Sep 18, 2025
Hyperion IMC Staining
This protocol is a draft, published without a DOI.
- Michael Haley1
- 1Bioimaging Core Facility, Faculty of Biology, Medicine and Health at University of Manchester
Protocol Citation: Michael Haley 2025. Hyperion IMC Staining. protocols.io https://protocols.io/view/hyperion-imc-staining-habib2akf
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: September 16, 2025
Last Modified: September 18, 2025
Protocol Integer ID: 227402
Keywords: imaging mass cytometry, hyperion, imc, staining protocol, detection of protein target, mass cytometry, using hyperion, standard biotool, ffpe tissue section, protein target, other tissue preparation
Abstract
The staining protocol is for detection of protein targets using Hyperion imaging mass cytometry. It is primarily written for working with FFPE tissue sections, though can be adapted to other tissue preparations. It is nearly identical to the protocol provided by Standard BioTools (see references), just with extra guidance and tips.
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Guidelines
Look for bullet points at the end of steps for specific tips or guidance on those steps.
Materials
- Xylene. Used pure straight from bottle. Note that two different changes of xylene are used, this is intentional. The first will accumulate more paraffin, with the second therefore being 'cleaner'.
- Ethanols. Either used pure, or diluted with water to appropriate percentage.
- Ultra pure or milliQ water. Either should be fine, I would just avoid using distilled water from a normal lab tap unless you are no completely confident in its quality. Historically, I have had doubts about the distilled water in the AV Hill building, especially when they have cleaned the pipes then not washed the excess cleaning reagents out properly!
- Tris-EDTA solution. 10 mM Tris base, 1 mM EDTA solution, pH 8.5. This is a fairly standard solution used for antigen retrieval, with several manufactures providing their own premade solutions or stocks that should be fine.
- PAP pen. Any brand should be fine, but once opened they can 'go off' with time, or dry out.
- Staining chamber. This should be humidified, usually with a water reservoir of some kind. The entire point is to keep the slides in a moist environment which will prevent even small volumes evaporating from the surface of the slide during the room temperature and over night incubations.
- 10% BSA stock. We used a 10% BSA stock supplied by ThermoFisher which is also stabilised with something antimicrobial. You can also make up your own stock, but beware that BSA stocks can become contaminated with bacteria, or form aggregates
- Block buffer. 3% bovine serum albumin (BSA) in PBS. Dilute from 10% BSA stock.
- PBS. I would suggest using bottled 1X PBS rather than making your own.
- 0.1% Triton X100-PBS. Add 500 μl of Triton X-100 to a 500 ml bottle of PBS and shake to mix.
- Iridium. Stock is Cell-ID‱ Intercalator-Ir—500 µM from Standard Biotools. Used 1:400 in PBS.
Troubleshooting
Safety warnings
Anyone familiar with immunohistochemistry should be familiar with most steps in this protocol. However, this protocol rewards being exact and careful, and can be very unforgiving if you are sloppy, or do not follow the protocol. The approximate cost to stain one slide with an entire panel is £300-400! Therefore, in even small pilot experiments can add up fairly quickly.
Before start
- Make sure you have everything in-place, as these experiments can be very expensive, as both human tissue and antibodies are very precious.
- Calculate your dilutions and volumes required for all antibodies in supplied spreadsheet. Assemble your panel of antibodies to ensure none are missing.
- Gather your tissues to ensure they are all present and fit for staining.
- FFPE sections should ideally be freshly cut and stored in the fridge. We have noticed sections older than 6 months start to see a loss of signal in some antigens.
Preparation
3h
Complete attached spreadsheet (Hyperion_Panel_Template_2025.xlsx attached to protocol, or create your own) detailing your panel de
Note
Your plan should include:
- The date you performed the staining
- What tissue you used, and exactly which slides
- What antibodies are in your panel (including unique identification numbers, if available)
- What concentrations you antibodies will be used at (e.g. 1 in 50)
- The total volume of antibody cocktail you need to stain all your tissue (usually 150 µL per section), preparing enough extra to allow for pipetting error (50 µL )
- For a complex experiment, allow time to double check everything!
1h
Make sure you have everything in-place, as these experiments can be very expensive, as both human tissue and antibodies are very precious
Assemble your panel of antibodies to ensure none are missing
Gather your tissues to ensure they are all present and fit for staining.
Note
FFPE sections should ideally be freshly cut and stored in the fridge. We have noticed sections older than 6 months start to see a loss of signal in some antigens.
Turn on the water bath and set it to 96 °C . Place the Tris-EDTA solution in the water bath to preheat for antigen retrieval.
If using FFPE (paraffin embedded) tissue, bake slides at 60 °C in an oven for 02:00:00 .
Note
This can help with tissue adherence, and isn't always required if tissue is very adherent. If you're unsure, then include this step, as it can only help.
2h
Deparaffinisation
33m
If using FFPE (paraffin embedded) tissue, slides must undergo deparaffinisation using xylene and graded alcohols.
Note
- Use separate reagents to those used for normal immunohistochemical staining to prevent contamination.
- Between reagents, briefly tap off excess to minimise the amount of carry over.
- Depending on how many slides you have, this can be performed in Falcon tubes, Coplin jars, or in racks in staining jars. Slide can be placed back-to-back, but be careful not to put them face-to-face.
- Incomplete removal of paraffin will ruin staining, as antibodies will not penetrate into tissue.
- Change reagents regularly - these reagents are cheap to replace, whereas antibodies from a failed run are not. Xylene will accumulate paraffin over time, and can become contaminated with water.
Xylene (100%) 00:10:00
10m
Xylene (100%) 00:10:00
10m
100% ethanol 00:02:00
2m
90% ethanol 00:02:00
2m
70% ethanol 00:02:00
2m
50% ethanol 00:02:00
2m
Water (ultrapure / Milipore) 00:05:00
Note
Can be longer if water bath isn't yet at temperature
5m
Antigen retrieval
48m
Place slides into pre-heated Tris-EDTA solution for 00:30:00
Note
- Depending on how many slides you have, this can be performed in Falcon tubes, Coplin jars, or in racks in staining jars. Slide can be placed back-to-back, but be careful not to put them face-to-face.
- In theory, alternative antigen retrieval buffers may be used. However, it is important to ensure that all antibodies are optimized using the same buffer, as different buffers can affect antigenicity in antibody-specific ways.
- During the antigen retrieval steps do NOT allow the Tris-EDTA buffer to dry out on the surface of the slides, which will happen very quickly if you deviate from this protocol and allow hot slides to air dry. All antigenicity in the tissue will be lost, and staining will be absent or patchy.
30m
Take containers (e.g. falcons, jars, etc, that contain the slides) out and let cool to 70 °C , monitoring using a thermometer. This usually takes ~10 min
Note
If you are using Falcon tubes, you can put the tubes in cold water to speed up the cooling. Do NOT do this with glass, as the rapid change in temperature can cause the glass to shatter.
10m
Replace liquid with PBS for 00:08:00
Note
Careful changing liquids not to pour the PBS directly onto the tissue face, and do allow the slides to sit without liquid and dry.
8m
Staining (day 1)
1h 16m
For each slide in-turn, we need to use a PAP pen to create a hydrophobic barrier to act as a reservoir for the antibody cocktail.
Note
- Complete these steps for each slide in-turn.
- Try and minimise the amount of drying, though some small amount of drying is sometimes hard to avoid, especially at the tissue edges. As there is only PBS present, this shouldn't cause any issues, but in future steps it is critical that tissue drying out is avoided.
- Prime the PAP-pen before use (usually by pressing down the nip), and make sure it's not dried out. The hydrophobic compounds in old pens can sometimes start to break down with age. You can diagnose this as it looks like the wash will get contaminated with an oily sheen, and/or it may not completely dry.
- Avoid touching the tissue with the PAP pen, it can leech into the tissue and cause non-specific binding on antibodies. Practice drawing with the PAP pen on spare slides if you are unfamiliar with their use.
30m
Remove slide from PBS and flick off excess.
Ensure there is a dry area around the tissue where we will be able to draw the PAP pen. You can use a cotton bud if required, being careful not to damage the tissue.
Draw around the tissue using the PAP pen, creating a complete ring. Leave a couple of mm gap between the pen and tissue.
Place slide flat in staining chamber (see materials).
Carefully add a few drops of blocking solution to the tissue. This should be enough to keep the tissue moist whilst the PAP pen dries.
Wait for PAP pen to dry
1m
Add block (3% BSA in PBS) for 00:45:00 at Room temperature
Note
Block can be left for longer if required, just be careful it doesn't dry out. Should be fine for 2 hours in a humidified staining chamber.
45m
Make up antibody panel
42m
This is a simple process of adding the antibodies in turn, but be careful and systematic in your approach! You can do this at any point, but I usually do it whilst the block is on.
Note
- Double check the staining volume and antibodies before you start.
- Include extra visual inspections of vials.
- Clearly track which antibodies you've added to the antibody cocktail. For example, I tick them off as I go, and move them from one rack into another.
- I take a picture of the top of the rack of the panel, so I have another record of the vials I used.
40m
Thoroughly vortex all antibodies for 00:02:00 , I usually do this with them all in the rack
2m
Following previously prepared spreadsheet, add appropriate amount of PBS and 10 % (v/v) BSA stock to microtube. The final concentration of BSA should be 0.5 % (v/v) once all antibodies are added.
Add each antibody in turn, pipetting from the top of each vial.
Keep antibody cocktail in the fridge until use. We will be centrifuging it immediately before use!
Staining (day 1)
34m
Vortex the antibody cocktail, then centrifuge 10000 x g, 00:02:00 , Any desktop microcentrifuge
Note
- The purpose of this is to spin down the aggregates (i.e. clumps of antibody that have stuck together) so that they are right at the bottom of the tube. These aggregates usually form over time in old vials, and are also more common in indium (113, 115) and platinum (194+) conjugated antibodies.
- Centrifugation should be done immediately prior to being added to slides.
- Alternatively, centrifuge the cocktail and carefully transfer the supernatant (i.e. pipette from the top of the liquid) to a new tube, leaving the final ~25 µL behind to avoid disturbing any aggregates.
- If aggregates are still present in your cocktail, they will show up as extremely bright specks that are smaller than cells. These can sometimes computationally removed, but can sometimes ruin the staining as they can be incredibly bright and spill through to several channels.
2m
Working through each slide in turn to minimise drying...
Flick off excess block (do not wash!)
Add antibody cocktail into the PAP-penned area, usually 150 µL per section. Pipette from the top of the liquid. Do not disturb the very bottom of the cocktail. Once you have added the stain to all slides, there should still be ~25 µL behind in the microtube.
Note
- If you already transferred the cocktail to a new tube after centrifugation, then you can just use the entire volume, as the aggregates should be left in the first tube.
- Make sure the entire surface of the tissue is covered, you can carefully move the liquid around with the tip of your pipette as you're adding it.
30m
Incubate Overnight in fridge at 4 °C in staining chamber.
2m
Staining (day 2)
1h 34m
Note
Washing off antibody cocktail...
- Each previous wash can be poured or flicked off, we want or don't need to entirely remove the previous wash, as that would risk the slide drying out.
Depending on how well adhered your tissue, you can adopt one of two strategies for washes:
- Use a pasteur pipette to carefully fill the entire surface of the slide with wash buffer, filling over the top of the PAP-pen areas. This gives a staining volume of 2-3 ml per wash. For well adhered tissue, this is usually fine, but adding the liquid can occasionally dislodge tissue.
- Place the entire slide into a container filled with wash for the duration of the wash. For example, back-to-back in Falcon tubes. This method allows for a much larger washing volume (40 ml), and lowering the slide into the wash is usually gentler and less likely to dislodge tissue.
Wash with 0.1% Triton X100-PBS for 00:08:00
8m
Wash with 0.1% Triton X100-PBS for 00:08:00
8m
Wash with PBS (1X) for00:08:00
8m
Add Iridium (1:400 in PBS) for 00:30:00 at Room temperature
- Make up 200 µL per slide
- Iridium stocks are kept in the freezer (-20 °C ), and can be freeze-thawed repeatedly
- Use 1 in 400 in PBs, e.g. 1 µL Iridium in 399 µL PBS
Note
WARNING: Without this step you will not be able to computationally analyse your data, as almost all methods depend on having strong and clear nuclear staining!
30m
Wash with PBS (1X) for 00:05:00
5m
Wash with deionised Millipore (or ultrapure water) by submerging entire slide (e.g. in Falcon) for 00:05:00
Note
We want to avoid PBS crystals, so submerging the entire slide is preferable
5m
Air dry and store at room temperature in a dry place - do NOT coverslip!
Note
Slides are ready to image once dry and are stable for years once stained
30m
Protocol references
This is adapted from (and is almost identical to) the protocol provided by Standard Biotools, available here:
https://www.standardbio.com/support/instrument-support/hyperion-support (under 'Protoocols')
The only differences are in timings of some of the washes thar we took from previous iterations of their online protocols. These should have little-to-no impact.