Name: the cryoapex method for electron microscopy analysis of membrane protein localization within ultrastructurally-preserved cells
Uploaded: 2020-02-27T13:00:00.000+00:00
Duration: 11 min 45 s
Description: 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 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 context.
The crux of the advancement of this method is the fact that the loss of ultrastructure experienced after preparation by traditional TEM methods comes primarily from the dehydration step rather than the fixation step40. It was previously believed that peroxidase-based methods were incompatible with HPF/FS because they require chemical fixation prior to the peroxidase reaction. To work around this, a protocol named CryoCHEM was recently developed in which samples are initially cryofixed, followed by rehydration and the peroxidase reaction49. This approach provides excellent target localization with significant improvements in sample staining and preservation. It has been shown to be useful for tissue samples and in cases where correlative fluorescence and electron microscopy is desired. In parallel to cryoCHEM, our method combines glutaraldehyde fixation with HPF and FS. CryoAPEX offers a streamlined protocol that works effectively even for small cellular samples.
Access to high pressure freezing and freeze substitution instruments is crucial to the cryoAPEX method. These instruments and skills are increasingly common in EM facilities. Even if HPF and FS equipment are not readily available, the chemically fixed sample is stable enough for a short amount of time to be transported modest distances40. We have found that samples can be stored after the DAB reaction at 4 &#176;C for at least 48 hours prior to HPF without significant loss of quality. Another critical aspect of the cryoAPEX protocol is the inclusion of controls, which are essential for a robust experiment and convincing results. Samples prepared by transient transfection with efficiency less than 100% will contain negative control cells as well as labeled cells within the same sample. If using cell lines with stable expression of APEX2, a separate negative control should be prepared by transfection of cells with the non-APEX2 labeled protein of interest. Several constructs that can serve as organellar controls are available through Addgene, and published images are available in this and other publications that can be used for verification33,34,36,48. In-depth discussion of experimental design and verification of new APEX2 fusion constructs has been provided by Martell et al.36
While cryoAPEX is broadly useful for the detection of membrane proteins, some limitations exist. Although APEX2 is a small 28 kDa protein, some proteins may not be able to incorporate the tag33,34. APEX2 is not considered useful for labeling soluble proteins in the cytosol, due to the diffuse reaction product33,36. Additionally, the detection of small quantities of protein poses a challenge due to the presence of staining in the surrounding cell. Preparation by HPF and FS preserves cellular components which are extracted by traditional fixation and dehydration. This leads to overall darker staining in the cell, potentially competing with low levels of APEX2 labeling.
The cryoAPEX technique is widely applicable to many proteins, with a limited number of steps that may require optimization. First, due to individual variability among proteins, the protein expression level and/or DAB reaction time may need to be adjusted in order for the signal to be visualized above the background staining of the cell. Helpful information and protocols for validation of new APEX2 fusion constructs and optimization of the expression and DAB staining are provided by Martell et al.36 From a cellular staining perspective, the FS protocol and/or chemicals may need to be adjusted for optimal visualization of different organelle membranes, within different cell types, tissues, or organisms50,51,52,53,54. In our experience, the FS conditions presented here have worked well for a variety of mammalian cell lines.
The hybrid approach of cryoAPEX has the potential to be applied to many other genetic labeling techniques. Replacing traditional alcohol dehydration with HPF/FS is expected to greatly improve the ultrastructural preservation and the protein localization information. Utilizing sapphire discs as a cell substrate to fix cells as a monolayer improves the preservation of the cell periphery, including the cytoskeleton and cell-cell contacts. Minor modifications to the protocol would be required to use sapphire discs. APEX technology can be used to detect green fluorescent protein (GFP) tagged proteins via a GFP-binding peptide35. This indirect method of detection opens up the potential to utilize APEX technology for the myriad proteins already tagged with GFP. The recently-introduced split APEX2 will be advantageous for proximity and interaction studies55. Additionally, existing HRP-based methods can be combined with HPF/FS to improve cellular preservation. One example is fluorescent indicator and peroxidase for precipitation with EM resolution (FLIPPER), in which individual cell markers have been fused with both a fluorescent tag and HRP, providing lumenal markers for Golgi or ER56. Use of improved peroxidase substrates in place of DAB is also possible with this method, including substrates which are optimized for RNA labeling 57. CryoAPEX also provides in-cell labeling and ultrastructural preservation necessary for three dimensional analysis of protein distribution through electron tomography, and potentially at high volumes through SBF-SEM or FIB-SEM48,58.
Overall, CryoAPEX is a robust method with wide applicability. In principle, it can be applied to any membrane protein, whether within the lumenal space of an organelle, on the cytoplasmic face, within vesicles, on the cell&#39;s plasma membrane or even in the extracellular space. For this vast range of membrane proteins, the cryoAPEX method provides the potential to see the localization and distribution of a protein with accuracy within its subcellular context.

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><tbody><tr><td>3,3&#039;-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&#039;s Modified Eagle&#039;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)34,42,43. OSER morphology includes regions of smooth, parallel, densely stacked membranes which serve as an optimal region to compare ultrastructural preservation. Preparation of the sample by traditional APEX methods resulted in clear labeling of OSER structures (Figure 2A&#8211;D). Upon inspection at high magnification, the stacked membranes appeared ruffled and non-uniform gaps were present between concentric membrane densities, indicating poor membrane preservation and lipid extraction (Figure 2D). The sample prepared by cryoAPEX also had clearly defined labeling of OSER structures; however, the membranes were smooth and parallel, and little to no lipid extraction was seen (Figure 2E&#8211;H). The results from cryoAPEX were of similar high quality preservation to those obtained from a sample which underwent HPF/FS without the additional chemical fixation and APEX2/DAB reaction steps (Figure 2I&#8211;K).
In addition to visually appreciable membrane preservation, the cryoAPEX method preserves the protein of interest such that aspects of protein distribution patterns may be observed in some cases. To illustrate this point, we used another ER-localized protein, huntingtin yeast interacting protein E (HYPE). HYPE is a membrane protein located on the luminal face of the ER membrane 44,45,46,47. HYPE-APEX2 constructs were overexpressed in HEK-293T cells. TEM analysis of 90 nm thin sections revealed that HYPE was present throughout the peripheral ER as well as the nuclear envelope (Figure 3A,B). Additionally, the HYPE density was able to be resolved into regularly spaced foci along the lumenal ER membrane (Figure 3C, arrows). HYPE distribution and foci were also visible in a sample prepared with traditional fixation and dehydration; however, extensive membrane disruption and extraction were present, making the sample suboptimal (Figure 3D,E).
To demonstrate the robust organellar specificity and applicability of the cryoAPEX method for a range of tagged proteins, we performed APEX2 labeling using three cellular markers. Mitochondria were labeled using mito-V5-APEX234. This marker of the mitochondrial matrix provided specific staining of mitochondria only (Figure 4A). Likewise, we assessed plasma membrane labeling using CAAX-APEX234, which produced distinct staining of the plasma membrane only (Figure 4C). No labeling was observed in intracellular organelles (Figure 4C). Additionally, we created a new construct as a marker for the Golgi lumen by fusing the first 118 amino acids of the mouse isoform of &#945;-mannosidase with the APEX2 gene48. The resulting MannII-APEX2 was transiently transfected into cells which were subsequently prepared by the cryoAPEX method. Stained Golgi stacks were clearly distinguishable from the surrounding organelles (Figure 4B). Individual stacks, cisternae, and some vesicles were labeled, typical of Golgi staining (Figure 4B). Altogether, these markers demonstrate that the cryoAPEX method provides specific labeling of membrane proteins within various organelles at high enough resolution to distinguish them from surrounding sub-cellular structures.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60677/60677fig1.jpg" /> 
Figure 1: Schematic of the essential steps in the CryoAPEX protocol. (A) Cells are grown and transfected with an APEX2 plasmid. (B) Cells are pelleted and fixed with glutaraldehyde, followed by (C) incubation with DAB and hydrogen peroxide to produce the peroxidase reaction product. (D) The pellet is cryofixed by HPF, (E) freeze substituted with heavy metals and acetone, and (F) embedded in resin. Thin sections are collected on the microtome. (G) TEM imaging is performed and additional contrast may be added by post-staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60677/60677fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60677/60677fig2.jpg" /> 
Figure 2: Comparison of OSER membrane preservation using traditional chemical fixation, cryoAPEX, and HPF/FS. The reorganized ER morphology in chemically fixed, DAB reacted ERM&#8211;APEX2-expressing cells that were processed via traditional chemical fixation and alcohol dehydration (A&#8211;D) or by cryoAPEX (E&#8211;H) was compared to ERM&#8211;APEX2 expressing cells that were cryofixed live and without the DAB reaction (I&#8211;K). The live cryofixed cells represent the best attainable ultrastructural preservation and serve here as the metric for evaluating membrane preservation obtained via the two APEX-based detection protocols (A&#8211;H). The evenly-spaced parallel lamellar stacking of the ER derived membranes obtained by cryoAPEX (exemplified in panels G and H), as opposed to the ruffled membranes obtained by traditional methods (panels C and D), highlights the superior membrane preservation obtained by cryoAPEX. This figure has been modified from Sengupta et al. 201948. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60677/60677fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60677/60677fig3.jpg" /> 
Figure 3: Protein localization of an APEX2-tagged ER membrane protein can be resolved into periodic foci. (A) An image of a thin section of a HEK-293T cell expressing HYPE&#8211;APEX2 and processed by cryoAPEX reveals staining of the ER in a well-preserved (dense) cytoplasmic background (B,C). Higher magnification images of a small section of the peripheral ER (demarcated by yellow box in A and shown in B, with further magnification of red box in B shown in C) exhibits periodic foci of APEX2-generated density (B, red box and C, white arrowheads showing periodicity between the HYPE foci). (D) Image of a thin section of a cell expressing HYPE-APEX2 and prepared by traditional chemical fixation and dehydration shows specific staining of the cortical ER and the nuclear envelope (red arrows). (E) At a higher magnification, periodic HYPE-specific foci were apparent within stretches of the ER (yellow box and white arrow heads in the inset), despite extensive membrane disruption, indicated by red arrows. NE = nuclear envelope. This figure has been modified from Sengupta et al. 201948. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60677/60677fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60677/60677fig4.jpg" /> 
Figure 4: Organelle markers show specificity of the signal obtained from APEX2-tagged proteins. APEX2-tagged protein constructs designed to localize to the mitochondrial matrix (mito-V5-APEX2; shown in A), or the Golgi lumen (&#945;-mannII-APEX2; shown in B), or the plasma membrane (CAAX-APEX2; shown in C) were transiently expressed in HEK293 cells and the samples processed by cryoAPEX. Each construct yielded organelle specific densities. Magnified views of two sections (yellow or red boxes) from the cells expressing &#945;-mannIIAPEX2 (panel B) or CAAX-APEX2 (panel C) are shown. This figure has been modified from Sengupta et al. 201948. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60677/60677fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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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Nanyang Technological University

Research

JoVE Journal

Biology

9.5K Views.  Purdue University. Deze methode combineert de mogelijkheid om vlekken voor een membraan eiwit, terwijl ook het behoud van membraan integriteit en subcellulaire architectuur. Dus het is echt krachtig voor het precies lokaliseren van een membraan geassocieerd eiwit. Het belangrijkste voordeel van deze techniek is dat het het gebruik van een robuuste genetisch kloonbare tag, APEX2, combineert met state-of-the-art methoden in cellulaire bewaring voor elektronenmicroscopie, inclusief cryofixatie en vriessubstitutie....

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

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

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

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

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

Biochemistry

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate neurotransmission that occurs within synapses because it is highly regulated by dynamic protein-protein interactions and phosphorylation events. Accordingly, when any method is used to study synaptic transmission, a major goal is to preserve these transient physiological modifications. Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments. In other words, the protocol describes a biochemical methodology to carry out protein enrichment from presynaptic, postsynaptic, and extrasynaptic compartments. First, synaptosomes, or synaptic terminals, are obtained from neurons that contain all synaptic compartments by means of a discontinuous sucrose gradient. Of note, the quality of this initial synaptic membrane preparation is critical. Subsequently, the isolation of the different subsynaptic compartments is achieved with light solubilization using mild detergents at differential pH conditions. This allows for separation by gradient and isopycnic centrifugations. Finally, protein enrichment at the different subsynaptic compartments (i.e., pre-, post- and extrasynaptic membrane fractions) is validated by means of immunoblot analysis using well-characterized synaptic protein markers (i.e., SNAP-25, PSD-95, and synaptophysin, respectively), thus enabling a direct assessment of the synaptic distribution of any particular neuronal protein.

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments.

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate ...

Expression and Purification of the Human Lipid-sensitive Cation Channel TRPC3 for Structural Determination by Single-particle Cryo-electron Microscopy

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium homeostasis, particularly store-operated calcium entry, which is critical to maintaining proper function of synaptic vesicle release and intracellular signaling pathways. Accordingly, TRPC channels have been implicated in a variety of human diseases including cardiovascular disorders such as cardiac hypertrophy, neurodegenerative disorders such as Parkinson&#39;s disease, and neurologic disorders such as spinocerebellar ataxia. Therefore, TRPC channels represent a potential pharmacologic target in human diseases. However, the molecular mechanisms of gating in these channels are still unclear. The difficulty in obtaining large quantities of stable, homogeneous, and purified protein has been a limiting factor in structure determination studies, particularly for mammalian membrane proteins such as the TRPC ion channels. Here, we present a protocol for the large-scale expression of mammalian ion channel membrane proteins using a modified baculovirus gene transfer system and the purification of these proteins by affinity and size-exclusion chromatography. We further present a protocol to collect single-particle cryo-electron microscopy images from purified protein and to use these images to determine the protein structure. Structure determination is a powerful method for understanding the mechanisms of gating and function in ion channels.

This protocol describes techniques used to determine ion channel structures by cryo-electron microscopy, including a baculovirus system used to efficiently express genes in mammalian cells with minimum effort and toxicity, protein extraction, purification, and quality checking, sample grid preparation and screening, as well as data collection and processing.

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...

Assessing Two-dimensional Crystallization Trials of Small Membrane Proteins for Structural Biology Studies by Electron Crystallography

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray crystallography to study structure-function questions of membrane proteins, as well as soluble proteins. Screening for two-dimensional (2D) crystals by transmission electron microscopy (EM) is the critical step in finding, optimizing, and selecting samples for high-resolution data collection by cryo-EM. Here we describe the fundamental steps in identifying both large and ordered, as well as small 2D arrays, that can potentially supply critical information for optimization of crystallization conditions.
By working with different magnifications at the EM, data on a range of critical parameters is obtained. Lower magnification supplies valuable data on the morphology and membrane size. At higher magnifications, possible order and 2D crystal dimensions are determined. In this context, it is described how CCD cameras and online-Fourier Transforms are used at higher magnifications to assess proteoliposomes for order and size.
While 2D crystals of membrane proteins are most commonly grown by reconstitution by dialysis, the screening technique is equally applicable for crystals produced with the help of monolayers, native 2D crystals, and ordered arrays of soluble proteins. In addition, the methods described here are applicable to the screening for 2D crystals of even smaller as well as larger membrane proteins, where smaller proteins require the same amount of care in identification as our examples and the lattice of larger proteins might be more easily identifiable at earlier stages of the screening.

Evaluating two-dimensional (2D) crystallization trials for the formation of ordered membrane protein arrays is a highly critical and difficult task in electron crystallography. Here we describe our approach in screening for and identifying 2D crystals of predominantly small membrane proteins in the range of 15 &#x2013; 90kDa.

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray ...

Optimizing Sample Preparation for Cryogenic Electron Microscopy

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native conditions, suspended in vitreous ice. This technique allows for the high-resolution visualization of proteins and other biomolecules without the need for crystallization, offering significant insights into their function and mechanism. Recent advancements in single-particle analysis, coupled with improved computational data processing, have made cryo-EM an indispensable tool in modern structural biology. Despite its growing adoption, cryo-EM faces persistent challenges that can limit its effectiveness, particularly uneven particle distribution. This issue often leads to poor resolution and reduced accuracy in reconstructed protein structures. This article outlines a simple, practical approach to address this challenge, using the small heat-shock protein from Methanocaldococcus jannaschii (MjsHSP16.5) as an example. The method optimizes sample preparation to minimize preferential adsorption, ensuring more homogeneous particle distribution and higher-quality protein cryo-EM structures. This technique offers valuable guidance for researchers aiming to overcome similar challenges in structural studies.

This protocol presents a practical method using Methanocaldococcus jannaschii (MjsHSP16.5) as an example to address uneven particle distribution through sample preparation optimization, providing a reference for researchers to efficiently elucidate macromolecular structures using cryogenic electron microscopy (cryo-EM).

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native ...

Do's and Don'ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state by transmission electron microscopy (TEM). This can be achieved with very low quantity of protein and in the buffer of choice, without the use of any stain, which is very useful to determine structure-function correlations of macromolecules. When combined with single-particle image processing, the technique has found widespread usefulness for 3D structural determination of purified macromolecules. 
The protocol presented here explains how to perform cryoEM and examines the causes of most commonly encountered problems for rational troubleshooting; following all these steps should lead to acquisition of high quality cryoEM images. The technique requires access to the electron microscope instrument and to a vitrification device. Knowledge of the 3D reconstruction concepts and software is also needed for computerized image processing. Importantly, high quality results depend on finding the right purification conditions leading to a uniform population of structurally intact macromolecules. 
The ability of cryoEM to visualize macromolecules combined with the versatility of single particle image processing has proven very successful for structural determination of large proteins and macromolecular machines in their near-native state, identification of their multiple components by 3D difference mapping, and creation of pseudo-atomic structures by docking of x-ray structures. The relentless development of cryoEM instrumentation and image processing techniques for the last 30 years has resulted in the possibility to generate de novo 3D reconstructions at atomic resolution level.

This paper describes the cryo-electron microscopy methodology, which is used to obtain high quality microscopic images of macromolecules in their near-native state. The method yields images suitable for further computerized processing using the single particle approach, devised to generate the 3D reconstruction of a macromolecule.

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein using the profiles of fluorescence intensity across the plasma membrane. Measured fluorescence profiles are fitted by a model for membrane and cytoplasm fluorescence distribution along a line applied perpendicularly to the cell periphery. This model is constructed from the fluorescence intensity values in reference cells expressing a fluorescently-tagged marker for cytoplasm and with FM 4-64-labeled plasma membrane. The method can be applied to various cell types and organisms; however, only plasma membranes of non-neighboring cells can be evaluated. This fast microscopy-based method is suitable for experiments, where subtle and dynamic changes of plasma membrane-associated markers are expected and need to be quantified, e.g., in the analysis of mutant versions of proteins, inhibitor treatments, and signal transduction observations. The method is implemented in a multi-platform R package that is coupled with an ImageJ macro that serves as a user-friendly interface.

Here, we present a protocol to perform a quantitative analysis of the level of plasma-membrane association for fluorescently-tagged peripherally-associated protein. The method is based on the computational decomposition of membrane and cytoplasmic component of signal observed in cells labeled with plasma membrane fluorescent marker.

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein ...

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

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