Indiana University adapted Akoya Phenocycler-Fusion Tissue Staining and Imaging Protocol for Fresh Frozen Kidney Samples

Prepare humidity chamber - we use an empty 1000uL tip box and place water and a wet paper towel under the tray.

Prepare Drierite absorbent beads - locate a second empty box with a lid and add an even layer of beads

Get an ice bucket - for use later when preparing Antibody Cocktail

Locate and/or label 5 plastic coplin jars for the following reagents:

1 x Acetone

2 x Hydration Buffer 

1 x Pre-stain fixative

1 x Staining Buffer

Prefill the acetone, hydration buffer, and staining buffer jars

Remove sample slides from the -80 freezer and immediately place on Drierite beads for 5 minutes

- make sure slide is tissue-side up

Remove slide from Drierite beads and place in Acetone coplin jar for 10 minutes

Place slide in the humidity chamber to dry for 2 minutes

Place slide in the first Hydration Buffer coplin jar and incubate for 2 minutes

- Dip slides 2-3 times to make sure acetone is rinsed off adequately 

Place slide in the second Hydration Buffer coplin jar and incubate for 2 minutes

During this incubation, prepare the Pre-staining Fixative solution

• For one coplin jar, use 36 mL of hydration buffer and 4 mL of fresh 16% PFA

Move slide into the Pre-staining fixative solution coplin jar and incubate for 10 minutes

Move slide back into the first Hydration Buffer coplin jar and dip slide 2-3 times to remove the fixative solution

- this is not an incubation step, just a quick rinse 

Move into the second Hydration Buffer coplin jar and dip slide 2-3 times to remove the fixative solution

- this is not an incubation step, just a quick rinse 

Place slide in Staining Buffer and incubate for 20-30 minutes

- timing depends on how quickly antibody cocktail can be made

During the Staining Buffer incubation, prepare the Antibody Cocktail

Remove the selected antibodies, spin down if necessary, and place on ice

Prepare appropriate volume of Blocking Buffer for the number of slides being stained

*See attached file for Blocking Buffer components

MasterMixTable.xlsx

Label one tube for each unique Antibody Cocktail being prepared

The FINAL volume for each tube should be 200 uL - counting antibodies. Remove 1 uL of Blocking Buffer per 1 uL of antibody being added.

- Amounts will vary depending on experiment and desired antibody concentration

Add the desired amount of antibodies to the appropriate tubes

- Our concentrations are listed in the attached table. Antibody clones and vendor available upon request 

MasterMixTable.xlsx

Pipette gently to mix the solution

Optional step: Cut a rectangular piece of parafilm that is approximately the same size as the sample slide

Pre-load a pipette with 190 uL of the prepared Antibody Cocktail

Remove slide from Staining Buffer and use a kimwipe to gently dry the slide, without touching the tissue sample(s)

Place slide on the humidity chamber, tissue side up, and dispense the Antibody Cocktail

- make sure the tissue sample is completely covered, and there are no air bubbles present 

Optional step: Gently place parafilm over tissue sample

Incubate Sample Slide overnight at 4C

- We place our samples in a cold room to ensure there is no disturbance 

Locate and/or label the following plastic coplin jars:

2 x Staining Buffer

1 x Post-Stain Fixative

1 x 100% Methanol

4 x PBS (1x concentration)

1 x CODEX Buffer + Buffer Additive (1x concentration)

Fill Methanol coplin jar and place in freezer until needed

If parafilm steps were included, gently remove parafilm from sample slide

Place sample slide in Staining Buffer coplin jar and incubate for 2 minutes

- Dip slide 2-3 times to ensure Antibody Cocktail is fully washed away

Place sample slide in the second Staining Buffer coplin jar and incubate for 2 minutes

During this incubation, prepare the Post-Staining fixative solution

- For one coplin jar, use 36 mL of storage buffer and 4 mL of fresh 16% PFA

Move slide into the Post-Stain coplin jar and incubate for 10 minutes

Remove slide from the Post-Stain fix and wash in the first PBS coplin jar

- this is not an incubation step, just a few quick dips to rinse the tissue

Repeat step 33 in the second and third PBS coplin jars

With slide still in the third PBS coplin jar, remove Methanol coplin jar from the freezer

Place slide in the ice cold Methanol for 5 minutes

After Methanol incubation, immediately place slide in the first PBS coplin jar to rinse

- As in step 33, this is not an incubation, just a quick rinse 

Repeat step 37 with the second and third PBS coplin jars

With the slide still in the third PBS coplin jar, prepare the final fixative solution

- for up to 5 slides, use 1000 uL of 1xPBS and 20 uL of Fixative Reagent  

Preload a pipette with 200 uL of final fixative solution

Remove sample slide from the third PBS coplin jar and place in the humidity chamber, tissue side up

Dispense 200 uL of Final Fixative solution and incubate for 20 minutes

After the 20 minute incubation, rinse the slides in each of the three PBS coplin jars, as was done previously.

There are two options for how to proceed next:

a) if imaging is not taking place immediately, sample slide(s) can be stored in Storage Buffer at 4C for up to 5 days.

b) if imaging is taking place the same day staining is finished, proceed to step 45

Place slide to be cover-slipped in a fresh jar of 1x PBS and incubate for 10 minutes

After the 10 minute incubation, carefully dry the slide with a kimwipe, so the coverslip can adhere properly

Place coverslip - sticky side up - on the stage. Make sure the coverslip is straight, as overhang will create an inadequate seal or cause the slide to not fit in the flow cell

Gently place the slide - tissue side facing the sticky side of the coverslip - on the stage, making sure the slide is lined up with the coverslip

Push the stage under the pressure arm, lower the arm, and let rest with for 30 seconds

Place the now cover-slipped slide into 1x CODEX Buffer for 10 minutes to allow for equilibration and proper adherence of coverslip to take place.

Prepare the reporter plate stock solution as described in the attached spreadsheet.

There will be one unique well per cycle, including the blanks. Each well gets a different volume of stock solution based on the amount of reporter to be added. Final volume of each well should total 250 uL.

In general, the volumes are as follows, per number of reporters being added

3 reporters -> 235 uL of stock solution

2reporters -> 240 uL of stock solution

1 reporter -> 245 uL of stock solution

0 reporters -> 250 uL of stock solution (typically the two blank cycles)

The stock solution for the first and last cycles go into wells H1 and H2, respectively.

Stock solution + reporters will start in A1 and continue in order until all cycles are accounted for.

*An example of a reporter plate set up is shown in the attached spreadsheet

MasterMixTable.xlsx

Once all the necessary wells are filled, cover wells with foil plate seal to protect from debris and prevent the wells from drying out

*we have prepared reporter plates up to a week in advance, but generally try to prepare them no sooner than the day before an imaging session

When reporter plates and slides are ready to use, follow the instructions on the Phenocycler-Fusion to begin the imaging experiment. An example of our experimental settings (exposures and cycle set up) is shown in the attached spreadsheet.

Imaging Parameters.xlsx

Analysis conducted by IU follows the pipeline(s) described in the following papers:

• Winfree et al 2017

• Ferkowicz et al 2020

• Winfree et al 2023

(links are listed in references tab)

Segmentation and cytometry analysis for each sample is conducted using Volumetric Tissue Exploration and Analysis (VTEA, https://vtea.wiki/ ).

A pipeline for combining individual sample datasets into one analytical space using R Studio will be available soon.