and quality control (based on RNA purity, integrity, and yield) from each of seven unique tissues to support disease mechanism investigation in the knee joint. With informed consent, samples were obtained from patients undergoing TKA for OA. Tissues were dissected, washed, and stored within 4 h of surgery by flash freezing for RNA or formalin fixation for histology. Collected tissues included articular cartilage, subchondral bone, meniscus, infrapatellar fat pad, anterior cruciate ligament, synovium, and vastus medialis oblique muscle. RNA extraction protocols were tested for each tissue type. The most significant modification involved the method of disintegration used for low-cell, high-matrix, hard tissues (considered as cartilage, bone, and meniscus) versus relatively high-cell, low-matrix, soft tissues (considered as fat pad, ligament, synovium, and muscle). It was found that pulverization was appropriate for hard tissues, and homogenization was appropriate for soft tissues. A proclivity for some subjects to yield higher RNA integrity number (RIN) values than other subjects consistently across multiple tissues was observed, suggesting that underlying factors such as disease severity may impact RNA quality. The ability to isolate high-quality RNA from primary human OA tissues provides a physiologically relevant model for sophisticated gene expression experiments, including sequencing, that can lead to clinical insights that are more readily translated to patients.

Primary tissues obtained from patients following total knee arthroplasty provide an experimental model for osteoarthritis research with maximal clinical translatability. This protocol describes how to identify, process, and isolate RNA from seven unique knee tissues to support mechanistic investigation in human osteoarthritis.

Osteoarthritis (OA) is a chronic and degenerative joint disease most often affecting the knee. As there is currently no cure, total knee arthroplasty ...

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.

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, ...

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 ...

Direct Synthesis of Gold Nanoparticles for Protein Localization Analysis

Direct Synthesis of EM-Visible Gold Nanoparticles in Cells for Protein Localization Analysis with Well-Preserved Ultrastructure

Analyzing the precise localization of protein molecules in cells with ultrastructural resolution is of great significance for the study of various physiological or pathological processes in all living organisms. Therefore, the development of clonable tags that can be used as electron microscopy probes is of great value, just as fluorescent proteins have played a crucial role in the field of optical imaging. The autonucleation suppression mechanism (ANSM) was recently uncovered, which allows for the specific synthesis of gold nanoparticles (AuNPs) on cysteine-rich tags, such as metallothionein (MT) and antifreeze protein (AFP).
Based on the ANSM, an electron microscopy labeling technology was developed, which enables the specific detection of tagged proteins in prokaryotic and eukaryotic cells with an unprecedented labeling efficiency. This study illustrates a protocol for the detection of MTn (an engineered MT variant lacking aldehyde-reactive residues) fusion proteins in mammalian cells with well-preserved ultrastructure. In this protocol, high-pressure freezing and freeze-substitution fixation were performed using non-aldehyde fixatives (such as tannic acid, uranyl acetate) to preserve near-native ultrastructure and avoid damage to the tag activity caused by aldehyde crosslinking.
A simple one-step rehydration was used prior to the ANSM-based AuNP synthesis. The results showed that the tagged proteins targeted various organelles, including the membranes and the lumen of the endoplasmic reticulum (ER), and mitochondrial matrices were detected with high efficiency and specificity. This research provides biologists with a robust protocol to address an enormous range of biological questions at the single-molecule level in cellular ultrastructural contexts.

Author Spotlight: Direct Synthesis of EM-Visible Gold Nanoparticles in Cells for Protein Localization Analysis with Well-Preserved Ultrastructure  

The present protocol describes a clonable electron microscopy labeling technology for detecting metallothionein-tagged proteins in cells using a novel autonucleation suppression mechanism-based gold nanoparticle synthesis technique.