Interaction experiments focus on the physical aspects of the separation as they relate to band broadening. Successful peptide
mapping depends, of course, on the interaction among the peptides, the mobile phase, and the column surface chemistry. Figure
5 (top) shows the separation of a tryptic digest of enolase with a 3.5 mm C18 HPLC column whose particles have 300 Å pores.
This is typical of the most common peptide separation columns. Figure 5 (bottom) is for a 1.7 μm UPLC column. Conditions are
the same for both columns.
Figure 5. Suitability of UPLC for peptide mapping in digestion of enolase is shown. Top plot is HPLC. Bottom plot is UPLC.
The peaks are normalized versus the tallest peak.
Note the greater number of peaks in the UPLC separation. The overall resolution and sensitivity are higher. In the UPLC map,
there are several small peaks that are difficult to discern with the HPLC run. This demonstrates that UPLC offers higher resolution
and sensitivity when compared with HPLC under the same gradient conditions. As is always observed when comparing two different
column chemistries, the separations are not identical in every detail. The overall appearance of the chromatograms is, however,
similar. This suggests that the selectivity of the UPLC column is suitable for peptide mapping.
The higher resolution and sensitivity with UPLC are particularly important when using the peptide map to detect modified peptides.
Higher resolution ensures that modified peptides are resolved from the unmodified form, as well as from other peptides in
the digest. Higher sensitivity means that modified peptides can be detected at lower levels. Figure 6 shows that UPLC is a
good way to separate a deamidated peptide from its unmodified form. UPLC should be the technique of choice for detecting all
the peptides in a sample.
Figure 6. Native and deamidated peptides can be separated. The upper graph is native peptide. The lower graph is of a sample
intentionally degraded before analysis. The peaks are normalized versus the tallest peak.
Scientists frequently interface peptide mapping with Electroscopy Ionization (ESI-MS) to provide additional information about
the eluting peptides, including molecular weight and sequence. MS can also identify modified peptides and glycosylation sites.
Therefore, it is important that a peptide mapping technique work well under conditions that are favorable for ESI-MS.
Mobile phases make a difference. TFA is commonly used as an acidic additive for peptide maps with UV detection, but it can
lead to suppression of ionization and reduced sensitivity in ESI-MS. Formic acid is more desirable for LC-MS, because it causes
less ion suppression than TFA. However, many reversed phase columns used for peptide mapping show lower retention and broader
peaks with formic acid than with TFA. Figure 7 compares the separation of several peptides with formic acid and with TFA on
a UPLC column with MS detection. With formic acid the peak heights are about three times higher. At the same time, there is
only a slight reduction in retention, corresponding to a difference of a few percent organic at the point of elution, and
a slight increase in peak width. This result indicates that the UPLC columns are compatible with formic acid to produce good
results for ESI-MS.
Figure 7. Plots show Influence of acidic additive in UPLC peptide mapping. Upper plot uses formic acid. Bottom plot uses
TFA. The peaks are normalized versus the tallest peak. Initially column mobile phase is A and the carrier is B.
Glycosylation is an important post-translational modification that plays a critical role in determining the efficacy and safety
of a therapeutic protein. Glycosylation can be analyzed on the intact protein by mass spectrometry, as released glycans by
a variety of techniques, or as glycopeptides in LC-MS peptide maps. When glycosylation can be characterized with LC-MS of
the glycopeptides, the site of attachment can be directly determined, and structural information can be obtained through MS-MS
experiments. This approach is limited, however, by the poor chromatographic peak shape of glycopeptides and incomplete resolution
of glycoforms with HPLC peptide mapping. The poor peak shape has been attributed to the large size of the glycopeptides and
their heterogeneous structure.
Figure 8. UPLC of tryptic fragments of a-1 acid glycoprotein. Selected ion chromatogram at m/z 657 of sialyted glycopeptides
shows many peaks. The peaks are normalized versus the tallest peak.
UPLC can do better. Figure 8 shows the UPLC-MS separation of a tryptic digest of α-1 acid glycoprotein. The MS detection was
performed with a Q-Tof mass spectrometer, which is well suited for glycopeptides because of its extended mass range. Data
are plotted as a selected-ion chromatogram for m/z 657, a signature ion for glycopeptides resulting from carbohydrate fragments.
The glycopeptides are detected as sharp, symmetrical peaks. These characteristics are important for minimizing spectral overlap
of different glycoforms of the same peptide. UPLC combined with ESI/TOF mass spectrometry will be a powerful tool for studying
the glycosylation state of proteins.
UPLC provides better peptide maps than can be obtained with HPLC. Better resolution is available in combination with improved
sensitivity. Multiple strategies are available for reducing run time without compromising resolution. Selectivity is comparable
to that of common reversed phase HPLC peptide mapping columns and can be easily transferred to alternative modifiers that
give better sensitivity in ESI-MS. The UPLC separations are proving highly suitable for separation of glycopeptides. With
the extension of available columns to various alternative chemistries and even smaller particles, UPLC will represent the
next generation tool for peptide mapping.
Jeff Mazzeo, Ph.D., is applied technology director at Waters Corp., 34 Maple Street, Milford, MA 01757, 508.482.3462, fax 508.482.4100, email@example.com
Tom Wheat, Ph.D., is life sciences application manager at Waters Corp., 508.482.8838, firstname.lastname@example.org
Beth Gillece-Castro is principal scientist at Waters Corp., 508.482.3007, email@example.com
Ziling Lu is applications chemist at Waters Corp., 508.482.4663, firstname.lastname@example.org
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