This research was supported by KBRI basic research program through Korea Brain Research Institute funded by Ministry of Science and ICT (19-BR-01-08), and National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT)(No. 2019R1A2C1010634). SCO1-APEX2 and HRP-KDEL plasmids were kindly provided by Hyun-Woo Rhee (Seoul National University). TEM data were acquired at Brain Research Core Facilities in KBRI.

1. Cell culture with patterned grid culture dish and cell transfection with SCO1-APEX2 and HRP-KDEL plasmid vector
<ol>
	<li>Seed 1 x 105 HEK293T cells by placing them into 35-mm glass grid-bottomed culture dishes in a humidified atmosphere containing 5% CO2 at 37 &#176;C in Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) supplemented with 10% fetal bovine serum, 100 U/mL penicillin, and 100 U/mL streptomycin.</li>
	<li>The day after seeding the cells, when they have grown to 50%-60% confluency, introduce the SCO1-APEX210 and HRP-KDEL12 plasmid to the cells using transfection reagent according to the manufacturer&#8217;s instructions (SCO1-APEX2 cDNA 0.5 &#181;g + HRP-KDEL plasmid DNA 0.5 &#181;g per 3 &#181;L transfection reagent).</li>
</ol>
2. Light microscopy of cells growing on patterned culture dishes and DAB staining for APEX2 and HRP
<ol>
	<li>At 16-24 h after transfection, remove all the culture media and immediately add 250 &#181;L of warm (30-37 &#176;C) fixation solution (Table 1) by gentle pipetting. Immediately remove the fixation solution and replace it with 1.5 mL of fresh fixation solution. Incubate on ice for 60 min, and then wash three times for 10 min each in 1 mL of ice-cold 0.1 M sodium cacodylate buffer (Table 1). 
	CAUTION: Aldehyde fumes are extremely toxic. Perform all work under a ventilated fume hood.</li>
	<li>Add 1 mL of cold (0-4 &#176;C) 20 mM glycine solution and incubate for 10 min on ice followed by three washes of 5 min each in 1 mL of cold 0.1 M sodium cacodylate buffer.</li>
	<li>Prepare a fresh 1x DAB solution (3.33 mL of 0.3 M cacodylate solution + 10 &#181;L of 30% H2O2 + 5.67 mL of cold water + 1 mL 10x DAB solution).</li>
	<li>Add 500 &#181;L of the freshly prepared 1x DAB solution (step 2.3) and incubate on ice for approximately 5-45 min until a light brown stain is visible under an inverted light microscope (Figure 1A).</li>
	<li>Gently remove the DAB solution and rinse three times with 1 mL of cold 0.1 M sodium cacodylate buffer for 10 min each.</li>
	<li>Use a phase-contrast inverted microscope (or a bright-field light microscope) to visualize the DAB staining at a magnification of 100x or higher. Use a marker pen to mark the bottom of the glass where the region of interest (ROI) is located (Figure 1B,C).</li>
</ol>
3. Sample preparation for the EM block
<ol>
	<li>Perform cell culture and DAB staining as described in steps 2.1-2.6.</li>
	<li>Post-fix the samples with 1 mL of 2% reduced OsO4 for 1 h at 4 &#176;C. 
	CAUTION: OsO4 fumes are highly toxic. Perform all work under a ventilated fume hood.</li>
	<li>Prepare a new TCH solution (Table 1) during step 3.2 and pass through a 0.22-&#181;m filter. 
	CAUTION: TCH fumes are highly toxic. Perform all work under a ventilated fume hood.</li>
	<li>Remove the fixative and rinse three times with 1 mL of distilled water for 5 min each at room temperature (RT).</li>
	<li>Place the cells in 1 mL of previously prepared and filtered TCH solution for 20 min at RT.</li>
	<li>Rinse the cells three times with 1 mL of distilled water for 5 min each at RT.</li>
	<li>Expose the cells a second time to 1 mL of 2% osmium tetroxide in distilled water for 30 min at RT.</li>
	<li>Remove the fixative and rinse three times with 1 mL of distilled water for 5 min each at RT. Add 1 mL of 1% uranyl acetate (aqueous), and leave overnight at 4 &#176;C in the dark.</li>
	<li>Wash the cells three times in 1 mL of distilled water for 5 min each at RT.</li>
	<li>Pre-warm Walton&#8217;s lead aspartate solution (Table 1) in an oven at 60 &#176;C for 30 min.</li>
	<li>Stain the cells with Walton&#8217;s lead aspartate solution by adding 1 mL of the pre-warmed lead aspartate solution, and then place in an oven for 30 min at 60 &#176;C.</li>
	<li>Rinse the cells three times with 1 mL of distilled water for 5 min each at RT.</li>
	<li>Incubate in a graded series of 2-mL ethanol aliquots (50%, 60%, 70%, 80%, 90%, 95%, 100%, 100%) for 20 min each at RT.</li>
	<li>Decant the ethanol and incubate for 30 min in 1 mL of 3:1 ethanol:low-viscosity embedding mixture medium at RT.</li>
	<li>Remove the medium and add 1 mL of 1:1 ethanol:low-viscosity embedding mixture medium. Incubate for 30 min at RT.</li>
	<li>Remove the medium and add 1 mL of 1:3 ethanol:low-viscosity embedding mixture medium. Incubate for 30 min at RT.</li>
	<li>Remove the medium and add 1 mL of 100% low-viscosity embedding medium and incubate overnight.</li>
	<li>Embed the sample in 100% low-viscosity embedding mixture and incubate for 24 h at 60 &#176;C.</li>
	<li>Prepare 90-nm thick sections using an ultramicrotome.</li>
	<li>Observe the grid under TEM at 200 kV.</li>
</ol>
4. Serial sectioning and mounting on indium-tin-oxide coated coverslips for SEM imaging
<ol>
	<li>Substrate preparation
	<ol>
		<li>Clean indium-tin-oxide (ITO)-coated glass coverslips (22 mm x 22 mm) by gentle agitation in isopropanol for 30-60 s.</li>
		<li>Remove the coverslips, drain off the excess isopropanol, and leave in a dust-free environment until dry.</li>
		<li>Treat the ITO-coated glass coverslips by glow discharge using a plasma coater for 1 min. 
		NOTE: Plasma activating confers a hydrophilic property on the substrate surface. It creates a very thin film of water on the substrate to prevent wrinkle formation in the sections when the section is attached to the substrate.</li>
		<li>Insert the ITO-coated glass coverslips into the substrate holder, and place into the knife boat.</li>
	</ol>
	</li>
	<li>Trimming of the sample block and serial sectioning
	<ol>
		<li>Insert the sample block into the sample holder of the ultramicrotome and set into the trimming block.</li>
		<li>Use a razor blade to trim away all excess resin around the target position (identified in step 2.6, Figure 1D-G). The shape of the block face should be trapezoid or rectangular. The leading edge and trailing edge must be absolutely parallel (Figure 1H,I).</li>
		<li>Insert the sample holder on the arm of the ultramicrotome and place the diamond knife in the knife holder. Insert the ITO glass coverslips into the ribbon carrier and clamp the carrier with the handle (Figure 1J). Set the ribbon carrier into the knife boat and fill the knife boat with filtered distilled water (Figure 1K).</li>
		<li>Adjust the carrier position with the slide of the knife by carefully pushing the handle of the holder to set the edge of the ITO glass close to the knife (Figure 1L).</li>
		<li>After cutting the section, stop the sectioning process, and slowly open the clamping screw of the tube and drain the water boat (flow rate of one drop of water per second).</li>
		<li>After completing the ribbon-collection process, remove the substrate with the handle of the clamping device and dry the ribbon (Figure 1M).</li>
	</ol>
	</li>
</ol>
5. Imaging in the SEM and alignment of the SEM image stack
<ol>
	<li>Mount the ITO-coated coverslip on aluminum stubs with sticky carbon tape. Seal the glass surface and the surface in the stub with sticky carbon tape, and then coat with a 10-nm thick carbon layer (Figure 1N).</li>
	<li>Observe the ITO-coated coverslip in a field emission SEM at a low acceleration voltage of 5 kV and a suitable working distance for the efficient collection of BSEs.</li>
	<li>Import the serial images into the Image J software (Fiji)13 using the virtual stack option. Open a new TrakEM14, and import the image stack into TrakEM. Click the right-mouse button and choose the align menu.</li>
	<li>Then select the image range (from the first image to the last image). Finish the auto-alignment, save the aligned dataset, and choose export to compile a flat image from the selected image range (from the first image to the last image). Finally, save the flat image data in AVI format in the Image J main menu. 
	NOTE: Supplemental Movie 1 and Supplemental Movie 2 show the SEM image stack and cropped image stack, respectively.</li>
</ol>
6. Segmentation of mitochondria and ER from serial images
<ol>
	<li>Start the 3dmod in IMOD15 software and open image files.</li>
	<li>In the ZaP window, draw the contour of the mitochondria and ER using middle-mouse button.</li>
	<li>To visualize the segmented volume, open the Model View window (Supplemental Movie 3).</li>
</ol>

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The authors have nothing to disclose.

Determining the cellular localization of specific proteins at a nanometer resolution using EM is crucial to understand the cellular functions of proteins. Generally, there are two techniques to study the localization of a target protein via EM. One is the immunogold technique, which has been used in EM since 1960, and the other is a technique using recently developed genetically encoded tags16. Traditional immunogold techniques have employed antibody-conjugated gold particles or quantum dots to show the location of the labeled protein. However, due to the requirement for high-quality antibodies and the penetration efficiency of antibodies affected by resin and fixative, this technique is significantly limited17. Specifically, because immunogold labeling is predominantly restricted to the surface of an ultrathin section without en bloc metal staining and strong osmium fixation, this technique is not directly applicable to modern 3DEM18. To use recent 3DEM methods, including SBEM and array tomography, with protein labeling, we utilized genetically encoded EM tags in this protocol. Genetically encoded tags do not require permeabilization, technically demanding ultra-cryosectioning, and immunostaining of individual sections because they localize to the site of interest prior to fixation.
The procedures for sample preparation in 3DEM generally include a combination of common chemical fixation and heavy metal staining methods because cells are composed mainly of C, H, O, and N, requiring staining with heavy metals to acquire contrast under EM9. Therefore, we employed reduced osmium fixation and metal staining to enhance contrast and conductivity for serial imaging. The procedure to stain samples before sectioning, known as en bloc staining, has been reported as an essential step for 3DEM methods such as SBEM and FIB-SEM19. We confirmed that the en bloc-stained cells in our protocol demonstrated clear SCO1-APEX2 and HRP-KDEL EM contrast exclusively in the mitochondrial IMS and ER, respectively, in TEM (Figure 3). Furthermore, the SEM images from 90-nm serial sections revealed a clear contrast in both organelles (Figure 4). Notably, the enhanced contrast by en bloc staining was distinctly distinguishable from the DAB signal, and the contrast and conductivity resulted in good-quality serial images (Figure 4). Additionally, this high contrast aids the facilitation of subsequent tasks such as alignment and segmentation with three-dimensional (3D) image software.
In recent years, volume electron microscopy techniques (dik-SBEM, FIB-SEM, and array tomography) have answered biological questions that required the observation of a large field of view and a 3D view. Dik-SBEM and FIB-SEM do not involve the physical handling of sections, so time consuming for alignment of images can be reduced. However, the sample has to be destroyed to obtain serial images, and the field of view is smaller than that of array tomography. Serial SEM imaging using array tomography is employed increasingly as an alternative to TEM serial sectioning, and the major advantage of this technique is its non-destructive manner and large field of view. Unlike other 3DEM techniques such as dik-SBEM and FIB-SEM, sections can be stored on a coverslip, an ITO-coated coverslip, a silicon wafer, or a tape and can be repeatedly imaged7.
APEX2 is easy to use and can give a wide range of staining densities without special equipment, unlike mini singlet oxygen generator20 or fluorescent protein21 techniques, generating DAB precipitation via a photooxidation. Its variable application has been tested in several cellular organelles including nucleus, plasma membrane, mitochondria matrix, mitochondrial cristae, ER, tubulin, and actin in COS 7 and HEK293T cell22. However, there are some limitations and several checkpoints for the use of genetically encoded tagging in electron micrographs. The expression levels of the exogenous genes should be controlled to achieve reasonable staining at EM level because if the expression of the genetic tag is too high, it may induce false-positive signals and perturb the ultrastructure of cells via membrane rupture and subcellular organelle aggregation23. Another possible problem is DAB overstaining, which has been reported to cause blurriness and membrane destruction22. To ensure the appropriate expression of genes, DAB-stained cells were compared with unstained cells using both light microscopy and EM (Figure 2 and Figure 3). Additionally, the fixation level should be well regulated to ensure that the endogenous oxidases are fully inactive. To prevent any artifacts from endogenous oxidases during processing, we fixed a monolayer of cells instead of using detached and pelleted cells24. This also helped to identify the cells of interest when we checked the DAB signal under light microscopy. Thus, our results indicate that the staining and fixation of cell monolayers are useful for CLEM using DAB staining (Figure 2). The serial images with array tomography and 3D model revealed that the ER&#8211;mitochondria contact sites occur on different z-planes (Supplemental Movie 2 and Supplemental Movie 3). The image produces a complete 3D visualization of the mitochondrial and ER networks in whole cells (Supplemental Movie 1). It suggests 3D volume analysis is essential for quantitative comparison of ER-mitochondria contacts. When studying the complex networks of intracellular organelles, this is an exceptionally useful technique.
In conclusion, this protocol was an efficient combination of CLEM and 3DEM techniques that allowed whole-cell investigation at EM level. Notably, two different tags at the same time and DAB signals in two different organelles were visible in a whole cell. In addition, labeled cells and unlabeled cells in same section can be compared, because of large scale EM. In this protocol, en bloc staining and DAB signals from genetically encoded tags were useful to investigate the interaction between membranous organelles in whole cells. This could be a suitable application for large-scale EM to investigate other cellular interactions.

Mitochondria and endoplasmic reticulum (ER) are membrane-bound cellular organelles. Their connection is necessary for their function, and proteins related to their network have been described1. The distance between the mitochondria and ER has been reported as approximately 100 nm using light microscopy2; however, recent super-resolution microscopy3 and electron microscopy (EM)4 studies have revealed it to be considerably smaller, at approximately 10-25 nm. The resolution achieved in super-resolution microscopy is lower than EM, and specific labeling is necessary. EM is a suitable technique to attain a sufficiently high-resolution contrast for structural studies of the connections between mitochondria and ER. However, a disadvantage is the limited z-axis information because the thin sections must be approximately 60 nm or thinner for conventional transmission electron microscopy (TEM). For sufficient EM z-axis imaging, three-dimensional electron microscopy (3DEM) can be used5. However, this involves the preparation of hundreds of thin serial sections of whole cells, which is very tricky work that only a few skilled technologists have mastered. These thin sections are collected on fragile formvar film-coated one-hole TEM grids. If the film breaks on one gird, serial imaging and volume reconstruction is not possible. Serial block-face scanning electron microscopy (SBEM) is a popular technique for 3DEM that uses destructive en bloc sectioning inside the scanning electron microscope (SEM) vacuum chamber with either a diamond knife (Dik-SBEM) or a focused ion beam (FIB-SEM)6. However, because those techniques are not available at all facilities, we suggest array tomography7 using serial sectioning and SEM. In array tomography, serial sections cut using an ultramicrotome are transferred to a glass coverslip instead of a TEM grid and visualized via light microscopy and SEM8. To enhance the signal for backscatter electron (BSE) imaging, we utilized an en bloc EM staining protocol employing osmium tetroxide (OsO4)-fixed cells with osmiophilic thiocarbohydrazide (TCH)9, enabling us to obtain images without post-embedding double staining.
Additionally, the mitochondrial marker SCO1 (cytochrome c oxidase assembly protein 1)&#8211; ascorbate peroxidase 2 (APEX2)10 molecular tag was used to visualize mitochondria at the EM level. APEX2 is approximately 28 kDa and is derived from soybean ascorbate peroxidase11. It was developed to show the detailed location of specific proteins at the EM level in the same way that green fluorescent protein-tagged protein is used in light microscopy. APEX2 converts 3,3&#39;&#160;-diaminobenzidine (DAB) into an insoluble osmiophilic polymer at the site of the tag in the presence of the cofactor hydrogen peroxide (H2O2). APEX2 can be used as an alternative to traditional antibody labeling in EM, with a protein localization throughout the depth of the entire cell. In other words, the APEX2-tagged protein can be visualized by specific osmication11 without immunogold labeling and permeabilization after ultra-cryosectioning. Horseradish peroxidase (HRP) is also a sensitive tag that catalyzes the H2O2-dependent polymerization of DAB into a localized precipitate, providing EM contrast after treatment with OsO4. The ER target peptide sequence HRP-KDEL (lys-asp-glu-leu)12 was applied to visualize ER within a whole cell. To evaluate our protocol of utilizing genetic tags and en bloc staining with reduced osmium and TCH (rOTO method), using the osmication effect at the same time, we compared the membrane contrast with and without the use of each genetic tag in rOTO en bloc staining. Although 3DEM with array tomography and DAB staining with APEX and HRP have, respectively, been utilized for other purposes, our protocol is unique because we have combined array tomography for 3DEM and DAB staining for mitochondria and ER labeling. Specifically, we showed five cells with and without APEX-tagged genes in the same section, which aided in investigating the effect of the genetic modification on cells.

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		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glutaraldehyde</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">16200</td>
			<td style="width:349px;">Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Paraformaldehyde</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">19210</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Sodium cacodylate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">12300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Osmium tetroxide 4 % aqueous solution</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">16320</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Epon 812</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">14120</td>
			<td>EMbed 812- 20 ml/ DDSA- 16 ml/ NMA- 8 ml/ DMP-30 - 0.8 ml</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Ultra-microtome Leica ARTOS 3D</td>
			<td style="width:123px;">Leica</td>
			<td style="width:123px;">ARTOS 3D</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Uranyl acetate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">22400</td>
			<td>Hazardous chemical</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Lead citrate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">17900</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">35mm Gridded coverslip dish</td>
			<td style="width:123px;">Mattek</td>
			<td style="width:123px;">P35G-1.5-14-CGRD</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glow discharger</td>
			<td style="width:123px;">Pelco</td>
			<td style="width:123px;">easiGlow</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Formvar carbon coated Copper Grid</td>
			<td style="width:123px;">Ted Pella</td>
			<td style="width:123px;">01805-F</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Hydrochloric acid</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">258148</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fugene HD</td>
			<td style="width:123px;">Promega</td>
			<td style="width:123px;">E2311</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glycine</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">G8898</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:275px;">3,3&#8242; -diaminobenzamidine (DAB)</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">D8001</td>
			<td>Hazardous chemical</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">30% Hydrogen peroxide solution</td>
			<td style="width:123px;">Merck</td>
			<td style="width:123px;">107210</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium hexacyanoferrate(II) trihydrate</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">P3289</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">0.22 um syringe filter</td>
			<td style="width:123px;">Sartorius</td>
			<td style="width:123px;">16534</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Thiocarbonyldihydrazide</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">223220</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Potassium hydroxide</td>
			<td style="width:123px;">Fluka</td>
			<td style="width:123px;">10193426</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">L-aspartic acid</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">A9256</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Ethanol</td>
			<td style="width:123px;">Merck</td>
			<td style="width:123px;">100983</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Transmission electron microscopy</td>
			<td style="width:123px;">FEI</td>
			<td style="width:123px;">Tecnai G2</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">Indium tin oxide (ITO) coated glass coverslips</td>
			<td style="width:123px;">SPI</td>
			<td style="width:123px;">06489-AB</td>
			<td>fragil glass</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Isopropanol</td>
			<td style="width:123px;">Fisher Bioreagents</td>
			<td style="width:123px;">BP2618-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Diamond knife</td>
			<td style="width:123px;">Leica</td>
			<td style="width:123px;">AT-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Inveted light microscopy</td>
			<td style="width:123px;">Nikon</td>
			<td style="width:123px;">ECLipse TS100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Scanning electron microscopy</td>
			<td style="width:123px;">Zeiss</td>
			<td style="width:123px;">Auriga</td>
			<td></td>
		</tr>
	</tbody>
</table>

mitochondria and endoplasmic reticulum imaging by correlative light and volume electron microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

Figure 1 describes the schedule and workflow for this protocol. The protocol requires 7 days; however, depending on the time spent on SEM imaging, this may increase. For cell transfection, the confluency of the cells should be controlled so as not to cover the bottom of the entire grid plate (Figure 1A). A high cell density could prevent the identification of the cell of interest during light microscope and EM observation. We used genetically tagged plasmids that expressed APEX2 and HRP to select efficiently transfected cells among the numerous cultured cells in the culture dish. We cultured HEK293T cells and confirmed the expression of SCO1-linked APEX2 (mitochondrial intermembrane space [IMS]) and HRP-conjugated KDEL (ER) in co-transfected cells. Under light microscopy, APEX2-transfected cells were stained a brown color, whereas cells without transfection remained unstained (Figure 1A and Figure 2). This allowed the identification of the transfected target cells, which were then used for correlative light-electron microscopy (CLEM), using DAB staining in a cultured cell population (Figure 3D-F and Figure 4B). It was helpful to mark the glass bottom (Figure 1B,C) to make it easy to identify the location of the target cell during the flat embedding step (Figure 1E,F). When the HEK293T cells were treated with an enhanced &#8220;double osmium&#8221; staining protocol (rOTO), whole cells were stained a dark color (Figure 1D). After removing the gridded coverslip from the culture dish, we identified the target location on the block surface under a stereo microscope (Figure 1G). We trimmed the cells in the ROI in a trapezoid shape, and the leading and trailing edges were made parallel (Figure 1H,I). To implement mitochondria and ER network reconstruction, we used a large diamond knife with a large water boat to serially section SCO1-APEX2 and HRP-KDEL-expressing HEK293T cells (Figure 1J-L). Serial 90-nm thick ribbon sections were successfully attached to the ITO-coated glass coverslip (Figure 1M), and the surface was coated with 10-nm thick carbon for observation via SEM (Figure 1N).
SCO1-APEX2 and HRP-KDEL proteins generate highly dense electronic signals derived from DAB conversion that are detectable in TEM (Figure 3) and SEM (Figure 4). The dark stain generated by SCO1-APEX2 was observed exclusively in the IMS and not in the matrix space of mitochondria (Figure 3D). Co-transfected cells (the left cell in Figure 3D,E) with both SCO1-APEX2 and HRP-KDEL plasmids expressed a highly dense electron signal in mitochondrial IMS and ER; however, we observed no ER staining in cells that were transfected only with SCO1-APEX2 (the right cell in Figure 3D,F). For serial images using SEM, first, we created an overview of the whole array image using the BSE detector over a large area (Figure 4A). Second, the ROI was placed in the first section and propagated to all other sections (Figure 4B). Finally, we visualized the ROI containing five target cells with 5-nm image pixels (Figure 4C). Zoomed-in images revealed detailed subcellular structures (Figure 4D) such as Golgi apparatus, mitochondria, nuclei, and ER. The serial images clearly showed that ER-mitochondria contacts were occurring on different z-planes (Supplemental Movie 2 and Supplemental Movie 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59750/59750fig1.jpg" /> 
Figure 1: Sample preparation workflow for SEM and TEM. (A) A culture dish containing gridded coverslips was seeded with cells and stained with DAB. (B,C) After DAB staining, a marker pen was used to mark the bottom of the glass where the target cells were located. (D) After OsO4 staining, the cells become a dark color. (E,F) Polymerase chain reaction (PCR) tubes (or any type of embedding capsules) were used to easily make an EM block that contained the marked positions. (E) Top view. (F) Bottom view. (G) A low-magnification stereo microscopy image of the surface of an EM block. (H) The ROI is in the middle of the flat surface. (I) A higher-magnification stereo microscopy image of the rectangular ROI. (J) ITO-coated glass coverslips (white asterisk) are inserted into the ribbon carrier (blue arrow), and the carrier is clamped by turning the handle clockwise. (K,L) The handle of the holder is carefully pushed, and the carrier is adjusted to position the edge of the ITO-coated glass coverslip close to the knife by sliding. (M) The ribbons are attached to the ITO-coated glass coverslips. (N) The ITO-coated glass coverslips are attached to the SEM stubs, and the residual glass is sealed with sticky carbon tape and coated with a 10-nm carbon thread layer. White asterisks indicate the ITO-coated glass coverslip, and black asterisks indicate the carbon tape. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/59750fig1large.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/59750/59750fig2.jpg" /> 
Figure 2: Inverted phase-contrast microscopy image of cultured HEK293T cells stained with DAB. (A) Staining of cells with DAB only (without OsO4 staining). White arrows indicate the unstained cells, and black arrows indicate DAB-stained cells. (B) Higher-magnification image of the ROI. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/59750fig2large.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/59750/59750fig3.jpg" /> 
Figure 3: TEM imaging of HEK293T cells exhibiting the targeted mitochondrial IMS (SCO1-APEX2) and ER (HRP-KDEL). (A-C) Untransfected HEK293T cells showing the double-membrane of mitochondria (M) and endoplasmic reticulum (ER). (D-F) APEX2 and HRP catalyze the polymerization of DAB into a local precipitate, which is subsequently stained with electron-dense OsO4. A dark contrast is apparent in the mitochondrial IMS (black arrowhead) and ER (black arrow); however, cells that were not transfected with HRP&#8211;KDEL exhibit unstained ER (white arrow). Scale bars: 1 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/59750fig3large.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/59750/59750fig4.jpg" /> 
Figure 4: Serial SEM imaging of HEK293T cells exhibiting the targeted mitochondrial IMS (SCO1-APEX2) and ER (HRP-KDEL). (A) Overview of the serial section ribbons observed using the BSE detector. (B) Correlation of low-magnification image (inset) with high-magnification BSE image (white dotted-line box indicates the ROI of DAB-stained cells). (C) High magnification of the ROI target cells with 5-nm image pixels. (D,E) A dark contrast is apparent in the mitochondrial IMS and ER but not the Golgi apparatus. (F-I) ER-mitochondria contacts (white arrow) occur on different z-planes. N, nucleus; M, mitochondria; ER, endoplasmic reticulum; G, Golgi apparatus. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/59750fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/59750/59750table1v2.jpg" /> 
Table 1: Solution recipes. 
<img alt="Movie 1" src="/files/ftp_upload/59750/59750movie1.jpg" /> 
Supplemental Movie 1: SEM image stack. Fiji13 with TrakEM14 software was used to align 91 images. The original aligned data set is 11 GB. To downsize the stack, resized and cropped image set was used. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/Supple_S1_stack.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 2" src="/files/ftp_upload/59750/59750movie2.jpg" /> 
Supplemental Movie 2: Cropped image stack. To visualize mitochondria, ER, and their contact sites in detail, images were cropped from original data set (5 nm/pixel). Scale bars: 1 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/Supple_S2_crop.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 3" src="/files/ftp_upload/59750/59750movie3.jpg" /> 
Supplemental Movie 3: 3D Reconstruction of mitochondria and ER. For 3D visualization, the contour of mitochondria and ER was segmented and visualized using IMOD15 software. Mitochondria were visualized as long tubular structures (red), and the ER networks (green) showed their complicated morphology. Yellow represented a large surface area of contact site between mitochondria and ER in different z-planes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59750/Supple_S3_3D.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy at JoVE.com

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

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Transmembrane Transport in Endoplasmic Reticulum and Peroxisomes

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12.8K Views.  Korea Brain Research Institute. We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Video: Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Single Particle Cryo-Electron Microscopy: From Sample to Structure

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The overall process involves i) vitrifying the specimen in a thin film supported on a cryoEM grid; ii) screening the specimen to assess particle distribution and ice quality; iii) if the grid is suitable, collecting a single particle dataset for analysis; and iv) image processing to yield an EM density map. In this protocol, an overview for each of these steps is provided, with a focus on the variables which a user can modify during the workflow and the troubleshooting of common issues. With remote microscope operation becoming standard in many facilities, variations on imaging protocols to assist users in efficient operation and imaging when physical access to the microscope is limited will be described.

Structure determination of macromolecular complexes using cryoEM has become routine for certain classes of proteins and complexes. Here, this pipeline is summarized (sample preparation, screening, data acquisition and processing) and readers are directed towards further detailed resources and variables that may be altered in the case of more challenging specimens.

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...