Here, we present two possible sources of errors that may both contribute to significant inaccuracy in peptide assay methods:
the filtration material (cellulose) used in sample clean up and the vial material (glass and plastic) used in liquid chromatographic
analysis. This study is based on β-endorphin but has relevance for most peptide assays.
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In recent years, significant attention has been paid to the use of peptides as therapeutic agents and this trend is expected
to continue (1,2). Because of their wide-ranging activities in many physiological processes such as acting as signaling molecules
and growth factors, peptides play a substantial role as therapeutic mediators, especially in the areas of neurology, endocrinology,
and hematology (3,4). Even though peptides have been shown to be promising therapeutics for many disease conditions, developing
them as drugs is a challenging process. Accurate quantification of peptides is one of these challenges and their poor stability,
losses during manufacturing and delivery process, and their high cost are other issues (4–6). Incorrect quantification of
peptides may lead to false conclusions in research and may also adversely affect the dosage that is prescribed for patients.
Quantification methods may vary depending on the nature of the sample and the analytical instruments used. If the analyte
is extracted from a natural source or biological matrix, sample purification is required before chemical analysis is undertaken
to remove molecules that may interfere with the analytical method or have detrimental effects on the instrumentation. Various
sample preparation methods such as solvent extraction, ultrafiltration, and precipitation are commonly applied to biological
samples. Analyte loss during these processes can be significant. Ultrafiltration devices are commonly used for the separation
of small peptides from proteins and other larger molecules. If the peptide is insufficiently stable to withstand acid precipitation
followed by solvent evaporation, ultrafiltration may then be the only option for sample purification. However, the ranges
of filter materials available have variable adsorption affinities toward different peptides (7). Although internal standards
are often used to correct for sample preparation losses, if the internal standard molecule is not sufficiently physicochemically
similar to the analyte molecule then inaccuracies due to losses will not be fully corrected.
The adsorption of peptides to container materials during the manufacturing process and subsequent storage is well-known (7,8).
However, the adsorption during analytical processes has been given scant attention. Even though the analysis time is relatively
short, the adsorption of peptides to sample vials during analysis may be a significant source of error. As liquid chromatography
(LC) analysis is routinely accomplished using autosamplers containing a large number of samples and standards, the contact
time of the analyte with the container may be sufficiently long to be problematic. The aim of this study was to investigate
the significance of the inaccuracy in peptide quantification caused by adsorption during the ultrafiltration and chromatographic
analysis procedures. Two different types of limited volume inserts (LVIs) commonly used in LC vials, plastic (polypropylene)
and glass, were used to evaluate the effect of sample vials during LC analysis. Commonly used microcentrifugal ultrafiltration
devices (10 kDa cut off) constructed from cellulose were evaluated for the effect of the sample preparation step. β-Endorphin
(β-END [1-31]) was quantified using liquid chromatography–mass spectrometry (LC–MS).
Experimental
β-END (1-31)–human was purchased from Sigma, USA, and β-END (1-31)-rat was purchased from Auspep Pty Ltd. Australia. An Agilent
Technologies high performance liquid chromatography (HPLC) system consisting of an Agilent 1100 LC pump and an Agilent 1100
well-plate autosampler with a 150 mm × 2.00 mm, 5-μm d
p Jupiter C4 (300 Å) reversed-phase HPLC column (Phenomenex) were used for separations. A binary solvent gradient consisting
of water 0.1% (v/v) formic acid in MilliQ (Millipore Corporation) water (solvent A) and 0.1% (v/v) formic acid in acetonitrile
(solvent B) was used for all separations. An API 3000 tandem mass spectrometer equipped with a turbo ion spray interface and
supported by Analyst 1.5 software (Applied Biosystems) were used for mass spectrometric detection in multiple reaction monitoring
(MRM) mode. The 694/136 pair (5+ charged endorphin molecular ion and fragment) was used for MRM detection of β-END (1-31).
To test peptide adsorption on filter material, a solution containing β-END (1-31)–rat (10 μM) in Milli-Q water was filtered
through the Microcon centrifugal devices (Millipore Corporation) using the recommended centrifugal force of 12,500 at 25 °C
for 10 min. The filtered sample was transferred to a plastic limited volume insert and injected onto the LC column. Triplicate
analyses were performed using LC–MS (MRM mode). The gradient used for this study started at 0% B and increased to 5% B during
the first 4 min, then to 70% B during the next 1 min. The mobile-phase composition was maintained at 70% B composition for
5 min and then returned to 0% B during the next 2 min. The column was re-equilibrated with 0% B for 10 min before the next
injection. Quantification was carried out using the peak areas. The same experiment was repeated in triplicate without the
filtering step.
To test peptide adsorption on LC LVIs, a solution containing β-END (1-31)–rat (0.5 μM) and β-END (1-31)–human (0.5 μM) in
Milli-Q water was prepared. Immediately following sample preparation, the solution was transferred to a glass LVI and injections
were made at 53-min intervals (run time for each injection) for 6 h and analyzed by the LC–MS (MRM mode). During the experiment,
the sample was maintained at 10 °C in the autosampler. The gradient started with 0% B for the first 3 min and changed to 50%
B over the next 30 min. The mobile-phase composition was then changed to 100% B within the next 5 min and maintained for 2
min. It was then returned to 0% B within the next 3 min and the column was re-equilibrated for 10 min before the next injection.
Quantification was carried out using the peak areas. The same experiment was repeated using a plastic (polypropylene) LVI
in place of glass inserts.
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