TITLE: MIBI staining
AUTHORS: Marc MB Bosse, Sean  Bendall, Mike  Angelo

[DESCRIPTION]
This protocol is the standard FFPE tissue staining procedure recommended for Multiplex Ion Beam Imaging Time of Flight instrument (MIBI_TOF) developed in the Sean C. Bendall and Michael R. Angelo labs. The protocol has been successfully used for MIBI and is the result of extensive optimization experiments. It is inspired from state-of-the art of immunohistochemistry staining procedures but differs in some very important steps, namely, glutaraldehyde fixation and final washes prior tissue dehydration. Failure to follow exactly all steps described in this procedure may result in inconsistencies in output data after MIBI_TOF acquisition.   


[BEFORE_START]
Verify the stocks of all reagents and place an order or prepare solutions, if some reagents are running low.


[GUIDELINES]
Staining tissue sections is fairly straightforward but there are few things to be cognizant of, when preparing samples:
Always try to limit mechanical damage to the sample surface. This can occur when moving the samples with forceps, particularly when the mounting substrates are small.
Once the samples have been rehydrated, they cannot dry out until the end of the protocol.
Be careful at all times to not touch the tissue, in order to not leave any residue.
Once the samples have been stained, fixed, and dehydrated, they have an indefinite shelf life and can be imaged at any time but need to be stored properly such as in a vacuum chamber or in a sealed vacuum bag
MIBI_TOF observes the basic principles of Mass spectrometry. Any contaminant ions present in water or in the air can potentially compromise the integrity of the sample.
Therefore to prevent potential contamination, it is important to always use single use lab ware containers to make the solutions. The protocol has been validated using the level of precision of graduated Nalgene bottles and 50 mL tubes. 
The use of washed beakers or graduated cylinders is not considered as good a practice, due possible introduction of contaminants (exemples of sources: barium from lab ware soap or calcium from air dried lab ware)

[STEPS]
SECTION: Slide for MIBI
1. FFPE or frozen sections should be deposited on special conductive slides for MIBI
It is recommended to use freshly cut tissue sections.  Otherwise tissue section slides should be stored properly using different state of the art methods (vacuum chamber, nitrogen chamber or vacuum sealed bags)
  

SECTION: Slide baking and PT module preparation
2. Bake the sections at 70 °C for 20 min in a dry incubator
    Optional : 60 min ; 60 min
   
 

SECTION: Slide baking and PT module preparation
2.1. Last 10 min place the slide (s) vertically with the label side up to allow drip down the paraffin
SECTION: Slide baking and PT module preparation
3. Prepare Target retrieval solution
2.5 mL  of target retrieval solution 10x (3-in-1), DAKO
in 22.5 mL of ultrapure (type 1,  >18 MOhms) water 

 

SECTION: Slide baking and PT module preparation
4. Put in the containers with the diluted target retrieval solution in the PT Module


   
 



SECTION: Slide baking and PT module preparation
5. PT Module Preheat 
Press RUN on digital screen and check for PREHEAT 75 on display

   




SECTION: Slide deparaffination
6. Linear Stainer
Pour out reagent containers and fill with fresh reagents:  

Xylene x 3, 100% Ethanol x 2, 95% Ethanol x 2, 80% Ethanol, 70% Ethanol, ddH2O x 2, exit stainless steel tank = ultrapure (type 1,  >18 MOhms) water 


 

  



SECTION: Slide deparaffination
6.1. Prior loading the slides on the Linear Stainer, immerse the slides in a container with fresh xylene. Make sure that the xylene solution covers all the paraffin. Incubate for a minimum of  5 min 
 


SECTION: Slide deparaffination
7. Insert slides into slide carriers

Place the slide carrier into first xylene container

SECTION: Slide deparaffination
8. Press on Menu 

Check for Processing time = 30 sec, Lift bar = 976, Number of dips = 3 

Continue to press Menu until the screen displays Start at: __

Set Start position corresponding to the first slide carrier position 
        
Exemple: If the first slide carrier is at position 4, use Plus (+) or Minus (-) button to increase or decrease to get Start at: 04 

SECTION: Slide deparaffination
8.1. Then press Enter 

SECTION: Slide deparaffination
8.2. Synchronize when the PT module temperature has reached 75˚C 

then Press Run on the Linear Stainer

SECTION: Slide deparaffination
8.3. Allow the rehydration process and wait until the slides have reached the stainless steel tank and stop
SECTION: Slide deparaffination
9. Bring the stainless steel tank with the slides in close to the PT module
SECTION: Antigen Retrieval
10. Open the PT Module and insert the slides in the warm Target retrieval solution container

Discard water immediately from the stainless steel tank 
SECTION: Antigen Retrieval
11. Press RUN again and check for first  WARMUP  then  HEAT on display, once the temperature has reached 97˚C
  

  
 

SECTION: Antigen Retrieval
12. Verify stock of 1x PBS wash buffer and prepare accordingly if running low


 

SECTION: Antigen Retrieval
13. Allow to run for 40 min at97 °C and then cool down for approximately 50 min and reach 65 °C

SECTION: Antigen Retrieval
14. When the alarm sounds Stop the PT module
SECTION: Antigen Retrieval
15. Take out the slides and let cool down at room temperature for at least 5 min 

SECTION: Antigen Retrieval
16. Prepare two coplin jars filled with MIBI 1x PBS wash buffer

SECTION: Antigen Retrieval
17. Transfer the slides in the first MIBI 1x PBS wash buffer and use orbital shaker set for 5 min, 70 rpm

SECTION: Antigen Retrieval
18. Transfer the slides to the second 1x PBS wash buffer and use orbital shaker set for 5 min, 70 rpm
SECTION: Hydrophobic barrier pen
19. Make sure to dry with a folded-tissue paper the slide, leaving a square of wet surface surrounding of the tissue section 
  
Note: Do not let AIR DRY the tissue section, this will result high background and false positive staining
SECTION: Hydrophobic barrier pen
20. Draw a square following the outside edges of the wet square with an hydrophobic barrier pen (ImmEdge pen)

  


SECTION: Optionnal: blocking endegenous biotin
21. If a biotinylated antibody or a probe is used, it is recommended to block endogenous biotin

Place the slides in the moist chamber

Add drops of Avidin solution (Avidin/Biotin blocking kit, Biolegend) sufficient to cover the sample and incubate for 10 min at Room temperature

SECTION: Optionnal: blocking endegenous biotin
21.1. Wash in coplin jar with MIBI 1x PBS wash buffer and use orbital shaker set for 5 min, 70 rpm
SECTION: Optionnal: blocking endegenous biotin
21.2. Add drops of Biotin solution sufficient to cover the sample and incubate for 10 min at Room temperature

SECTION: Optionnal: blocking endegenous biotin
21.3. Wash in coplin jar with MIBI 1x PBS wash buffer and use orbital shaker set for 5 min, 70 rpm
SECTION: Optionnal: blocking endegenous biotin
21.4. Next day, use anti-biotin metal-labeled antibody (clone 1D4-C5) in Stain 2 panel
SECTION: Blocking
22. Add 100 µL of Blocking Buffer for 18 mm2

For blocking solution preparation refer to MIBI and IHC solutions protocols




   

 


SECTION: Blocking
23. Place the slides in a moist chamber at Room temperature and incubate  20 min to  60 min 

SECTION: Multiplex Antibody mix
24. Prepare antibody mix based on the putative multiplex antibody panel

Make sure that all the antibodies are ready to use BEFORE starting to build the panel

It is highly recommended to prepare all the antibodies, ready to use, a day before the panel is built

SECTION: Multiplex Antibody mix
24.1. Evaluate the total volume of multiplex antibody mix by counting the number of slides and the surface area per slide

Refer to the chart for the volume of antibody to apply according to the estimated surface area
 

 
 
SECTION: Multiplex Antibody mix
24.2. Build an antibody mix table information to make the antibody panel as follow:


Conjugation ID, Target name, Channel, Antibody concentration, Titer, Volume 

Exemple:
 

 
For Antibody Diluent (NHS 3%)  solution preparation refer to MIBI and IHC solutions protocols





SECTION: Multiplex Antibody mix
25. Add 400 µL of antibody diluent (NHS 3%) to a Centrifugal 0.1 µm filter unit (Millipore, UFC30VV00)

SECTION: Multiplex Antibody mix
25.1. 10000 rcf, 1 min 
SECTION: Multiplex Antibody mix
25.2. Discard flow through

SECTION: Multiplex Antibody mix
26. Add antibody mix to the filter unit 

SECTION: Multiplex Antibody mix
26.1. 10000 rcf, 1 min 
SECTION: Stain 1 (Overnight)
27. Remove the blocking solution by tapping the slide on a side

Immediately add the filtered multiplex antibody mix


SECTION: Stain 1 (Overnight)
28. Place the moist chamber at 4˚C  , preferably in a place with low disturbance (e.g. a designated area in a cold room)

SECTION: Wash buffer
29. Prepare two Coplin jars filled with 1x PBS wash buffer
SECTION: Wash buffer
29.1. Transfer the slides into the first Coplin jar and use orbital shaker set for 5 min, 70 rpm
SECTION: Wash buffer
29.2. Transfer the slides into the second Coplin jar and use orbital shaker set for 5 min, 70 rpm
SECTION: Stain 2 (1h)
30. 
Add adequate volume of the selected sub-panel of antibody mix

Refer to the chart for the volume of antibody to apply

 


SECTION: Stain 2 (1h)
31. Place sample in a sealed humidity chamber, transfer to 4oC refrigerator, and incubate 60 min 
SECTION: Wash buffer
32. After 1h incubation 
   than go to step 33

SECTION: Prepare solutions
33. Prepare fresh glutaraldehyde fixing solution

Glutaraldehyde fixing solution

Add 30 mL of 1x PBS low barium in a 50 mL tube
Break the glass glutaraldehyde 8% (amber vial) 
Add the content of the glutaraldehyde (10 mL) by inverting it and tapping the bottom of the vial in the 50 mL tube
Transfer the content in a linear stainer container

SECTION: Prepare solutions
34. Set the linear stainer containers

Fill containers with the following solution and order

Glutaraldehyde x 1, PBS low barium x 1, TRIS 100 mm pH 8.5 x 3, ddH2O x 2, 70% Ethanol x1, 80% Ethanol x1, 95% Ethanol x 2, 100% Ethanol x 2, exit stainless steel tank = empty
SECTION: Glutaraldehyde fixation
35. Mount the slides on the linear slide holder

Fix for 5 min 

SECTION: Glutaraldehyde fixation
36. Rinse briefly with 1x PBS low barium 
SECTION: Dehydration and Storage
37. Press on Menu 

Check for Processing time = 30 sec, Lift bar = 976, Number of dips = 3 

Continue to press Menu until the screen displays Start at: __

Set Start position corresponding to the first slide carrier position 
        
Exemple: If the first slide carrier is at position 3, use Plus (+) or Minus (-) button to increase or decrease to get Start at: 03 

SECTION: Dehydration and Storage
37.1. Then press Enter 
SECTION: Dehydration and Storage
37.2. Press Run on the Linear Stainer
SECTION: Dehydration and Storage
37.3. Allow the dehydration process and wait until the slides reached the empty stainless steel tank and stop
SECTION: Dehydration and Storage
37.4. Store the slides immediately under vacuum until MIBI acquisition

Alternatively, the stained slides can be stored in a vacuum sealed bag for longterm storage pre and post MIBI acquisition