Protocol for Immunogold Labelling of Tissue for TEM (adapted by Jeannie Mui)
1. Quenching and Washing
- Float each grid, sample side down, on three drops of 50 µL 0.2 M glycine for 3 minutes each to quench residual aldehydes.
- Float each grid, sample side down, on three drops of 100 µL PBS for 5 minutes each.
2. Blocking
- Float each grid on a drop of PBS containing 1% BSA for 5 minutes.
3. Primary Antibody Incubation
- Incubate the grid with 20 µL primary antibody diluted in PBS + 0.1% BSA.
- Incubate 1 hour at room temperature or overnight at 4°C in a humidified chamber.
4. Washing
- Wash the grid on five drops of DPBS (or PBS) for 5 minutes each.
5. Secondary Antibody Incubation
- Block again with 1% BSA in PBS for 5 minutes.
- Incubate with 20 µL gold‑conjugated secondary antibody (typically 1:20 in 0.1% BSA) for 1 hour at room temperature.
6. Final Washing
- Wash the grid on five drops of DPBS (or PBS) for 5 minutes each.
- Wash on five drops of ddH₂O for 2 minutes each.
7. Post‑Fixation (Optional to improve tissue integrity only if not previously fixed)
- Float the grid on a drop of 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer for 2 minutes.
- Wash with 5 drops of ddH₂O for 2 minutes each.
8. Post Staining
- Stain with 4% uranyl acetate for 5 minutes.
- Stain with Reynold's Lead for 5 minutes.
11. Imaging
- Image grids using a transmission electron microscope (TEM).
- Alternatively, store labelled grids in a labelled TEM grid box until ready for analysis.
Notes
- Perform all antibody incubations in a humidified environment to prevent grid drying.
- Use freshly prepared fixation and staining solutions for optimal labelling quality.
- Handle grids carefully to avoid contamination and mechanical damage.
Reagent Preparation Tables
General Buffers and Solutions
PBS (1× Phosphate‑Buffered Saline)
| Component | Final Concentration | Amount for 1 L |
|---|---|---|
| NaCl | 137 mM | 8.0 g |
| KCl | 2.7 mM | 0.2 g |
| Na₂HPO₄ | 10 mM | 1.44 g |
| KH₂PO₄ | 1.8 mM | 0.24 g |
| pH | 7.2–7.4 | Adjust with HCl/NaOH |
DPBS (Dulbecco’s PBS)
- Use premade DPBS without calcium and magnesium, or equivalent.
- Store at RT.
0.2 M Glycine Quench Solution
| Component | Amount |
|---|---|
| Glycine | 1.5 g |
| PBS | Bring to 100 mL |
Stable at RT for 1 day. Can be frozen at -20 °C.
Blocking Solution (1% BSA in PBS)
| Component | Amount |
|---|---|
| BSA (Fraction V) | 1 g |
| PBS | Bring to 100 mL |
Filter‑sterilize (0.22 µm) for best results.
Primary Antibody Dilution Buffer
| Component | Final % |
|---|---|
| PBS | — |
| BSA | 0.1% |
| Optional: Tween‑20 | 0.01% (reduces nonspecific binding; test first) |
Secondary Antibody Dilution (Gold‑Conjugated)
| Component | Amount |
|---|---|
| Secondary antibody | 1:20 (typical; follow manufacturer) |
| Dilution buffer | PBS + 0.1% BSA |
Prepare fresh; avoid vortexing (damages gold conjugates).
2.5% Glutaraldehyde in 0.1 M Sodium Cacodylate Buffer (Post-Fixation Optional)
| Component | Amount |
|---|---|
| 25% Glutaraldehyde | 10 mL |
| 0.1 M Cacodylate Buffer | 90 mL |
Prepare under a hood. Store aliquots at 4°C.
4% Uranyl Acetate (Post Stain)
| Component | Amount |
|---|---|
| Uranyl acetate | 0.4 g |
| ddH₂O (filtered) | 10 mL |
| pH | Should be ~4.3 |
Spin at 10,000×g for 1 min before use to remove particulates.
Store in amber tubes at 4°C (1–3 months).
Troubleshooting Guide
Below are the most common issues encountered during immunogold labelling of TEM sections, along with causes and solutions.
1. High Background Labelling
Possible Causes
- Insufficient blocking
- Antibody concentration is too high
- Inadequate washing between steps
- Sticky grids or contaminated reagents
- Gold secondary binding nonspecifically to Fc receptors or charged sites
Solutions
- Increase blocking time to 10–15 minutes or use 2% BSA.
- Add 0.05–0.1% Tween‑20 to washes (optional; test first).
- Increase the number and duration of washes (e.g., 7–10 drops, 5 min each).
- Use fresh blocking solution and filtered buffers.
- Reduce primary antibody concentration (e.g., 1:200 → 1:500).
- Try fish-skin gelatin (0.2–0.5%) as an alternative blocker for sticky samples.
- Ensure grids are handled only with clean forceps.
2. Weak or No Gold Labelling
Possible Causes
- The primary antibody is too dilute or has expired
- Poor antigen preservation (fixation too strong or too old)
- Inadequate permeabilization for tissue
- Secondary antibody not binding (wrong species pairing)
- Gold particles aggregated or degraded
Solutions
- Increase primary antibody concentration or incubation time (overnight at 4°C).
- Reduce fixative strength if possible (e.g., 2% glutaraldehyde instead of 2.5%).
- Validate the antibody on known‑positive control tissue.
- Ensure correct species pairing (e.g., mouse → anti‑mouse gold).
- Gently flick the tubes rather than vortexing to avoid destroying gold conjugates.
- Check gold conjugate by spotting a drop on a grid—intact particles appear monodisperse.
3. Patchy or Uneven Labelling
Possible Causes
- Grids drying during incubation
- Uneven spreading of antibody droplets
- Bad Parafilm surface or environmental contamination
- Variation in glow‑discharge quality
Solutions
- Maintain humid-chamber conditions throughout all antibody steps.
- Apply droplets gently and ensure even contact with the grid surface.
- Replace Parafilm frequently; avoid dust.
- Confirm glow discharge settings (carbon side up, 20 µA, 30 sec).
- Discard hydrophobic grids—they cause poor wetting.
4. Gold Particle Aggregation
Possible Causes
- The antibody solution is too salty
- Mechanical agitation (vortexing)
- Contaminants or particulate dust
Solutions
- Use PBS or DPBS, not Tris buffers (which can destabilize some gold conjugates).
- Mix by slow inversion only.
- Filter buffers (0.22 µm) before use.
- Spin gold solution at 10,000×g, 2–3 min to remove aggregates (carefully avoid pellet).
5. Poor Negative Staining / Low Contrast
Possible Causes
- Uranyl acetate is too old or contaminated
- Grid washed insufficiently (residual salts crystallize)
- Stain exposure is too short
Solutions
- Spin the uranyl acetate before use.
- Increase the number of ddH₂O washes to reduce salt crystals.
- Increase staining time to 1.5–2 minutes if needed.
- Ensure grids fully dry before imaging to prevent stain pooling.