Name: the cryoapex method for electron microscopy analysis of membrane protein localization within ultrastructurally-preserved cells
Uploaded: 2020-02-27T13:00:00
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 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 ...

This method combines the ability to stain for a membrane protein while also preserving membrane integrity and subcellular architecture. So it's really powerful for precisely localizing a membrane associated protein. The main advantage of this technique is that it combines the use of a robust genetically cloneable tag, APEX2, with state of the art methods in cellular preservation for electron microscopy, including cryofixation and freeze substitution.Together, these allow precise protein localization within a well-preserved cell. This protocol can be used to dissect mechanisms by which viruses replicate inside the host cell. Viruses often will hijack host proteins during their infection scheme.Viruses are the small genomes, which they utilize to replicate inside these host cells. Recently, we've been able to in unpublished data, label these proteins using this technology to understand how viruses replicate. This technique proved really critical in a recent collaborative study on a bacterial toxin that causes golgi breakdown.Using cryoAPEX, we were able to precisely localize the toxin with the fragmented golgi pieces, something that was really difficult to see otherwise using normal immunofluorescence techniques. Demonstrating this technique will be Elaine Mihelc, a graduate student, and research associate, Stephanie Angel, from our laboratories. After seeding HEK293 cells in the dish, transfect the cells with APEX2 tagged mammalian expression plasmids using transfection reagent according to the manufacturer's directions.At 12-15 hours post transfection, wash the cells once with PBS. Then, wash off the cells with PBS from the dish into a 15 milliliter conical tube. Centrifuge at 500 x g for five minutes.Carefully remove the supernatant and re-suspend the pellet in two milliliters of 2%glutaraldehyde and 0.1 molar sodium cacodylate buffer at pH 7.4 at room temperature. Place the sample on ice to incubate for 30 minutes. Then, pellet the sample at 500 x g for five minutes at four degrees Celsius.Wash the pellet three times for five minutes each with two milliliters of 0.1 molar sodium cacodylate buffer by centrifuging at four degrees Celsius. After centrifugation, remove the supernatant. To carry out the peroxidase reaction, wash the pellet by re-suspending in three milliliters of DAB solution followed by pelleting at 500 x g for five minutes.Then, re-suspend the pellet in three milliliters of DAB solution with 5.88 millimolar hydrogen peroxide. Incubate for 30 minutes at room temperature. The pellet becomes visibly brown-colored, indicating the presence of the insoluble DAB reaction product.After washing again with sodium cacodylate buffer in DMEM, according to the manuscript, re-suspend the cell pellet in 500 microliters of a cryoprotectant solution of DMEM containing 10%fetal bovine serum and 15%bovine serum albumin. Transfer the cell suspension to a 0.5 milliliter microcentrifuge tube. Pellet again, slightly increasing the centrifuge speed from 500 x g.Discard the majority of the supernatant, ensuring that enough liquid is left so that the pellet will not dry out. Wick away any remaining liquid from the cell pellet using the corner of a laboratory wipe or paper towel. Let enough liquid remain that the pellet forms a paste, similar in consistency to toothpaste.Aspirate two to three microliters 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. 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. Keeping the cartridges immersed in liquid nitrogen, remove the membrane carrier from its cartridge. Place into a plastic capsule and place the plastic capsule into a Cryovial full of liquid nitrogen.In a chemical hood, prepare one milliliter per sample of the solution of 0.2%weight by volume tannic acid and 5%DI water in acetone in a Cryovial. Place them into liquid nitrogen to freeze. Place the freeze substitution mix one vials and the Cryovials containing the frozen cell pellets into the freeze substitution units sample chamber.Transfer the inner capsule containing the membrane carrier from the liquid nitrogen vial into the corresponding vial containing freeze substitution mix one. Start a freeze substitution protocol at 90 degrees Celsius. After 24 hours, pause the freeze substitution and wash the samples three times for five minutes with acetone that has been cooled to 90 degrees Celsius.Next, in a chemical hood, prepare one milliliter per sample of a solution of 1%osmium tetroxide, 0.2%uranyl acetate, and 5%DI water in acetone in a Cryovial and place them in liquid nitrogen to freeze. Place the Cryovials with the freeze substitution mix two into the freeze substitution unit and transfer the capsules from the third acetone wash into the freeze substitution mix two vials. Incubate them and freeze substitution mix two for 72 hours at 90 degrees Celsius, followed by gradual warming to zero degrees Celsius over 12-18 hours.Keep the temperature at zero degrees Celsius and add pre-cooled acetone into the vials to wash three times for 10 minutes each. Prepare a resin mixture of choice in a plastic beaker according to the manufacturer's directions and add 2%4%and 8%of resin into the vials to immerse the capsules. Incubate for two hours each at zero degrees Celsius.Then, incubate in 15%30%60%90%and 100%resin components of A, B, and D only for four hours each at room temperature. After the 20 hours, prepare a mixture of resin components A, B, C, and D and incubate for four hours. Place the membrane carriers with cell pellet side up into flat embedding molds and fill with resin mixture A, B, C, and D.Place them in an oven to polymerize at 60 degrees Celsius for 24-36 hours.After polymerization, remove the blocks from the mold and place the sample in the vertical chuck of the ultra microtome 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.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 components A, B, C, and D.Polymerize at 60 degrees Celsius for 24-36 hours. 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 ultra microtome.Using a glass or diamond knife, trim the block into a trapezoidal shape closely surrounding the cell pellet. Obtain 90 nanometer ultra thin sections of the cell pellet using a glass or diamond knife. Pick up a ribbon of sections on a TEM grid.Dry the grid by blotting the edge on a piece of filter paper and store it in a TEM grid storage box. Mount the grid on the TEM holder and place the holder into the microscope. Use a Tecnai T12 at 80 kilovolts for screening cryoAPEX samples.Acquire images of cells and subcellular structures of interest with APEX2 labeling. In this study, preparation of the sample by traditional APEX methods resulted in clear labeling of organized, smooth endoplasmic reticulum structures. 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.The sample prepared by cryoAPEX also had clearly-defined labeling of organized, smooth endoplasmic reticulum structures, however, the membranes were smooth and parallel and little to no lipid extraction was seen. TEM analysis of 90 nanometer thin sections revealed that Huntington yeast interacting protein E was present throughout the peripheral endoplasmic reticulum as well as the nuclear envelope. Additionally, the Huntington yeast interacting protein E density was resolved into regularly-spaced foci along the luminal endoplasmic reticulum membrane.Huntington yeast interacting protein E 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. APEX2 labeling was performed using three cellular markers, of which mito-V5-APEX2 provided specific staining of mitochondria only and CAAX-APEX2 produced distinct staining of the plasma membrane only.No labeling was observed in intracellular organelles. Staining for the golgi lumen was also assessed using an Alpha MAN2-APEX2 marker as described in our original Sengupta et al manuscript. Following this procedure, three-dimensional methods such as electron tomography or focused ion beam scanning electron microscopy may be performed in order to investigate the 3D distribution of a protein of interest.Many of the chemicals used in this protocol are toxic, so before use, safety data sheets should be consulted to ensure that the chemicals are handled using proper personal protective equipment and disposed of according to institutional guidelines.

Research

JoVE Journal

Biology

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 ...

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

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 ...

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 ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

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 ...

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 ...

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 ...

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 ...

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, ...

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