Western Blot Protocol: A Complete Guide for Antibody-Based Detection

The western blot protocol is the workhorse method of antibody-based protein detection. Since Towbin, Staehelin, and Gordon's 1979 description of protein transfer from SDS-PAGE to nitrocellulose and Burnette's 1981 nomenclature ("Western blotting" as a play on Southern blots for DNA), the technique has remained the standard laboratory method for confirming target protein identity, quantifying abundance changes, and detecting post-translational processing. Well-run, a western blot resolves proteins by molecular weight, discriminates target from non-target signal, and provides semi-quantitative abundance comparisons between conditions.

This western blot antibody protocol covers the full workflow — sample preparation, gel electrophoresis, transfer, blocking, primary and secondary antibody incubation, detection, quantification, and troubleshooting — with attention to the proteinase-specific considerations that matter when the target is an MMP, cathepsin, ADAM, granzyme, or other proteinase where zymogen and mature forms coexist. Family-specific western blot protocols for the four major mechanistic classes are linked in Pillar Resources at the end.

Overview of the western blot workflow

A western blot has six primary stages, each with decisions that affect the final result:

1. Sample preparation — cell or tissue lysis in an appropriate buffer, protein quantification, denaturation with SDS and reductant, heat denaturation.
2. Gel electrophoresis — separation by molecular weight in a polyacrylamide gel. Gel percentage matched to target size.
3. Transfer — proteins moved from gel to membrane (PVDF or nitrocellulose) by wet, semi-dry, or dry transfer.
4. Blocking — non-target sites on the membrane blocked with protein solution (BSA or milk) to reduce background.
5. Antibody incubation — primary antibody binds target, secondary antibody (conjugated to HRP or fluorophore) binds primary.
6. Detection and quantification — chemiluminescent or fluorescent signal imaged and band intensity measured.

Each stage has its own optimisation and its own failure modes. The following sections walk through them in order, with proteinase-specific notes at each stage.

Sample preparation

Sample preparation determines what you can detect. A protease inhibitor cocktail is mandatory when the sample contains proteases (which for proteinase research is by definition), otherwise your target and other proteins in the lysate are degraded before they reach the gel. Sample buffer composition, reduction, and denaturation choice all affect what forms of the target are visible.

Lysis buffer selection

Common choices: RIPA (radioimmunoprecipitation assay, contains SDS + deoxycholate + Triton — most stringent, extracts membrane and nuclear proteins but may denature antibody epitopes); Triton X-100 buffer (milder, for cytosolic and lightly membrane-associated proteins); NP-40 buffer (very mild, preserves native complexes); Laemmli sample buffer directly (harshest — extracts nearly everything but no further processing possible). For proteinase research: RIPA works well for total protein detection; milder buffers are appropriate if downstream IP or activity assays are planned.

Protease inhibitors — mandatory for proteinase research

Include a commercial protease inhibitor cocktail (e.g., Roche cOmplete, Thermo Halt) at manufacturer-recommended concentration. Add EDTA (5-10 mM) to inhibit metalloproteinases specifically. For calpain research, add EGTA (chelates calcium). For serine protease research, PMSF (1 mM, added fresh) or AEBSF is standard. For cysteine protease research, E-64 is standard. Keep samples cold throughout.

Protein quantification

BCA or Bradford assay against a BSA standard curve. Load equal micrograms of protein per lane (typically 10-40 μg for whole-cell lysates). Loading too little produces weak signal that fails to distinguish specific from background; loading too much produces smearing and poor resolution.

Denaturation and reduction

Add SDS-containing sample buffer (typically 4× Laemmli), heat to 95°C for 5 minutes with a reducing agent (β-mercaptoethanol or DTT). Reduction breaks disulphide bonds. For proteinases with disulphide-dependent structure, some antibodies raised against native folds will not detect the reduced form — verify your antibody's compatibility with reducing conditions before assuming a negative result is target-absent.

Gel electrophoresis

SDS-PAGE separates proteins by molecular weight. Gel percentage should be matched to the target: 8% for proteins 60-200 kDa (most MMPs, ADAMs), 10% for 30-80 kDa (many cathepsins, TIMPs, granzymes), 12% for 15-40 kDa (small mature-form fragments, cleaved caspases), 15% for smaller than 25 kDa. Gradient gels (4-12% or 4-20%) are practical when the target and its cleaved products span a wide size range — common for proteinases where the zymogen, mature form, and small cleavage products all need to be visualised on the same blot.

Molecular weight markers

Include a pre-stained MW ladder spanning the expected band size. Two lanes of ladder (leftmost and rightmost of the gel) is standard practice to allow accurate size interpolation across the entire blot.

Loading and running

Load 10-40 μg total protein per lane. Include a positive control lane (known target-positive lysate or recombinant target) and, for antibody validation blots, a negative control lane (target-negative lysate or knockout material). Run at 100-150 V until the dye front reaches the bottom (~1 hour for standard mini-gels, longer for gradient or larger-format gels).

Transfer

Transfer moves proteins from the gel to a membrane where they become accessible to antibody. Three methods dominate:

After transfer, briefly stain the membrane with Ponceau S (1-2 minutes, then rinse) to confirm even transfer across the gel. Ponceau-stained proteins are visible, distinct from lane to lane, and the ladder should be sharp. If the transfer is uneven, blocking will not fix it — start over.

Blocking

Blocking saturates non-specific protein-binding sites on the membrane so that primary antibody only binds its target. Standard blocking solutions: 5% non-fat dry milk in TBS-T (Tris-buffered saline + 0.1% Tween-20), or 5% BSA in TBS-T. Milk is cheaper and typically more effective for most targets. BSA is required for phospho-specific antibodies (milk contains casein phosphoproteins that generate background) and for streptavidin-based detection (milk contains endogenous biotin).

Block for 1 hour at room temperature or overnight at 4°C. Under-blocking produces high background; over-blocking is not a problem in practice. See the western blot blocking buffer protocol for the detailed recipe and buffer variations.

Primary antibody incubation

The primary antibody is diluted in blocking buffer (or in TBS-T alone for some fluorescent detection workflows) at a working concentration determined by titration. Standard dilutions range from 1:500 to 1:10,000 depending on antibody titre — for TPB rabbit polyclonals, 1:1000 is a typical starting point.

Incubation time and temperature

Two standard protocols: 1 hour at room temperature (fast, higher background), or overnight at 4°C (better signal-to-noise, standard for most publications). For faint targets, overnight at 4°C is essentially always the correct choice. See the primary antibody incubation time guide for detailed workflow options.

Titration is mandatory for every new antibody

Manufacturer-recommended dilutions are a starting point. Every new antibody must be titrated in the researcher's own sample type across at least 4-5 dilutions (e.g., 1:500, 1:1000, 1:2500, 1:5000, 1:10000). The optimal dilution is the one giving the highest signal-to-background ratio — not necessarily the highest absolute signal. See the dilution optimization protocol and the calculate-antibody-dilution protocol.

Washing and secondary antibody

After primary antibody, wash the membrane 3-5 times, 5 minutes each, in TBS-T. Insufficient washing is the most common source of high background — always err on the side of more and longer washes.

Secondary antibody: an anti-species antibody (e.g., goat anti-rabbit for a rabbit primary) conjugated to horseradish peroxidase (HRP) for chemiluminescent detection or to a fluorophore (Cy3, Cy5, IRDye 680/800) for fluorescent detection. Dilute the secondary in blocking buffer at 1:5,000 to 1:20,000 depending on manufacturer recommendation. Incubate 1 hour at room temperature. Wash 3-5 times, 5 minutes each, in TBS-T after secondary. See the choose-secondary-antibody guide and the why-use-secondary-antibody guide.

Detection systems

Two dominant detection systems: chemiluminescence (ECL) and fluorescence.

Band interpretation for proteinases

Interpreting bands on a proteinase Western blot requires knowing what forms of the target exist and what molecular weights they run at. Unlike housekeeping proteins that produce a single band at a predictable size, proteinases typically produce multiple bands — the zymogen, the mature form after propeptide cleavage, further processed products, and sometimes complexes with endogenous inhibitors.

Example: MMP-9. proMMP-9 runs at ~92 kDa. Active MMP-9 (propeptide-cleaved) runs at ~82-84 kDa. Truncated catalytic domain runs smaller (~68 kDa). TIMP-1 complex runs higher (~120 kDa). A single lane of stimulated macrophage lysate may show all four bands. Distinguishing them requires knowing the expected sizes and, ideally, running a recombinant standard as a size marker. See the family-specific western blot protocols in Pillar Resources for the expected band patterns for each proteinase class.

Loading controls

A loading control is a stably expressed housekeeping protein (GAPDH, actin, tubulin, vinculin) detected on the same blot as the target. Loading controls confirm equal protein loading and provide the denominator for quantification. Choose a loading control whose molecular weight is well separated from the target — GAPDH (37 kDa) is a good choice for targets in the 60-100 kDa range but overlaps with cleaved caspase-3 or activated cathepsins. See the store-western-blot-membrane guide for reprobing strategies when the loading control needs to run on the same membrane.

Quantification

Quantitative Western blot requires linear signal capture (fluorescence preferred), non-saturated bands, and normalisation to a loading control. Use ImageJ, Image Studio (LI-COR), or Image Lab (BioRad) to measure integrated band density. Normalise target density to loading control density in the same lane. Compare normalised values across conditions.

For accurate quantification, ensure: (1) all bands are within the linear dynamic range of the detection system — no saturation, no baseline signal below detection limit; (2) the loading control does not itself change under the experimental conditions (verify with a second loading control if uncertain); (3) technical replicates from at least 3 independent experiments before drawing conclusions.

Troubleshooting overview

Four dominant western blot failure modes and their most common root causes:

Adapting the standard protocol to proteinase-specific considerations

The standard western blot workflow above works for most targets. Proteinases have three specific issues that require workflow adaptation.

Zymogen vs mature form resolution

Every proteinase has a zymogen form and one or more processed forms. Running on a gradient gel (4-12% or 4-20%) gives resolution across the size range of interest. Use antibodies that either detect all forms (mid-catalytic epitope, mature-form N-terminus not required) or use domain-specific antibodies in separate lanes or channels to distinguish them.

Sample handling to preserve activation state

If the experimental question is about activation state (what fraction of the target is zymogen vs mature), samples must be handled to prevent ex-vivo activation during lysis. Include EDTA in the lysis buffer for MMPs (prevents ex-vivo activation by contaminating metalloproteinases). Include broad-spectrum protease inhibitors for other classes. Snap-freeze samples immediately after lysis; do not incubate on ice for extended periods before running.

Multi-domain detection with superpooled antibodies

For proteinases where detecting all functional forms simultaneously matters more than paralogue selectivity (a common experimental need), superpooled antibodies (multi-domain polyclonal pools raised against the propeptide, catalytic domain, and C-terminal region separately) detect all forms in a single antibody with domain-resolved signal in a multi-channel readout. See the superpooled antibody usage protocol.

Family-specific western blot protocols

The general workflow above applies to any target. Family-specific protocols cover the details that differ between proteinase classes — appropriate gel percentages, expected band sizes, activation-state considerations, and paralogue cross-reactivity notes. Individual family protocols are linked in Pillar Resources at the end of this page.

Frequently Asked Questions

Explore related topics in this cluster

• Aspartic proteinase Western blot protocol— BACE, cathepsin D, renin
• Cysteine proteinase Western blot protocol— cathepsins, calpains, caspases
• Metalloproteinase Western blot protocol— MMPs, ADAMs, ADAMTSs
• Serine proteinase Western blot protocol— coagulation, complement, kallikreins
• Western blot blocking buffer— milk and BSA recipes
• Western blot stripping buffer— glycine-SDS stripping recipe
• Antibody dilution optimization— titration methodology
• Troubleshooting weak signal— no-signal diagnostic tree
• Troubleshooting non-specific bands— multi-band diagnostic tree
• How much antibody for Western blot— starting concentrations by class
• Calculate antibody dilution— dilution factor math
• Primary antibody preparation— dilution buffer and mixing
• Primary antibody incubation time— overnight vs 1-hour incubation
• Choose a primary antibody for WB— selection guidance for WB
• Choose a secondary antibody for WB— host, conjugate, dilution
• Why use a secondary antibody— signal amplification rationale
• Optimize antibody dilution— iterative dilution optimisation
• Store WB membrane after secondary— short and long-term storage