Cox Systems Biology

University of Toronto

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Fluorescent Immunohistochemistry (IHC)

Author: Genlei Lu

Contents

  1. Materials
  2. Day One
  3. Day Two

Materials

materials printable
  • Xylenes
    • Ensure xylenes are handled under sufficient ventilation (fume hood).
  • 100% ethanol
  • 95% ethanol
    • If more 95% ethanol is needed, dilute by adding 41.2mL distilled water to unopened 4L jug of 100% ethanol
  • 70% EtOH
  • Distilled water
  • Antigen retrieval solution
    • Sodium citrate solution (IHCWorld): 
      • Add 2.94g trisodium citrate to 1000 mL distilled water (milliQ).
      • Adjust pH to 6.0 or 9.0 (application dependent) using test strips by adding very small amounts of HCl (if too acidic, recover with very tiny amount of NaOH).
      • Add 0.5mL Tween.
  • Casein blocking buffer
  • Tween 20
  • Unconjugated primary antibodies
  • Fluorophore-conjugated secondary antibodies
  • Humidity chamber
  • PBS (Stock or 10x)
  • TrueBlack Lipofuscin Autofluorescence Quencher (TLAQ)(Cell Signaling Technology #92401)
    • Optional depending on the autofluorescence characteristics of your tissue of interest.
  • DAPI
  • Antifade aqueous mounting media (Invitrogen P36965)
    • ProLong™ Diamond Antifade Mountant
  • Nail polish
    • Brighter colors are easier to see on glass.

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Day One

day one printable
  1. Label all slides w/ pencil.
    • For organoids: do not use slide warmer as it disrupts organoid integrity.
    • For tissues: slide warmer can be used but may cause sections to fall from slides more easily.
  2. Dewax slides by incubate slides in series of 2 buckets of xylenes for 10 minutes each, 2 buckets of 100% ethanol for 5 minutes each, and 2 buckets of 95% ethanol for 5 minutes each.
  3. Incubate slides in bucket of distilled water for 10 minutes
  4. Move slides to a bucket of appropriate antigen retrieval solution*, place bucket in steamer, and steam for 20 minutes to 1hr.
  5. After antigen retrieval steam, allow slides to cool in freezer, then transfer to a bucket of room temperature distilled water.
    • Monitor slides/bucket during cooldown for freezing/ice formation.
    • While slides cool, dilute antibodies in diluent of casein buffer.
      • Casein blocks nonspecific antibody binding.
      • Aim for 200mL of diluted antibody per slide/sample.
    • Place moistened paper towels inside of humidity chamber and flatten them within grooves to accommodate slides.
  6. Fill 3 buckets w/ 250mL 1x PBS (dilute from 10x stock). Add 1% Tween to the first bucket. Transfer slides in rack & wash for 5 minutes in each bucket (PBS-Tween → PBS → PBS)
    • If using TLAQ: detergents will wash away TLAQ and therefore are incompatible. All steps involving detergents should be completed prior to TLAQ application.
    • If using TLAQ: during washes,prepare 200mL of TLAQ working solution per slide/sample by diluting stock solution 1:20 in 70% EtOH.
  7. Working one slide at a time:
    1. Blot water off of slides at a site far from the sample.
    2. Draw a wide circle (at least 1cm gap between circle + organoids) around sample with hydrophobic pen.
      1. Do not press too hard with pen.
      2. Ensure each circle is complete.
    3. Allow pen to dry very briefly then transfer slide into humidity chamber.
    4. If using TLAQ:
      1. Starting with 1 slide: set timer to keep track of intervals between each slide; prepare 250mL of PBS for wash.
      2. Add 200mL of TLAQ to slide.
        1. If necessary, gently disperse antibody solution with pipette tip just contacting the solution itself, not touching the sample/slide!
        2. TLAQ is light sensitive. Slides should be kept from light sources as much as possible once TLAQ has been applied.
      3. Repeat for all slides while being mindful of the interval between each slide.
        1. Specific interval does not matter as long as they are consistent between slides.
        2. Keep the slides in a line in the order they TLAQ for ease of tracking.
      4. Following incubation of 3-5 minutes w/ TLAQ, transfer each slide into PBS per your time intervals and order.
        1. This ensures that each slide incubates with secondary antibody for a full 1 hour.
        2. Prepare 2 additional buckets of 250mL PBS.
      5. Wash for 5 minutes in each of the 3 PBS buckets.
      6. Repeat step 7.1 for each slide prior to step 7.5
    5. Apply primary antibodies:
      1. Add 200mL of diluted primary antibodies per slide/sample.
  8. Place lid on humidity chamber and carefully move the chamber to the +4oC fridge for overnight incubation of primary antibodies.

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Day Two

day two printable
  1. Prepare three 250mL wash buckets by adding 25mL of 10X PBS to each bucket, then topping each bucket up to the 250mL line with distilled water.
  2. Take slides out of the humidity chamber and check to make sure none of the slides became dry overnight.
    • If this occurred, make a note of which samples this happened to and consider removing from analysis.
  3. Move slides to a slide rack while holding the rack above the first wash bucket so that the liquid on the slides is caught by the bucket. Wash slide in this and subsequent buckets for 5 minutes each.
    • Dilute fluorophore-conjugated secondary antibodies in Casein blocking buffer during washes.
      • Ensure diluted antibodies are kept from light using containers/foil as much as possible.
    • Aim for 200mL of diluted antibody per slide/sample.
  4. Set a timer for 1 hour. Blot the first slide and apply 200mL ofdiluted fluorophore-conjugated secondary antibody onto it. Start the timer and apply secondary antibody to each subsequent slide in regular intervals (30 seconds, 1 minute, whatever works) keeping the slides in a line in the order they received secondary antibody.
    • Incubate at room temperature & in the dark (lid on humidity chamber is sufficient)
    • Disperse antibodies with pipette tip if necessary.
    • While secondary antibodies incubate, prepare all materials needed to finish slide prep:
      • Prepare 3 more 250mL 1x PBS washes.
      • Prepare DAPI working solution (1:1000 to 1:5000 dilution in PBS)
        1. Aim for 200mL of diluted DAPI per slide/sample.
  5. Transfer slides into rack and into the first wash bucket at the same time interval and order as you applied secondary antibody.
    • This ensures that each slide incubates with secondary antibody for a full 1 hour.
  6. When each slide has been placed in the rack in the first wash bucket, perform all 3 washes for 5 minutes each. 
  7. Reset timer. Apply 200mL of diluted DAPI w/ tracked-intervals similar to TLAQ/secondary antibodies. Incubate each slide w/ DAPI in the dark for 5 minutes.
  8. Transfer slides into rack and into the first wash bucket at the same time interval and order as you applied DAPI. Wash for 5 minutes each in the 3 buckets.
  9. Working one slide at a time, use kimwipe to remove as much residue PBS as possible without contacting the tissue. Using provided dropper, apply antifade aqueous mounting media to slides and apply slide cover.
    • Press down on slide cover to disperse mounting media on top of sample and to push any bubbles away.
    • Keep in the fume hood.
  10. Apply nail polish to seal gaps between slide cover & slide.
  11. Wait a few hours more until nail polish cures before imaging.
    • If sealed correctly, stained slides may be stored for up to 7 days in +4oC before imaging.

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Last updated: 06/2025