Understanding the impact of matrix effects on multiply charged compounds.
Its widespread adoption resulted in the discovery of a phenomenon first observed with the API technique, in which a change in response was observed for a given concentration of a target analyte in the presence of other sample components. (1) This phenomenon became known as 'matrix effects.' There are two basic types of matrix effects: ion suppression, which occurs when the signal is suppressed; and ion enhancement, which occurs if the signal is enhanced. (2) Both effects occur in the presence of matrix components. According to numerous studies, matrix effects are a result of a disrupted ESI process by sources such as endogenous, non-volatile and exogenous components that affect ion ejection from the liquid droplet surface.
Stable label internal standards (IS) have been found to effectively account for matrix effects when used in conjunction with adequate sample clean-up procedures. Matrix factors (MF) are calculated for the analyte and the IS separately and a ratio of the two factors yields the IS-normalized MF for the analyte. Usually, six different lots of matrix are tested to determine the IS-normalized MF. The assessment of matrix effects is now a standard criterion and the submission of the relevant data is a standard requirement for the investigational new drug (IND) application submission process. (3), (4)
Overcoming matrix effects for peptide-based new molecular entities (NME) is a novel challenge in preclinical studies. As stable IS are rarely made available during the early preclinical development stages, and chemistry-based sample clean-up methods may not deliver the required efficiency, determination of the MF becomes challenging. Immuno-affinity purification is often preferred for sample clean-up when stable label IS is unavailable; it is, however, expensive and therefore not always available during early development.
Hydrophobic peptides are of particular interest to researchers. One of their characteristics is that they elute late in the mobile phase gradient, usually around the same time as endogenous plasma components such as phospholipids. This fact, combined with the lack of immuno-affinity based clean-up techniques in early preclinical studies, makes bioanalytical method development challenging. Endogenous phospholipids are present to some extent in most extraction techniques used in bioanalysis and are major causes of ion suppression. Although the impact of endogenous phospholipids has been extensively studied for singly charged small molecules, their effect on multiply charged compounds such as peptides has not been fully characterized yet. In addition, although chemistry based sample clean-up techniques such as solid phase extraction (SPE) exploit the large difference between the chemistry of a small molecule and cellular debris--such as endogenous proteins, peptides and phospholipids--this difference in chemistries becomes smaller when trying to purify a hydrophobic peptide, limiting the extent of the clean-up. To address these challenges, pharmaceutical companies can outsource to companies with experience in developing, validating and utilizing assays for a range of drug development projects.
In this study, the multiply charged ions were generated using four peptide standards; Porcine Growth Hormone Releasing Factor (GRF) FW:5108.76, Glucagon-Like Peptide (GLP) FW: 3297.63, C-Type Natriuretic Peptide FW:2197.6 (Sigma Chemicals, St Louis, Missouri, USA) and a proprietary peptide, FW:1282. Intermediate solutions of 100 ng/mL were prepared using 10 mM ammonium acetate. A caffeine standard of 1 mg/mL solution in methanol (Sigma Chemicals) was also used, as well as K-2 EDTA human and rat plasma (Rockland Immunochemicals, Gilbertsville, Pennsylvania, USA). An Agilent 1100 Binary Pump HPLC system was operated at a flow rate of 0.5 mL/min. Water (0.1% formic acid) was used in mobile phase A and acetonitrile (0.1% formic acid) was used in mobile phase B. The gradient was set at 10% B for 1 minute; it was ramped to 90% B for 3 minutes and held for 1 minute, for a total run time of 5 minutes. Samples were injected on a Waters Corp 2.1 x 50 mm, 3.5 [micro]m C4 BEH column using a CTC PAL autosampler with an injection volume of 20 [micro]L.
The quantitative analysis of the peptides was done using a Thermo Scientific TSQ Vantage triple stage quadrupole mass spectrometer with a heated electrospray ionization probe. The instrument was operated as follows: 4.2 kV spray voltage, 300 [degrees]C vaporizer temperature, 350 [degrees]C ion transfer tube temperature, 65 au sheath gas pressure, 15 au auxiliary gas pressure and an Argon filled collision gas pressure of 1.5 mTorr. Resolution at full width half maxima (FWHM) for Q1 was set to 1.0 FWHM and to 0.7 FWHM for Q3 using a total scan time of 0.1 s and a scan width of 0.002 u.
Results and Discussion
Figure 1 illustrates the chromatographic elution profile for the proprietary hydrophobic peptide normalized against the elution profiles of the common endogenous phospholipids and lysophospholipids. The common phosphonate product ion (m/z 184) was monitored from precursor ions at m/z 496, 524, 704, 758, 786 and 804 as a general indication of phospholipid elution profiles. As demonstrated in Figure 1, a significant amount of late-eluting phospholipids were present. These can accumulate on the column and elute randomly, a common issue when implementing a protein precipitation clean-up method. Despite the use of C4 column chemistry, the hydrophobic peptide still eluted in a zone close to the elution of the common phospholipids, possibly indicating that more sample clean-up was required to develop a more robust method in case ion suppression became an issue.
[FIGURE 1 OMITTED]
The results of the ion suppression infusion experiment are presented in Figures 2a and 2b. Whereas the reference small molecule (caffeine) trace clearly showed ion suppression, the trend for the multiply charged compounds that were infused individually exhibited signs of ion enhancement. Although the extent of enhancement was not confirmed quantitatively, the differences in the matrix effect trend indicated a difference between singly charged small molecules and multiply charged biomolecules. One explanation of this phenomenon, based on this initial observation, could be that the multiple charges present on biomolecules created a higher electrical imbalance within the droplet, thus enhancing ejection of gas phase ions when compared with similar droplet stoichiometry for singly charged species.
[FIGURE 2 OMITTED]
To conclude the study and ensure a robust bioanalytical method for the proprietary hydrophobic peptide, the clean-up was changed to SPE and ultra performance liquid chromatography (UPLC) was used. This is shown in Figure 3. As the method utilized a chemical analogue IS, clean-up became important to facilitate a high quality bioanalytical method. The limit of quantitation was enhanced to 0.05 ng/mL using SPE/UPLC compared with 0.125 ng/mL using protein precipitation-based sample clean-up as demonstrated in Figure 3.
[FIGURE 3 OMITTED]
Inadequate sample clean-up, the presence of endogenous, non-volatile and exogenous compounds and poor chromatography are the main factors that contribute to the occurrence of matrix effects, negatively impacting accuracy in bioanalytical studies. Multiply charged compounds such as hydrophobic peptides are of particular interest as they represent an analytical challenge for traditional clean-up techniques based on extraction chemistry, such as SPE and LLE, particularly when no stable label IS are available. To meet both the sensitivity and the validated bioanalysis requirements, clean-up techniques such as microelution SPE coupled to modern UPLC-MS/MS techniques using high resolution accurate mass may help to reduce the impact of matrix effects during the bioanalysis of biologics.
(1.) D.L. Burhman, et al., JASMS 7, 1099-1105 (1996).
(2.) L.L. Jessome, et al., LCGC North America 24(5), 498-511 (2006).
(3.) US Department of Health and Human Services, Food and Drug Administration, Center for Drug Evaluation and Research (CDER), "Guidance for Industry Bioanalytical Method Validation," (May 2001). www.fda.gov
(4.) C.T. Viswanathan, et al., "Workshop/Conference Report--Quantitative Bioanalytical Methods Validation and Implementation: Best Practices for Chromatographic and Ligand Binding Assays," AAPS J. 9(1), E30-E42 (2007).
(5.) R. King, et al., JASMS 11, 942-950 (2000).
For more information
Rohan Thakur Taylor Technology Princeton, New Jersey, USA.
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|Date:||Mar 1, 2010|
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