IF Protocol Hub

General IF Protocol - Standard Procedure for Immunofluorescence

Standard step-by-step immunofluorescence protocol ready for immediate use. Complete procedure from sample preparation to imaging with verification steps.

Created: 12/13/2025Updated: 12/13/2025By: IF Protocol Hub

General IF Protocol: Standard Procedure

This is the standard, ready-to-use immunofluorescence protocol that can be applied to most IF experiments. Follow each step carefully and verify results at each stage.

When to Use This Protocol

This general protocol works for most immunofluorescence applications. For specific sample types (cultured cells, tissue sections, frozen sections), see the specialized protocols linked below.

Pre-Experiment Preparation

Reagents & Materials

Essential Reagents

  • Fixative: 4% paraformaldehyde (PFA) in PBS, pH 7.4
  • Permeabilization: 0.2% Triton X-100 in PBS
  • Blocking: 3-5% BSA in PBS (or 5-10% normal serum)
  • Primary antibody: Specific to your target antigen
  • Secondary antibody: Fluorophore-conjugated, species-matched
  • Nuclear stain: DAPI (1 μg/ml) or Hoechst 33342
  • Mounting medium: Antifade mounting medium (ProLong Gold, Vectashield)

Essential Equipment

  • Microscope slides and coverslips (22 × 22 mm)
  • Humidified chamber (or petri dish with wet paper towels)
  • Fine forceps
  • Fluorescence microscope with appropriate filter sets
  • Pipettes and tips
  • Parafilm or aluminum foil (for light protection)

Before You Start

  • Ensure all reagents are at room temperature (unless specified otherwise)
  • Prepare fresh fixative solution (PFA degrades over time)
  • Verify antibody compatibility and recommended dilutions
  • Have all controls ready (negative control, isotype control)

Step-by-Step Procedure

1

Sample Preparation

For Cultured Cells:

  1. Grow cells on sterile glass coverslips to 50-70% confluence
  2. Coat coverslips with poly-L-lysine for better attachment (optional)
  3. Apply experimental treatments as required
  4. Ensure cells are in optimal condition before fixation

For Tissue Sections:

  1. Prepare 4-10 μm thick sections from frozen or paraffin-embedded tissue
  2. Mount on charged or poly-L-lysine-coated slides
  3. For paraffin sections: deparaffinize and rehydrate first
2

Fixation

Procedure:

  1. Remove culture medium and gently rinse once with PBS
  2. Add 4% PFA in PBS (pre-warmed to room temperature)
  3. Incubate for 15-20 minutes at room temperature
    • For tissue sections: 20-30 minutes
  4. Work in a fume hood when handling PFA

Alternative Fixatives

  • Methanol: Ice-cold 100% methanol, 5-10 min at -20°C (no permeabilization needed)
  • Acetone: Ice-cold 100% acetone, 5 min at -20°C (for certain antigens)
3

Washing

  1. Wash samples 3 times with PBS, 5 minutes each wash
  2. Use gentle agitation or rocking motion
  3. Ensure complete removal of fixative to prevent background staining

Verification

After washing, samples should be clear of fixative. No visible residue should remain.

4

Permeabilization

Skip this step if using methanol or acetone fixation.

Procedure:

  1. Prepare 0.2% Triton X-100 in PBS (standard concentration)
  2. Incubate samples for 10-15 minutes at room temperature
  3. For nuclear antigens: use 0.5% Triton X-100 for 15 minutes
  4. For membrane proteins: use 0.1% Triton X-100 for 5 minutes or skip

Permeabilization Alternatives

  • Saponin (0.1-0.5%): Reversible, good for membrane proteins
  • Digitonin (10-50 μg/ml): Selective for plasma membrane
  • Tween-20 (0.1-0.5%): Milder than Triton X-100
5

Blocking

Purpose: Reduce non-specific antibody binding

Procedure:

  1. Prepare blocking solution: 3-5% BSA in PBS (or 5-10% normal serum)
  2. Use serum from the same species as your secondary antibody
  3. Incubate samples for 30-60 minutes at room temperature
  4. Do not wash after blocking - proceed directly to primary antibody

Verification

Blocking solution should completely cover the sample. No bubbles should be present.

6

Primary Antibody Incubation

Antibody Preparation:

  1. Dilute primary antibody in blocking solution
  2. Typical dilutions: 1:50 to 1:500 (check manufacturer's recommendations)
  3. Perform antibody titration to determine optimal concentration

Incubation:

  1. Apply 100-200 μl of diluted antibody to each sample
  2. Incubate in a humidified chamber to prevent drying
  3. Option 1: 1-2 hours at room temperature
  4. Option 2: Overnight at 4°C (often provides better signal)
  5. Protect from light during incubation

Controls Required

  • Negative control: No primary antibody (only blocking solution)
  • Isotype control: Isotype-matched control antibody
  • Positive control: Known positive sample (if available)
7

Washing After Primary Antibody

  1. Wash 3 times with PBS, 5 minutes each wash
  2. For high background: add 0.1% Tween-20 to PBS
  3. Ensure complete removal of unbound antibody

Verification

Wash buffer should be clear. No visible antibody solution should remain.

8

Secondary Antibody Incubation

Antibody Preparation:

  1. Dilute secondary antibody in blocking solution
  2. Typical dilutions: 1:200 to 1:1000
  3. Use species-matched secondary antibody
  4. Protect from light (fluorophores are light-sensitive)

Incubation:

  1. Apply 100-200 μl of diluted secondary antibody
  2. Incubate for 1 hour at room temperature in the dark
  3. Or incubate for 30 minutes if using high-quality antibodies

Critical

  • Always protect from light during secondary antibody incubation
  • Use aluminum foil or work in a dark room
  • Light exposure will quench fluorescence
9

Final Washing

  1. Wash 3 times with PBS, 5 minutes each wash
  2. For high background: add 0.1% Tween-20 to PBS
  3. Final wash should be thorough to remove all unbound secondary antibody
10

Nuclear Counterstaining (Optional)

Procedure:

  1. Prepare DAPI (1 μg/ml) or Hoechst 33342 in PBS
  2. Incubate for 5-10 minutes at room temperature
  3. Wash once with PBS (5 minutes)
  4. This step helps visualize cell nuclei and assess cell morphology
11

Mounting

Procedure:

  1. Place a drop of antifade mounting medium on a clean slide
  2. Invert coverslip (sample side down) onto mounting medium
  3. Gently press to remove air bubbles
  4. Seal edges with nail polish (optional, for long-term storage)
  5. Store slides at 4°C in the dark until imaging

Mounting Tips

  • Use enough mounting medium to prevent air bubbles
  • Avoid pressing too hard (can damage cells)
  • Let mounting medium cure for 30 minutes before imaging
12

Imaging

Procedure:

  1. Allow mounted slides to cure for 30 minutes at room temperature
  2. Image using fluorescence microscope with appropriate filter sets
  3. Use appropriate exposure times (avoid overexposure)
  4. Capture images immediately or store slides at 4°C in the dark

Expected Results

  • Positive signal: Bright, specific fluorescence at target location
  • Negative control: Minimal to no fluorescence
  • Nuclear stain: Clear, distinct nuclei
  • Background: Low, uniform background fluorescence

Verification Checklist

Before considering your experiment complete, verify:

  • Negative control shows minimal fluorescence
  • Positive signal is specific and localized
  • Background is low and uniform
  • Nuclear counterstain (if used) is clear
  • No air bubbles in mounting medium
  • Images are properly exposed (not overexposed)

For specific applications, see: