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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

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.

Abstract

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.

Introduction

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' 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 °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.

Protocol

1. Cell Culture and Transfection

  1. Seed HEK-293T cells on a 60 mm diameter or larger tissue culture dish and grow to 60%–90% confluence in a cell culture incubator at 37 °C and 5% CO2.
  2. Transfect cells with APEX2-tagged mammalian expression plasmids using transfection reagent (see Table of Materials) according to the manufacturer's directions.
  3. At 12–15 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.

2. Chemical Fixation and Peroxidase Reaction

  1. 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 °C. From this point until step 2.3.3, keep the sample and solutions on ice, and perform centrifugation at 4 °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.
  2. 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.
  3. Carry out the peroxidase reaction
    1. Prepare a fresh solution containing 1 mg/mL of 3,3'-diaminobenzidine tetrahydrochloride (DAB) in 0.1 M sodium cacodylate buffer. Dissolve the DAB by vigorous vortexing for 5–10 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.
    2. Wash pellet by resuspending in 3 mL of DAB solution followed by pelleting at 500 x g for 5 min.
    3. 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 15–45 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.
    4. Pellet the cells, then wash 2x for 5 min with 0.1 M sodium cacodylate buffer, followed by one wash in Dulbecco's modified Eagle's medium (DMEM) or cell media of choice.
  4. Resuspend the cell pellet in 500 µL 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.

3. High Pressure Freezing

  1. 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.
  2. 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 µL pipet tip.
  3. Aspirate 2–3 µL 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.
  4. 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.
  5. Inspect the temperature vs. time and pressure vs. time graphs to verify that the pressure reached 2100 bar and the temperature reached -196 °C within 200 ms, and both parameters remained steady for the 600 ms of measurement.
  6. Repeat steps 3.3 to 3.5 until the cell pellet has been used or the desired number of samples has been frozen.
  7. 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.

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.

  1. Fill the automated freeze substitution unit with LN2. Bring the temperature to -90 °C.
  2. Prepare FS Mix 1 and begin FS.
    1. 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.
    2. Place the FS Mix 1 vials and the cryo-vials containing the frozen cell pellets into the FS unit's sample chamber. Transfer the inner capsule containing the membrane carrier from the LN2 vial into the corresponding vial containing FS Mix 1.
    3. Start a FS protocol with its first step being 24 h at -90°C. After the 24 h, pause the FS, and wash the samples 3x for 5 min with acetone that has been cooled to -90 °C.
  3. 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.
    1. 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.
    2. 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 °C, followed by gradual warming to 0 °C over 12-18 h.
  4. Keep the temperature at 0 °C and wash 3x for 30 min with pre-cooled acetone from a freshly opened bottle.

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.

  1. 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's directions, and incubate samples in the following resin concentrations: 2%, 4%, and 8% for 2 h each at 0 °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.
  2. 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.
  3. Polymerize in an oven at 60 °C for 24-36 h.
    NOTE: The protocol can be paused after the polymerization.
  4. 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.
  5. 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 °C for 24–36 h.
    NOTE: The protocol can be paused after the polymerization.

6. Sectioning

  1. 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.
  2. Obtain 90 nm ultrathin sections of the cell pellet using a glass or diamond knife.
  3. 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.

7. TEM Imaging

  1. 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.
  2. 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–K) and lead post-stained samples (Figure 2A–H).
    1. 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.
    2. Float grids on a drop of Sato'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.
  3. 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.

Results

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...

Discussion

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...

Disclosures

The authors declare no conflict of interest.

Acknowledgements

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.

Materials

NameCompanyCatalog NumberComments
3,3'-Diaminobenzidine tetrahydrochloride hydrateSigma-AldrichD5637-1G
Acetone (Glass Distilled)Electron Microscopy Sciences10016
Beakers; Plastic, Disposable 120 ccElectron Microscopy Sciences60952
Bovine Serum AlbuminSigma-AldrichA7906-100G
Cryogenic Storage Vials, 2 mLVWR82050-168
Dulbecco's Modified Eagle's MediumCorning10-017-CV
Durcupan ACM Fluka, single component A, M epoxy resinSigma-Aldrich44611-500ML
Durcupan ACM Fluka, single component B, hardener 964Sigma-Aldrich44612-500ML
Durcupan ACM Fluka, single component C, accelerator 960 (DY 060)Sigma-Aldrich44613-100ML
Durcupan ACM Fluka,single component DSigma-Aldrich44614-100ML
Embedding mold, standard flat, 14 mm x 5 mm x 6 mmElectron Microscopy Sciences70901
Embedding mold, standard flat, 14 mm x 5 mm x 4 mmElectron Microscopy Sciences70900
Fetal Bovine Serum; Nu-Serum IV Growth Medium SupplementCorning355104
Glass Knife Boats, 6.4 mmElectron Microscopy Sciences71008
Glass KnifemakerLeica MicrosystemsEM KMR3
Glutaraldehyde 10% Aqueous SolutionElectron Microscopy Sciences16120
HEK 293 CellsATCCCRL-1573
High Pressure Freezer with Rapid Transfer SystemLeica MicrosystemsEM PACT2Archived Product Replaced by Leica EM ICE
Hydrogen Peroxide 30% SolutionFisher Scientific50-266-27
Lipofectamine 3000 Transfection ReagentThermoFisher ScientificL3000015
Membrane carrier for EM PACT2, 1.5 mm x 0.1 mmMager Scientific16707898
Osmium Tetroxide, crystallineElectron Microscopy Sciences19110
Phosphate Buffered Saline (PBS) 20X, Ultra Pure GradeVWR97062-950
Plastic Capsules for AFS/AFS2, 5 mm x 15 mmMager Scientific16702738
Slot grids, 2 x 1 mm copper with Formvar support filmElectron Microscopy SciencesFF2010-Cu
Sodium Cacodylate Buffer, 0.2 M, pH 7.4Electron Microscopy Sciences102090-962
Sodium Hydroxide, Pellet 500 G (ACS)Avantor Macron Fine Chemicals7708-10
Tannic AcidElectron Microscopy Sciences21710
Tissue Culture Dishes, Polystyrene, Sterile, Corning, 100 mmVWR25382-166
Ultra Glass Knife StripsElectron Microscopy Sciences71012
UltramicrotomeLeica MicrosystemsEM UC7
Uranyl Acetate DihydrateElectron Microscopy Sciences22400

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