While wearing proper PPE, set up surgical area in fume hood

Once a suitable period post-surgery has occurred (3+ weeks) the animal can be perfused

Set up surgical area

Load ketamine-xylazine cocktail into injection syringe (0.2mL/mouse)

In physiological fume hood (separate from PFA-exposed hood) load isoflurane into holding chamber

Place mouse in isoflurane holding chamber until mouse breathing has slowed and running has stopped

Inject 0.2mL ketamine/xylazine IP and place animal back into original cage

Monitor breathing and foot-pinch twitch reflex

Once mouse reflex has stopped (should still be breathing) quickly move mouse to the surgical area

Restrain mouse using PrecisionGlide needles to hold down limbs

Cut mouse skin from to reveal fascia layer, do not yet make incision to reveal pleural space

Using fine surgical scissors, cut fascia to reveal organ space

Cut diaphragm and up sides of rib cage to reveal heart

Using resistance clippers, cut right atrium of the heart

Grip heart and insert cold PBS flat needle into left ventricle from the apex of the heart, begin to perfuse 10mL of PBS by hand at ~2mL/min

Begin flow of PFA at 120mL/hr, switch PBS needle for PFA needle using the same hole formed from the initial insertion

Set timer for 10min, when expires reduce flow to 100mL/hr for 15min

Set timer for 15 min, when expires reduce flow to 90mL/hr until 50mL is reached, 25min

Remove needle, turn mouse over (should be rigid due to PFA perfusion) and use thick scissors to sever head

Using fine surgical scissors, remove scalp to reveal skull

Use resistance clippers to gently cut skull without damaging brain, make incisions on skull at most rostral section of brain to enable skull to be peeled from brain

Use spatula to delicately remove brain and place into glass vial labelled with mouse ID, 4% PFA, initials and date

Refrigerate brain at 4°C for 24 hours

Set up work-station by gathering 6-well plate with PBS, a petri dish, and 6/12 glass vials (depending on if collecting both hemispheres)

Fill all of these with PBS, label the glass vials with the mouse ID, hemisphere genotype, initials, and date

Remove vibratome blade from manufacturer packaging and wash with ethanol followed by distilled water to remove protective oils

Use hex-tool to tighten blade to the blade-holder so that it is straight and extends several millimeters beyond the black blade-guards

Do not attach blade to vibratome at this time

Plug in vibratome and turn on, check settings: Slice thickness is 70um, slice frequency is set at 9, slicing speed should be between “4” and “5”

Retrieve brain from 4°C Fridge, wash in PBS x 3 and dispose of waste in Liquid PFA waste disposal unit

Place brain into holding chamber and using a brush, gently orient the brain to be equally distributed along the sagittal axis

For sagittal slicing, take one Personna 0.012 HD Heavy Duty Single Edge Razor blade and insert into the sagittal guide slits, push down through brain while maintaining even force between sides of the blade, this will cut the brain into two halves

Take one hemisphere and place into a petri dish filled with PBS, put the other half back into glass vial of PBS and store at 4°C until ready to slice

Equip the scalpel with its blade and prepare to use

Using spatula, pick up the hemisphere by the bisected plane on the flat edge of the spatula

Using filter paper, dry the bisected plane, absorbing residual PBS

Paint a thin line of super glue onto the vibratome stage, then quickly use the flat side of the scalpel blade to push the onto the stage so that the bisected plane comes into full contact with the glue

Place stage in stage holder and fill with PBS

Attach vibratome blade to vibratome

Raise the stage so that the blade is several millimeters above the slice

Using V-Max, set front and back of continuous cutting using the limit-set button, should be set a few millimeters in front of and behind the most forward and back parts of the brain

Begin slicing, placing the first brain slice in well 1, and the following in well 2, going in 6 slice groups (slice 7 will be in well 1) to generate a 1/6 series of the brain per vial

Once complete, take slices from ONE WELL that should contain your ROI and observe for viral expression using the epifluorescent microscope (if applicable)

Return slices to their original well

Transfer slices

Place each series in a glass vial and rinse sections with PBS 3 times

Add 1 mL of PBS-T with 2% Normal Donkey Serum (20 µL/mL) to each series and swirl briefly

Optional blocking step: leave slices in PBS-T and 2% Normal Donkey Serum at room temperature for 45–60 min

Add primary antibody to each series (see table 2)

Shake gently for 48–72 h at 4 ◦C (sections should barely revolve around the vial)

Rinse sections with PBS 3 times before starting secondary reactions

Create necessary “Secondary Antibody Cocktail” consisting of 1 mL of PBS-T with 2% Normal Donkey Serum (20 µL/mL) and corresponding secondary antibody (refer to suggested antibody concentration). Volume of cocktail should be +1 to all reagents to avoid lack of volume due to pipette error (meaning 12 vials = 13mL PBS-T, 20uL NDS x 13, 4uL 2° x 13)

Protect from light for all remaining steps.

Shake gently for 90 min at room temperature (sections should barely revolve around the vial)

Rinse sections with PBS 3 times before mounting

Using a glass petri dish, gently remove brain slices from 1 vial at a time and mount onto “name of glass slide”

Mount sections serially on slides with Prolong Diamond Anti-fade mounting media; protect slides from light and keep at 4 ◦C after 24 h drying at room temperature

Proper slide labelling format is

mouse ID – hemisphere

series

primary antibody

secondary antibody

genotype

initials – date

Insert new slide data and into slide census spreadsheet