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.

Related Protocols
-----------------

- :doc:`labeling-tissue-samples`
- :doc:`agarose-cube-cleared-samples`