Western blotting (immunoblotting) is one of the most widely used techniques in molecular biology and biochemistry. It detects specific proteins in a sample using antibodies. Understanding it step-by-step — not just memorising it — is what separates good students from great ones.
What is Western Blotting?
- Definition: A technique used to detect specific proteins in a complex sample by separating them by size, transferring to a membrane, and probing with antibodies
- Named after: W. Neal Burnette (1981), who coined "Western blot" as a play on the "Southern blot" (DNA) and "Northern blot" (RNA) techniques
- Based on: SDS-PAGE (gel electrophoresis) for separation + antibody-antigen interaction for detection
- Used for: HIV diagnosis (confirmatory test), protein expression studies, COVID-19 serology, cancer biomarker research, food safety testing
- Key advantage: Can detect a single specific protein out of thousands in a complex mixture
- Time required: Typically 2–3 days for a complete run from sample to result
A western blot result showing protein bands detected by antibodies — darker bands indicate higher protein amounts. Image: Wikimedia Commons
The Underlying Principle
- Step 1 — Separation: Proteins are separated by molecular weight using SDS-PAGE. SDS (sodium dodecyl sulphate) denatures proteins and gives them a uniform negative charge, so they migrate through the gel based on size alone — smaller proteins move faster
- Step 2 — Transfer: Separated proteins are transferred from the gel to a solid membrane (PVDF or nitrocellulose) using an electric current — this is the "blotting" step
- Step 3 — Blocking: Non-specific binding sites on the membrane are blocked (usually with milk or BSA) to prevent antibodies from sticking non-specifically
- Step 4 — Antibody probing: A primary antibody specific to the target protein is applied, binds to its antigen, then a secondary antibody (carrying a detection label) binds to the primary antibody
- Step 5 — Detection: The label on the secondary antibody (enzyme or fluorophore) produces a signal that is visualised as a band at the molecular weight of the target protein
Key Concept
SDS (Sodium Dodecyl Sulphate) is a detergent that binds to proteins at a constant ratio (~1.4g SDS per gram of protein). This coats all proteins with negative charge proportional to their mass, ensuring migration through the gel is based on size — not on the protein's natural charge or shape.
Step-by-Step Procedure
🔬 STAGE 1: Sample Preparation
- 1Lyse cells using appropriate lysis buffer (RIPA buffer for total protein extraction) on ice
- 2Centrifuge at 12,000–14,000 × g for 10–15 minutes at 4°C to remove cell debris
- 3Collect the supernatant (protein lysate)
- 4Quantify protein concentration using Bradford assay, BCA assay, or Lowry method
- 5Mix equal amounts of protein (typically 20–50 µg per lane) with Laemmli sample buffer (contains SDS, β-mercaptoethanol, glycerol, bromophenol blue)
- 6Boil at 95–100°C for 5 minutes to fully denature proteins and reduce disulfide bonds
- 7Cool on ice and centrifuge briefly before loading
💡 Tip: β-mercaptoethanol (BME) reduces disulfide bonds between cysteine residues, ensuring proteins are fully linear. This is essential for accurate size separation.
⚡ STAGE 2: SDS-PAGE (Gel Electrophoresis)
- 1Cast the gel: pour resolving gel first (8–15% acrylamide depending on target protein size — higher % for smaller proteins)
- 2Allow resolving gel to polymerise (~30 min), then pour stacking gel (4–5% acrylamide) on top with comb inserted
- 3Once stacking gel is set, remove comb. Place gel in electrophoresis tank with Tris-glycine running buffer
- 4Load molecular weight marker (ladder) in first lane — this gives size reference for your bands
- 5Load samples carefully into wells using a gel loading tip
- 6Run at 80V through stacking gel, then 120V through resolving gel until dye front reaches the bottom
- 7Total run time: approximately 1–2 hours
⚠️ Important: The stacking gel concentrates all proteins into a sharp starting band regardless of volume differences. The resolving gel is where actual size separation occurs. Never skip the two-gel system.
SDS-PAGE gel showing protein bands separated by molecular weight. Image: Wikimedia Commons
📋 STAGE 3: Transfer (Blotting)
- 1Activate PVDF membrane by soaking in methanol for 15 seconds, then equilibrate in transfer buffer (nitrocellulose membrane does not need methanol activation)
- 2Build the transfer "sandwich" in this exact order (anode to cathode): + Anode → Fibre pad → Filter paper → Membrane → Gel → Filter paper → Fibre pad → Cathode −
- 3Remove all air bubbles by rolling a glass rod over each layer — bubbles prevent transfer
- 4Transfer at 100V for 60–90 minutes (wet transfer) or 25V overnight (semi-dry transfer options vary by system)
- 5Confirm transfer success: reversibly stain membrane with Ponceau S (red staining of bands) or stain gel to check protein depletion
Why PVDF or Nitrocellulose?
These membranes bind proteins non-covalently through hydrophobic interactions. PVDF has higher protein binding capacity and mechanical strength. Nitrocellulose is more fragile but gives lower background. Choice depends on protein of interest and detection method.
🛡️ STAGE 4: Blocking
- 1Rinse membrane in TBST (Tris-buffered saline + 0.1% Tween-20) for 5 minutes
- 2Incubate in 5% non-fat dry milk in TBST OR 5% BSA in TBST for 1 hour at room temperature with gentle rocking
- 3">Use milk for most antibodies; use BSA if probing for phosphorylated proteins (milk contains casein which is phosphorylated and interferes)
- 4Blocking saturates non-specific protein-binding sites on the membrane — preventing antibodies from sticking everywhere and creating high background
🔎 STAGE 5: Antibody Incubation
- 1Dilute primary antibody in blocking buffer (e.g., 1:1000) — consult datasheet for optimal dilution
- 2Incubate membrane with primary antibody: 2 hours at room temperature OR overnight at 4°C (preferred — gives stronger, more specific signal)
- 3Wash membrane 3× with TBST, 10 minutes each — removes unbound primary antibody
- 4Dilute secondary antibody in blocking buffer (e.g., 1:5000–1:10000). Secondary must match the species of the primary antibody (e.g., if primary is rabbit anti-human, use anti-rabbit secondary)
- 5Incubate with secondary antibody for 1 hour at room temperature
- 6Wash membrane 3× with TBST, 10 minutes each
💡 Key Concept: The primary antibody recognises your target protein (antigen-specific). The secondary antibody recognises the primary antibody and carries the detection label (usually HRP enzyme or fluorophore). This indirect detection amplifies the signal significantly.
✨ STAGE 6: Detection & Visualisation
- 1ECL (Enhanced Chemiluminescence): Most common. HRP enzyme on secondary antibody reacts with ECL substrate to produce light. Expose membrane to X-ray film or use a digital imager
- 2Apply ECL reagent evenly to membrane surface, incubate 1–5 minutes
- 3In darkroom: place membrane on X-ray film, expose for 30 seconds to 5 minutes, develop film
- 4">Bands appear at the molecular weight of your target protein — compare position to the ladder to confirm size
- 5Densitometry: Use ImageJ or similar software to quantify band intensity for relative protein expression analysis
- 6Loading control: Always probe for a housekeeping protein (β-actin, GAPDH, tubulin) to confirm equal loading across lanes
Troubleshooting Common Problems
- No bands visible: Primary or secondary antibody concentration too low; protein not transferred; wrong antibody species; protein degraded — check antibody datasheets and include positive control
- High background (bands everywhere): Blocking insufficient; antibody concentration too high; washing steps too short — increase blocking time, reduce antibody concentration, wash longer
- Band at wrong molecular weight: Post-translational modifications (glycosylation, phosphorylation) alter apparent MW; protein may run differently under denaturing conditions — always verify with positive control
- Smeared bands: Too much protein loaded; gel ran too fast; sample not fully denatured — reduce protein load, slow down run voltage
- Bands only in some lanes: Uneven loading; pipetting error; bubbles during transfer — recheck quantification and transfer sandwich
- White bands on dark background (overexposed): Too much antibody or too much ECL — reduce antibody concentration or exposure time
🎓 Exam Practice Questions
- What is the role of SDS in Western blotting? Why is it added?
- Why is β-mercaptoethanol added to the sample buffer?
- What is the purpose of the stacking gel in SDS-PAGE?
- Why must the transfer sandwich be assembled in a specific order?
- Why should BSA be used instead of milk when probing for phosphoproteins?
- What is the role of the secondary antibody? What is the advantage of using it over a directly labelled primary?
- What is a loading control and why is it essential?
- Differentiate between Western blot, Northern blot, and Southern blot.