Peptide Adsorption to Labware: Why Dilute Solutions Lose Peptide

What adsorption is and why concentration matters

Adsorption is the binding of dissolved molecules to a surface. Peptides carry charged groups and hydrophobic side chains, so they can bind to the silanol groups of glass and to the hydrophobic surface of plastics. Nothing happens to the peptide chemically: it is still intact, it is just on the wall instead of in the liquid, and a measurement of the liquid reads low.

The key point is that a surface fills up. In a study of the decapeptide cetrorelix, adsorption followed a Langmuir isotherm and reached a plateau of about 0.4 µg per cm² of vial surface.^[1] A worked example with illustrative numbers shows what that means. Suppose 1 mL of solution wets about 5 cm² of a tube wall, and the wall can take up to 0.4 µg/cm². At most about 2 µg can be lost to the wall. In a 1 mg/mL stock (1,000 µg in the tube) that is 0.2%, too small to notice. In a 2 µg/mL working solution (2 µg in the tube) it could be essentially all of the peptide. The same container is harmless at one concentration and decisive at another.

What the recovery studies measured

The cetrorelix work began with a practical problem: HPLC analysis of solutions in the range of 0.2 to 4 µg/mL gave a poor and variable response. Rinsing the emptied vials showed that the missing peptide was on the vial walls. Adsorption decreased in the order glass, then polypropylene and polyethylene (about equal), then PTFE, and differed considerably between glass vials from different manufacturers.^[1]

A study of three cationic membrane-active peptides (mastoparan X, melittin and magainin 2) used analytical HPLC to measure losses to common glass and plastic sample containers. At typical experimental concentrations, 90% or more of the peptide could be lost from solution, and the adsorption was rapid.^[2]

A broader comparison incubated eight radiolabelled endocrine peptides, chosen to span a wide range of net charge, size, end groups and modifications (among them ghrelin, GLP-1, insulin and peptide YY), for 48 hours in glass and plastic tubes, either uncoated or coated with siliconizing fluid. Recovery differed widely between surfaces, siliconizing lowered recovery rather than raising it, and adding 1% bovine serum albumin improved it. With albumin, and with each peptide in its best-suited tube, recovery was above 89% for all eight.^[3]

The time course was measured for colistin, a cyclic lipopeptide antibiotic, in protein-free medium. In large glass, polypropylene and polystyrene tubes, starting concentrations were 44 to 102% of the expected value and 8 to 90% after 24 hours, with half-lives of loss between 0.9 and 12 hours. Low-protein-binding microtubes started at 63 to 99 percent and held 59 to 90 percent at 24 hours. The authors favoured low-protein-binding polypropylene, discouraged polystyrene, and advised keeping the number of dilution steps to a minimum.^[4]

Factors that make losses larger

The studies above point to the same handful of variables. None of them predicts the outcome for a new peptide on its own, which is why reviews of the subject conclude that there is no general approach and that conditions have to be optimised for each analyte.^[6]

• Low concentration. Losses are a fixed amount per surface area, so they matter most in dilute working solutions and in the last steps of a serial dilution.
• Surface area to volume. Small volumes in a given tube expose relatively more wall per microlitre. In a study of amyloid-β peptides in cerebrospinal fluid stored in polypropylene tubes, 0.1 mL aliquots measured lower than 0.5 mL aliquots of the same sample.^[5]
• Charge and hydrophobicity. Cationic peptides bind the negatively charged surface of glass; hydrophobic and lipidated peptides tend to bind plastics.^[2]
• Material and coating. Glass, polypropylene, polystyrene and PTFE behave differently, and coatings do not always help.^{[1, 3]}
• Time and number of transfers. Loss continues over hours, and every new tube, plate well and pipette tip is another fresh surface.^{[4, 6]}

How laboratories reduce adsorption

The options fall into two groups: change the surface the peptide meets, or change the solution so the peptide prefers to stay dissolved. A review of strategies for peptide and protein analysis by LC-MS describes both, including a systematic approach to choosing the dissolution solvent and steps to limit binding in autosampler parts, columns and tubing, where it shows up as carryover between runs.^[6]

1. Keep stock solutions concentrated and make dilute working solutions shortly before use, in as few steps as possible. The dilution calculator helps plan a short series.
2. Use containers that have been shown to give good recovery for the peptide in question. Low-retention polypropylene tubes performed best in the colistin and amyloid-β studies.^{[4, 5]}
3. Where the assay tolerates it, add a carrier protein such as bovine serum albumin, which occupies binding sites on the wall.^[3] It also adds a large amount of other protein, which some measurements cannot accept.
4. Consider a surfactant, an organic cosolvent or a more acidic solvent. For cetrorelix, ethanol, acetic acid and micellar surfactant solutions each raised and steadied the HPLC response, up to fivefold.^[1] Tween-20 raised amyloid-β recovery but did not remove the difference between aliquot volumes.^[5]
5. Make aliquots large enough that wall losses are a small fraction of the volume, and record the tube type with each one.

Testing a container for losses

A simple recovery test answers the question for a specific peptide, concentration and container. Prepare a solution of known concentration, measure part of it at once, leave the rest in the candidate container for the time it would normally stand, and measure it again. Compare both readings with a reference prepared in a container known to give good recovery, or in a solvent that suppresses adsorption. A fall over time, or a reading below the reference, indicates loss to the wall.

Two further checks are useful. Rinsing the emptied container with a stronger solvent, such as one containing organic solvent or acid, and finding peptide in the rinse confirms adsorption rather than degradation, which is how the cetrorelix losses were first identified.^[1] And in a calibration series, points at the low end that fall below the line, or poor repeatability only at low concentrations, are a typical sign of adsorption rather than an instrument problem.^[6] Measurement methods are covered in the HPLC guide and the mass spectrometry guide.

Adsorption is one of several ways a peptide can leave solution; aggregation and poor solubility are covered in the solubility and aggregation guide, and aliquot planning in the freeze-thaw and aliquoting guide. This guide describes laboratory practice only and is not a statement about any particular material. Terms used here are defined in the glossary, and questions can be sent to support.