Source pressure. The transmission of ions into the mass spectrometer, and the maintenance of
tertiary and quaternary protein structure, is heavily influenced by the so-called “backing”
pressure in the source region of the instrument (Fig. 14.18). An increased source pressure
results in collisional cooling and collisional focusing of ions, and can aid in the transmission of
ions with high m/z. The optimal source pressure should be determined empirically for each
sample, and can be adjusted using the isolation valve (Speedivalve) of the roughing pump, which
changes the conductance of the vacuum line between the source and the pump. In our experience, source pressures in the region of 3–5 mbar were required to observe well-resolved spectra
of APol-solubilized MPs. Pressures up to ~8 mbar can be achieved on the instrument, but care
should be taken when adjusting the Speedivalve at such pressures to ensure the instrument does
not automatically vent.
Sample cone and extraction cone. Increased cone voltages (sample and extraction cone,
Fig. 14.18) can improve desolvation (leading to increases in signal-to-noise and mass accuracy)
and the transmission of high m/z ions. Additionally, increasing the cone voltages imparts higher
internal energies on analyte ions. This can be considered a double-edged sword for MP
analysis, as increased energy can aid the dissociation of APols and ejection of “naked” MPs,
but also potentially lead to gas phase unfolding and charge stripping of the protein. Unfolding
can be discerned by analysis of the charge state distribution observed (more highly charged ions
are more unfolded) or the CCS values of the ion species observed (increased CCSs indicate
protein unfolding) (Benesch 2009). Cone voltages should be optimized to ensure optimal ion
transmission and APol dissociation, without resulting in protein unfolding. In our experience,
sampling cone voltages >100 V are required to observe APol-trapped MPs. The extraction cone
should be set between 0 and 10 V.
Trap DC bias. The voltages in the IM sector of the instrument can also have a significant
influence on ion heating. Again this can be considered a double-edged sword, as described
above for the cone voltages. The trap DC bias voltage, the voltage which accelerates ions into
the higher-pressure IM sector of the instrument from the lower-pressure trap cell (Fig. 14.18), is
particularly important to optimize. For soluble proteins, the trap DC bias is typically set to 20 V
for optimal transmission without inducing unfolding. In our experience, voltages of 80–100 V
are required to observe APol-trapped MPs.
Trap/transfer collision voltages. The trap and transfer sectors of the Synapt HDMS
(Fig. 14.18) are filled with an inert buffer gas, typically argon. Analyte ions are accelerated
into the trap/transfer cells and undergo collisions with the buffer gas. This can be used to
dissociate the MP from the APol. The trap and transfer collision voltages should be optimized
empirically for each sample, but typically the trap collision voltage was set to 50–100 V, while
the transfer collision voltage was set to 10–150 V.
IM parameters. IM separates ions based on their mass, shape, and charge. More extended
ions experience more collisions with the buffer gas (in our case nitrogen, but helium can also be
used; see Bush et al. 2010), relative to more compact ions, and therefore exit the ion mobility cell
more slowly. The IM sector of the Synapt HDMS comprises the trap collision cell, ion mobility
cell, and transfer collision cell, each of which is a stacked-ring ion guide (Fig. 14.18). Ion
packets are pulsed periodically into the central ion mobility cell from the trap for IM separation.
The mobility (drift time) of each ion is recorded, and the value of m/z is determined by time-offlight (TOF) analysis. For each ion species, an arrival time distribution (ATD) can be extracted
from the three-dimensional m/z, drift time, and intensity data set (Fig. 14.19). The width of ATDs
of a protein or protein assembly can be used to study conformational heterogeneity and
conformational changes (Konijnenberg et al. 2014; Zhou et al. 2014; Martens et al. 2016;
Schiffrin et al. 2016). Because each conformation presents a Gaussian distribution, ATDs
652
14 The Use of Amphipols in Mass Spectrometry
tertiary and quaternary protein structure, is heavily influenced by the so-called “backing”
pressure in the source region of the instrument (Fig. 14.18). An increased source pressure
results in collisional cooling and collisional focusing of ions, and can aid in the transmission of
ions with high m/z. The optimal source pressure should be determined empirically for each
sample, and can be adjusted using the isolation valve (Speedivalve) of the roughing pump, which
changes the conductance of the vacuum line between the source and the pump. In our experience, source pressures in the region of 3–5 mbar were required to observe well-resolved spectra
of APol-solubilized MPs. Pressures up to ~8 mbar can be achieved on the instrument, but care
should be taken when adjusting the Speedivalve at such pressures to ensure the instrument does
not automatically vent.
Sample cone and extraction cone. Increased cone voltages (sample and extraction cone,
Fig. 14.18) can improve desolvation (leading to increases in signal-to-noise and mass accuracy)
and the transmission of high m/z ions. Additionally, increasing the cone voltages imparts higher
internal energies on analyte ions. This can be considered a double-edged sword for MP
analysis, as increased energy can aid the dissociation of APols and ejection of “naked” MPs,
but also potentially lead to gas phase unfolding and charge stripping of the protein. Unfolding
can be discerned by analysis of the charge state distribution observed (more highly charged ions
are more unfolded) or the CCS values of the ion species observed (increased CCSs indicate
protein unfolding) (Benesch 2009). Cone voltages should be optimized to ensure optimal ion
transmission and APol dissociation, without resulting in protein unfolding. In our experience,
sampling cone voltages >100 V are required to observe APol-trapped MPs. The extraction cone
should be set between 0 and 10 V.
Trap DC bias. The voltages in the IM sector of the instrument can also have a significant
influence on ion heating. Again this can be considered a double-edged sword, as described
above for the cone voltages. The trap DC bias voltage, the voltage which accelerates ions into
the higher-pressure IM sector of the instrument from the lower-pressure trap cell (Fig. 14.18), is
particularly important to optimize. For soluble proteins, the trap DC bias is typically set to 20 V
for optimal transmission without inducing unfolding. In our experience, voltages of 80–100 V
are required to observe APol-trapped MPs.
Trap/transfer collision voltages. The trap and transfer sectors of the Synapt HDMS
(Fig. 14.18) are filled with an inert buffer gas, typically argon. Analyte ions are accelerated
into the trap/transfer cells and undergo collisions with the buffer gas. This can be used to
dissociate the MP from the APol. The trap and transfer collision voltages should be optimized
empirically for each sample, but typically the trap collision voltage was set to 50–100 V, while
the transfer collision voltage was set to 10–150 V.
IM parameters. IM separates ions based on their mass, shape, and charge. More extended
ions experience more collisions with the buffer gas (in our case nitrogen, but helium can also be
used; see Bush et al. 2010), relative to more compact ions, and therefore exit the ion mobility cell
more slowly. The IM sector of the Synapt HDMS comprises the trap collision cell, ion mobility
cell, and transfer collision cell, each of which is a stacked-ring ion guide (Fig. 14.18). Ion
packets are pulsed periodically into the central ion mobility cell from the trap for IM separation.
The mobility (drift time) of each ion is recorded, and the value of m/z is determined by time-offlight (TOF) analysis. For each ion species, an arrival time distribution (ATD) can be extracted
from the three-dimensional m/z, drift time, and intensity data set (Fig. 14.19). The width of ATDs
of a protein or protein assembly can be used to study conformational heterogeneity and
conformational changes (Konijnenberg et al. 2014; Zhou et al. 2014; Martens et al. 2016;
Schiffrin et al. 2016). Because each conformation presents a Gaussian distribution, ATDs
652
14 The Use of Amphipols in Mass Spectrometry