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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A., Shroff, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+microscopy:+A+concise+guide+to+current+imaging+methods.">Fluorescence microscopy: A concise guide to current imaging methods.</a> Current Protocols in Neuroscience. 79, 1-25 (2017).</li><li>Schermelleh, L., Heintzmann, R., Leonhardt, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+super-resolution+fluorescence+microscopy.">A guide to super-resolution fluorescence microscopy.</a> Journal of Cell Biology. 190 (2), 165-175 (2010).</li><li>Chen, F., Tillberg, P. W., Boyden, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expansion+microscopy.">Expansion microscopy.</a> Science. 347 (6221), 543-548 (2015).</li><li>Tillberg, P. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein-retention+expansion+microscopy+of+cells+and+tissues+labeled+using+standard+fluorescent+proteins+and+antibodies.">Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies.</a> Nature Biotechnology. 34 (9), 987-992 (2016).</li><li>Chozinski, T. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expansion+microscopy+with+conventional+antibodies+and+fluorescent+proteins.">Expansion microscopy with conventional antibodies and fluorescent proteins.</a> Nature Methods. 13 (6), 485-488 (2016).</li><li>Ku, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiplexed+and+scalable+super-resolution+imaging+of+three-dimensional+protein+localization+in+size-adjustable+tissues.">Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues.</a> Nature Biotechnology. 34 (9), 973-981 (2016).</li><li>Chen, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanoscale+imaging+of+R.N.A.+with+expansion+microscopy.">Nanoscale imaging of R.N.A. with expansion microscopy.</a> Nature Methods. 13 (8), 679-684 (2016).</li><li>Tsanov, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SmiFISH+and+FISH-quant+-+A+flexible+single+R.N.A.+detection+approach+with+super-resolution+capability.">SmiFISH and FISH-quant - A flexible single R.N.A. detection approach with super-resolution capability.</a> Nucleic Acids Research. 44 (22), 165 (2016).</li><li>Wang, G., Moffitt, J. R., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiplexed+imaging+of+high-density+libraries+of+R.N.A.s+with+MERFISH+and+expansion+microscopy.">Multiplexed imaging of high-density libraries of R.N.A.s with MERFISH and expansion microscopy.</a> Scientific Reports. 8 (1), 4847 (2018).</li><li>Wen, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+direct+grafting+strategies+via+trivalent+anchoring+for+enabling+lipid+membrane+and+cytoskeleton+staining+in+expansion+microscopy.">Evaluation of direct grafting strategies via trivalent anchoring for enabling lipid membrane and cytoskeleton staining in expansion microscopy.</a> ACS Nano. 14 (7), 7860-7867 (2020).</li><li>Sun, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Click-ExM+enables+expansion+microscopy+for+all+biomolecules.">Click-ExM enables expansion microscopy for all biomolecules.</a> Nature Methods. 18, 107-113 (2021).</li><li>White, B. M., Kumar, P., Conwell, A. N., Wu, K., Baskin, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+expansion+microscopy.">Lipid expansion microscopy.</a> Journal of the American Chemical Society. 144 (40), 18212-18217 (2022).</li><li>Truckenbrodt, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=X10+expansion+microscopy+enables+25-nm+resolution+on+conventional+microscopes.">X10 expansion microscopy enables 25-nm resolution on conventional microscopes.</a> EMBO Reports. 19 (9), e45836 (2018).</li><li>Chang, J. -. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Iterative+expansion+microscopy.">Iterative expansion microscopy.</a> Nature Methods. 14 (6), 593-599 (2017).</li><li>Park, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epitope-preserving+magnified+analysis+of+proteome+(eMAP).">Epitope-preserving magnified analysis of proteome (eMAP).</a> Science Advances. 7 (46), (2021).</li><li>Klimas, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnify+is+a+universal+molecular+anchoring+strategy+for+expansion+microscopy.">Magnify is a universal molecular anchoring strategy for expansion microscopy.</a> Nature Biotechnology. , (2023).</li><li>Gao, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expansion+stimulated+emission+depletion+microscopy+(ExSTED).">Expansion stimulated emission depletion microscopy (ExSTED).</a> ACS Nano. 12 (5), 4178-4185 (2018).</li><li>Zwettler, F. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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)&#160;</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 
			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&#8242;,N&#8242;- 
			Tetramethylethylenediamine (TEMED)</td>
			<td>Sigma Aldrich</td>
			<td>T9281</td>
			<td>Accelerator</td>
		</tr>
		<tr>
			<td>N,N&#8242;-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&#8482; 15mL&#160;Conical</td>
			<td>Thermo Fisher</td>
			<td>339651</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc&#8482; 50mL&#160;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&#160; 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&#160;</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

7.0K Views.  Carnegie Mellon University. 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.

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

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current state-of-the-art cryoET combined with subtomogram averaging analysis enables the high-resolution structural determination of macromolecular complexes that are present in multiple copies within tomographic reconstructions. Tomographic experiments usually require a vast amount of tilt series to be acquired by means of high-end transmission electron microscopes with important operational running-costs. Although the throughput and reliability of automated data acquisition routines have constantly improved over the recent years, the process of selecting regions of interest at which a tilt series will be acquired cannot be easily automated and it still relies on the user's manual input. Therefore, the set-up of a large-scale data collection session is a time-consuming procedure that can considerably reduce the remaining microscope time available for tilt series acquisition. Here, the protocol describes the recently developed implementations based on the SerialEM package and the PyEM software that significantly improve the time-efficiency of grid screening and large-scale tilt series data collection. The presented protocol illustrates how to use SerialEM scripting functionalities to fully automate grid mapping, grid square mapping, and tilt series acquisition. Furthermore, the protocol describes how to use PyEM to select additional acquisition targets in off-line mode after automated data collection is initiated. To illustrate this protocol, its application in the context of high-end data collection of Sars-Cov-2 tilt series is described. The presented pipeline is particularly suited to maximizing the time-efficiency of tomography experiments that require a careful selection of acquisition targets and at the same time a large amount of tilt series to be collected.

The increasing demand for large-scale data collection in cryogenic electron tomography requires high-throughput image acquisition routines. Described here is a protocol that implements the recent developments of advanced acquisition strategies aimed at maximizing the time-efficiency and throughput of tomographic data collection.

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current ...

Longitudinal Intravital Microscopy Using a Mammary Imaging Window with Replaceable Lid

The branched structure of the mammary gland is highly dynamic and undergoes several phases of growth and remodeling after birth. Intravital microscopy in combination with skin flap surgery or implantation of imaging windows has been used to study the dynamics of the healthy mammary gland at different developmental stages. Most mammary imaging technologies are limited to a time frame of hours to days, whereas the majority of mammary gland remodeling processes occur in time frames of days to weeks. To study mammary gland remodeling, methods that allow optical access to the tissue of interest for extended time frames are required. Here, an improved version of the titanium mammary imaging window with a replaceable lid (R.MIW) is described that allows high-resolution imaging of the mammary gland with a cellular resolution for up to several weeks. Importantly, the R.MIW provides tissue access over the entire duration of the intravital imaging experiment and could therefore be used for local tissue manipulation, labeling, drug administration, or image-guided microdissection. Taken together, the R.MIW enables high-resolution characterization of the cellular dynamics during mammary gland development, homeostasis, and disease.

This protocol describes a novel mammary imaging window with a replaceable lid (R.MIW). Intravital microscopy after implantation of the R.MIW allows for longitudinal and multi-day imaging of the healthy and diseased mammary gland with a cellular resolution during the different developmental stages.

The branched structure of the mammary gland is highly dynamic and undergoes several phases of growth and remodeling after birth. Intravital microscopy in ...