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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<li>Kijanka, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+immuno-gold+labeling+protocol+for+nanobody-based+detection+of+HER2+in+breast+cancer+cells+using+immuno-electron+microscopy%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">A novel immuno-gold labeling protocol for nanobody-based detection of HER2 in breast cancer cells using immuno-electron microscopy.</a> Journal of Structural Biology. 199 (1), 1-11 (2017).</li>
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<li>Hua, Y., Laserstein, P., Helmstaedter, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Large-volume+en-bloc+staining+for+electron+microscopy-based+connectomics%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Large-volume en-bloc staining for electron microscopy-based connectomics.</a> Nature Communications. 6, 7923(2015).</li>
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<li>Martell, J. D., Deerinck, T. J., Lam, S. S., Ellisman, M. H., Ting, A. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electron+microscopy+using+the+genetically+encoded+APEX2+tag+in+cultured+mammalian+cells%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">Electron microscopy using the genetically encoded APEX2 tag in cultured mammalian cells.</a> Nature Protocols. 12 (9), 1792-1816 (2017).</li>
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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 sh...

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.

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

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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12.9K 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

Low-Cost Cryo-Light Microscopy Stage Fabrication for Correlated Light/Electron Microscopy

The coupling of cryo-light microscopy (cryo-LM) and cryo-electron microscopy (cryo-EM) poses a number of advantages for understanding cellular 
dynamics and ultrastructure. First, cells can be imaged in a near native environment for both techniques. Second, due to the vitrification process, samples are
 preserved by rapid physical immobilization rather than slow chemical fixation. Third, imaging the same sample with both cryo-LM and cryo-EM provides correlation of
 data from a single cell, rather than a comparison of "representative samples". While these benefits are well known from prior studies, the widespread use of correlative 
 cryo-LM and cryo-EM remains limited due to the expense and complexity of buying or building a suitable cryogenic light microscopy stage. Here we demonstrate the 
 assembly, and use of an inexpensive cryogenic stage that can be fabricated in any lab for less than $40 with parts found at local hardware and grocery stores. This 
 cryo-LM stage is designed for use with reflected light microscopes that are fitted with long working distance air objectives. For correlative cryo-LM and cryo-EM 
 studies, we adapt the use of carbon coated standard 3-mm cryo-EM grids as specimen supports. After adsorbing the sample to the grid, previously established protocols 
 for vitrifying the sample and transferring/handling the grid are followed to permit multi-technique imaging. As a result, this setup allows any laboratory with a 
 reflected light microscope to have access to direct correlative imaging of frozen hydrated samples.

We demonstrate the fabrication of a low-cost cryogenic stage designed to fit most reflected light microscopes. This lab-built cryogenic stage enables efficient and reliable correlative imaging between cryo-light and cryo-electron microscopy.

The coupling of cryo-light microscopy (cryo-LM) and cryo-electron microscopy (cryo-EM) poses a number of advantages for understanding cellular 
dynamics ...

Bioengineering

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

Correlative Light- and Electron Microscopy Using Quantum Dot Nanoparticles

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using widefield fluorescence light microscopy and transmission electron microscopy (TEM). To demonstrate the protocol we have immunolabeled ultrathin epoxy sections of human somatostatinoma tumor using a primary antibody to somatostatin, followed by a biotinylated secondary antibody and visualization with streptavidin conjugated 585 nm cadmium-selenium (CdSe) quantum dots (QDs). The sections are mounted on a TEM specimen grid then placed on a glass slide for observation by widefield fluorescence light microscopy. Light microscopy reveals 585 nm QD labeling as bright orange fluorescence forming a granular pattern within the tumor cell cytoplasm. At low to mid-range magnification by light microscopy the labeling pattern can be easily recognized and the level of non-specific or background labeling assessed. This is a critical step for subsequent interpretation of the immunolabeling pattern by TEM and evaluation of the morphological context. The same section is then blotted dry and viewed by TEM. QD probes are seen to be attached to amorphous material contained in individual secretory granules. Images are acquired from the same region of interest (ROI) seen by light microscopy for correlative analysis. Corresponding images from each modality may then be blended to overlay fluorescence data on TEM ultrastructure of the corresponding region.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of epoxy embedded human pathology tissue. We employ commercial antibody fragment conjugated QDs that are visualized by widefield fluorescence light microscopy and transmission electron microscopy.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using ...

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.

Correlative Light Electron Microscopy CLEM for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

A correlative light electron microscopy (CLEM) method is applied to image virus-induced intracellular structures via electron microscopy (EM) in cells that are previously selected by light microscopy (LM). LM and EM are combined as a hybrid imaging approach to achieve an integrated view of virus-host interactions.

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing ...

Correlative Light and Electron Microscopy (CLEM) as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity necessary for proper cellular function. Understanding how the enzymes, proteins, and cytoskeletal components govern and maintain this biochemical segregation is therefore of paramount importance. The use of fluorescently tagged molecules to localize to and/or perturb subcellular compartments has yielded a wealth of knowledge and advanced our understanding of cellular regulation. Imaging techniques such as fluorescent and confocal microscopy make ascertaining the position of a fluorescently tagged small molecule relatively straightforward, however the resolution of very small structures is limited 1.
On the other hand, electron microscopy has revealed details of subcellular morphology at very high resolution, but its static nature makes it difficult to measure highly dynamic processes with precision 2,3. Thus, the combination of light microscopy with electron microscopy of the same sample, termed Correlative Light and Electron Microscopy (CLEM) 4,5, affords the dual advantages of ultrafast fluorescent imaging with the high-resolution of electron microscopy 6. This powerful technique has been implemented to study many aspects of cell biology 5,7. Since its inception, this procedure has increased our ability to distinguish subcellular architectures and morphologies at high resolution. 
Here, we present a streamlined method for performing rapid microinjection followed by CLEM (Fig. 1). The microinjection CLEM procedure can be used to introduce specific quantities of small molecules and/or proteins directly into the eukaryotic cell cytoplasm and study the effects from millimeter to multi-nanometer resolution (Fig. 2). The technique is based on microinjecting cells grown on laser etched glass gridded coverslips affixed to the bottom of live cell dishes and imaging with both confocal fluorescent and electron microscopy. Localization of the cell(s) of interest is facilitated by the grid pattern, which is easily transferred, along with the cells of interest, to the Epon resin used for immobilization of samples and sectioning prior to electron microscopy analysis (Fig. 3). Overlay of fluorescent and EM images allows the user to determine the subcellular localization as well as any morphological and/or ultrastructural changes induced by the microinjected molecule of interest (Fig. 4). This technique is amenable to time points ranging from &le;5 s up to several hours, depending on the nature of the microinjected sample.

Correlative Light and Electron Microscopy CLEM as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

The CLEM technique has been adapted to analyze ultrastructural morphology of membranes, organelles, and subcellular structures affected by microinjected molecules. This method combines the powerful techniques of micromanipulation/microinjection, confocal fluorescent microscopy, and electron microscopy to allow millimeter to multi-nanometer resolution. This technique is amenable to a wide variety of applications.


The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity ...

Biology

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Study of Endoplasmic Reticulum and Mitochondria Interactions by In Situ Proximity Ligation Assay in Fixed Cells

Structural interactions between the endoplasmic reticular (ER) and mitochondrial membranes, in domains known as mitochondria-associated membranes (MAM), are crucial hubs for cellular signaling and cell fate. Particularly, these inter-organelle contact sites allow the transfer of calcium from the ER to mitochondria through the voltage-dependent anion channel (VDAC)/glucose-regulated protein 75 (GRP75)/inositol 1,4,5-triphosphate receptor (IP3R) calcium channeling complex. While this subcellular compartment is under intense investigation in both physiological and pathological conditions, no simple and sensitive method exists to quantify the endogenous amount of ER-mitochondria contact in cells. Similarly, MAMs are highly dynamic structures, and there is no suitable approach to follow modifications of ER-mitochondria interactions without protein overexpression. Here, we report an optimized protocol based on the use of an in situ proximity ligation assay to visualize and quantify endogenous ER-mitochondria interactions in fixed cells by using the close proximity between proteins of the outer mitochondrial membrane (VDAC1) and of the ER membrane (IP3R1) at the MAM interface. Similar in situ proximity ligation experiments can also be performed with the GRP75/IP3R1 and cyclophilin D/IP3R1 pairs of antibodies. This assay provides several advantages over other imaging procedures, as it is highly specific, sensitive, and suitable to multiple-condition testing. Therefore, the use of this in situ proximity ligation assay should be helpful to better understand the physiological regulations of ER-mitochondria interactions, as well as their role in pathological contexts.

Study of Endoplasmic Reticulum and Mitochondria Interactions by In Situ Proximity Ligation Assay in Fixed Cells

Here, we describe a procedure to visualize and quantify with high sensitivity the endogenous interactions between the endoplasmic reticulum and mitochondria in fixed cells. The protocol features an optimized in situ proximity ligation assay targeting the inositol 1,4,5-triphosphate receptor/glucose-regulated protein 75/voltage-dependent anion channel/cyclophilin D complex at the mitochondria-associated membrane interface.

Structural interactions between the endoplasmic reticular (ER) and mitochondrial membranes, in domains known as mitochondria-associated membranes (MAM), ...

Visualizing Retinal Synaptic Structures in 3D with Volume Electron Microscopy

Preparation of Retinal Samples for Volume Electron Microscopy

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level precision. Within the retina, various types of neurons establish synaptic connections in the inner and outer plexiform layers. While conventional EM techniques have yielded valuable insights into retinal subcellular organelles, their limitation lies in providing 2D image data, which can hinder accurate measurements. For instance, quantifying the size of three distinct synaptic vesicle pools, crucial for synaptic transmission, is challenging in 2D. Volume EM offers a solution by providing large-scale, high-resolution 3D data. It is worth noting that sample preparation is a critical step in Volume EM, significantly impacting image clarity and contrast. In this context, we outline a sample preparation protocol for the 3D reconstruction of photoreceptor axon terminals in the retina. This protocol includes three key steps: retina dissection and fixation, sample embedding processes, and selection of the area of interest.

Author Spotlight: Retinal Neuroscience Studies with Volume Electron Microscopy

This protocol describes the detailed steps for preparing retinal samples for volume electron microscopy, focusing on the structural features of retinal photoreceptor terminals.

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level ...

Neuroscience

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for structural analysis, light microscopy is essential for the visualization of dynamic organelles by target-specific labeling. The following protocol describes a method that combines dual-color correlative light and electron microscopy (CLEM) to observe the interactions between mitochondria and lysosomes. In this study, mitochondria were labeled with mEosEM (Mito-mEosEM) and lysosomes with TMEM192-V5-APEX2. The results obtained from CLEM images enable us to observe the changes in the interactions between mitochondria and lysosomes under external stress conditions. Treatment with bafilomycin (BFA), which inhibits lysosomal function, resulted in an increase in contact between mitochondria and lysosomes, leading to the formation of fragmented mitochondria trapped inside lysosomes. Conversely, treatment with U18666A, which inhibits cholesterol export from lysosomes, caused lysosomes to be surrounded by mitochondria, indicating a distinct form of interaction. This study presents an effective method for observing the interactions between mitochondria and lysosomes in fixed cells. Furthermore, CLEM imaging with dual-color probes offers a powerful tool for future investigations of organelle dynamics and their implications for cell function and pathology.

This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for ...

Ultrastructural Localization of Endogenous LC3 by On-Section Correlative Light-Electron Microscopy

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal details that are important for understanding the autophagic process. However, EM methods often lack molecular information, obstructing the correlation of ultrastructural information obtained by EM to fluorescence microscopy-based localization of specific autophagy proteins. Furthermore, the rarity of autophagosomes in unaltered cellular conditions hampers investigation by EM, which requires high magnification, and hence provides a limited field of view.
In answer to both challenges, an on-section correlative light-electron microscopy (CLEM) method based on fluorescent labeling was applied to correlate a common autophagosomal marker, LC3, to EM ultrastructure. The method was used to rapidly screen cells in fluorescence microscopy for LC3 labeling in combination with other relevant markers. Subsequently, the underlying ultrastructural features of selected LC3-labeled spots were identified by CLEM. The method was applied to starved cells without adding inhibitors of lysosomal acidification.
In these conditions, LC3 was found predominantly on autophagosomes and rarely in autolysosomes, in which LC3 is rapidly degraded. These data show both the feasibility and sensitivity of this approach, demonstrating that CLEM can be used to provide ultrastructural insights on LC3-mediated autophagy in native conditions-without drug treatments or genetic alterations. Overall, this method presents a valuable tool for ultrastructural localization studies of autophagy proteins and other scarce antigens by bridging light microscopy to EM data.

Here, we present a protocol for optimized on-section correlative light-electron microscopy based on endogenous, fluorescent labeling as a tool to investigate the localization of rare proteins in relation to cellular ultrastructure. The power of this approach is demonstrated by ultrastructural localization of endogenous LC3 in starved cells without Bafilomycin treatment.

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal ...

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

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Low-Cost Cryo-Light Microscopy Stage Fabrication for Correlated Light/Electron Microscopy

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Correlative Light- and Electron Microscopy Using Quantum Dot Nanoparticles

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Correlative Light and Electron Microscopy (CLEM) as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

Study of Endoplasmic Reticulum and Mitochondria Interactions by In Situ Proximity Ligation Assay in Fixed Cells

Preparation of Retinal Samples for Volume Electron Microscopy

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

Ultrastructural Localization of Endogenous LC3 by On-Section Correlative Light-Electron Microscopy