Brain Clearing and Expansion Workshop University of British Columbia Koerner Pavilion 1st floor conference centre July 12th 2017 Sponsored by: Leica Microsystems, special thanks to the UBC faculty of medicine for supporting the “Dynamic Brain Circuits and Connections in Health and Disease. Organizers Tim Murphy and Ann Marie Craig. Special help from UBC trainees Matilde Balbi, Claire Bomkamp, and Eli York Faculty Josh Vaughan UW, Kwanghun Chung MIT, and Jonathan Epp Univ. Calgary.
Transcript
Brain Clearing and Expansion Workshop University of British Columbia Koerner Pavilion 1st floor conference centre July 12th 2017
Sponsored by: Leica Microsystems, special thanks to the UBC faculty of medicine for supporting the “Dynamic Brain Circuits and Connections in Health and Disease. Organizers Tim Murphy and Ann Marie Craig. Special help from UBC trainees Matilde Balbi, Claire Bomkamp, and Eli York Faculty Josh Vaughan UW, Kwanghun Chung MIT, and Jonathan Epp Univ. Calgary.
Brain Clearing and Expansion Workshop University of British Columbia July 12th, 2017
9:00 –9:05 AM Tim Murphy UBC and Ann Marie Craig UBC welcome to workshop.
9:00-10:15 AM Kwanghun Chung MIT CLARITY, SHIELD, SWITCH, and MAP and how to combine these
techniques. CLARITY example protocol detail line by line (why you do certain steps).
10:15-10:30AM Coffee break
10:30-11:15 Josh Vaughan Univ. of Washington Expansion microscopy and super-resolution techniques
11:15-12:00 Jonathan Epp Univ. of Calgary Brain-wide analysis approaches and open source clearing
methods
12:00-1:00 lunch conference room Koerner
1:00-4:00 form 3 groups and rotate through cleared samples Kwanghun Chung lab (Demo Group 1),
software for wide scale analysis Epp (Demo Group 2), and expansion demo Vaughan (Demo Group 3),
after 1 h switch to next demo.
Start in demo Group 1 Jan-Apr birthdays, Group 2 May-Aug, and Group 3 Sept-Dec.
A station will also be available to examine commercial Brain Clearing systems LifeCanvas (Jeff Stillman).
4:00-5:00 wrap-up and questions everyone together, probably the most informative part!
Take elevator from ground up to 1st floor follow signs to double doors for imaging center
Chung lab CLARITY protocol
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CLARITY Protocol Please check www.chunglabresources.org for more information.
ORIGINAL ARTICLES Sung-Yon Kim*, Jae Hun Cho*, Evan Murray, Naveed Bakh, Heejin Choi, Kimberly Ohn, Sara Vassallo, Luzdary Ruelas, Austin Hubbert, Meg McCue, Philipp Keller and Kwanghun Chung. Stochastic electrotransport selectively enhances the transport of highly electromobile molecules, PNAS, 2015 Nov 17: 112(46): E6274-83. doi: 10.1073/pnas.1510133112. Epub 2015 Nov 2. PubMed PMID: 26578787; PubMed Central PMCID: PMC4655572. Kwanghun Chung, Jenelle Wallace, Sung-Yon Kim, Sandhiya Kalyanasundaram, Aaron Andalman, Tom J. Davidson, Kelly A. Zalocusky, Joanna Mattis, Sally Pak, Viviana Gradinaru, Hannah Bernstein, Julie Mirzabekov, Charu Ramakrishnan, and Karl Deisseroth, Structural and molecular interrogation of intact biological systems, Nature, 2013, 497, 332-337
RELEVANT ARTICLES Taeyun Ku*, Justin Swaney*, Jeong-Yoon Park*, Alexander Albanese, Evan Murray, Jae Hun Cho, Young-Gyun Park, Vamsi Mangena, Jiapei Chen, and Kwanghun Chung. Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues, Nature Biotechnology, 2016, doi:10.1038/nbt.3641. Evan Murray*, Jae Hun Cho*, Daniel Goodwin*, Taeyun Ku*, Justin Swaney*, Sung-Yon Kim, Heejin Choi, Jeong-Yoon Park, Austin Hubbert, Meg McCue, Young-Gyun Park, Sara Vassallo, Naveed Bakh, Matthew Frosch,, Van J. Wedeen, H. Sebastian Seung, and Kwanghun Chung. Simple, scalable proteomic imaging for high-dimensional profiling of intact systems, Cell, Dec 3:163(6): 1500-14. doi: 10.1016/j.cell.2015.11.025. PubMed PMID: 26638076.
REAGENTS Anesthetics Beuthanasia-D (Schering-Plough Animal Health Corp.) Hydrogel Monomer Solution 32% Paraformaldehyde (Electron Microscopy Sciences, #15714-S) 40% Acrylamide Solution (Bio-Rad, #161-0140) Azo-initiator (Wako, #VA-044) 10X PBS (Invitrogen, #70011-044) UltraPure Distilled Water (Invitrogen, #10977-015) Caution: PFA is a hazardous chemical. All work involving PFA must be conducted in a certified fume hood and in compliance with governmental/institutional regulations. SDS Clearing Solution
Boric Acid (Sigma-Aldrich, #B7901) Sodium Dodecyl Sulfate (Sigma-Aldrich, #L3771) Lithium Hydroxide Monohydrate (Sigma-Aldrich, #254274)) Washing Solution Triton-X (Sigma-Aldrich, #T8787) Sodium Azide (Sigma-Aldrich, #S2002) 1X PBS (Invitrogen, #10010-023) Optical clearing solution This solution consists of 23.5% (w/v) n-methyl-d-glucamine, 29.4% (w/v) diatrizoic acid, and 32.4% (w/v) iodixanol in water. Use a stir bar (or shake if necessary) to fully dissolve the powders at each step. Do not use heat when mixing the solution, as this will cause a color change. This solution should be stored carefully to ensure that no water is lost, as just a small amount of evaporation will result in precipitation. Teflon tape can be used to increase the security of the bottle’s seal, and parafilm can be used around the cap. It may be necessary to use a 60% iodixanol solution (see reagents list) rather than iodixanol powder, as it is not cheaply available. Therefore, an example recipe for the optical clearing solution is as follows: Dilute 60% iodixanol solution to 47% iodixanol. To achieve this, roughly 2.75 mL water should be added for every 10 mL of 60% iodixanol solution. Then, for every 10 mL of the resulting solution, 3.39 g n-methyl-d-glucamine and 4.24 g diatrizoic acid should be added in order. Be sure to take into account that the final volume will be significantly larger than the starting volume. In addition to our optical clearing solution (termed PROTOS), there are proprietary optical clearing solutions available as well as other published recipes that other groups have developed for their tissue clearing protocols. The proprietary formulations are prohibitively expensive (RapiClear and FocusClear) and at least FocusClear is known to result in the formation of precipitate within samples during long-term storage. PROTOS is the most cost-effective option for high quality optical clearing, which is an absolute must for thick tissue imaging. CUBIC-mount (Lee, 2016) and sRIMS (Yang, 2014) are cheaper than PROTOS, but they are noticeably less effective. Solution Recipe Cost per 500
mL PROTOS 29.4% diatrizoic acid $565.80 23.5% n-methyl-d-
Premade PROTOS is available from LifeCanvas technologies (EasyIndex, http://www.lifecanvastech.com/) Tissue In principle, any tissue type from any animals of any ages with or without fluorescence can be used. In the previous paper (Chung et al., 2013), we demonstrated that CLARITY is compatible with whole adult mouse brain, whole adult zebrafish brain and even extensively formalin-fixed postmortem human brain section (without the perfusion step and further optimization in this case). Tissues with strong fluorescent protein expression can undergo CLARITY processing described in this protocol and then can be directly imaged; tissues without fluorescent proteins can be labeled with antibodies or RNA probes (Chung et al., 2013) for subsequent imaging. Caution: Work with animals must be conducted in compliance with governmental and institutional regulations. EQUIPMENT Transcardial perfusion of fixatives and hydrogel monomers Dissection board (styrofoam lid is fine) 20 ml syringes with luer lock ends (Fisher Scientific, 14-820-19) 1 ml syringes (Terumo, SS-01T) Winged infusion sets (Terumo, SV-25BLK) Needles (Fisher Scientific, BD 305109) Absorbant pads (VWR, 56616-032) 50 mL Falcon tubes (BD Falcon, #352070) Guillotine, for sacrificing larger animals (Kent Scientific, #DCAP) Surgical scissors (Fine Science Tools, #14130-17) Fine scissors (Fine Science Tools, #14137-10) Hemostats (Fine Science Tools, #13011-12) Forceps (Fine Science Tools, #11050-10, #11251-10) Spatula (Fine Science Tools, #10092-12) Hydrogel-tissue hybridization To build your own hydrogel-tissue hybridization system, you need the following items.
Dessicator with 3-way stopcock (VWR, #24988-197) Vacuum Pump (Buchi, #071000) Compressed Nitrogen Tank (AirGas, #NI UHP300)
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Compressed Gas Tank Pressure Regulator (AirGas, #Y11215B580) Teflon Tape (McMaster-Carr, #4591K11) 3/8” tubing (McMaster-Carr, #5155T36) 3/8” to 1/4" barbed tubing connector (McMaster-Carr, #5463K633)
All-in-one system is available from LifeCanvas (www.lifecanvastech.com, EasyGel)
ETC clearing To build your own ETC clearing system, you need the following items.
Buffer Filter with Light-Blocking Blue Bowl (McMaster-Carr, #4448K35) Platinum wire with 0.5mm diameter (Sigma, #267201) Bottle for Chamber fabrication (Nalgene via Amazon, #2118-0002) Nalgene Straight Side Jar – Poly, 32oz (Nalgene via Amazon) Single barbed tube fitting (7/16” hex for 1/4” tubing) (McMaster-Carr, #5463K245) Tube to tube coupling for 3/32” to 1/16” tubing (McMaster-Carr, #5117K51) 3M Duo adhesive dispenser (McMaster-Carr, #7467A43) 3M Duo adhesive-mixing applicators (McMaster-Carr, #7467A12) 3M Duo adhesive cartridges (McMaster-Carr, #746A17) Sample holder (BD Falcon, #352340) Bio-Rad HC PowerPac System (Bio-Rad, #164-5052) Banana to Large Alligator Test Lead Set (Elenco, #TL16) Clear 1/4" tubing (Mcmaster, #5155T26) Clear 5/8” tubing (McMcaster, #5155T46) 1/4" wye connector (McMaster, #53055K155) 4x Chemical resistant stopcock 1/4" to 1/4" (Mcmaster, #48285K24) 5/8" to 1/4" tubing connection (McMaster-Carr, #2974K271) Elbow connection 1/4" male pipe to 1/4" barbed fitting (McMaster-Carr, #5463K489) Elbow connection 1/4" barbed fitting (McMaster-Carr, #5463K594) Rubber Grounding Plug (Leviton via Amazon, #L00-515PR-000) Magnetic Water Pump (Pan World via Premium Aquatics, #NH-10PX)
All-in-one system with advanced features is available from LifeCanvas (www.lifecanvastech.com, SmartClear) Imaging KWIK-SIL (World Precision Instruments, #KWIK-SIL) Willco-Dish (Ted Pella, #14032-120) Blu-Tack Reusable Adhesive (Blu-Tack via Amazon) REAGENT SETUP Hydrogel Monomer Solution Keeping all reagents on ice, prepare a 10% stock solution of initiator by dissolving 1 g of initiator in 10 mL UltraPure water. Using this stock, prepare a solution (w/w) of 4% acrylamide, 0.25% initiator, 1X PBS, and 4% PFA in UltraPure water. For 40 mL, this is 26 mL UltraPure Water, 4 mL 40% acrylamide solution, 1 mL initiator solution, 4 mL 10X PBS, and 5 mL 32% PFA. For
each tissue sample being processed, 80 mL will be needed. Always add water first to keep the other components dilute, and add the reagents in the order listed here. The solution can be stored at -20° C indefinitely. CAUTION: Make sure to keep all reagents and the final solution on ice at all times. The hydrogel polymerization reaction is triggered by heat. SDS Clearing Solution Create a solution consisting of 20 mM lithium hydroxide monohydrate and 200 mM SDS. Next, add boric acid until the pH reaches 8.5. PBST Make a solution consisting of 0.1% Triton-X and 0.1% Sodium Azide using 1X PBS. For a 500mL PBS bottle, this is 500 μL Triton-X and 500 mg Sodium Azide. EQUIPMENT SETUP Hydrogel-Tissue Hybridization Mount the nitrogen tank with an appropriate tank bracket and attach the regulator to the tank outlet, using Teflon tape if necessary to prevent leaking. Run 3/8” tubing from the regulator outlet to the stopcock of the desiccator using a 3/8” to 1/4” barbed tube fitting. Connect the vacuum pump to the desiccator by simply connecting the supplied tubing to the barbed fitting on the stopcock. ETC System Create the measurement reservoir in a similar manner using a 32oz Nalgene bottle and 1/4" barbed elbow connectors. Be sure to place the connections on opposite sides of the bottle, and angle them slightly to maximize mixing in the chamber. Apply epoxy to both the inside and outside parts of the connection and allow to dry overnight. Caution: Don’t allow any epoxy to enter the tubing connectors, as this will impede flow within the system. Create 2 more holes in the lid of the bottle, large enough for insertion of a pH probe and a thermometer for data acquisition. These holes should be left unsealed. Create a heat exchange module by measuring out two pieces of around 2 ft of 1/4" tubing. Connect these to the system in parallel using wye connectors and submerge in water. Connect the water filter to the system using 1/4" male pipe thread to 1/4" tubing elbow connectors. Tube all the components of the system together using 1/4" tubing, though 5/8” tubing will be needed for the pump connection. A reducing fitting should be used to connect this larger tubing to the rest of the system. Critical: The system should be connecting in the following order (in the direction of flow): Pump, Filter, ETC Chamber, Measurement Reservoir, Heat Exchanger. Placing the measurement reservoir direction after the ETC chamber allows for direct readouts of the temperature and pH as they are in the ETC chamber. It is also important that the reservoir is only separated from the pump inlet by tubing, as the reservoir is necessary to start the system. If desired, drain valves can be created using wye connectors and stopcocks and placed between any elements of the system. Additionally, when connecting the system with 1/4" tubing, 1/4" stopcocks should be
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added to the system on either side of the ETC chamber, so that it can periodically be isolated from the system to check the samples without draining the entire system. PROCEDURES Perfusion and Tissue Preparation
1. Make a fresh batch of hydrogel monomer solution, or thaw frozen stock solution at 4 ⁰C or on ice. After the solution is completely thawed and transparent (but still ice-cold), gently invert to mix. Ensure no precipitation or bubbles are seen in the solution.
2. Deeply anesthetize an animal with Beuthanasia-D (0.5 ml per 1 kg of body weight intraperitoneally) and surgically open the chest cavity with a midline abdominal incision that bifurcates rostrally into a Y-shape. Punch a small hole in the right atrium and insert an injection needle into the left ventricle to allow perfusion.
Caution: Experiments involving animals must be conducted in accordance with governmental and institutional regulations. Animals must be fully anesthetized before making incisions: deep anesthesia can be confirmed by absence of corneal reflex (eye blink) or by any other overt signs of response to physical stimuli. 3. Prepare two syringes filled with ice-cold PBS and hydrogel monomer solution, respectively, each with winged needle sets for each solution. In the case of mouse, perfuse first with 20 mL of ice-cold phosphate-buffered saline (PBS) at a rate of less than 5 mL/min, carefully take the needle out and perfuse with 20 mL of the ice-cold hydrogel solution. Rats require about 200 mL of each solution at the rate of 20 mL/min.
Critical Step: Maintain a slow rate of perfusion: we found that injecting less than 5 mL per minute for both solutions in the case of mouse yields better results. Use extreme caution not to introduce bubbles to the vasculature (especially when introducing needles), as this decreases the quality of perfusion. 4. Carefully harvest the organs of interest and place them immediately in a 50 mL conical tube containing 20 mL of the ice-cold hydrogel monomer solution for both post-fixation and even infiltration of monomers. Keep this on ice until it can be transferred to a 4 ⁰C refrigerator.
Caution: always keep the temperature low to prevent thermal initiation of the hydrogel-formation reaction.
5. Incubate the sample for 1 day at 4 ⁰C to allow for further distribution of monomer and initiator molecules throughout the tissue.
Caution: if the sample contains fluorophores, cover the tube containing the sample in aluminum foil to prevent photobleaching.
Caution: If the tissues are left in the hydrogel solution for more than one day, enough protein will diffuse out of the tissue to act as a cross-linker, causing rigid gel to form around the sample. This will result in a slower rate of lipid clearing. Critical Step: Uniform penetration of monomers throughout the tissue is critical for 1) even polymerization throughout the tissue and 2) keeping the macro- and microstructure intact. Parts
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of the region of cellular structures that are not infiltrated with monomers may not be bound to the hydrogel mesh even after hybridization, and subsequent electrophoresis will result in loss of the unbound biomolecules. Furthermore, uneven distribution of monomers may cause anisotropic expansion and reduction in volume during the electrophoretic tissue clearing and refractive index matching steps.
Hydrogel Tissue Embedding
6. After the tissues have been allowed to incubate in the hydrogel monomer solution for 1 day, move the samples to 10 mL of fresh hydrogel monomer solution. The tubes that the tissues are transferred to should have Teflon tape applied to them BEFORE the solution is added.
7. Place the conicals in a desiccation chamber on a tube rack and unscrew the caps about halfway. The desiccator should have a 3-way stopcock. Removal of oxygen is necessary for hydrogel-tissue hybridization because oxygen radicals may terminate the polymerization reaction. Critical Step: If the caps are not unscrewed, there will be no gas exchange in the desiccator and oxygen will not be removed from the conicals. 8. Connect nitrogen gas and a vacuum pump to the desiccator via the 3-way stopcock. Open flow in all three directions and turn on the nitrogen gas. Allow the gas to flow for about 5 seconds. This step is necessary to flush oxygen from all the tubing in the system. 9. Without turning off the nitrogen flow, turn on the vacuum pump and adjust the stopcock so that flow is only open to the desiccator and the vacuum pump. Allow the vacuum pump to run for at least 10 minutes. The nitrogen should not be shut off because the tubing is gas-permeable. If nitrogen flow is stopped, oxygen will diffuse back into the tubing. 10. Turn the stopcock VERY SLOWLY so that flow is only open to the nitrogen gas and the desiccator, and then turn off the vacuum pump. Allow the desiccation chamber to fill with nitrogen gas. 11. Very quickly, lift the lid of the desiccator and tighten the caps of the conicals inside. It helps to have two people—one to hold the lid slightly open and another to close the tubes. The nitrogen gas can now be shut off. Critical Step: If the lids are not closed quickly enough, oxygen will re-enter the conicals and impede the polymerization reaction. If at this stage you find that the lids were already closed, open them slightly and repeat the de-gassing procedure. 12. Gently shake the samples in a 37 °C warm room for 2 hours. This temperature will trigger radical initiation by the azo-initiator. 13. To remove unreacted PFA, wash the samples in 50 mL of clearing solution at 37 °C for 24 hours, with gentle shaking. Do this a total of three times. Caution: This clearing solution with PFA must be discarded as hazardous waste according to government and institutional regulations.
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Pause Point: Tissues may be stored in clearing solution indefinitely following this step. If the sample contains fluorescence, be sure to cover with aluminum foil. Electrophoretic Tissue-Clearing
14. At this point, you should have already constructed an ETC system as detailed in the section EQUIPMENT SETUP. Add the sample to the ETC chamber and close the lid. Connect any remaining unconnected tubing. 15. Fill the system with clearing solution by first filling the measurement reservoir and placing it on a surface a few inches higher than the level of the heat exchanger and pump. This will allow buffer to fill the tubing. Start the pump and add more buffer to the measurement reservoir as needed to fill the system. 16. Connect the electrodes to the lead cables and start the power supply. Use around 40 V. Caution: Never start the power supply unless you have confirmed that the flow rate is satisfactory. The flow rate can be adjusted by slightly turning one of the stopcocks that surrounds the ETC chamber. A high flow rate may result in physical damage to the tissue, whereas low flow rate may result in inadequate cooling and damage the sample. Be sure to stop the voltage before stopping the pump when you shut the system off. Caution: pH below 7 and temperatures above 37 °C can result in loss of fluorescence and damage to the tissue. Be sure to check the system regularly to ensure that the temperature is not too high and that the pH has not dropped below about 7.3. If the pH is low, drain the current buffer and add new clearing solution. If the temperature is too high, lower the voltage to reduce resistive heating. 17. Check the samples regularly to determine that the system is working properly and that clearing is progressing. The entire process should take several days. 18. Remove the cleared samples from the ETC system and wash them twice with PBST for 24 hours each. Pause Point: Samples can be stored indefinitely in PBST at room temperature. 19. Place the sample in a volume of PROTOS that is sufficient to cover the tissue completely and allow it to incubate for 2 days. After the first day, move the sample to a container of fresh PROTOS. Caution: Make sure that the container holding the sample and PROTOS is completely sealed and air-tight. Evaporation of water from PROTOS will cause the refractive index of the solution to change and will thus lower the effectiveness of optical clearing. 20. To image the cleared sample, it must be mounted between a glass slide and a black Willco dish. Roll up a piece of Blu-Tack adhesive into cylinder shapes of a thickness slightly more than the thickness of your sample. Place them horizontally on the glass slide. Press down the edge of Blu-Tack to close up the gap between the Blu-Tack adhesive and the glass slide (Shown in pictures).
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21. Carefully place the sample in between the Blu-Tack pieces and add about 20 μL of PROTOS to the sample.
22. With the lipped side facing up, firmly press a Willco dish down onto the adhesive until it just comes into contact with the sample. Using a pipette, add more PROTOS to the gaps between adhesive until the imaging chamber is filled.
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23. KWIK-SIL is an adhesive that cures rapidly. Carefully add it to the gaps between the Blu-Tack to build a wall and seal in the sample. Take care not to introduce any bubbles, and make sure the chamber is completely filled with PROTOS.
24. Cover this construction with aluminum foil and store it away safely to cure. After about 20 minutes, the sample is ready for imaging.
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Chung lab MAP protocol
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MAP protocol Please check www.chunglabresources.org for more information.
• Blu-Tack – Bostik, Blu-Tack Reusable Adhesive 75g • Slide glasses (1 mm-thick) • Cover slips #1 • 8 mm Round Cover Slip German Glass, #1.5 - EMS Cat # 72296-08 • Silicone isolators 8 well Press-to-Seal (0.5 mm x 9 mm) - Sigma S1810 • Parafilm M – Bermis PM996
A. Preparing the coverslips:
All steps in this subsection are performed under sterile conditions
1. Using forceps, gently place 8 mm circular glass cover slips in 24 or 48 well plates. If the slides are opened in the biosafety cabinet, there is no need to sterilize them in my experience.
• Add 250 μL 0.1% gelatin solution to each well (take aliquot from big bottle to avoid contamination).
• Incubate RT for 15 min. • Aspirate solution and let dry in the biosafety cabinet with the lid off for 15 min.
• This step ensures a gelatin layer on top of the glass to help cells adhere. It also allows for effective detaching during cell embedding (section B). This step might also work with poly-Lysine instead of gelatin, but I have not tried it.
2. Seed HeLa cells at desired density. For a 24 well plate 25,000–100,000 cells per well is a useful range. 100k will give you mostly confluent coverslips, which is not great for imaging tubulin, might be ok for FISH. I typically use 50,000 cells per well.
3. Let the cells bind overnight. I have left them for overnight (can leave longer, but keep in mind the cell density will increase, so plate accordingly.)
B. Embedding the coverslips:
From this point on sterility is not a concern for staining, might be important for FISH to minimize DNA degradation and bacterial/viral contamination.
4. Fix cells in 200 μL freshly prepared 3% PFA + 0.1% glutaraldehyde (GA) for 10 min at RT.
• 500 μL 10X PBS • 470 μL 32% PFA • 10 μL 50% GA • 4 mL ddH2O
[This fixation buffer is essential for good tubulin staining, but this protocol also works with 4% PFA fixation or 1% GA fixation (always buffered in PBS).]
4b. Finish the fixation protocol:
• Remove fixation buffer, • wash once with PBS, • reduce sample with 0.1% NaBH4 for 7 min, • quench sample with 100 mM glycine for 10 min, • wash twice with PBS, • and add PBS and keep at 4°C until samples are ready to be processed.
5. Prepare 5–10 mL Cell-MAP monomer solution (I use this solution for a week if stored at 4°C with aluminum foil to protect from light):
• 20% w/v acrylamide • 10% w/v sodium acrylate • 0.05% bis-acrylamide • 4% PFA • 0.67% TEMED (do this step under a fume hood!) • 10% 10X PBS
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6. Prepare 5% ammonium persulfate (APS) solution (0.05 g in 1 mL ddH2O). Do not add this to the Cell-MAP solution since it will cause it to polymerize quickly! I keep this solution at 4°C and use for up to a week.
7. Add 200 μL Cell-MAP monomer solution to each well. Incubate at RT overnight. [This step causes PFA-adducts to react with the amide groups in acrylamide]
8. When ready to embed your sample. Set up your “work area” for polymerization.
• Ethanol spray and clean your lab bench. • Add Press-and-seal silicone spacers to the glass slide • Place two folded-over Kim wipes on the bench. • Cut 2 squares of Parafilm-M (use their squares as a reference), peel off the wax paper and place
them on your work bench clean side (wax paper) up.
9. Use forceps to grab the cover slip containing the cells. Do not scratching the surface or break the glass. I usually use the forceps to tilt the coverslip upright and grab it by the edge.
• Pipette 90 μL of Cell-MAP solution onto Parafilm-M (will form a droplet). • Pipette 10 μL of 5% APS solution onto Parafilm-M (1 cm away from Cell-MAP solution). Do not
mix the APS with the Cell-MAP solution just yet! • Tap the cover slip on the Kim wipes and place it face up to remove liquid on the bottom of the
coverslip to remove excess liquid. • Set P200 pipette to 25 μL, use tip to drag APS droplet into the Cell-MAP solution droplet. Pipette
to mix 6–8 times. • Add 25 μL into the well of the silicone spacer, and use the pipette tip to spread the volume
evenly around the glass. Count to five Mississippi (!) and place the coverslip with cells face down (cells embedded in monomer solution). It is possible to do 2–3 samples at a time with some practice, maybe more. [The trick here is getting enough polymerization to increase the viscosity of the monomer solution before adding the coverslip face down. If done too early, the coverslip will sink to the bottom of the solution and monomer will polymerize above the cells)
• Wait 5 min for solution to polymerize (use leftover MAP + APS droplet to check consistency). • Use forceps to peel off polymerized MAP. • You can also do this entire embedding directly on the coverslip with the cells. After mixing the
APS and monomer solution, simply place 25 μL on the coverslip with the cells facing up. This works really well, but the curvature of the droplet can cause some annoyances during sample mounting and imaging. This approach can work if a razor is used to trim the polymer drop/disk into smaller sections.
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10. Place droplet in 15 mL tube with 10 mL PBST (PBS + 0.1% Triton X-100 + 0.02% sodium azide) to rehydrate for 30 min.
11. Decant PBST and use the metal spatula at the opening of the tube to prevent gel from escaping. Fill tube with 10 mL denaturation solution (200 mM SDS + 200 mM NaCl + 50 mM Tris, pH 9, store at 37°C or SDS will precipitate).
12. Denature gel for 30–60 min at 95°C.
13. Place sample in a large volume of ddH2O and let expand for 30 min. Take picture to record expansion.
14. Place sample in 10 mL PBST on shaker for 15 min. Repeat 3 times in total.
15. Stain with antibodies, etc. I typically stain with my primary and secondary for 2 h at 37°C. I preform 3 × 5 min washes after each antibody.
16. Before imaging, expand sample at least 4 h in ddH2O (I usually do this overnight). Longer incubation insures full sample expansion.
17. Mount polymer embedded cells as you would a thin tissue section using Blu-Tack (Bostik), glass slide and a Willco dish (Willco Wells, HBSB-5030). Mount in water and depending on the lens use water immersion whenever possible.
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MAP protocol Please check www.chunglabresources.org for more information.
Original Article Taeyun Ku, Justin Swaney, Jeong-Yoon Park, Alexander Albanese, Evan Murray, Jae Hun Cho, Young-Gyun Park, Vamsi Mangena, Jiapei Chen, and Kwanghun Chung. "Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues", Nature Biotechnology, 2016, doi:10.1038/nbt.3641.
This protocol is optimized for 1 mm-thick or thinner mouse brain slices.
• Fixation solution – 4% PFA in PBS • Washing solution – 0.02% NaN3 (w/v) in PBS • Low AA solution – 4% AA (w/v), 4% PFA in PBS • Inactivation solution – 1% acetamide (w/v), 1% glycine (w/v), 0.02% NaN (w/v) in DI water.
Titrate the pH to 9.0 with NaOH • MAP solution – 30% AA (w/v), 0.1% BA, 10% SA (w/v), 0.05% V-50 (w/v) in PBS
Make 10% V-50 stock solution, and aliquot in Eppendorf tubes and freeze them at -20°C before use.
Make 38% SA stock solution. SA can have oily phase separation and black particles. Once making the stock solution, centrifuge it at 1,000 g for 5 minutes. Use only the top fresh solution, and aliquot in 50 ml conical tubes and freeze them before use.
MAP solution can be prepared and kept at 4°C if light-protected. But V-50 initiator must be freshly added before use.
• Denaturation solution – 200 mM SDS, 200 mM NaCl, 50 mM Tris in DI water. Titrate the pH to 9.0 with NaOH.
Steps:
i. Perfusion ii. Sectioning
iii. Post-fixation iv. AA integration v. Inactivation
vi. Monomer incubation vii. Mounting
viii. Gel embedding ix. Denaturation, clearing & expansion
i. Perfusion
Perfusion is performed as usual.
1. Keep PBS and “Fixation solution” on ice. 2. Perfuse an anesthetized mouse transcardially with PBS.
Ensuring that blood is eliminated is important since remnant blood may cause remarkable nonspecific signal during antibody staining of MAP-processed tissue.
3. Switch to “Fixation solution”. Six minutes for each of perfusion step with 20 mL solution is recommended.
4. After perfusion, extract the brain and put it in a 50 mL conical tube with 30 mL “Fixation solution”.
5. Incubate at 4°C overnight with gentle shaking. All further incubations in solution are performed with gentle shaking.
6. Next morning, move the tube to 37°C and incubate for 3 h.
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ii. Sectioning
Depending on the final slice thickness to be used, the brain can be sliced in either way: it can be sectioned to the desired final thickness; or sliced to 1 mm-thick blocks and then sliced again into the final thickness after gel embedding. The latter is a better way to obtain multiple samples at once, but the bottom part can be lost due to the use of glue during the second sectioning.
1. Carefully discard the “Fixation solution” in the tube. 2. Wash with 30 mL “Washing solution”. You can either proceed to sectioning or store the sample
in the solution. 3. Use a vibratome to slice the brain. Fill the chamber with cold “Washing solution”.
According to the embedding step, the most practical thicknesses recommended are 170 μm and 1 mm. See the “Mounting” part.
iii. Post-fixation
Additional fixation allows the tissue to maintain its protein integrity for favorable staining.
1. Put slices to be processed into well-plate or conical tube with ice-cold “Fixation solution”. The amount of solution can be 1.5 mL for a 24 well-plate and 10 mL for a 15 mL conical tube.
2. Incubate at 4°C overnight and then at 37°C for 2 h. 3. Wash with “Washing solution” twice at 37°C for 3 h each.
iv. AA-integration
AA should be crosslinked to proteins to link the final poly-AA mesh with the tissue. This prior AA integration step helps control the tissue-gel interaction by limiting the amount of AA residues attached to the tissue that will participate in the gelling step later. Integration with 4% AA will give good tissue-gel properties both for mechanical rigidity and stainability (antibody penetration).
1. Incubate slices in “Low-AA solution” at 4°C overnight and then at 37°C for 3 h. The solution should be light-protected.
2. Wash with “Washing solution” three times at 37°C for 2–3 h each.
v. Inactivation
Inactivation quenches remaining methylol group formed by PFA and not reacted with AA. Also, this step together with AA-integration step completes the quenching to prevent further inter- and intra-protein crosslinking.
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1. Incubate slices in “Inactivation solution” at 37°C for 4–10 h depending on the slice thickness. Protect the samples from light.
2. Wash with “Washing solution” three times at 37°C for 3 h each.
vi. Monomer incubation
Note that PFA is not used in the “MAP solution”. The solution should be light-protected in the entire further steps.
1. Freshly add V-50 to make “MAP solution”. 2. Incubate slices in the “MAP solution” at 4°C. One day is enough for 170 μm-thick slices, and 1
mm-thick slices can be incubated for 2 d.
vii. Mounting
A gel formed with high (30%) AA is very rigid. The use of such high AA provides an expanded tissue the required mechanical stability. However, an excess gel layer formed outside the top and bottom surfaces of the tissue hinders lipid clearing and antibody delivery. Minimizing the excess gel layer is, therefore, important and a key consideration in the mounting step.
1. Prepare required materials for mounting: 1-mm-thick slide glasses, cover slips #1 (for tissue slides with 170 μm or related thicknesses), Blu-Tack, a fine paint brush
2. Shape a piece of Blu-Tack using gloved hands into a long and thin tube (rolling and pulling). The thickness can be roughly twice the tissue thickness. Put the Blu-Tack at the bottom region of a slide glass (Pic 1) and gently press with your gloved finger to make the Blu-Tack adhere to the glass. Use a razor blade to trim excess Blu-Tack (very important) to get the desired thickness. Add a small Blu-Tack ball of a similar thickness at the top region. Prepare multiple slides because this preparation will take the longest time. 3. Wet the center position of the Blu-Tack well on the slide glass using a tiny volume of “MAP solution” with a paint brush.
4. Place a tissue slice at the position. Avoid making bubbles. 5. Wet the top surface of the slice using a paint brush. This is to make a convex top and avoid bubbles when covering.
6. If you are mounting a 170 μm-thick slice, place three cover slips (Pic 2). If you are mounting a 1 mm-thick slice, place three glass slides.
7. Place a slide glass in parallel on top of the bottom slide glass.
Pic 1. Blu-Tack shaping on a slide glass
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8. Start pressing the slide glass gently using three fingers at the locations of inserted cover slips/slide glasses. Avoid pressing the tissue or bottom Blu-Tack region. This may squash the tissue.
9. Completely press the cover. The top ball region should also be pressed. This flattened ball will maintain the entire thickness consistent during embedding. Some resistance should be felt from all the three glasses inserted. You can check if pressing is completely done by the tissue trapped between two glasses and not moving when you stand it up.
10. Remove the inserted glasses. 11. Hold the sandwich block vertically, and slowly add some volume of
“MAP solution” into the space between the two slide glasses using a micropipette (Pic 3).
12. Once mounting is finished, immediately go ahead for “Gel embedding” step.
viii. Gel embedding
A nitrogen gas is required to lower the oxygen that disturbs polymerization. An Easy-Gel (www.lifecanvastech.com) device can be used for this embedding procedure.
1. Prepare a 45°C heating environment or set the temperature of Easy-Gel well to 45°C.
2. Insert the sandwich block into a 50 mL conical tube. 3. Purge the tube with a nitrogen gas for 10 s. 4. Cap the tube. Maintaining a positive pressure from a nitrogen gas tank
is preferred. 5. Embed for 2 h. Ensure that there is no leakage of nitrogen gas when
using Easy-Gel. 6. Take the sandwich block out. 7. Add a small volume of “Denaturation solution” on top of the formed
gel to hydrate the polymer. 8. Using a razor blade, carefully separate the two slide glasses. Be cautious not
to tear the tissue. 9. Cut out the excess gel around the tissue using a razor blade (Pic 5).
The excess gel does not need to be completely excised at current step if the tissue boundary region must not be lost. Softened tissue after expansion is easier to remove the gel. See next section.
10. You can also slice the embedded tissue at this step with a vibratome if you need thinner sections.
ix. Denaturation, clearing & expansion
Pic 2. Sandwiching a brain section with the aid of three spacers
Pic 3. Completion of mounting by filling the well with “MAP solution”
Pic 4. After embedding
Pic 5. Trimming an embedded slice using a razor blade
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1. Immerse the embedded tissue in a “Denaturation solution” in a conical tube. 10 mL or 20 mL can be used for 15 mL or 50 mL conical tube, respectively.
2. Incubate at 37°C. The incubation time can be 2 h for 170 μm-thick slices and overnight for 1 mm-thick slices.
3. For 1 mm-thick slices, incubate at 70°C for 5 h in a water bath or using EasyClear (LifeCanvas Technologies). Otherwise, skip to step #4.
4. Incubate at 95°C for either 45 min (170 μm) or 1 h (1 mm). Shaking is not required for this step. 5. Dispose the solution with being careful not to lose the tissue. 6. Transfer the tissue to a Petri dish or other flat container that is good for gentle horizontal
shaking. 7. Carefully wash the tissue with a small amount of DI water. 8. Fill the dish with DI water. Adjust the volume not to flood on a shaker. 9. Incubate at 37°C for 2–8 h depending on the slice thickness. 10. Exchange DI water and incubate more until the tissue fully expands (e.g., 4 h for 170 μm,
overnight for 1 mm). 11. If the excess gel restricts the expansion of tissue region, the gel can be removed carefully using
paint brushes and/or a razor blade. 12. Measure dimensions of the expanded tissue to estimate the expansion ratio. 13. For further handling such as staining or chopping into small pieces, immerse the tissue in a
desired solution like PBS + 0.1% Triton X-100 + 0.02% NaN3. 14. For imaging after staining, the sample can be expanded repeatedly in DI water either in a well-
plate or a Petri dish.
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SWITCH protocol Please check www.chunglabresources.org for more information.
ORIGINAL ARTICLES Evan Murray*, Jae Hun Cho*, Daniel Goodwin*, Taeyun Ku*, Justin Swaney*, Sung-Yon Kim, Heejin Choi, Jeong-Yoon Park, Austin Hubbert, Meg McCue, Young-Gyun Park, Sara Vassallo, Naveed Bakh, Matthew Frosch,, Van J. Wedeen, H. Sebastian Seung, and Kwanghun Chung. Simple, scalable proteomic imaging for high-dimensional profiling of intact systems, Cell, Dec 3:163(6): 1500-14. doi: 10.1016/j.cell.2015.11.025. PubMed PMID: 26638076.
RELEVANT ARTICLES Taeyun Ku*, Justin Swaney*, Jeong-Yoon Park*, Alexander Albanese, Evan Murray, Jae Hun Cho, Young-Gyun Park, Vamsi Mangena, Jiapei Chen, and Kwanghun Chung. Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues, Nature Biotechnology, 2016, doi:10.1038/nbt.3641. Sung-Yon Kim*, Jae Hun Cho*, Evan Murray, Naveed Bakh, Heejin Choi, Kimberly Ohn, Sara Vassallo, Luzdary Ruelas, Austin Hubbert, Meg McCue, Philipp Keller and Kwanghun Chung. Stochastic electrotransport selectively enhances the transport of highly electromobile molecules, PNAS, 2015 Nov 17: 112(46): E6274-83. doi: 10.1073/pnas.1510133112. Epub 2015 Nov 2. PubMed PMID: 26578787; PubMed Central PMCID: PMC4655572. Kwanghun Chung, Jenelle Wallace, Sung-Yon Kim, Sandhiya Kalyanasundaram, Aaron Andalman, Tom J. Davidson, Kelly A. Zalocusky, Joanna Mattis, Sally Pak, Viviana Gradinaru, Hannah Bernstein, Julie Mirzabekov, Charu Ramakrishnan, and Karl Deisseroth, Structural and molecular interrogation of intact biological systems, Nature, 2013, 497, 332-337 REAGENTS
REAGENT SETUP Perfusion solution Create a solution with a final concentration of 1X PBS, 4% paraformaldehyde (PFA), and 1% glutaraldehyde (GA). As 40 mL of this solution is necessary for each perfusion, a typical recipe is: 4mL 10X PBS, 5 mL 32% PFA, 0.8 mL 50% GA, and 30.2 mL water. This solution should be made fresh immediately prior to performing perfusion and kept on ice at all times. It is recommended to chill all of the separate ingredients before mixing the components. Fixation-OFF solution Titrate a bottle of PBS to pH 3 using HCl. Create solutions of 0.1 M HCl in water and 0.1 M potassium hydrogen phthalate (KHP) in water. Finally, mix these solutions in a ratio of 2:1:1 (pH 3 PBS):(0.1 M HCl):(0.1 M KHP). To this new solution, add a stock solution of GA to make a final concentration of 4% GA. Ensure that this solution stays cold at all times. It is recommended to chill the solution before adding GA. Fixation-ON solution Add a stock solution of GA to PBS (pH 7.4) to make a final concentration of 1% GA. Ensure that this solution stays cold at all times. It is recommended to chill the PBS before adding GA. PBST To PBS, Add Triton-X 100 (TX) to a final concentration of 0.1% (v/v). Also, add sodium azide to a final concentration of 0.02% (w/v). Practically, this is achieved by adding 1 mL of TX and 0.2 g of sodium azide to 1 L of PBS. Inactivation solution To PBS, add acetamide to a final concentration of 4% (w/v) and glycine to a final concentration of 4% (w/v). Thermal clearing solution To water, add sodium dodecyl sulfate (SDS) to a final concentration of 200 mM, sodium sulfite to a final concentration of 20 mM, sodium hydroxide to a final concentration of 10mM. Titrate the solution to pH9 using boric acid. This solution should be made fresh frequently, as the sulfites tend to degrade over time in solution. DiD-OFF solution
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To PBS, add SDS to a final concentration of 10 mM. Dissolve 1mg of DiD powder per 200 µL. This solution should be kept protected from light. Note: molecules similar to DiD can be used if other excitation/emission wavelengths are desired, so long as the molecule is sufficiently lipophilic. Antibody-OFF solution To PBS, add SDS to a final concentration of 0.5 mM. This is most easily accomplished by diluting a stock solution of SDS. When adding large proportions of antibody to this solution (say, >1:10), care should be taken to account for the resulting change in SDS concentration. Optical clearing solution This solution consists of 23.5% (w/v) n-methyl-d-glucamine, 29.4% (w/v) diatrizoic acid, and 32.4% (w/v) iodixanol in water. Use a stir bar (or shake if necessary) to fully dissolve the powders at each step. Do not use heat when mixing the solution, as this will cause a color change. This solution should be stored carefully to ensure that no water is lost, as just a small amount of evaporation will result in precipitation. Teflon tape can be used to increase the security of the bottle’s seal, and parafilm can be used around the cap. It may be necessary to use a 60% iodixanol solution (see reagents list) rather than iodixanol powder, as it is not cheaply available. Therefore, an example recipe for the optical clearing solution is as follows: Dilute 60% iodixanol solution to 47% iodixanol. To achieve this, roughly 2.75 mL water should be added for every 10 mL of 60% iodixanol solution. Then, for every 10 mL of the resulting solution, 3.39 g n-methyl-d-glucamine and 4.24 g diatrizoic acid should be added in order. Be sure to take into account that the final volume will be significantly larger than the starting volume. In addition to our optical clearing solution (termed PROTOS), there are proprietary optical clearing solutions available as well as other published recipes that other groups have developed for their tissue clearing protocols. The proprietary formulations are prohibitively expensive (RapiClear and FocusClear) and at least FocusClear is known to result in the formation of precipitate within samples during long-term storage. PROTOS is the most cost-effective option for high quality optical clearing, which is an absolute must for thick tissue imaging. CUBIC-mount (Lee, 2016) and sRIMS (Yang, 2014) are cheaper than PROTOS, but they are noticeably less effective. Solution Recipe Cost per 500
mL PROTOS 29.4% diatrizoic acid $565.80 23.5% n-methyl-d-
Premade PROTOS is available from LifeCanvas technologies (EasyIndex, http://www.lifecanvastech.com/) PROCEDURES All samples must be preserved through use of either procedure 1a or 1b below and then inactivated through procedure 2 and cleared through procedure 3 in order. Procedures 4a and 4b are optional, but it is not recommended to perform both in the same round of staining. Samples thicker than 50-100 µm must undergo procedure 5 in order to be imaged fully, but it is optional for very thin samples. After procedure 6, you may go back to procedure 4a or 4b to complete another round of staining. Processing times at each step will vary depending upon the tissue type and size of the sample. Unless otherwise noted, the parameters given below were optimized for adult mouse brain samples. 1a. Perfusion If it is possible, perfusion is the preferred method of tissue preservation. Using the perfusion technique of your choice, first perfuse 20 mL of ice-cold PBS through the beating heart of an anesthetized mouse, followed by 20 mL of the ice-cold perfusion solution described above. Take care not to introduce any bubbles during the procedure, and use a flow rate slow enough to avoid damage to the vasculature or brain sample (<5 mL/min). After both solutions have been perfused, carefully remove the brain from the skull using any technique you are comfortable with. The dura membrane should also be removed during the process. Place the sample into 20 mL of perfusion solution and incubate at 4 ˚C with gentle shaking for 3 days. 1b. SWITCH-mediated tissue preservation If perfusion is not possible, the sample must be preserved using SWITCH. The sample should be first fixed with PFA for several days before proceeding. Incubate the sample in 40 mL fixation-OFF solution at 4 ˚C with gentle shaking for 2 days. The sample should then be moved to fixation-ON solution at 4 ˚C with gentle shaking for an additional 2 days. NOTE: the timing for the Fixation OFF and ON steps is dependent on the sample size and may need to be optimized from these starting values on a case-by-case basis. We found that these parameters worked well for banked human samples of roughly 0.5-1.0 cm thickness. 2. Fixative inactivation
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After fixation via either perfusion or SWITCH, the sample must be washed in PBST to remove unbound fixative molecules. For mouse brains, 2 washes of 6 hrs each at RT with gentle shaking was sufficient. To inactivate remaining fixative molecules, the sample must then be washed in inactivation solution at 37 ˚C O/N using a water bath or EasyClear. If the solution turns yellow, the inactivation solution should be replaced with fresh solution and the sample incubated for several more hours. Note: if the sample needs to be cut, this should take place now before the sample is cleared. 3. Thermally-assisted lipid clearing Inactivated samples must next be incubated in thermal clearing solution to wash away remaining inactivation solution and to distribute sodium sulfite through the sample. After 2 washes of 6 hrs each at 37 ˚C, the samples should be placed in a tube of fresh thermal clearing solution, which should then be placed in a water bath heated to 70 ˚C or in EasyClear. Other temperatures or methods of consistent heating may be used, but samples may deteriorate over time at higher temperatures. If a sample contains fluorophores that were genetically-encoded, introduced through viral injection, etc., then the sample may be cleared at 37 ˚C to preserve this fluorescence. The clearing process will take longer at this low temperature, but temperatures higher than this will result in loss of fluorescence during clearing. 4a. SWITCH-mediated myelinated fiber labeling After a sample has been clearing, SWITCH-mediated labeling is possible. Myelinated fibers can be readily visualized with the lipophilic DiD fluorescent molecule. The sample should be equilibrated in a solution of 10 mM SDS in PBS in order to distribute SDS molecules throughout the sample. The sample should then be placed in a volume of DiD-OFF solution just large enough to cover the sample and incubated at 37 ˚C with gentle shaking for 12 hrs to 7 days depending on the size of the sample (1 mm-thick section to whole mouse brain). The sample should then be moved to 40 mL of PBST and incubated at 37 ˚C for 12 hrs to 2 days. We have also observed that tomato lectin and nuclear stains such as DAPI or Syto16 can be used with this SWITCH approach. 4b. SWITCH-mediated immunolabeling After lipid clearing of a sample, SWITCH-mediated immunolabeling is possible. The sample should be equilibrated in antibody-OFF solution in order to distribute SDS molecules throughout the sample. The sample should then be placed in a fresh volume of antibody-OFF solution just large enough to cover the sample, and then antibodies should be added in the desired proportions. The sample should then be incubated at 37 ˚C with gentle shaking for 12 hrs to 7 days depending on the size of the sample (1 mm-thick section to whole mouse brain). The sample should then be moved to 40 mL of PBST (antibody-ON solution) and incubated at 37 ˚C for 12 hrs to 2 days to initiate antibody binding and wash out unbound antibodies. If secondary antibody labeling is required after primary staining, incubate the sample in a fresh volume of PBST with secondary antibodies. It is important to use enough secondary antibodies to saturate all primary antibodies within the sample. 5. Optical clearing After labeling, the sample must be equilibrated in PROTOS, refractive index-matching solution, or in EasyIndex to achieve maximal optical clearing. Incubate the stained and fully washed sample in PROTOS with proper shaking at 37°C using EasyClear or shaking water bath. Use 500 µl, 2 ml, 25 ml, 50ml of PROTOS for clearing 100 µm, 1 mm, intact mouse hemisphere, intact mouse brain, respectively.
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The sample may incubate in PROTOS or EasyIndex longer as needed depending on the thickness. 100 µm slice can be cleared within 10 min, whereas intact mouse brains require overnight incubation. After optical clearing, the sample should be clear enough to easily see through by eye. If the solution immediately surrounding the sample seems inhomogeneous, it suggests that the sample has not yet fully equilibrated with the solution. The sample must be further incubated in a fresh PROTOS solution with proper shaking until it reaches complete equilibrium. 6. Sample Mounting 1. Form a blue tack “worm” only slightly thicker(~0.1mm) than the samples. Lay this “worm” in a circle around the center of a slide glass (Fig. 1A). 2. Use a pipet tip to seal the blue tack and slide glass surface by pressing down the blue tack along the outside edge of the circle (Fig. 1B). CRITICAL S TEP Sealing blue tack can prevent PROTOS from slipping between the blue tack and slide glass, and also from the evaporation and drying during storage. 3. Place slices on the center of the slide glass inside this circle (Fig. 1C). Ensure that the slices are wet with PROTOS. The entire circle does not need to be filled with protos, unless samples are to be stored for an extended period of time CRITICAL S TEP if long term storage of the sample is intended, see the end of this procedure for details. 4. Place a clean “covering dish”(wilco dish) on top of the blue tack circle, pressing down until it comes into contact with the surface of the samples (Fig. 1D). 5. Place the slide under the microscope’s water objective and fill the “covering dish” with enough water so that a column of water forms between the objective and dish upon lowering the objective (Fig. 1E). CRITICAL STEP Ensure no bubbles are formed between the objective and dish, because they may distort images. Note: This set up works for microscopes with low NA→ these microscopes allow changes in medium between the objective and final sample. In this case, a 10X water objective on a confocal microscope was used to image 1mm slices. Note: For extended sample preservation- Leave the circle of blue tack disconnected and place the samples at the center of this circle. Seal the blue tack to the slide glass surface, except for a small opening as described above. Press on the “covering dish” until it makes contact with the samples. Use a pipet to fill the space between the slide glass and “covering dish” with PROTOS through the small opening. Use glue to seal this opening. These samples can now be stored for an extended time.
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Fig. 1 Sample Mounting Diagram. A) Place a blue tack ring slightly thicker than the sample on a glass slide. B) Seal the blue tack to the slide using a pipet tip. C) Place the samples in the center, keeping them wet with PROTOS. D) Press on a covering dish until it comes in contact with the sample surface. E) Place the slide under the water objective, filling the covering dish with water until it forms a column of water with the lowered objective.
7. Molecular probe elution After imaging, the optical clearing solution should be washed out of the sample with thermal clearing solution. After the sample has equilibrated, the sample should be placed in a 70 ˚C water bath or EasyClear for 2 hrs to O/N depending on the size of the sample. Labeling can proceed again after this step is completed
A B
C D
E
The following software can be useful for analyzing large 3D data files. This list is by no means inclusive but provides a user friendly, open source/cross platform starting point for data analysis. Fiji (ImageJ) Fiji is primarily designed for 2D image analysis but incorporates many tools and plugins that can still be useful for 3D analysis. Fiji is able to handle large data files provided the system has adequate RAM. Although many people will be familiar with the use of FIJI the following will highlight some features that are useful for 3D processing. Stitching There are a number of stitching plugins in FIJI. I find the Stitch sequence of grids of images plugin very useful to stitch together a sequence of files and works well with Lavision lightsheet files. Plugins à Stitching à deprecated à Stitch sequence of grids of images Input the number of X and Y tiles as well as the number of Z steps and overlap between tiles. Point the plugin to the folder with the image files Copy and paste the image file name and alter to indicate XYZ file structure For example, File Name 10-09-34_TF 20_3L LS_Ultra[00 x 00]_C00_xyz-Table Z0000_Ultra Filter0000.ome.tif Change to 10-09-34_TF 20_3L LS_Ultra[{xx} x {yy}]_C00_xyz-Table Z{zzzz}_Ultra Filter0000.ome.tif Set an output directory Start XYZ from 0 Uncheck compute overlap if using a file format with embedded coordinates (e.g. OME Tif). If the image positioms are unknown this can be computed by the plugin. Linear blending usually works well. This will automatically perform the XY stitching for each Z plane.
3D Visualization To view a data file in 3D there are a few options. 2D projection. The simplest way to view a 3D file is to generate a projection of the 3D image to view the file in 2D space Image à Stacks à Z project Choose max intensity (standard deviation also works well for some images) This will generate a 2D image for display purposes. Depth coded projection. A modification of this is to produce a depth coded projection image that displays different z planes as different colors in a 2D image. With an image stack open, go to Image à hyperstacks à stacks to hyperstacks. Then once the stack is converted go to Image à hyperstack à Temporal color code. Chooses spectrum as the LUT
3D Projection Fiji also has a built in 3D projection module to visualize an image volume. TO use this feature open a stack or 3D file. Then go to Image à stack à 3D project.
Start with brightest point as the projection method. Input the Z spacing. The rotation angle increment can be left at 10 but for a smoother rotation set to 1 (slower rendering). Check the interpolate box. This can be fairly slow depending on the file size. The 3D projection can be saved as an image stack or compiled as an avi video file. 3D viewer. There is also a 3d viewer plugin incorporated into FIJI. Go to Plugins à 3D viewer. Add the image and display as Volume. The resulting volume can be recorded ad an animation. The volume can also be displayed as a surface rendering or multi-orthoslice Orthogonal Views. The image stack can be viewed simultansously in XY, YZ and XZ planes. Go to Image à stack à orthogonal views. Moving the yellow line changes the plane in all 3 windows. This can be useful for colocalization.
Ilastik One of the problems with generating large data sets from cleared tissue is that the quantification can easily become overwhelming to handle. For example, cell counting which can be handled fairly easily in individual regions or subsampled tissue is now a huge task. To overcome this issue automated quantification is required. Ilastik allows users to perform simplified machine learning to identify objects of interest and batch process this across large tissue volumes. Pixel classification + object classification To begin open Ilastik and create a new project using Pixel classification + object classification Input data - Import an image file (i.e., a single z plane from your stack) Feature selection - First, select features that are suitable for your image. Training – you will need to create at least 2 label classes (more for multichannel images) that differentiate signal from background. Once the 2 labels are created begin to draw on the image in the areas that correspond to your signal and areas that correspond to background. The more input you give the program the better it will learn the rules so make sure to sample various areas of the image. Click on live update to see real time progress of the classifier. Thresholding - Set the input to the channel that corresponds to your signal. From here there will be a bit of trial and error. This panel can be used as a first pass to clean up our cell detection but don’t worry about getting the classification perfect here. We will still do an object based classification later. Better to over select rather than under select objects at this point. The intensity based threshold value can be changed between 0-1. Higher values will be more restrictive and lower values more permissive. Setting the value too low will likely result in multiple objects being lumped together. Too high of a setting will result in too little signal being detected. The size filter can be used to eliminate objects that are too small or too large to be considered objects of interest. Object feature selection - So far we have told the software what constitutes signal versus background. Now we will further classify what part of the signal constitutes an object of interest. Select the feature sets based on size, shape intensity etc that are useful for describing your objects of interest. Object classification – you will need to define at least 2 label classes again. This time they should correspond to objects of interest versus irrelevant signal. Click on live update to see the classification in real time. Then begin to click on objects that you consider real and on objects
that you consider to be irrelevant. Again, the more training you provide the better the classification will be. Correct items that the classifier gets wrong by clicking on them with the opposite label. You should end up with an accurate classification of the cells of interest. This training data that you have provided can then be used to batch process the rest of your image stack. Cell density counting An alternative method in ilastik is to use the cell density counting approach. This is particularly useful for high density images with cells that are closely clustered. To begin open Ilastik and create a new project using cell density counting Input data - Import an image file (i.e., a single z plane from your stack) Feature selection - First, select features that are suitable for your image. Counting - Here you will find 2 classification labels. Foreground and background. Background will be painted on using a pen tool as was done in the pixel classification method. The Foreground will be selected with a point selection tool. To use this tool click in the middle of your cells. Set the sigma value so that the marker almost fills the cells but does not spill beyond the cell boundaries. Add a box to see the density in a given area or click on update total density to see the density score for the entire image. This technique can also be used to batch process an entire image stack. Keep in mind that this method usually needs more training input than the pixel/object method so make sure to select many cell and multiple regions of the background.
Vaa3D Vaa3D is similar to Fiji in the sense that it is based on plugins which can be used to expand the functionality of the software. However, Vaa3D as the name suggests is more suitable for 3D image processing. There are many plugins for a variety of purposes. A couple of these functions are discussed below to help you begin using the software. To use, first open a 3D file in Vaa3D. You will initially be greeted with a 2D orthoslice or your image. On the bottom right of the screen you can click on see in 3D to view your image volume. Once in the 3D window you have a control panel on the right side of the screen. Here you can change the projection method and threshold your image if desired. The LUT can be changed by using the volume color map tab. The volume cut controls allow you to isolate certain parts of the image volume. At the botoom right there are controls to zoom, shift and rotate your volume. This can also be done using the mouse. There are a number of option available using right mouse clicks on the image. Markers can be defined by a single right click to define a cell or dendrite or by 2 sequential right clicks (performed in different orientations) for better accuracy. In the right click menu you can also zoom in to a region by drawing a box around the ROI. A new 3D window will be generated. There are a number of ways to trace neurons in Vaa3D. PluginsàNeuron_tracing but you can also trace neurons directly from the right click menu. Place a marker first on your cell body and then on the ends of each dendrite. Then Right click to select trace from marker 1 to all other marker positions. To make an animated movie of your volume Go to plug-in à movies and snapshots à Zmovie maker à generate…
use the volume cuts, zoom, shift and rotation controls to move your image around. Click Add an anchor point after each move. When done click on preview. If you’re happy with the preview save the file.
Neutube Neutube is an open source platform for neuron tracing. I have found that it performs very well and has a simple interface. The automatic tracing function is also available within Vaa3D as a plugin. However, I prefer to use the manual tracing option in most cases. Begin by opening an image stack. Tracing can be performed on the 3D image or the 2D stack. I find it easier to use the 2D stack. Simply click on branches of the neuron and you will see a skeleton path traced even in areas that are out of the current z-plane. Continue to click on branches as you scroll through the stack. The final tracing can be saved as a .swc file for further analysis in Vaa3D or numerous other platforms.
