Dec 13, 2023

Public workspaceCODEX FFPE Staining and Fixation 

DOI
https://dx.doi.org/10.17504/protocols.io.n92ldm6y9l5b/v1
  • wbei 1
  • 1Stanford University
  • Human BioMolecular Atlas Program (HuBMAP) Method Development Community
    Tech. support email: [email protected]
Joanna Bi
Icon indicating open access to content
QR code linking to this content
DOI: https://dx.doi.org/10.17504/protocols.io.n92ldm6y9l5b/v1
Protocol Citation: wbei 2023. CODEX FFPE Staining and Fixation . protocols.io https://dx.doi.org/10.17504/protocols.io.n92ldm6y9l5b/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: December 13, 2023
Last Modified: December 13, 2023
Protocol Integer ID: 92211
Keywords: codex ffpe staining, akoya flowcells in the akoya phenocycler, akoya phenocycler, akoya flowcell, fixing ffpe slide, ffpe slides for use, akoya protocol, codex, slide
Abstract
Detailed protocol for preparing, staining, and fixing FFPE slides for use with Akoya flowcells in the Akoya phenocycler (CODEX). Slides are ready to be used with the Akoya phenocycler and the Akoya protocol following this protocol.
Troubleshooting
Staining
4h 44m 30s
Bake slides at Temperature70 °C for at least 1 hour in an oven/incubator

Note
Work on 8 slides at a time

1h
Deparaffinize and rehydrate the slides


Incubate slides for Duration00:21:00 in xylene in a coplin jar

21m
Place slides in ST4020 Linear Staining vial and start the staining protocol
Each step is 3 minutes
Xylene x3 -> 100% EtOH x2 -> 95% EtOH x2 -> 80% EtOH x1 -> 70% EtOH x1 -> ddH2O x3

Note
It is okay for some of the slides to only have 2 xylene and/or 2 ddH2O steps. If using the ST4020 Linear Stainer, you can move the slides in the front vial behind the slides in the back vial for the xylene, and remove the slides in the front vial once it has 3 ddH2O steps. Keep slides in ddH2O until the next step


36m
After starting the linear staining step, fill slide chamber with HIER 1X buffer and incubate HIER buffer at Temperature75 °C (Temperature170 °F ) in pressure cooker filled with enough water (cover chamber with aluminum foil)

Transfer slides to chamber containing heated HIER buffer
Put the chamber back in the pressure cooker, heat to Temperature97 °C (Temperature205 °F ) and incubate for Duration00:17:30 min
Note
Temperature and incubation time is crucial here


17m 30s
Stop the pressure cooker and turn it off, leave the chamber in the water bath for Duration00:20:00 to cool down slowly.
20m
Take the chamber out of pressure cooker, cool down at RT for about Duration00:30:00 until around RT

30m
Wash tissue
Place slide in coplin jar containing 80 mL of 1X TBS IHC Wash Buffer with Tween 20
(https://www.cellmarque.com/ancillaries/CM/2087/TBS-IHC-Wash-Buffer-Tween-20)
Incubate on a shaker for Duration00:10:00 at around 100 rpm

10m
Make blocking buffer solution (amount depends on sample area. ~120-140 uL/slide)
1 mL = 780 uL of S2 (RT), 50 uL of B1, 50 uL of B2, 50 uL of B3, 70 uL of BC4
NOTE: can store remaining buffer at 4°C
Block
Tap off excess wash buffer, wipe edges and back with Kim Wipes, and place slides in humidity chamber (or use pipette box with water and paper towel underneath)
Add 120-140 ul (depending on the sample area) of blocking buffer and incubate for Duration01:00:00 at RT in humidity chamber
(Can leave much longer and just need to add solution so tissue does not dry out)

1h
Dilute antibodies in blocking buffer to a total of 120 ul (ratio of blocking buffer: antibody cocktails should be >= 1 v/v).
Antibody Staining
After 1 hour of blocking, tap off excess buffer and add 120 ul of conjugated antibody solution. Cover tissue area with Parafilm

Incubate DurationOvernight at Temperature4 °C in humidity chamber on a shaker
30m
Fixation
42m
Wash tissue.
Place slide in chamber containing S2 buffer.
Incubate for Duration00:04:00 on a shaker

4m
Fix tissue. Prepare 1.6% PFA (dilute from 16% PFA) solution in S4 buffer (1:10 (v/v)). NOTE: Use fresh vial of PFA every 1-2 weeks. 
Place slide in humidity chamber and add 100 uL of PFA solution or enough to cover the tissue
Incubate for Duration00:10:00 .

10m
Wash tissue.
Place slide in chamber containing 1x PBS for Duration00:01:00 on a shaker

1m
Ice-cold methanol incubation. Place slide chamber on an ice and fill with cold methanol (Temperature4 °C ).

Remove slide from chamber containing 1x PBS and place in chamber containing ice-cold methanol.
Incubate for Duration00:05:00 .

5m
Wash tissue.
Remove slide from cold methanol and place in chamber containing 1x PBS (ok to use same PBS as from step #10).
Incubate for Duration00:01:00 on a shaker

1m
Fix tissue. Prepare final fixative solution. Remove FIX aliquot from -20°C freezer right before use and let it melt. Add entire contents (~20 ul) to 1 ml of 1x PBS. Mix fully.
Add 100 uL of fixative solution (or enough to cover the tissue), taking care not to pipette directly onto tissue.
Incubate for Duration00:20:00 in a humidity chamber.

20m
Wash tissue.
Place slide in chamber containing 1x PBS for Duration00:01:00 on a shaker

1m
Assemble the Akoya flowcells to the slides directly or store slides in S4 buffer, and assemble later.