This work was supported by Carnegie Mellon University and the D.S.F. Charitable Foundation (Y.Z. and X.R.), the National Institutes of Health (N.I.H.) Director&#39;s New Innovator Award DP2 OD025926-01, and the Kauffman Foundation.

All the experimental procedures involving animals were conducted in accordance with the National Institutes of Health (NIH) guidelines and were approved by the Institutional Animal Care and Use Committee at Carnegie Mellon University. Human tissue samples were commercially obtained.
1. Preparation of the stock reagents and solutions
NOTE: Refer to the Table of Materials for a list of the reagents used.
<ol>
	<li>Prepare the gelling stock solutions. These will be combined immediately before the gelling step.
	<ol>
		<li>Prepare the monomer stock solution (4 g/100 mL N,N- dimethylacrylamide acid [DMAA], 50 g/100 mL sodium acrylate [SA], 66.7 g/100 mL acrylamide [AA], 0.10 g/100 mL N,N&#8242;-methylenebisacrylamide [Bis], 33 g/100 mL sodium chloride [NaCl], 1x phosphate-buffered saline; PBS) using the components listed in Table 1. Dissolve or dilute all the stock solutions in water. The monomer stock solution can be stored at 4 &#176;C for at least 3 months.</li>
		<li>Make the following stock solutions separately in water: 0.5% (w/w) 4-Hydroxy-TEMPO (4HT), which inhibits gelation during monomer diffusion into the tissues, and 10% (w/w) tetramethylethylenediamine (TEMED), which accelerates the radical generation initiated by ammonium persulfate (APS) and is prepared at 10% (w/w). 
		&#8203;NOTE: The stock solutions can be distributed into 1-2 mL aliquots and stored at 4 &#176;C for up to 3 months; however, APS is made immediately before preparing the gelling solution.</li>
	</ol>
	</li>
	<li>Prepare the homogenization buffer (1% w/v S.D.S., 8 M urea, 25 mM EDTA, 2x PBS, pH 8, at room temperature [R.T.]) by combining 1 g of SDS, 48.048 g of urea, 5 mL of EDTA (0.5M, pH 8), 20 mL of 10x PBS, 25 mL of Tris (2 M), and 0.75 g of glycine. Add water for a total volume of 100 mL. The solution can be scaled up or down as needed and can be stored at R.T.</li>
</ol>
2. Tissue preparation for archived and freshly prepared clinical tissue slides
NOTE: The tissue pre-processing steps differ based on how the specimens are prepared.
<ol>
	<li>For formaldehyde-fixed paraffin-embedded (FFPE) clinical samples, follow the steps below.
	<ol>
		<li>Place the samples in a sequential series of solutions (a glass slide staining jar or 50 mL conical tube may be used): xylene, 95% ethanol, 70% ethanol, and 50% ethanol, followed by doubly deionized water. Use each solution twice for at least 3 min each time. Use forceps to place the slide into the container, and add 15 mL of the respective solution (enough to cover the sample). Place the capped conical tube horizontally on a shaker at room temperature.</li>
	</ol>
	</li>
	<li>For samples stained and mounted on permanent slides, follow the steps below.
	<ol>
		<li>Place the slide briefly in xylene, and carefully remove the coverslips using an appropriate tool, such as a razor blade. If the coverslip remains stuck, return the slides in xylene to room temperature until the coverslip loosens.</li>
		<li>Then, process the samples as FFPE samples (step 2.1.1). 
		NOTE: For hematoxylin and eosin (H&#38;E)-stained slides, the hematoxylin and eosin stain is lost during the expansion process.</li>
	</ol>
	</li>
	<li>For unfixed frozen tissue slides in optimum cutting temperature (OCT) solution, fix the tissues in acetone at &#8722;20 &#176;C for 10 min before washing the samples with 1x PBS solution three times for 10 min each time at room temperature.</li>
	<li>For previously fixed frozen clinical tissue slides, melt the OCT by incubating the slides for 2 min at room temperature, and then wash with 1x PBS solution three times for 5 min each time at room temperature.</li>
</ol>
3. Tissue preparation for paraformaldehyde-fixed mouse brain
<ol>
	<li>Follow this protocol for mouse brain sections that are 30-50 &#181;m thick. 
	NOTE: Success may be found with larger sections, but longer times may be required for monomer solution diffusion and homogenization.</li>
	<li>If the sections are not currently stored in a 1x PBS solution (for instance, in a 30% [v/v] glycerol and 30% [v/v] ethylene glycol solution in 1x PBS), wash three times for at least 10 min each time at room temperature in 1x PBS in a 6-to-24-well plate.</li>
	<li>Obtain an uncoated glass microscopy slide. If one side of the glass slide is coated, use a wet paintbrush to check for the uncoated side (often, the label of the coated side will be made of ground glass and will become less opaque when wet).</li>
	<li>Use a paintbrush to wet the uncoated slide. Remove the mouse brain tissue from the well plate with the paintbrush, and carefully place it onto the slide, using a rolling motion to ensure the tissue is flat. Allow excess PBS to remain on the tissue until all the sections are mounted. 
	NOTE: While a single glass slide may be used for multiple tissue sections, when more tissue sections are gelled at once (even on separate slides), more care must be taken to ensure they do not completely dry out.</li>
	<li>Using a laboratory paper cloth, remove the majority of the excess PBS from the slide. Leave a small liquid ring around each tissue section, but leave the rest of the slide mostly dry. This remaining liquid will cover the tissue while the gel monomer solution is being prepared.</li>
</ol>
4. Gelling
NOTE: This protocol is suitable for all tissue types prepared for use with this technique.
<ol>
	<li>Apply spacers on either side of the tissue section (Figure 2A) to prevent tissue compression. To do so, make the spacers by thinly cutting pieces of cover glass using a diamond-tipped pen, and then secure them to the slide using small amounts of water or 1x PBS, or fix them using a small amount of super glue. Next, gently place the spacers on either side of the tissue.</li>
	<li>Prepare the final gelling solution. Generally, ~200 &#181;L is sufficient per tissue section, but the volume should not exceed 800 &#181;L per slide. Any more gelling solution will result in spillover. 
	NOTE: Methacrolein is used to anchor biomolecules into the gel. However, no separate anchoring step is required, and methacrolein may be added directly to the final gelling solution.
	<ol>
		<li>Per section, combine the following in order: 200 &#181;L of monomer solution, 0.5 &#181;L of 0.5% 4HT stock solution (1:400 dilution), 0.2-0.5 &#181;L of methacrolein (1:400-1:1,000 dilution depending on the tissue type; generally, use a 1:1,000 dilution for paraformaldehyde [PFA]-fixed samples and 1:400 for FFPE clinical specimens), 2 &#181;L of 10% TEMED stock solution (1:100 dilution), and 5 &#181;L of 10% A.P.S. stock solution (1:40 dilution). 
		NOTE: Prepare the final gelling solution immediately before use. To prevent premature gelling, add the APS solution last.</li>
	</ol>
	</li>
	<li>Remove the excess solution from the tissue section, and place the slide in a Petri dish. Add all the freshly prepared, cold gelling solution to the sample, and incubate the mixture on the tissue for 30 min at 4 &#176;C to allow diffusion into the tissue.</li>
	<li>Place a second uncoated glass slide carefully over the tissue and gel solution. Air bubbles should be avoided (Figure 2B). 
	NOTE: If air bubbles become trapped over the tissue, the glass lid may be carefully lifted and placed back down. Additionally, any monomer solution that spills out may be trimmed off after polymerization and will not cause any problems for the tissue.</li>
	<li>Ensure that polymerization is completed. This polymerization can be done in two ways, each producing similar results. First, incubate the samples at 37 &#176;C in a humidified environment (such as a closed Petri dish with a damp paper towel) overnight. Alternatively, for more rapid polymerization, incubate the samples in the humidified atmosphere for 2 h at 37 &#176;C followed by 1 h at 60&#176;C.</li>
</ol>
5. Sample digestion and tissue expansion
NOTE: This protocol is suitable for all tissue types.
<ol>
	<li>Wearing eye protection, slide a razor blade between the two glass slides of the gelling chamber to separate them. 
	NOTE: The glass slides should separate very easily. If different parts of the gel are stuck to both slides, a paintbrush may be used to guide the gel to one or the other. If the gel has become particularly dry during polymerization, 1x PBS may be applied to the outside of the gel chamber, but do not submerge the gel fully, as this will cause it to expand before the tissue is homogenized, which will result in cracking.</li>
	<li>Trim the blank gel around the tissue to minimize the volume. Cutting the gel asymmetrically allows for the tracking of the orientation of the gel after homogenization, as the sample will be completely transparent.</li>
	<li>Gently lift the tissue-containing gel from the slide with a razor blade, and gently transfer it into a 2 mL centrifuge tube with a paintbrush.</li>
	<li>Fill the tube to the top with homogenization buffer (Table 2) pre-heated to 80 &#176;C.</li>
	<li>Incubate the sample with shaking at 80 &#176;C for 8 h (for PFA-fixed mouse brain) or 60 h (most FFPE clinical samples; see Table 3). 
	NOTE: The homogenization time is dependent on the tissue type and fixation method. Many of these conditions have been validated already, but for others, optimization may be necessary.</li>
	<li>Pour the contents of the centrifuge tube into a single well of a 6-well plastic cell culture plate.</li>
	<li>Remove the denaturation buffer with a transfer pipet.</li>
	<li>To completely remove the remaining SDS from the hydrogel, wash the sample in a 1% non-ionic surfactant (C12E10) solution as follows: at least three times for 10 min each time at room temperature; for 60 min at 60 &#176;C; and then another three washes for 10 min each time at room temperature.</li>
	<li>Store the sample in 1x PBS + 0.02% sodium azide at 4 &#176;C until post-expansion staining and imaging need to be performed. 
	&#8203;NOTE: At this point, the sample should be ~3-4.5-fold larger in each dimension, depending on the tissue type.</li>
</ol>
6. Post-expansion biomolecule profiling
<ol>
	<li>Antibody staining 
	NOTE: This follows a typical immunofluorescence (IF)/immunohistochemistry (I.H.C.) staining protocol. The amounts of primary and secondary antibodies used depend on the concentrations suggested by the manufacturer or by optimization for the specific experiment. Individual buffers may be substituted at the user&#39;s discretion.
	<ol>
		<li>With the sample in a single well of a 6-well cell culture plate, wash the expanded samples three times for 10 min each time in 1x PBS at R.T., transferring the liquid with a pipette.</li>
		<li>Perform an optional blocking step (for instance, 3% bovine serum albumin/0.1% Triton-X100 in 1x PBS) for at least 1 h at room temperature or overnight at 4 &#176;C.</li>
		<li>Incubate with the primary antibodies in staining buffer (for instance, 0.1% Triton-X100 1x PBS or 9x PBS/1% TritonX/10 mg/L heparin) overnight at 37 &#176;C. Depending on the sample size, 0.5-2 mL should adequately cover the gel.</li>
		<li>Wash the sample three times for at least 10 min each time in 1x PBS at room temperature.</li>
		<li>Incubate in the corresponding secondary antibodies for 3 h at 37 &#176;C in the staining buffer of choice.</li>
		<li>Wash the sample three times for at least 10 min each time in 1x PBS at room temperature.</li>
	</ol>
	</li>
	<li>NHS-ester pan-protein staining
	<ol>
		<li>With the sample in a 6-well plate, pour 1-2 mL of polyethylene glycol (PEG 200) onto the sample (enough to fully cover it). 
		NOTE: The tissue should shrink and return to its original pre-expansion size (or close to it).</li>
		<li>Stain with fluorophore-conjugated NHS-ester diluted to 13.3-80 &#181;g/mL for 3 h at R.T. in 1-2 mL of staining buffer (for instance, 1x PBS, 2x S.S.C., or 100 mM sodium bicarbonate solution).</li>
		<li>Wash the sample three times for at least 10 min each time in 1x PBS at room temperature.</li>
	</ol>
	</li>
	<li>Lipid staining 
	NOTE: Post-expansion lipid staining is not effective on FFPE clinical samples, as lipids are lost during the fixation process.
	<ol>
		<li>Wash the sample three times for at least 10 min each time in 1x PBS at room temperature.</li>
		<li>Apply a conventional lipophilic dye (DiO, DiI, or DiD) diluted 200-fold in 2 mL of 0.1% TritonX-100/1x PBS for 72-96 h at room temperature.</li>
		<li>Wash at least three times with 1x PBS</li>
	</ol>
	</li>
	<li>Fluorescence in situ hybridization (FISH)
	<ol>
		<li>Place homogenized gel samples in hybridization buffer pre-heated to 60 &#176;C for 30 min.</li>
		<li>Prepare the hybridization buffer for FISH: Combine 2x S.S.C. (300 mM NaCl, 30 mM sodium citrate, pH 7.0), 10% (v/v) dextran sulfate, 20% (v/v) ethylene carbonate, and 0.1% (v/v) Tween20.</li>
		<li>Incubate the gel samples with a hybridization buffer containing 10 pM (per oligo) of FISH probes against the target gene (at least 20 probes per 10 kb) at 45 &#176;C overnight.</li>
		<li>Wash with stringency wash buffer (2x S.S.C. and 0.1% Tween20) two times for 15 min each time at 45 &#176;C.</li>
		<li>Wash with stringency wash buffer another two times for 10 min each time at 37 &#176;C.</li>
		<li>Finally, wash with 1x PBS at least three times for 10 min each time at room temperature.</li>
		<li>The samples are ready for imaging or storage in 1x PBS + 0.02% sodium azide at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Full tissue expansion and fluorescence imaging 
	NOTE: The expansion factor attainable with Magnify varies with the tissue type and expansion method (a full summary can be found in Table 3). However, as a general estimate, a Magnify sample will expand 3-4.5-fold in 1x PBS, 5-7-fold in PBS diluted to 1:50 in ddH2O, and 8-11-fold in ddH2O. The optimal expansion factor for imaging depends on each experiment&#39;s needs. The expansion factor is highly tunable, such that a single sample may be expanded and shrunk many times for imaging at various scales.
	<ol>
		<li>Move the samples expanded 4-fold in PBS to a glass-bottom 6-well imaging plate using a paintbrush. Move any sample expanded further by cutting it into smaller pieces or placing gently into a large glass-bottom imaging plate with a thin piece of flexible plastic.</li>
		<li>Perform fluorescence imaging using a conventional wide-field microscope, a confocal microscope, or another imaging system of choice. Removing excess liquid around the gel with a paper laboratory cloth may prevent gel movement during imaging. The gels can also be immobilized by applying 0.1% poly-L-lysine to the glass surface before imaging. 
		NOTE: The application of poly-L-lysine will make the gel entirely immobile upon contact. For this reason, caution must be taken that the gel is applied to the glass without folding or stretching. Additionally, the gel should not be allowed to dry completely on poly-L-lysine-coated glass, as this could make removal difficult, and the gel should not be shrunk in PBS while it is still attached to the glass with poly-L-lysine, as this could cause gel or tissue damage.</li>
		<li>After imaging, shrink the samples in 1x PBS with at least three washes for 10 min each time, as it is not recommended to store the samples long-term in ddH2O due to the degradation of the fluorescence signal. In particular, fully expanded samples stained with lipophilic dyes should be imaged as close to the time of expansion as possible to prevent dissociation of the dye in water.</li>
		<li>Store the samples in 1x PBS + 0.02% sodium azide at 4 &#176;C.</li>
	</ol>
	</li>
</ol>

Expansion Microscopy for Robust 11-Fold Expansion

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The authors declare the following competing financial interest(s): Y.Z., A.K., Z.C., and B.R.G. invented several inventions related to Magnify and ExM.

Here, we present the Magnify protocol17, an ExM variant that can retain multiple biomolecules with a single chemical anchor and expand challenging FFPE clinical specimens up to 11-fold with heat denaturation. The key changes in this protocol that distinguish it from other ExM protocols include the use of a reformulated gel that remains mechanically robust even when fully expanded, as well as the use of methacrolein as the biomolecule anchor. The most critical steps in this protocol are as follows:...

Biological systems&#160;exhibit structural heterogeneity, from the limbs and the organs down to the levels of proteins at the nanoscale. Therefore, a complete understanding of the operation of these systems requires visual examination across these size scales. However, the diffraction limit of light causes challenges in visualizing structures smaller than ~200-300 nm on a conventional fluorescence microscope. In addition, optical super-resolution methods1,2,3, such as stimulated emission depletion (STED), photo-activated localization microscopy (PALM), stochastic optical reconstruction microscopy (STORM), and structured illumination microscopy (SIM), though powerful, present their own challenges, as they require expensive hardware and reagents and often have slow acquisition times and a poor ability to image large volumes in 3D.
Expansion microscopy4 (ExM) provides an alternative means of circumventing the diffraction limit of light by covalently anchoring biomolecules into a water-swellable polymer gel and physically pulling them apart, thus rendering them resolvable on conventional optical microscopes. A multitude of ExM protocol variants have been developed since the original publication of&#160;ExM less than a decade ago, and these protocols allow the direct incorporation of proteins5,6,7, RNA8,9,10, or lipids11,12,13 into the gel network by altering the chemical anchor or expanding the sample further (thus improving the effective resolution) either in a single step14 or multiple iterative steps15,16. Until recently, no single ExM protocol could retain these three biomolecule classes with a single commercially available chemical anchor while providing a mechanically sturdy gel that could expand ~10-fold in a single expansion round.
Here, we present Magnify17, a recent addition to the ExM arsenal that uses methacrolein as the biomolecule anchor. Methacrolein forms covalent bonds with tissue like that of paraformaldehyde, ensuring that multiple classes of biomolecules can be retained within the gel network without requiring various specific or custom anchoring agents. Additionally, this technique can expand a broad spectrum of tissues up to 11-fold, including notoriously challenging samples such as formalin-fixed paraffin-embedded (FFPE) clinical samples. Previous methods for expanding such mechanically rigid samples required harsh protease digestion, rendering the antibody labeling of proteins of interest impossible after the sample had been expanded. In contrast, this technique achieves the expansion of FFPE clinical samples using a hot denaturing solution, thus preserving whole protein epitopes within the gel, which can be targeted for post-expansion imaging (Figure 1).

<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>Acrylamide</td><td>Sigma Aldrich</td><td>A8887</td><td>Gel Monomer component</td></tr><tr><td>Ammonium persulfate (APS)&amp;nbsp;</td><td>Sigma Aldrich</td><td>A3678</td><td>Initiatior</td></tr><tr><td>DAPI (1 mg/mL)</td><td>Thermo Scientific</td><td>62248</td><td></td></tr><tr><td>Decaethylene glycol mono dodecyl ether (C12E10)</td><td>Sigma Aldrich</td><td>P9769</td><td>Non-ionic surfactant</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>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>Homogenization Buffer Component</td></tr><tr><td>Forceps</td><td></td><td></td><td></td></tr><tr><td>Glycine</td><td>Sigma Aldrich</td><td>G8898</td><td>Homogenization Buffer Component</td></tr><tr><td>Heparin</td><td>Sigma Aldrich</td><td>H3393</td><td></td></tr><tr><td>Methacrolein</td><td>Sigma Aldrich</td><td>133035</td><td>Anchoring Agent</td></tr><tr><td>Micro cover Glass #1 (24x60mm)</td><td>VWR</td><td>48393 106</td><td></td></tr><tr><td>Micro cover Glass #1.5 (24x60mm)</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 (Bis)</td><td>Sigma Aldrich</td><td>M7279</td><td>Gel Monomer component</td></tr><tr><td>N,N-dimethylacrylamide (DMAA)</td><td>Sigma Aldrich</td><td>274135</td><td>Gel Monomer component</td></tr><tr><td>Nunclon 4-Well x 5 mL MultiDish Cell Culture Dish</td><td>Thermo Fisher</td><td>167063</td><td></td></tr><tr><td>Nunclon 6-Well Cell Culture Dish</td><td>Thermo Fisher</td><td>140675</td><td></td></tr><tr><td>Nunc&amp;trade; 15mL&amp;nbsp;Conical</td><td>Thermo Fisher</td><td>339651</td><td></td></tr><tr><td>Nunc&amp;trade; 50mL&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 Scientific</td><td>BP399-1</td><td></td></tr><tr><td>Polyethylene glycol&amp;nbsp; 200</td><td>Sigma Aldrich</td><td>P-3015</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 (SA)</td><td>AK Scientific</td><td>R624</td><td>Gel Monomer component</td></tr><tr><td>Sodium azide</td><td>Sigma Aldrich</td><td>S2002</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>Sodium dodecyl sulfate (SDS)</td><td>Sigma Aldrich</td><td>L3771</td><td>Homogenization Buffer Component</td></tr><tr><td>Tris Base</td><td>Fischer Scientific</td><td>BP152-1</td><td>Homogenization Buffer Component</td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T8787</td><td></td></tr><tr><td>Urea</td><td>Sigma Aldrich</td><td>U5378</td><td>Homogenization Buffer Component</td></tr><tr><td>Xylenes</td><td>Sigma Aldrich</td><td>214736</td><td></td></tr><tr><td>20x SSC</td><td>Thermo Scientific</td><td>AM9763</td><td></td></tr><tr><td>Tween20</td><td>Sigma Aldrich</td><td>P1379</td><td></td></tr><tr><td>poly-L-lysine&amp;nbsp;</td><td>Sigma Aldrich</td><td>P8920</td><td></td></tr></tbody></table>

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.

If the protocol has been successfully completed (Figure 1), the sample will appear clear and flat after heat denaturation; any folding or wrinkling indicates incomplete homogenization. A successfully expanded sample will be 3-4.5-fold larger than before expansion in 1x PBS and 8-11-fold larger when fully expanded in ddH2O. Figure 3 shows example pre- and post-expansion images of 5 &#181;m thick FFPE human kidney sample processed using this protocol an...

Watch this Scientific Journal Video about Universal Molecular Retention with 11-Fold Expansion Microscopy at JoVE.com

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.

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 ...

universal-molecular-retention-with-11-fold-expansion-microscopy

Research

JoVE Journal

Biology

7.1K Views.  Carnegie Mellon University. We proberen eenvoudige en goedkope manieren te vinden voor onderzoekers om hun biologische monsters in beeld te brengen met resoluties onder de diffractielimiet, met name op het gebied van neurowetenschappen en klinische pathologie. Veel recente ontwikkelingen op het gebied van expansiemicroscopie omvatten het aanpassen van gelchemie om monsters verder uit te breiden, het verankeringsmolecuul te veranderen om verschillende doelen te visualiseren en verwarmingsverandering te gebruiken om eiwit...

Video: Auteur in de schijnwerpers: Universele moleculaire retentie met 11-voudige expansiemicroscopie  

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 ...

Developmental Biology

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Bioengineering

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

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.

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, ...

Medicine

Label-Retention Expansion Microscopy (LR-ExM) Enables Super-Resolution Imaging and High-Efficiency Labeling

Expansion microscopy (ExM) is a sample preparation technique that can be combined with most light microscopy methods to increase the resolution. After embedding cells or tissues in swellable hydrogel, samples can be physically expanded three-to sixteen-fold (linear dimension) compared to the original size. Therefore, the effective resolution of any microscope is increased by the expansion factor. A major limitation of the previously introduced ExM is reduced fluorescence after polymerization and the digestion procedure. To overcome this limitation, label-retention expansion microscopy (LR-ExM) has been developed, which prevents signal loss and greatly enhances labeling efficiency using a set of novel trifunctional anchors. This technique allows one to achieve higher resolution when investigating cellular or subcellular structures at a nanometric scale with minimal fluorescent signal loss. LR-ExM can be used not only for immunofluorescence labeling, but also with self-labeling protein tags, such as SNAP- and CLIP-tags, thus achieving higher labeling efficiency. This work presents the procedure and troubleshooting for this immunostaining-based approach, as well as discussion of self-labeling tagging approaches of LR-ExM as an alternative.

Label-Retention Expansion Microscopy LR-ExM Enables Super-Resolution Imaging and High-Efficiency Labeling

A protocol of label retention expansion microscopy (LR-ExM) is demonstrated. LR-ExM uses a novel set of trifunctional anchors, which provides better labeling efficiency compared to previously introduced expansion microscopies.

Expansion microscopy (ExM) is a sample preparation technique that can be combined with most light microscopy methods to increase the resolution. After ...

Ultrastructural Expansion Microscopy in In Vitro Life Cycle of Trypanosoma cruzi

Ultrastructural Expansion Microscopy in Three In Vitro Life Cycle Stages of Trypanosoma cruzi

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial resolution of a cell or tissue for microscopic imaging. This is performed by physically expanding a sample with off-the-shelf chemicals and common lab equipment.
Chagas disease is a widespread and pressing public health concern caused by T. cruzi. The disease is prevalent in Latin America and has become a significant problem in non-endemic regions due to increased migration. The transmission of T. cruzi occurs through hematophagous insect vectors belonging to the Reduviidae and Hemiptera families. Following infection, T. cruzi amastigotes multiply within the mammalian host and differentiate into trypomastigotes, the non-replicative bloodstream form. In the insect vector, trypomastigotes transform into epimastigotes and proliferate through binary fission.The differentiation between the life cycle stages requires an extensive rearrangement of the cytoskeleton and can be recreated in the lab completely using different cell culture techniques. 
We describe here a detailed protocol for the application of U-ExM in three in vitro life cycle stages of Trypanosoma cruzi, focusing on optimization of the immunolocalization of cytoskeletal proteins. We also optimized the use of N-Hydroxysuccinimide ester (NHS), a pan-proteome label that has enabled us to mark different parasite structures.

This study shows a detailed protocol to perform ultrastructure expansion microscopy in three in vitro life cycle stages of Trypanosoma cruzi, the pathogen responsible for Chagas disease. We include the optimized technique for cytoskeletal proteins and pan-proteome labeling.

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial ...

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 ...

Immunological Synapse Formation Between Helper T Cells and Antigen-Presenting Cells

Source: Bello-Gamboa, A. et al., <a href="https://app.jove.com/v/60312">Imaging the Human Immunological Synapse.</a>&#160;J. Vis. Exp.&#160;(2019) This video demonstrates the in vitro study of immunological synapse formation between an antigen-presenting cell and helper T lymphocytes using a fluorescence microscope. T cell receptors of helper T lymphocytes interact with an antigen-MHC-II complex in antigen-presenting cells and form immunological synapses, followed by secretory vesicle movements toward the synapse, which is visualized by fluorescent microscopy.

Source: Bello-Gamboa, A. et al., Imaging the Human Immunological Synapse.&#160;J. Vis. Exp.&#160;(2019) This video demonstrates the in vitro study of ...

JoVE Encyclopedia of Experiments

Immunology

Immune Systems and Components

Assays and Functional Profiling

Bimolecular Fluorescence Complementation-Coupled Photoactivated Localization Microscopy

To prepare a sample for imaging, plate about 5.5 x 104 stable expression cells per well of an 8-well glass-bottomed chamber slide, in 350 microliters of phenol red-free DMEM. Use fresh paraformaldehyde with glutaraldehyde to fix the cells, and after replacing the fixative with PBS or imaging buffer, vortex 100-nanometer gold particles, and add 35 microliters per well for tracking stage drift during imaging.
Approach the microscope and power on the 405 and 561-nanometer lasers. Keeping the shutters closed at this point, ensure the 561-nanometer dichroic mirror and notch filter are in place. Open the image acquisition software.
Turn on the EMCCD camera and allow it to cool down, and set the exposure time to 100 milliseconds and the EMCCD gain to an appropriate value. Add immersion oil to the objective, and secure the sample on the microscope stage. Next, with either brightfield or the 561-nanometer laser, bring the sample into focus.
For imaging Ras and other membrane proteins, use a 60X apochromat TIRF objective with a 1.49 numerical aperture, and bring the microscope into TIRF configuration.
Adjust the excitation laser so that it is off-centered when hitting the back aperture of the TIRF objective, which will cause the laser to deflect upon reaching the coverslip buffer interface. Keep adjusting the laser incident position until the critical angle is reached and the laser is being reflected back.
Search for a cell to image with several gold particles in view. Then, set a region of interest that encloses the cell or a region of it and the gold particles.
In case sufficient activation is already occurring because of high expression levels, begin acquisition with the 405-nanometer laser off and the 561-laser on. Otherwise, turn on the 405-nanometer laser at the lowest factory power setting, and increase it gradually until there are tens of molecules per frame or so that single molecules are well-separated.
As data acquisition continues, gradually increase the 405-nanometer laser power to keep the spot density roughly constant. Continue image acquisition until high 405 power does not initiate any more activation events.

Universal Molecular Retention with 11-Fold Expansion Microscopy

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