Expansion Microscopy for Beginners: Visualizing Microtubules in Expanded Cultured HeLa Cells (original) (raw)

Curr Protoc Neurosci. 2020 Jun; 92(1): e96.

Chi Zhang,1 ,2 ,† Jeong Seuk Kang,3 ,† Shoh M. Asano,1 ,2 ,7 Ruixuan Gao,1 ,2 and Edward S. Boydencorresponding author1 ,2 ,4 ,5 ,6

Chi Zhang

1Media Lab, Massachusetts Institute of Technology (MIT), Cambridge Massachusetts

2McGovern Institute, MIT, Cambridge Massachusetts

Jeong Seuk Kang

3John A. Paulson School of Engineering and Applied Sciences, Harvard University, Cambridge Massachusetts

Shoh M. Asano

1Media Lab, Massachusetts Institute of Technology (MIT), Cambridge Massachusetts

2McGovern Institute, MIT, Cambridge Massachusetts

7Current address, Internal Medicine Research Unit, Pfizer Inc., Cambridge Massachusetts

Ruixuan Gao

1Media Lab, Massachusetts Institute of Technology (MIT), Cambridge Massachusetts

2McGovern Institute, MIT, Cambridge Massachusetts

Edward S. Boyden

1Media Lab, Massachusetts Institute of Technology (MIT), Cambridge Massachusetts

2McGovern Institute, MIT, Cambridge Massachusetts

4Department of Biological Engineering, MIT, Cambridge Massachusetts

5Department of Brain and Cognitive Sciences, MIT, Cambridge Massachusetts

6Koch Institute for Cancer Research, MIT, Cambridge Massachusetts

1Media Lab, Massachusetts Institute of Technology (MIT), Cambridge Massachusetts

2McGovern Institute, MIT, Cambridge Massachusetts

3John A. Paulson School of Engineering and Applied Sciences, Harvard University, Cambridge Massachusetts

4Department of Biological Engineering, MIT, Cambridge Massachusetts

5Department of Brain and Cognitive Sciences, MIT, Cambridge Massachusetts

6Koch Institute for Cancer Research, MIT, Cambridge Massachusetts

7Current address, Internal Medicine Research Unit, Pfizer Inc., Cambridge Massachusetts

corresponding authorCorresponding author.

†Contributed equally

This is an open access article under the terms of the http://creativecommons.org/licenses/by/4.0/ License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

Abstract

Expansion microscopy (ExM) is a technique that physically expands preserved cells and tissues before microscope imaging, so that conventional diffraction‐limited microscopes can perform nanoscale‐resolution imaging. In ExM, biomolecules or their markers are linked to a dense, swellable gel network synthesized throughout a specimen. Mechanical homogenization of the sample (e.g., by protease digestion) and the addition of water enable isotropic swelling of the gel, so that the relative positions of biomolecules are preserved. We previously presented ExM protocols for analyzing proteins and RNAs in cells and tissues. Here we describe a cookbook‐style ExM protocol for expanding cultured HeLa cells with immunostained microtubules, aimed to help newcomers familiarize themselves with the experimental setups and skills required to successfully perform ExM. Our aim is to help beginners, or students in a wet‐lab classroom setting, learn all the key steps of ExM. © 2020 The Authors.

Keywords: antibody, beginner, clearing, expansion microscopy, fluorescence microscopy, HeLa cells, hydrogel, imaging, immunocytochemistry, immunohistochemistry, microscopy, microtubules, super‐resolution microscopy

INTRODUCTION

The resolution of traditional optical microscopes is limited to a few hundred nanometers, a value set by the diffraction limit of light. As a result, nanoscale objects, such as immunostained microtubule filaments (estimated filament width ∼60 nm), are not clearly resolved when imaged using conventional, diffraction‐limited microscopes. Expansion microscopy (ExM) is an emerging technology that uses common lab equipment and commercially available, inexpensive chemicals to physically magnify biological specimens so that they can be imaged with nanoscale resolution under conventional, diffraction‐limited microscopes. During ExM, a swellable polymer network (or hydrogel) is uniformly synthesized throughout a biological specimen (with polymer threads winding their way around and between biomolecules, throughout and around cells). Biomolecules of interest (e.g., proteins, RNAs, and/or their markers) are anchored to the polymer network using commercially available crosslinking chemicals. Then, specimens undergo a process of softening, or mechanical homogenization (e.g., by proteolysis, or denaturation), followed by a process of isotropic three‐dimensional physical expansion when water is added, causing the polymer, and thus the specimen in which the polymer is embedded, to swell. As a result, biomolecules of interest are spatially separated from each other, and the effective resolution of the microscope is increased. For a recent review of the history of the field, with a broad survey of ExM methods and examples of applications to different parts of biology and medicine, please see Wassie, Zhao, & Boyden (2019).

After a typical ∼4.5× linear expansion (∼100× volumetric expansion) of a specimen, the effective resolution of a microscope objective lens with ∼300‐nm diffraction‐limited resolution becomes ∼70 nm (300 nm divided by ∼4.5). Since the basic concept behind ExM was established a few years ago (Chen, Tillberg, & Boyden, 2015), many different versions of ExM have been introduced by our and other labs. To name a few, protein‐retention forms of expansion microscopy (Chozinski et al., 2016; Gambarotto et al., 2019; Ku et al., 2016; Tillberg et al., 2016; Truckenbrodt et al., 2018) enable the expansion of cells and tissues labeled with standard fluorescent proteins and antibodies; expansion fluorescence in situ hybridization (ExFISH; Chen et al., 2016) allows the expansion and visualization of both proteins and RNAs; iterative expansion microscopy (iExM; Chang et al., 2017) allows repetitive expansion of the specimen through multiple rounds of polymerization and expansion (e.g., two rounds of expansion results in ∼4.5 × 4.5, or ∼20×, physical magnification); expansion pathology (ExPath) optimizes ExM for preserved human specimens (Zhao et al., 2017); and lattice light‐sheet microscopy applied to ExM‐processed specimens (Gao et al., 2019) enables fast imaging with nanoscale resolution and molecular contrast across large volumes.

We earlier described best‐practice, step‐by‐step ExM protocols for proExM and ExFISH for both cell culture and tissue specimens, as well as detailed procedures for handling and mounting expanded samples and for imaging them with confocal and light‐sheet microscopes (Asano et al., 2018). To complement these previous protocols, we here present a beginner‐friendly cookbook‐style protocol for expanding HeLa cells with immunostained microtubules using proExM. The idea here was to create a tutorial exercise that newcomers to ExM can try out, and that is simple enough to be practiced on one's own but comprehensive enough that all the core skills and practices of ExM will be explored in the process of completing the exercise. This exercise may be useful in wet‐lab classroom settings, and indeed we have used it in workshop settings for cohorts of a few dozen students, as it allows students to make rapid progress, even as they learn all the core steps of ExM and acquire hands‐on practice.

We chose to expand HeLa cells with immunostained microtubules in this exercise because microtubules are widely used as model cellular structures for super‐resolution microscopy, the improved resolution enabled by ExM can be immediately appreciated by a beginner performing the protocol, and there are established cell fixation methods to preserve microtubule structures for super‐resolution imaging (Bates, Huang, Dempsey, & Zhuang, 2007), well‐validated commercial antibodies for tubulins, and successful examples and expected outcomes for proExM of cultured cells with microtubules labeled (Tillberg et al., 2016). As befits the tutorial/exercise nature of this protocol, we choose to utilize cultured HeLa cells as the biological specimens because they are readily available from commercial sources (see the Basic Protocol), relatively easy to culture and maintain, and highly amenable to antibody staining after fixation and permeabilization.

In this 4‐day protocol (Fig. ​1), we first culture HeLa cells on round coverslips placed in 24‐well plates (Day 1), and then perform cell fixation, immunostaining, and protein anchoring (Day 2). Next, we subject the cells to gelation using an optimized chamber design to minimize the physical handling and transfer of the hydrogel‐embedded sample, and then use proteinase K digestion to soften, or mechanically homogenize, the hydrogel‐embedded cells (Day 3). Finally, we expand the sample by water dialysis, and then mount the sample on a poly‐L‐lysine‐coated glass plate before fluorescence microscope imaging (Day 4).

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Key steps in expanding fixed HeLa cells with immunostained microtubules. AcX, 6‐((acryloyl)amino)hexanoic acid, succinimidyl ester; ProK, proteinase K.

We have tested this protocol with many beginners, including in the context of a hands‐on ExM training workshop at a medical school, and all of the participants in the workshop, with no prior experience with ExM, were able to successfully reproduce the protocol. Thus, we recommend that users consider using this protocol to learn the essential experimental setups and skills of ExM before applying and optimizing other, more flexible but more advanced ExM protocols (Asano et al., 2018, and protocols posted at http://expansionmicroscopy.org) to analyze their biological systems of interest. At the time of the writing of the current paper, more than 150 experimental papers and preprints involving some form of ExM have been published, suggesting that the field is poised for rapid growth. We hope that this protocol paper will help beginners get going with nanoimaging, a previously equipment‐ and/or complexity‐rich enterprise, but one that can now be reduced to a simple set of chemical processing steps.

VISUALIZATION OF MICROTUBULES IN HeLa CELLS BY EXPANSION MICROSCOPY

Key steps involved in the expansion microscopy of fixed and immunostained HeLa cells are shown in Figure ​1, and step‐by‐step instructions are provided below. Note that all steps up to the gelation (step 31) are performed on cells cultured on 13‐mm‐diameter coverslips placed in wells of a 24‐well plate. All the measures described below are for one well of a 24‐well plate; the measures should be scaled up linearly when handling multiple wells. Because of the time‐sensitive nature of the gelation process (steps 29 and 30), we recommend that beginners perform this protocol working with no more than two wells simultaneously. Please refer to the Reagents and Solutions section for details on the preparation of reagents and solutions described in the protocol.

Materials

Day 1: Cell culture

IMPORTANT NOTE: Perform all cell culture steps in a biosafety cabinet to ensure sterility. Prewarm cell culture medium, trypsin‐EDTA solution, and DPBS (without calcium and magnesium) to 37°C before cell culture experiments. The cells need to be passaged or plated when a plate reaches ∼70% confluency. A detailed mammalian cell culture protocol can be found in Phelan (2007).

The estimated experimental time on Day 1 is 2 hr.

Passaging HeLa cells

Plating HeLa cells on coverslips

Day 2: Cell fixation, immunostaining, and AcX treatment

The following cell fixation and immunostaining steps are optimized for microtubules in HeLa cells and may not be applicable to other samples. See the introduction for references to other protocols for fixation and immunostaining of other types of specimens.

The estimated experimental time on Day 2 is 8 hr.

Day 3: Gelation and proteinase K digestion

The estimated experimental time on day 3 is 3 hr.

Rarely, during step 40, the cell culture coverslip might be pried off while the gel remains attached to the Parafilm (Fig, ​3, part 1b). In such cases, follow the steps below for sample processing.

  1. Pry off and remove the two spacer stacks, and then trim the gel, while on the Parafilm, using a razor blade into an asymmetric shape mirroring that shown in Figure ​3, part 1a (see Figure ​3, part 1b, for details).
  2. Use a razor blade to cut the Parafilm around the gel, and then transfer and immerse the gel together with the attached piece of Parafilm in the petri dish containing the ProK‐containing digestion solution, with the gel facing up and the Parafilm at the bottom.
  3. Keep the sample on an orbital shaker at 60 rpm at room temperature overnight (>6 hr) in the dark (e.g., by wrapping the dish with aluminum foil).

Day 4: Expansion, mounting, and imaging

The estimated experimental time on day 4 is 3 hr.

REAGENTS AND SOLUTIONS

AcX stock solution

AcX stock solution Amount Final concentration
AcX (Thermo Fisher, cat. no. A20770) 5 mg 10 mg/ml
Anhydrous DMSO (Thermo Fisher, cat. no. D12345) 500 μl
Total 500 μl

The AcX stock solution can be divided into 10‐ to 20‐μl aliquots and stored up to 2 months at −20°C. The aliquots should be stored in a sealed container with drying agents (e.g., Drierite) or in a desiccator to avoid degradation by hydration.

APS stock solution

APS stock solution Amount Stock solution concentration (g/100 ml solution)
APS (Thermo Fisher, cat. no. 17874) 1 g 10
Water 9 ml
Total 10 ml

Divide APS stock solution into 1‐ml aliquots and store up to 2 weeks at −20°C.

Cell culture medium

Cell culture medium Amount (ml) Final concentration
Dulbecco's Modified Eagle Medium (DMEM) 445
Heat‐inactivated fetal bovine serum (FBS) 50 10% (v/v)
10,000 U/ml penicillin‐streptomycin 5 100 U/ml

Store DMEM (Corning, cat. no. 10013CV) at 4°C per manufacturer's guidelines. Store heat‐inactivated FBS (Thermo Fisher, cat. no. A3840001) and penicillin‐streptomycin (Thermo Fisher, cat. no.15140122) at −20°C per manufacturer's guidelines. Thaw heat‐inactivated FBS and penicillin‐streptomycin before usage. Prepare the cell culture medium in a biosafety cabinet. Store the cell culture medium up to 12 months at 4°C.

Cytoskeleton extraction buffer

Cytoskeleton extraction buffer Stock solution concentration Amount (ml) Final concentration
Triton X‐100 5% (w/v) 4 0.5% (w/v)
1,4‐Piperazinediethanesulfonic acid (PIPES) 1 M (pH 7.0) 4 0.1 M
Ethylene glycol‐bis(2‐aminoethylether)‐_N,N,N′,N′_‐tetraacetic acid (EGTA) 10 mM 4 1 mM
Magnesium chloride (Thermo Fisher, cat. no. AM9530G) 1 M 0.04 1 mM
Water 27.96
Total 40

Prepare 20 ml of 5% (w/v) Triton X‐100 by dissolving 1 g Triton X‐100 (Sigma Aldrich, cat. no. T9284‐100ML) in 19 ml water. Prepare 20 ml of 1 M PIPES buffer by dissolving 6 g PIPES (Sigma Aldrich, cat. no. P6757) in 15 ml of water, adjusting pH to 7.0 with 5 M sodium hydroxide solution, and adding water to a total volume of 20 ml. Prepare 20 ml of 10 mM EGTA solution by dissolving 76 mg EGTA (Sigma Aldrich, cat. no. 03777) in 20 ml water. Store the Triton X‐100, PIPES, and EGTA stock solutions and the prepared cytoskeleton extraction buffer up to 6 months at room temperature.

Digestion buffer

Digestion buffer Stock solution concentration Amount (ml) Final concentration (/100 ml solution)
Triton X‐100 5% (w/v) 10 0.50 g
Ethylenediaminetetraacetic acid (EDTA; Thermo Fisher, cat. no. 15575020) 0.5 M, pH 8.0 0.2 0.2 ml (1 mM)
Tris(hydroxymethyl)aminomethane (Tris; Thermo Fisher, cat. no. AM9855G) 1 M, pH 8.0 5 5 ml (50 mM)
Sodium chloride 5 M 20 4.67 g (1 M)
Water 64.8
Total 100

Store digestion buffer up to 2 weeks at room temperature. For long‐term storage, store 10‐ml aliquots of digestion buffer up to 12 months at −20°C.

Microtubule fixation solution

Microtubule fixation solution Amount (ml) Final concentration
16% (w/v) paraformaldehyde (Electron Microscopy Sciences, cat. no. 15710) 1.5 3% (w/v)
8% (w/v) glutaraldehyde (Electron Microscopy Sciences, cat. no. 16019) 0.1 0.1% (w/v)
10× PBS (Corning, cat. no. 21040CV)) 0.8 1×
Water 5.6
Total 8

Prepare microtubule fixation solution in a chemical fume hood. Use microtubule fixation solution when fresh, fixing cells immediately after making the solution.

Monomer solution (colloquially known as Stock X)

Monomer solution (“Stock X”) Stock solution concentration (g/100 ml solution) Amount (ml) Final concentration after addition of APS and TEMED stock solutions (g/100 ml solution)
Sodium acrylate (see vendors below) 38 (33% (w/w) in water) 2.25 8.6
Acrylamide (Sigma Aldrich, cat. no. A9099‐25G) 50 0.5 2.5
_N,N_′‐Methylenebisacrylamide (Sigma Aldrich, cat. no. 146072‐100G) 2 0.75 0.15
Sodium chloride (Thermo Fisher, cat. no. AM9759) 29.2 (5 M) 4 11.7 (2 M)
PBS 10× 1 1×
Water 1.3
Total 9.8

It takes time to fully dissolve sodium acrylate, acrylamide, and _N,N_′‐methylenebisacrylamide into water. Vortexing and sonication can help to speed up this process, but do not heat the solutions, because monomers may prematurely gel into polymer form at higher temperatures. Store the acrylamide and _N,N_′‐methylenebisacrylamide stock solutions up to 6 months at 4°C. The sodium acrylate stock solution must be used immediately after making. The Stock X solution can be divided into aliquots of 980 µl each and stored for at least a month at −20°C. Note: Sodium acrylate batches from different vendors, or from different lots, can vary in quality. Low‐quality sodium acrylate may not completely dissolve in water at the relatively high concentration of 33% (w/w), or may appear yellow or orange when dissolved in water. If the solution is cloudy, or appears yellow or orange, discard the solution and switch to a new bottle of sodium acrylate. Clear or slightly yellow solutions of sodium acrylate can be used in ExM experiments. We have recently tried out sodium acrylate from the vendors below, but we always recommend checking the quality through the aforementioned checks, because quality can vary over time, even from a given vendor.

Sodium acrylate vendor Cat. no.
Combi‐Blocks QC‐1489
AK Scientific R624
BLDpharm BD151354
Santa Cruz Biotechnology sc‐236893B
Fisher Scientific 50‐750‐9773

Quenching solution

Quenching solution Amount Final concentration
Glycine (Sigma Aldrich, cat. no. G7126‐100G) 750 mg 100 mM
10× PBS 10 ml 1×
Water 90 ml
Total 100 ml

Store quenching solution up to 12 months at 4°C.

Reduction solution

Reduction solution Amount Final concentration
Sodium borohydride (Sigma Aldrich, cat. no. 213462‐25G) 10 mg 0.1% (w/v)
PBS 10 ml 1×
Total 10 ml

The reduction solution must be made fresh before use. Store sodium borohydride at room temperature in a desiccated container. Sodium borohydride is reactive and should be handled with care. Weigh out 10 mg sodium borohydride into a 50‐ml tube, add 10 ml PBS, and pipet up and down ten times, using ∼5 ml of volume each time, to mix and dissolve the sodium borohydride. The dissolution of sodium borohydride in PBS will produce hydrogen gas bubbles; if there are no bubbles, the sodium borohydride solution has gone bad.

TEMED stock solution

TEMED stock solution Amount Stock solution concentration (g/100 ml solution)
TEMED (Thermo Fisher, cat. no. 17919) 1 g (1.29 ml) 10
Water 9 ml
Total 10 ml

Divide TEMED stock solution into 1‐ml aliquots and store at −20°C for up to 2 weeks.

COMMENTARY

Background Information

Optical microscopy is instrumental for studying biological structures in cells and tissues. Due to the diffraction of light by lenses, fluorophores within a few hundred nanometers (e.g., ∼300 nm) of each other are not resolvable when imaged with conventional, diffraction‐limited optical microscopes. Optical super‐resolution microscopy techniques (reviewed in Huang, Bates, & Zhuang, 2009) greatly improve the imaging resolution by separately detecting objects residing within a diffraction‐limited volume, so that their locations can be accurately determined; these techniques enable researchers to image many biological structures that are smaller than a few hundred nanometers, but they require specialized hardware.

Complementary to, but conceptually different from, optical super‐resolution microscopy techniques, we recently developed expansion microscopy (ExM, reviewed in Wassie et al., 2019) as a sample processing technique to physically enlarge preserved biological specimens. After ExM processing, biological objects within a diffraction‐limited volume are physically separated from one another and can thus be detected with conventional, diffraction‐limited microscopes. ExM realizes such physical separation by synthesizing a polyelectrolyte hydrogel throughout the sample, with biomolecules or their markers crosslinked to the hydrogel network. After softening the mechanical properties of the hydrogel‐embedded sample by protease digestion or biomolecule denaturation, the sample can be isotropically expanded via treatment with water.

ExM builds on discoveries going back several decades, including the study of the physics of polyelectrolyte hydrogels that swell vastly when immersed in water (Tanaka, Sun, Nishio, Swislow, & Shah, 1980), and the use of hydrogel embedding of biological specimens to improve staining and imaging (Hausen & Dreyer, 1981). In ExM, the hydrogel used is a swellable polyelectrolyte hydrogel, explicitly designed to be synthesized densely throughout a biological specimen (the polymer spacing may be as small as a few nanometers; Cohen, Ramon, Kopelman, & Mizrahi, 1992)); furthermore, the hydrogel is synthesized evenly throughout the sample, so that biomolecules that are crosslinked to the gel network can be separated isotropically in 3D, thus faithfully preserving their relative positions after expansion. In practice, the expansion process introduces errors of a few percent over length scales comparable to a microscope's field of view (Wassie et al., 2019), but these are negligible for the vast majority of biological questions currently being investigated, which are primarily concerned with the relative organization of molecules within cells and tissues.

We believe users can learn the principles of, and experimental skills for, the core processes of ExM through this exercise of visualizing immunostained microtubules in expanded HeLa cells. By practicing this protocol, users will get a head start on ExM and pave a smooth path toward using the evolving suite of ExM techniques (reviewed in Tillberg & Chen, 2019; Wassie et al., 2019; protocols in Asano et al., 2018, and posted at http://expansionmicroscopy.org) for their own applications.

Troubleshooting

Potential problems and solutions are listed in Table ​1.

Table 1

Troubleshooting the Protocol

Step Potential problem Solution
Steps 1‐10 Cells are dead (e.g., appearing as round, non‐adherent cells) or contaminated, or other problems associated with cell culture occur Learn and practice aseptic techniques. Please refer to Phelan, (2007) for detailed instructions on aseptic techniques. We also found video resources, such as Cell Culture Basics (https://www.thermofisher.com/us/en/home/references/gibco‐cell‐culture‐basics.html), to be educational and useful.
Steps 11‐14 Immunostained microtubules, before expansion, show discontinuous and/or distorted morphologies in step 24 Distorted or discontinuous morphologies of microtubules suggest failed fixation steps. We suggest that the users use the fixative vendors recommended in this protocol. Always use fresh fixatives from the ampules provided by the manufacturer; these ampules are intended for one‐time use after opening. The microtubule fixation solution (see recipe) must be used freshly and cannot be stored. Measure the pH of the PBS solution to confirm that the value is ∼pH 7.4. If the above troubleshooting attempts fail, discard all solutions and purchase new ones from the recommended vendors.
Steps 11‐28 Liquid aspiration from a 24‐well plate damages cells on the coverslip When aspirating liquid from a well, use one hand to hold the 24‐well plate and the other to hold the pipet. First push and hold the pipet plunger to dispense air out of the pipet tip so that the pipet is ready for drawing up liquid. Then, carefully move the pipet tip to the bottom edge of the well, and carefully tilt the plate ∼15° toward the pipet tip. Finally, slowly release the plunger to draw the liquid into the pipet tip. Do not aspirate the liquid directly from the cell layer, as that might damage the cells.
Cells dry out between steps To prevent drying of cells, we recommend that beginners simultaneously handle no more than two wells at a time, to minimize the time between liquid aspiration and addition steps.
Step 13 Immunostained microtubules, before expansion, show significant background fluorescence signals in step 24 The sodium borohydride reduction solution (see recipe), used to quench the fixatives and reduce background fluorescence, must be made fresh before use. Sodium borohydride, when fresh, should generate hydrogen gas bubbles when dissolved in PBS. If no hydrogen gas bubbles observed, use a new bottle of sodium borohydride.
Steps 15‐24 Immunostained microtubules, before expansion, show very weak immunostaining signals in step 24 Antibodies must be stored per manufacturer's instructions to ensure good quality. Secondary antibodies must be stored in the dark to prevent dye photobleaching. Use brightfield imaging to inspect the morphology of the cells to confirm successful fixation. If fixation is successful, the likely cause of weak immunostaining signals is a bad lot of antibodies. Replace both primary and secondary antibodies with newly purchased ones using the catalog numbers and vendors provided in this protocol.
Step 26 Immunostained microtubules show good imaging outcomes before expansion in step 24, but very weak or even no fluorescence signals after expansion in step 54 AcX, which crosslinks proteins to the hydrogel network, is an NHS ester and is prone to degradation by hydration. Therefore, AcX powder must be stored in a container with drying reagents (e.g., Drierite) at −20°C. AcX must be dissolved in anhydrous DMSO rather than regular DMSO, and the aliquoted AcX solution must be stored in a desiccating container at −20°C. Also, measure the pH of the PBS solution to confirm that the value is around pH 7.4, as high pH (> 9) will quickly hydrolyze AcX before it reacts with proteins. If the above troubleshooting attempts fail, discard all AcX solutions and purchase new materials from the recommended vendors.
Steps 29‐30 Gelation solution gels prematurely before step 37 The speed of polymerization significantly increases at higher temperature (e.g., at human body temperature of ∼37°C) or with high concentrations of APS and TEMED. To prevent premature gelation, first, all solutions (Stock X, APS, and TEMED) must be chilled on ice before mixing. APS, the radical polymerization initiator, must be added last. The gelation solution must be immediately placed back on ice after adding APS and vortexing. Keep the gelation solution on ice when transferring the solution to the 24‐well plate in step 30. The 24‐well plate must be kept on ice during the 5‐min incubation period in step 30. If the above instructions were strictly followed, but the gelation solution still prematurely gelled, it is possible that the concentration of the APS stock solution was mistakenly high. Make a fresh APS stock solution and redo the experiment.
Step 31 Coverslips or glass slides are broken during chamber construction Use forceps to carefully pick up and discard broken coverslips or glass slides into a sharp waste container. Be very careful and avoid getting wounded by the broken glass. Then, redo step 31 with new glass slides and coverslips.
Steps 32‐34 Cell culture coverslip dries before the chamber is filled with gelation solution in step 35 Steps 33 and 34 must be finished within 2 min so that the cells will not dry out. If the user finds these steps to be demanding, we suggest practicing them with clean cell culture coverslips placed in a clean 24‐well plate before performing the actual protocol.
Cell culture coverslip is broken Use forceps to carefully pick up and discard broken coverslips into a biological sharp waste container. Be very careful and avoid getting wounded by the broken glass. Then, redo the experiments.
Step 35 Air bubbles are trapped in the gelation chamber Use a pair of forceps to slowly lift one edge of the cell culture coverslip at a slight angle, until the air bubbles migrate to the edge of the coverslip and disperse, and then slowly place the cell culture coverslip back to rest on top of the two spacer stacks. Refill the chamber with gelation solution if necessary.
Step 40 Gel fails to form Check the incubator and make sure the temperature is at 37°C. If the temperature is correct, the most likely next reason for failed gelation is bad reagents. Prepare new Stock X, APS, and TEMED solutions (ordering fresh reagents if needed) and redo the experiments.
Step 44 Gel remains attached to the coverslip or the Parafilm after ProK digestion If the gel remains attached to the coverslip or Parafilm, it is most likely that the digestion step failed. Check the pH of the digestion buffer to make sure that it is ∼8.0. The ProK enzyme must be stored at −20°C. The gelled sample must be fully immersed in the ProK‐containing digestion buffer during the overnight digestion in step 43. If the above instructions were strictly followed, but the problem still persists, purchase fresh reagents and enzymes to make new ProK‐containing digestion solution.
Gel cannot be located or is lost during removal of the digestion buffer At this step, the gel is still relatively small and difficult to locate. First, we recommend illuminating the petri dish from different angles using a flashlight or a lamp to locate the gel. The gel will slightly scatter incident light and so should be visible when illuminated from different angles. Second, we recommend dispensing the aspirated digestion solution into another clean petri dish, rather than discarding it right away, so that if the gel is accidently drawn into the transfer pipet, it can possibly be recovered from the dispensed digestion solution. After the gel is secured, then discard the aspirated digestion solution appropriately.
Steps 44‐53 Gel is damaged Always use a wet paint brush to handle the gel if needed. Be slow and careful when handling the gel. The most likely cause of gel damage is aspiration or solution addition in steps 46, 47, 48, and 51. During these steps, first locate the gel using the instructions listed above, and then slowly aspirate solution from around the gel, avoiding touching the gel itself. Tilt the petri dish to move liquids to the side of the dish opposite to the gel, so as to avoid potential contact with the gel when aspirating liquids. If only a small part of the gel is damaged, use a clean razor blade to cut off the damaged part, provided that the remaining part is big enough for downstream processing and imaging.
Steps 46‐48 and 51 Gel cannot be located As the gel expands, it will be easier to locate. We recommend illuminating the petri dish from different angles using a flashlight or lamp to locate the gel. The gel will slightly scatter incident light and so should be visible when illuminated from different angles. Slightly tilt the petri dish at different angles to help with this process, as needed.
Step 53 Air bubbles are trapped between the bottom of the gel and the glass bottom of the 6‐well plate Use a wetted paintbrush to gently press the top of the gel to expel large air bubbles. In the case of small air bubbles that were not removed, image regions of the sample that do not include these air bubbles during microscopy. Mounting smaller gels (e.g., ∼10 mm × 10 mm, which is still sufficiently large for imaging) is less likely to introduce air bubbles.
Other Can I use this protocol for other types of specimens, for example, tissues? No, this protocol is designed and optimized for visualizing immunostained microtubules in expanded HeLa cells. Please refer to other, more flexible and comprehensive protocols (Asano et al., 2018, and protocols posted at http://expansionmicroscopy.org) for imaging of other types of specimens.
Can I store the expanded gel before mounting? Yes, the expanded gel can be stored in water in the petri dish for up to 1 week at 4°C, in the dark to avoid photobleaching. For long‐term storage, remove the water from the petri dish containing the expanded gel, add 10 ml PBS solution, and store the petri dish for up to a few months at 4°C. The gel will shrink in PBS and can be re‐expanded, mounted, and imaged by first removing the PBS solution and then following steps 46‐54 of the protocol.
Can I store the expanded and mounted gel? Yes, the expanded and mounted gel can be stored for up to 1 week at 4°C in the dark.
How should I properly dispose of used chemicals, gels, and solutions? Follow instructions on the Material Safety Data Sheet (MSDS) provided by the manufacturers. Consult the Environment, Health and Safety (EHS) office of your institution for questions related to chemical and biological waste disposal.

Understanding Results

When imaged under fluorescence microscopes, successfully fixed and immunostained HeLa cells will show bright, continuous, blurry lines indicating microtubule signals (Fig. ​4A). The background fluorescence should be low.

Because cultured HeLa cells are transparent, hydrogel‐embedded samples will appear transparent throughout the entire ExM process. After gelation, the sample will look like a thin layer of transparent gel attached to the cell culture coverslip or, rarely, to the Parafilm (Fig. ​3, part 1). After digestion with ProK, the sample will be detached from the coverslip or Parafilm and linearly expanded to ∼1.5 times its original size (Fig. ​3, part 3). Then, after expansion with water, the gel will linearly expand to ∼4.5 times its original size.

When imaged under fluorescence microscopes, expanded HeLa cells will show bright, continuous, sharp lines representing microtubule signals (Fig. ​4B). In expanded HeLa cells, immunostained microtubule filaments will be more visible, and resolved from one another, than those in HeLa cells before expansion (Fig. ​4A) when imaged using the same microscope settings. Because microtubules are structured throughout the entire cell volume, one might estimate the size of the expanded HeLa cell by imaging a single xy plane at the bottom of the cell. The expanded HeLa cell should have a size of ∼100‐600 µm.

Acknowledgments

We thank Paul Reginato, Timothy Su, and Patrick Flynn for helpful comments. For funding, we acknowledge Lisa Yang, John Doerr, the Open Philanthropy Project, NIH 1R01EB024261, the HHMI‐Simons Faculty Scholars Program, NIH grant 1R01MH110932, U.S. Army Research Laboratory and the U.S. Army Research Office under contract/grant number W911NF1510548, and NIH grants 1RM1HG008525, R011NS102727, U01MH114819, and 1U19MH114821.

Notes

Zhang, C. , Kang, J. S. , Asano, S. M. , Gao, R. , & Boyden, E. S. (2020). Expansion microscopy for beginners: Visualizing microtubules in expanded cultured HeLa cells. Current Protocols in Neuroscience, 92, e96. doi: 10.1002/cpns.96[PMC free article] [PubMed] [CrossRef] [Google Scholar]

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