Oct 22, 2024
Yale University adapted Akoya Phenocycler-Fusion Tissue Staining and Imaging Protocol for Fresh Frozen Human Lymph node Samples adapted from Indiana University
- Negin Farzad1,
- Archibald Enninful1,
- Rong Fan1
- 1Yale University
- Cellular Senescence Network (SenNet) Method Development Community
Protocol Citation: Negin Farzad, Archibald Enninful, Rong Fan 2024. Yale University adapted Akoya Phenocycler-Fusion Tissue Staining and Imaging Protocol for Fresh Frozen Human Lymph node Samples adapted from Indiana University . protocols.io https://dx.doi.org/10.17504/protocols.io.ewov19erylr2/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: October 22, 2024
Last Modified: October 22, 2024
Protocol Integer ID: 110545
Funders Acknowledgement:
Yale TMC for Cellular Senescence in Lymphoid Organs
Grant ID: 1U54AG076043-01
Abstract
This protocol presents the Yale University TMC Akoya Phenocycler-Fusion Tissue Staining & Imaging Protocol for Fixated Fresh Frozen Human Lymph Node Samples. This protocol was adapted from Indiana University adapted Akoya Phenocycler-Fusion Tissue Staining and Imaging Protocol for Fresh Frozen Kidney Samples.
This protocol has been used for FF human lymph node samples for the Cellular Senescence Network (SenNet) Program and the Human BioMolecular Atlas Program consortia. The size of the marker panels has ranged between 35 to 47 targets and includes cell type markers for B cells, T cells, macrophages, and other immune cell types. that will label various cell types Multiple sections have been placed on a single slide and imaged simultaneously.
Materials
Akoya Biosciences Phenocycler-Fusion and Staining Kit
Antibodies of choice
Setup - Day 1
Setup - Day 1
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
Preparing Tissue for Staining
Preparing Tissue for Staining
1h 2m
1h 2m
Remove sample slides from the -80 freezer and immediately place on Drierite beads for 5 minutes
- make sure slide is tissue-side up
5m
Remove slide from Drierite beads and place in Acetone coplin jar for 10 minutes
10m
Place slide in the humidity chamber to dry for 2 minutes
2m
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
2m
Place slide in the second Hydration Buffer coplin jar and incubate for 2 minutes
2m
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
10m
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
30s
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
30s
Place slide in Staining Buffer and incubate for 20-30 minutes
- timing depends on how quickly antibody cocktail can be made
30m
During the Staining Buffer incubation, prepare the Antibody Cocktail
Preparing Antibody Cocktail
Preparing 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
MasterMix_Table.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
MasterMix_Table.xlsx Pipette gently to mix the solution
Tissue Staining
Tissue Staining
1d
1d
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
1d
Set Up - Day 2
Set Up - Day 2
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
Post Staining
Post Staining
51m
51m
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
2m
Place sample slide in the second Staining Buffer coplin jar and incubate for 2 minutes
2m
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
10m
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
30s
Repeat step 33 in the second and third PBS coplin jars
30s
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
5m
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
20m
After the 20 minute incubation, rinse the slides in each of the three PBS coplin jars, as was done previously.
1m
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
Cover-slipping the Slide for Imaging
Cover-slipping the Slide for Imaging
10m 30s
10m 30s
Place slide to be cover-slipped in a fresh jar of 1x PBS and incubate for 10 minutes
10m
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
30s
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.
10m
Preparing the Reporter Plate
Preparing the Reporter Plate
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
MasterMix_Table.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
Imaging the Slides
Imaging the Slides
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.xlsx Analysis Options Available
Analysis Options Available
Segmentation and analysis for each sample are conducted using Qupath software (https://qupath.github.io/).
Downstream analysis following cell segmentation and phenotyping is done using Seurat in a custom R Studio script that can be made available.
Protocol references
Black, S., Phillips, D., Hickey, J.W. et al. CODEX multiplexed tissue imaging with DNA-conjugated antibodies. Nat Protoc 16, 3802–3835 (2021). https://doi.org/10.1038/s41596-021-00556-8