Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion

Summary

Nanoscale imaging of clinical tissue samples can improve understanding of disease pathogenesis. Expansion pathology (ExPath) is a version of expansion microscopy (ExM), modified for compatibility with standard clinical tissue samples, to explore the nanoscale configuration of biomolecules using conventional diffraction limited microscopes.

Abstract

In modern pathology, optical microscopy plays an important role in disease diagnosis by revealing microscopic structures of clinical specimens. However, the fundamental physical diffraction limit prevents interrogation of nanoscale anatomy and subtle pathological changes when using conventional optical imaging approaches. Here, we describe a simple and inexpensive protocol, called expansion pathology (ExPath), for nanoscale optical imaging of common types of clinical primary tissue specimens, including both fixed-frozen or formalin-fixed paraffin embedded (FFPE) tissue sections. This method circumvents the optical diffraction limit by chemically transforming the tissue samples into tissue-hydrogel hybrid and physically expanding them isotropically across multiple scales in pure water. Due to expansion, previously unresolvable molecules are separated and thus can be observed using a conventional optical microscope.

Introduction

Investigating the molecular organization of tissues in a three-dimensional (3D) context can provide new understanding of biological functions and disease development. However, these nanoscale environments are beyond the resolution capabilities of conventional diffraction limited microscopes (200−300 nm), where the minimal resolvable distance, d is defined by d α λ/NA. Here λ is the wavelength of light and NA is the numerical aperture (NA) of the imaging system. Recently, direct visualization of fluorescently labeled molecules has been made possible by newly developed super-resolution imaging techniques^1,2,3, including stimulated emission depletion (STED), photo-activated localization microscopy (PALM), stochastic optical reconstruction microscopy (STORM), and structured illumination microscopy (SIM). Although these imaging techniques have revolutionized understanding of biological function at the nanoscale, in practice, they often rely on expensive and/or specialized equipment and image processing steps, can have slower acquisition time comparing to conventional optical imaging, require fluorophores with specific characteristics (such as photo-switching capability and/or high photostability). In addition, it remains a challenge to perform 3D super-resolution imaging on tissue specimens.

Expansion microscopy (ExM), first introduced in 2015⁴, provides an alternative means of imaging nanoscale features (<70 nm) by physically expanding preserved samples embedded in a swellable polyelectrolyte hydrogel. Here, key biomolecules and/or labels are anchored in situ to a polymer network that can be isotopically expanded after chemical processing. Because the physical expansion increases the total effective resolution, molecules of interest can then be resolved using conventional diffraction-limited imaging systems. Since the publication of the original protocol, where custom synthesized fluorescent labels were anchored to the polymer network⁴, new strategies have been used to directly anchor proteins (protein retention ExM, or proExM)^5,6,7,8,9 and RNA^9,10,11,12 to the hydrogel, and increase physical magnification through iterative expansion¹³ or adapting gel chemistry^8,14,15.

Here we present an adapted version of proExM, called expansion pathology (ExPath)¹⁶, which has been optimized for clinical pathology formats. The protocol converts clinical samples, including formalin-fixed paraffin-embedded (FFPE), hematoxylin and eosin (H&E) stained, and fresh-frozen human tissue specimens mounted on glass slides, into a state compatible with ExM. Proteins are then anchored to the hydrogel and mechanical homogenization is performed (Figure 1)¹⁶. With a 4-fold linear expansion of the samples, multicolor super-resolution (~70 nm) images can be obtained using a conventional confocal microscope having only a ~300 nm resolution and can also be combined with other super-resolution imaging techniques.

Protocol

1. Preparation of Stock Reagents and Solutions

1. Prepare gelling solution components.
   NOTE: Solution concentrations are given in g/mL (w/v percent).
   1. Make the following stock solutions: 38% (w/v) sodium acrylate (SA), 50% (w/v) acrylamide (AA), 2% (w/v) N,N′-methylenebisacrylamide (Bis), and 29.2% (w/v) sodium chloride (NaCl). Dissolve the compounds in doubly deionized water (ddH₂O). Use the amounts in Table 1 as a reference; prepared solutions can be scaled up or down in volume as needed. For example, to make 10 mL of a 38% (w/v) SA solution, add 1.9 g SA to a graduated 10 mL cylinder and add ddH₂O to a volume of 5 mL.
   2. Prepare 9.4 mL of monomer solution at a 1.06x concentration as shown in Table 1.
      NOTE: This will result in a 1x concentration after addition of the initiator, accelerator, and inhibitor. The monomer stock can be stored at 4 °C for up to 3 months, or at -20 °C for long-term storage.
   3. Prepare the following stock solutions separately in ddH₂O: 0.5% (w/v) of the inhibitor 4-hydroxy-2,2,6,6-tetramethylpiperidin-1-oxyl (4HT), which inhibits gelation to enable diffusion of the gelling solution into tissues, 10% (v/v) of the initiator tetramethylethylenediamine (TEMED), which accelerates radical generation by ammonium persulfate (APS), and 10% (w/v) APS which initiates the gelling process.
      NOTE: Stock solutions of 4HT and TEMED can be prepared in 1 mL aliquots and stored at -20 °C for at least 6 months. APS has been found to lose efficacy after long-term storage and is best prepared in small quantities (<0.1 mL) immediately before gelling.
2. Prepare digestion buffer (50 mM Tris pH 8.0, 25 mM EDTA, 0.5% [w/v] nonionic surfactant, 0.8 M NaCl) by combining 25 mL of 1 M Tris pH 8 (3.03 g of Tris base in 25 mL of ddH₂O), 25 mL of EDTA (0.5 M pH 8), 2.25 g of nonionic surfactant, and 23.38 g of NaCl. Add ddH₂O for a total volume of 500 mL.
   NOTE: The solution can be scaled up or down as needed and be stored in at 4 °C. Proteinase K (ProK) will be added immediately before the digestion step.
3. Prepare 20 mM sodium citrate solution by combining 2.941 g of sodium citrate tribasic dihydrate with 500 mL of ddH₂O and adjusting pH to 8.0 at room temperature (RT). Scale the volume of stock as needed.
4. Prepare a stock solution of 6-((acryloyl)amino)hexanoic acid, succinimidyl ester (acryloyl-X, SE; AcX), the anchoring compound. Dissolve AcX in 500 µL of anhydrous dimethyl sulfoxide (DMSO) for a final concentration of 10 mg/mL.
   NOTE: The solution can be stored in a desiccated environment at -20 °C in 20 µL aliquots.
5. If not using commercially available buffers for immunostaining, prepare blocking buffer. Use a blocking buffer of 5% (v/v) normal animal serum and 0.1% (w/v) nonionic surfactant in 1x phosphate-buffered saline (PBS) and select the serum based on the host animal of the secondary antibodies. For example, to prepare 500 mL blocking buffer for antibodies raised in goat, combine 25 mL of goat serum, 0.45 g of nonionic surfactant, and 1x PBS to a volume of 500 mL.

2. Preparation of Archived and Freshly Prepared Clinical Tissue Slides for ExPath

1. Convert the tissue into an ExPath compatible format. Choose one of the four following steps (2.1.1−2.1.4) based on how the specimen was prepared: FFPE slides, stained FFPE slides, or unfixed or fixed frozen tissue slides in optimum cutting temperature (OCT) solution.
   NOTE: These are based on standard recovery steps for pathology samples and are not specific to the ExPath protocol.
   1. FFPE clinical samples
      1. Prepare 30 mL of 95% ethanol, 70% ethanol, and 50% ethanol. Measure out 30 mL of xylene, 100% ethanol, and ddH₂O.
      2. Place the slide with the sample in a 50 mL conical using forceps and add 15 mL of xylene. Cap the tube and place it horizontally on an orbital shaker at approximately 60 rpm and incubate at RT for 3 min for each solution. Repeat with the remaining 15 mL xylene.
      3. Repeat step 2.1.1.2 with 100% ethanol, 95% ethanol, 70% ethanol, 50% ethanol, and ddH₂O in place of xylene.
   2. Stained and mounted permanent slides
      1. Place the slide in a 100 mm Petri dish and cover with xylene. Carefully remove the coverslip using a razor blade. If the coverslip is not easily removed, return the slide to the xylene until the coverslip loosens.
      2. Process using the steps for FFPE samples (steps 2.1.1.1−2.1.1.3).
         NOTE: In the case of H&E stained slides, the stains are eliminated during the expansion process.
   3. Unfixed frozen tissue slides in OCT solution
      1. Fix the tissue in acetone at -20 °C for 10 min.
      2. Wash the samples with 1x PBS solution 3 times for 10 min each at RT.
   4. Previously fixed, frozen clinical tissue slides
      1. Incubate the slides for 2 min at RT to melt the OCT solution.
      2. Wash the sample with 1x PBS solution 3 times for 5 min each at RT.
2. Perform heat treatment for antigen retrieval on all samples after format conversion.
   1. Add 20 mM citrate solution (pH 8 at RT) in a heat resistant container, such as a slide staining jar.
      NOTE: There should be enough solution to cover the tissue mounted on the slide (50 mL for a standard slide staining jar).
   2. Heat the citrate solution to 100 °C in the microwave and place the slide in the solution. Immediately transfer the container to an incubation chamber and incubate at 60 °C for 30 min.
      NOTE: The protocol can be paused here. Slides can be placed in Petri dishes and covered in 1x PBS and stored at 4 °C.
3. Stain the sample using standard immunofluorescence (IF)/immunohistochemistry (IHC) staining protocols.
   NOTE: Specific primary and secondary antibody concentrations and staining durations are dependent on the concentrations suggested by the manufacturer or by optimization for the specific experiment.
   1. Use a hydrophobic pen to draw a boundary around the tissue section(s) on the slide to minimize the volume of solution needed to cover the tissue. Place the slide in a dish large enough to fit the slide. For a standard 3-inch slide, use a 100 mm Petri dish.
      NOTE: The hydrophobic pen does not interfere with the polymerization of the sample nor the digestion process.
   2. Incubate the tissue with blocking buffer for 1 h at 37 °C, 2 h at RT, or 4 °C overnight to reduce nonspecific binding.
   3. Dilute primary antibodies to the desired concentration in the appropriate amount of prepared blocking buffer (or other preferred staining buffer). Incubate the tissues with the primary antibody solution for at least 3 h at RT or 37 °C, or overnight at 4 °C.
      NOTE: Samples should be placed in a humidified container (such as a Petri dish with a damp wipe) to prevent the tissue from drying out. Typically, antibodies have been diluted to 1:100−1:500 in 200−500 µL of buffer, depending on the tissue size and antibody used.
   4. Wash the tissue with prepared blocking buffer (or other preferred washing buffer) 3 times for 10 min at RT.
   5. Dilute secondary antibodies (and 300 nM 4′,6-diamidino-2-phenylindole [DAPI] if desired), in prepared blocking buffer (or other preferred staining buffer) to a concentration of approximately 10 µg/mL. Incubate the tissue in the secondary antibody solution for at least 1 h at RT or 37 °C.
      NOTE: Timing may be adjusted depending on the antibodies used and the thickness of the tissue. Secondary antibodies containing cyanine dyes (Cy3, Cy5, Alexa 647) are not compatible with the ExM protocol when applied pre-polymerization. Suggested dyes include Alexa 488 (green), Alexa 546 (orange/red), and Atto 647N or CF633 (far-red). DAPI must be reapplied after expansion, as it is washed away during the expansion process.
   6. Wash the tissue with prepared blocking buffer (or other preferred washing buffer) 3 times for 10 min each at RT.
      NOTE: The protocol can be paused here. Slides can be placed in Petri dishes and covered in 1x PBS and stored at 4 °C.
   7. Perform fluorescent imaging using a conventional wide-field microscope, confocal microscope, or other imaging system of choice.
      NOTE: This step is required to determine biological length using the expansion factor by comparing pre- and post-expansion images. To facilitate post-expansion imaging, easily identifiable regions of interest should be selected and images at both low and high magnification should be collected.

3. In Situ Polymerization of Specimens

1. Incubate the specimen in anchoring solution.
   1. Prepare the anchoring solution (typically 250 µL is enough to cover the tissue section) by diluting the AcX stock solution in 1x PBS to a concentration of 0.03 mg/mL for samples fixed with non-aldehyde fixatives or 0.1 mg/mL for samples fixed with aldehyde fixatives, which have fewer free amines available to react with AcX.
   2. Place the slide in a 100 mm Petri dish and pipette the anchoring solution over the tissue. Incubate for at least 3 h at RT or overnight at 4 °C.
2. Incubate the samples in gelling solution.
   1. Prepare at least 100-fold excess volume of gelling solution. Per 200 µL, combine the following, in order: 188 µL of monomer solution, 4 µL of 0.5% 4HT stock solution (1:50 dilution, final concentration: 0.01%), 4 µL of 10% TEMED stock solution (1:50 dilution, final concentration 0.2%), and 4 µL of 10% APS stock solution (1:50 dilution, final concentration 0.2%).
      NOTE: Gelling solution should be made immediately before use. The solution should be kept at 4 °C and the APS solution should be added last, to prevent premature gelling.
   2. Remove excess solution from the tissue section and place the slide in a 100 mm Petri dish. Add fresh, cold gelling solution to the sample and incubate the mixture on the tissue for 30 min at 4 °C, to allow diffusion of solution into the tissue.
3. Construct a chamber on the slide around the sample (Figure 2A) without disturbing the gelling solution.
   1. Make spacers for the gelling chamber by thinly cutting pieces of cover glass using a diamond knife.
      NOTE: To facilitate imaging post expansion, the spacers should be close in thickness to the tissue specimen to reduce the amount of blank gel above the tissue. Number 1.5 glass can be used for standard clinical samples (5−10 µm). Cover glass pieces can be stacked for thicker samples.
   2. Secure the spacers on either side of the tissue using droplets of water (~10 µL).
   3. Carefully place a cover glass lid over the slide, making sure to avoid trapping air bubbles over the tissue (Figure 2B).
4. Incubate the sample at 37 °C in a humidified environment (such as a closed Petri dish with a damp wipe) for 2 h.
   NOTE: The protocol can be paused here. The slide chamber can be stored inside a sealed Petri dish at 4 °C.

4. Sample Digestion

1. Remove the lid of the gelling chamber by gently sliding a razor blade under the coverslip and slowly lifting the coverslip off the gel surface. Trim the blank gel around the tissue to minimize volume. Cut the gel asymmetrically to track the orientation of the gel after homogenization, since the sample will become transparent.
   1. Dilute ProK by 1:200 in digestion buffer (final concentration 4 U/mL) before use. Prepare enough solution to completely submerge the gel; a single well of a four-well plastic cell culture plate requires at least 3 mL per well.
   2. Incubate the sample in a closed container containing the digestion buffer for 3 h at 60 °C. If the sample does not detach from the slide during digestion, use a razor blade to gently remove the sample.
      NOTE: The specimen should be completely submerged in digestion buffer to prevent the sample from drying out and placed in a covered container (small slide box, plastic well, Petri dish, etc.) that can be sealed with film.

5. Sample Expansion and Imaging

1. Use a soft paint brush to transfer the specimen into 1x PBS in a container compatible with the desired imaging system and large enough to accommodate the fully expanded gel. Make sure that the tissue is placed with the sample-side down if imaging on an inverted system or up if imaging on an upright system to minimize the distance from the imaging objective to the sample. Flip the gel using a soft paint brush if needed.
   NOTE: Side-illumination from an LED can be used to make them visible in liquid. A standard 6-well plate can accommodate samples that have a pre-expanded diameter less than 0.6 cm. A glass bottom well plate should be used for imaging on an inverted system.
2. Wash the samples in 1x PBS at RT for 10 min. If desired, re-stain the sample with 300 nM DAPI as the digestion process washes away the DAPI stain. Remove PBS and stain with 300 nM DAPI diluted in 1x PBS for 20 min at RT, followed by a 10 min wash with 1x PBS at RT.
   NOTE: The samples can be covered with 1x PBS and stored at 4 °C before proceeding to the next step.
3. To expand the samples, replace the PBS and wash with an excess volume of ddH₂O (at least 10x the final gel volume) 3−5 times for 10 min each, at RT.
   NOTE: After the 3^rd or 4^th wash, the specimen’s expansion should begin to plateau. For storage, to prevent bacterial growth, the ddH₂O can be supplemented with 0.002%−0.01% sodium azide (NaN₃). In this case, the final expansion factor is reversibly reduced by 10%.
4. Perform fluorescence imaging using a conventional wide-field microscope, confocal microscope, or other imaging system of choice.
   NOTE: To prevent gels from drifting, excess liquid can be removed from the well. Gels can also be immobilized with 1.5−2% low-melt agarose. Prepare 1.5−2%(w/v) low-melt agarose in water in a container 2−4 times the volume of solution. Warm the solution in a 40 °C water bath or in a microwave for 10−20 s to melt the solution. Pipette the melted agarose around the edges of the gel. After allowing the agarose to harden at RT or 4 °C, add water to the sample to prevent dehydration.

Results

If the protocol has been successfully carried out (Figure 1), samples will appear as a flat and transparent gel after mechanical homogenization (Figure 3A) and can expand by a factor of 3−4.5x in water (Figure 3B), providing an effective resolution of ~70 nm depending on the final expansion factor and imaging system used^5,16.

Discussion

Here, we present the ExPath protocol¹⁶, a variant of proExM⁵ that can be applied to the most common types of clinical biopsy samples used in pathology, including FFPE, H&E stained, and fresh-frozen specimens on glass slides. Format conversion, antigen retrieval, and immunostaining of the specimens follow commonly used protocols that are not specific to ExPath. Unlike the original proExM protocol⁹, ExPath relies on a higher concentration of EDTA i...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
4-hydroxy-TEMPO (4HT)	Sigma Aldrich	176141	Inhibitor
6-well glass-bottom plate (#1.5 coverglass)	Cellvis	P06-1.5H-N
Acetone	Fischer Scientifc	A18-500
Acrylamide	Sigma Aldrich	A8887
Acryloyl-X, SE (AcX)	Invitrogen	A20770
Agarose	Fischer Scientifc	BP160-100
Ammonium persulfate (APS)	Sigma Aldrich	A3678	Initiatior
Anti-ACTN4 antibody produced in rabbit	Sigma Aldrich	HPA001873
Anti-Collagen IV antibody produced in mouse	Santa Cruz Biotech	sc-59814
Anti-Vimentin antibody produced in chicken	Abcam	ab24525
Aqua Hold II hydrophobic pen	Scientific Device	980402
Breast Common Disease Tissue Array	Abcam	ab178113
DAPI (1 mg/mL)	Thermo Scientific	62248	Nuclear stain
Diamond knife No. 88 CM	General Tools	31116
Ethanol	Pharmco	111000200
Ethylenediaminetetraacetic acid (EDTA) 0.5 M	VWR	BDH7830-1
FFPE Kidney Sample	USBiomax	HuFPT072
Forceps
Goat Anti-Chicken IgY (H+L), Highly Cross-Adsorbed CF488A	Biotium	20020
Goat Anti-Chicken IgY (H+L), Highly Cross-Adsorbed CF633	Biotium	20121
Goat anti-Rabbit IgG (H+L) Highly Cross-Adsorbed Alexa Fluor 546	Invitrogen	A11010
MAXbind Staining Medium	Active Motif	15253	Can be substituted with non-commercial staning buffer of choice.
MAXblock Blocking Medium	Active Motif	15252	Can be substituted with non-commercial blocking buffer of choice.
MAXwash Washing Medium	Active Motif	15254	Can be substituted with non-commercial washing buffer of choice.
Micro cover Glass #1 (24 mm x 60 mm)	VWR	48393 106
Micro cover Glass #1.5 (24 mm x 60 mm)	VWR	48393 251
N,N,N′,N′- Tetramethylethylenediamine (TEMED)	Sigma Aldrich	T9281	Accelerator
N,N′-Methylenebisacrylamide	Sigma Aldrich	M7279
Normal goat serum	Jackson Immunoresearch	005-000-121	For preparing blocking buffer. Dependent on animal host of secondary antibodies.
Nunclon 4-Well x 5 mL MultiDish Cell Culture Dish	Thermo Fisher	167063	Multi-well plastic culture dish
Nunclon 6-Well Cell Culture Dish	Thermo Fisher	140675
Nunc 15 mL Conical	Thermo Fisher	339651
Nunc 50 mL Conical	Thermo Fisher	339653
Orbital Shaker
Paint brush
pH Meter
Phosphate Buffered Saline (PBS), 10x Solution	Fischer Scientifc	BP399-1
Plastic Petri Dish (100 mm)	Fischer Scientifc	FB0875713
Proteinase K (Molecular Biology Grade)	Thermo Scientific	EO0491
Razor blade	Fischer Scientifc	12640
Safelock Microcentrifuge Tubes 1.5 mL	Thermo Fisher	3457
Safelock Microcentrifuge Tubes 2.0 mL	Thermo Fisher	3459
Sodium acrylate	Sigma Aldrich	408220
Sodium chloride	Sigma Aldrich	S6191
Sodium citrate tribasic dihydrate	Sigma Aldrich	C8532-1KG
Tris Base	Fischer Scientifc	BP152-1
Triton X-100	Sigma Aldrich	T8787
Wheat germ agglutinin labeled with CF640R	Biotium	29026
Xylenes	Sigma Aldrich	214736