Reconstituting lyophilised peptides for in-vitro work

Opening a cold vial condenses atmospheric moisture onto and into the cake. Let the vial equilibrate fully to room temperature while sealed — for a small vial this is a matter of minutes, not seconds — then open it.

Before adding anything, spin or tap the vial so that material lodged in the stopper or on the shoulder ends up at the bottom. Lyophilised cake is light and static-prone, and a meaningful fraction of a small fill can sit somewhere other than the base of the vial. Anything left in the closure is mass you weighed but will not recover.

Solvent selection follows from the sequence. Peptides with a net charge and a hydrophilic character usually dissolve directly in sterile water. Basic peptides often go into dilute acetic acid; acidic peptides into dilute ammonium bicarbonate or a mildly basic buffer. Hydrophobic and aggregation-prone sequences may need a small volume of DMSO or acetonitrile to dissolve first, followed by dilution into aqueous buffer.

The order of operations matters for the difficult cases: dissolve in the minimum volume of the strong solvent, confirm the solution is clear, and only then dilute. Adding water first to a peptide that will not dissolve in it produces a suspension that is very hard to rescue, and adding organic solvent afterwards rarely fixes it.

Where a solution will be drawn from more than once over a period of days, bacteriostatic water is used in laboratory settings for its preservative; where the material is destined for cell culture, the preservative itself is usually unwanted and sterile-filtered aqueous buffer is the better choice. Match the solvent to the downstream assay, and record which one you used.

Run the solvent slowly down the inner wall of the vial rather than jetting it directly onto the cake. Peptides are surface-active; a forceful stream generates foam, and foam is peptide adsorbed at an air-water interface where it can denature and aggregate.

Then leave it alone. Most lyophilised material dissolves within a few minutes with occasional gentle swirling or inversion. Do not vortex vigorously and do not shake — both are efficient foam generators. Where dissolution is slow, brief bath sonication or a few minutes at room temperature achieves more than agitation.

The solution should end up clear. Persistent cloudiness, visible particulates or a faint gel means the peptide has not fully dissolved or has aggregated, and the concentration you calculated is not the concentration you have.

Concentration is compound mass over volume, and the trap is that the mass on the label is powder mass, not compound mass. If a net peptide content figure is available, apply it: a vial nominally containing 10 mg of powder at 80% net peptide content holds 8 mg of compound, and reconstituting in 2 mL gives 4 mg/mL, not 5.

For molar concentrations, divide by the molecular weight of the free peptide rather than the salt form, and be explicit in your notes about which you used. This is a common source of discrepancy between labs working on the same compound.

Write the resulting figure on the tube along with the solvent, the lot number and the date. A concentration reconstructed from memory a fortnight later is a guess.

Freeze-thaw cycling degrades peptides in solution through several mechanisms at once — concentration of solutes in the unfrozen fraction, pH shifts as buffer components crystallise, and interface stress. Each cycle costs something.

The fix is to divide the stock into single-use aliquots at the moment of preparation, so that each tube is thawed exactly once. Size the aliquots to your actual experiment rather than to a round number, because a tube half-used and refrozen defeats the purpose.

Low-binding tubes are worth using for dilute solutions. At low concentrations, adsorption to polypropylene walls removes a non-trivial fraction of the peptide from solution, and the effect is worst precisely where you can least afford it. A small amount of carrier protein serves the same purpose where the assay tolerates it.