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Expansion Pathology (ExPath), Version 1.1Stock SolutionBlocking buffer: MAXblock? Blocking Medium (Active Motif)Staining buffer: MAXbind? Staining Medium (Active Motif)Washing buffer: MAXwash? Washing Medium (Active Motif)Monomer solution:ComponentStock concentration*Amount (mL)Final concentration*Sodium acrylate382.258.6Acrylamide500.52.5N,N′-Methylenebisacrylamide20.500.10Sodium chloride29.2411.7PBS10x11xWater1.15Total9.4***All concentrations in g/100 mL except PBS**9.4/10 mL (1.06x), with the remaining 6% volume brought up by initiator, accelerator and inhibitor.Storage: We have been mixing up the monomer solution at 4°C and storing it at -20°C for long term storage. (We also have seen no issues storing it at 4°C for up to 3 months.) TEMED, ammonium persulfate (APS), and 4-Hydroxy-TEMPO (4HT inhibitor) stock solutions (see below) can be kept at -20 degrees C for at least 6 months. Slice Gelling Solution: Mix the following 4 solutions on ice in the following order:Monomer solutionsTEMED accelerator (from stock, below)4HT inhibitor solution (from stock, below) Ammonium persulfate (APS) initiator solution (from stock, below)The initiator solution needs to be added last, to prevent premature gelation. The mixture should be vortexed to ensure full mixing.Each slice needs ~200?l of gelling solution. For 200?l gelling solution, mix the following:Monomer solution (1.06x) (188?l) (keep at 4°C, throughout, to prevent premature gelation)Inhibitor solution (4?l): 4-hydroxy-TEMPO (4HT stock solution made up at 0.5% in water, final concentration 0.01%, thus, dilution ratio is 1:50), which inhibits gelation to enable diffusion into tissue sectionsAccelerator solution (4?l): TEMED (TEMED stock solution made up at 10% in water, final concentration 0.2%, or 1:50 dilution), which accelerates radical generation by APSInitiator solution (4?l): APS (APS stock at 10%, final concentration 0.2%, or 1:50 dilution), which initiates the gelling process, so it needs to be added lastDigestion Buffer (can be stored as aliquots in the fridge at 4°C): 50 mM Tris pH 8.025 mM EDTA 0.5% Triton X-1000.8 M NaClAdd: Proteinase K (Molecular Biology Grade NE Biolabs, P8107S, 1:200, final concentration 4 units/mL) to digestion buffer before use. (There is probably no difference in results between 8 unit/mL proteinase K, as used in our previously published protocols, or 4 units/mL which saves money. But using 8 units/mL will not cause any harm. It may be possible to use even less than 4 units/mL, however.)For the expansion step, the cut gel is placed in a 6-well multi-well black-walled plate with clear bottom (, Cat# P06-1.5H-N)ExPath procedures for clinical archived tissue slidesStep 1, Sample pre-processingFor formaldehyde-fixed paraffin-embedded (FFPE) clinical samples, place sample in a series of solutions sequentially, for 3 mins for each step: 2× xylene, 2× 100% ethanol, 95% ethanol, 70% ethanol, 50% ethanol, and finally doubly deionized water. Do all the steps at room temperature (RT), 3 mins each.For stained and mounted permanent slides, place samples briefly placed in xylene. Then remove coverslips carefully with appropriate tools, such as a razor blade. If the coverslip is difficult to remove, further incubate the slides in xylene at RT until the coverslip is loosened. Then wash slides in a series of solutions sequentially: 2× 100% ethanol, 95% ethanol, 70% ethanol, 50% ethanol, and finally doubly deionized water. Do all the steps at RT, 3 mins each.Note: For H&E stained slides, hematoxylin and eosin are eliminated during the expansion process.For unfixed frozen tissue slides in optimum cutting temperature (OCT) solution (Tissue-Tek), fix the tissues for 10 min in cold acetone at -20 oC before washing with 1x PBS solution 3 times for 10 min at RT. For already fixed, frozen clinical tissue sections, leave slides at RT for 2 mins to let the OCT, melt and then wash 3x with PBS solution at RT for 5 min each. Step 2, Sample heat treatment. Place specimens in 20 mM sodium citrate solution (pH 8, 100oC) in a heat-resistant container, and then transfer the whole container to a 60 oC incubation chamber for 30 mins.Step 3, Primary antibody staining. This is similar to a typical staining protocol (immunofluorescence (IF)/immunohistochemistry (IHC)):Treat the tissue on slides with the blocking buffer, for 1 hour at 37 °C. (Note: alternatively, the incubation could be at RT for 2 hours or 4 °C overnight, according to the product manual.)Incubate the tissue with primary antibodies in staining buffer, for 3 hours at RT or 37 °C, or overnight at 4°C, depending on the antibodies being used. (Note: suggested incubation periods and temperatures are given as a guide only. It is recommended that the user optimize these parameters for use in their own experiment. Also note, the sample needs to be placed in a humidified container during this incubation period, to prevent drying out.)Wash the tissue with washing buffer, 3 times, ~10 min each, at RT.Step 4, Secondary antibody stainingIncubate the tissue on slides with secondary antibodies at a concentration of approximately 10 ?g/mL together with 300 nM DAPI (if desired; DAPI can be obtained from Thermo Fisher Scientific) in the staining buffer, for at least 1 hour at RT or 37 °C (Note: suggested incubation periods and temperatures are given as a guide only. It is recommended that the user optimize these parameters for use in their own experiment), for 5 ?m thick tissue (further optimization of incubation duration or temperature may be needed for thicker tissues).Note: Do not use cyanine dyes (Cy3, Cy5, Alexa 647) on your secondary antibodies (since the ExM protocol is not compatible with applying these pre-polymerization). We suggest Alexa 488 for green staining, Alexa 546 for orange-red staining, and Atto 647N or CF633 (the latter from Biotium) for far-red staining.Wash the tissue on slides with washing buffer, 3 times, ~10 min each, at RT.Step 5, Anchoring treatmentCover tissue section with 1x PBS and take pre-expansion images on a microscope, so that expansion factor and thus biological units of length can be established later. Dissolve Acryloyl-X, SE (Life Technologies, A20770) in 500?L anhydrous DMSO, to result in a 10 mg/mL stock solution. Aliquot this stock solution in 20 uL aliquots, and store in a desiccated environment at -20 C.Adjust AcX to a concentration of 0.03-0.1 mg/ml (that is, 0.03 mg/ml for samples fixed with non-aldehyde fixatives; 0.1 mg/ml for samples fixed with aldehyde fixatives) in 1x PBS. We roughly added 250 microliters to each slide.Treat stained slices for >3 hours at RT (if desired, this reaction can be run overnight).Step 6, GellingMake sure to remove excess solution from the tissue section before quickly adding fresh, cold gelling solution. Place gelling solution on top of the tissue section and make sure the whole tissue section is immersed in the solution. (One can use a hydrophobic pen to draw a hydrophobic boundary around the tissue section, to confine the solution and prevent leaking.) Incubate the mixture on the tissue for 30 mins at 4 °C. Use freshly prepared gelling solution, immediately after adding APS at 4 °C. Make sure at least a 100-fold excess volume of monomer solution is used, e.g., ~200 ?l of gelling solution for each tissue section on the slide. Gel chambers are constructed by sandwiching the slice between a slide and a coverslip, with spacers on either side of the tissue section to prevent compression of the tissue slice (see schematic below, Figure 1). Spacers are made from cut coverslips using a diamond knife. For most human tissue sections in clinical settings (5-10 ?m thick), pieces of cover glass (VWR micro cover glass, 24x60mm, No.1 or 1.5) can be used for spacers. Avoid air bubbles trapped inside the chamber.Assemble the tissue slide into a gel chamber (Figure 1) as described in the previous paragraph, and then incubate at 37°C in a humidified environment for 2 hours.After incubation you can proceed to digestion and expansion (below) or place the slide chamber at 4°C inside a Petri dish (or other container as appropriate) sealed with Parafilm for storage. Frost Glass SlideGel Chamber (side view)Top CoverglassSpacer (Coverglass #1/1.5)5 ?m Tissue sectionFrost Glass SlideGel Chamber (side view)Top CoverglassSpacer (Coverglass #1/1.5)5 ?m Tissue sectionFigure 1. Gel chamber schematic.Step 7, Digestion and expansion Take off the top cover of the gel chamber using a razor blade placed at the edge of the coverslip, sliding the blade along the coverslip side touching the gel surface and then gently using the blade to lift the coverslip off the gel surface. Trim the tissue-containing-gel to minimize volume, using a sharp razor blade, and cut a corner in an off-angle fashion for tracking of orientation throughout later steps (when the gel is transparent and orientation sometimes hard to gauge).Submerge the gel, still on the slide, in 3 mL of freshly made digestion buffer (including proteinase K), then incubate for 3 hours at 60 °C. (Make sure to completely submerge the slice in the digestion buffer, and make sure it does not dry out by sealing the container with Parafilm. Any container with a cover, such as a small slide box, or a plastic well, can be used to incubate the gel and digestion buffer.) Normally the sample will detach from the glass slide by itself after digestion. If needed, use a razor blade to gently shave the sample off the slide into a well of a 6-well plate (CellVis) filled with PBS (Note: If the gel is already floating in the digestion buffer, use a fine, soft paint brush to pick it up and transfer it into a well of a 6-well plate filled with PBS.) The user can use side-illumination from a green LED light source, to help visualize the cut pieces, if needed. Make sure to place each piece into the well with the tissue side down. If the gel piece is flipped over, add PBS and slide the small paint brush gently underneath the gel piece, and flip it over in the well. Wash the samples once with 1x PBS buffer for 10 min at RT and stain with 300 nM DAPI in PBS buffer for 20 mins at RT, then wash them once with 1x PBS for 10 min at RT. For expansion, remove the PBS and wash the samples with excess volume of ddH2O (we usually use at least 10x the final gel volume; optionally, one can add to the water 0.002% - 0.01% NaN3 to prevent bacterial growth, although the final expansion factor is reduced by 10%), 3-5 times, for 10 minutes each time at RT. Slice expansion should reach a plateau after about the 3rd or 4th wash. Note: the expansion chamber needs to be of adequate size for the sample. You might need to trim the gel. The sample might need to be trimmed into smaller pieces if no chamber of proper size can be obtained. In general, an expanded gel containing a tissue with diameter less than 0.6 cm pre-expansion fits nicely in a glass bottom 6 well plate. Gels can be immobilized with 1.5-2% low melt agarose in water to prevent drift during imaging. Final step: image with conventional fluorescent, confocal microscope, or other desired scopesOption: Expansion immunoFISH. For ExPath samples being processed for immunohistochemistry followed by expansion and then DNA FISH probing:Place digested gel samples in a hybridization buffer made of 1× PBS, 15% ethylene carbonate, 20% dextran sulfate, 600 mM NaCl and 0.2 mg/ml single stranded salmon sperm DNA at 85°C for 30 minsMix the samples with 30 ?L of hybridization buffer containing user-selected SureFISH probes (Agilent/Dako) pre-heated at 85 °C for 10 mins. Incubate the mixtures at 45°C overnight. Wash the samples with stringency wash buffer made of 1×SSC (150 mM NaCl, 15 mM sodium citrate, pH 7.0) and 20% ethylene carbonate at 45°C for 15 min, followed by washes with 2× SSC at 45 °C, 3 times for 10 mins each. Finally, wash the gel samples with 0.02× SSC multiple times at RT (5 min each) until the expansion is completed (compared to the gel samples expanded in water, the size of the same gel sample is reduced by about 20%, because of the salt of the 0.02x SSC). ................
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