Direct detection of S-palmitoylation by mass spectrometry

Article abstract of DOI:10.1021/ac402850s

Direct detection and quantification of protein/peptide palmitoylation by mass spectrometry (MS) is a challenging task because of the tendency of palmitoyl loss during sample preparation and tandem MS analysis. In addition, the large difference in hydrophobicity between the palmitoyl peptides and their unmodified counterparts could prevent their simultaneous analysis in a single liquid chromatography-MS experiment. Here, the stability of palmitoylation in several model palmitoyl peptides under different incubation and fragmentation conditions was investigated. It was found that the usual trypsin digestion protocol using dithiothreitol as the reducing agent in ammonium bicarbonate buffer could result in significant palmitoyl losses. Instead, it is recommended that sample preparation be performed in neutral tris buffer with tris(2-carboxyethyl)phosphine as the reducing agent, conditions under which palmitoylation was largely preserved. For tandem MS analysis, collision-induced dissociation often led to facile palmitoyl loss, and electron capture dissociation frequently produced secondary side-chain losses remote from the backbone cleavage site, thus discouraging their use for accurate palmitoylation site determination. In contrast, the palmitoyl group was mostly preserved during electron transfer dissociation, which produced extensive inter-residue cleavage coverage, making it the ideal fragmentation method for palmitoyl peptide analysis. Finally, derivatization of the unmodified peptides with a perfluoroalkyl tag, N-[(3-perfluorooctyl)propyl] iodoacetamide, significantly increased their hydrophobicity, allowing them to be simultaneously analyzed with palmitoyl peptides for relative quantification of palmitoylation.

Full text of DOI:10.1021/ac402850s

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Article
Direct Detection of S-Palmitoylation by Mass Spectrometry
YUHUAN JI, Nancy Leymarie, Dagmar Haeussler, Markus
Michael Bachschmid, Catherine E Costello, and Cheng Lin
Anal. Chem., Just Accepted Manuscript • Publication Date (Web): 26 Nov 2013
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Analytical Chemistry
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Direct Detection of SꢀPalmitoylation by Mass Spectrometry
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Yuhuan Ji,1,2 Nancy Leymarie,1 Dagmar J. Haeussler,3 Marcus M. Bachschmid,3 Catherine E.
Costello,1,2 Cheng Lin1,2*
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1. Center for Biomedical Mass Spectrometry, 2. Department of Biochemistry, 3. Department of
Medicine, Boston University School of Medicine, Boston, MA 02118
* To whom correspondence should be addressed
Phone: +1 (617) 638 6705
Fax: +1 (617) 638 6761
Email: chenglin@bu.edu
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Abstract
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Direct detection and quantification of protein/peptide palmitoylation by mass spectrometry (MS) is a
challenging task because of the tendency of palmitoyl loss during sample preparation and tandem MS
analysis. In addition, the large difference in hydrophobicity between the palmitoyl peptides and their
unmodified counterparts could prevent their simultaneous analysis in a single liquid chromatographyꢀ
MS experiment. Here, the stability of palmitoylation in several model palmitoyl peptides under different
incubation and fragmentation conditions was investigated. It was found that the usual trypsin digestion
protocol using dithiothreitol as the reducing agent in ammonium bicarbonate buffer could result in
significant palmitoyl losses. Instead, it is recommended that sample preparation be performed in neutral
Tris buffer with tris(2ꢀcarboxyethyl)phosphine as the reducing agent, conditions under which
palmitoylation was largely preserved. For tandem MS analysis, collisionꢀinduced dissociation often led
to facile palmitoyl loss, and electron capture dissociation frequently produced secondary sideꢀchain
losses remote from the backbone cleavage site, thus discouraging their use for accurate palmitoylation
site determination. In contrast, the palmitoyl group was mostly preserved during electron transfer
dissociation, which produced extensive interꢀresidue cleavage coverage, making it the ideal
fragmentation method for palmitoyl peptide analysis. Finally, derivatization of the unmodified peptides
with a perfluoroalkyl tag, Nꢀ[(3ꢀperfluorooctyl)propyl] iodoacetamide, significantly increased their
hydrophobicity, allowing them to be simultaneously analyzed with palmitoyl peptides for relative
quantification of palmitoylation.
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Introduction
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SꢀPalmitoylation is a postꢀtranslational modification (PTM) which results from the attachment of
a palmitoyl group to a cysteine residue through a thioester linkage. Palmitoylation is reversible and
dynamic; the cycling between the palmitoylation and depalmitoylation states regulates several
intracellular events such as protein trafficking, sorting, tethering, and proteinꢀprotein interaction.1
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The regulatory role of the palmitoylation/depalmitoylation cycle has been implicated in many
studies. ElꢀHusseini et al. showed that PSDꢀ95, a major component of the postsynaptic density (PSD) at
glutamatergic synapses, is important for AMPA receptor trafficking. Pharmacological interruption of the
palmitoylation/depalmitoylation cycle of PSDꢀ95 results in a change in the number of AMPA receptors
at synapses, attenuating the synaptic signaling.2 Wedegaertner et al. demonstrated that agonist activation
of the β2ꢀadrenergic receptor induces increased palmitate turnover on Gαs.3 The depalmiltoylated Gαs
loses its function in signal transduction after its release from the plasma membrane to the cytosol.4
Robinson et al. found that bradykinin, a G proteinꢀcoupled receptor ligand, induces depalmitoylation
and translocation of endothelial nitric oxide synthase 3 (eNOS). They proposed that this regulation may
be important for NOꢀrelated signaling pathway in endothelial cells.5 Burgoyne et al. showed that bovine
aortic endothelial cells suffered apoptosis under oxidative stress, coincident with decrease in HꢀRas
palmitoylation, abnormal intracellular distribution of HꢀRas, and reduced survival signaling from HꢀRas.
They further proposed that direct competition from oxidative PTMs of the Cꢀterminal reactive cysteines
prevents HꢀRas palmitoylation, leading to endothelial dysfunction.6
In order to better understand the role of palmitoylation in biological systems, it is necessary to
develop analytical methods that can localize palmitoylation sites as well as quantitatively measure the
change in protein palmitoylation under various conditions. In early days, during decades of research,
radioactive labeling was used for detection of palmitoylation. Although radioactive labeling is effective
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for quantitation, the procedure is tedious, potentially toxic and time consuming.7 Moreover, it is
incapable of specifying the palmitoylation sites, unless mutations are made 8
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or antibodies that target
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regions of interest are used in conjunction with enzymatic digestion.9 A mass spectrometry (MS)
compatible method for palmitoylation analysis has recently been developed, which utilizes the acylꢀ
biotinyl exchange (ABE) chemistry to irreversibly replace the thioesterꢀlinked palmitoyl modifications
with stable biotin tags.10 This is achieved by blockage of all free thiol groups with Nꢀethylmaleimide
(NEM) followed by removal of the palmitoyl group from cysteines via hydroxylamineꢀinduced
hydrolysis, and labeling of the newly freed thiol groups with biotin. The biotinylated proteins are
affinity purified by streptavidinꢀagarose beads, enzymatically digested, and subjected to MS analysis.
One advantage of the ABE approach is the achievement of palmitoyl peptide/protein enrichment from
complex samples. However, false positives could arise due to incomplete blockage of free cysteines, and
false negatives could occur due to inadequate thioester hydrolysis by hydroxylamine, or inefficient
biotinꢀlabeling. Further, palmitoylation is not the only form of thioester linkages, and the ABE method
could lead to false assignments of palmitoylation when the cysteine is occupied by some other
modification.
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An alternative approach utilizing metabolic labeling with the palmitic acid analogue, 17ꢀ
octadecynoic acid (17ꢀODYA), and click chemistry has shown great promise in largeꢀscale profiling of
protein palmitoylation using cell lines.11 Click chemistry can be used to attach different functional
groups to the palmitoyl mimic, allowing enrichment and detection of palmitoyl proteins using both MS
and nonꢀMS methods. In conjunction with mass spectrometry, the metabolic labeling approach can be
used for identification of the palmitoylation sites, and characterization of the palmitoylation dynamics.12
However, it is difficult and costly to extend its use to study palmitoylation in large organisms. Moreover,
relative quantification of the protein palmitoylation remains challenging, as the unpalmitoylated proteins
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are not retained. As discussed below, care must also be taken to minimize loss of palmitoyl groups
during multiple reaction and sample cleanup steps.
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Direct detection of Sꢀacylation by MS has also been reported.13ꢀ18 This usually starts with the
purification of the proteins of interest, followed by reductive alkylation, and tryptic digestion either inꢀ
gel or inꢀsolution. MS analysis, typically using a matrixꢀassisted laser desorption/ionizationꢀtimeꢀofꢀ
flight (MALDIꢀTOF) or a MALDIꢀTOF/TOF instrument, can then be performed for palmitoyl peptide
identification. Such an approach is well suited for discovery of novel palmitoylation sites, but could be
problematic in quantitative studies, as palmitoyl loss may occur during sample preparation, because of
the labile thioester linkage. In particular, controversy exists over the stability of palmitoylation when
dithiothreitol (DTT) is used as the reducing agent.18ꢀ20 Additionally, palmitoyl groups may be lost,
through prompt or metastable fragmentation, during tandem MS analysis, leading to uncertainty in
palmitoylation site determination. Finally, the large difference in the hydrophobicity of the
palmitoylated and unpalmitoylated peptides makes relative quantification a challenging task.
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Here, the stability of palmitoylation in several palmitoyl peptide standards under various
experimental conditions was investigated in order to establish a sample preparation protocol that retains
palmitoylation for MS analysis. The potential of a derivatization strategy for relative quantification of
palmitoyl peptides and their unmodified counterparts was also explored. Lastly, fragmentation behaviors
of palmitoyl peptides under several dissociation modes were studied to evaluate their applicability for
characterization of palmitoyl peptides.
Experimental
Preparation of Palmitoyl Peptides
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The palmitoylation reaction was performed as previously reported21 with some modifications.
Each peptide standard (200 ꢁg) was allowed to react with 1 ꢁL of palmitoyl chloride in 10 ꢁL of 100%
TFA for 10 min at room temperature. The resulting mixture was dried under a nitrogen flow.
Purification of the products was performed on an Agilent 1200 series HPLC system (Agilent
Technologies, Santa Clara, CA) using a Vydac 214ms5215 column (C4, 5 ꢂm, 300 Å, 2.1 mm ID x 150
mm). Mobile phase A consisted of 95:5 water/ACN with 0.1% TFA and mobile phase B consisted of
85:10:5 ACN/IPA/water with 0.1% TFA. The sample was suspended in 400 ꢁL of 30% B (singly
palmitoylated peptides) or 40% B (doubly palmitoylated peptides), sonicated for 1 min, centrifuged for
10 min at 21,000 RCF. The supernatant was collected and centrifuged for another 10 min before HPLC
injection. A linear gradient of 30−100% B (singly palmitoylated peptides) or 40ꢀ100% B (doubly
palmitoylated peptides) over 20 min was employed with a flow rate of 0.3 mL/min. UV detection was
performed at 214 nm. The palmitoyl peptide fractions were collected, aliquoted, and dried. The amount
of palmitoyl peptide in each aliquot was determined by a micro BCA protein assay kit. Aliquots were
frozen at ꢀ80 °C for later use.
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Stability Test of Palmitoyl Peptides
Aliquots of HPLCꢀpurified palmitoyl peptides were incubated in 100 mM ABC (pH 8.0), 50 mM
Tris (pH 7.4), or 50 mM AA buffer (pH 4.0), with or without the presence of DTT or TCEP in different
concentrations, at either 37 oC or 55 oC. At several time points, a 0.5ꢀꢁL aliquot was taken and diluted in
5 ꢁL of 50% ACN/0.1% TFA. The diluted sample (0.5 ꢁL) was coꢀcrystallized with 0.5 ꢁL of DHB (10
ꢁg/ꢁL in 50% ACN/0.1% TFA) on a steel MALDI target plate, and later analyzed on either a Reflex IV
MALDIꢀTOF mass spectrometer (Bruker Daltonics, Billerica, MA) or an ultrafleXtreme MALDIꢀ
TOF/TOF mass spectrometer (Bruker Daltonics, Bremen, Germany) with 25~50% laser power. For each
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spectrum, 200~2000 laser shots were accumulated. The MALDIꢀTOF mass spectra were analyzed using
the FlexAnalysis 3.4 software.
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Preparation of IAMꢀ and FIAMꢀLabeled Peptides
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Alkylation with IAM was performed according to the manufacturer’s protocol.22 FIAMꢀlabeling
was performed following the protocol as described previously23 with slight modifications. The peptide
standard was first dissolved in 50 mM Tris buffer (pH 7.4) with 0.5 mM TCEP and kept at 37 °C for 30
min; the resulting solution was incubated with equal amount of 10 mM FIAM dissolved in THF at 37 °C
for 30 min in the dark. The mixture was dried under a nitrogen flow. Purification was performed using a
linear gradient of 30−100% B over 20 min at a flow rate of 0.3 mL/min. The amount of IAMꢀ and
FIAMꢀlabeled peptides in each fraction was determined using a micro BCA protein assay kit. Aliquots
were frozen at ꢀ80 °C for later use.
HPLC Separation
HPLC behavior of IAMꢀlabeled, FIAMꢀlabeled and palmitoyl peptides was investigated on an
Agilent 1200 series HPLC system using a Vydac 214ms5215 column with the same solvent system as
described above. The samples were introduced at 20% B and analyzed with a linear gradient of 20−100%
B over 20 min at a flow rate of 0.3 mL/min. Peaks were detected by the UV absorption at 214 nm, and
fractions were collected and analyzed by MALDIꢀTOF MS for identification.
LCꢀMS Quantification
Quantification was achieved by LCꢀMS on an LTQꢀOrbitrap XL instrument (Thermo Fisher
Scientific, San Jose, CA) equipped with a nanoACQUITY UPLC (Waters, Milford, MA) and a Triversa
Nanomate system (Advion Biosystems, Inc., Ithaca, NY). A Waters BEH300 C4 column (1.7 ꢂm, 150
ꢂm ID x 100 mm) was used for separation. Mobile phase A consisted of 95:5 water/ACN with 0.1% FA
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and mobile phase B consisted of 95:5 ACN/water with 0.1% FA. Palmitoyl peptides and FIAMꢀlabeled
peptides were combined at different ratios and introduced at 40% B with a flow rate of 0.5 ꢂL/min. The
gradient was held at 40% B for 20 min, followed by a ramp to 100% B over 30 min. It was then held at
100% B for 5 min, followed by a ramp to 40% B over 2 min, and maintained at 40% B for 23 min for
column reꢀequilibration. All mass spectra were acquired in the Orbitrap and analyzed by the Xcalibur
software.
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Tandem MS Analyses
Offꢀline tandem MS analyses were performed on a 12ꢀT solariX hybrid QhꢀFTICR mass
spectrometer (Bruker Daltonics, Bremen, Germany). HPLCꢀpurified palmitoyl peptides were dissolved
in 50:50 water/ACN with 0.1% FA to a concentration of 1 pmol/ꢂL and directly infused into the mass
spectrometer. Precursor ions of interest were isolated by the frontꢀend quadrupole and fragmented by
different dissociation methods. Collisionꢀinduced dissociation (CID) was performed with the collision
voltage set to 7 to 20 V; electron capture dissociation (ECD) was achieved with irradiation of ~1.5ꢀeV
electrons for 50 ms; electron transfer dissociation (ETD) was performed with a 400ꢀms reagent
accumulation and a 50 to 100ꢀms reaction time. A 1ꢀs transient was acquired for each scan and each
spectrum was the result of summing 100 transients. Fluoranthene anions were used as the reagent for
ETD.
Onꢀline LCꢀMS/MS analyses were performed on the LTQꢀOrbitrap XL mass spectrometer. Dataꢀ
dependent acquisition was performed by switching between one MS scan (r = 60,000 at m/z 400) and
three MS/MS events (r = 7,500). The three most abundant ions with charge state ≥ 2 were isolated with
a window of ±3 m/z for CID, highꢀenergy CID (HCD), and ETD (with fluoranthene anions). The
normalized collision energy was set at 35% for CID, and 30% for HCD. ETD reaction time was set at 80
ms with supplemental activation set at 15.
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Results and Discussion
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Stability of Palmitoyl Peptides
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Synthetic peptides PDFRIAFQELLCLR, MGCVQCKDKEA, and ARAWCQVAQKF were
chosen as model systems because their sequences contain palmitoylation motifs from proteins that are
known to undergo in vivo palmitoylation: betaꢀ2 adrenergic receptor,8 tyrosineꢀprotein kinase Fyn,24 and
glutamate decarboxylase 2, respectively.25 The chosen sequences do not contain serine, threonine, or
tyrosine residues, which are targets of Oꢀpalmitoylation that could complicate the analysis. The in vitro
palmitoylation was performed in 100% TFA, under which condition all basic groups, including the Nꢀ
terminal amine and the side chains of the lysine and arginine residues, were protonated and not
palmitoylated. Tandem MS analysis showed that palmitoylation occurred at the cysteine and tryptophan
residues, producing PDFRIAFQELLCpalmLR, MGCpalmVQCpalmKDKEA, and ARAWpalmCpalmQVAQKF.
Tryptophan palmitoylation likely occurred at the C2 position (Supplemental Scheme S1) via an
electrophilic substitution reaction. The stability of palmitoylation in various buffers was investigated by
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incubating HPLCꢀpurified palmitoyl peptides at 37 C for 1 hr, 3 hr, 6 hr, and 16 hr. The MALDIꢀTOF
mass spectra (Supplemental Figure S1) show that 6ꢀhr incubation of MGCpalmVQCpalmKDKEA in the
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standard tryptic digestion buffer (100 mM ABC, pH 8.0) at 37 C already led to significant palmitoyl
loss, whereas all three palmitoyl peptides were stable after overnight incubation in either the neutral
buffer (50 mM Tris, pH 7.4) or the acidic buffer (50 mM AA, pH 4.0). Further investigation revealed
that even incubation in the pH 7.4 ABC buffer resulted in complete depalmitoylation within 16 hours,
and this was attributed to the gradual pH increase of the ABC buffer (Supplemental Figure S2),
presumably because CO2 is more volatile than NH3. Since most proteases used in proteomics research
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attain their highest enzymatic activities at or near physiological pH, Tris buffer was used in the
following studies.
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To test the effect of the reducing agents on the palmitoyl stability, palmitoyl peptide standards
were incubated in Tris buffer in the presence of DTT or TCEP at 37 oC or at 55 oC for 30 min and 1 hr.
The results are summarized in Figures 1 and S3, which show the relative abundances of the palmitoyl
peptides and their depalmitoylated forms as a function of the incubation time and temperature, with DTT
or TECP. DTT, a reducing agent commonly used for reduction of disulfide bonds and other reversible
oxidative modifications of cysteines, accelerated the depalmitoylation process for all three peptides, but
the other widelyꢀused reducing reagent, TCEP, did not cause appreciable loss of palmitoylation. Further,
the depalmitoylation rate was significantly increased at higher temperature, as one would expect for any
reaction with an activation barrier.
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The mechanisms for disulfide bond reduction by DTT and by TCEP are illustrated in
Supplemental Scheme S2. For DTT, disulfide reduction proceeds via sequential thiolꢀdisulfide exchange
reactions, where one of the two disulfide sulfur atoms is attacked by a thiolate group of DTT, releasing
one cysteine residue and creating a mixed disulfide species; the subsequent nucleophilic attack by the
remaining thiolate of DTT releases the other cysteine and forms cyclic oxidized DTT. Although thiolꢀ
disulfide exchanges are reversible, disulfide bond reduction by DTT is quite unidirectional, as the
byproduct, the oxidized DTT, is highly stable due to its 6ꢀmembered ring structure. DTT is reactive only
at pH >7 since the negatively charged thiolate is the only reactive form. Furthermore, DTT must be used
in at least 20ꢀfold molar excess to the disulfide bond to ensure complete reduction.26 However, thiolate
is a strong nucleophile that can also attack the carbonyl of the thioester linkage in Sꢀpalmitoylated
peptides, resulting in disruption of the palmitoylation, as observed here.
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The reactive group on TCEP is the free electron pair on the phosphine group, which can attack
the disulfide bond, releasing one cysteine and forming an intermediate phosphorusꢀsulfur linkage. The
subsequent nucleophilic attack to the positively charged phosphorus by the oxygen of a water molecule
releases the second cysteine, and forms a phosphine oxide. This reaction is irreversible because of the
formation of a strong phosphorusꢀoxygen double bond in the oxidized TCEP27 and proceeds at nearly
stoichiometric ratio. TCEP is a potent reducing agent, and works under a broad pH range since
phosphine is a very weak base and retains its nucleophilic character even in acidic solutions. In addition,
phosphine does not react with a thioester, making TCEP an ideal reducing agent for studying Sꢀ
palmitoylation.
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Analysis of Palmitoyl Peptides by RPꢀHPLC
Several HPLC methods have been developed to analyze very hydrophobic peptides28 and
peptides with lipid modifications.29,30 Due to their dramatic difference in hydrophobicity, it is
challenging to analyze palmitoyl peptides and their unmodified counterparts in a single HPLC run.
While it is possible to analyze lipid modified peptides on a C18 column, it requires the use of strong
organic solvent systems. For example, Gustafsson et al. reported that SPꢀC, a 35ꢀresidue palmitoyl
peptide, can be separated on a C18 column by using 60~75% methanol/ethanol as the initial solvent
followed by elution with isopropanol,30 under which conditions most unmodified peptides cannot be
retained. Meanwhile, C4 columns retain palmitoyl peptides through weaker interactions, allowing
separation and elution of palmitoyl peptides using mild organic solvents such as acetonitrile. However,
C4 columns are not suitable for analysis of unmodified peptides which are usually not retained. Figure
2a shows the chromatogram of a mixture of the three palmitoyl peptide standards and their IAMꢀlabeled
counterparts, acquired on a C4 column with the gradient program shown in the inset. Although all three
palmitoyl peptides were retained and well separated, two of the three IAMꢀlabeled peptides flowed
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through with the initial solvent. One way to overcome this difficulty is to increase the hydrophobicity of
unmodified peptides through derivatization. FIAM, with its structure shown in Supplemental Scheme S3,
is a cysteine alkylation reagent with a hydrophobic perfluoroalkyl moiety, that our laboratory has
demonstrated to be useful for enrichment of native cysteineꢀcontaining peptides and the RPꢀHPLC
analysis of the mixtures of these products with their irreversibly modified analogs.23 For the quantitative
studies undertaken herein, we rationalized that this selective hydrophobic FIAM labeling of the cysteine
thiol group in the unmodified peptides would allow them to be analyzed simultaneously with the
palmitoyl peptides. Figure 2b shows the chromatogram of a mixture of the palmitoyl peptides and their
FIAMꢀlabeled counterparts, acquired on a C4 column using the same gradient program as shown in
Figure 2a. All six peptides were retained on the C4 column in the initial phase, and were well resolved
by gradient elution with mild organic solvent.
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Relative Quantification of Palmitoyl Peptides by Onꢀline LCꢀMS
Siteꢀspecific quantification of protein palmitoylation can be achieved by utilizing either the ABE
chemistry or metabolic labeling. However, these methods are not suitable for relative quantification of
palmitoyl peptides with respect to their unpalmitoylated forms, as the unpalmitoylated peptides are
usually not retained under the HPLC conditions employed. As demonstrated above, with FIAMꢀlabeling,
all the cysteineꢀcontaining peptides, both with and without palmitoylation, can be analyzed in a single
LC run. Here, an onꢀline LCꢀMS study of palmitoyl and FIAMꢀderivatized peptide mixtures was
performed on an LTQꢀOrbitrap XL mass spectrometer using the Waters BEH300 C4 column to explore
the potential of using extracted ion chromatograms (EICs) for the relative quantification of peptide
palmitoylation.
Figure 3a shows the total ion chromatogram of a mixture of palmitoyl and FIAMꢀlabeled peptide
standards containing equal amounts of each peptide. The six peptides were well separated from one
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another and eluted between 40% and 100% B. Note that the differences in the peak areas observed for
these peptides likely results from differences in their ionization efficiencies. LCꢀMS analyses were then
performed on mixtures of these peptides at seven different ratios (FIAM/palm = 10:1, 5:1, 2:1, 1:1, 1:2,
1:5, and 1:10). The ion abundance of each peptide was calculated as the sum of the chargeꢀstate
normalized integrated peak areas of all observed charge states in their respective EICs using the
Xcalibur Quan browser. The calibration curve for relative quantification was generated by plotting the
ratio of the calculated ion abundance as a function of the ratio of the amount of peptides injected. As
shown in Figures 3bꢀd, for all three pairs of palmitoyl and FIAMꢀlabeled peptides, a satisfying linear
correlation exists between these two ratios, thus establishing the validity of relative quantification based
on the integrated EIC peak areas of the palmitoyl and FIAMꢀderivatized peptides. It is important to note
that all samples must be analyzed under the same LC conditions as those used to generate the calibration
curve. A change in the column, solvent system, and/or gradient program would lead to changes in the
solvent composition during elution of peptides of interest, and consequently, their ionization efficiencies.
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Tandem MS Analysis of Palmitoyl Peptides
Tandem mass spectrometry is a powerful tool for identification and localization of PTMs, and
has been applied to characterize lipidꢀmodified peptides. 23, 31ꢀ33 In general, the thioether linkage is stable
under lowꢀenergy CID conditions, as illustrated by the retention of the perfluoroalkyl group in most CID
fragment ions from the FIAMꢀlabeled peptides.23 Hoffman and Kast reported a CID study of peptides
with various lipid modifications, including Nꢀmyristoylation, farnesylation, and Sꢀpalmitoylation.31
Whereas Nꢀmyristoylation was fairly stable under CID, Sꢀpalmitoylation was labile upon collisional
activation, producing abundant b – 238 ions as the result of the thioester bond breakage either before or
after the backbone amide cleavage. On the other hand, labile modifications are often preserved under
ECD, which is considered nonꢀergodic and directional towards the backbone NꢀCα bond cleavage. Guan
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studied the CID and ECD fragmentation behaviors of ghrelin,32 a peptide with Oꢀacylation, and found
that ECD produced far more extensive backbone fragmentation without breaking the ester bond, while
the esterꢀlinked octanoyl group was lost in many CID fragments. Kaczorowska et al. recently studied the
CID and ECD fragmentation behaviors of Sꢀdipalmitoylated peptides, and found that both CID and ECD
could provide structural information on the peptide sequence and the modification.33 However, in the Sꢀ
dipalmitoylated peptides reported in this study, the palmitoyl groups were not directly attached to the
cysteine thiol group, but were connected via ester linkages to a glycerol, which was linked to the
cysteine via a thioether bond. Since the thioester linkage is more labile than the ester linkage, we felt
that it would be important to study the fragmentation behaviors of Sꢀacylated peptides under different
dissociation conditions. A systematic investigation of the fragmentation behaviors of Sꢀpalmitoyl
peptides was performed here with CID, ECD, and ETD as the dissociation modes. Tandem mass spectra
were acquired either onꢀline on the LTQꢀOrbitrap instrument (CID, HCD, and ETD), or offꢀline on the
hybrid QhꢀFTICR instrument (CID, ECD, and ETD). Representative tandem mass spectra of the three
palmitoyl peptide standards and their corresponding cleavage maps are shown in Figures 4, 5, and S4,
respectively.
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For the triply charged peptide, PDFRIAFQELLCpalmLR, lowꢀenergy CID spectra acquired on
both instruments provided complete interꢀresidue cleavage coverage, generating abundant bꢀ and yꢀtype
ions, as well as a few aꢀions (Figure 4a, b). Some yꢀions, labeled with an asterisk (y*), were also
observed with loss of the palmitoyl group (C16H30O, 238.23 Da). In general, y* ions were in much lower
abundance than normal yꢀions, and should not have a significant impact on palmitoylation site
determination. Similar sequence coverage was obtained by HCD (Figure 4c). However, with its higher
energy input, HCD also resulted in increased palmitoyl losses from many yꢀions. Nonetheless, the
palmitoylation site could still be localized to the cysteine residue based on the presence of a highꢀ
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abundance y3 ion with the palmitoyl group attached (m/z 629.44) and absence of any smaller palmitoyl
groupꢀcarrying yꢀions.
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The ECD spectrum (Figure 4d) of the same precursor ion was characterized by extensive cꢀ and
zꢀ ion series, providing complete interꢀresidue cleavage coverage. Surprisingly, ECD also produced
many z ions with loss of a palmitoyl group, labeled as z* ions, some of which were in very high
abundance. Considering that insignificant palmitoyl loss was observed in the CID spectra of this peptide,
formation of these z* ions was unlikely an ergodic process. It has been previously reported that the alpha
carbon radical formed upon NꢀCα bond cleavage in ECD can initiate further backbone and/or sideꢀchain
cleavages, with or without radical migration.34ꢀ36 Supplemental Scheme S4A illustrates a possible
mechanism for the z* ion formation, in which the backbone alpha carbon radical abstracts a hydrogen
from the alpha carbon of the palmitoyl group, and the subsequent radicalꢀinduced alpha cleavage
releases a tetradecylketene (C16H30O, 238.23), leaving the radical on the sulfur atom of the cysteine
residue. Palmitoyl loss here was driven by the stability of the sulfurꢀcentered radical formed, whereas in
the case of Oꢀacylation, no acyl loss was observed because it would have required the unfavorable
formation of an oxygenꢀcentered radical.32 The ECD spectrum also contains another series of ions,
labeled as z§ ions, which correspond to z ions with partial loss of the cysteine sideꢀchain (C15H31COS•,
271.21 Da). The z§ ions were likely formed as the result of radical induced alpha cleavage following
radical migration to the alpha carbon of the cysteine residue (Supplemental Scheme S4B). Loss of the
C15H31COS• group was so energetically favored, once the radical was formed at the alpha carbon of the
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*
cysteine residue, that neither the z3 nor the z3 ion was observed in the ECD spectrum. The NꢀCα bond
cleavage Nꢀterminal to the palmitoylated cysteine residue was always followed by the partial cysteine
§
side chain loss, leading to the formation of an abundant z3 ion. Overall, we found that ECD generated
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too many siteꢀnonspecific side chain loss product ions as a result of radical migration to be useful for
palmitoylation site determination.
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Similar to ECD, ETD of the triply charged PDFRIAFQELLCpalmLR precursor also produced
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extensive cꢀ and zꢀ ion series, with complete interꢀresidue cleavage coverage. However, unlike ECD, no
§
palmitoyl loss was observed in any of the z ions formed. Moreover, only one z§ ion was detected: a z3
§
ion. Note that the z3 ion is essentially the w3 ion, and its formation did not require radical migration. It
appears that radical migration was suppressed in ETD, either due to its lower energy input, or because of
radical stabilization by collisional cooling. The palmitoylation site could be confidently assigned to the
cysteine residue based on the mass difference between the c11 and c12 ions. The absence of the z3 ion and
the presence of the w3 ion provided additional evidence for cysteine palmitoylation.
The tandem MS fragmentation behavior of the doubly palmitoylated peptide,
MGCpalmVQCpalmKDKEA, was similar to that of the PDFRIAFQELLCpalmLR peptide, but also showed
some distinct differences. For the triply charged MGCpalmVQCpalmKDKEA precursor ion, lowꢀenergy
CID (Figure 5a) produced only a limited number of bꢀ and yꢀ type ions, and the palmitoylation site at the
second cysteine residue could not be determined because no backbone cleavage at either side of that
cysteine residue was observed. Although its HCD spectrum (Figure 5b) was more informative, and
provided 100% cleavage coverage, most palmitoylꢀcontaining fragment ions also underwent extensive
palmitoyl loss during the fragmentation process, including some with loss of two palmitoyl groups, e.g.
**
the y10 ion, thus preventing reliable palmitoylation site localization. For this peptide, unlike in the
previous case, ECD (Figure 5c) produced complete cleavage coverage with minimum palmitoyl loss.
Note that the PDFRIAFQELLCpalmLR peptide contains polar residues (D, E, and R) near both termini,
allowing it to adopt a folded gasꢀphase conformation, in which the palmitoylꢀcysteine residue was close
to several potential backbone cleavage sites. Upon the NꢀCα bond cleavage by ECD, the radical could
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migrate from the initial Cα position to the spatially adjacent palmitoylꢀcysteine residue even if it was
distant in sequence, initiating further side chain losses and formation of z§ and z* ions. Such spatial
proximity may not be present in the MGCpalmVQCpalmKDKEA peptide, because all of its polar residues
(K, D, and E) were located on one side of the palmitoylꢀcysteine residues, and the two bulky
hydrophobic palmitoylꢀcysteine side chains further prevented formation of a compact structure.
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§
§
Consequently, radical migration was not prevalent here, resulting in only two lowꢀabundance z7 and z8
ions. As in the previous case, z ions with an Nꢀterminal cysteine, z6 and z9, were not observed, instead,
§
§
cleavages Nꢀterminal to the two cysteine residues led to the formation of the z6 (w6) and z9 (w9) ions,
which could be used to confirm the two cysteine palmitoylation sites. The ETD spectrum of
MGCpalmVQCpalmKDKEA (Figure 5d) was very similar to its ECD spectrum, producing a complete
series of NꢀCα bond cleavages, with no palmitoyl loss. Partial cysteine side chain loss was only observed
§
for z ions with an Nꢀterminal cysteine. Interestingly, two c ions with partial cysteine side chain loss, c9
§
and c10 , were also observed. As c ions are even electron species, radical migration was unlikely to play
§
a role here. Since both c9 and c10§ ions were singly charged ions produced from triply charged precursor
ions with all three potential protonation sites residing within the first nine residues, these two ions were
likely formed as a result of two electron transfer processes, with one accounting for the cysteine side
chain loss, and the other leading to the backbone NꢀCα bond cleavage.
The general features of the tandem mass spectra (Figure S4) of the triply charged peptide,
ARAWpalmCpalmQVAQKF, resemble those of the other two peptides. Abundant palmitoyl losses were
observed in its lowꢀenergy CID (Figure S4a) and HCD spectra (Figure S4b). For ECD (Figure S4c), in
addition to the palmitoyl loss and cysteine side chain loss, abundant tryptophan side chain loss remote
from the ECD cleavage site was observed, in agreement with findings from a previous study.37 Again,
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the ETD spectrum (Figure S4d) was the most informative with minimal secondary side chain losses,
except for the characteristic cysteine side chain loss at the cysteine cleavage site.
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Conclusions
Sꢀpalmitoylation is a labile modification both in solution and in the gas phase. In order to
minimize palmitoyl loss during proteomic sample preparation, it is recommended that palmitoyl (and
other acylated) proteins and peptides be processed under neutral or slightly acidic conditions and at
room temperature. Use of DTT should be avoided; instead, TCEP is the preferred disulfide reducing
agent for palmitoyl protein analysis. The drastic difference in hydrophobicity between palmitoyl
peptides and their unpalmitoylated forms can be reduced by derivatization of free cysteine residues with
a perfluoroalkyl tag, enabling relative quantification of palmitoylation by LCꢀMS. CID of Sꢀ
palmitoylated peptides can lead to facile loss of palmitoyl groups, whereas ECD can result in extensive
radical migration and secondary side chain loss driven by the formation of stable sulfurꢀcentered radicals.
ETD appeared to be the best fragmentation method for tandem MS analysis of palmitoyl peptides
because it produced extensive backbone fragmentation with minimum palmitoyl loss. In summary, this
study presents a comprehensive strategy, including sample preparation, LCꢀMS and tandem MS analysis,
for direct detection and quantification of Sꢀpalmitoyl peptides.
Acknowledgements
This research is supported by the NIH grants P41 RR10888/GM104603, P01 HL068758, R37
HL104017, S10 RR015942, S10 RR025082, S10 RR020946, S10 OD010724, and NIH/NHLBI contract
HHSN268201000031C.
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Figures and legends
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Figure 1. Stability of three palmitoyl peptides, as represented by the relative abundance of the palmitoyl
peptides and their various depalmitoylated forms after 30 min or 60 min of incubation in Tris buffer (50 mM, pH
7.4) at 37 oC or at 55 oC, in the presence or absence of DTT or TCEP.
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Figure 2. (a) Chromatogram of a mixture of three palmitoyl
peptides and their IAM-labeled counterparts; (b)
Chromatogram of a mixture of three palmitoyl peptides and
their FIAM-labeled counterparts.
Figure 3. Relative quantification by UPLC-MS analysis on an
LTQ-Orbitrap XL mass spectrometer. (a) The total ion
chromatogram of a mixture of palmitoyl and FIAM-labeled
peptides; (b), (c), and (d) the relative quantification
calibration curves of the PDFRIAFQELLCFIAMLR and
PDFRIAFQELLCpalmLR peptide pair, the MGCFIAMVQCFIAMKDKEA
and MGCpalmVQCpalmKDKEA peptide pair, and the
ARAWCFIAMQVAQKF and ARAWpalmCpalmQVAQKF peptide pair,
respectively.
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Figure 4. Tandem MS spectra of PDFRIAFQELLCpalmLR: (a) CID on solariX, (b) CID on Orbitrap, (c) HCD on Orbitrap,
(d) ECD on solariX, and (e) ETD on solariX. * indicates loss of C16H30O (238.23 Da); § indicates loss of C15H31COS•
(271.21 Da).
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Figure 5. Tandem MS spectra of MGCpalmVQCpalmKDKEA: (a) CID on Orbitrap, (b) HCD on Orbitrap, (c) ECD
on solariX, (d) ETD on Orbitrap. * indicates loss of C16H30O (238.23 Da); § indicates loss of C15H31COS• (271.21
Da).
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