BABB 2.0 Protocol

Materials and Reagents

  • PBS 0.02% sodium azide

  • 4% PFA

  • Blocking or Staining buffer - PBS + 0.5%NP40 +10% DMSO + 5% serum* + 0.5% Triton X-100

  • Wash Buffer (WB) - PBS + 0.5%NP40 + 10% DMSO

  • Methanol - 25%, 50%, 75%, 100%

  • DCM - dichloromethane

  • BABB - 1:2 mixture (benzyl alcohol: benzyl benzoate)

Preparation notes

  • Place 45mL of BABB in 50mL conical and add 5g of activated aluminum oxide (to remove peroxide contaminants), rotate for at least 1 hour at room temperature. Then centrifuge the tube at 3000rpm for 10 mins to pellet the aluminum oxide.

Procedure

Day 1

  • Collect tissue samples and place in 4% PFA for fixation at 4˚C for less than 24 hours.

Day 2

  • Remove 4% PFA by washing tissues with PBS 0.02% sodium azide at least 3 x 2 hours for 6 hours in total.

  • Let the samples overnight in PBS 0.02% sodium azide at 4˚C.

  • Samples can be stored in this solution for future processing.

Day 3

  • Cut the tissues ~2mm or ~1mm using the “matrix”.

  • Place each tissue in its own tube (1.5mL) and add 1.0mL.

Pre-treatment

  • Incubate samples in 25% QUADROL at 37˚C with gentle shaking.

  • Refresh the solution until supernatant is not green if the sample is non-perfused.

  • Perfused samples: Incubate in 25% QUADROL at 37˚C with gentle shaking overnight.

Day 4 or 5

  • Wash out QUADROL with two washes with PBS for 30 mins.

  • Incubate tissues in Blocking Buffer and rotate at room temperature for 1 day.

Day 5 or 6

  • Remove blocking buffer and add primary antibody in Staining Buffer.

  • Rotate for 72 hours (about 3 days) at room temperature.

Day 8 or 9

  • Remove primary antibody and wash samples with Wash Buffer.

  • Change WB every 2 hours at least for 6 hours.

  • Leave samples overnight in 4% PFA for primary post fixation.

Day 9 or 10

  • Wash samples twice for 5 minutes with PBS to remove 4% PFA.

  • Add secondary antibody in staining buffer and rotate for 72 hours (about 3 days) at room temperature.

Day 12 or 13

  • Remove secondary antibodies by washing samples with Wash Buffer.

  • Change WB every 2 hours at least for 6 hours.

  • Leave tissues rotating at room temperature in WB for the next day.

Day 12

  • Refresh Wash Buffer.

Day 13

  • Place samples in nuclear dye overnight.

  • Incubation time depends on the dye used in the experiment.

  • Example: SYTOX Green 488 (1uM) should stay for 48 hours (about 2 days) in PBS buffer.

Day 14

  • Wash samples twice for 15 minutes with PBS to remove the nuclear dye (If the nuclear dye is SYTOX Green 488).

  • Dehydrate lungs in methanol gradient (25%,50% ,75%) at least 1 hour in each %.

  • Finally placed sample in 100% methanol for 45 minutes.

  • Then, change out methanol and rotate for another 45 minutes.

  • Meanwhile, prepare fresh BABB.

  • After dehydration, incubate samples in DCM for delipidation.

  • 1st wash for 30-45 minutes, refresh DCM (after 45 minutes, verify if the tissues sink into the bottom of the tube), then proceed to clearing step.

Clearing

  • Add fresh BABB, mix, and remove BABB 3 times.

  • Then, remove BABB and add fresh BABB, rotate 15 minutes at room temperature.

  • Finally, remove BABB and add fresh BABB, rotate overnight at room temperature, protected with foil.

Day 15

  • Samples ready for imaging.

  • Longer incubation in BABB (at least 24 hours) improved imaging resolution.

Optional: Agarose Embedding

  • Prepare fresh 1% agarose low gelling temperature (more clarity and lower gelling temperature 24-28°C) Sigma A9414.

  • Let it cold for few minutes before adding to the tissue.

  • Place sample in bottom of the metal holder mold and fill until completely cover the tissue.

  • Let the sample solidify for at least 10-15 minutes before removing the agarose block from the mold.

  • Transfer sample to an 50mL or 15mL conical tube.

  • Proceed to dehydrate and match refractive index as describe above.

  • Make sure the block is completely immersed in the solution.

  • Shake the samples while dehydrating and clearing.

Notes and Cautions

  • We usually work with ~2mm or ~1mm tissue slices chopped with TED PELLA tissue slicer (0.5mm).

  • Cover samples with aluminum foil.

  • Protect tubes with aluminum foil.

  • SYTOX should not be prepared in phosphate buffers, but we used it and it works.

  • Important: prevent bubbles while pipetting agarose.