The protocol described here stems from a publication by Sengupta et al., Journal of Cell Science, 132 (6), jcs222315 (2019)48. This work is supported by grants R01GM10092 (to S.M.) and AI081077 (R.V.S.) from the National Institutes of Health, CTSI-106564 (to S.M.) from Indiana Clinical and Translational Sciences Institute, and PI4D-209263 (to S.M.) from the Purdue University Institute for Inflammation, Immunology, and Infectious Disease.

1. Cell Culture and Transfection
<ol>
	<li>Seed HEK-293T cells on a 60 mm diameter or larger tissue culture dish and grow to 60%&#8211;90% confluence in a cell culture incubator at 37 &#176;C and 5% CO2.</li>
	<li>Transfect cells with APEX2-tagged mammalian expression plasmids using transfection reagent (see Table of Materials) according to the manufacturer&#39;s directions.</li>
	<li>At 12&#8211;15 h post-transfection, wash cells once with phosphate buffered saline (PBS). Remove cells from the dish by gentle washing with PBS. A dissociation reagent such as trypsin may be used if required for a given cell type. Centrifuge at 500 x g for 5 min to form a pellet.</li>
</ol>
2. Chemical Fixation and Peroxidase Reaction
<ol>
	<li>Carefully remove the supernatant and resuspend the pellet in 2 mL of 2% glutaraldehyde (v/v) in 0.1 M sodium cacodylate buffer, pH 7.4, at room temperature. Place sample on ice and incubate for 30 min. Pellet the sample at 500 x g for 5 min at 4 &#176;C. From this point until step 2.3.3, keep the sample and solutions on ice, and perform centrifugation at 4 &#176;C. 
	CAUTION: Both glutaraldehyde and sodium cacodylate buffer (containing arsenic) are toxic. Proper safety procedures and personal protective equipment should be used during handling. Solutions containing glutaraldehyde and/or sodium cacodylate buffer should be disposed of as hazardous chemical waste.</li>
	<li>Wash the pellet 3x for 5 min with 2 mL of 0.1 M sodium cacodylate buffer. For these as well as subsequent washes, gently resuspend the cell pellet in the required solution, then centrifuge for 5 min at 500 x g and carefully remove and discard the supernatant. Care should be taken with the repeated pelleting and resuspension steps, in order to minimize sample loss.</li>
	<li>Carry out the peroxidase reaction
	<ol>
		<li>Prepare a fresh solution containing 1 mg/mL of 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) in 0.1 M sodium cacodylate buffer. Dissolve the DAB by vigorous vortexing for 5&#8211;10 min. 
		CAUTION: DAB is toxic and a potential carcinogen and should be handled with proper safety procedures and personal protective equipment. Solutions containing DAB should be treated as hazardous chemical waste.</li>
		<li>Wash pellet by resuspending in 3 mL of DAB solution followed by pelleting at 500 x g for 5 min.</li>
		<li>Resuspend the pellet in 3 mL DAB solution to which hydrogen peroxide has been added to achieve a final concentration of 5.88 mM. Incubate for 30 min at room temp. The pellet becomes visibly brown-colored indicating the presence of the insoluble DAB reaction product. 
		NOTE: The DAB incubation time may need to be optimized for each sample. The color change can be monitored on the light microscope. In our experience, a 15&#8211;45 min incubation is adequate for most proteins. Hydrogen peroxide should be obtained from a freshly opened bottle or one that has been kept well-sealed after opening.</li>
		<li>Pellet the cells, then wash 2x for 5 min with 0.1 M sodium cacodylate buffer, followed by one wash in Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) or cell media of choice.</li>
	</ol>
	</li>
	<li>Resuspend the cell pellet in 500 &#181;L of a cryo-protectant solution of DMEM (or other cell media of choice) containing 10% fetal bovine serum and 15% bovine serum albumin. Pellet again, slightly increasing the centrifuge speed from 500 x g if required to achieve a pellet in the thick cryo-protectant solution. Discard the majority of the supernatant, ensuring that enough liquid is left so that the pellet will not dry out. Transport the cell pellet to the high pressure freezing instrument.</li>
</ol>
3. High Pressure Freezing
<ol>
	<li>Fill the high-pressure freezer reservoir with liquid nitrogen (LN2) and start the pump to fill the sample chamber with LN2. 
	CAUTION: Use proper safety procedures and personal protective equipment when working with liquid nitrogen. 
	NOTE: These steps are specific to the Leica EMPACT2 high pressure freezer.</li>
	<li>Wick away any remaining liquid from the cell pellet using the corner of a laboratory wipe or paper towel. Enough liquid should remain that the pellet forms a paste similar in consistency to toothpaste. It should be thin enough to be aspirated into a 20 &#181;L pipet tip.</li>
	<li>Aspirate 2&#8211;3 &#181;L of the cell pellet and deposit it onto a membrane carrier. Fill the well of the membrane carrier completely, so that the surface tension creates a slight dome on top, but the liquid does not spill out of the well. No air bubbles should be present.</li>
	<li>Slide the membrane carrier into the cartridge and secure. Place the cartridge into the HPF machine that has been prepped and primed, and press Start to freeze.</li>
	<li>Inspect the temperature vs. time and pressure vs. time graphs to verify that the pressure reached 2100 bar and the temperature reached -196 &#176;C within 200 ms, and both parameters remained steady for the 600 ms of measurement.</li>
	<li>Repeat steps 3.3 to 3.5 until the cell pellet has been used or the desired number of samples has been frozen.</li>
	<li>Keeping the cartridges immersed in LN2, remove each membrane carrier from its cartridge, place into a plastic capsule, and place the plastic capsule into a cryo-vial full of LN2. 
	NOTE: The protocol may be paused here. The cryo-vials with samples can be stored in a LN2 dewar in cryo-canes or cryo-boxes.</li>
</ol>
4. Freeze Substitution
CAUTION: Use proper safety procedures and personal protective equipment when working with liquid nitrogen. Additionally, many of the chemicals utilized in step 4 are toxic, including tannic acid, osmium tetroxide, and uranyl acetate. These chemicals must be handled according to proper safety procedures and disposed of as hazardous chemical waste.
<ol>
	<li>Fill the automated freeze substitution unit with LN2. Bring the temperature to -90 &#176;C.</li>
	<li>Prepare FS Mix 1 and begin FS.
	<ol>
		<li>In a chemical hood, prepare a solution of 0.2% tannic acid (w/v) and 5% DI water in acetone and aliquot 1 mL per sample into cryo-vials. Place into LN2 to freeze.</li>
		<li>Place the FS Mix 1 vials and the cryo-vials containing the frozen cell pellets into the FS unit&#39;s sample chamber. Transfer the inner capsule containing the membrane carrier from the LN2 vial into the corresponding vial containing FS Mix 1.</li>
		<li>Start a FS protocol with its first step being 24 h at -90&#176;C. After the 24 h, pause the FS, and wash the samples 3x for 5 min with acetone that has been cooled to -90 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare FS Mix 2 and complete FS 
	CAUTION: Osmium tetroxide is a highly toxic and oxidizing chemical that should only be handled by trained individuals according to established safety protocols. Protocols for the storage and disposal of osmium-containing solutions must be followed, as well as labeling of lab areas where osmium tetroxide is in use. Osmium tetroxide should be handled in a chemical hood with personal protective equipment including eye protection, a lab coat providing full arm protection, double Nitrile gloves, and an optional respirator.
	<ol>
		<li>In a chemical hood, prepare a solution of 1% osmium tetroxide, 0.2% uranyl acetate, and 5% DI water in acetone. Aliquot 1 mL per sample into cryo-vials and place in LN2 to freeze. 
		NOTE: Stock solutions of tannic acid (10% w/v in acetone), osmium tetroxide (10% w/v in acetone) and uranyl acetate (8% w/v in methanol) may be prepared and stored in cryo-vials in a LN2 dewar for ease of preparation of FS Mixes.</li>
		<li>Place the cryo-vials with FS Mix 2 into the FS unit and transfer the capsules from the third acetone wash into the FS Mix 2 vials. Incubate in FS Mix 2 for 72 h at -90 &#176;C, followed by gradual warming to 0 &#176;C over 12-18 h.</li>
	</ol>
	</li>
	<li>Keep the temperature at 0 &#176;C and wash 3x for 30 min with pre-cooled acetone from a freshly opened bottle.</li>
</ol>
5. Resin Infiltration and Embedding
CAUTION: The resin used here (see Table of Materials) is toxic prior to polymerization, and should be handled with proper safety procedures and personal protective equipment. Any unpolymerized resin should be disposed of as hazardous chemical waste.
<ol>
	<li>Infiltrate the samples with increasing concentrations of resin dissolved in acetone from a newly opened bottle. Prepare a mixture of resin components A, B, and D in a plastic beaker according to the manufacturer&#39;s directions, and incubate samples in the following resin concentrations: 2%, 4%, and 8% for 2 h each at 0 &#176;C. Incubate in 15%, 30%, 60%, 90%, and 100% resin for 4 h each at room temperature. Incubate for 4 h in a mixture of components A, B, C, and D.</li>
	<li>Place the membrane carriers with cell pellet side up into flat embedding molds and fill with resin (A, B, C, and D). Paper labels for the samples can be added to the wells at this time.</li>
	<li>Polymerize in an oven at 60 &#176;C for 24-36 h. 
	NOTE: The protocol can be paused after the polymerization.</li>
	<li>Remove the blocks from the mold and let cool. To remove the membrane carrier, first place the sample in the vertical chuck of the ultramicrotome where it can be visualized with magnification. Separate the membrane carrier from the block by a combination of dabbing liquid nitrogen on the membrane carrier to separate the metal from the plastic, and using a razor blade to chip away the resin around the membrane carrier. When separated, gently lift away the membrane carrier leaving the cell pellet dome on the face of the block.</li>
	<li>Place the block with the exposed cell pellet facing upward in a flat embedding mold that is slightly deeper than the first mold, and fill with resin. Polymerize at 60 &#176;C for 24&#8211;36 h. 
	NOTE: The protocol can be paused after the polymerization.</li>
</ol>
6. Sectioning
<ol>
	<li>Trim the block around the cell pellet using a razor blade. Then place the block in the sample chuck on the sectioning arm of an ultramicrotome. Using a glass or diamond knife, trim the block into a trapezoidal shape closely surrounding the cell pellet.</li>
	<li>Obtain 90 nm ultrathin sections of the cell pellet using a glass or diamond knife.</li>
	<li>Pick up a ribbon of sections on a TEM grid. Formvar-coated copper slot grids (1 x 2 mm2 slot) are useful for imaging serial sections. Dry the grid by blotting the edge on a piece of filter paper, and store in a TEM grid storage box. 
	NOTE: The protocol can be paused after sectioning.</li>
</ol>
7. TEM Imaging
<ol>
	<li>Mount the grid on the TEM holder and place into the microscope. We routinely use a Tecnai T12 at 80 kV for screening cryoAPEX samples. Acquire images of cells and subcellular structures of interest with APEX2 labeling.</li>
	<li>If desired, obtain additional membrane contrast by the use of lead post-staining. See Figure 2 for comparison of non-post-stained samples (Figure 2I&#8211;K) and lead post-stained samples (Figure 2A&#8211;H).
	<ol>
		<li>Float dry grids section-side down on a drop of dilute sodium chloride solution (~1.5 mM), 2x for 1 min each, then 1x for 10 min.</li>
		<li>Float grids on a drop of Sato&#39;s lead solution for 1 min. Wash by floating on sodium chloride solution 3x for 1 min, then on DI water 3x for 1 min. Blot excess liquid from the grids and store in a grid box. 
		CAUTION: Lead is a toxic chemical and should be handled with proper safety procedures and personal protective equipment. Solutions containing lead should be disposed of as hazardous chemical waste.</li>
	</ol>
	</li>
	<li>Image post-stained samples on the TEM. 
	NOTE: In traditional sample preparation for TEM, the lead contrasting step is performed prior to TEM imaging. However, it is recommended that for cryoAPEX samples, imaging is carried out first on non-contrasted samples. This ensures that the signal from the tag can be easily located by its strong contrast with the more lightly stained cellular structures. For many samples, no further staining will be required; however, if additional membrane contrast is desired, lead post-staining can be performed (Step 7.2) and the sample re-imaged.</li>
</ol>

The authors declare no conflict of interest.

The cryoAPEX protocol presented here provides a robust method to characterize the localization of membrane proteins within the cellular environment. Not only does the use of a genetically encoded APEX2 tag provide precise localization of a protein of interest, but the use of cryofixation and low-temperature dehydration provides excellent preservation and staining of the surrounding cellular ultrastructure. Combined, these approaches are a powerful tool for localizing a protein with high precision within its subcellular c...

Biological studies often include questions of resolving subcellular protein localization within cells and organelles. Immunofluorescence microscopy provides a useful low-resolution view of protein localization, and recent advances in super-resolution imaging are pushing the bounds of resolution for fluorescently tagged proteins1,2,3. However, electron microscopy (EM) remains the gold standard for imaging high-resolution cellular ultrastructure, though the labeling of proteins is a challenge.
Historically, several EM methods have been used to approach questions of ultrastructural protein localization. One of the most commonly utilized methods is immunoelectron microscopy (IEM), where antigen-specific primary antibodies are used to detect the protein of interest. EM signal is generated by the application of secondary antibodies conjugated with electron-dense particles, most commonly colloidal gold4,5. Alternately, antibodies conjugated with enzymes such as horse radish peroxidase (HRP) can be used to produce an electron-dense precipitate6,7,8. Two main approaches exist for IEM, termed pre-embedding and post-embedding labeling. In pre-embedding IEM, antibodies are introduced directly into cells, which necessitates light fixation and permeabilization of the cells9,10,11. Both steps can damage ultrastructure12,13. Development of significantly smaller antibodies consisting of an antibody Fab fragment conjugated with 1.4 nm nanogold allows very gentle permeabilization conditions to be used; however, nanogold is too small for direct visualization under TEM and requires additional enhancement steps to become visible14,15,16. In post-embedding IEM, antibodies are applied on thin sections of cells which have been fully processed by fixation, dehydration, and embedding in resin17. While this approach avoids the permeabilization step, preserving the epitope of interest throughout sample preparation is challenging18,19,20. The Tokuyasu method of light fixation followed by freezing, cryo-sectioning, and antibody detection provides improved epitope preservation21,22. However, the technical requirements of cryo-ultramicrotomy, as well as the sub-optimal contrast achieved in the cell, are disadvantages23.
The use of genetically encoded tags eliminates many of the difficulties of IEM related to detection of the protein of interest. A variety of tags are available, including HRP, ferritin, ReAsH, miniSOG, and metallothionein24,25,26,27,28,29,30,31,32. Each of these has advantages over previous methods, but each also has drawbacks preventing widespread use. These drawbacks range from inactivity of HRP in the cytosol to the large size of the ferritin tag, light sensitivity of ReAsH, and small size and lack of compatibility with cellular staining of metallothionein. Recently, a protein derived from ascorbate peroxidase has been engineered as an EM tag, named APEX233,34. As a peroxidase, APEX2 can catalyze the oxidation of 3,3&#39; diaminobenzidine (DAB), producing a precipitate that reacts with osmium tetroxide to provide local EM contrast with minimal diffusion from the protein of interest (less than 25 nm)33,35. Unlike traditional HRP-based methods, APEX2 is extremely stable and remains active in all cellular compartments33. Samples can be processed for TEM using traditional EM sample staining and methods that allow good visualization of the surrounding structures33,34,36. Because of its small size, stability, and versatility, APEX2 has emerged as an EM tag with great potential.
Many of the approaches discussed above either cannot be or have not yet been combined with the current state of the art in ultrastructural preservation, cryofixation and low-temperature freeze-substitution. Thus, they suffer from a lack of membrane preservation and/or cell staining to determine accurate protein localization. This necessarily limits the resolution and interpretation of the data that can be obtained. Cryofixation by high pressure freezing (HPF) involves rapid freezing of samples in liquid nitrogen at a high pressure (~2,100 bar), which causes vitrification rather than crystallization of aqueous samples, thus preserving cells in a near-native state37,38,39. HPF is followed by freeze substitution (FS), a low temperature (-90 &#176;C) dehydration in acetone combined with incubation with typical EM stains such as osmium tetroxide and uranyl acetate. HPF and FS together provide a distinct advantage over traditional chemical fixation (a longer process which can lead to artefacts) and alcohol dehydration at room temperature or on ice (which can lead to extraction of lipids and sugars), and thus are desirable to combine with the best EM tags for protein detection.
One reason that HPF/FS has not been combined with APEX2 labeling is that light chemical fixation is a prerequisite for the peroxidase reaction, limiting the diffusion of the DAB reaction product. In APEX2 studies thus far, fixation and peroxidase reaction are followed by traditional EM methods for staining and alcohol dehydration33,36. However, it has been shown that following chemical fixation with HPF/FS provides a distinct advantage in preservation over traditional chemical fixation and alcohol dehydration alone40. The loss of ultrastructural integrity seen in traditional TEM samples appears less connected to fixation than to dehydration, which is typically done using alcohol at room temperature or on ice, and can lead to extraction of lipids and sugars40,41. To develop the cryoAPEX method, we hypothesized that chemical fixation and peroxidase reaction, followed by HPF and FS, would produce an optimal result in terms of ultrastructural preservation.
Here we present the cryoAPEX protocol, which combines APEX2 tagging with cryofixation and freeze substitution methods (Figure 1). This straightforward protocol consists of transfection of an APEX2-tagged protein of interest, chemical fixation of cells, and the peroxidase reaction. HPF and FS are then performed followed by typical resin embedding and thin sectioning. TEM imaging reveals excellent preservation of ultrastructure using this method. Additionally, high-resolution subcellular localization and spatial distribution of an endoplasmic reticulum (ER) lumenal protein were observed. This method is widely useful for detection of membrane protein localization within cells for electron microscopy analysis. In our hands, the method has worked successfully for a variety of cell lines grown in tissue culture, including HEK-293T (human embryonic kidney), HeLa (human cervical cancer), Cos7 (African green monkey kidney fibroblast), and BHK (baby hamster kidney). Detailed instructions are described below using HEK-293T cells.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3,3&#39;-Diaminobenzidine tetrahydrochloride hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>D5637-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone (Glass Distilled)</td>
			<td>Electron Microscopy Sciences</td>
			<td>10016</td>
			<td></td>
		</tr>
		<tr>
			<td>Beakers; Plastic, Disposable 120 cc</td>
			<td>Electron Microscopy Sciences</td>
			<td>60952</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A7906-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryogenic Storage Vials, 2 mL</td>
			<td>VWR</td>
			<td>82050-168</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium</td>
			<td>Corning</td>
			<td>10-017-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Durcupan ACM Fluka, single component A, M epoxy resin</td>
			<td>Sigma-Aldrich</td>
			<td>44611-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Durcupan ACM Fluka, single component B, hardener 964</td>
			<td>Sigma-Aldrich</td>
			<td>44612-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Durcupan ACM Fluka, single component C, accelerator 960 (DY 060)</td>
			<td>Sigma-Aldrich</td>
			<td>44613-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Durcupan ACM Fluka,single component D</td>
			<td>Sigma-Aldrich</td>
			<td>44614-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding mold, standard flat, 14 mm x 5 mm x 6 mm</td>
			<td>Electron Microscopy Sciences</td>
			<td>70901</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding mold, standard flat, 14 mm x 5 mm x 4 mm</td>
			<td>Electron Microscopy Sciences</td>
			<td>70900</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum; Nu-Serum IV Growth Medium Supplement</td>
			<td>Corning</td>
			<td>355104</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Knife Boats, 6.4 mm</td>
			<td>Electron Microscopy Sciences</td>
			<td>71008</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Knifemaker</td>
			<td>Leica Microsystems</td>
			<td>EM KMR3</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde 10% Aqueous Solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>16120</td>
			<td></td>
		</tr>
		<tr>
			<td>HEK 293 Cells</td>
			<td>ATCC</td>
			<td>CRL-1573</td>
			<td></td>
		</tr>
		<tr>
			<td>High Pressure Freezer with Rapid Transfer System</td>
			<td>Leica Microsystems</td>
			<td>EM PACT2</td>
			<td>Archived Product Replaced by Leica EM ICE</td>
		</tr>
		<tr>
			<td>Hydrogen Peroxide 30% Solution</td>
			<td>Fisher Scientific</td>
			<td>50-266-27</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 3000 Transfection Reagent</td>
			<td>ThermoFisher Scientific</td>
			<td>L3000015</td>
			<td></td>
		</tr>
		<tr>
			<td>Membrane carrier for EM PACT2, 1.5 mm x 0.1 mm</td>
			<td>Mager Scientific</td>
			<td>16707898</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium Tetroxide, crystalline</td>
			<td>Electron Microscopy Sciences</td>
			<td>19110</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS) 20X, Ultra Pure Grade</td>
			<td>VWR</td>
			<td>97062-950</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic Capsules for AFS/AFS2, 5 mm x 15 mm</td>
			<td>Mager Scientific</td>
			<td>16702738</td>
			<td></td>
		</tr>
		<tr>
			<td>Slot grids, 2 x 1 mm copper with Formvar support film</td>
			<td>Electron Microscopy Sciences</td>
			<td>FF2010-Cu</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Cacodylate Buffer, 0.2 M, pH 7.4</td>
			<td>Electron Microscopy Sciences</td>
			<td>102090-962</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide, Pellet 500 G (ACS)</td>
			<td>Avantor Macron Fine Chemicals</td>
			<td>7708-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Tannic Acid</td>
			<td>Electron Microscopy Sciences</td>
			<td>21710</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Culture Dishes, Polystyrene, Sterile, Corning, 100 mm</td>
			<td>VWR</td>
			<td>25382-166</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra Glass Knife Strips</td>
			<td>Electron Microscopy Sciences</td>
			<td>71012</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome</td>
			<td>Leica Microsystems</td>
			<td>EM UC7</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl Acetate Dihydrate</td>
			<td>Electron Microscopy Sciences</td>
			<td>22400</td>
			<td></td>
		</tr>
	</tbody>
</table>

the cryoapex method for electron microscopy analysis of membrane protein localization within ultrastructurally-preserved cells

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

In order to compare the ultrastructural preservation using the cryoAPEX method with traditional fixation and dehydration, we prepared samples in which an endoplasmic reticulum membrane (ERM; ER membrane) peptide was tagged with APEX2 and transfected into HEK-293T cells. ERM-APEX2 localizes to the cytoplasmic face of the ER and remodels the ER structure into morphologically distinct structures known as organized smooth ER (OSER)34,42,4...

Watch this Scientific Journal Video about The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells at JoVE.com

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

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

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

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9.4K Views.  Purdue University. This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

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

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

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

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

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

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

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current state-of-the-art cryoET combined with subtomogram averaging analysis enables the high-resolution structural determination of macromolecular complexes that are present in multiple copies within tomographic reconstructions. Tomographic experiments usually require a vast amount of tilt series to be acquired by means of high-end transmission electron microscopes with important operational running-costs. Although the throughput and reliability of automated data acquisition routines have constantly improved over the recent years, the process of selecting regions of interest at which a tilt series will be acquired cannot be easily automated and it still relies on the user's manual input. Therefore, the set-up of a large-scale data collection session is a time-consuming procedure that can considerably reduce the remaining microscope time available for tilt series acquisition. Here, the protocol describes the recently developed implementations based on the SerialEM package and the PyEM software that significantly improve the time-efficiency of grid screening and large-scale tilt series data collection. The presented protocol illustrates how to use SerialEM scripting functionalities to fully automate grid mapping, grid square mapping, and tilt series acquisition. Furthermore, the protocol describes how to use PyEM to select additional acquisition targets in off-line mode after automated data collection is initiated. To illustrate this protocol, its application in the context of high-end data collection of Sars-Cov-2 tilt series is described. The presented pipeline is particularly suited to maximizing the time-efficiency of tomography experiments that require a careful selection of acquisition targets and at the same time a large amount of tilt series to be collected.

The increasing demand for large-scale data collection in cryogenic electron tomography requires high-throughput image acquisition routines. Described here is a protocol that implements the recent developments of advanced acquisition strategies aimed at maximizing the time-efficiency and throughput of tomographic data collection.

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current ...

Imaging Replicative Domains in Ultrastructurally Preserved Chromatin by Electron Tomography

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

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

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

Visualizing Subcellular Localization of a Protein in the Heart Using Quantum Dots-Mediated Immuno-Labeling Followed by Transmission Electron Microscopy

The subcellular localization is critical to delineating proper function and determining the molecular mechanisms of a particular protein. Several qualitative and quantitative techniques are used to determine the subcellular localization of proteins. One of the emerging techniques in determining the subcellular localization of a protein is quantum dots (QD)-mediated immunolabeling of a protein followed by imaging them with transmission electron microscopy (TEM). QD is a semiconductor nanocrystal with a dual property of crystalline structure and high electron density, which makes them applicable to electron microscopy. This current method visualized the subcellular localization of Sigma 1 receptor (Sigmar1) protein using QD-TEM in the heart tissue at ultrastructural level. Small cubes of the heart tissue sections from a wild-type mouse were fixed in 3% glutaraldehyde, subsequently osmicated, stained with uranyl acetate, followed by sequential dehydration with ethanol and acetone. These dehydrated heart tissue sections were embedded in low-viscosity epoxy resins, cut into thin sections of 500 nm thickness, put on the grid, and subsequently subjected to antigen unmasking with 5% sodium metaperiodate, followed by quenching of the residual aldehydes with glycine. The tissues were blocked, followed by sequential incubation in primary antibody, biotinylated secondary antibody, and streptavidin-conjugated QD. These stained sections were blot dried and imaged at high magnification using TEM. The QD-TEM technique allowed the visualization of Sigmar1 protein&#39;s&#160;subcellular localization at the ultrastructural level in the heart. These techniques can be used to visualize the presence of any protein and subcellular localization in any organ system.

The present protocol describes a method of immuno-labeling a protein in the heart tissue sections using quantum dots. This technique provides a useful tool to visualize any protein&#39;s subcellular localization and expression at the ultrastructural level.

The subcellular localization is critical to delineating proper function and determining the molecular mechanisms of a particular protein. Several ...

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 ...