Below roughly 1 µg/mL, a meaningful fraction of a peptide can end up stuck to the plasticware instead of dissolved in the buffer. The solution looks identical. The concentration is not what the calculation says. And because the loss scales with surface area rather than with the amount present, it gets proportionally worse the more dilute you go — exactly where accuracy usually matters most.
Why surfaces take peptide out of solution
Polypropylene is hydrophobic. So are large parts of most peptides — the non-polar residues that sit buried when the molecule is folded and exposed when it is not. Put the two together and adsorption is thermodynamically favourable: the peptide trades an unfavourable water contact for a favourable plastic one.
Charge contributes as well. Plastic surfaces carry some charge from manufacturing and handling, and basic peptides — arginine- and lysine-rich sequences — interact electrostatically on top of the hydrophobic effect. This is why two peptides at identical concentration in identical tubes can lose very different fractions.
Why it is a low-concentration problem specifically
A tube wall has a roughly fixed capacity: the surface adsorbs approximately the same absolute quantity regardless of what is in solution, until it saturates. That quantity is a rounding error against a milligram and a catastrophe against a nanogram.
| Concentration in a 1 mL tube | Peptide present | If the wall takes ~100 ng |
|---|---|---|
| 1 mg/mL | 1,000,000 ng | 0.01% lost — invisible |
| 10 µg/mL | 10,000 ng | 1% lost — negligible |
| 1 µg/mL | 1,000 ng | 10% lost — starts to matter |
| 100 ng/mL | 100 ng | up to 100% lost — the experiment is gone |
The figures are illustrative rather than universal — actual capacity depends on the peptide, the surface and the buffer — but the shape of the problem is real and it is the shape that matters.
What actually helps
Does a carrier protein work?
It is the most effective single measure. Adding a carrier — bovine serum albumin at around 0.1%, or an equivalent — saturates the binding sites on the surface so your peptide stays in solution. The caveat is that it has to be compatible with the assay: a carrier protein is ruinous in mass spectrometry, can interfere with protein quantification, and may bind the peptide itself in some systems. Where it is tolerated, it is the answer.
Do low-binding tubes help?
Yes, and they are the easy win where a carrier is not an option. Low-binding or protein-lobind plasticware is surface-treated to reduce exactly this interaction, and it costs a little more than standard tubes. Use them for working dilutions rather than for stocks, since stocks are concentrated enough not to care.
What about glass?
Glass is not a general escape. Silanised glass adsorbs less than untreated glass, but untreated borosilicate can be as bad as polypropylene for basic peptides because the silanol surface carries negative charge. Swapping to glass without treating it often moves the problem rather than solving it.
Does the buffer composition matter?
It does. A small amount of non-ionic surfactant — polysorbate at low concentration, for instance — competes for the surface effectively, and raising ionic strength can reduce electrostatic adsorption. Both change the solution chemistry, so both need to be compatible with what comes next; neither is free.
Does pre-rinsing the tip help?
Pre-wetting a pipette tip with the solution before the actual transfer is standard technique for good reason: it saturates the tip surface and improves the accuracy of the volume that follows. It helps, it costs nothing, and it does not substitute for the measures above.
Which peptides are most affected?
Three kinds, for three different reasons. Acylated peptides — the incretin class and anything else carrying a fatty acid chain to bind albumin — are surfactant-like by design and concentrate at any hydrophobic surface; that is a direct consequence of how they were engineered. Strongly basic peptides such as Selank, with several arginine and lysine residues, bind negatively charged surfaces electrostatically, which makes untreated glass worse for them than plastic. And very short, unstructured sequences — TB-500 at seven residues, GHK at three — carry so little mass that a fixed surface loss is a large fraction of what was there. Folded proteins such as IGF-1 LR3 are less exposed per molecule but are worked with at nanogram concentrations, where a carrier protein is routine rather than optional.
Does adsorption affect a stock solution too?
In principle, but not in a way that matters. At 1 mg/mL the wall's capacity is a rounding error against the amount in solution, which is why the problem is specific to dilutions rather than stocks. The place a stock does lose material is different: repeated freeze–thaw cycles drive aggregation, and aggregated peptide leaves solution by a mechanism that has nothing to do with the tube. Keep stocks concentrated and aliquoted; reserve the low-binding tubes and carrier protein for the working dilutions, where the surface actually competes.
How to know whether it is happening to you
The signature is a concentration–response curve that flattens or shifts at the dilute end while behaving normally at the concentrated end, or results that vary with how long a dilution sat before use. If a fresh dilution gives a different answer from one prepared an hour earlier, adsorption is the first thing to suspect — the wall is still taking material while the tube sits.
The direct test is to prepare a dilution, hold it, and quantify it against a freshly prepared one. If the aged sample reads low, the peptide is on the plastic.
Whatever you conclude, record the plasticware and the buffer in the write-up. Two laboratories running the same nominal concentration in different tubes are not running the same experiment, and this is one of the more common reasons a result does not reproduce.
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