Aldehyde Complexes with Protonated Peptides in the Gas Phase

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Aldehyde Complexes with Protonated Peptides in the Gas Phase Xiangguo Shi,†,§ Jianhua Ren,‡ and Joel H. Parks*,† † ‡

Rowland Institute at Harvard, 100 Edwin H. Land Boulevard, Cambridge, Massachusetts 02142, United States University of the Pacific, 3601 Pacific Avenue, Stockton, California 95211, United States

bS Supporting Information ABSTRACT: This Article presents a study of aldehyde complexes with peptide ions formed by bimolecular collisions in the gas phase. Desolvated ions generated by electrospray ionization are stored within a radio frequency (RF) ion trap and exposed to aldehyde vapor. Mass spectrometry measurements were performed on the resulting aldehyde complexes formed with single amino acids (LysH+, HisH+, and ArgH+) and polypeptides [Pron-Lys+2H]2+ and [(Gly-Ser)m-Lys+2H]2+. These data identify several interesting and unexpected aspects of the aldehyde complex kinetics. It is observed that the formation of stable complexes requires the presence of water vapor. The formation kinetics of aldehydepeptide complexes exhibits multiexponential time dependence that is modeled by interactions in the presence of structural heterogeneity. Aldehyde binding appears to involve a competition between conformers with unhindered access to protonation sites and conformers with intramolecular solvation of these sites. Proton transfer to the aldehyde ligand is responsible for the loss of the complexes. This is supported by proton affinity calculations and identified by reaction products exhibiting loss of protonation by the parent ion accompanied by the appearance of aldehyde cations.

1. INTRODUCTION Evidence presented by Williams and coworkers,1 Clemmer and coworkers,2,3 and Jarrold4 indicates that an ensemble of gasphase biomolecular ions can exhibit structural heterogeneity expressed by a distribution of conformations. Although the relationship between gas-phase and solution-phase conformations is not well understood, distinct correlations are found in electrospray mass spectrometry measurements (ESI MS), suggesting that gas-phase biomolecular ions retain a “memory” of their solution-phase conformation. The relationship of gas-phase structure to the solution environment implies that biomolecules in the gas phase are not characterized by a common structure.1 Consequently, the structural heterogeneity observed in gasphase measurements of cytochrome c hydrogen/deuterium exchange reactions by McLafferty and coworkers5 and gas-phase ion mobility measurements by Jarrold and coworkers4 and Clemmer and coworkers6 are consistent with the structural heterogeneity identified in the solution folding kinetics of cytochrome c measured in time-resolved ESI MS by Konermann and coworkers.7 Additional measurements of structural heterogeneity arising in solution phase experiments are presented in reviews of protein folding kinetics8 and conformational dynamics.9 There have been several recent studies10 of protein complexes in the gas phase exhibiting the presence of structural heterogeneity. Measurements by Williams and coworkers10a investigated the binding sites of water molecules to protonated Phe and derivatives by infrared photodissociation (IRPD). Multiexponential decays observed in kinetics data characterized different r 2011 American Chemical Society

binding sites of competing structural isomers by determining the relative populations of ions with water molecules attached at the various binding sites. Klassen and coworkers10b performed timeresolved blackbody infrared radiative dissociation (BIRD) measurements of the dissociation kinetics of proteinfatty acid complexes. Multiexponential decays of the dissociating complexes identified multiple noninterconverting structures. In both of these experiments, the complexes are initially formed in the electrospray solution, and the dissociation process is then performed within a Fourier transform ion cyclotron resonance (FTICR) mass spectrometer. It is assumed that the complex soformed includes the lower energy conformations of the biomolecule within the solvent. After dissociation, it is not certain that the final biomolecule states are low-energy conformations of the bare biomolecular ion. This Article describes the interactions of aldehydes, CH3(CH2)nCHO, with trapped protonated biomolecules within a radio frequency (RF) ion trap.11 In this case, complexes can be formed on lower energies of the potential energy surface corresponding to conformations of the bare biomolecular ion. Complexes formed by aldehydes12 with isolated gas-phase biomolecular ions offer the capability to investigate the effects of polymeric environments on biomolecule structure and dynamics. It is important to consider the possibility of relating such data to the conformation and dynamics of proteins that reside Received: December 18, 2010 Revised: July 20, 2011 Published: August 11, 2011 11183

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within cell membranes. The present experiments measure the mass spectra and kinetics of aldehydepeptide complexes with single amino acids and peptide sequences. In each biomolecular species, aldehyde binding appears to involve a competition between conformers with unhindered access to protonation sites and conformers with intramolecular solvation of charge sites. These measurements demonstrate that the kinetics of aldehydebinding displays the presence of structural heterogeneity consistent with results obtained in the broad array of gas-phase studies of biomolecular ions.

2. EXPERIMENTAL AND COMPUTATIONAL METHODS 2.1. Trapped Ion Mass Spectrometry. Details of the RF trap instrumentation have been published elsewhere11 and will be briefly summarized here. Experiments are performed on biomolecules stored in a home-built quadrupole ion trap by exposing the ions to neutral gas-phase molecules carried into the trap with a background helium gas. Biomolecule ions were formed in the gas phase by nanoelectrospray using a 1:1 acetonitrile/water solution. The hydrated ions were desolvated in a stainless-steel capillary held at 60 °C and passed through an octupole ion guide into a quadrupole ion trap. The desolvated ions of interest were isolated by stored waveform inverse Fourier transform (SWIFT) ion excitation. The trap background helium gas pressure was pulsed for ∼1 to 2 s at ∼2 103 Torr for ion loading at qz = 0.50 and thermal relaxation to the trap temperature of 298 K. A constant helium pressure was maintained at ∼3 105 Torr during ion storage and exposure to the aldehyde species. The helium gas entered the trap through an aperture in the ring electrode. After exposure to reactant vapors, ions were ejected, and a mass spectrum was obtained using an electron channeltron multiplier. In these experiments an independent gas handling system formed a water seeded helium flow by bubbling helium through a glass water cell, and this seeded flow was combined with the background helium flow during exposure. The trap design provided low conductance paths for gas flow out of the trap volume, and we estimated from residual gas analyzer measurements that the water vapor pressure inside the trap was e104 Torr. During exposure, the helium pressure in the trap was maintained at or near the background pressure by controlling the flow with a leak valve (Granville-Philips 203). Mass spectra obtained after exposure of trapped peptide ions to the water-seeded helium flow indicated the presence of aldehydepeptide complexes. Aldehydes, CH3(CH2)nCHO, are a ubiquitous contaminant in the neutral gas handling apparatus arising from forepump oil vapor13 condensed in the foreline oil trap. This vapor is present at sufficiently low levels that it is not detectable by the residual gas analyzer. In our gas handling apparatus, the aldehyde vapor was most likely entrained in the helium flow and admitted directly into the trap mixed with the helium and water vapor flow. The average aldehyde pressure inside the trap was estimated by leak valve settings to be e107 Torr, and this was consistent with the measured kinetic rates; however, this value was only a rough order of magnitude estimate. Kinetic measurements of reaction rates were obtained by varying the time delay between exposure of trapped ions to aldehyde and water vapors and ejection of all ions to obtain a mass spectrum. Depending on the rates, 1316 different exposure times were used in each run covering the range 0.590 s.

Figure 1. Mass spectrum displaying the Lys+ parent ion, (M+H)+, and a band of aldehydeLys complexes denoted by (M+H+2H2O+Aldn)+. The aldehyde species are indicated by n, the number of (CH2) methylene groups. The mass spectra were obtained for a 10 s exposure to aldehyde vapor at 300 K.

The data indicate no significant loss of trapped ions during exposures, and ion ejection occurs without significant loss of the ion complexes. One of the more intriguing aspects of the complex formation process is that stable complexes occur when water vapor and aldehyde species are present simultaneously during the aldehyde interaction with biomolecules. If the helium flow is admitted into the trap without the water vapor, then complex formation is below the noise level. 2.2. Materials. Peptides were commercially synthesized (BioMer Technology, Concord, CA) and purified by reversedphase HPLC to a stated purity of >70% prior to shipment. Protein and aldehyde samples were obtained from Sigma. Electrospray solutions contained peptides at 10 μM in 50% acetonitrile/50% water. Acetonitrile (>99.9%) and distilled, deionized water were obtained from Fisher Scientific (Fair Lawn, NJ) and VWR (West Chester, PA), respectively. Electrospray solutions were buffered by 10 mM NH4OAC (pH 7.0). 2.3. Computational Methods. The geometries of all neutral and charged peptides as well as aldehydes were optimized using the AM1 semiempirical method.14 For all peptide species, the initial guess structures were helices. The charged peptides were formed by placing a proton at the N-terminus proline residue, the C-terminus lysine residue, or both. Vibrational frequencies were also calculated using AM1 to yield the zeropoint energies and the thermal corrections to the enthalpy at 298 K. True energy minima were determined by checking the absence of imaginary frequencies from the set of obtained frequencies. Following geometry optimizations, single-point energies were calculated using density functional theory at the B3LYP/6-31+G(d) level.1517 The enthalpy for each species was obtained by combining the electronic energy calculated at the B3LYP level and the zero-point energy plus the thermal correction calculated at the AM1 level. The enthalpy values were used to calculate the proton affinities of the peptides using an isodesmic proton transfer reaction with lysine as the reference. Calculations of the aldehyde proton affinities were performed using an isodesmic proton transfer reaction with acetaldehyde as the reference, as discussed in Section 3.2 and in the Supporting Information. All calculations were performed using the Gaussian 03 suite of programs.18

3. RESULTS AND DISCUSSION This section presents the results of measurements of neutralion collisions leading to the formation of aldehyde complexes 11184

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Table 1. Aldehyde Masses from Lys+ Mass Spectrum (CH2)n n

ΔMa

Mexptb

MAldc

butanal

2

107.6

71.6

72.11

pentanal

3

123.5

87.5

86.13

hexanal

4

138.8

102.8

100.16

heptanal

5

151.6

115.6

114.18

octanal

6

165.5

129.5

128.21

nonanal

7

179.0

143.0

142.24

decanal

8

194.0

158.0

156.2

undecanal dodecanal

9 10

205.5 219.6

169.5 183.6

170.29 184.32

tridecanal

11

234.0

198.0

198.34

aldehyde

ΔM mass spectrum distance from parent ion (Da). b Mexpt = (ΔM 2Mwater) (Da). c Mald aldehyde molecular weight (Da). a

Figure 2. (a) Mass spectrum of the peptide [Pro2-Lys+H]+ displays only the parent ion after a 50 s of exposure at 300 K. (b) Mass spectrum of the doubly protonated peptide [Pro2-Lys+2H]2+ displays two bands of doubly protonated aldehydePro2 complexes in addition to the parent ion in both the singly and doubly protonated charge states after an exposure of 1 s at 300 K.

with trapped, desolvated amino acids (LysH+, HisH+, and ArgH+) and polypeptides [Pron-Lys+2H]2+, [(Gly-Ser)n-Lys+ 2H]2+. Mass spectrometry data identify the aldehyde species forming complexes with [Pro4-Lys+2H]2+, and the kinetics of these complexes will be described by reactions that model their time dependence. The presence of structural heterogeneity will be shown to play an important role in the evolving kinetics of [Pron-Lys+2H]2+ for n = 210. Data and calculations will be presented supporting the presence of proton transfer reactions contributing to the kinetics of [Pro4-Lys+2H]2+. The increased importance of proton transfer for the flexible structure of [(GlySer)3-Lys+2H]2+ will be discussed. 3.1. AldehydeAmino Acid Complexes. (a). Mass Spectra. Figure 1 displays a mass spectrum after exposure of trapped LysH+ ions for 10 s at 300 K showing the parent ion at M = 147.2 Da and a mass band spanning 250400 Da. The mass band represents populations of single aldehydes, CH3(CH2)nCHO, bound to LysH+. Each aldehydeLysH+ complex is denoted by (M+H+2H2O+Aldn)+, where n indicates the number of CH2, methylene groups. The sum of parent and complex ions is measured to be constant, independent of exposure time or trap storage time. The range of n values extends over 2 e n e 11, exhibiting a particularly stable complex for butanal, n = 2. The aldehyde distribution measured13 in mass spectra of Inland 19 forepump oil vapor is remarkably close both in abundance and

Figure 3. Parent ion fraction for amino acids is plotted versus exposure time for ArgH+, HisH+, and LysH+ amino acids. The solid curves each represent a double exponential fit to the data.

species to the measured distribution of aldehydeLysH+ complexes shown in Figure 1. Table 1 lists ΔM, the mass separation of each peak from the peak of the LysH+ parent ion. An experimental value for the aldehyde mass, Mexpt, associated with each mass peak is obtained from the relationship Mexpt = ΔM 2MH2O. These experimental mass values are equal to the corresponding aldehyde molecular weights, Mald, to within experimental uncertainty expressed by the slope of Mexpt versus MAld = 0.99 ( 0.01. The presence of two water molecules in each complex indicated by these mass spectra and the observation that water vapor is required for complex formation are most probably related to details of the formation dynamics. This will be discussed further in the section on kinetics measurements. (b). Sequence and Charge State Dependence. The mass spectrum in Figure 2a for [Pro2-Lys+H]+ at 300 K and exposure of 50 s displays a result observed for all singly protonated sequences [Pron-Lys+H]+ with n g 2. These sequences display a complete absence of complex formation. Figure 2b shows the mass spectrum for a 1 s exposure of the identical sequence, but doubly protonated, [Pro2-Lys+2H]2+. This spectrum displays a broad array of complexes formed in two mass bands and, in addition, a peak corresponding to the singly protonated parent ion, [Pro2-Lys+H]+. Increasing the length of the proline sequence increases the number of backbone carbonyl groups available to solvate a charge site, which would sterically hinder availability of the site to an aldehyde bond. Increased length also reduces the Coulomb repulsion between the protonation sites, which increases the probability that solvation substructures can form. Consequently, the result shown in Figure 2a is most probably the result of a competition between intramolecular solvation and aldehyde binding interactions for accessible protonation sites. For the higher charge state of [Pro2-Lys+2H]2+, the probability for exposed protonation sites increases, yielding the dramatic change in complex formation displayed in Figure 2b. A detailed analysis of the mass spectrum features measured for doubly protonated peptides will be presented below in Section 3.2. This competition for exposed charge sites implies that the aldehyde interacts predominantly with the peptide protonation site through the carbonyl group at the aldehyde terminus. The dependence on protonation site accessibility provides a strong sensitivity to the structural heterogeneity of the biomolecule ensemble, which will become more evident in the results discussed below. (c). Kinetics. Figure 3 compares loss of the parent ion, n1, resulting from the formation of the aldehyde complexes, na, for 11185



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Table 2. Amino Acid Kinetics Fit Parametersa Af

kf (s1)

As

ks (s1)

Ap

kb/kf

lysine

0.63 (0.02)

3.6 (0.3)

0.25 (0.01)

0.1 (0.01)

0.11 (0.01)

0.12 (0.01)

histidine

0.34 (0.03)

1.0 (0.1)

0.36 (0.03)

0.08 (0.01)

0.28 (0.01)

0.40 (0.02)

arginine

0.11 (0.02)

1.1 (0.4)

0.19 (0.01)

0.05 (0.01)

0.7 (0.02)

2.2 (0.2)

peptide

a

Parentheses indicate standard deviation.

Figure 4. Mass spectrum of the peptide [Pro4-Lys+2H]2+ for the indicated exposure times at 300 K. Peaks for the parent ion (2+) and singly protonated parent (1+) are indicated. The lower mass band represents complexes with a single aldehyde having peaks denoted by n, the number of (CH2) methylene groups. The higher mass band represents complexes composed of two aldehydes having peaks denoted by n,m, the number of (CH2) methylene groups in each species. Each spectrum peak in the higher mass band contains contributions from a sum of complexes, as explained in the text.

amino acids LysH+, HisH+, and ArgH+. Values for the number of parent ions, n1, and the aldehyde complexes, na, are obtained by integrating over the mass spectrum peak for the parent ion and the broad mass band (Figure 1), respectively. To eliminate effects of fluctuations on the number of trapped parent ions, we normalized both n1 and na abundances by the initial ion number, n1/n0, na/n0. The total number of ions, n0, was observed to remain constant (n0 = n1 + na) within experimental uncertainty for each exposure time. Figure 3 displays the decay of the parent ion fraction for exposure times up to 90 s and indicates that the decay asymptotes to a constant plateau for each amino acid. Both the decay rate and the plateau value are dependent on the amino acid species. Each decay of ion population in Figure 3 is fit by a double exponential, n1/n0 = Af exp(kft) + As exp(kst) + Ap, in which kf (ks) is the fast (slow) decay component. Multiexponential decays in ensemble measurements of biomolecule properties are generally associated with populations of different conformer structures.19 In particular, the observation of plateaus in Figure 3 identifies the presence of a back reaction resulting from a group of structures for which complex formation is strongly inhibited. For example, the dominant back reaction would be expected if there were structures for which intramolecular solvation of the charge was sufficiently probable to prevent stabilization of the complex. The constant plateau for each of these amino acids indicates that they arise from a population of noninterconverting structures. In this model, the decay parameters in Table 2 represent an ensemble fraction, Ai, having similar structures that

yield a decay rate of the parent ion given by ki. The parameters in Table 2 indicate that lysine is dominated by structures yielding the fast rate kf; histidine by roughly equal contributions from rates kf, ks; and arginine by the constant plateau, Ap. The steady-state indicated by the plateau describes an equilibrium population of parent ions and complexes determined by the forward rate of bimolecular collisions, kf, and the back rate, kb, of the complex dissociation. The steady state n1 population given by n0Ap is expressed by dn1/dt = kfn0Ap + kbna = 0 with n0 = n1 + na. This steady-state yields a ratio of backward-to-forward rates given by kb/kf = Ap/(1 Ap), which was measured to be roughly independent of the background helium pressure in the trap. Table 2 lists this ratio for each amino acid and indicates that Arg+ has the largest population of conformer structures that do not form aldehyde complexes. Assuming structures contributing to complex formation differ primarily in the degree of intramolecular solvation, as suggested by the model above, it remains to specify the details of such structures. The lowest energy structures of both LysH+ and ArgH+ and their hydrated forms have been calculated,20 and differences in the importance of electrostatic interactions were noted. Different hydration sites are indicated in structures calculated20 for LysH+ 3 (H2O)n and ArgH+ 3 (H2O)n, and result in part from greater charge localization on the ammonium group of Lys compared with the guanidinium group of Arg. The differences in water positions for these calculated structures suggest differences in steric availability of the charge sites for aldehyde binding. The presence of two water molecules per aldehyde in the mass spectrum of the Lys complexes identified in Table 1 is found for the Arg and His complexes as well. It is reasonable to consider that each complex was formed by aldehyde addition to a hydrated amino acid with at least one to two waters and possibly more. In this case, a hydrated parent ion ensemble would be composed of different structures with relative populations depending on the specific amino acid. These aldehydeamino acid complexes suggest that water can play a critical role in the complex formation process. Calculations are planned to consider the stability of aldehyde complexes on hydrated amino acids of LysH+ 3 (H2O)n and ArgH+ 3 (H2O)n to better understand how these structures relate to the measured kinetics. Finally, it must be pointed out that the dependence of formation on the number of methylene groups has not been considered here and further experiments on single aldehyde species could address this issue. 3.2. AldehydePolypeptide Complexes. (a). [Pro4-Lys+2H]2+ Mass Spectra. Figure 4 displays mass spectra measured for the peptide [Pro4-Lys+2H]2+ obtained at 300 K for several exposures to a mixture of water and aldehyde vapors between 0.5 and 10 s. The charge sites are assumed to be on the lysine residue and on the N-terminus. Note that features of the mass spectrum are similar to those observed for [Pro2-Lys+2H]2+ in Figure 2b and will be analyzed here in detail for this larger peptide. 11186



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bands denoted by na. For each exposure, the spectrum peaks corresponding to the parent ions and the aldehyde complex bands were integrated and normalized by the total ion number measured for that exposure. In Figure 5, the n2+ population decays to a plateau of ∼2%, and na increases to a peak at a comparable time and rate. Formation of n+ begins with complex formation at a rate that is comparable to the decay rate of na. The sum of all species is constant, independent of exposure time, but as a result of n+ formation, the total charge is not constant. The reactions dominating aldehydepeptide kinetics include Figure 5. Fraction of ions obtained from mass spectra for [Pro4-Lys+ 2H]2+ versus exposure time. Ion fractions representing the doubleprotonated parent peptide, n2+, the singly protonated parent n+, and the sum of all aldehyde complexes, na, are plotted. The legend identifies markers for these curves. The solid curves represent fits to solutions derived from the kinetic model discussed in the text.

ka

ð1Þ

where na = ∑n[M + 2H + Aldn]2+ + ∑n,m[M + 2H + Aldn + Aldm]2+. Decay of the complex is assumed to involve proton transfer to the aldehyde ligands kþ

Peptide Peaks. Two mass peaks associated with bare ions are evident in the spectrum. In addition to the parent ion peak near 270 Da, [Pro4-Lys+2H]2+, the mass spectrum (Figure 4) displays a peak near 540 Da identified as singly protonated [Pro4-Lys +H]+. As exposure of the peptide to aldehyde flux increases, the parent ion peak is observed to decrease, and the singly protonated ion peak increases. These variations will be shown to arise from the formation and loss of the aldehyde complex, respectively, and are discussed in the Kinetics section below. Small abundances of proline fragmentation are formed during ejection of the parent ion. Such fragmentation has been identified21 to occur routinely for doubly charged polyproline peptides, and these peaks are sufficiently small in these spectra that they do not interfere with kinetics measurements. Mass Bands of Aldehyde Complexes. Two broad mass bands centered near 350 and 450 Da and extending over 300480 Da are observed to evolve as the parent ion peak decreases. These bands shift to higher masses with increasing exposure, as observed in Figure 4. The peaks in these bands are identified as masses of aldehydepeptide complexes and are indicated in the spectra by the number of methylene groups in the aldehyde species. The lower mass band corresponds to a distribution of complexes formed by a single aldehyde species n denoted by (M+ 2H+Aldn)2+. The higher mass band is composed of complexes formed by two aldehyde species n and m denoted by (M+2H+ Aldn+Aldm)2+. Each mass peak in the double aldehyde band that is indicated by n/n in Figure 4 represents a sum of complexes formed by all of those aldehyde pairs (n1, n2) having a total of n1 + n2 = 2n methylenes. For simplicity, these peaks are indicated by the aldehyde pair having n1 = n2. Each intermediate peak in the double aldehyde band denoted by n/m represents a sum of complexes formed by all those aldehyde pairs having a total of n1 + n2 = 2n + 1 methylenes. Water molecules are not observed to be included in the complexes for this polypeptide. This suggests that although the presence of water vapor was required to stabilize formation, the aldehydepeptide interaction is sufficient to maintain the stability without hydration. (b). [Pro4-Lys+2H]2+ Kinetics. Ion abundances were measured from mass spectra obtained at 300 K for exposures of the parent ion to aldehyde and water vapors over a range of 0.590 s. Figure 5 shows plots of normalized ion abundance versus exposure time for the parent ion, n2+, the singly charged parent ion, n+, and the sum of adsorption complexes in the two mass

∑n Aldn fs na

n2þ þ

na sf

∑ ½AldnHþ þ Aldm Hþ þ ðAldnAldm ÞHþ

n, m

þ nþ

ð2Þ

Data and calculations discussed below support the presence of proton transfer and, in particular, the dimer formation channel indicated by (AldnAldm)H+. These reactions are modeled by rate equations, eq (A2), discussed in the Supporting Information. The decay of n2+ is fit by a double exponential, and similar to the model applied to the kinetics of single amino acid complexes, the superscript i = (f, s) represents the rates and populations associated with two groups of noninterconverting structures that yield the fast and slow decay rates. The total populations are then given by n2þ ¼ np þ nf0 exp½kfa ðt t 0 Þ þ ns0 exp½ksa ðt t 0 Þ na ¼

∑i nia ,

nþ ¼

∑i niþ

ð3Þ

The rates kia and ki+ characterize the net formation of nai and ni+, respectively. The populations naf(nas) and nf+(ns+) are each calculated and summed to obtain the total populations of na and n+. The plateau, np, is not included in the calculation for na and n+ because np is assumed to represent a structure that does not form complexes. Solutions of the rate equations are given in the Supporting Information, and Figure 5 displays a fit of these solutions as smooth curves through the data. The close fits for na and n+ cannot be obtained if the double exponential fit for n2+ is simply used in the rate equation to calculate na and n+. The rate equations do not include back-reaction contributions from na f n2+, implying that aldehyde loss by the complex has a rate significantly slower than the formation rates. This assumption is found to be consistent with data and analysis of the individual complex mass bands discussed below. The parameters used to fit the peptide data are listed in Table 3. The minimal plateau of ∼2% of the ions is consistent with the expected accessibility of charge sites for the 2+ structure, as suggested by the calculated 2+ structure shown in the Supporting Information, Figure S3. The fast association rate, kfa, is associated with the dominant structure population (Af = 0.83) forming the aldehyde complex. An upper bound to this rate can be estimated by considering the chargedipole and charge-induced dipole interactions2224 between 11187



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Table 3. [Pro4-Lys+2H]2+ Kinetics Fit Parametersa

a

Af

kfa (s1)

As

ksa (s1)

Ap

kb/kf

k+ (s1)

0.83 (0.01)

3.18 (0.09)

0.15 (6 103)

0.75 (0.05)

0.02 (2 103)

0.02(2 103)

3 103 (3 104)


Table 4. Mass Band Kinetics Parameters: na1 a na2 mass band

na0

R

k12

k21

ka+

na1 na2

0.80 0.20

0.72 (0.06) 0.82 (0.05)

0.20 (0.02) 0.21 (0.02)

0.14 (0.03) 0.17 (0.03)

0.005 (0.001) 0.003 (0.001)

Table 5. Mass Band Kinetics Parameters: na2 a na1a

a

Figure 6. (a) Fraction of complex ions and singly protonated parent ions measured in mass spectra of [Pro4-Lys+2H]2+ are plotted versus exposure time. Spectra are obtained after aldehyde complex ions having a single aldehyde, na1, are isolated and exposed to aldehyde vapor for the indicated times. The mass bands representing single aldehyde complexes, na1, double aldehyde complexes, na2, and the singly charged parent ion, n+ are indicated. The solid curves represent fits to solutions derived from a kinetic model of complex interconversion discussed in the text. (b) Graph similar to part a for spectra obtained after aldehyde complex ions having two aldehydes, na2, are isolated and exposed to aldehyde vapor for the indicated times.

mass band

na0

R

k12

k21

ka+

na1

0.08

0.31 (0.01)

0.51 (0.05)

0.16 (0.02)

0.006 (0.001)

na2

0.92

0.41 (0.01)

0.47 (0.05)

0.19 (0.03)

0.003 (0.001)


selecting na2 ions after na1 was completely converted to na2, allowing the inverse kinetics, na2na1, to be monitored. The mass spectra obtained are shown in Figures S2a and S2b in the Supporting Information. Figure 6a displays the normalized ion abundances for na1, na2, and n+ obtained from mass spectra that display interconversion kinetics for exposures of 0.590 s following the initial conditions, na10 = 0.8, na20 = 0.2. Note that interconversion reaches an equilibrium after ∼57 s predicted by the rate equation solutions in eq (B3) of the Supporting Information. However, the n+ ion abundance increases monotonically throughout the entire range of aldehyde vapor exposure. Consequently, both na1 and na2 bands are contributing to n+ formation. Similar behavior is displayed for interconversion kinetics in Figure 6b following the initial conditions, na10 = 0.08, na20 = 0.92. Kinetics reactions include the formation and dissociation of na2 during exposure na1 þ

[Pro4-Lys+2H]2+ and the aldehyde carbonyl group. Consider bimolecular collisions between the peptide ion and the aldehyde species hexanal, CH3(CH2)4CHO, having a polarizability,25 11.9 Å3, and carbonyl dipole moment,26 2.5 D. A rate constant of 1.4 109 cm3/molecule-s is calculated assuming localized charge and point-dipole approximations. A collision rate of ∼4.4 s1 is estimated assuming a density corresponding to an aldehyde vapor pressure of ∼107 Torr in the ion trap. This rate is comparable in magnitude to the experimentally measured decay rate of the parent ion, kfa ≈ 3.18 s1 (Table 3). (c). Interconversion between Complexes. Ion manipulation in the RF trap allows complex ions with a single aldehyde, na1, or two aldehydes, na2, to be individually selected and exposed to the aldehyde/water vapors. This technique was applied to measure the interconversion kinetics of the complex species, na1na2. Mass spectra were obtained by selecting complex ions contained in the na1 band at a time after the parent ion was completely depleted. Additional mass spectra were then obtained that monitor the time evolving kinetics for mass bands, na1 = ∑n(M + 2H + Aldn)2+, na2 = ∑n,m(M + 2H + Aldn + Aldm)2+, and n+ for exposure times 0.550 s at 300 K. Similarly, mass spectra were obtained by

s s na2 ∑n Aldn f r k k12

ð4Þ

21

and the decay of both single and double aldehyde complexes to the singly protonated parent ion, n+, by proton transfer k1þ

na1 sf nþ þ

k2þ

na2 sf

∑n Aldþn

ð5aÞ

½Aldn Hþ þ Aldm Hþ þ ðAldn Aldm ÞHþ ∑ n, m

þ nþ

ð5bÞ

These reactions are modeled by rate equations and their solutions given in the Supporting Information (eqs B1B5). The rates k12 and k21 are conversion rates for na1f na2 and na2f na1, respectively; k1+ and k2+ are the net proton transfer rates for na1 and na2 to form n+, respectively. Figure 6a,b includes best-fit solutions as smooth curves through the data. Note that back reactions of na1 to the parent ion are not included in the rate equation, consistent with the mass spectra in Figure S2a in the 11188



Figure 7. Percentage of [Pron-Lys+2H]2+ ions forming aldehyde complexes versus peptide length (left axis) obtained from mass spectra, as discussed in the text. The ratio of back-to-forward reaction rates (right axis) is also plotted versus peptide length. The legend identifies markers for these data. The solid curve is a sigmoidal fit to the data, and the dashed curve is a guide to the eye.

Supporting Information that do not exhibit a mass peak near 270 Da corresponding to the parent ion. Table 4 lists the rate parameters k12, R = k21/k12, and k1+, k2+ resulting from fitting rate equation solutions to both the na1 and na2 data independently. The column of initial abundance, na0, lists na10 and na20 for the kinetics fits shown in Figure 6a. Note that the independent fits yield essentially identical sets of rate parameters within experimental uncertainty, indicating that proton transfer is the limiting rate for aldehyde loss from both na1 and na2 aldehyde complexes. The average of the rates k1+, k2+ weighted by the na1 and na2 fractions is equal to the k+ rate previously found for the sum (na1 + na2) within experimental uncertainty. Table 5 lists the rate parameters resulting from fitting rate equation solutions to both the na1 and na2 data independently for different initial conditions, as shown in the column of initial abundance, na0, which lists na10 and na20 for the kinetics fits shown in Figure 6b. In principle, if experimental conditions were identical except for the initial conditions on na10 and na20, then best fit parameters in Figure 4b would be the same as those in Figure 4a within experimental uncertainty. Note that only the values for R and k12 in Table 5 exhibit significantly different parameters from those in Table 4, k12 differing by a factor of 2.3 and R by a factor of (2.3)1. Because the rate k12 depends linearly on the aldehyde flux and R depends on 1/k12, it is apparent that there was a variation of aldehyde flux between the data shown in Figures 6a and 6b. The aldehyde flux is unfortunately a weakly controlled parameter in this experiment, and a factor of 2 variation from different experiments is not surprising. This is the only case in which best-fit parameters for different experiments are compared, and changes in flux were not observed for data within an experimental run. (d). Sequence Dependence. Mass spectra exhibiting aldehyde complex formation on polyproline peptides [Pron-Lys+2H]2+ were obtained for n = 210 at 300 K. Spectra are shown in Figure S1 of the Supporting Information for peptides n = 5, 6, 7, and 9 for an exposure of 5 s. These spectra display a decrease in the complex formation rate as the proline number increases, suggesting a decrease in the fraction of structures that can form complexes. The total number of ions that have formed a complex is given by the sum naT = (na + n+) because n+ ions were derived from complexes by proton transfer. Figure 7 displays a plot of naT versus n (number of proline residues) and indicates that naT decreases in a sigmoidal curve, characterizing a transition in the population of structures of the ion ensemble near n = 7.

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Figure 8. Low mass portion of the [Pro4-Lys+2H]2+ spectrum shown in Figure 4. Low abundance peaks clustered around the parent ion are identified as singly protonated aldehyde dimers. Each dimer peak contains contributions from a sum of dimers similar to Figure 4 and explained in the text.

A steady-state analysis similar to that performed for amino acid plateaus in Section 3.1 yields an expression for the ratio of backward and forward reaction rates, kb and kf, given by kb/kf = (1 AaT)/AaT, where AaT = naT/n0 normalized by the total ion number. The ratio of these rates is observed to vary from kb/kf ≈ 0 for n = 26 to ∼35 for n = 10. The basis for such a large variation in this rate is most probably associated with an increasing probability of intramolecular solvation as the peptide length increases. These measurements support the hypotheses that accessible protonations sites are required for complex formation and that structural heterogeneity is a dominant characteristic in the formation of aldehydepeptide complexes. (e). Proton Transfer Formation of (M+H)+. The application of proton transfer reactions to investigate the structure and structural change of gas-phase biomolecules has been wellestablished.27 The following discussion will support the assertion that a proton transfer reaction is responsible for the singly protonated parent ion observed in mass spectra characterizing aldehyde complex formation with [Pron-Lys+2H]2+. The presence of proton transfer will be identified by reaction products observed in the mass spectra and also supported by calculations of the relevant proton affinities. Finally, measurements of [(Gly-Ser)m-Lys+2H]2+ that exhibit greater flexibility than the polyproline peptide will be shown to accelerate the proton transfer process, as expected. [Pro4-Lys+H]+ Formation. Proton transfer reactions in doubly protonated complexes formed with a single aldehyde result in protonation of the aldehyde ligand, AldnH+ (eq 5a). The intermediate product ion will rapidly dissociate in the presence of strong Coulomb repulsion, producing a bare singly protonated parent ion. Transfer in doubly protonated complexes formed with two aldehydes can result in three separate product channels (eq 5b): formation of a protonated aldehyde, AldnH+ or AldmH+, and the formation of a protonated dimer, AldnAldmH+. Following the loss of an aldehyde ion from the complex, the remaining neutral aldehyde species dissociates consistent with observed mass spectra and probably related to thermal energy deposited during the transfer reaction. In the dimer channel, both aldehydes dissociate as a dimer ion. Consequently, proton transfer in the doubly protonated peptide results in the formation of a bare, singly protonated parent ion. Reaction products of protonated aldehydes and aldehyde dimers provide evidence of these kinetics. Figure 8 displays the low mass portion of the spectrum obtained for the 10 s of exposure of [Pro4-Lys+2H]2+ shown in Figure 4. The mass peaks identified as singly protonated aldehyde dimers are composed of aldehydes indicated by the methylene 11189



Figure 9. Low mass portion of the mass spectrum of [Gly-Ser-Lys+ 2H]2+. Low abundance peaks near the parent ion are identified as singly protonated aldehyde monomers. The spectrum peaks for aldehyde ions are indicated in blue by the number of aldehyde methylene groups.

numbers. The dimer ion peaks indicated by n,n include contributions from all aldehyde pairs having a methylene sum of 2n. Dimer ion peaks indicated by n,m include contributions from all aldehyde pairs having a methylene sum of 2n + 1. The dissociation channel of single aldehyde ions is not detectable in these mass spectra because their masses are below the minimum mass for trap stability. Intramolecular proton transfer in these aldehyde complexes from the protonated peptide, H+Pro4-LysH+, to an aldehyde species will depend on the relative proton affinities of the peptide and the aldehyde. Here the peptide sequence has been written in a form that indicates proton sites on the Lys ammonium group and the N-terminus. Calculations of all trans polyproline structures and enthalpies were performed as described in the Computational Methods. The Supporting Information describes results for the peptide structures shown in Figure S3 and for proton affinities of both peptides and aldehydes in Figures S4 and S5, respectively. In particular, a proton affinity of PA ≈ 205 kcal/mol was calculated for the reaction H+Pro4-LysH+ f Pro4-LysH+ + H+ (Figure S4). Note that the PA calculated for the doubly protonated peptide is less than that of the singly protonated peptide (Figure S4) by ∼50 kcal/mol due to the presence of electrostatic repulsion. The calculation was performed for the static helical structure of H+Pro4-LysH+ shown in Figure S3 in which the protons are separated by 13.35 Å. The proton affinities of aldehyde species (Figure S5) are found to be PA ≈190 kcal/ mol, roughly independent of the number of methylene groups.28 The difference in proton affinities of the peptide and aldehyde of ∼15 kcal/mol could be compensated for by fluctuations that reduce the charge separation to ∼8 to 9 Å. Polyproline structures composed of a random mixture of trans and cis isomers have been shown29 to be more compact than the extended structures of all trans isomers. This also can lead to a PA for H+Pro4-LysH+ which is closer to the aldehyde PA. [(Gly-Ser)m-Lys+H]+ Formation. The possibility that fluctuations can play an important role determining the rate of intramolecular proton transfer can be investigated by measurements of aldehyde complex formation in [(Gly-Ser)m-Lys+2H]2+. Mass spectrometry measurements were performed by the same procedures used to obtain the polyproline data, and similar methods were applied to reduce the data for kinetics analysis. The mass spectrum for [Gly-Ser-Lys+2H]2+, m = 1, shown in Figure 9 was obtained for an exposure of 0.5 s at 300 K. In this case, the mass spectrum falls in a mass range that allows the single aldehyde ions to remain trapped in stable ion trajectories. As a result, mass peaks are detected for the singly protonated aldehyde ions, Aldn+, n = 57. The aldehyde mass peaks in both

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Figure 10. Fraction of ions obtained from mass spectra for [(Gly-Ser)3Lys+2H]2+ versus exposure time. Ion fractions representing the doubleprotonated parent peptide, n2+, the singly protonated parent, n+, and the aldehyde complexes, na, are plotted versus exposure time. The legend identifies markers for these data. The solid curve represents a single exponential fit to the data for na.

Figures 8 and 9 are significantly weaker than other peaks in the spectra. Note, however, that the integrated abundance of the singly protonated parent ion is a relatively small fraction of the total ions, and the aldehyde product peaks in Figures 8 and 9 represent a small number of the total aldehyde ion products. In addition, the branching ratios among the single and dimer ion reaction channels are convolved with the observed abundances. Nevertheless, these spectra provide convincing evidence that proton transfer reactions are present in the kinetics, determining the formation and loss of aldehydepeptide complexes. An indication that fluctuations can be associated with increased proton transfer rates is shown in Figure 10 which exhibits the kinetics for aldehyde complexes with [(Gly-Ser)3-Lys+2H]2+ peptides. The large flexibility11,30 of (Gly-Ser)m peptides relative to the polyproline peptides is expected to increase the rate that fluctuations lead to close encounters of the protonation sites. Figure 10 displays the kinetics for the [(Gly-Ser)3-Lys+2H]2+ peptide exhibiting a formation rate of the singly protonated parent ion of k+ = 0.14 ( 0.01 s1, a factor of ∼50 greater than that measured for [Pro4-Lys+2H]2+, (Table 3). Kinetics data were also obtained for [(Gly-Ser)5-Lys+2H]2+ and are shown in Figure S6 of the Supporting Information. These data indicate that the formation rate of the singly protonated parent decreases for [(Gly-Ser)5-Lys]2+ to k+ = 0.010 ( 0.001 s1, a factor of ∼3 greater than that measured for [Pro4-Lys+2H]2+. These results demonstrate that a more flexible, doubly protonated peptide increases the rate of intramolecular proton transfer in these measurements. As expected, a longer peptide length reduces the rate enhancement because fluctuations in [(Gly-Ser)5-Lys]2+ sample a larger volume that reduces the rate that close encounters of the protonation sites will occur.

4. SUMMARY AND CONCLUSIONS The experiments presented in this Article introduced aldehyde vapor into an RF trap and studied the collisional interactions of a distribution of aldehydes with unsolvated ions of amino acids and polypeptides. Mass spectrometry measurements were performed to follow the kinetic reactions responsible for the formation and loss of the observed aldehydeion complexes. The motivation for these measurements is the possibility that such complexes could provide a simple model of the hydrophobic environment experienced by biomolecules residing within the cell membrane. 11190


The Journal of Physical Chemistry B The question is whether such a model offers the opportunity to study changes in structure experienced by membrane proteins. These measurements yielded reproducible mass spectra and kinetic rates; however, they did not provide rate constants as a result of uncertainties in the aldehyde vapor densities. These data identified the basic kinetics processes dominating formation and loss of aldehyde complexes for different amino acids and polypeptide lengths. In each species, formation appears to involve a competition between intramolecular solvation of protonation sites within the isolated gas-phase peptide and solvation by the carbonyl group on the aldehyde molecule. This suggests the aldehyde binding results from a noncovalent interaction with biomolecule protonation sites. As a result, the availability or access to these charge sites is a critical prerequisite to complex formation, and this requirement is the basis for the observed sensitivity to structural heterogeneity. Complex formation is also found to require the presence of water vapor, and it is not clear at this point how the water plays a role in stabilization of the aldehyde bond. The wateraldehyde interaction clearly requires further experiments with single aldehydes and possibly calculations to understand the basis for the observed results. Initial measurements of aldehydeprotein complexes on bovine pancreatic trypsin inhibitor, cytochrome c, and ubiquitin have each displayed a strong formation dependence on protonation state, but inclusion here is beyond the scope of this Article. Kinetic models describing complex formation are based on the assumption that there are noninterconverting groups of conformer structures that vary in their ability to stabilize the complex. It is suggested that this variation depends on the degree that these conformer structures exhibit intramolecular solvation of the charge sites. Although these models are consistent with the observed kinetic behavior, they remain hypotheses awaiting further analysis and experiments. However, it appears clear that complex formation is intimately related to structure and yields another metric by which to identify structural change. The loss of complexes through proton transfer to an aldehyde, followed by dissociation is identified by observed reaction products and supported by proton affinity calculations. Note that the rapid dissociation of the second aldehyde in a complex after proton transfer is consistent with thermal dissociation of a weak noncovalent bond. Measurements with flexible peptides that include (Gly-Ser)m units indicate that proton transfer probability is increased by amplitude fluctuations. Such fluctuations can result in a decrease in protonation site separations, leading to decreased proton affinity, as shown by calculations.

’ ASSOCIATED CONTENT

bS

Supporting Information. Rate equation calculations performed to analyze the kinetics of aldehyde complexes: (a) the kinetics governing the exposure time dependence of formation and decay of aldehydepeptide complexes and (b) the association and dissociation processes leading to interconversion among different complex species. Calculations of the structures and proton affinities of doubly charged polyproline peptides and of neutral aldehydes. These calculations helped to consider the possibility that proton transfer within the complex is responsible for the presence of both the singly protonated peptide and aldehyde cation products in the mass spectra. Various figures have been included to support the various kinetic analyses. The full listing for ref 18 is also included. This material is available free of charge via the Internet at http://pubs.acs.org.

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’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Tel: (617) 497-4653. Fax: (617) 497-4627. Present Addresses §

Aileron Therapeutics, Inc., Cambridge, MA 02139.

’ ACKNOWLEDGMENT We gratefully acknowledge helpful discussions with Dr. Ryan Danell (Danell Consulting), Dr. James Foley (Rowland), and Dr. Michael Burns (Rowland) and assistance with experimental electronics and data acquisition software by Chris Stokes, Winfield Hill, and Dr. Alan Stern (Rowland). We gratefully acknowledge the generous financial support by the National Science Foundation (grant CHE-0962680) and during the early stage of these experiments by The Rowland Institute at Harvard. ’ REFERENCES (1) Rodriguez-Cruz, S. E.; Klassen, J. S.; Williams, E. R. J. Am. Soc. Mass Spectrom. 1999, 10, 958–968. (2) Hoaglund-Hyzer, C. S.; Counterman, A. E.; Clemmer, D. E. Chem. Rev. 1999, 99, 3037–3079. (3) Koeniger, S. L.; Merenbloom, S. I.; Clemmer, D. E. J. Phys. Chem. B 2006, 110, 7017–7021. (4) Jarrold, M. F. Annu. Rev. Phys. Chem. 2000, 51, 179–207. (5) McLafferty, F. W.; Guan, Z.; Haupts, U.; Wood, T. D.; Kelleher, N. L. J. Am. Chem. Soc. 1998, 120, 4732–4740. (6) Badman, E. R.; Hoaglund-Hyzer, C. S.; Clemmer, D. E. Anal. Chem. 2001, 73, 6000–6007. (7) Konermann, L.; Collings, B. A.; Douglas, D. J. Biochemistry 1997, 36, 5554–5559. (8) Konermann, L.; Simmons, D. A. Mass Spectrom. Rev. 2003, 22, 1–26. (9) Kaltashov, I. A.; Eyles, S. J. Mass Spectrom. Rev. 2002, 21, 37–71. (10) (a) Prell, J. S.; Chang, T. M.; O’Brien, J. T.; Williams, E. R. J. Am. Chem. Soc. 2010, 132, 7811–7819. (b) Liu, L.; Bagal, D.; Kitova, E. N.; Schnier, P. D.; Klassen, J. S. J. Am. Chem. Soc. 2009, 131, 15980– 15981. (11) Iavarone, A. T.; Meinen, J.; Schulze, S.; Parks, J. H. Int. J. Mass Spectrom. 2006, 253, 172–180. (12) Shi,X.; Ren, J.; Danell, R. M.; Parks, J. H. Adsorption of Aldehydes on Trapped Biomolecular Ions. In Proceedings of the 58th ASMS Conference On Mass Spectrometry and Allied Topics, Salt Lake City, Utah, May 2327, 2010 ; ASMS: San Francisco, 2010. (13) Scientific Instrument Services, Inc. http://www.sisweb.com/ referenc/applnote/app-42.htm (accessed December 18, 2010). (14) Dewar, M. J. S.; Zoebisch, E. G.; Healy, E. F.; Stewart, J. J. P. J. Am. Chem. Soc. 1985, 107, 3902–3909. (15) Parr, R. G.; Yang, W. Density-Functional Theory of Atoms and Molecules; Oxford University Press: New York, 1989. (16) Becke, A. D. J. Chem. Phys. 1993, 98, 5648–5652. (17) Lee, C.; Yang, W.; Parr, R. G. Phys. Rev. B 1988, 37, 785–789. (18) Frisch, M. J.; et al. Gaussian 03, revision A.1; Gaussian, Inc.: Pittsburgh, PA, 2003. Full reference is listed in the Supporting Information. (19) (a) Bai, C.; Wang, C.; Xie, X. S.; Wolynes, P. G. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 11075–11076. (b) Xie, X. S. Annu. Rev. Phys. Chem. 1998, 49, 441–480. (c) Stayton, P. S.; Sligar, S. G. Biochemistry 1991, 30, 1845–1851. (20) Gao, B.; Wyttenbach, T.; Bowers, M. T. J. Phys. Chem. B 2009, 113, 9995–10000. (21) Unnithan, A. G.; Myer, M. J.; Veale, C. J.; Danell, A. S. J. Am. Soc. Mass Spectrom. 2007, 18, 2198–2203. 11191



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Aldehyde Complexes with Protonated Peptides in the Gas Phase

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