Western blotting turns a messy protein mix into a readable picture. When it works, you see clear bands at the expected molecular weights; when it doesn’t, you’re staring at foggy backgrounds or missing bands and wondering what went wrong. This guide gives you a complete, practical western blot protocol, cover sample prep, gels, transfer, blocking, antibodies, detection (ECL, fluorescent, alkaline phosphatase), stripping and reprobing, and the common causes of variability in western blot protocols.
What You’ll Need (Western Blotting Protocol Overview)
Core reagents and plastics
- Lysis buffer (RIPA or mild NP-40/Triton; add fresh protease/phosphatase inhibitors)
- Sample buffer (Laemmli with SDS; reducing agent such as DTT or β-ME)
- Precast or cast polyacrylamide gels (SDS-PAGE), 7.5–15% depending on target size
- Running buffer (Tris-Glycine-SDS)
- Transfer buffer (Tris-Glycine with methanol; add SDS only when needed)
- PVDF or nitrocellulose membrane (PVDF requires methanol activation)
- Blocking reagent (5% milk in TBST, or 3–5% BSA for phospho targets; fish gelatin for fluorescence)
- Primary antibody (validated for WB)
- Secondary antibody (HRP-conjugated or fluorescent dye-conjugated)
- Wash buffer (TBST 0.05–0.1% Tween-20)
- Chemiluminescent substrate (ECL western blot protocol) or fluorescent imaging setup
- Protein ladder (molecular weight markers)
Equipment
- Microcentrifuge, mini-centrifuge for spin-downs
- Vortex, heat block/thermocycler at 95 °C
- Gel tank and power supply
- Wet transfer tank or semi-dry transfer unit
- Rocker for antibody incubations
- Imaging system (chemiluminescence imager or LI-COR/Odyssey for fluorescent western blot protocol)
Western Blot Sample Preparation Protocol
Getting the sample right decides the rest of your day. Poor extraction or overloaded lanes are the top reasons blots look bad.
Protein extraction protocol for western blot
- Keep everything cold. Chill tubes, buffers, and a microcentrifuge.
- Add lysis buffer (e.g., RIPA with inhibitors) to your cells or tissue (10× volume for cell pellets; 10–20× for tissues).
- Disrupt thoroughly: pipette up/down, brief sonication, or mechanical homogenization (especially for tough tissues).
- Incubate on ice 15–30 min with mixing.
- Spin at 12–16k × g for 10–15 min at 4 °C; collect the supernatant.
- Quantify protein (BCA recommended; compatible with many detergents).
Western blot cell lysis protocol (contextual notes)
- Adipose tissue western blot protocol: supplement with extra detergents and nuclease; expect lipids—clarify supernatant carefully.
- Serum western blot protocol: deplete high-abundance proteins if targeting low-abundance analytes.
- Histone western blot protocol: use acid extraction or nuclear lysis buffers.
- Exosome western blot protocol: isolate exosomes (ultracentrifugation/kit), then lyse gently to preserve membrane proteins.
Sample preparation
- Mix lysate with 4× sample buffer (final 1×), add reducing agent.
- Heat at 95 °C for 5 min (some membrane proteins prefer 65–70 °C to preserve epitopes).
- Brief spin; hold on ice until loading.
Controls and normalization
- Choose a loading amount (10–30 µg per lane for most cell lysates).
- Prefer total protein normalization (Ponceau S or stain-free) or stable housekeeping proteins (GAPDH western blot protocol, beta actin western blot protocol). Validate that the control is constant across conditions.
Gel Electrophoresis
Picking gel percentage
- 7.5–10%: high MW proteins
- 10–12%: mid-range proteins (25–100 kDa)
- 12–15%: small proteins/peptides
Running conditions
- Pre-run gels for a few minutes to warm and flush wells.
- Load ladder and samples carefully—avoid bubbles.
- Run at ~120 V until the dye front reaches the bottom (mini gels ~45–70 min).
Common gel issues (and fixes)
- Smiling bands: buffer getting hot → lower voltage or run in the cold room.
- Curved lanes: uneven polymerization or crooked wells → use fresh gels/align combs.
Protein Transfer: Wet vs Semi-Dry
Western blot wet transfer protocol
- Membrane choice: PVDF (activate in methanol 15–30 s, rinse in water, equilibrate in transfer buffer) or nitrocellulose (no activation).
- Build the sandwich: sponge → filter → gel → membrane → filter → sponge; remove all bubbles with a roller.
- Transfer at 100 V for 60–90 min (cold transfer buffer, stir bar, and ice pack help maintain 4–10 °C).
- For very large proteins, extend time or reduce current to prevent blow-through.
Semi dry western blot protocol
- Use pre-soaked filter papers and the manufacturer’s buffer system.
- Transfer at constant current (e.g., 0.8 mA/cm²) for 10–45 min depending on MW.
- Semi-dry is fast and convenient, but wet transfer often handles high MW proteins better.
PVDF vs nitrocellulose
- PVDF: higher binding capacity, robust for stripping; must be methanol-activated; can increase background if blocking is poor.
- Nitrocellulose: low background, easy handling; can be brittle and less amenable to harsh stripping.
Blocking and Antibody Incubations
Blocking
- Standard: 5% nonfat dry milk in TBST for most targets (not ideal for phospho-proteins due to casein).
- For phospho-epitopes: 3–5% BSA in TBST.
- For fluorescent western blot protocol: use milk-free blockers compatible with NIR dyes (LI-COR/Odyssey recommends specialized buffers).
- Incubate 30–60 min at room temp with gentle rocking.
Primary antibody
- Dilute in blocking buffer or TBST with 1–5% blocker.
- Typical dilution 1:500 to 1:2000 (follow datasheet; titrate for your samples).
- Incubate 1–2 h at room temp or overnight at 4 °C.
Washes
- Three washes in TBST, 5–10 min each, gentle rocking. Insufficient washing is the most common cause of background.
Secondary antibody western blot protocol
- HRP-conjugated (for ECL): 1:5000–1:20,000 in blocking buffer.
- Fluorescent secondary (700/800 nm): 1:10,000 (check supplier guidance).
- Incubate 45–60 min at room temp; protect fluorescent blots from light.
- Wash 3–4× in TBST.
Detection Methods (Pick One, Do It Well)
ECL western blot protocol (HRP/chemiluminescence)
- Equilibrate membrane in TBS (no Tween) for better signal consistency.
- Mix equal parts substrate; cover the membrane (0.1 mL/cm² approx).
- Incubate 1–3 min; drain excess.
- Image immediately; capture a series of exposures (5 s to 5 min) to avoid saturation.
Pros: Sensitive, inexpensive, widely supported.
Cons: Narrow dynamic range; film is hard to quantify—use a digital imager if possible.
Fluorescent western blot protocol
- Use NIR secondaries (e.g., 700 and 800 nm) on LI-COR/Odyssey or equivalent.
- Block with dye-compatible buffers; avoid milk if it increases background.
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Multiplex two targets on the same membrane (saves time and samples).
Pros: Very wide dynamic range, true multiplexing, linear quantification.
Cons: Requires a fluorescence imager and careful buffer selection.
Alkaline phosphatase western blot protocol
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AP-conjugated secondaries with chemifluorescent or colorimetric substrates.
Pros: Long signal stability (great for long exposures).
Cons: Slower signal development than ECL; less common in many labs.
Biotin–Streptavidin / Streptavidin-HRP western blot protocol
- Useful when signal amplification is needed or primary antibodies are biotinylated.
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Sequence: primary (biotinylated) → streptavidin-HRP → substrate.
Pros: Extra sensitivity.
Cons: Endogenous biotin can cause background; block appropriately.
Stripping and Reprobing (When One Blot Must Do More)
Western blot stripping buffer protocol
- Mild stripping (e.g., glycine pH ~2.2, brief SDS at RT) for sensitive epitopes.
- Harsh stripping (e.g., SDS + β-ME at 50–56 °C) for stubborn antibodies.
- Rinse thoroughly; re-block; re-probe.
Western blot membrane stripping protocol—cautions
- Phospho-proteins and some membrane proteins lose signal after harsh stripping.
- If the target is precious, consider running duplicate gels instead of aggressive stripping.
Stripping protocol western blot: when to use
- To probe the same membrane for a different isoform or control.
- To replace a suboptimal antibody with an improved one.
Causes of Variability in Western Blot Protocols (and How to Control Them)
Sample variability
- Different lysis buffers change what you extract. Use the same buffer for comparisons.
- Protease/phosphatase activity rises quickly at room temp—keep samples cold.
Loading and gel factors
- Unequal loading or transfer produces fake “expression changes.”
- Choose gel % that resolves your protein; wrong percentage smears bands.
Transfer efficiency
- Over-transfer (“blow-through”) leads to faint bands; under-transfer yields heavy gel retention.
- Add 0–0.05% SDS to transfer buffer for high MW proteins; reduce methanol to improve pore size. Do the opposite for small proteins.
Blocking and antibody conditions
- Milk vs BSA matters: casein in milk can block phospho epitopes.
- Too concentrated antibodies = background; too dilute = missing bands. Titrate.
Wash stringency
- Under-washing leaves haze; over-washing can reduce weak signals. Standardize TBST concentration and wash times.
Detection chemistry
- ECL substrates vary in sensitivity and stability; fluorescent systems are more linear but require clean buffers and proper blocking.
Documentation
- Keep a simple run sheet (gel %, transfer settings, antibody lots/dilutions, exposure times). Reproducibility improves dramatically when you can copy conditions.
Brand Protocols and PDFs (Use as Reference Entities)
Many labs start with a vendor’s recommended conditions, then tune them. If you need product-specific instructions or troubleshooting trees, check these entities:
- Bio-Rad western blotting protocol
- Thermo Fisher western blot protocol
- Invitrogen western blot protocol
- Abcam western blot protocol PDF
- Cell Signaling Technology western blot protocol
- Sigma western blot protocol
- LI-COR western blot protocol / Odyssey western blot protocol
Western Blot Protocol Step by Step (Compact Checklist)
- Prepare lysates on ice; quantify protein (BCA).
- Mix with 4× sample buffer + reducer; heat 95 °C 5 min.
- Load 10–30 µg per lane; run SDS-PAGE at ~120 V to completion.
- Activate PVDF (methanol), assemble sandwich; transfer (wet: 100 V, 60–90 min; semi-dry: per unit’s specs).
- Block (5% milk in TBST, or 3–5% BSA for phospho) 30–60 min.
- Incubate primary (optimized dilution) 1–2 h RT or overnight 4 °C.
- Wash TBST 3×, 5–10 min each.
- Incubate secondary (HRP or fluorescent) 45–60 min RT.
- Wash TBST 3–4×; detect via ECL or fluorescence; capture multiple exposures.
- Normalize (housekeeping or total protein), quantify bands, and report with ladder and replicates.
- Optional: strip and reprobe if needed, with caution.
Frequently Asked Questions (FAQ)
1) What is the western blot protocol in simple terms?
Lyse cells → run proteins on a gel → transfer to a membrane → block → add primary antibody → add secondary antibody → detect signal (chemiluminescence or fluorescence).
2) Which membrane is better, PVDF or nitrocellulose?
PVDF binds more protein and tolerates stripping; nitrocellulose tends to give lower background. Pick based on target and whether you plan to reprobe.
3) How much protein should I load?
Start with 10–30 µg per lane for cell lysates. Optimize per target; too much causes fat bands and background.
4) What blocking buffer should I use?
Use 5% milk for most targets. Use BSA for phospho-proteins. For fluorescent detection, use milk-free, dye-compatible blockers.
5) Why is my background so high?
Likely causes: too much antibody, insufficient washes, cross-reactive blocking buffer, or contaminated buffers. Titrate antibody, increase wash time, switch blocker, and make fresh buffers.
6) My high-molecular-weight protein is weak. What can I change?
Extend wet transfer time, reduce methanol (to open gel pores), add a trace of SDS (0–0.05%) to transfer buffer, and keep the stack cold.
7) Can I strip and reprobe the same blot?
Yes—use a mild western blot membrane stripping protocol for sensitive epitopes. Harsh stripping can damage some targets, especially phospho-sites.
8) When should I use fluorescent western blot protocol instead of ECL?
Choose fluorescence if you need accurate quantitation or multiplexing (two targets on one blot). Choose ECL for everyday sensitivity with simple equipment.
9) Do housekeeping proteins always work as loading controls?
Not always. GAPDH or β-actin can change under some treatments. Consider total protein normalization (Ponceau S/stain-free) for robust comparisons.
