I produced a fluorescent protein fused to a cell-penetrating peptide and purified it using anti-His magnetic beads. Elution was done with 50 mM Tris, 500 mM NaCl, and 250 mM imidazole at pH 7.4. I then dialyzed the eluted protein into a buffer containing 50 mM Tris and 300 mM NaCl, pH 7.4.
When I ran SDS-PAGE to assess purity, the sample before dialysis showed a band at the expected molecular weight. However, after dialysis, the protein appeared as a much larger band, suggesting aggregation or multimer formation. I have lowered the protein concentration for SDS-PAGE, still encountered the same issue.
Has anyone encountered this before? What could be the cause, and how can I prevent or reverse it?
Any suggestions are welcome. Thanks!
how do you know that the higher band is really your protein? mass spec? western blot? it is rare that multimers can resist to the denaturation conditions of a SDS page and boiled in reducing laemmli however if there is not enough Beta mercapto in your laemmli (old buffer) some disulfide bond scould have been formed during dialysis and are not denatured by your laemmli buffer +100°C 10min...
@Didier Thank you for your response. I haven't done mass spec or WB for the protein in question. But I have reproduced this aggregation with 3 different batches of sample. The positive control for SDS-PAGE indicated that reducing condition was ok. I will try WB to confirm its actually the fluorescent protein. If it's confirmed, do you have any suggestions for me to try next? I would love to hear more from you.
you produce your fusion protein in E. Coli and is it soluble? your dialysis buffer have a lower NaCl concentration... maybe your protein precipitates. Signal peptide have hydrophobic stretchs maybe it induces multimerization of your fluorescent protein? is your protein preparation fluorescent after renaturation/dialysis (this would indicate taht it is the fusion protein expected.)
Dear Khai Nguyen
Is this cell penetrating peptide hydrofobic?
Since generally soluble protein aggregates though non covalent interaction appear as monomer in SDS-page while the aggregates are shown only in non denaturing approaches as SEC, DLS; NMR.
The fact that aggregates are present in SDS-page it seems to be due or hydrofobic interactionstipical of membrane proteins or covalent (eg S-S bond formation) but only if you are running non reducing SDS-page.
best regards
Manuele
Didier Poncet Thank you for your follow-up. Yes, my protein is soluble. The signal peptide has a couple of hydrophobic amino acids close to each other. My protein was fluorescent after dialysis. Do you have any suggestions on how to remove the salt without causing aggregation?
Manuele Martinelli Thank you for your response. The signal peptide has a hydrophobic stretch in it. But the protein itself (without the signal peptide) also aggregated, forming a smear instead of a distinct band in SDS-PAGE. The SDS-PAGE was done under a reducing condition. I really appreciate if you have any suggestions on how to avoid this problem.
if the protein is fluorescent after dialysis this mean that it is (at least in part) refolded but if the protein itself (without the signal peptide) also aggregated, it does not seems normal to me; try a brand new laemmli buffer (one that smell Beta mercapto!!!) and boil your samples 10 min. For the salt I woud suggest to start with the same concentration in the dialysis buffer than in the solubilization buffer.
Khai Nguyen The appearance of a high molecular weight band on SDS-PAGE after dialysis of your purified His-tagged fluorescent protein may indicate protein aggregation or multimer formation, possibly due to incomplete denaturation or improper folding during or after dialysis. This can result from inadequate SDS or reducing agent in the sample buffer, residual imidazole or salts affecting protein solubility, or storage conditions that promote oligomerization. To resolve this, ensure thorough denaturation with SDS and a reducing agent like DTT or β-mercaptoethanol, optimize dialysis conditions (e.g., buffer composition and pH), and consider including mild detergents or additives to maintain protein solubility.
Annie Huang
Thank you for your reply. Do you have a protocol that I could reference for my future dialysis attempt? I would love to know more about details such as the concentrations of imidazole and salt in dialysis buffers, the amount of Triton X-100 to be added, and when Triton X-100 should be added. Any recommendation is welcome.
Maybe your protein captures some Ni(2+) or Co(2+) from the beads, and keeps it bound to the His-tag? Then, the protein might remain monomeric when imidazole is still around but polymerize after its removal via His-tag - metal bridges. Try adding some EDTA (like, around 5 mM) to the first portion of the dialysis buffer and see if it helps.
Actually, metal ions are known to promote Histidine oxidation, and oxidized His is a good crosslinker by itself. It can make a covalent bond with His, Arg, Lys and Cys residues. Naturally, the lower is concentration of your protein, the lower would be the extent of intramolecular crosslinking.
- Badges
- Science topic
- Similar topics
- Engineering
- Materials Engineering
- Powders