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Western Blot Woes: Why Is My Protein Running at the Wrong Molecular Weight?
- CLYTE research team
- 3 days ago
- 8 min read
It's one of the most common and frustrating moments at the lab bench: you've run your Western Blot, your controls look good, but your protein of interest is showing a strong band at 50 kDa when its calculated sequence weight is 40 kDa. Or worse, your 120 kDa protein is appearing as a 50 kDa fragment.
You might also be interested in this WB protocol! Ask Sophie about Protein Running at the Wrong Molecular Weight!
Before you throw out your samples, your antibody, or your gel, it's important to understand that this is a very common scenario. The "calculated" molecular weight (derived from the amino acid sequence) is purely theoretical. The "observed" molecular weight (where it migrates on a gel) is a physical reality affected by a host of biological and technical factors.
Understanding why they don't match is the first step in troubleshooting your experiment and, in many cases, discovering important new details about your protein.
Is It Your Protein or Your Gel? The Two Main Culprits
When a band shows up at the wrong size, the cause almost always falls into two categories:
Biological Reasons: The protein in your sample is not the simple, theoretical amino acid chain you expect. It has been modified, cut, or is part of a complex.
Technical Reasons: The protein is fine, but your experimental conditions (the gel, the buffers, the standards) are causing it to migrate in an unexpected way.
Biological Reasons: Why Your Protein's "Actual" Weight is Different
These are often the most interesting reasons, as they reveal the true post-translational life of your protein inside the cell.
Post-Translational Modifications (PTMs): The Protein's "Accessories"
PTMs are chemical additions to the protein after it's been synthesized, and they are a major cause of upward shifts in molecular weight.
Glycosylation: This is the most common culprit for proteins running higher than expected. The covalent addition of bulky sugar moieties (N-linked or O-linked) can significantly slow a protein's migration. For example, the protein PD-L1 has a calculated weight of ~33 kDa but is often observed at 45-70 kDa due to heavy glycosylation.
Ubiquitination: The addition of one or more ubiquitin proteins (which are ~8.6 kDa each) will cause a distinct and significant increase in molecular weight.
Phosphorylation: While a single phosphate group is too small to notice, many proteins are phosphorylated at multiple sites. This cumulative addition of negative charges can alter how the protein binds to SDS and slow its migration, making it appear slightly higher.
Cleavage and Processing: The Protein is Being Cut
If your protein is running lower than expected, it's almost certainly because it has been cut.
Protein Precursors: Many proteins are synthesized as inactive "pro-proteins" or with "signal peptides" that direct them to the correct cellular location. These extra sequences are then cleaved off to create the mature, functional protein. The full-length sequence in the database may represent the precursor, while your antibody is detecting the smaller, mature form. A classic example is Caspase-3, which exists as a 32 kDa proenzyme until it is cleaved into its active 17/19 kDa subunits.
Proteolytic Degradation: This is an experimental artifact. When you lyse your cells, you release proteases that can "chew up" your protein of interest. This results in distinct bands at a lower molecular weight. This is especially common if samples are not kept on ice or if protease inhibitors are not added to the lysis buffer.
Splice Variants & Isoforms: Different Versions of the Same Gene
Your cell line may not be expressing the single "canonical" version of the protein. Alternative splicing of the mRNA can create multiple different isoforms from the same gene. You may be detecting a shorter or longer splice variant that is perfectly valid, but different from the sequence you used for your calculation.
Multimers & Aggregates: The Protein Isn't Running Solo
If your protein is appearing at a weight that is 2x, 3x, or significantly higher than expected, it may not be fully denatured.
Dimers and Multimers: Incomplete sample preparation (e.g., old or insufficient reducing agents like DTT or 2-mercaptoethanol, or not boiling long enough) can fail to break the disulfide bonds that hold protein complexes together. You may be observing a dimer (2x weight) or trimer (3x weight).
Aggregates: Transmembrane proteins or those with hydrophobic domains are notoriously "sticky" and can aggregate, especially during lysis, causing them to run much higher on the gel.
Technical Reasons: Why Your Experiment is Giving the Wrong Weight
Sometimes the protein is exactly as expected, but the experiment itself is flawed.
The Ladder is Lying: Inaccurate Molecular Weight Markers
This is a critical and often-overlooked factor. The pre-stained protein standards (ladders) you use to judge size are not always accurate. Studies have shown that there can be significant variation between different commercial brands, and even between lots. These standards can be particularly unreliable for very high molecular weight proteins. Your 130 kDa protein might be running perfectly, but the "130 kDa" band on your ladder might be running at 140 kDa, throwing off your entire analysis.
SDS-PAGE Anomalies: When the Gel Itself is the Problem
SDS-PAGE works by coating proteins in a uniform negative charge (from SDS) so they migrate based on size alone. But this assumption can fail.
Amino Acid Composition: Proteins with an unusual number of highly charged residues (e.g., many acidic or basic amino acids like lysine and arginine) may not bind SDS uniformly. This causes them to migrate abnormally. The tumor suppressor p53 is a famous example, with a calculated weight of 43.7 kDa but often running at ~53 kDa.
Intrinsically Disordered Proteins: Proteins that lack a stable 3D structure can also run at anomalous weights.
Antibody Problems: Detecting the Wrong Thing
The band you see might not even be your protein. It could be a non-specific "parasite band" that your antibody is accidentally binding to. This is common if the antibody concentration is too high or if you are using a lower-quality (e.g., non-affinity-purified) antibody.
A Western Blot Troubleshooting Protocol: What to Do Next
Check the Literature: Before anything else, search for your protein (and the specific antibody you're using) in published papers. Is it known to have PTMs, isoforms, or run at an odd weight? The manufacturer's datasheet often provides an expected vs. observed weight.
Validate Your Sample Prep: Remake your samples. Use a fresh protease inhibitor cocktail in your lysis buffer and always keep samples on ice. Add fresh DTT or β-mercaptoethanol to your loading buffer and boil your samples for 5-10 minutes to ensure complete denaturation.
Validate Your Reagents:
Antibody: Try optimizing the antibody concentration (use less). If possible, use a blocking peptide or a knock-out (KO) cell line to confirm the band is your target.
Ladder: Run your sample next to two different brands of protein ladders to see if the "observed" weight changes relative to the markers.
Test a Hypothesis:
Suspect Glycosylation? Treat your lysate with an enzyme like PNGase F (which removes N-linked glycans). If your higher band shifts down to its calculated weight, you've found your answer.
Suspect Degradation? Compare a fresh lysate (with inhibitors) to an old one. If the lower band is gone in the fresh sample, degradation was the problem.
Conclusion: A Mismatched Band is a Clue, Not a Failure
An unexpected molecular weight in a Western Blot is not usually a sign of a completely failed experiment. More often, it is a critical clue pointing toward the complex biological processing of your protein—from PTMs and cleavage to isoforms. By troubleshooting the technical and biological possibilities, you can solve the puzzle and gain a much deeper understanding of your target.
Frequently Asked Questions (FAQ)
What happens if you load too much protein in a Western blot?
Loading too much protein in a single lane can cause several problems that ruin your results.
Band Distortion: The most common issue is "smiling" or "frowning" bands, where the band is warped instead of flat, making it impossible to analyze. Overloaded protein can also cause the lane to "bleed" or "smear" into adjacent lanes.
Poor Separation: The gel's separating power gets overwhelmed, and bands that are close in size may merge into one large, unresolved smear.
High Background: Excess protein can non-specifically bind to the membrane, leading to high background noise and making it difficult to see your specific band.
Signal Saturation: If you are using a chemiluminescent (ECL) substrate, the signal from an overloaded band will be "blown out" (completely blacked out on film or maxed out on a digital imager). This makes quantification impossible, as you can't tell the difference between "a lot" of protein and "way too much" protein.
How is it possible that the wrong protein could be formed during protein synthesis?
This is a great question. While it's rare for a cell to make a completely random, "wrong" protein, several common mechanisms produce different versions of the expected protein:
Alternative Splicing: This is the most common reason. When a gene is transcribed from DNA to mRNA, it can be "spliced" in different ways. This process can include or exclude certain sections (exons), resulting in multiple distinct protein isoforms from a single gene. These isoforms are all "correct" but will have different lengths and, therefore, different molecular weights.
Mutations: A mutation in the gene's DNA sequence can cause an error. The most relevant for size changes is a nonsense mutation, which creates a premature "stop" signal, resulting in a shorter, truncated protein.
Ribosomal Frameshifting: This is a rarer event where the ribosome "slips" during translation, changing the reading frame. This leads to a completely different (and usually non-functional) amino acid sequence from that point on.
How can I separate proteins with similar molecular weights?
This is a common challenge when you're looking at proteins that are very close in size (e.g., a phosphorylated vs. unphosphorylated form, or two different isoforms). Here are the best strategies:
Optimize Your Gel Percentage: Use a higher percentage acrylamide gel (e.g., 12% or 15%) for better resolution of smaller proteins, or a lower percentage (e.g., 8%) for larger ones.
Use a Gradient Gel: A 4-20% gradient gel is often the best solution. The gradually decreasing pore size sharpens the bands and expands the separation distance between them.
Change Your Buffer System: Standard Tris-Glycine gels are good, but Bis-Tris gels (run with MOPS or MES buffer) often provide much sharper bands and better resolution. For very small proteins (<15 kDa), a Tris-Tricine system is superior.
Run a Longer Gel: A physically longer gel (e.g., a 15-well, 15 cm gel) gives the proteins more "runway" to separate from each other.
Why is the actual band size different from the predicted?
This is the central question of the main article! The predicted (or calculated) weight is a theoretical number based only on the amino acid sequence. The actual (or observed) weight is what you see on the gel, which is affected by numerous real-world factors:
Post-Translational Modifications (PTMs): Biological "accessories" like glycosylation add significant bulk, making the protein run higher (slower).
Protein Cleavage: The protein may be synthesized as a large precursor and then "cut" into its smaller, mature form (e.g., removing a signal peptide). This makes it run lower than the full-length predicted sequence.
Splice Variants: Your antibody might be detecting a shorter or longer isoform than the "canonical" sequence you used for your calculation.
Gel Anomalies: Some proteins (like highly acidic or basic ones) don't bind SDS uniformly and migrate at an unexpected speed.
Inaccurate Ladders: The pre-stained molecular weight markers themselves can be imprecise, throwing off your comparison.