This work was supported by the Faculty Start-up fund from the Carnegie Mellon University (YZ) and NIH Director&#8217;s New Innovator Award (DP2 OD025926-01 to YZ).

1. Preparation of Stock Reagents and Solutions
<ol>
	<li>Prepare gelling solution components. 
	NOTE: Solution concentrations are given in g/mL (w/v percent).
	<ol>
		<li>Make the following stock solutions: 38% (w/v) sodium acrylate (SA), 50% (w/v) acrylamide (AA), 2% (w/v) N,N&#8242;-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.</li>
		<li>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 &#176;C for up to 3 months, or at -20 &#176;C for long-term storage.</li>
		<li>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 &#176;C for at least 6 months. APS has been found to lose efficacy after long-term storage and is best prepared in small quantities (&#60;0.1 mL) immediately before gelling.</li>
	</ol>
	</li>
	<li>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 &#176;C. Proteinase K (ProK) will be added immediately before the digestion step.</li>
	<li>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.</li>
	<li>Prepare a stock solution of 6-((acryloyl)amino)hexanoic acid, succinimidyl ester (acryloyl-X, SE; AcX), the anchoring compound. Dissolve AcX in 500 &#181;L 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 &#176;C in 20 &#181;L aliquots.</li>
	<li>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.</li>
</ol>
2. Preparation of Archived and Freshly Prepared Clinical Tissue Slides for ExPath
<ol>
	<li>Convert the tissue into an ExPath compatible format. Choose one of the four following steps (2.1.1&#8722;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.
	<ol>
		<li>FFPE clinical samples
		<ol>
			<li>Prepare 30 mL of 95% ethanol, 70% ethanol, and 50% ethanol. Measure out 30 mL of xylene, 100% ethanol, and ddH2O.</li>
			<li>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.</li>
			<li>Repeat step 2.1.1.2 with 100% ethanol, 95% ethanol, 70% ethanol, 50% ethanol, and ddH2O in place of xylene.</li>
		</ol>
		</li>
		<li>Stained and mounted permanent slides
		<ol>
			<li>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.</li>
			<li>Process using the steps for FFPE samples (steps 2.1.1.1&#8722;2.1.1.3). 
			NOTE: In the case of H&#38;E stained slides, the stains are eliminated during the expansion process.</li>
		</ol>
		</li>
		<li>Unfixed frozen tissue slides in OCT solution
		<ol>
			<li>Fix the tissue in acetone at -20 &#176;C for 10 min.</li>
			<li>Wash the samples with 1x PBS solution 3 times for 10 min each at RT.</li>
		</ol>
		</li>
		<li>Previously fixed, frozen clinical tissue slides
		<ol>
			<li>Incubate the slides for 2 min at RT to melt the OCT solution.</li>
			<li>Wash the sample with 1x PBS solution 3 times for 5 min each at RT.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Perform heat treatment for antigen retrieval on all samples after format conversion.
	<ol>
		<li>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).</li>
		<li>Heat the citrate solution to 100 &#176;C in the microwave and place the slide in the solution. Immediately transfer the container to an incubation chamber and incubate at 60 &#176;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 &#176;C.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Incubate the tissue with blocking buffer for 1 h at 37 &#176;C, 2 h at RT, or 4 &#176;C overnight to reduce nonspecific binding.</li>
		<li>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 &#176;C, or overnight at 4 &#176;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&#8722;1:500 in 200&#8722;500 &#181;L of buffer, depending on the tissue size and antibody used.</li>
		<li>Wash the tissue with prepared blocking buffer (or other preferred washing buffer) 3 times for 10 min at RT.</li>
		<li>Dilute secondary antibodies (and 300 nM 4&#8242;,6-diamidino-2-phenylindole [DAPI] if desired), in prepared blocking buffer (or other preferred staining buffer) to a concentration of approximately 10 &#181;g/mL. Incubate the tissue in the secondary antibody solution for at least 1 h at RT or 37 &#176;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.</li>
		<li>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 &#176;C.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
3. In Situ Polymerization of Specimens
<ol>
	<li>Incubate the specimen in anchoring solution.
	<ol>
		<li>Prepare the anchoring solution (typically 250 &#181;L 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.</li>
		<li>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 &#176;C.</li>
	</ol>
	</li>
	<li>Incubate the samples in gelling solution.
	<ol>
		<li>Prepare at least 100-fold excess volume of gelling solution. Per 200 &#181;L, combine the following, in order: 188 &#181;L of monomer solution, 4 &#181;L of 0.5% 4HT stock solution (1:50 dilution, final concentration: 0.01%), 4 &#181;L of 10% TEMED stock solution (1:50 dilution, final concentration 0.2%), and 4 &#181;L 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 &#176;C and the APS solution should be added last, to prevent premature gelling.</li>
		<li>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 &#176;C, to allow diffusion of solution into the tissue.</li>
	</ol>
	</li>
	<li>Construct a chamber on the slide around the sample (Figure 2A) without disturbing the gelling solution.
	<ol>
		<li>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&#8722;10 &#181;m). Cover glass pieces can be stacked for thicker samples.</li>
		<li>Secure the spacers on either side of the tissue using droplets of water (~10 &#181;L).</li>
		<li>Carefully place a cover glass lid over the slide, making sure to avoid trapping air bubbles over the tissue (Figure 2B).</li>
	</ol>
	</li>
	<li>Incubate the sample at 37 &#176;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 &#176;C.</li>
</ol>
4. Sample Digestion
<ol>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Incubate the sample in a closed container containing the digestion buffer for 3 h at 60 &#176;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.</li>
	</ol>
	</li>
</ol>
5. Sample Expansion and Imaging
<ol>
	<li>Use a soft paint brush to transfer the specimen into 1x PBS in&#160;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.</li>
	<li>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 &#176;C before proceeding to the next step.</li>
	<li>To expand the samples, replace the PBS and wash with an excess volume of ddH2O (at least 10x the final gel volume) 3&#8722;5 times for 10 min each, at RT. 
	NOTE: After the 3rd or 4th wash, the specimen&#8217;s expansion should begin to plateau. For storage, to prevent bacterial growth, the ddH2O can be supplemented with 0.002%&#8722;0.01% sodium azide (NaN3). In this case, the final expansion factor is reversibly reduced by 10%.</li>
	<li>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&#8722;2% low-melt agarose. Prepare 1.5&#8722;2%(w/v) low-melt agarose in water in a container 2&#8722;4 times the volume of solution. Warm the solution in a 40 &#176;C water bath or in a microwave for 10&#8722;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 &#176;C, add water to the sample to prevent dehydration.</li>
</ol>

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YZ and OB are two of the inventors who have filed for and obtained patent protection on a subset of the technologies described here (US patents US20190064037A1, WO2018157074A1, and WO2018157048A1).

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&#38;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...

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&#8722;300 nm), where the minimal resolvable distance, d is defined by d &#945; &#955;/NA. Here &#955; 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 techniques1,2,3, 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 (&#60;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 RNA9,10,11,12 to the hydrogel, and increase physical magnification through iterative expansion13 or adapting gel chemistry8,14,15.
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&#38;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.

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

nanoscopic imaging of human tissue sections via physical and isotropic expansion

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.

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&#8722;4.5x in water (Figure 3B), providing an effective resolution of ~70 nm depending on the final expansion factor and imaging system used5,16. <strong c...

Watch this Scientific Journal Video about Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion at JoVE.com

Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion

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.

In modern pathology, optical microscopy plays an important role in disease diagnosis by revealing microscopic structures of clinical specimens. However, ...

nanoscopic-imaging-human-tissue-sections-via-physical-isotropic

Research

JoVE Journal

Medicine

7.5K Views.  Carnegie Mellon University. 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.

Video: Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion

Registered Bioimaging of Nanomaterials for Diagnostic and Therapeutic Monitoring

Nanomedications can be carried by blood borne monocyte-macrophages into the reticuloendothelial system (RES; spleen, liver, lymph nodes) and to end organs. The latter include the lung, RES, and brain and are operative during human immunodeficiency virus type one (HIV-1) infection. Macrophage entry into tissues is notable in areas of active HIV-1 replication and sites of inflammation. In order to assess the potential of macrophages as nanocarriers, superparamagnetic iron-oxide and/or drug laden particles coated with surfactants were parenterally injected into HIV-1 encephalitic mice. This was done to quantitatively assess particle and drug biodistribution. Magnetic resonance imaging (MRI) test results were validated by histological coregistration and enhanced image processing. End organ disease as typified by altered brain histology were assessed by MRI. The demonstration of robust migration of nanoformulations into areas of focal encephalitis provides '"proof of concept" for the use of advanced bioimaging techniques to monitor macrophage migration. Importantly, histopathological aberrations in brain correlate with bioimaging parameters making the general utility of MRI in studies of cell distribution in disease feasible. We posit that using such methods can provide a real time index of disease burden and therapeutic efficacy with translational potential to humans.

Bioimaging methods used to assess cell biodistribution of nanoparticles are applicable for therapeutic and diagnostic monitoring of nanoformulated compounds. The methods described herein are sensitive and specific when assessed by histological coregistration. The methodologies provide a translational pathway from rodent to human applications.

Nanomedications can be carried by blood borne monocyte-macrophages into the reticuloendothelial system (RES; spleen, liver, lymph nodes) and to end ...

Bioengineering

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Neuroscience

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a combination of light and electron microscopy. Samples embedded in resin are sectioned into arrays consisting of ribbons of hundreds of ultrathin sections and deposited on pieces of silicon wafer or conductively coated coverslips. Arrays are imaged at low resolution using a digital consumer like smartphone camera or light microscope (LM) for a rapid large area overview, or a wide field fluorescence microscope (fluorescence light microscopy (FLM)) after labeling with fluorophores. After post-staining with heavy metals, arrays are imaged in a scanning electron microscope (SEM). Selection of targets is possible from 3D reconstructions generated by FLM or from 3D reconstructions made from the SEM image stacks at intermediate resolution if no fluorescent markers are available. For ultrastructural analysis, selected targets are finally recorded in the SEM at high-resolution (a few nanometer image pixels). A ribbon-handling tool that can be retrofitted to any ultramicrotome is demonstrated. It helps with array production and substrate removal from the sectioning knife boat. A software platform that allows automated imaging of arrays in the SEM is discussed. Compared to other methods generating large volume EM data, such as serial block-face SEM (SBF-SEM) or focused ion beam SEM (FIB-SEM), this approach has two major advantages: (1) The resin-embedded sample is conserved, albeit in a sliced-up version. It can be stained in different ways and imaged with different resolutions. (2) As the sections can be post-stained, it is not necessary to use samples strongly block-stained with heavy metals to introduce contrast for SEM imaging or render the tissue blocks conductive. This makes the method applicable to a wide variety of materials and biological questions. Particularly prefixed materials e.g., from biopsy banks and pathology labs, can directly be embedded and reconstructed in 3D.

This protocol targets specific cells in tissue for imaging at nanoscale resolution using a scanning electron microscope (SEM). Large numbers of serial sections from resin-embedded biological material are first imaged in a light microscope to identify the target and then in a hierarchical manner in the SEM.

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a ...

Developmental Biology

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick tissue samples and organs in health and disease. By controlling the scanning pattern in multi-photon microscopy and applying appropriate numerical algorithms, we developed a striped-illumination approach, which enabled us to achieve 3-fold better axial resolution and improved signal-to-noise ratio, i.e. contrast, in more than 100 &#181;m tissue depth within highly scattering tissue of lymphoid organs as compared to standard multi-photon microscopy. The acquisition speed as well as photobleaching and photodamage effects were similar to standard photo-multiplier-based technique, whereas the imaging depth was slightly lower due to the use of field detectors. By using the striped-illumination approach, we are able to observe the dynamics of immune complex deposits on secondary follicular dendritic cells &#8211; on the level of a few protein molecules in germinal centers.

High-resolution intravital imaging with enhanced contrast up to 120 &#181;m depth in lymph nodes of adult mice is achieved by spatially modulating the excitation pattern of a multi-focal two-photon microscope. In 100 &#181;m depth we measured resolutions of 487 nm (lateral) and 551 nm (axial), thus circumventing scattering and diffraction limits.

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick ...

Immunology and Infection

Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte foot processes contain actin bundles that bind to cytoskeletal adaptor proteins such as podocin. Those adaptor proteins, such as podocin, link the backbone of the glomerular slit diaphragm, such as nephrin, to the actin cytoskeleton. Studying the localization and function of these and other podocytic proteins is essential for the understanding of the glomerular filter&#39;s role in health and disease. The presented protocol enables the user to visualize actin, podocin, and nephrin in cells with super resolution imaging on a conventional microscope. First, cells are stained with a conventional immunofluorescence technique. All proteins within the sample are then covalently anchored to a swellable hydrogel. Through digestion with proteinase K, structural proteins are cleaved allowing isotropical swelling of the gel in the last step. Dialysis of the sample in water results in a 4-4.5-fold expansion of the sample and the sample can be imaged via a conventional fluorescence microscope, rendering a potential resolution of 70 nm.

The presented method enables visualization of fluorescently labeled cellular proteins with expansion microscopy leading to a resolution of 70 nm on a conventional microscope.

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte ...

Biochemistry

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged molecules within complex biological samples. While traditional confocal microscopes allow one to resolve two adjacent fluorescent point sources located a few hundred nanometers apart, observing the finer details of subcellular biology requires the ability to resolve signals in the order of tens of nanometers. Numerous hardware-based methods for super-resolution microscopy have been developed to allow researchers to sidestep such resolution limits, although these methods require specialized microscopes that are not available to all researchers. An alternative method for increasing resolving power is to isotropically enlarge the sample itself through a process known as expansion microscopy (ExM), which was first described by the Boyden group in 2015. ExM is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules. The expanded sample can then be observed at an effectively increased resolution using a traditional confocal microscope. Here, we describe a protocol for implementing ExM in whole-mount Drosophila embryos, which is used to examine the localization of Par-3, myosin II, and mitochondria within the surface epithelial cells. This protocol yields an approximately four-fold increase in sample size, allowing for the detection of subcellular details that are not visible with conventional confocal microscopy. As proof of principle, an anti-GFP antibody is used to distinguish distinct pools of myosin-GFP between adjacent cell cortices, and fluorescently labeled streptavidin is used to detect endogenous biotinylated molecules to reveal the fine details of the mitochondrial network architecture. This protocol utilizes common antibodies and reagents for fluorescence labeling, and it should be compatible with many existing immunofluorescence protocols.

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Here, a protocol for the implementation of expansion microscopy in early Drosophila embryos to achieve super-resolution imaging using a conventional laser-scanning confocal microscope is presented.

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged ...

3D Imaging of Soft-Tissue Samples using an X-ray Specific Staining Method and Nanoscopic Computed Tomography

We demonstrate a laboratory-based method combining X-ray microCT and nanoCT with a specific X-ray stain, which targets the cell cytoplasm. The described protocol is easy to apply, fast and suitable for larger soft-tissue samples. The presented methodology enables the characterization of crucial tissue structures in three dimensions and is demonstrated on a whole mouse kidney. The multiscale approach allows to image the entire mouse kidney and supports the selection of further volumes of interest, which are acquired with higher resolutions ranging into the nanometer range. Thereby, soft-tissue morphology with a similar detail level as the corresponding histological light microscopy images is reproduced. Deeper insights into the 3D configuration of tissue structures are achieved without impeding further investigations through histological methods.

A protocol for 3D visualization of microscopic tissue structures by using an X-ray specific staining method designed for X-ray computed tomography is presented.

We demonstrate a laboratory-based method combining X-ray microCT and nanoCT with a specific X-ray stain, which targets the cell cytoplasm. The described ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

Biology

Expansion Microscopy for Robust 11-Fold Expansion

Universal Molecular Retention with 11-Fold Expansion Microscopy

The nanoscale imaging of biological specimens can improve the understanding of disease pathogenesis. In recent years, expansion microscopy (ExM) has been demonstrated to be an effective and low-cost alternative to optical super-resolution microscopy. However, it has been limited by the need for specific and often custom anchoring agents to retain different biomolecule classes within the gel and by difficulties with expanding standard clinical sample formats, such as formalin-fixed paraffin-embedded tissue, especially if larger expansion factors or preserved protein epitopes are desired. Here, we describe Magnify, a new ExM method for robust expansion up to 11-fold in a wide array of tissue types. By using methacrolein as the chemical anchor between the tissue and gel, Magnify retains multiple biomolecules, such as proteins, lipids, and nucleic acids, within the gel, thus allowing the broad nanoscale imaging of tissues on conventional optical microscopes. This protocol describes best practices to ensure robust and crack-free tissue expansion, as well as tips for handling and imaging highly expanded gels.

Author Spotlight: Universal Molecular Retention with 11-Fold Expansion Microscopy  

Presented here is a new version of expansion microscopy (ExM), Magnify, that is modified for up to 11-fold expansion, conserving a comprehensive array of biomolecule classes, and is compatible with a broad range of tissue types. It enables the interrogation of the nanoscale configuration of biomolecules using conventional diffraction-limited microscopes.

The nanoscale imaging of biological specimens can improve the understanding of disease pathogenesis. In recent years, expansion microscopy (ExM) has been ...

Expansion Microscopy for Visualizing Intracellular Sialylated Glycans via MOE

Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes within cells through click and bioorthogonal chemistry. Metabolic oligosaccharide engineering (MOE) has attracted considerable interest due to the essential role of glycosylation in numerous biological processes that involve molecular recognition and its impact on pathologies ranging from cancer to genetic disorders to viral and bacterial infections.
Although MOE is better known for the detection of cell surface glycoconjugates, it is also a very important methodology for the study of intracellular glycans in physiological and pathological contexts. Such studies greatly benefit from high spatial resolution. However, super-resolution microscopy is not readily available in most laboratories and poses challenges for daily implementation. Expansion microscopy is a recent alternative that enhances the resolution of microscopy by physically enlarging biological specimens labeled with fluorescent markers. By embedding the sample in a swellable gel and causing it to expand isotropically through chemical treatment, subcellular structures can be visualized with enhanced precision and resolution without the need for super-resolution techniques.
In this work, we illustrate the capacity of expansion microscopy to visualize intracellular sialylated glycans through the combined use of MOE and click chemistry. Specifically, we propose a procedure for bioorthogonal labeling and expansion microscopy that employs a reporter targeting sialylation, which may be associated with immunofluorescence for co-localization studies. This protocol enables localization studies of sialoconjugate biosynthesis, intracellular trafficking, and recycling.

Here, we propose a simple protocol combining metabolic oligosaccharide engineering, click chemistry, and expansion microscopy that allows bioimaging of intracellular sialylated N-glycoproteins with improved resolution using routine microscopy equipment.

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes ...

Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion

Summary

Explore More Videos

Chapters in this video

Registered Bioimaging of Nanomaterials for Diagnostic and Therapeutic Monitoring

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

3D Imaging of Soft-Tissue Samples using an X-ray Specific Staining Method and Nanoscopic Computed Tomography

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Universal Molecular Retention with 11-Fold Expansion Microscopy

Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy