Immunohistochemistry Protocols

ABC Immunohistochemistry Protocol – Antigen Retrieval from Paraffin Sections

Day 1
Deparaffinization and tissue rehydration

  • Melt paraffin/sections in oven at 65˚C for 30 min. Remove slides to cool to RT. (Fill glass Coplin jars with citrate antigen retrieval buffer, place in water bath, and start warming bath so it has time to reach 92-94˚C.)
  • Begin rehydration while bath warms up.
  • Xylene – 2 x 5 min 
  • 100% EtOH – 2 x 2 min 
  • 95% EtOH – 2 x 2 min 
  • 75% EtOH – 1 x 2 min
  • 50% EtOH – 1 x 2 min
  • dH2O – 2 x 5 min
  • 0.1 M PBS (pH 7.3) – 1 x 5 min

 

Antigen Retrieval

  • Incubate slides in preheated citrate buffer (10 mM sodium citrate, 0.05% tween 20, pH 6.0 OR use Retrieve-All) for 40 min at 92-94˚C.
  • Remove Coplin jars and allow slides to cool to RT
  • Transfer slides to Coplin jar and wash in dH2O – 3 x 5 min
  • Using the PAP Pen, carefully draw a water barrier circle around the tissue sections on the slide – allow this circle to dry for several seconds or up to approx. one min. (add drop of dH2O to the tissue sections during this process so the tissue does not dry out)

 

Immunostaining

  • Rinse slides with PBS (pH 7.3-7.4): 4 x 5 min each
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 in PBS):  1 x 10 min
  • Wash slides with 1.0% H2O2 in PBS:  1 x 15 min (2 ml of 30% H2O2 in 60 ml of PBS-This amount fills one coplin jar)
  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Remove slides one at a time and using a clean Kimwipe, carefully wipe around the tissue sections to dry the slide
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with BLOCKING BUFFER (3 drops of normal serum provided in the ABC kit + 10 ml of 1.0% BSA + 0.4% Triton X-100+ PBS)
  • Allow the sections to remain in blocking buffer for 1.5-2 hrs. at RT
  • Pour off the blocking buffer
  • Replace with primary antibody solution (antibody of choice diluted in the SAME BLOCKING BUFFER as used above)
  • Incubate overnight at RT

 

 

Day 2

  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • While rinsing slides above, prepare the biotinylated secondary antibody solution. (Blocking buffer = 3 drops of normal serum (provided in the ABC kit) + 10 ml of solution containing 1.0% BSA + 0.4% Triton X-100 + PBS. Add 1 drop of biotinylated secondary antibody from the ABC kit to the Blocking Buffer Solution).  
  • Cover the tissue with the secondary antibody solution and incubate for 2 hrs. at RT in the humidity box
  • Prepare the ABC reagent from the Vector Kit at least 30 min prior to using. ABC reagent is made by adding 1 drop of solution A + 1 drop of solution B to 2.5 ml of solution containing 1.0% BSA + 0.4% Triton X-100 + PBS. Let this sit at RT while performing the following rinses
  • **Add 1% solution first, then add solutions A and B to it. ORDER MATTERS (can render a non-reactive solution if A and B added first)**
  • Pour off the secondary antibody solution
  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Cover the tissue with the ABC reagent and incubate at RT for 1-1.5 hrs. in the humidity box
  • Prepare 50 mM Tris buffer (Trizma, pH 7.6) for the next rinse stage – (1.49 g Trizma (pH 7.6, Sigma T7943) + 200 ml dH2O
  • Pour off the ABC reagent from the slides and place slides into a Coplin jar
  • Rinse the slides with the Tris buffer:  4 x 5 min
  • During the last Tris rinse, prepare the VIP development solution – must be at RT for proper use (Vector ImmPACT VIP Kit, SK4605). Use 5 ml VIP diluent + 3 drops each of solutions 1,2, 3, and 4 from the kit.
  • Place the slides onto a light-colored background to monitor color development.
  • Add the VIP color development solution to the tissue sections.
  • Monitor development of color, which should take from 2 to 20 min to reach maximum intensity. (Use the bench scope to monitor)
  • Stop color development by submersing slides into dH2O in a Coplin jar
  • Rinse with dH2O – 2 x 2 min
  • Before attaching coverslips to each slide, the tissue must be dehydrated via the reverse process of rehydrating at the beginning of this protocol (i.e., take tissue from 50% EtOH through the proper EtOH washes to Xylene)
  • Attach coverslips using Cytoseal
  • Allow to air dry overnight

 

ABC Immunohistochemistry Protocol – (VIP chromogen and  frozen sections)

Day 1
Immunostaining

  • Using the PAP Pen, carefully draw a water barrier circle around the tissue sections on the slide – allow this circle to dry for several seconds or up to approx. one min. (the tissue sections should not be allowed to dry out)
  • Rinse slides with PBS (pH 7.3-7.4): 4 x 5 min each
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 in PBS):  1 x 10 min
  • Wash slides with 1.0% H2O2 in PBS:  1 x 15 min (2 ml of 30% H2O2 in 60 ml of PBS-This amount fills one coplin jar)
  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Remove slides one at a time and using a clean Kimwipe, carefully wipe around the tissue sections to dry the slide
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with BLOCKING BUFFER (3 drops of normal serum provided in the ABC kit + 10 ml of 1.0% BSA + 0.4% Triton X-100+ PBS)
    •  Goat Kit: Use rabbit normal serum
    •  Rabbit Kit: Use goat normal serum
  • Allow the sections to remain in blocking buffer for 1.5-2 hrs. at RT
  • Pour off the blocking buffer
  • Replace with primary antibody solution (antibody of choice diluted in the SAME BLOCKING BUFFER as used above)
  • Incubate overnight at RT

 

Day 2

  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • While rinsing slides above, prepare the biotinylated secondary antibody solution. (Blocking buffer = 3 drops of normal serum (provided in the ABC kit) + 10 ml of solution containing 1.0% BSA + 0.4% Triton X-100 + PBS. Add 1 drop of biotinylated secondary antibody from the ABC kit to the Blocking Buffer Solution). 
    • Goat Kit:  Biotinylated rabbit anti-goat
    •  Rabbit Kit: Biotinylated goat anti-rabbit 
  • Cover the tissue with the secondary antibody solution and incubate for 2 hrs. at RT in the humidity box
  • Prepare the ABC reagent from the Vector Kit at least 30 min prior to using. ABC reagent is made by adding 1 drop of solution A + 1 drop of solution B to 2.5 ml of solution containing 1.0% BSA + 0.4% Triton X-100 + PBS. Let this sit at RT while performing the following rinses
  • **Add 1% solution first, then add solutions A and B to it. ORDER MATTERS (can render a non-reactive solution if A and B added first)**
  • Pour off the secondary antibody solution
  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Cover the tissue with the ABC reagent and incubate at RT for 1-1.5 hrs. in the humidity box
  • Prepare 50 mM Tris buffer (Trizma, pH 7.6) for the next rinse stage – (1.49 g Trizma (pH 7.6, Sigma T7943) + 200 ml dH2O
  • Pour off the ABC reagent from the slides and place slides into a Coplin jar
  • Rinse the slides with the Tris buffer:  4 x 5 min
  • During the last Tris rinse, prepare the VIP development solution – must be at RT for proper use (Vector ImmPACT VIP Kit, SK4605). Use 5 ml VIP diluent + 3 drops each of solutions 1,2, 3, and 4 from the kit.
  • Place the slides onto a light-colored background to monitor color development.
  • Add the VIP color development solution to the tissue sections.
  • Monitor development of color, which should take from 2 to 20 min to reach maximum intensity. (Use the bench scope to monitor)
  • Stop color development by submersing slides into dH2O in a Coplin jar
  • Rinse with dH2O – 2 x 2 min
  • Before attaching coverslips to each slide, the tissue must be dehydrated via the reverse process of rehydrating at the beginning of this protocol (i.e., take tissue from 50% EtOH through the proper EtOH washes to Xylene)



 

  • Xylene – 2 x 5 min 
  • 100% EtOH – 2 x 2 min 
  • 95% EtOH – 2 x 2 min 
  • 75% EtOH – 1 x 2 min
  • 50% EtOH – 1 x 2 min

 

  • Attach coverslips using Cytoseal
  • Allow to air dry overnight

Fluorescence Immunohistochemistry Protocol-For Mounted Slides

Day 1

  • Using the PAP Pen, carefully draw a water barrier circle around the tissue sections on the slide – allow this circle to dry for several seconds or up to approx. one min. 
  • Rinse slides with PBS (pH 7.3-7.4): 4 x 5 min each
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 in PBS):  1 x 10 min
  • Remove slides one at a time and using a clean Kimwipe, carefully wipe around the tissue sections to dry the slide.
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with blocking buffer (10% normal donkey serum in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Allow the sections to remain in blocking buffer for 2 hrs. at RT
  • Pour off the blocking buffer
  • Replace with primary antibody solution (antibody of choice diluted in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Incubate tissue with primary antibody overnight in incubation box.    

 

 

Day 2

  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with blocking buffer (10% normal donkey serum in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Allow the sections to remain in blocking buffer for 2 hrs. at RT
  • Prepare fluorescent secondary antibody (secondary antibody should be diluted in 1.0% BSA + 0.4% Triton X-100 + PBS).
  • Cover the tissue with the secondary antibody solution and incubate for 2 hrs. at RT in the incubation box. *From this point on, use low light and/or cover tissues.*
  • Rinse slides with PBS:  4 x 5 min
  • Remove excess PBS with a Kimwipe.
  • Carefully add a drop of mounting medium to the center of the tissue and apply cover glass.
  • Seal cover glass with clear nail polish. For thicker tissue, add a weight before sealing. 

 

Biotin-Streptavidin Fluorescent Immuno Protocol-For Mounted Slides

 

Day 1

  • Using the PAP Pen, carefully draw a water barrier circle around the tissue sections on the slide – allow this circle to dry for several seconds or up to approx. one min. 
  • Rinse slides with PBS (pH 7.3-7.4): 4 x 5 min each
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 in PBS):  1 x 10 min
  • Remove slides one at a time and using a clean Kimwipe, carefully wipe around the tissue sections to dry the slide.
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with blocking buffer (10% normal donkey serum in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Allow the sections to remain in blocking buffer for 2 hrs. at RT
  • Pour off the blocking buffer
  • Replace with primary antibody solution (antibody of choice diluted in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Incubate tissue with primary antibody overnight in incubation box 

 

 

Day 2

  • Rinse slides with PBS:  4 x 5 min
  • Rinse slides with 0.5% BSA + 0.4% Triton X-100 + PBS:  1 x 10 min
  • Place the slides into a black, covered slide incubation box/humidity box
  • Cover the tissue sections with blocking buffer (10% normal donkey serum in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Allow the sections to remain in blocking buffer for 1.5 hrs. at RT
  • Prepare fluorescent secondary antibody (secondary antibody should be diluted in 1.0% BSA + 0.4% Triton X-100 + PBS) (use Biotin-SP for the antibody that will later add Streptavidin to; if there is a second primary then use the normal secondary for it).
  • Cover the tissue with the secondary antibody solution and incubate for 2 hrs. at RT in the incubation box. *From this point on, use low light and/or cover tissues.*
  • Rinse slides with PBS:  4 x 5 min
  • Remove excess PBS with a Kimwipe.
  • Prepare fluorescent Streptavidin secondary antibody (should be diluted in 1.0% BSA + 0.4% Triton X-100 + PBS)
  • Cover the tissue with the fluorescent Streptavidin antibody solution and incubate for 2 hrs. at RT in the incubation box.
  • Rinse slides with PBS:  4 x 5 min
  • Remove excess PBS with a Kimwipe.
  • Carefully add a drop of mounting medium to the center of the tissue and apply cover glass.
  • Seal cover glass with clear nail polish. For thicker tissue, add a weight before sealing.

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