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STUDIES OF THE OXIDATIVE REFOLDING OF CYSTINYL
PROTEINS BY HYDROGEN/DEUTERIUM EXCHANGE-MASS
SPECTROMETRY

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Xue Li

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2/05 p:lCIRC/DateDue.indd-p.1

STUDIES OF THE OXIDATIVE REFOLDING OF CYSTINYL PROTEINS BY
HYDROGEN/DEUTERIUM EXCHANGE-MASS SPECTROMETRY

By

Xue Li

A DISSERTATION
Submitted to
Michigan State University
In partial fulﬁllment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Department of Chemistry

2005

ABSTRACT

STUDIES OF THE OXIDATIVE REF OLDING OF CYSTINYL PROTEINS BY
HYDROGEN/DEUTERIUM EXCHANGE-MASS SPECTROMETRY

By
Xue Li

Hydrogen/deuterium exchange (HDX) with accurate mass measurement provides
important information about the conformation of a polypeptide chain. When a protein is
dissolved in D20, hydrogens located at peptide amide linkages can exchange with
deuteriums ﬁom the solvent at easily measurable rates. The rates depend on features
directly related to protein structure such as whether the amide hydrogens are participating
in intramolecular hydrogen bonding, and the extent to which they are shielded from the
solvent. Thus, conformational information about a protein can be obtained by assessing
the extent of deuterium incorporation during HDX.

Various parameters for HDX experiments were investigated to minimize back-
exchange and to accurately measure the mass of the deuterium-labeled protein by matrix
assisted laser desorption/ionization time of ﬂight mass spectrometry (MALDI-TOF-MS).
Optimized protocols for different types of HDX experiments were then proposed.
Disulﬁde bonds, an important post-translational modiﬁcation of proteins, play a key role
in stabilizing a protein’s 3-dimensional structure and controlling its activity. The novel
cyanylation/cleavage methodology, developed previously in this laboratory, has been
applied in several studies of protein folding because of its effectiveness in trapping,
identifying, and preserving the disulﬁde structure of a given folding intermediate of a
cystinyl protein. This methodology was integrated with HDX-MS to study the

conformational changes of a mode] cystinyl protein, long-R3 insulin-like growth factor-I

(LR3IGF-I), as a function of disulﬁde bond formation during its oxidative refolding. The
integrated approaches establish a novel strategy for studying the refolding of cystinyl
proteins.

Conventionally, HDX-MS has been used to characterize large proteins. We
investigated the applicability and advantages of HDX-MS in the conformational study of
small cystinyl peptides. The conformations of ﬁve peptide models derived from bovine
pancreatic trypsin inhibitor (BPTI) were characterized to study the effect of long-range
interactions (induced by the formation of disulﬁde bonds) in stabilizing their local folded
structure. Collisionally induced dissociation-tandem mass spectrometry (CID-MS/MS)
was employed to assess the extent of deuterium incorporation at the single amino acid
residue level.

During the oxidative refolding of LR3IGF-I, only one of the ﬁfteen possible one-
disulﬁde intermediates (the one containing the disulﬁde bond Cys31—Cys74) accumulates
to a signiﬁcant extent. A structural explanation is proposed for this phenomenon:
during the folding of LR3IGF-I, long-range interactions induced by the formation of the
Cy531-CYS74 disulﬁde bond may stabilize the local folded structure around it; the local
folded structure, in turn, protects this disulﬁde bond against attack from reducing
reagents in the solution such as free sulﬂtydryl groups. This hypothesis was tested by

probing the structural features of synthetic peptide models that mimic those of the intact

Cys3l—Cys74 l-S intermediate.

ACKNOWLEDGMENTS

I would like to express my gratitude to my advisor, Dr. J. Throck Watson, for his
support, advice, and encouragement. I would also like to thank Dr. William J.
Wedemeyer for his guidance and help; Dr. Victoria L. McGufﬁn and Dr. David P.
Weliky for helpful discussions and for serving on my committee.

I want to acknowledge Dr. Honggao Yan and Dr. Leslie A. Kuhn for helpful
discussions; Dr. Robin J. Hood, Dr. Yi-Te Chou, Dr. Rhonda Husain and Dr. Jiong Yu
for assistance and discussion; Dr. Steven P. Gygi for access to the LCQDeca XP ion-trap
mass spectrometer at Harvard Medical School and Dr. Babak Borhan for access to the
CD spectrometer at Michigan State University. To all the Watson group members, past
and present, thank you for your ﬁ'iendship and cooperation. I also owe many thanks to
the members of the Mass Spectrometry Facility: Dr. Daniel A. Jones, Lijun Chen,
Beverly Chamberlin and Susanne Hoffmann-Benning.

I especially would like to thank my mother, Shuying Xue, for her love, patience,
and support, without which I would not have been able to ﬁnish this Ph.D. project. At
last, but not the least, I would like to thank my husband, Gabriel D. Musk, and my

Chinese and American families for their love and support.

iv

TABLE OF CONTENTS

LIST OF TABLES ............................................................................................................ vii
LIST OF FIGURES ......................................................................................................... viii
ABBREVIATIONS .......................................................................................................... xii
CHAPTER 1
INTRODUCTION .............................................................................................................. l
I. Introduction ................................................................................................................... 1
II. Disulﬁde Mapping of Cystinyl/Cysteinyl Proteins ....................................................... 3
A. Conventional Proteolytic Fragmentation Strategy .................................................. 4
B. Novel Cyanylation/ Cleavage Methodology ........................................................... 9
HI. Oxidative Folding of Cystinyl/Cysteinyl Proteins and Trapping of Folding
Intermediates ............................................................................................................... 14
A. Disulﬁde Formation in Oxidative Folding of Cystinyl/Cysteinyl Proteins .......... 15
B. Trapping of the Folding Intermediates ................................................................. 18
IV. Conformational Study of Proteins and Peptides ......................................................... 24
A. Conventional Technologies in Protein/Peptide Conformational Studies .............. 24
B. HDX—MS ............................................................................................................... 31
V. Desorption/Ionization Mass Spectrometry ................................................................. 34
A. Matrix-Assisted Laser Desorption/Ionization Time of Flight Mass Spectrometry
(MALDI-TOF-MS) ..................................................................................................... 35
B. Electrospray Ionization Ion Trap Mass Spectrometry (ESI-Ion Trap-MS) .......... 45
C. The Comparison of MALDI and E81 .................................................................... 51
D. ESI-CID-MS/MS .................................................................................................. 53
VI. References ................................................................................................................... 60
CHAPTER 2
OPTIMIZATION OF CONDITIONS FOR MONITORING HYDROGEN/DEUTERIUM
EXCHANGE IN PROTEINS BY MALDI-TOF-MS ....................................................... 73
I Introduction ................................................................................................................. 73
11. Experimental Section .................................................................................................. 75
III. Results and Discussion ............................................................................................... 79
A. Complete Reduction of RNase A .......................................................................... 79
B. Effect of Deuterated and Non-deuterated Matrix ................................................. 83
C. Use of a Deuterium-labeled Internal Standard ...................................................... 88
D. Ablation of Sodium Ion Adducts Early in the Analysis at a Given Spot on the
MALDI Target ............................................................................................................ 89
E. Effectiveness of Quenching In-Exchange at pD 2.5 ............................................. 91
IV. Conclusion .................................................................................................................. 92
V. References ................................................................................................................... 93

CHAPTER 3
INTEGRATION OF HYDROGEN/DEUTERIUM EXCHANGE AND
CYANYLATION-BASED METHODOLOGY FOR CONFORMATIONAL STUDIES

OF CYSTINYL PROTEINS ............................................................................................. 96
I. Introduction ................................................................................................................. 96
11. Experimental Section .................................................................................................. 99
III. Results and Discussion ............................................................................................. 103
A. Experimental Design for the Application of HDX-MS to Cystinyl/Cysteinyl
Proteins .................................................................................................................... 103
B. HDX Behavior of Different Segments of LR3IGF-I ........................................... 106
C. Comparison of HDX-MS Results with NMR Data on the Native Structure of
LR3IGF-I ................................................................................................................... 111
D. Some Figures of Merit for the Integrated Methodologies .................................. 114
IV. Conclusion ................................................................................................................ 121
V. Reference .................................................................................................................. 122
CHAPTER 4

CHARACTERIZATION OF PEPTIDE FOLDING NUCLEI BY
HYDROGEN/DEUTERIUM EXCHANGE-MASS SPECTROMETRY AND

CIRCULAR DICHROISM ............................................................................................. 126
I. Introduction ............................................................................................................... 126
H. Materials and Methods .............................................................................................. 129
1]]. Results and Discussion ............................................................................................. 132
A. Criteria for Selection of Peptide Models ............................................................ 132
B. Conformational Study of Selected Peptide Models by CD Spectroscopy .......... 137
C. Study of the Conformational Features of Selected Peptide Models by HDX-
MALDI-MS .............................................................................................................. 143
IV. Conclusions ............................................................................................................... 157
V. References ................................................................................................................. 159
CHAPTER 5
IDENTIFICATION AND CHARACTERIZATION OF AN AUTONOMOUS FOLDING
UNIT FOR LR3IGF-I ...................................................................................................... 163
I. Introduction ............................................................................................................... 163
H. Materials and Methods .............................................................................................. 166
HI. Results and Discussion ............................................................................................. 169
A. Design of the peptide models .............................................................................. 169
B. CD results ............................................................................................................ 172
C. HDX results ........................................................................................................ 177
IV. Conclusion ................................................................................................................ 199
V. Future Work .............................................................................................................. 200
VI. Reference .................................................................................................................. 201

vi

Table 1.

Table 2.

Table 2.

Table 2.

Table 2.

Table 3.

Table 3.

Table 4.

Table 4.

Table 5.

Table 5.

2

l

2

LIST OF TABLES

Comparison of ESI—MS and MALDI-MS [147] ............................................ 52

Calculated and detected m/z value (average) of protonated cleavage products
from RN ase A following complete reduction, cyanylation, and cleavage. . 81

Comparison of observed deuterium incorporation level in reduced (unfolded)
RNAse A using deuterated and non-deuterated matrix. .............................. 84

Comparison of the HDX results using internal and external calibration. ...... 88
Deuterium incorporation in native (folded) RNase A after “quenching”. ..... 91
Summary of HDX results on identiﬁed peptic ﬁ'agments. .......................... 110
Summary of mass shifts for two peptic fragments from different red-ox
species after 10—sec HDX. (Results are the average from three independent
experiments; plus/minus terms indicate the 95% conﬁdence interval). 115
HDX results for oc- and B-subunits as assessed by MALDI-TOF-MS. ....... 144
HDX results for the Pp subunit as assessed by CID-MS/MS. ...................... 153
HDX results of each intact peptide model. .................................................. 178

HDX-CID-MS results for each peptide model. ........................................... 196

vii

LIST OF FIGURES

Figure 1. 1 Cysteine and cystine. ...................................................................................... 3
Figure 1. 2 Conventional proteolytic ﬁagmentation strategy for disulﬁde bond
assignment of a protein. ................................................................................. 4
Figure 1. 3 Disulﬁde/sulﬂrydryl exchange in proteins ...................................................... 5
Figure 1. 4 Conceptual representation of conventional protocol for disulﬁde-bond
assignment based on mass spectrometry ........................................................ 8
Figure 1. 5 Chemical overview of the cyanylation/cleavage strategy. Partial reduction
means that the protein of interest is transformed into a mixture of isofonns,
each of which corresponds to reduction of only one of its disulﬁde bonds;
for a protein containing 11 cysteines, n isomers of the singly reduced protein
will result. Itz stands for iminothiazolidinyl carboxyl residue at the amino
terminus [redrawn from [36]]. ..................................................................... 11
Figure 1. 6 Chemistry involved in the cyanylation and cleavage reactions .................... 13
Figure 1. 7 The formation of a disulﬁde bond upon adding a disulﬁde reagent to a
reduced protein ............................................................................................. 16
Figure 1. 8 Trapping folding intermediates by acidiﬁcation. ......................................... 19
Figure 1. 9 Chemical trapping of folding intermediates by alkylation. .......................... 20
Figure 1. 10 Reversible trapping of folding intermediates by AEMTS ........................... 22
Figure 1. 11 Representative CD spectra of a-helical, B-sheet, and random coil structure
in proteins (redrawn from [89]). .................................................................. 31
Figure 1. 12 Highly charged clusters are formed during disintegration of the
analyte/matrix solid [117]. M stands for the neutral analytes, H” for protons,
and B' for anions (i.e., triﬂuoroacetate). ...................................................... 37
Figure l. 13 Chemical structure of compounds commonly used as matrices for MALDI.
................................................................................................................................... 39
Figure 1. l4 MALDI-TOF-MS instrument [123]. ........................................................... 42
Figure 1. 15 Schematic diagram of ion trajectories in a reﬂectron-TOF m/z analyzer
[129]. ............................................................................................................ 44

viii

Figure 1.

Figure 1.

Figure 1.

Figure 1.

Figure 1.

Figure 1.

Figure 1.

Figure 2.

Figure 2.

Figure 2.

Figure 2.

Figure 2.

Figure 3.

Figure 3.

Figure 3.

16

17

18

19

20

21

22

Schematic of the electrospray ionization process [136]. ............................. 47

ESI-MS instrument interface. ...................................................................... 48
Schematic diagram of ion-trap mass spectrometer [146]. ........................... 50
Conceptual representation of the MS/MS technique (redrawn from [149]).54

Segment of a peptide showing the residues of alanine and lysine linked by
an amide bond, the peptide bond. ................................................................ 56

Conceptual pathway of forming the y-ions by expulsion of residues from the
amino terminus, thereby leaving the charge on detectable fragment ions
retaining the C-terminus (redrawn from [156]). .......................................... 57

Conceptual pathway of forming the b-ions by expulsion of protonated
residues plus the hydroxyl group at the C- terminus, leaving the charge on
N-terminal fragment ions (redrawn from [156]); for convenience, the b-ion
here is shown as an acylium ion, however, the actual structure is believed to
be an oxazolone species ([157, 158]) ........................................................... 58

Conceptual chemical scheme for complete reduction of bovine pancreatic
RNase A, and conﬁrmation of complete reduction by subsequent
cyanylation, cleavage,and analysis of cleavage reaction mixture ................ 80

MALDI mass spectrum of the cleavage products of RNase A. ................... 82

Mass spectra of completely reduced RNase A after incubation with D20 and
then mixed with (A) deuterated matrix or (B) non-deuterated matrix before
analysis by MALDI-TOF-MS ...................................................................... 85

Time course of HDX on reduced RNase A. ................................................. 86

Mass spectra of fully deuterated myglobin (dzéo—Mb) obtained with
deuterated sinapinic acid as matrix (A) Cumulative result from the ﬁrst 50

laser shots; (B) Cumulatlast 50 laser shots. ................................................. 90
Amino acid sequence of IGF-I and LR3IGF-I; the sequence of IGF-I starts at

residue 14 of LR3IGF-I with a Glu at position 16. ...................................... 98
MALDI-TOF mass spectrum of peptic digest of LR3IGF-I. ...................... 108

Average NMR structure of LR3IGF—I [8]. N-terminus and C-tenninus of the
protein are labeled by N and C, respectively. Three helixes are labeled
according to inclusive residues. Three disulﬁde bonds are represented by
dotted lines between the designated cysteines. The dark-gray region is the

ix

Figure 3. 4

Figure 4. 1

Figure 4. 2

structurally ordered part of the protein; the light-gray segments are highly
ﬂexible parts of the protein (in which the heavy atom r.m.s.d. is larger than
the average r.m.s.d. of the ordered part of the protein). The ﬁgure was
drawn using the RasMol software [33]. ..................................................... 112

Mass spectra of (A) the segment 67 — 72 (LRRLEM, 6-mer) from different
redox species and (B) the segment 38 - 55 (FNKPTGYGSSSRRAPQTG,
18-mer) from different redox species. The ﬁrst row shows the mass spectra
of the peptides before deuteration; the second row shows the mass spectra
after sustained HDX to achieve a high degree of deuteration of the peptides.
The remaining four rows are mass spectra of indicated redox species (R:
reduced; l-S: one—disulﬁde intermediate; 2-S: two-disulﬁde intermediate, N:
native) after a 10-s exposure to D20 at pD 6.8. ......................................... 118

Amino acid sequence of BPTI. ................................................................... 134

Ribbon diagram of BPTI [35], with peptides Pm and Pp depicted in dark and
light gray, respectively. The three native disulﬁde bonds are also shown:
Cyss-Cy855 (left), Cysl4-Cys33 (right), and Cy53o-Cy551 (middle). The ﬁgure
was drawn using the MOLMOL software [36]. ......................................... 135

Figure 4. 3 (A) CD spectra of Pap at 0°C, Par; at 60°C and the summed spectrum of PO!

Figure 4. 4

Figure 4. 5
Figure 4. 6

Figure 4. 7

and Pp at 0°C. (B) CD spectra of Pa at 0°C, P,m at 0°C, P,m at 60°C, and the
difference spectrum of P,m and Pa at 0°C. (C) CD spectra of P5 at 0°C, Ppp
at 0°C, PM; at 60°C, and the difference spectrum of P33 and PB at 0°C. (D)
Temperature dependence of the CD signal at 217 nm for Pap. .................. 139

Mass spectra of (A) Pug before HDX, (B) Pap after HDX, (C) Pa after HDX,
(D) PB after HDX, (E) P.m after HDX, (F) Ppp after HDX. The disulﬁde
bond in each dimer was present during HDX, but was reduced before the

mass measurement. The cluster of peaks corresponding to the Pa-chain is
on the left, while that for the PB-chain is on the right in (A) and (B).
Likewise, P.m and P55 were exposed to HDX as dimers, but reduced to the

monomer prior to analysis by MALDI-MS. .............................................. 145
CID-MS/MS spectrum of the PB sequence. ................................................ 151
CID-MS/MS spectrum of the P,x sequence. ................................................ 152

Ribbon diagram of the natively structured PB peptide; Asn24 is indicated in
light gray and relevant hydrogen bonds are shown in dark gray. The
mainchain—mainchain H-bond donor/acceptor pairs from bottom:
Phe33/Argzo, P hezz/ Gln31, Gln31/Phe22, AS1124/L81129, Leuzg/Asn24, and
Gly23/Asn24. The H-bonds between the amide hydrogens of Ly57 and A183

Figure 5. 1

Figure 5.2

Figure 5. 3

to the sidechain atom 0°l of Asns are also shown. The ﬁgure was drawn
using the MOLMOL software [36] ............................................................. 155

HPLC chromatogram showing the time-dependent distribution of CDAP-
trapped intermediates during the refolding of LR3IGF-I [13]. The disulﬁde
structure of each species is: I’ (Cysn — Cys74), II’ (Cysn — Cys74, Cyslg -
Cy36l)a III’ (Cys31 - CYSM, CY519 - Cysw, Cysts: - Cyses), N (Cy331 - CyS74,
CYS19 — Cysm , Cys6o — Cysés), R (completely reduced). (experimental
details on page 99 in Chapter 3) ................................................................ 164

The natively structured P1 and P11 [24]. P. is depicted in dark gray, with its
initial residue, Argn, shown at the far left and its ﬁnal residue, Ser39, shown
in the back on the right. Similarly, P11 is depicted in light gray, with its
initial residue, Arg54, shown at the bottom and its ﬁnal residue, Gln75, shown
at the far right. The crossing-linking disulﬁde bond, CyS31-Cys74, is shown
in the upper right. The side chains of the substituted residues are also shown
in dark gray (in P1) and light gray (in P"). The ﬁgure was drawn using the
MOLMOL software [25]. The side-chains were drawn using SCWRL
software [26]. ............................................................................................. 171

(A) CD spectra of PIP" at 0 °C, 60 °C, 80 °C, and the summed spectra of P1
and P11 at 0 °C. (B) CD spectra of P1P] at 0 °C, 60 °C, 80 °C, and the
spectrum of P; at 0 °C. (C) CD spectra of PUP" at 0 °C, 60 °C, 80 °C, and

the spectrum of P" at 0 °C. ......................................................................... 174

Figure 5. 4 CID-MS/MS spectra of each peptide model: (A) the P1 monomer, (B) the PIP]

homodimer, (C) the P” monomer, (D) the ................................................. 180

Figure 5. 5 CID-MS3 spectra of the two dominant MS/MS ions from P; (A) (B) and Pu

(C) (D). The charge state of doubly charged ions is shown as (+2). ........ 186

Figure 5. 6 CID-MS3 spectra of the two dominant MS/MS ions from P1P“ (A) (B), PIP]

(C)(D), and PnPu (E) (F). The charge state of doubly charged ions is shown
as (+2). ....................................................................................................... 190

xi

ABBREVIATIONS

a—CHCA: a-Cyano-4-hydroxycinnamic acid
l-S: l-disulﬁde

2, 5-DI—IB: 2,5-dihydroxybenzoic acid

2-S: 2-disulﬁde

3-D: 3-dimensional

ACN: acetonitrile

AEMTS: 2-aminoethyl methanethiosulfonate
AFU: autonomous folding unit

BPTI: Bovine pancreatic trypsin inhibitor
BPTI: bovine pancreatic trypsin inhibitor
CAD/CID-MS: collisionally induced/activated dissociation mass spectrometry
CD: Circular Dichroism

CDAP: 1-cyano—4-dimethylamino-pyridinium
CI: chemical ionization

CRM: charge residue model

DTT: dithiothreitol

ESI: electrospray ionization

FA: formic acid

FAB: fast atom bombardment

FT -MS: Fourier transform-mass spectrometry

Gdn-HCl: Guanidine hydrochloride

xii

GSSG: Glutathione

H-bonds: hydrogen bonds

HDX-MS: hydrogen deuterium exchange-mass spectrometry
HPLC: high-performance liquid chromatography

HX: hydrogen exchange

ICR: ion-cyclotron resonance

IGF-I: insulin-like grth factor-I

LC-ESI-MS: liquid chromatography-electrospray ionization-mass spectrometry
LR3IGF-I: Long R3 insulin-like grth factor-I

MALDI-MS: matrix assisted laser desorption/ionization-mass spectrometry
MS: mass spectrometry

N: native

NMR: Nuclear Magnetic Resonance

NMR: nuclear magnetic resonance

NOE: nuclear Overhauser enhancement

PID: photo-induced dissociation

ppm: parts per million

PSD: post-source decay analysis

R: completely reduced

RNase A: Ribonuclease A

SLD: soft laser desorption

TCEP: tris(2-carboxyethyl) phosphine

TFA: triﬂuoroacetic acid

xiii

TOF: time-of-ﬂight

xiv

CHAPTER 1

INTRODUCTION

1. Introduction

Biological macromolecules such as proteins are the “main actors” in the makeup
of life [1]. Mass spectrometry (MS) has long been used as an important tool for the study
of small and medium-sized biomolecules. The revolutionary breakthrough that makes
this technique an indispensable tool for macromolecule analysis is the discovery and
development of two ionization techniques — electrospray ionization (E81) [2] and
matrix-assisted laser desorption/ionization (MALDI) [3]. ESI uses a strong electric ﬁeld
to facilitate spraying the sample, and, thus, produce free-hovering ions [2]; in contrast,
MALDI subjects the sample molecules to an intense laser pulse that releases them as free
ions in the form of protonated molecules [3]. These developments partly made chemical
biology the “leading science” of our time. ESI-MS and MALDI-MS have become among
the most popular tools for the understanding of the basis of life — the identity, 3-
dimensional (3-D) structure, biofunction, and interaction of proteins and other biological
macromolecules [1 ].

Proteins are macromolecules built of combinations of 20 different amino acids
linked together by amide bonds in long chains. They are the major functional molecules
in a living system [4]. For a protein chain to achieve its speciﬁc bioactivity, it must be
folded into a very speciﬁc globular form [5, 6]. This phenomenon naturally gives rise to
the question, “What is the source of the information responsible for this speciﬁc folding

of the peptide chain?” Ever since C.B. Anﬁnsen’s Nobel Prize-winning work that

showed that the linear sequence of amino acids in a protein determines its biologically
active conformation [7], great efforts have been made to understand the rules that govern
the folding (and unfolding) pathway of various proteins. Although different mechanisms
for the folding of various proteins have been proposed during the last few decades, a
general solution to the protein-folding problem is still unknown [8].

The most popular strategy to study the folding mechanism of a protein is to trap
and characterize the folding intermediates that appear during the folding process.
However, such efforts are often hampered because many intermediates appear only
transiently during the course of folding and are therefore difﬁcult to isolate and
characterize [9-11]. In the case of disulﬁde-containing proteins, some intermediates can
be trapped by chemical modiﬁcation of free sulfhydryl groups (-SH) before their isolation
and characterization [12-15]. During the last decade, a cyanylation/cleavage
methodology was developed and applied to study the oxidative refolding of a few
biologically important cystinyl/cysteinyl proteins [16, 17]. In these studies, a picture of
disulﬁde-bond formation along the oxidative folding pathway was proposed based on the
determined disulﬁde structure of each trapped intermediate.

The folding process of cystinyl/cysteinyl proteins involves both disulﬁde-bond
formation and conformational changes. During the last decade, hydrogen/deuterium
exchange coupled with mass spectrometry (HDX-MS) has become more and more
popular in the study of protein conformation and conformational dynamics [18-20]. The
main aim of this dissertation is to describe the application of HDX-MS to study the
conformational changes during the oxidative refolding of cystinyl proteins as a ﬁrnction

of their disulﬁde-bond formation. In Chapter 2, various experimental conditions will be

compared to optimize the conditions for monitoring HDX in proteins by MALDI-MS. In
Chapter 3, using Long R3 insulin-like growth factor-I (LR3IGF-I) as a model protein, the
feasibility and advantages of integrating the cyanylation/cleavage methodology and
HDX-MS to study the oxidative refolding of cystinyl proteins will be demonstrated. In
Chapter 4, the application of HDX-MS to the characterization of small peptides will be
investigated. In Chapter 5, the identiﬁcation and characterization of an autonomous
folding unit (AFU) for LR3IGF-I will be discussed, and the folding process of LR3IGF-I
will be ﬁrrther investigated by designing peptide models and then characterizing these

peptide models using our established methodologies.

II. Disulfide Mapping of Cystinyl/Cysteinyl Proteins

Among the 20 different amino acids that compose proteins, cysteine is the one
with unique properties. In the reduced form, cysteine contains a free sulfhydryl group;
while in its oxidized form, cystine, a disulﬁde bond is formed by linking two sulﬂrydryl

groups together (Figure 1.1).

 

 

C02“ . , H020 _ c0211
2 HS oxrdatron > Y S S
< .
reductlon
NHz NHz NH2

cysteine cys tine

Figure l. l Cysteine and cystine.

The oxidation status of cysteine residues is very important to the bioactivity of a
protein. For example, the free sulfhydryl groups in cysteine proteases usually work as the
active sites for enzyme catalysis; in addition, they are often chelating sites for metals in
proteins [21]. Disulﬁde bonds, on the other hand, usually help to maintain the proper 3-D
structure of a protein, and, therefore, its bioactivity. For instance, although the two
isomers produced during the refolding of insulin-like growth factor-I (IGF-I) bear exactly
the same amino acid sequence and are only different in their disulﬁde structure, they
show signiﬁcant difference in their 3-D structure and bioactivity [22]. Due to the
important roles that free cysteine sulﬂrydryls and cystine disulﬁde bonds play, the
unambiguous determination of disulﬁde structure for a protein is a very important aspect
in protein analysis.

A. Conventional Proteolytic Fragmentation Strategy

Modiﬁcation of ﬁee sulﬂrydryl groups
Enzymatic cleaiiage of a protein
Fractionation of peptides by HPLC
Reduction of disulﬁde bonds
Modiﬁcation of nascent free sulfhydryl groups

Separation of the resulting peptides

Sequencing by Edrnan degradation/mass spectrometry

Figure l. 2 Conventional proteolytic fragmentation strategy for disulﬁde bond

assignment of a protein.

Figure 1.2 illustrates a well-established proteolytic fragmentation strategy for
disulﬁde-bond assignment in cysteine-containing proteins [23, 24]. First, a protein is
cleaved by enzymes between half-cystinyl residues under conditions that minimize
disulﬁde scrambling (or disulﬁde bond exchange, Figure 1.3). The resulting digests are
then separated using reversed-phase high performance liquid chromatography (rp-
HPLC). Finally, the amino acid sequence of the resulting peptides is determined by
Edman degradation or mass spectrometry, and the identiﬁed peptides are related to
speciﬁc segments of the protein. The characteristic aspects of the strategy are discussed

below.

 

   

Figure 1. 3 Disulﬁde/sulﬂrydryl exchange in proteins.

Ideally, the chosen enzymes for cleavage of cystinyl/cysteinyl proteins should
have strict speciﬁcity to produce well-deﬁned fragments, thus simplifying the
identiﬁcation process. Second, the chosen enzymes should have maximum activity under
conditions favoring minimal disulﬁde scrambling. Third, the chosen enzymes should
have the capability to cleave between every half-cystinyl residue to obtain peptides
containing no more than one disulﬁde bond. In practice, however, compromise usually
has to be made among these three requirements. Trypsin has been the most frequently

used enzyme so far due to its high speciﬁcity for Arg and Lys residues. Unfortunately,

the activity of this enzyme is maximized at pH 8.3, and thus, disulﬁde scrambling often
occurs during the digestion [25, 26]. If a protein cannot be cleaved by speciﬁc enzymes,
it must be cleaved by non-speciﬁc enzymes. Pepsin (with maximum activity around pH
3.0), for example, is one of the most popular non-speciﬁc enzymes used for these studies
[27-30]. However, when using non—speciﬁc enzymes, it is difﬁcult to relate the disulﬁde-
bridged peptides to speciﬁc segments of the protein simply by determining their
molecular masses by MS; in this case, more complicated and tedious procedures such as
Edman degradation have to be used to identify the proteolytic fragments.

Following the digestion of a protein, the next step is to purify and to identify each
disulﬁde-bonded peptide. Rp-HPLC under acidic conditions is usually used for this
purpose. This is because this method is rapid and employs conditions where disulﬁde
scrambling is minimized (i.e., a gradient of acetonitrile (ACN) in 0.1% triﬂuoroacetic
acid (TFA)). The puriﬁcation of digests from large proteins to homogeneity may still be
challenging because many peptide fragments may be produced during digestion.

Conventionally disulﬁde-containing peptides can be identiﬁed using HPLC by
comparing chromatograms of digests before and after reduction of the disulﬁde bonds. A
peak corresponding to a disulﬁde-containing proteolytic fragment in the chromatogram of
the unreduced digests should be replaced by two or more new peaks in the chromatogram
of the reduced digest. Each new peak corresponds to a component peptide of the original
unreduced proteolytic fragment. In the case that a proteolytic fragment has an internal
disulﬁde bond, the hope is that there will be at least a small shift in the elution time upon
reduction. The amino acid sequence of the peptides can be conﬁrmed by conventional

methods such as amino acid analysis [31]. This procedure, however, not only is tedious

and time consuming, but also can be unreliable because it depends too much on the
changes of peptide hydrophobicity upon reduction of the disulﬁde bonds.

Modern mass spectrometric techniques such as MALDI-MS and ESI-MS have
made the task of disulﬁde-bond assignment in proteins more tractable [32, 33]. As
shown in Figure 1.4, disulﬁde-linked peptides are identiﬁed based on their molecular
masses, and by comparing the mass spectra of the sample before and after reduction.
Speciﬁcally, in the spectrum of the reduced sample, the signal corresponding to the
disulﬁde-linked peptide(s) is replaced by two signals corresponding to each of the
component cysteine-containing peptides; if a proteolytic fragment contains an intra—
molecular disulﬁde bond, a mass shift of two mass units should be observed. A major
advantage of mass spectrometry in locating disulﬁde bonds in proteins is that it does not
require rigorous puriﬁcation of peptides before identiﬁcation. Moreover, modern mass
spectrometric techniques such as tandem mass spectrometry and post-source decay
analysis (PSD) permit fast and simple sequence identiﬁcation that can conﬁrm a speciﬁc

peptide segment in a protein.

 

Disulﬁde-bonded protein

1 Enzymatic digestion

 

Mixture of peptides
8“ x/
A /;\ \‘S:S7/ s . s
\ ' \\ l + other peptides
\\ \ /
\

\

 

Mass spectrum of unreduced digest

 

/
\ l // / + other peptides

Mass spectrum of reduced digest

SH X/
/ A \‘SH H31 SH. SH
c ' I
\ l I /

\

 

Figure 1. 4 Conceptual representation of conventional protocol for disulﬁde-bond

assignment based on mass spectrometry.

B. Novel Cyanylation/ Cleavage Methodology

There are two major problems in the conventional proteolytic fragmentation
strategy for locating disulﬁde bond linkages. One is disulﬁde scrambling that often
occurs in mild alkaline solutions. Because in practice it is often necessary to use a
speciﬁc enzyme to produce recognizable fragments, and because most speciﬁc enzymes
only work under alkaline conditions, the scrambling problem is usually difﬁcult to avoid
[25, 26]. The second problem is that peptides from which the locations of disulﬁde
bonds can be deduced are not always produced after digestion. This is especially a
problem for proteins containing adjacent cysteines [25, 34]. For instance, trypsin cleaves
speciﬁcally at the C-terminal side of Lys or Arg unless followed by Pro. Therefore, if
two half-cystinyl residues are not separated by Lys or Arg, tryptic ﬁ'agments containing
more than one disulﬁde bond will be produced, and thus unambiguous disulﬁde
assignment is very difﬁcult. In addition, the -Cys-Cys- structure in a protein is usually
resistant to any enzyme [33]. Due to these two problems, disulﬁde structure assignment
by the conventional approach remains a challenge for protein chemists.

The cyanylation/cleavage strategy is a novel analytical approach for the
assignment of disulﬁde bond linkages in proteins, and it is much simpler and more
efﬁcient than conventional methodologies. All steps before blocking of free sulﬂrydryl
groups (and thus, “freezing” the disulﬁde structure of the protein) are under acidic
conditions (pH 3), and, therefore, minimize, if not completely avoid, the problem of
disulﬁde scrambling. In addition, this method is especially useful for highly knotted,

cysteine-rich peptides such as sillucin [35], which are not readily analyzed by

conventional methodology. Figure 1.5 illustrates a chemical overview of the
methodology [36].

The basic idea of the cyanylation/cleavage analytical strategy is as follows:

First, the protein of known sequence is partially reduced by a reducing reagent
such as tris (2-carboxyethyl) phosphine (TCEP) hydrochloride. The nascent partially
reduced protein isomers are immediately cyanylated by 1-cyano-4-dimethylamino-
pyridinium (CDAP) tetraﬂuoroborate. The cyanylation products are then separated and
collected using HPLC. Finally, the peptide is cleaved on the N-terminal side of the
cyanylated cysteines, and any residual disulﬁde bonds that may connect some of the
cleavage products are reduced. A speciﬁc partially reduced isoform usually produces a
speciﬁc set of cleavage products; therefore, the disulﬁde structure of the isoform can be
deduced by identifying its cleavage products, which usually can be achieved simply by
measuring their molecular weights. The chemistries of cyanylation and cleavage
reactions are illustrated in Figure 1.6.

In the last several years, the disulﬁde-bond structure of various peptides and
proteins has been determined by this strategy [35-37]. This, in turn, demonstrated the

feasibility of the cyanylation/cleavage approach under various conditions.

10

Figure 1. 5 Chemical overview of the cyanylation/cleavage strategy. Partial reduction
means that the protein of interest is transformed into a mixture of isoforms, each of which
corresponds to reduction of only one of its disulﬁde bonds; for a protein containing n
cysteines, n isomers of the singly reduced protein will result. Itz stands for

iminothiazolidinyl carboxyl residue at the amino terminus [redrawn from [36]].

11

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

40
1 10 30 I 50
1" S
¢ Partial reduction;
TCEP at pH 3
H H
F 20 F 40 F 20 F 40
1 10 I 30 I 50 1 10 I 30 50
S S SH SH
1 Cyanylation; CDAP at pH 3
CN CN
20 40 20 40
1 10 30 I so 1 10 I 30 I 50
n S SCN SCN
* HPLC fractionation + MALDI-MS
* Cleavage; 1 M NH4OH
1 20 9 11240 1 0 50
I I
11210 S\ 29 1 S\ 19
S S
I I
11230 40 50 11220 30 39
Reduction; TCEP
I 20 9 11240 20 50
Itle 'SH 29 1 [H 19
I“ I”
112 30 40 50 11220 30 39
(Cysro “ Cys3o) (Cyszo ‘ CYS40)
Figure 1. 5

12

- /
Cyanylatlon N C y<:\>—NMe2
pH 3; 25°C, 10' _ _
BF4
CDAP

N=C_S

Cleavage
1 M NH4OH, 1hr, 25°C

/0 “Y S o
C\ II

NH2 H

 

 

truncated peptide iminothiazolidine (itz)-blocked
peptide

Figure 1. 6 Chemistry involved in the cyanylation and cleavage reactions.

13

III. Oxidative Folding of Cystinyl/Cysteinyl Proteins and Trapping of Folding

Intermediates

Protein folding is a subject that has been under investigation for several decades
[8]. Although a few folding mechanisms have been proposed and each of them has some
supporting experimental evidence, none can yet explain all the experimental
observations. Thus, no single folding theory can be regarded as authoritative [8]. At the
current stage, research in this area is still focused on the folding study of individual
proteins, in the hope that investigation on the folding pathway of various proteins will
lead to the ﬁnal establishment of a general folding mechanism. The major difﬁculty in
studies of protein folding is that many folding intermediates are short-lived and/or exist
as a distribution of possible conformers, which usually makes the isolation and
characterization of folding intermediates a challenge [9-11].

Protein folding is the process by which a polypeptide chain acquires the proper 3-
D structure to achieve its biologically active native state [5], and the folding of a
cystinyl/cysteinyl protein involves both changes in its 3-D structure and evolution in its
disulﬁde bond connections. Observations in numerous experiments have illustrated that
during the folding process of cystinyl/cysteinyl proteins, the formation of disulﬁde bonds
is usually coordinated with protein conformational changes [38-42]. In other words,
disulﬁdes reshufﬂe (break and reform) readily in order that a protein (or a segment of the
polypeptide chain) can adopt conformations that are more energetically favorable.
Disulfrde folding intermediates have been proven to be amenable to chromatographic
puriﬁcation, which makes possible the further characterization of these intermediates for

information about the folding pathway [12, 16, 17]. To prevent further disulﬁde

14

rearrangement after the folding process is quenched, folding intermediates are usually
trapped by chemical modiﬁcation of its free sulﬂrydryl groups before separation and
puriﬁcation.

A. Disulfide Formation in Oxidative Folding of Cystinyl/Cysteinyl Proteins

Even when two free sulﬂiydryl groups are in close proximity, they do not form a
disulﬁde bond spontaneously without an appropriate oxidant. Two types of oxidants are
usually used for this purpose — oxygen (with the presence of trace amount of heavy
metals) [43, 44] or a low molecular weight disulﬁde [17, 45]. Both types of oxidants are
commonly used in practice, although oxygen-assisted disulﬁde-bond formation is usually
much slower, and its mechanism is still not clear. When a low molecular weight
disulﬁde is used, it is proposed that the disulﬁde bond formation is a sulﬂ'rydryl/disulﬁde
exchange process between the oxidant and the protein [46]; Figure 1.7 illustrates this
process.

First, one sulﬂiydryl group in the reduced protein is deprotonated to produce a
nucleophilic thiolate anion group. This group (Sf) then attacks one of the sulfurs of the
disulﬁde bond in the low molecular weight disulﬁde. In the transition state, there is
approximately equal bond formation to the nucleophilic sulfur (S 1') and the leaving sulfur
(RLS'). In the next step, a mixed disulﬁde bond forms between the attacking thiolate
anion group (8") and the central sulfur (RCS), and the leaving group (RLS') leaves the
complex as a thiolate anion. The third step is an intra-molecular disulﬁde rearrangement.
In this step, a second sulﬂrydryl group in the reduced protein is deprotonated to form a

new attacking nucleophilic group (S 2'); it attacks sulfur S I in the mixed diSulﬁde bond.

15

 

 

 

 

 

+ ?—
2 SH RC
- ll
88’ s I
1 I 55-
RC
2 SH
1 S—S RL
I + _|
RC 3
2 SH ¢T
, "‘1
RC
2 S-
- II -.
I _$ ..... f5
S RC
_ 2 5‘ J
I S "
| + I
2 S C

Figure 1. 7 The formation of a disulﬁde bond upon adding a disulﬁde reagent to a
reduced protein.

16

Then a disulﬁde bond between sulfurs S2 and S2 forms through a transition state
analogous to that in the second step [46-48].

The observed rate of disulﬁde formation depends on several factors. The rate
increases with pH because sulfhydryl/disulﬁde exchange starts with the attack of a
nucleophilic thiolate anion to an existing disulﬁde bond, and more sulﬂrydryl groups
exist as reactive thiolate anions at higher pH. The rate constant of the nucleophilic
substitution reaction increases as the basicity of the attacking thiolate anion increases or
that of the leaving group decreases. In addition, the stability of the mixed-disulﬁde
intermediate and the tendency of thiolate anions of different cysteine residues to come
into proximity of the mixed disulﬁde (which is mainly controlled by the conformation
adopted by the segment of the polypeptide chain) are also important factors that control
the observed rate of disulﬁde formation [46-48].

Two types of low molecular weight disulﬁdes are usually used, linear disulﬁde
and cyclic disulﬁde. Glutathione (GSSG) and dithiothreitol (DTT) are representative of
each type. The GSSG/GSH (reduced glutathione) system plays an important role in
disulﬁde bond formation in proteins in vivo [49], and they distribute universally in nature
(i.e., in tissues of animals, plants, and microorganisms) [50]. Glutathione is often used in
disulﬁde bond formation in vitro; however, the formation of signiﬁcant amounts of non-
native stable mixed disulﬁde species usually complicate the experiments [17, 45]. DTT,
on the other hand, produces energetically unfavorable mixed disulﬁdes that dissociate
rapidly to produce disulﬁde bonds between cysteine residues in the polypeptide chain.

DTT, therefore, is the preferred oxidant in some oxidative folding experiments [45].

17

Once a disulﬁde bond is formed during the folding process, it can undergo intra-
molecular rearrangements via a process analogous to the intramolecular step illustrated in
Figure 1.7. The rate at which such rearrangements transpire is determined by the extent
to which the conformation of the polypeptide chain favors a nucleophilic attack of a
thiolate anion on the existing disulﬁde bond (Figure 1.3). Therefore, this step provides
very useful information about protein conformation and folding.

B. Trapping of the Folding Intermediates

' To study the oxidative refolding mechanism of a cystinyl/cysteinyl protein, the
folding process is quenched at pre-determined time intervals, and various folding
intermediates are trapped in chemically stable forms that can be separated, puriﬁed and
characterized. Both inter- and intra- molecular disulﬁde rearrangements can take place
only if there are attacking nucleophilic thiolate anions in the system. The general idea
behind all kinds of trapping strategies is to deactivate all the reactive thiolate anions in
the system by modifying/blocking them chemically so that they cannot attack any
existing disulﬁde bonds in the system. This prevents any further disulﬁde rearrangement.
A good trapping reagent should be able to block ﬁee sulﬂrydryl groups rapidly and
completely without modifying other sites in the protein [51, 52]. Several common
trapping strategies and reagents are discussed below.
1. Acid Trapping

The pKa of the typical sulﬂrydryl group is near 8.7 [12]. Therefore,
sulfhydryl/disulﬁde exchange can be signiﬁcantly slowed by protonating reactive thiolate
anions, which is usually done by adding acid to the folding solution until its pH value

reaches 3 or even less (Figure 1.8).

18

HS
pH<3 S
»

 

 

 

Figure 1. 8 Trapping folding intermediates by acidiﬁcation.

Acid trapping has been applied to the folding study of several proteins [12, 17, 41,
43, 53, 54]. This method is rapid and occurs at the diffusion control rate (109 M'IS'I)
[55]. Its major advantage is that it does not require the modiﬁcation of the free
sulﬂrydryl groups in folding intermediates, thus conformational changes induced by any
blocking group are not a concern. Another advantage of acid quenching is its
reversibility. In other words, an acid-quenched intermediate can be puriﬁed, and then
further disulﬁde rearrangement or folding of this isolated intermediate can be resumed
after appropriate adjustment of pH. This is an invaluable tool for protein folding study
because it can provide some detailed insight into the folding mechanism that is difﬁcult
to obtain otherwise. This strategy has been applied to a few folding studies [12, 17, 54,
56].

The rate constant for disulﬁde fonnation/rearrangement decreases ten times for
each one-unit decrease in the pH value of the system. Therefore, the major disadvantage
of this method is that it only slows down, but does not completely stop, the dynamics of
disulﬁde bond formation in a cystinyl/cysteinyl protein. Thus, samples should be

analyzed in a reasonably timely manner after the folding process is quenched by

19

acidiﬁcation [55]. In addition, if the disulﬁde structure of an acid-trapped species needs
to be determined, then its free sulﬂrydryl groups must be chemically modiﬁed.
2. Trapping by Alkylation

Alkylating all free thiol groups in the system is by far the most widely used
trapping strategy. Many alkylation reagents, such as iodoacetate, iodoacetarnide, and 4-
vinyl pyridine have been used to block reactive thiolate anions [57-60]. Iodoacetate is
most commonly used. No further sulfhydryl/disulﬁde exchange can occur after all the

free thiol groups in the folding intermediates and the reduced protein are alkylated

(Figure 1.9).

ICH2COO' ———->

   

Figure 1. 9 Chemical trapping of folding intermediates by alkylation.

The half-life for the reaction between 1 mM GSSG and a sterically accessible
sulﬂrydryl group is about 30 s at pH 8.7 [61]. Using 0.2 M iodoacetate, the half-life of
the alkylation reaction of an exposed thiol group is about 0.3 s [62]. The alkylation
reaction is usually much faster than many inter- and some intra- molecular thiol/disulﬁde
exchange reactions. However, in the folding study of some proteins such as bovine
pancreatic trypsin inhibitor (BPTI) and ribonuclease, rearrangement of intermediates

during trapping with iodoacetate has been reported [12, 52]. In these cases, the rate of

20

alkylation is signiﬁcantly slower than that of some intramolecular disulﬁde
rearrangements. In quantitative analysis, this will cause a problem of underestimation of
the intermediates that react comparatively slowly with the alkylation reagent (or
overestimation of those intermediates that react comparatively fast with the alkylation
reagent). A much higher concentration of alkylation reagents can be used to accelerate
the alkylation reaction with the risk that functional groups other than thiols may also be
modiﬁed [63].
3. Trapping by AEMTS

In order to solve the problems of the conventional alkylation reagents mentioned
above, a novel reagent, 2-aminoethyl methanethiosulfonate [(NH2)C2H5SSO2CH3]
(AEMTS) was proposed [51, 64]. This reagent reacts with free sulﬂiydryls rapidly, and
one or two minutes are usually long enough to completely alkylate all the free thiol
groups without modifying other sites in a protein. Similar to the case of acid quenching,
this reagent blocks thiol groups reversibly, and thus, the folding process can be resumed
for an isolated intermediate. In addition, AEMTS brings one positive charge to each free
thiol group in an intermediate (Figure 1.10); therefore, the more free thiol groups a
disulﬁde species has, the more positive charges it bears after modiﬁcation. This has been
exploited in the separation of complicated folding intermediates by ion-exchange

chromatography [65, 66].

21

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22

AEMTS has been widely used in the folding study of cystinyl proteins in recent
years. Characterization of the 3-D structure of folding intermediates trapped by this
reagent has also been reported [67]. In this case, it seems that this rather large blocking
group, the 2-aminoethyl methanethiol group [(NH2)C2H5S-], did not signiﬁcantly effect
the 3-D structure and bioactivity of the protein. However, it should still be a concern
whether or not this reagent is the best choice if the study focuses on the 3-D structure of
folding intermediates.

4. Trapping by CDAP

The cyanylation reagent, CDAP, has also been used to trap various folding
intermediates in a few studies [16, 17]. After pre-determined time intervals, the folding
process is quenched by adding 20-fold more CDAP reagent to the solution and adjusting
the pH value is adjusted to 3 by the hydrochloric acid (HCl) in the CDAP solution. The
trapped folding intermediates then can be separated and puriﬁed using r'p-I-IPLC, and
their disulﬁde structure can be determined using the cyanylation/cleavage methodology
(see section II. B.).

The major advantage of this method is that the cyanylation of free sulﬂrydryl
groups in the intermediates not only stops any further disulﬁde exchange, but is also the
ﬁrst step for disulﬁde assignment based on the cyanylation/cleavage methodology. The
cyano- (CN-) group is rather small compared with most blocking groups, and it has been
reported not to interfere with the 3-D structure of the trapped intermediates [68]. Thus,
CDAP is a good choice if the study is aimed at investigating conformations of the folding

intermediates.

23

Using CDAP, the cyanylation of all free sulﬂrydryl groups is usually complete
within 5 to 10 minutes. Thus, similar to the conventional alkylation reagents (as
discussed in section 111.82.), the rather slow blocking reaction could cause bias in

quantitative analysis.

IV. Conformational Study of Proteins and Peptides

A polypeptide chain must take a certain 3-D structure for the protein to have its
proper bioactivity [6]. During the folding of a cystinyl/cysteinyl protein, the
conformation of the polypeptide chain usually changes cooperatively with its disulﬁde
bond formation [38-42]. Therefore, conformational study of the trapped intermediates is
an indispensable aspect in the folding study of cystinyl proteins. Some commonly used
tools for protein conformational studies are discussed below.
A. Conventional Technologies in Protein/Peptide Conformational Studies
1. The X-ray Diffraction Methods

X-ray crystallography is among the most powerful and commonly used tools in
the investigation of protein 3-D structure. In X—ray crystallography, a crystal of the
molecules to be visualized is exposed to a collirnated beam of monochromatic X-rays,
and the consequent diffraction pattern is recorded by a radiation counter or on
photographic ﬁlm. The distribution of the electron density, and, therefore, the structure
of the crystal, may then be calculated from the diffraction pattern by Fourier
transformation [69, 70].

X-ray crystallography has been the most important method in enzyme studies

since it has provided the experimental tools to provide our current knowledge of the 3-D

24

structure of proteins [69]. By May 2002, about 80% of atomic coordinate sets deposited
in the Protein Data Bank had been determined by this technique [1]. At low resolution (4
to 6 A), the electron density maps usually can only provide information on the overall
shape of the molecule; on the other hand, with a high resolution of about 1.9 A and very—
well ordered crystals, the positions of atoms can be ﬁtted with an accuracy of :t 0.2 A
[69]. The conformation of numerous proteins has been characterized using this
technique, and it is almost always the ﬁrst choice when only the static 3-D structure of a
protein needs to be elucidated [71].

Crystallization of the protein molecules of interest is an uncircumventable
requirement of X-ray crystallography; thus, for those proteins whose crystals are still not
available, other methods have to be used to characterize their conformation [72]. In
addition, the 3-D structure of a protein in the crystalline state could differ signiﬁcantly
from that in solution state [71, 73]. Because both in vitro and in viva reactions usually
take place in a solution environment, the appropriateness of explaining observed
biological phenomena using the static protein structure obtained from X-ray
crystallography is always a concern. Furthermore, X-ray crystallography is not suitable
for investigation of various dynamic processes such as those that occur during protein
folding [71, 73, 74].

2. Nuclear Magnetic Resonance (NMR) Spectroscopy

Nuclear magnetic resonance (NMR) refers to the absorption of electromagnetic
radiation of a speciﬁc frequency by certain atomic nuclei placed in a static magnetic ﬁeld
and simultaneously exposed to electromagnetic irradiation with certain frequency [75-

78]. The NMR phenomenon results from the quantum mechanical property of nuclear

25

spin. Only those nuclei with nuclear spin I :e 0 (i.e., the nuclei of 1H or 13C) can absorb
electromagnetic radiation under certain conditions, and, thus, produce a NMR signal.
Common biological nuclei such as those of 12C, and I°O have I = 0, and, therefore, are
not detectable by NMR [77].

NMR spectroscopy is the use of the NMR phenomenon to study physical,
chemical, and biological properties of compounds [75]. The development of high-ﬁeld
NMR methods is the most important advance in protein structural determination in recent
years [79]. Currently, NMR and X-ray crystallography are the major tools for
conformational studies of macromolecules such as proteins [1].

The local magnetic ﬁeld in a nucleus is sensitive to the types and closeness of its
neighboring nuclei. The strategy of solving 3-D structures by NMR is to assign the many
resonances in an NMR spectrum to their respective nuclei in the macromolecule and,
thereby, determine which atoms are close to each other. The overall 3-D structure of a
protein is calculated principally from constraints on the distances between the nuclei
under investigation [79]. Some NMR parameters useﬁrl for this purpose, as well as
different NMR methods, are discussed below.

In a one-dimensional experiment, the sample is subjected to a constant magnetic
ﬁeld and to irradiation by a radio frequency electromagnetic ﬁeld to generate an NMR
spectrum consisting of a series of resonance lines that vary in many ways, including their
positions and ﬁne structure. The location of a resonance line is deﬁned by a “chemical
shift” that depends on the chemical environment around the nucleus of interest. This
parameter is measured in parts per million (ppm) as a fraction of the static magnetic ﬁeld

by which the signal is displaced from that of a reference compound. The spins of nuclei

26

that are separated by a small number of covalent bonds (5 3) interact and split the
resonance lines. These “through-bond” effects are called spin-spin (or J) coupling and
are the cause of ﬁne structure in an NMR signal.

Nuclear Overhauser effect refers to NMR signal intensity changes produced by
dipolar cross-relaxation from neighboring spins by radio frequency perturbation of the
spin population [80] (named after American physicist Albert Overhauser, who
hypothesized it in the early 19505), and the degree of increase in the signal is known as
the nuclear Overhauser enhancement (NOE) [81]. This parameter measures “through
space” interaction between nuclei. The effect is caused by the coupling of dipoles, the
magnitude of which falls off as r'° (r is the inter-atomic distance between nuclei). Thus,
information about inter-atomic distance between nuclei close in space can be obtained by
studying this parameter. NOEs are usually classiﬁed as “strong”, “intermediate”, or
“weak”. The assignments are made through identiﬁcation of the “through-bond” and
“through-space” interactions. Observation and quantiﬁcation of NOE effects between
nuclei in close proximity are the basis of determination of protein conformation by NMR
[1, 79].

In multidimensional NMR, the sample is irradiated by two radio frequency ﬁelds
in various sequences and pulses [1, 74, 79]. The 3-D structure of a protein is deduced
from a combination of the restraints on the distances between nuclei and energy
calculation. Certain mathematical methods (i.e., a method based on metric matrix
distance geometry [82]) are used to calculate the 3-D structure for a protein based on the
structural constraints obtained from NMR experiments, and a family of possible

structures is usually obtained. In regions where considerable consistent information has

27

been obtained, the different solutions usually give essentially superimposable structures;
on the other hand, the regions that seem particularly disordered are often highly mobile.
The dynamics of a protein molecule is described by parameters such as the varying
mobility, timescales, and amplitudes of motions of the mobile segments along the
polypeptide chain. Knowledge of the conformation and the dynamics of a protein are
essential for understanding how the protein molecule interacts with other molecules in its
environment [ 1 ].

The major advantage of NMR, compared with X-ray crystallography, is its ability
to yield insight into the dynamics of a molecule [1, 71, 73, 74]. In addition, NMR can be
used to characterize the 3-D structure of proteins that fail to crystallize. However, NMR
analysis is usually limited to moderate-sized proteins in solution (typically < 50 kDa)
[83]. The quality of NMR data is usually compromised if the sample contains
paramagnetic ligands [84]. Moreover, NMR analysis is very time-consuming, especially
compared with the techniques discussed below.

3. Circular Dichroism (CD) Spectroscopy

Plane-polarized light is composed of two vectorial components that are circularly
polarized: one is right-handed (R) and polarized in a clock-wise direction; the other is
left-handed (L) and polarized counterclockwise. When plane-polarized light is shone
through an optically active sample at a wavelength at which the light is absorbed, the left-
and right-handed circularly polarized light may not be equally absorbed by the sample.
In other words, the L and R lights have different molar absorptivities, £2 and ﬁg,

respectively. This phenomenon is called circular dichroism [85].

28

The molar absorptivity (a) is deﬁned by Equation 1.1:

5 = — (Equation 1.1)

The CD is the difference in molar absorptivity, As.
A6: 82 - 63 (Equation 1.2)
Thus,

_ (AL — AR)
cl

Ar: (Equation 1.3)

Where AL and A R are the absorbance of the left- and right-handed polarized light,

respectively; c is the concentration of the solution, and l is the length of the light path. A L

— AR is usually in the region of 0.005% to 0.1% of A2 or AR. The unit of As is cm M'l or

103 cmzmol'I. The output of many CD instruments is in ellipticity, 9, measured in
degrees.

6= 33(AL — A R) (Equation 1.4)

In most scientiﬁc papers, CD data are reported in mean residue weight ellipticity,

[@IMRWI

[@]mw = % (Equation 1.5)
c

Where N is the number of residues in the protein [85].

The backbone of a protein is optically active in the far ultraviolet range (170 —
250 nm), and speciﬁc secondary structures produce characteristic CD spectra. For
example, the CD spectrum of or-helical structure is typically characterized by strong
negative signals at around 208 nm and 222 nm, and a strong positive signal at around 192

nm. The characteristic CD signal of B-sheet structure is a positive signal near 195 nm

29

and a negative signal near 217 nm [86]. Typical CD spectra of a-helical and B-sheet
structures are shown in Figure 1.11. In addition to qualitative analysis, in principle,
quantitative results such as the amount of the helical content in a polypeptide chain can
be obtained by comparing the ellipticity of the sample to a standard reference at a
characteristic wavelength for helical structure.

In addition to the contribution from the backbone of a protein, signals from
natural chromophores sometimes also can be observed in a CD spectrum. This is because
some natural chromophores exhibit idiosyncratic CD behavior when placed in a
heterogeneous environment in a folded protein. For instance, tryptophan can give CD
signals in the near-ultraviolet region (~270 — 300 nm) [85], and tyrosine in the 270 — 300
nm region [85] as well as in the far-ultraviolet region (~230 nm) [87]. CD spectroscopy
has been widely used in conformational study of proteins and peptides [88]. It is a
convenient tool for investigation of the 3-D structure of a polypeptide chain, as well as
conformational changes of a protein upon binding or during the folding process. The
major advantage of this technique is that CD experiments are simple, straightforward, and
fast. However, the resolution of this technique is limited by the size of the polypeptide
chain under investigation; in other words, there is no way to determine the location of the
structure along the entire polypeptide chain solely based on CD results. In addition,
although a-helical structure usually produces a strong and distinctive CD signal, the CD

technique is not a highly effective tool for the study of B-sheet or other secondary

structure. This is because the CD signal of B-sheet and other secondary structure is
usually much weaker and less distinctive [85]. Moreover, the accuracy of quantitative

CD analysis is usually disappointing. CD spectroscopy is often used for preliminary

30

experiments to obtain a rough idea of a protein’s conformation before starting more

detailed and more time-consuming analysis such as NMR analysis.

   
  

or-helix

B—sheet
Random coil

  

 

 

190 200 210 220 230 240 250
Wavelength (nm)

 

Figure 1. 11 Representative CD spectra of a-helical, B-sheet, and random coil structure

in proteins (redrawn from [89]).

B.

HDX-MS

The so-called hydrogen/deuterium exchange technique has for a long time been

used to study the 3-D structure of proteins and peptides, as well as their conformational

changes [18-20]. Among all hydrogens in a polypeptide chain, those bound to carbon

atoms

essentially do not undergo exchange, while those bound to oxygen, sulfur, or side-

31

chain nitrogen undergo isotopic exchange too rapidly to be measured by current
analytical techniques. In contrast, hydrogens located on the nitrogen atoms in the peptide
amide linkages exchange with deuterated solvent at a rate suitable for analytical
measurement. When a protein is dissolved in D20, amide hydrogens are replaced slowly
by deuteriums if they are located either in well-deﬁned secondary structures such as or-
helixes and B-sheets (where the amide hydrogens are usually involved in intramolecular
hydrogen bonds (H-bonds)), or in the hydrophobic core of the protein (where access to
solvent is limited). In contrast, amide hydrogens located at unfolded regions exchange
much more rapidly due either to weakened H-bonds or to much better access to the
deuterated solvent. Thus, conformational information of a polypeptide chain can be
obtained by assessing the deuterium incorporation level of the protein after HDX; the
fewer deuteriums incorporated by the polypeptide chain (or a segment of the polypeptide
chain), the more folded the polypeptide chain (or the segment) is, and vice versa [90].
When an amide hydrogen is replaced by a deuterium, its corresponding NMR
signal disappears. Thus, NMR can be used to monitor deuterium incorporation. For
more than two decades, HDX has been coupled with NMR in many folding studies to
investigate conformational changes of proteins and peptides [91]. The resolution of
HDX-NMR is at the single amino acid residue level. This method usually requires
milligram-scale samples for analysis; however, in protein folding studies to harvest
folding intermediates at that scale is not always possible. Its other disadvantages include
molecular mass limitations (usually less than 50 kDa) and difﬁculties in analyzing

complexes containing paramagnetic ligands [92, 93].

32

The atomic mass of hydrogen is ~1 Da, while that of deuterium is ~2 Da.
Therefore, a mass shift of ~1 Da should be observed if an amide hydrogen is substituted
by a deuterium. MS has long been applied to measure molecular weights and mass
changes of proteins and peptides [94, 95]. In recent years, HDX-MS has been
successfully used to study the conformational changes in a number of protein systems
[18-20]. Due to the superior detection limit of MS, the sample quantity required for
HDX-MS is at least a thousand times less than that for HDX-NMR analysis. In many
cases, this feature permits the study of intermediates of very low concentration, which
might be very important for refolding kinetics. In addition, the upper mass limit of this
approach is yet to be established, and paramagnetic ligands present no difﬁculty to MS.
In HDX-MS, conventionally the deuterated protein is often dissected by proteolytic
digestion before the assessment of deuterium incorporation. (Pepsin is usually the
enzyme of choice for reasons that will be discussed in Chapter 3). Compared with HDX-
NMR, the major disadvantage of HDX-MS is that the resolution of this methodology is
normally limited by the size of peptic fragments (5 — 12 residues, on average) [92].
However, in a few cases, the resolution has been improved to the single amino acid
residue level with the application of tandem mass spectrometry (i.e., collisionally induced
dissociation-mass spectrometry/mass spectrometry (CID-MS/MS), see details in the next

section) [96-99].

33

V. Desorption/Ionization Mass Spectrometry

Since J .J . Thompson’s pioneering description of the ability to separate molecules
based on different mass and charge in 1912, many different theories were proposed and
various types of mass spectrometers were invented. All MS process consists of a
chemical preparation of charged molecules in the gas phase and then physical separation
of ions in vacuum. Although MS has long been used in the analysis of small and
medium-sized molecules, the goal of analyzing large macromolecules remained elusive
for over 70 years. The efforts were hampered by the idea that one always had to
volatilize the molecules ﬁrst and then ionize them, and by the fact that many
macromolecules such as proteins are polar and thermally labile, and, thus, are not suitable
for thermal volatilization. The major problem to be solved was how to produce intact
gas-phase ions of a molecule from compounds that are usually solids and that degrade or
decompose when heated [1].

The use of chemical ionization (CI) made it possible, for the ﬁrst time, to ionize
thermo-labile biomolecules [100]. In CI, a reagent gas is ﬁrst ionized to form abundant
reagent gas ions, which then collide with sample molecules for proton transfer. Another
technique, plasma desorption, uses high-energy ions to desorb and ionize molecules of
interest [101]. A milestone was passed when the fast atom bombardment (FAB)
technique was reported, where a non-volatile chemical protection environment (the
matrix, often glycerol) was used to dissolve the sample molecules to enable thermally
labile compounds to survive the ionization process [102]. Despite some success with
these techniques in mass determination of small biomolecules (molecular weight < 10

kDa), the challenge of how to analyze high-molecular—weight compounds by mass

34

spectrometry and to make mass spectrometry into a powerful detector for liquid
separation techniques was not solved until the late 19805 when two novel ionization
techniques, MALDI [3] and ESI [2], were reported. These soft desorption/ionization
techniques have overcome the problem of the desorption and ionization of large and
thermo-labile biomolecules, and, thus, have become indispensable tools for the
identiﬁcation and characterization of these compounds. The last decade has seen
explosive grth in the applications of these two techniques in biochemical sciences [l].
A. Matrix-Assisted Laser Desorption/Ionization Time of Flight Mass

Spectrometry (MALDI-TOF-MS)

The use of laser light as an energy source to volatilize/ionize macromolecules was
the focus of several key research groups in the 19805 [1]. In late 19805, M. Karas and F.
Hillenkamp showed that a UV-absorbing matrix could be used to volatilize small analyte
molecules [103], and the major breakthrough in the application of laser desorption to
large biomolecules was reported by K. Tanaka in 1987 [104, 105]. Tanaka presented the
ionization of proteins (12 — 35 kDa) using a low-energy (nitrogen) laser and a matrix of
glycerol containing colloidal particles. The technique developed by Tanaka is called soft
laser desorption (SLD). The currently predominant MALDI technique is a fast growing
version of SLD, and its application to proteins was reported shortly after Tanaka’s initial
breakthrough. The key to the MALDI process is to incorporate the analyte
macromolecules in a low-molecular-weight crystalline matrix whose absorption
maximum matches the wavelength of the laser pulse. The matrix molecules absorb

energy from the laser pulse and transfer the energy to the analyte molecules, which

35

results in a soft desorption process of the analyte molecules; in other words, little or no
fragmentation/decomposition of the analyte molecule occurs during desorption [106].

The major advantage of the MALDI technique is its ability to transfer intact low-
charged macromolecules to the gas phase as protonated species. In addition, it also yields
good results with contaminated samples. Although the mechanism of sample desorption
and ionization has not been completely understood, MALDI, especially when combined
with time-of-ﬂight mass spectrometric detection, has found numerous applications in the
study of biological macromolecules.

1. The MALDI Process

Despite the widespread of analytical applications of MALDI, the mechanism for
ionization and desorption of the analyte is still under investigation [107-116]. A
comprehensive and general model for MALDI was proposed in recent years that can
successfully explain some of the major phenomena observed in experiments [117]. It is
assumed that the charge state of the analytes as acquired in solution due to solvent and
pH is roughly conserved within the matrix crystals, and that some water is preserved in
the surrounding of the charge site of the analyte, and, thus, counter ions may be only
roughly associated with the analyte charge site to give overall electroneutrality. One role
of the matrix is to separate the analyte molecules (by dilution) to prevent analyte-analyte
molecular (or ionic) interactions during the ionization process. The key role of the matrix
is in absorbing the radiation, thereby protecting the analyte from radiation damage.
Provided that the energy density achieved is large enough, a sudden and explosive phase
transition occurs, which is best described by the formation of a gas jet of matrix neutrals,

which entrains analyte molecules. Initial species formed as a result of laser

36

desorption/ablation are clusters. They consist of matrix, analyte and ionic species
embedded in the matrix crystal, all held together by H-bonds and coulombic interactions.
These clusters are highly excited owing to the laser energy absorbed by the matrix and
contain enough excess energy to evaporate/desolvate to molecular and ionic species
either directly by evaporation of neutral species or after chemical reactions resulting in
more stable neutral or ionic species. The charging and, thus, ionization process is the

statistical occurrence of clusters with a deﬁcit/excess of anions or cations (Figure 1.12).

 

0 Matrix
O Analyte

Figure 1. 12 Highly charged clusters are formed during disintegration of the
analyte/matrix solid [117]. M stands for the neutral analytes, H+ for protons, and B' for

anions (i.e., triﬂuoroacetate).

37

The initially generated charged clusters evaporate on a very short time-scale by
loss of neutral matrix molecules or neutral reaction products. Upon matrix evaporation,
ion—ion and neutral—ion chemical reactions take place, e. g., proton-transfer reactions and
proton neutralization to neutral species; the successive steps of evaporation (e.g., HCl
from C1' and xH+) are driven by the reaction enthalpy.

2. Matrix and Sample Preparation

The choice of proper matrix is the key to successful analysis of high-molecular-
weight macromolecules using the MALDI technique. The chosen matrix should exhibit
maximum absorbance at the wavelength of the laser used. The low-energy nitrogen laser
(337 nm) is the most widely used in commercially available MALDI mass spectrometers.
Most currently available matrix compounds show strong UV absorption around this
wavelength. In addition, the chosen matrix must have good solubility in the solvent
required for dissolution of the analyte and be inert to the analyte and solvents. It must be
miscible with the analyte in the solid phase, and be vacuum compatible (with low vapor
pressure), and possess a chemical composition that promotes the ionization of matrix
substituents that can donate protons to the analyte. Moreover, the proper matrix should
crystallize readily and have a low heat of sublimation [118].

According to these criteria, among the hundreds of organic compounds that have
been investigated, only a few are suitable for peptide and protein analysis [119, 120].
The structure of some of the most widely used matrices are shown in Figure 1.13.
Empirically a-Cyano-4-hydroxycinnamic acid (or-CHCA) is usually the ﬁrst choice for

proteins/peptides less than 10 kDa, 2,5-dihydroxybenzoic acid (2, S-DHB) for peptide

38

analysis, and 3,5-dimethoxy—4-hydroxycinnamic acid (sinapinic acid) for proteins larger

than 10 kDa [121].

OH
CO H
\CN
HO
OH
or-CHCA 2,5-DHB
CH=CH-C02H
CH3. OCH3

0H

Sinapinic acid

Figure 1. 13 Chemical structure of compounds commonly used as matrices for MALDI.

Several sample preparation methods have been developed for various analytes.
Some of the most widely used methods are discussed below. In the dried-droplet method
[103], a droplet of the matrix solution (1 —— 10 mmol) and a droplet of the solution

containing the protein analyte (1 -— 10 umol) are dried on the sample plate (usually a

39

piece of metal). A common problem with this method is the aggregation of higher
amounts of analyte/matrix crystals in a ring around the edge of the drop; these crystals
are usually heterogenous and irregularly distributed. Another problem often observed
during crystallization is segregation, which refers to the exclusion of salts and some of
the analyte from the matrix as the solvent evaporates and the matrix crystallizes.
Component segregation generates a heterogeneous mixture of analyte throughout the
solidiﬁed sample, which results in highly variable analyte ion production as the laser
moves across the sample surface. The “dried-droplet” method is the oldest and simplest
sample preparation method. Despite its many disadvantages, it has remained the most
common method of sample preparation in the MALDI community.

The fast evaporation method is a variant of the “dried droplet” method. This
method was developed in 1994 with the main goal of improving the resolution and mass
accuracy of MALDI measurements [121]. First, a thin layer of the matrix is placed onto
the sample plate. A droplet of the analyte solution is then applied on top of the dried
matrix bed. As the droplet evaporates, it dissolves some of the matrix that then
crystallizes onto the previous layer of matrix. This method, however, does not provide
reproducible sample-to-sample data for peptide and protein mixtures, and, thus, it is not
the ﬁrst choice if the protein/peptide sample contains more than one component.

In the year 2000, a novel method called the ultrathin layer method was reported
by B. Chait [122]. In this method, saturated matrix solution is ﬁrst applied over the
whole surface of a sample plate. The matrix is then gently wiped with a tissue, leaving
only a faint layer of matrix visible as a yellowish reﬂection when the plate is examined at

an angle. The sample is dissolved in saturated matrix solution directly, and a small

40

aliquot (usually 0.5 11L) is spotted onto the pre-coated sample plate. A protein-matrix
crystalline ﬁlm forms readily on the ultrathin layer of previously crystallized matrix,
leaving most of the salts, detergents and other components in the excess liquid to be
removed by vacuum aspiration. The spot can be further washed with dilute acidic
solution to remove residual salts and detergents. This ultrathin-layer method presents a
homogeneous matrix-protein co-crystallized surface, allowing for reproducible data
acquisition over the entire surface area. Compared with the conventional dried droplet
method, this method shows higher sensitivity and accuracy, as well as better resolution.
It is a robust, detergent-ﬁiendly method for a variety of proteins and mixtures including
membrane proteins.

Until today the choice of matrix and sample preparation method is still an
empirical process due to a lack of understanding of the desorption/ionization mechanism.
The quality of MALDI analysis results is controlled by many factors such as the choice
of matrix and solvent, the additive used, and the crystallization conditions employed.
Parallel analyses under different conditions are required for optimization of the MALDI
experimental conditions.

3. Instrumentation

A MALDI ion source is usually interfaced with a time-of-ﬂight (TOF) mass-to-
charge (m/z) analyzer. Figure 1.14 is a schematic drawing of a MALDI-TOP mass
spectrometer. The sample consisting of co-crystallized analyte and matrix is irradiated
by a pulsed laser that is directed and focused by a prism and optical lens. The intensity of

the laser is controlled by an attenuator, and it is increased gradually until the threshold for

41

 

 

 

 

 

Beam Splitter
QII 55-3:- Laser
/ |

 

 

 

 

 

 

 

 

 

 

 

 

/ / . T '
/ ransrent
ll / Trrgger C: Recorder
/
//
’/
Sample I/ I I I I I-
Probews L 633 @665 603 }
I, \ 5 ‘\ Flight Tube Li
I 1011‘ K ‘\‘ Detector
I SourceI \\
\ Protein
\\
Matrix
Probe Tip

Figure 1. l4 MALDI-TOF-MS instrument [123].

ionization is reached. The interaction of photons with the matrix and analyte results in
the desorption and ionization of the co-crystallized sample/matrix from the metal surface
of the sample plate. A static electric ﬁeld is then applied to the generated ions by
applying a two-stage high voltage (typically i 25 kV, depending on positive or negative
ion detection mode) to the sample probe with regard to a closely spaced accelerating
electrode. The ions are thus accelerated to the same kinetic energy (assuming the initial

kinetic energy is zero) by the electric ﬁeld toward a long (1 — 2 m) ﬁeld-free TOF

analyzer [123].

42

The TOF m/z analyzer is simple, inexpensive, and well suited to the pulsed nature
of laser desorption in MALDI [124]. MALDI can produce ions with very high m/z
values, and the TOF m/z analyzer has virtually no upper mass limit. Therefore, the
combination of these two techniques (MALDI-TOF-MS) is very popular, especially in
the analysis of macromolecules such as proteins and peptides.

The time required for an ion to traverse the ﬂight tube is dependent on its m/z

value and is described by Equation 1.6.

l
TOF = E = L(—"-‘-eV)2 (Equation 1.6)
U 22

Where L is the length of the ﬂight tube, 1) and m are the velocity and the mass of the ion,
respectively, and V is the acceleration potential [125]. According to Equation 1.6, ions
with lower m/z value should have a shorter ﬂight time than heavier ions. Thus, ions with
different m/z are separated into a series of spatially discrete ion packets, each of which
travels with a velocity characteristic of its m/z ratio.

A detector is positioned at the end of the ﬂight tube and produces a signal as each
ion packet strikes it. A TOF spectrum is a record of the detector signal as a ﬁrnction of
time. The difference between the start time, which is triggered by the laser pulse and is
common to all ions, and the arrival time of each ion at the detector is proportional to
(m/z)”. This is the basis for the conversion of ion ﬂight time (TOF) into an m/z value
along the x-axis in a TOF spectrum [125].

Ions of different m/z values produced by a single laser shot arrive at the detector
at different times. Therefore, in contrast to a conventional scanning m/z analyzer, a TOF
mass spectrometer can generate an entire spectrum from every single laser shot; thus, at

least in theory, no information is lost. However, in practice, the spread of initial

43

conditions of ions with the same m/z value, such as the time/location of ion formation and
initial kinetic energy distribution, is usually large, which results in poor resolving power
of the linear TOF m/z analyzer [126].

Two techniques were introduced to solve the problem of poor resolving power,
reﬂectron-TOF [127, 128] and delayed extraction of the ions [126]. The reﬂectron-TOF
consists of two linear ﬁeld-free regions and an ion mirror (Figure 1.15); the ion mirror
compensates for the spread of initial kinetic energy of ions with the same m/z value. Ions
of the same m/z value, but with a distribution of energies, travel at different velocities.
After traversing the ﬁeld—ﬁ'ee region, they reach a series of grids and ring electrodes of
increasing potential (the mirror or reﬂectron) that create a retarding ﬁeld that acts as a
mirror by deﬂecting the ions and sending them back through the ﬁeld-free ﬂight tube.
Ions with more kinetic energy penetrate more deeply into the reﬂector region and spend
more time in that area, while ions with less kinetic energy penetrate less deeply and spend

less time. Thus, both types of ions arrive at the detector at the same time.

Ion source

f;::" it::::

 

% 3%: W

A

 

 

......
......
......
......
......
......
......
......
......
IIIIII
IIIIII
llllll

 

 

 

 

 

Detector Reﬂector region

Figure 1. 15 Schematic diagram of ion trajectories in a reﬂectron-TOF m/z analyzer

[129].

44

The delay time is the interval between the laser pulse, which ionizes the sample,
and application of the acceleration potential. During this time (usually nanoseconds),
ions disperse owing to their initial velocity in the source region (the acceleration region
just above the sample surface). Ions with higher initial kinetic energy move farther into
the extraction ﬁeld than ions having less kinetic energy, and, thus, acquire less kinetic
energy when the extraction ﬁeld is turned on; on the other hand, ions of lower initial
kinetic energy, having moved not as far into the extraction region, traverse a longer
segment of the electric ﬁeld after the extraction potential is applied, and, thus, acquire
slightly more kinetic energy than the other group, which allow them to “catch up” with
the ions of more initial kinetic energy. The “time delay” dramatically improves the
resolving power achievable in MALDI-TOF-MS.

B. Electrospray Ionization Ion Trap Mass Spectrometry (ESI-Ion Trap-MS)

Although the electrospray phenomenon has been studied for over two centuries,
the use of electrospray as an ionization method for macromolecules was introduced by M.
Dole only about four decades ago [130, 131]. He proposed the ﬁrst model of the
electrospray process, including the charge residue model (CRM) that has survived as a
main explanation for the enigmatic ESI process. In 1984, the ﬁrst combination of
electrospray and mass spectrometry was reported by J. Fenn [132]. The well-deﬁned
breakthrough of ESI for MS came in 1988, when he presented an identiﬁcation of intact
proteins of 40 kDa with a molecular-weight accuracy of 0.01% [133]. Since then, the
application of ESI-MS to macromolecule analysis has expanded explosively. Today,
different types of commercial instruments are available, and ESI-MS has become a

routine and indispensable tool in bioanalytical laboratories in both academia and industry.

45

1. The Electrospray Process in ESI-MS

In ESI-MS, an analyte solution is pumped through a capillary biased at a high
potential (2 — 5 kV). This potential provides the electric-ﬁeld gradient required to
produce charge separation at the surface of the liquid. As a result, the liquid protrudes
from the capillary tip and forms a “Taylor cone” (Figure 1.16). Finally, droplets
containing an excess of positive or negative charge leave the tip when the solution that
comprises the Taylor cone reaches the Rayleigh limit, which refers to the point at which
coulombic repulsion of the surface charge exceeds the surface tension of the solution
[134]. These droplets travel through the electric ﬁeld at atmospheric pressure toward the
entrance to the mass spectrometer; the droplets diminish in size due to solvent
evaporation and coulombic explosions with the production of gas phase ions (charged
analyte molecules). A few mechanisms have been proposed to describe this process.

According to the CRM (also known as coulomb ﬁssion mechanism), a charged
droplet evaporates during the electrospray process, and the coulombic repulsion among
the charges on the surface of the droplet becomes larger and larger until a Rayleigh
explosion occurs, and a number of smaller dr0plets whose surface also contains
electrostatic charges are produced. This process takes place over and over again, and the
droplet is divided into smaller and smaller droplets that eventually consist only of single
ions [130]. On the other hand, the ion evaporation theory assumes that the release of ions
directly from the surface of the droplet takes place when the coulombic repulsion is large

enough to overcome the liquid’s surface tension [135].

46

Reduction

 

 

Taylor cone

«5
2 col

 

 

 

 

6?
High Voltage
Power Supply 5
Electrons

* roc

 

 

Figure 1. 16 Schematic of the electrospray ionization process [136].

Regardless of the mechanism by which ions are produced, the ESI process
generates gas-phase ions that can be analyzed for m/z ratio in the mass spectrometer. The
transition of ions from the liquid phase to the gas phase is very gentle, and the analyte
molecule is likely still surrounded by at least one solvation shell when it occurs.
Fragmentation during this event has only rarely been observed, and aspects of the tertiary

structure of a protein might survive into the gas phase under certain circumstances [137].

2. Instrumentation

Figure 1.17 is a schematic diagram of a typical electrospray interface to a mass
spectrometer. The electrospray capillary is typically a metal or glass capillary with a
small i.d. (<100 pm). A high voltage (1 — 5 kV relative to the desolvating capillary) is
applied to the capillary. An analyte solution is forced into the electrospray capillary at a
ﬂow rate of 1 — 100 uL/min and is nebulized by the resulting electrospray plume. A
mixture of ACN and water (or methanol and water) is usually used as solvent, with a
small percentage (< 5%) of acetic acid or formic acid (FA). The protein/peptide
concentration is usually at 10’5 — 10'7 M. The most important application of ESI is as an
interface between a mass spectrometer and separation equipment such as electrophoresis

[138] or HPLC [139].

Ion

Optics

 

 

 

 

Electrospray 4:”: Sklmmer
Capillary $22: ................... .
""51..;EZZZZIIIijj' Heated \ l”\

 

 

 

Capillary

 

 

 

 

+ High Voltage m/z Analyzer

Figure l. 17 ESI-MS instrument interface.

48

Also worth mention is the growing interest in the use of nanospray in LC-ESI-MS
analysis due to the greatly improved sensitivity, a result of the use of very small spraying
oriﬁce (l — 2 pm inner diameter) and very low ﬂow rate usually at nl/min scale).
Nanospray also achieves more efﬁcient ionization and desolvation than does
conventional ESI [140, 141].

Once the ions are formed and sprayed into the heated capillary, they need to be
moved to the m/z analyzer. During this progression, the instrument needs to focus the
ions of interest through several stages of differential pumping of decreasing vacuum
pressure regions to reach the very high vacuum analyzer region. This is achieved in the
assembly of heated capillary, Skimmers, and ion optics. The typical internal diameter of
the heated capillary is between 400 pm and 1mm; the length is usually 10 — 25 cm. Its
purpose is both to complete the desolvation process of the charged analytes and to bafﬂe
atmospheric pressure on the source side to about 1 torr. The heated capillary is
electrically biased (with the same polarity as the ions of interest) to prevent analyte ions
from contacting with the inside walls. In the next step, Skimmers are used to transmit
ions from the heated capillary as much as possible while decreasing the pressure in each
subsequent vacuum stage of the chamber. The Skimmers are also electrically biased to
focus the ion beam and deﬁne its average kinetic energy. The use of ion optics can
ﬁrrther enhance ion transmission and focus the ion beam into the m/z analyzer [142].

ESI is commonly coupled with the ion-trap mass spectrometer [143]. As shown
in Figure 1.18, a quadrupole ion trap consists of a ring electrode and two endcap
electrodes, between which is a small volume to conﬁne ions. A sinusoidal potential (RF

at a ﬁxed frequency) is applied to the ring electrode, while the endcap electrodes is

49

usually maintained at an oscillating (AC) potential. Under a certain storage condition
(speciﬁc instrumental parameters), only ions within a given range of m/z value can be
stored in the ion trap; upon change in the storage conditions, the trapped ions can be
expelled from the trap. Ions striking the detector transfer charge that registers as a
current that is proportional to ion abundance. The high sensitivity and ability to be used
for tandem MS experiments have resulted in the wide use of the ion-trap mass

spectrometer [144, 145].

RF generator

      

Entrance Exit
endcap endcap
Ion Injection .................... ’ .................... > Ion Ejection

Ring
Electrode

L AC generator

Figure 1. 18 Schematic diagram of ion-trap mass spectrometer [146].

C. The Comparison of MALDI and ESI

As shown in Table 1.1, MALDI and ESI have different and complementary
natures [147]. Together, they make a formidable set of tools with high levels of
sensitivity, accuracy, and mass range. They are widely used for routine identiﬁcation and
quantitation of synthesized small molecules and proteins, as well as compounds obtained
directly from biological matrices. In recent years, these techniques also have been
applied to novel ﬁelds such as the study of non-covalent interactions, protein folding, in

vitro drug analysis, and drug discovery.

51

Table 1. 1 Comparison of ESI-MS and MALDI-MS [147].

 

 

 

 

 

 

ESI-MS MALDI-MS
Mass limit, Da >1,000,000 Da > 1,000,000 Da
Ionization Continuous Pulsed
Common mass Quadrupole, quadrupole ion TOF, FTICR,
analyzer trap, FTICR, TOF, quadrupole ion trap
magnetic sector
Advantages HPLC/MS capable Tolerant of mM concentrations
Multiple charging of salts
Capable of observing Highest mass capability
noncovalent complexes directly Tolerant of mixtures
from solution Limitation of detection:
Detection limit: attomole-to- zeptomole-to-femtomole
femtomole
Being developed as a tool for
sequence analysis
Disadvantages Multiple charging can be Typically low resolution

confusing with mixtures
Typically requires < mM salt
concentration for good signals

Not tolerant of mixtures

(< 500) for linear TOF without
delayed extraction
Difﬁcult to couple with LC/MS

 

 

Suitable compounds

 

Peptides
Proteins
Carbohydrates
Nucleotides
Oligonucleotides
Phosphoproteins
Small chargable molecules
Glycoproteins

 

Peptides
Proteins
Carbohydrates
Nucleotides
Oligonucleotides
Phosphoproteins
Small chargable molecules
Glycoproteins
Heterogeneous samples

 

52

 

D. ESI-CID-MS/MS
1. Concept and the Dissociation Process

MS/MS can be thought of as a means by which to obtain the mass spectrum of a
mass spectrum. Among these various modes of ion dissociation, CID is probably most
widely used, especially in ion-trap instruments [145]. Figure 1.19 is a conceptual
representation of CID-MS/MS. First, a given molecule is ionized and allowed to
fragment with mass selection to obtain a conventional mass spectrum (the top spectrum
in Figure 1.19). Second, ions of a particular m/z value (ions with m/z 129 in this case) are
selected as precursor ions for a dissociation process followed by a second mass analysis
to produce the product-ion spectrum (the bottom spectrum in Figure 1.19). The chemical
structure of the precursor ion can be characterized by determining the products of the
dissociation process. As illustrated in Figure 1.19, the key of a MS/MS process is that a
signiﬁcant population of the precursor ions undergoes some form of dissociation between
the two stages of mass selection. In CID, the precursor ion collides with a gas molecule
(usually an inert gas such as argon) to obtain internal energy, which leads to its

decomposition [ 148].

53

Ionization

J m/z analysis

41 MS 1

 

129

[- .;.—. 1

/ / Collision

 

 

 

 

 

\\

 

 

 

 

 

 

 

 

— Dissociation in collision cell—— Cell
‘m/z analysis l
I I MS 2
100 114 129
m/z ’

Figure l. 19 Conceptual representation of the MS/MS technique (redrawn from [149]).

2. Instrumentation

There are two types of MS/MS instrumentation, MS/MS-in-space and MS/MS-in-
time. The ﬁrst type of instrument use a mass-selective process to select a given precursor
ion at a given set of coordinates in space, transfer the ion to a different set of coordinates
for ion dissociation, and transfer the products of that dissociation to yet another set of
coordinates for secondary m/z analysis. The triple-quadrupole mass spectrometer is
representative of instruments that are arranged in this manner. One disadvantage of
MS/MS-in-space is that it requires many different points of ion focus that must be

optimized at the same time to achieve a successful experiment [150].

54

In MS/MS-in-time [151], ions are formed in a given set of coordinates in time,
and then at a later time some of the ions are expelled from those coordinates so that only
the ions of interest remain. At a later time, the residual ions are exposed to some
dissociation process again at the same coordinates, and this is followed by yet a later
episode of m/z analysis so that the entire MS/MS process is conducted in the same
coordinates of space, but at different times. This kind of MS/MS analysis is adopted by
ion-cyclotron resonance (ICR), Fourier-transform (FT) mass spectrometers [152], as well
as the ion-trap mass spectrometers [145, 153, 154]. MS/MS-in-time is much simpler to
operate than MS/MS-in-space because it only has one set of ion optics that must be
optimized and merely controlled sequentially to allow the ions of interest to reside in the

ion volume to achieve the desired results [155].

3. Nomenclature for Classifying N-terminal and C-terminal Fragment Ions of
Peptides

Most proteins are constructed from peptides consisting of 20 common amino acid
residues. As shown in Figure 1.20, the amino acids are condensed into the peptide bond
(an amide bond) by connecting the carboxyl group of one amino acid with the amino
group of the second amino acid. When a peptide is subjected to CID ﬁ'agmentation, b-
ions and/or y-ions are most often observed; these are produced through cleavage at the
peptide bond. A y-ion is one that is formed by cleavage at a peptide bond with charge
retention on the C-terminal end of the molecule (Figure 1.21), while a b-ion is one that is

a fragment of the peptide with charge retention on the amino terminal side of the cleaved

55

amide bond (Figure 1.22). The sequence identiﬁcation of an unknown peptide mainly

depends on recognition of peaks representing a particular series of ions [156].

1’ l’

----- N H-(liH-C—NH — CH — C

CH3 ((I3H2)4
NH2
peptide bond

Figure 1. 20 Segment of a peptide showing the residues of alanine and lysine linked by

an amide bond, the peptide bond.

56

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3. Application

MS/MS and higher stage MSn (n > 2) can be used to great advantage in qualitative
and quantitative analysis by supplying addition information on fragments that can be
related to the structure of a precursor ion (the ion of interest) [159, 160]. Ion trap
instruments have the advantages of high sensitivity and high capacity for fragment ions.
In recent years, ESI-ion trap-MS/MS has become an indispensable tool in biomedical
research [161-165]. It has been widely used in proteomics, as well as in the investigation
of small molecules such as fatty esters and ketones. The coupling of HPLC and ion trap-
MS/MS through the ESI interface made possible on-line separation, and, thus, direct

analysis of biological samples is allowed.

59

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72

CHAPTER 2
OPTIMIZATION OF CONDITIONS FOR MONITORING

HYDROGEN/DEUTERIUM EXCHANGE IN PROTEINS BY MALDI-TOF—MS

I. Introduction

The 3-D structure or conformation of a protein is the key to its bioactivity [1].
Many diseases stem from impairment of structural properties of speciﬁc proteins (e.g.,
muscular dystrophy [2] and Creutzfeld-Jakob disease [3]). The ultimate goal of protein
science is to be able to predict the 3-D structure and biofunction of a protein de novo and
how it will bind to ligands [2]. In spite of considerable efforts over the past 25 years,
principles that govern the transitions of biopolymers from totally unstructured to highly
ordered states remain one of the greatest mysteries in structural biology. Furthermore,
adequate experimental methodology is still being developed to provide a practical and
effective way to determine a protein’s structure and dynamics [4, 5]. Several
experimental methods have contributed to our current understanding in this area, among
which amide hydrogen exchange (HX) has played a particularly important role [6, 7].
About 50 years ago, it was recognized that different amide hydrogens in different protein
structures undergo isotope exchange at different rates [8, 9], which was followed by the
widespread use of hydrogen exchange as a tool to study protein structure and dynamics.
HX has been used to characterize protein folding [IO-12], functionally different forms of
- proteins [13-15], and protein-protein interactions [16, 17].
Among the many different kinds of hydrogens in a protein, only those located at

peptide amide linkages have exchange rates in a range that can be easily measured, and

73

the rates depend on whether they are participating in intra-molecular H-bonding and on
the extent to which they are shielded from the solvent. Because intra-molecular H-
bonding and solvent shielding are directly related to protein structure, it follows that
amide HX rates can be used to probe protein structure [18].

Historically, the monitoring of HX rates depends on qualitative infrared
spectroscopic measurements [19] and quantitative density-gradient measurements [20].
With the new techniques developed in NMR, replacement of amide hydrogens by
deuteriums (HDX) as monitored by NMR became one of the most practical and powerful
tools in protein conformational studies since the 19708 [21]. During the last two decades,
detecting HDX by MS has become a powerful and popular new approach for
investigating protein structure and dynamics due to the unmatched detection limit and the
ability to analyze large proteins (when coupled with peptic digestion). Furthermore, this
approach has the power to decipher heterogeneity within an ensemble of a particular
species, an aspect not readily amenable to conventional techniques [21-23].

Both MALDI-MS [24-26] and LC-ESI-MS [27-30] have been used to monitor
HDX. LC-ESI-MS provides preliminary separation of mixtures, and, thus, relieves the
problem of signal suppression that usually occurs in the analysis of a complicated
mixture such as the peptic digest of a large protein. In addition, the tandem mass
spectrometry technique (MS", 11 > 1), which is routinely coupled with ESI-CID-MS
makes single-amino-acid-residue resolution achievable in HDX-MS. This method,
however, requires signiﬁcant modiﬁcation of the instrument, and the operation of the
instrument requires considerable expertise. HDX-MALDI-MS, on the other hand,

provides a simple and rapid approach to obtaining data about the exchange of deuteriums

74

with amide hydrogens in a protein. No modiﬁcations of the instrument are required, and
the exchange data for many components of a mixture can be obtained from a single
spectrum. The experiments do not require complicated handling, and are usually less
time consuming than similar experiments with LC-ESI-MS. HDX-MALDI-MS has
found broad utility for analyzing HDX of proteins in studies of protein folding and
protein-ligand interactions.

This chapter describes the results of investigating various conditions for protocol
optimization of HDX experiments on proteins. MALDI-MS was used to measure the
mass shift of bovine pancreatic ribonuclease A (RNase A) aﬁer HDX under various
conditions. This protein is a medium-sized protein that consists of 124 amino acid
residues, including eight cysteines in the form of four intra-molecular disulﬁde bonds.
RNase A has considerable secondary structure (three or-helices and six B-sheets) as
stabilized by four disulﬁde bonds [31]; thus, its native and denatured states provide

diverse conformations for our assessment of HDX by MALDI-T OF mass spectrometry.

11. Experimental Section
Materials

RNase A (type I-AS), TCEP hydrochloride, sodium citrate, or-CHCA, CDAP
tetraﬂuoroborate, and TF A (HPLC grade) were obtained from Sigma and used without
further puriﬁcation. Guanidine hydrochloride (Gdn-HCI) and ACN (HPLC grade) were
purchased from Invitrogen Corporation (Carlsbad, CA) and EMD Chemicals Inc.,
respectively. The 0.10 M TCEP solution in 0.10 M citrate buffer at pH 3.0 was freshly

prepared before use.

75

Preparation of Model Proteins

RNase A (20 nmol) was dissolved with 0.1 M citrate buffer, pH 3.0, containing 6
M Gdn-HCl and 0.1 M TCEP reducing agent to a ﬁnal concentration of 0.6 nmol/pL.
The denaturation and reduction of RNase A were carried out at 37 °C for 2 hr. The
reduced protein was puriﬁed by HPLC with linear gradient of 10% to 40% B in one hour,
where solvent A was H20 containing 0.1% TFA, and solvent B was ACN containing
0.1% TFA. The HPLC fraction was collected, dried under reduced pressure, and stored
at —80 °C. The reduced/denatured protein was reconstituted in HzO (pH ~7.0) right
before the HDX experiment. Native RNase A was dissolved and equilibrated in H20 (pH
~7.0) at room temperature for about one hour to achieve an equilibrium distribution of
the native state in solution.
Conﬁrmation of Complete Reduction

To the dried and completely reduced RNase A was added 0.2 M CDAP solution
(in 0.1 M citrate buffer, pH 3.0) to a ﬁnal concentration of l nmol/uL protein. The
cyanylation of free thiol groups by CDAP proceeded at room temperature for 15 min
under nitrogen. The cyanylated protein was then desalted using HPLC and the proper
fraction was dried. To cleave the polypeptide chain on the N-terminal side of each
cyanylated cysteine residue, 4 [1L of 6 M Gdn-HCl in 1 M NH40H aqueous solution was
added to the dried protein to dissolve it, and then 10 uL of 1 M NH.;OH was added.
Cleavage of the polypeptide chain was performed at room temperature for one hour.
Excess ammonia was removed in a vacuum system, and the cleavage products were

analyzed by MS for identiﬁcation.

76

Mass Spectrometry

For the conﬁrmation of complete reduction, MALDI mass spectra were obtained
with a Voyager DE-STR TOF mass spectrometer (Applied Biosystems Inc.,
Frarningham, MA) using an accelerating voltage of 25 kV with the grid voltage at 93%,
guide wire at 0.002%, and extraction delay time at 300 ns. The mass spectrometer was
equipped with a 337-nm nitrogen laser. Time-of-ﬂight to mass conversion was achieved

with the use of internal standards of bradykinin (average m/z ratio for [M + H]+ =

1061.22), bovine pancreatic insulin (average calculated m/z ratio for [M + H]+
5734.56). A saturated solution of or-CHCA, prepared in ACN / water / TFA (50:50:0.05),
was used as the matrix. The sample spots for MALDI were prepared by the “ultrathin
layer method” [32] as described in Section V. A. 1. of Chapter 1.

For the comparison of deuterated and non-deuterated matrix, instrument settings
were the same as described above. The matrix used in these experiments was a solution
containing 5 mg/mL of sinapinic acid in 0.1% deuterated TFA (in D20)/ acetonitrile 1:2
by volume (pD 2.5). The matrix was incubated with D20 overnight before use. The
conventional non-deuterated matrix was prepared exactly the same way as for the
deuterated matrix, except that H20 and non-deuterated TFA were used instead of D20
and deuterated TFA (dl-TFA) (TFA: CF 3CO2H; dl-TFA: CF3CO2D). Time-of-ﬂight to
mass conversion was achieved with the use of external calibration of horse skeletal
myoglobin (average calculated m/z ratio for [M + H]+ = 16,952.56, [M + 2H]2+ =
8476.78).

For all the other experiments using deuterated matrix, a Voyager DE MALDI-

TOF mass spectrometer equipped with a 337-nm nitrogen laser (Perseptive Biosystems

77

Inc., Framingham, MA) was used. The accelerating voltage was set at 25 kV, grid
voltage at 93%, and guide wire at 0.02%. The extraction delay time was 600 ns. The
matrix used in these experiments was in a solution containing 10 mg/mL of sinapinic acid
in 0.1% deuterated TFA (in D20)/ acetonitrile 1:2 by volume (pD 2.5). The matrix was
incubated with D20 overnight before use. Time-of-ﬂight to mass conversion was
achieved with the use of internal or external calibration of fully deuterated myoglobin
(d260-Mb) (average calculated m/z ratio for [d26o-Mb + D]+ = 17214.18, [d26o-Mb + 2D]2+
= 8608.09). The fully deuterated myglobin (d260—Mb) was prepared by incubating
apomyoglobin (1 nmol/uL) in buffered D20 (pD 7.0) at 45 0C overnight. Ion current
from 200 laser shots was accumulated for each spectrum.
Deuterium Exchange Experiments

The deuterium exchange was initiated by diluting 1 [1L of the protein solution (-
750 pmol/uL) with 20 uL D20 containing 20 mM phosphate buffer (pH 7.0). After
different time intervals, the exchange reaction was quenched by adding a 120-uL aliquot
of matrix solution (pD or pH 2.5). An aliquot of 0.5 uL of the resulting solution was
spotted on the MALDI sample plate and analyzed by MALDI-TOF-MS. All pH or pD
values were measured using a Beckman (D 40 pH meter or ColorpHast pH indicator
strips, pH 0-14 (EM Science, Gibbstown, NJ). pD values were calculated by adding 0.4

to the corresponding pH meter readings [33].

78

III. Results and Discussion
A. Complete Reduction of RN ase A

Previously, reduction of the native RNase A was achieved by adding reduced
DTT in pH 8.0 buffer [34]; our strategy was to reduce the protein with TCEP in aqueous
solution at pH 3.0. Complete reduction is possible under such conditions and re-
oxidation of the reduced protein is minimized due to the low pH.

To conﬁrm complete reduction, the reduction product was cyanylated by CDAP
followed by cleavage into the nine cleavage products represented at the bottom of Figure
2.1. Table 2.1 lists the calculated m/z values of expected cleavage fragments and the
experimentally observed values. The MALDI spectrum is shown in Figure 2.2. Seven
peaks corresponding to seven out of the nine possible fragments appear in the mass
spectrum, which conﬁrms the reduced state of seven cysteine residues in the completely
reduced RNase A. The missing two peaks corresponding to fragments with m/z 774.86
and 789.83 (corresponding to fragment Itz-58-64 and Itz-65-71, respectively) lead to
ambiguity in the redox state of Cysés. However, if all the other seven cysteine residues
are reduced in RNase A, Cysos also must be in the reduced state because it would have no
partner for forming a disulﬁde bond. The signal corresponding to these two fragments is
probably suppressed by ionization of the matrix or obscured by peaks representing matrix
ions.

The assignment of some peaks in Figure 2.2 (those labeled with “?”) has not yet
been determined. They probably correspond to products of unidentiﬁed side reactions
occurring during the cyanylation and cleavage process. These experimental results

conﬁrm the complete reduction of RNase A under our experimental conditions.

79

(D

S

i i_ls S

1 26 40 58 65 72 84 95 110 124

 

 

 

 

 

 

l TCEP, 0.1M, pH3, 37°C, 2Hrs

1 26 40 58 65 72 84 95 110 124

l CDAP, 0.2M, pH3, rt, 15min

I] ll

SCN jCN ISCNSCNSCN SCN SCN ISCN
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1

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1-25 , ItzZ6-39 Itz40-57 It258-64 Itz65-71

Itz72-83 Itz84-94 Itz95-109 Itzl 10-124

Expected Cleavage Products

Itz = iminothiazolidinyl carboxyl residue at the amino terminus of the cleavage products

Figure 2. 1 Conceptual chemical scheme for complete reduction of bovine pancreatic
RNase A, and conﬁrmation of complete reduction by subsequent cyanylation,

cleavage,and analysis of cleavage reaction mixture.

80

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B. Effect of Deuterated and Non-deuterated Matrix

As described in most current papers, a matrix solution with H20 and organic
reagents (i.e., ACN) is usually used for MALDI-MS analysis of deuterated proteins [24-
26]. However, H20 is introduced to the deuteration reaction system when the non-
deuterated matrix solution is mixed with the deuterated protein. As a result, a large
portion of the incorporated deuteriums on the protein will be back-exchanged by
hydrogens from H2O or carboxyl hydrogens of the matrix molecules (most matrix
molecules used in MALDI-MS are carboxylic acids). It is assumed that back-exchange
only happens to deuteriums attached to oxygen, sulﬁrr, or side-chain nitrogen, while all
the amide deuteriums incorporated are retained. 0n the other hand, the use of deuterated
matrix, as prepared with deuterated solvents and matrix that has been incubated in D20
overnight, has been reported [35]. The use of conventional non-deuterated matrix
solution and deuterated matrix solution was compared using the ﬁrlly reduced RNase A
as a model protein; the experimental results are discussed below.

A comparison of observed deuterium incorporation using the deuterated and non-
deuterated matrix is shown in Table 2.2. The spectra are shown in Figure 2.3. As
shown in Figure 2.3 (A), using the deuterated matrix, after incubating with D20 (pH 7.0)
for 10 s, the fully reduced RNase A showed an exchange level of over 95%. Longer
incubation time with D20 to even 90 min does not improve the deuterium incorporation
level (Figure 2.4). This implies that under the conditions described above, an equilibrium
of H/D exchange can be achieved within 10 s, which is in agreement with the result
obtained by theoretical calculation [36]. On the other hand, using conventional non-

deuterated matrix, an exchange level of 50% (Table 2.2) was detected as shown in Figure

83

Table 2. 2 Comparison of observed deuterium incorporation level in reduced (unfolded)

RNAse A using deuterated and non-deuterated matrix.

 

 

 

 

 

 

 

 

 

RNase A
Proteins (Completely reduced)
Number of amino acid residues 124 (4 Pro)
Average molecular weight (Da) 13690
Number of amide bonds 123
Number of amide hydrogens 119
(Expected mass shift if the amide hydrogens (120)
are replaced by deuteriums)
Number of side-chain hydrogens 126
Total number of hydrogens 245
(Expected mass shift if all hydrogens (246)
are replaced by deuteriums)
Observed mass shift with deuterated matrix 13927 — 2 - 13690
(Da) = 235
(Percent deuteration) ( _2_3_5 = 95.5%)
246
Observed mass shift with non-deuterated matrix (Da) 13814 - 1 - 13690
= 123
(Percent deuteration) (5% = 50.0%)

 

 

 

 

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2.3 (B). This corresponds to an extent of back-exchange of about 50%, which is also in
agreement with previously reported results in similar experiments [24-26].

The extent of back-exchange is greatly reduced by employing deuterated matrix.
In addition, the use of deuterated matrix facilitates the use of a fully deuterated internal
standard to improve the accuracy and precision of mass measurement (see details in the
following section). The use of non-deuterated matrix with fully deuterated internal
standard is not feasible because it is difﬁcult to control the back-exchange of the fully
deuterated standard once it is mixed with non-deuterated matrix solution. Therefore,
deuterated matrix could be a good choice in the study of intact proteins in systems with a
low concentration of salts. However, if the polypeptide chain is dissected by peptic
digestion before mass measurements, then it is probably more practical to use a non-
deuterated matrix because, otherwise, all the reagents and chemicals involved in sample
treatment after quenching HDX, including pepsin and desalting reagents, should be
completely deuterated to match the use of the deuterated matrix. In practice, it is not
always easy to satisfy this requirement. In the following sections of this chapter, a
deuterated matrix was used in all experiments because all of them are focused on intact
RNase A. In the other chapters, however, the conventional non-deuterated matrix was

used when assessing deuterium incorporation of various peptic fragments of a protein.

87

C. Use of a Deuterium-labeled Internal Standard

In TOF-MS measurements, time-of-ﬂight to mass conversion is usually achieved
with the use of external or internal standards, and the latter approach normally provides
the highest accuracy and precision. In HDX-MS experiments, a fully deuterated internal
standard is preferred over non-deuterated internal standards. This is because the non-
deuterated species may be partly deuterated when it is mixed with the deuterated sample,
and, thus, unpredictable change in the molecular weight of the non-deuterated standard
will result in inaccuracy in mass determination. In this section, we compared the use of
fully deuterated myglobin (d26o-Mb) for internal and external calibration with deuterated
matrix. The results are summarized in Table 2.3. Native RNase A was used as the model
protein in the experiments; it was exposed in D20 for various periods before mass

measurement.

Table 2. 3 Comparison of the HDX results using internal and external calibration.

 

 

Incubation time 10 15 30
(S)
Observed m/z with internal calibration 13879 13883 13883
(standard deviation ) (s = 3) (s = 3) (s = 2)

 

Observed m/z with external calibration 13851 13864 13862
(standard deviation ) (s = 10) (s = 5) (s = 7)

 

 

 

 

 

 

. Each value is the average of ﬁve repeated experiments

88

As expected, the results obtained by using an internal standard have much better
precision than the corresponding results from the use of an external standard. In addition,
the measured m/z value is lower using an external standard rather than an internal
standard, which could be a result of systematic error caused by the use of an external

standard.

D. Ablation of Sodium Ion Adducts Early in the Analysis at a Given Spot on the

MALDI Target

When analyzing relatively large proteins (> 10 kDa) in the low-resolution linear
mode, each isotope peak cannot be resolved, and a single broad peak is usually observed
that consists of all the unresolved isotope peaks. A symmetric bell-shaped peak is
usually required to determine the average mass of the molecule accurately and precisely.
However, in our experiments, a tail in the distribution is usually observed early in the
analysis at a given spot on the MALDI target (Figure 2.5 (A)). The tail might be caused
by Na+-adducts of the protein under investigation (d26o-Mb in this case). Due to the low
resolution of this type of analysis, the Na+-adducts peak cannot be resolved from the
analyte peak. The sodium ions may come from various sources. For example, the H20
used in the experiments was stored in a glass bottle, and a trace amount of sodium ions
could be dissolved from the glass into the water. In addition, the matrix and other
chemicals used may contain a trace amount of salts including some sodium salts. The
sodium ion adducts may cause overestimation of the m/z value of the analyte or poor

reproducibility for repeated experiments.

89

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The sodium ion adducts can be dissipated during the early stage of the laser
irradiation. As shown in Figure 2.5 (B), the peak resulting from the last 50 laser shots, is
signiﬁcantly more symmetric than the one from the ﬁrst 50 laser shots and the tail is
much less obvious in the latter case. In all the experiments reported here, to improve the
accuracy and precision of MALDI measurements, ablation of Na+-adducts at a given spot
on the target was conducted by discarding the spectrum ion current ﬁ'om the ﬁrst 50 laser
shots before starting data acquisition. The smaller peak following the peak of the analyte

likely represents an adduct of the matrix (sinapinic aicd) in this case.

E. Effectiveness of Quenching In-Exchange at pD 2.5

The acidity and temperature have important effects on the HDX rate. The
exchange rate decreases 10-fold with a 20-°C decrease in temperature. The exchange rate
is lowest at around pH 3 [37]. Thus, it is usually assumed that the H/D exchange is
“quenched” after decreasing the temperature to 0 °C and adjusting the pH of the reaction
system to 2.5 [25, 28]. To test this assumption, after adding deuterated matrix to the
HDX mixture of native RNase A, the solution was incubated on ice for different time
intervals before analysis by MALDI-MS. The temperature of the system was decreased
from 25 °C to 0 °C and the pH was decreased from 7.0 to 2.5 upon adding the matrix

solution (Table 2.4).

Table 2. 4 Deuterium incorporation in native (folded) RNase A after “quenching”.

 

 

 

 

 

 

 

 

Time interval 0.25 5 15 60
(min)

Mass shiﬁr 197.8 196.8 197.8 209.7
(Da)

 

'mass shiﬁ is the average of duplicate experiments

91

No signiﬁcant in-exchange (substitution of hydrogens by deuteriums from solvent
during HDX) occurred during a 5-15 min exposure to deuterated sinapinic acid matrix at
pD 2.5 and 0°C. This result is important because to obtain detailed information about
protein conformation, the protein will be digested by pepsin at 0°C, pH 2.5 for 5 — 15
min. The above experimental result indicates that no signiﬁcant back-exchange would be

expected to occur during the digestion process.

IV. Conclusion

0 Internal calibration rather than external calibration should be used whenever
possible to improve the accuracy and precision of assessment of the deuterium
incorporation level by MALDI-TOF-MS. The strategy of using a completely
deuterated protein for internal calibration overcomes the problem of unpredictable
in-exchange of a non-deuterated protein as a standard mixed into a deuterated
matrix.

0 A Na+-adduct of the analyte may affect the symmetry of the mass spectral peak;
the accuracy and precision of the MALDI-MS measurement may be improved by
ablation of Na+-adducts at a given spot on the target before starting data
acquisition.

0 HDX can be effectively quenched by adjusting the pD of the system to 2.5 at 0°C.

92

V. References

[1] T. E. Creighton, in Mechanisms of Protein Folding (Pain, R. H., Ed.), Oxford
University Press, Oxford (1994) pl.

[2] A. Fersht, in Structure and Mechanism in Protein Science: a guide to enzyme
catalysis and protein folding, W. H. Freeman and Company, New York (1999)

pl.

[3] A. L. Horwich, J. S. Weissman, Deadly conformations--protein misfolding in prion
disease, Cell 89 ( 1997) 499-510.

[4] R. J aenicke, Protein self-organization in vitro and in vivo: partitioning between
physical biochemistry and cell biology, Biol Chem 379 (1998) 237-43.

[5] S. E. Radford, C. M. Dobson, From computer simulations to human disease:
emerging themes in protein folding, Cell 97 (1999) 291-8.

[6] S. W. Englander, L. Mayne, Y. Bai, T. R. Sosnick, Hydrogen exchange: the modern
legacy of Linderstrom-Lang, Protein Sci 6 (1997) 1101—9.

[7] T. M. Raschke, S. Marqusee, Hydrogen exchange studies of protein structure, Curr
Opin Biotechnol 9 (1998) 80-6.

[8] K. U. Linderstrom-Lang, in Symposium on Protein Structure (N euberger, A., Ed),
Methuen, London (1958).

[9] K. U. Linderstrom-Lang, in Symposium on Peptide Chemistry, Vol. 2, Chem. Soc.
Spec. Publ. (1955) pl-20.

[10] H. Roder, G. Wagner, K. Wuthrich, Individual amide proton exchange rates in
thermally unfolded basic pancreatic trypsin inhibitor, Biochemistry 24 (1985)
7407-11.

[11] H. Roder, G. A. Elove, S. W. Englander, Structural characterization of folding
intermediates in cytochrome c by H-exchange labelling and proton NMR, Nature
335 (1988) 700-4.

[12] M. F. Jeng, S. W. Englander, G. A. Elove, A. J. Wand, H. Roder, Structural
description of acid-denatured cytochrome c by hydrogen exchange and 2D NMR,
Biochemistry 29 (1990) 10433-7.

[13] A. J. Wand, H. Roder, S. W. Englander, Two-dimensional 1H NMR studies of

cytochrome 0: hydrogen exchange in the N-terminal helix, Biochemistry 25
(1986) 1107-14.

93

[14] A. Muga, H. H. Mantsch, W. K. Surewicz, Membrane binding induces
destabilization of cytochrome c structure, Biochemistry 30 (1991) 7219-24.

[15] S. W. Englander, J. J. Englander, R. E. McKinnie, G. K. Ackers, G. J. Turner, J. A.
Westrick, S. J. Gill, Hydrogen exchange measurement of the free energy of
structural and allosteric change in hemoglobin, Science 256 (1992) 1684-7.

[16] P. Brandt, C. Woodward, Hydrogen exchange kinetics of bovine pancreatic trypsin
inhibitor beta-sheet protons in trypsin-bovine pancreatic trypsin inhibitor,
trypsinogen-bovine pancreatic trypsin inhibitor, and trypsinogen-isoleucylvaline-
bovine pancreatic trypsin inhibitor, Biochemistry 26 (1987) 3156-67.

[17] Y. Paterson, S. W. Englander, H. Roder, An antibody binding site on cytochrome c
deﬁned by hydrogen exchange and two-dimensional NMR, Science 249 (1990)
755-9.

[18] S. W. Englander, Hydrogen exchange and structural dynamics of proteins and
nucleic acids, Quarterly Review of Biophysics 16 (1984) 521-655.

[19] H. Lenormant, E. R. Blout, Origin of the absorption band at 1,550 cm.-1 in proteins,
Nature 172 (1953) 770-1.

[20] A. Hvidt, K. Linderstrom-Lang, Exchange of hydrogen atoms in insulin with
deuterium atoms in aqueous solutions, Biochim Biophys Acta 14 (1954) 574-5.

[21] I. A. Kaltashov, S. J. Eyles, Studies of biomolecular conformations and
conformational dynamics by mass spectrometry, Mass Spectrom Rev 21 (2002)
37-71.

[22] T. E. Wales, J. R. Engen, Hydrogen exchange mass spectrometry for the analysis of
protein dynamics, Mass Spectrom Rev (2005).

[23] J. R. Engen, D. L. Smith, Investigating protein structure and dynamics by hydrogen
exchange MS, Anal Chem 73 (2001) 256A-265A.

[24] J. G. Mandell, A. M. Falick, E. A. Komives, Measurement of amide hydrogen
exchange by MALDI-TOF mass spectrometry, Anal Chem 70 (1998) 3987-95.

[25] G. S. Anand, C. A. Hughes, J. M. Jones, S. S. Taylor, E. A. Komives, Amide H/2H
exchange reveals communication between the cAMP and catalytic subunit-
binding sites in the R(Dalpha subunit of protein kinase A, J Mol Biol 323 (2002)
377-86.

[26] J. G. Mandell, A. Baerga-Ortiz, A. M. Falick, E. A. Komives, Measurement of

solvent accessibility at protein-protein interfaces, Methods Mol Biol 305 (2005)
65-80.

94

[27] J. R. Engen, W. H. Gmeiner, T. E. Smithgall, D. L. Smith, Hydrogen exchange
shows peptide binding stabilizes motions in Hck SH2, Biochemistry 38 (1999)
8926-35.

[28] J. R. Engen, E. M. Bradbury, X. Chen, Using stable-isotope-labeled proteins for
hydrogen exchange studies in complex mixtures, Anal Chem 74 (2002) 1680-6.

[29] J. R. Engen, Analysis of protein complexes with hydrogen exchange and mass
spectrometry, Analyst 128 (2003) 623-8.

[30] Y. Deng, H. Pan, D. L. Smith, That Hydrogen Exchange at Individual Peptide
Amide Linkages Can Be Determined by Collision-Induced Dissociation Mass
Spectrometry, Journal of the American Chemical Society 121 (1999) 1966-7.

[31] A. Wlodawer, L. A. Svensson, L. Sjolin, G. L. Gilliland, Structure of phosphate-free
ribonuclease A reﬁned at 1.26 A, Biochemistry 27 (1988) 2705-17.

[32] M. Cadene, B. T. Chait, A robust, detergent-ﬁiendly method for mass spectrometric
analysis of integral membrane proteins, Anal Chem 72 (2000) 5655-8.

[33] P. K. Glasoe, F. Long, Use of glass electrodes to measure acidities in deuterium
oxide, Notes 64 (1960) 188-90.

[34] D. M. Rothwarf, H. A. Scheraga, Regeneration of bovine pancreatic ribonuclease A.
1. Steady-state distribution, Biochemistry 32 (1993) 2671-9.

[35] J. Villanueva, F. Canals, V. Villegas, E. Querol, F. X. Aviles, Hydrogen exchange
monitored by MALDI-TOF mass spectrometry for rapid characterization of the
stability and conformation of proteins, FEBS Lett 472 (2000) 27-33.

[36] Y. Bai, J. S. Milne, L. Mayne, S. W. Englander, Primary structure effects on peptide
group hydrogen exchange, Proteins 17 (1993) 75-86.

[37] S. W. Englander, A. Poulsen, Biopolymers 7 (1969) 379.

95

CHAPTER 3

INTEGRATION OF HYDROGEN/DEUTERIUM EXCHANGE AND
CYANYLATION-BASED METHODOLOGY FOR CON FORMATIONAL

STUDIES OF CYSTINYL PROTEINS

I. Introduction

HDX-MS has been widely used to assess the conformation of proteins in a variety
of conditions ranging from isolated states to non-covalent interactions [1-5]. The
relatively “open” conformation of folding intermediates compared to that of the native
protein allows greater deuterium incorporation due either to weakened H-bond
interactions or to exposure of normally buried residues to solvent. Thus, the progress of
protein refolding can be monitored by diminished HDX as detected by MS as the
denatured protein recovers to its native structure [5, 6]. In an effort to recognize the
approximate location of the incorporated deuterium atoms along the polypeptide
backbone, after the protein is exposed to HDX, it is digested by pepsin before analysis by
MS. Pepsin is the desirable protease for this purpose because it ﬁmctions at low pH, a
condition that minimizes the HDX rate, thereby helping to preserve whatever deuterium
atom incorporation was achieved during HDX; also important is the fact that most peptic
fragments are small (5 — 12 residues, on average), thereby allowing reasonably good
resolution of the sites of deuterium incorporation along the polypeptide backbone. NMR
has also been used to assess HDX in proteins [7-10]; this technique has the advantage of
single-residue resolution of deuterium incorporation, although it usually requires nearly

micromole quantities of protein.

96

Progressive regeneration of various disulﬁde bonds in a reduced cystinyl protein
has also been used to monitor the progress of protein refolding [1 1-13]. We have used
our cyanylation/cleavage methodology to determine the disulﬁde structure of cystinyl
proteins [14, 15] and to trap and identify late-forming (seconds to minutes) refolding
intermediates of human epidermal growth factor (h-EGF) [16], IGF, and one of its
mutants [17], each of which contain three disulﬁde bonds in their native structure. The
folding intermediates that have some free sulﬂiydryls can be captured directly from
aliquots of the refolding solution by reaction with CDAP tetraﬂuoroborate, a reagent that
adds a CN-group to a ﬁee sulfhydryl group, thereby covalently modifying it to a
thiocyanate. The CDAP reaction is performed at pH 3, a condition that minimizes
disulﬁde scrambling and that quenches the folding process as the cyanylation reaction
goes to completion in 10min. The cyanylated folding intermediates are stable and can be
separated by rp-HPLC.

IGF-I has been reported to partly control carbohydrate metabolism in living
systems, as well as to be mitogenic to several cell types [18-20]. IGF-I is a small protein
that consists of 70 residues including 6 cysteines in the form of three intra-molecular
disulﬁde bonds, two of which involve adjacent cysteines (Figure 3.1). LR3IGF-I is a
variant of IGF-I that also is a highly potent growth factor [21, 22]. LR3IGF-I retains all
of the residues of IGF-I except Glu3, which is replaced by Arg and a 13-residue extension

appears at the N-terminus (Figure 3.1).

97

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Here, we use our cyanylation/cleavage methodology to trap the early forming 1-
disulﬁde (l-S) intermediate and a later forming 2-disulﬁde (2-S) intermediate of
LR3IGF-I as we reported previously [17]. The cyanylated l-S intermediates
(Cys31-Cys74) and 2-S intermediates (Cys31—Cys74, Cyslg—Cysm) (disulﬁde structure was
determined by our cyanylation/cleavage methodology) are isolated by rp-HPLC and then
exposed to D20 at pD 6.8 for HDX. The intermediate is then digested with pepsin and
analyzed by MALDI-MS to determine the extent of deuterium incorporation at
approximate locations (limited by the size of the peptic fragments) of various amides
along the polypeptide backbone. In this way, we can indirectly monitor both the progress
of the refolding process as a ﬁmction of disulﬁde bond formation and the conformational
status of these trapped species by the degree of exposure of various amide hydrogens as
assessed by HDX. Progressive “snap shot” results during these studies provide insight to

the dynamic properties of the protein.

II. Experimental Section

The HDX experiments were carried out generally according to procedures
described previously for assessment of deuterium incorporation by LC-MS [2], but in our
work modiﬁed to allow removal of Gdn-HCl and other salts prior to analyses by MALDI-
MS as described below. Our application also involves reduction of disulﬁde bonds, a
step that complicates the sample processing relative to those previously reported
applications of HDX as monitored by MALDI for proteins that do not contain disulﬁde

bonds [23].

99

Materials

All reagents were of the highest purity available from commercial sources; they
were used without further puriﬁcation unless described speciﬁcally. NaHzPO4 was
purchased from Mallinckrodt Baker (Phillipsburg, NJ). Gdn-HCl, tris (hydroxymethyl)
aminomethane-hydrochloride (Tris-HCl), and sodium citrate were obtained from
Invitrogen (Carlsbad, CA) and Spectrum Quality Products (Gardena, CA), respectively.
ACN (HPLC grade) was from EMD Chemicals (Gibbstown, NJ). Immobilized pepsin
was obtained from Pierce (Rockford, IL). C18 ZipTip® was obtained from Millipore
(Bedford, MA). All other chemicals were purchased from Sigma-Aldrich. The TCEP
solutions were freshly prepared before use. All pH or pD values were measured using a
Beckman CD 40 pH meter or ColorpHast pH indicator strips, pH 0 — 14 (EM Science,
Gibbstown, NJ). pD values were calculated by adding 0.4 to the corresponding pH meter
readings [24].

Harvest of folding intermediates

The oxidative folding, trapping, and separation of intermediates and the
determination of their disulﬁde structure are done essentially as described previously
[17]. LR3IGF-I (0.1 mg) was dissolved in 0.5 m1 of citrate buffer, pH 3.0, containing 6
M Gdn-HCl and 0.1 M TCEP. Reduction of the proteins was carried out at 37 °C for 2
hr; the reduced/denatured LR3IGF-I was puriﬁed by rp-HPLC, dried under reduced
pressure, and stored at -70 °C.

The refolding of reduced protein was initiated by diluting the reduced/unfolded
protein sample with 0.10 M Tris-HCl buffer (pH 8.7), containing 1 mM oxidized

Glutathione (GSSG), 10 mM GSH, 0.2 M KCl, and 1 mM EDTA, to a ﬁnal protein

100

concentration of 0.1 mg/ml. The refolding intermediates were trapped in a time-course
manner by removing aliquots (0.1 m1) of protein solution and mixing with 1.0 M HCl
containing freshly prepared 0.2 M CDAP to give a solution of pH 2 — 3. Cyanylation of
free thiol groups by the CDAP proceeded at room temperature for 10 min. The trapped
intermediates were immediately separated and puriﬁed by HPLC. The column was a
Vydac C13 (catalog no. 218TP54; S-um particle size, 300-A pore, 4.6 x 250 mm).
Solvent A was 0.1% aqueous TFA. Solvent B was ACN containing 0.1% TFA. The
linear gradient was 30 — 50% solvent B in 45 min at a ﬂow rate of 1 ml/min. The HPLC
fractions were collected manually, dried under reduced pressure, and stored at —70 °C for
ﬁsrther use.

The identiﬁcation of each intermediate was accomplished previously using the
cyanylation/cleavage methodology [17].
HDX

Each protein sample was equilibrated in aqueous phosphate buffer (20 mM, pH
7.0) for about an hour before the HDX experiment. HDX was initiated by diluting 1 uL
of aqueous protein solution (100 pmol/uL) with 20 [LL of 20 mM phosphate buffer/D20,
pD 6.8 at room temperature. After 10 8, exchange was “quenched” by decreasing pH and
temperature of the reacting system to about pH 2.5 at 4 °C by adding 19 uL of aqueous
TCEP solution (0.5 M, in 0.1 M citrate buffer containing 2 M Gdn-HCl, pH 2.5, 4 0C) to
reduce the residual disulﬁde bonds. The proteins were exposed to D20 for 10 s, a period
long enough to exchange deuteriums for “free” amide hydrogens [25], but short enough

to avoid signiﬁcant exchange with amide hydrogens involved in H-bonding or

101

“protected” by interior folds. Labeling time of less than 10 s requires the use of fast
mixing and quenching instruments.
Pepsin digestion and reduction of residual disulfide bonds

Immobilized pepsin slurry (80 uL) was washed three times with 400 uL of chilled
0.1% TFA (pH 2.5) in a 0.65-mL siliconized tube and stored on ice. “Quenched” protein
samples (40 uL) were added to the pepsin slurry and digested for l min on ice with gentle
- mixing. The immobilized pepsin was removed by a 30-s centrifugation.
Analysis by MALDI-TOF-MS

Mass spectra were acquired on a PerSeptive Biosystems Voyager DE STR
instrument (Applied Biosystems Inc., Framingham, MA). Data were acquired in the
positive reﬂectron mode of operation. Accelerating voltage was 20 kV. Grid voltage and
guide wire voltage were 76% and 0.04% of the accelerating voltage, respectively. The
mirror voltage ratio was 1.12. Extraction delay time was 150 ns. Matrix was 5 mg/mL
a-CHCA in a solution containing ACN, ethanol, and H20 (1:1 :1, v:v:v); the ﬁnal pH was
adjusted to 2.5 with TFA. During the analysis to obtain the data reported herein, the
instrument was internally calibrated using the monoisotopic mass of two well-
characterized peptic fragments (sequences LRRLEM and
LSSLFVNGPRTLCGAELVDALQF; monoisotopic masses are 816.4640 and 2449.2675
Da, respectively). Typically, ion current from 256 laser shots was accumulated for one
spectrum. The average mass of a peptide was calculated by determining the centroid of
its distribution of isotope peak intensities using Data Explorer TM 4.0 installed by the

manufacturer of the MALDI instrument.

102

The sample was desalted using a C13 ZipTip® (Millipore, MA) pipette tip before
being eluted with matrix solution and spotted on a chilled MALDI target. The target was
immediately placed in a desiccator under a moderate vacuum action of a mechanical
vacuum pump such that the spots would dry within 1 — 2 min. Then, the target was
transferred to the MALDI mass spectrometer as soon as possible. The completely
deuterated protein standard was prepared by dissolving LR3IGF-I in DzO at pD 7.0
containing 8 M d4-urea and incubating at room temperature for over 48 hr; this fully
deuterated species was treated by the same sample-handling protocol as all the other
proteins to determine the extent of back-exchange during sample processing after HDX.
The extent of back-exchange during all the experimental procedures described ranged

from 38% to 47%.

III. Results and Discussion
A. Experimental Design for the Application of HDX-MS to Cystinyl/Cysteinyl
Proteins

The key procedure leading to optimal HDX results is to minimize back-exchange
during sample treatment after deuterium incorporation is “quenched”. If all (or most of)
the incorporated deuterimns are substituted by hydrogens from aqueous solution before
mass measurement, then the HDX results will be meaningless. For the most part, HDX-
MS has been widely used in the conformational study of proteins containing no disulﬁde
bonds [1, 23, 26]. Few researchers, if any, have applied this approach to cystinyl

proteins. This is probably because the special features of cystinyl proteins require special

103

sample treatment before analysis by MS, where signiﬁcant back-exchange could easily
occur.

To investigate the extent of deuterium incorporation in different segments along
the polypeptide chain of the protein, deuterated proteins are usually digested before mass
measurement. Proteolytic digestion of cystinyl proteins is usually very slow, if not
impossible [27, 28], unless the protein is denatured and its disulﬁde bonds are reduced.
Therefore, the disulﬁde bonds in a deuterated cystinyl protein should be reduced to
facilitate digestion and peptide identiﬁcation. This leads to two problems that may cause
signiﬁcant back-exchange. The ﬁrst is that only reducing reagents effective under the
conditions required for minimum back-exchange (pH ~3.0 and low temperature) can be
used, and even then, the reaction has to be rapid enough to avoid signiﬁcant back-
exchange. The other problem is that salts introduced to the system have to be removed
after reduction to facilitate analysis by MS. Signiﬁcant back-exchange could occur
during this desalting step both because it is time consuming and large amounts of
aqueous solvents are used. This is less of a problem in HDX-LC-MS experiments
because most LC-MS instruments include an automatic sample-desalting step prior to
injection of the sample into the mass spectrometer. After the sample is loaded, it is
usually ﬂushed with copious inorganic solvents to remove salts and other inorganic
contaminants, which normally takes only 1 or 2 min. When pre-cooled solvent of proper
pH (~ 3.0) is used, the extent of back-exchange can be held to less than 20% [29, 30]. In
the case of HDX-MALDI-MS, however, even for non-cystinyl proteins that usually do
not require desalting, the extent of back-exchange is usually around 50% [23, 31]. The

desalting has to be done manually before loading the MALDI plate to the mass

104

spectrometer, and, thus, special sample treatment protocols have been developed to
minimize possible back-exchange during this step.

To solve the ﬁrst problem, TCEP, which is active at pH ~3.0, was used for the
reduction of disulﬁde bonds. Its concentration was high (0.5 M) to accelerate reduction
(within a few minutes) at low temperature (4 oC). Gdn-HCl was added to the system to
denature the protein, thus facilitating the reduction of disulﬁde bonds buried in the native
protein. To shorten digestion and reduction time, TCEP solution (0.5 M, in 0.1 M citrate
buffer containing 2 M Gdn-HCl, pH 2.5, 4° C) was added to the HDX system at
appropriate time intervals to “quench” deuterium incorporation by decreasing the pH of
the system to ~30. The reduction of disulﬁde bonds starts immediately after adding
TCEP. The mixture is then promptly transferred to the pepsin slurry so that the digestion
and reduction occur simultaneously. A large amount of pepsin (Enzyme : Substrate = 30
: 1) was used to achieve signiﬁcant digestion within a short time. Under our
experimental conditions, complete reduction and digestion are achieved within 1 min,
even shorter than the reported time required for most non-cystinyl proteins (usually
between ﬁve and ten minutes [23, 29]). Efﬁcient reduction/digestion minimizes back-
exchange during this step.

TCEP, Gdn-HCl and Tris-HCI are salts introduced to the system during reduction
and digestion; they must be removed before the sample is mixed with matrix for MALDI-
MS analysis. The Zip-Tip® is a novel desalting tool developed by Millipore (Billerica,
MA), and can be thought of as a HPLC column at the tip of a pipette tip. The Zip-Tip®

pipette tip used in our experiments was a lO-uL pipette tip with a bed of chromatography

media (C13 material) embedded at its end. To use a Zip-Tip® pipette tip for desalting,

105

ﬁrst the pipette tip is wetted and equilibrated using ﬁrst organic and then aqueous
solvent. Peptides/proteins are bound to the C13 bed by aspirating and dispensing the
sample solution for a few cycles. The analytes bound to the tip are then washed by
aspirating water into the tip and then dispensing the water. Finally, desalted
proteins/peptides are eluted from the tip by ﬂushing the C13 bed with an elution solution
that usually contains 50% organic solvent (i.e., ACN). To minimize back-exchange, the
Zip-Tip® and all the desalting solvents used were pre-cooled to 4 °C, and the desalting
solvents were adjusted to pH ~3.0. Aﬁer some practice, the whole desalting procedure
can be performed within one minute. The designed procedures serve to minimize back-
exchange.

The extent of back-exchange during the entire experimental procedure described
ranged from 38% to 47%, typical in HDX-MALDI-MS experiment [23, 31]. The
variation of back-exchange among different peptides is presumably due to side-chain
effects, which can alter the amide hydrogen exchange rate under quenching conditions by
as much as 10-fold [32]. Because the back-exchange rate of different peptic fragments
varies slightly [2, 32], the adjustment of back-exchange for each peptide was based on its
speciﬁc back-exchange value (between 38% and 47%) as determined during the
“control” experiment with the completely deuterated LR3IGF-I.

B. HDX Behavior of Different Segments of when

Ten peptic fragments of LR3IGF-I (corresponding to 78% sequence coverage)
were identiﬁed by molecular weight after analysis of the peptic digest mixture by
MALDI-MS, using the results of an earlier report on the peptic digestion of IGF-I as a

guide [2]. The mass spectrum of the digest is shown in Figure 3.2. The sequences of the

106

ten peptic fragments examined by MS following HDX are represented by the arrows
below the following sequence of LR3IGF-I, which also shows the three disulﬁde bonds of
the native structure (Figure 3.1).

It should be pointed out that some of the sites for peptic cleavage are different in
the oxidized and reduced forms of LR3IGF-I; this is because the selectivity of pepsin is
conformation dependent. Of the ten peptic fragments that were common to most of the
oxidized and reduced forms of LR3IGF-I, seven were identiﬁed by having a calculated
mass that was within i10 ppm of the m/z value for the corresponding observed mass
spectral peak; further, no other possible peptic fragment had a calculated mass within
i500 ppm of the experimentally observed m/z values for these seven peptides. Each of
the other three peptic fragments had alternative candidates having a calculated mass
within :50 ppm of the experimentally observed m/z value. In each of these three cases,
the peptic fragment chosen was the one that agreed with the results of a published study
of the peptic digest of IGF-I [2], and each had a calculated mass within 115 ppm of the
observed values.

As discussed in the following sections, the HDX behavior of each identiﬁed
peptic fragment is consistent with its known conformational features, which, in turn,

conﬁrms the correct identiﬁcation of these peptic fragments.

107

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A summary of the mass shifts during HDX (corrected for back-exchange) for
these ten peptic fragments (listed according to residue number in the sequence of
LR3IGF-I) is presented in Table 3.1 for the completely reduced (R), l-S intermediate (1S:
Cys31—Cys74), 2-S intermediate (28: Cys31—Cys74, Cyslg—Cysm), and native forms (N:
Cys31—Cys74, CYS19—CYS61, Cys6o—Cys65) of LR3IGF-I. After HDX of the completely
reduced protein, the mass shift of most peptic peptides is approximately the same as the
number of exchangeable amide hydrogens, suggesting that these amide hydrogens are
solvent accessible and not engaged in H-bonds. Additionally, the sequential formation of
the three disulﬁde bonds in LR3IGF-I appears to afford progressively increasing global
protection against hydrogen exchange as reﬂected in the corresponding diminution in
mass shift for most of the peptic fragments; these coordinated trends indicate a mode of
folding dependent on the formation of disulﬁde bonds.

At the stage of having formed only one disulﬁde bond (Cy531—Cys74), the mass
shift of each peptic fragment is slightly decreased, except in the fragment consisting of
residues 38 - 55. Thus, the connection of Cys31 and Cysu may initiate partial secondary
structural conformations that involve H-bonds or reduce solvent accessibility as deduced
from only slightly decreased mass shifts. The mass shift data from analysis of peptic
fragments of the 2-S intermediate shows that the number of exchangeable amide
hydrogens declined signiﬁcantly after the formation of disulﬁde bonds Cys31—Cy574 and
Cyslg—Cysm, but the amide hydrogens in fragment 38 — 55 are still freely exchangeable.
Finally, peptic fragments of the native protein (three disulﬁde bonds) after HDX show

mass shifts similar to those in the results for the 2-S intermediate. Certain fragments that

109

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110

have their amide hydrogens further protected may consist of residues involved in the
formation of secondary and tertiary structure as discussed in the following section.
C. Comparison of HDX-MS Results with NMR Data on the Native Structure of
LR3IGF-I

The HDX results reported in Table 3.1 are consistent with NMR data on the
native structure of LR3IGF-I (Figure 3.3) and IGF-1, including those from studies using
HDX, which provide an alternate means of studying the dynamics of cystinyl proteins [7-
10]. LR3IGF-I is an or-helical orthogonal bundle protein composed of three or helixes and
three disulﬁde bonds. The l-S intermediate in our study has a disulﬁde bond within
Cys31-Cys74 that connects the C termini of helix 1 (21 - 31) and helix 3 (67 — 73) as
illustrated in Figure 3.3. The differences observed in our MS study between the mass
shift of peptic fragments from the l-S intermediate and the number of constituent amide
hydrogens occur mostly for fragments that are in the vicinity of either Cys31, Cys74, or the
helixes in the NMR structure. This observation suggests that the Cys31—Cys74 disulﬁde
bond induces the early formation of secondary structures involving H-bonds and/or
stabilizes the local conformational ﬂuctuations that allow transient solvent penetration.
The additional disulﬁde bond Cyslg—Cysm in 2-S seems to close up the hydrophobic core
within the three helixes of the NMR structure (Figure 3.3), further reducing the
accessibility of solvent; this development apparently promotes the development of
secondary and tertiary structure and stabilizes the conformation of the protein. In this
regard, the NMR data are in reasonable agreement with our MS HDX results. The mass-
shift data from MS analysis of peptic fragments 67 — 72, 73 — 83, and 10 - 22/11 — 23

from the native protein suggest that the CYSGO‘CySQSS disulﬁde bond reinforces the

111

coupled conformation of helix 2 and helix 3 and increases their contact with helix 1,

further diminishing access of the solvent to amide hydrogens.

     

  

Helix 2 '5

(56-63)

    

Helix 3
(67-73)

Figure 3. 3 Average NMR structure of LR3IGF-I [8]. N-terminus and C-terminus of the
protein are labeled by N and C, respectively. Three helixes are labeled according to
inclusive residues. Three disulﬁde bonds are represented by dotted lines between the
designated cysteines. The dark-gray region is the structurally ordered part of the protein;
the light-gray segments are highly ﬂexible parts of the protein (in which the heavy atom
r.m.s.d. is larger than the average r.m.s.d. of the ordered part of the protein). The ﬁgure

was drawn using the RasMoI software [33].

112

In the HDX-NMR studies [7, 9, 10], all the helixes in native LR3IGF-I and IGF-1
were found to contain slowly exchanging amide hydrogens, indicating some local
structure; these NMR data are similar to our MS results. However, the HDX-NMR study
of LR3IGF-I [7] also found slowly exchanging amide hydrogens among the ﬁrst ten
residues of the N-terrninal tail and in the segment consisting of residues 43 — 51; this
indication of some structure among residues 43 — 51 contradicts the results from the
study of IGF—I [9, 10], although results from another study by the same group [8] indicate
that this segment is highly ﬂexible as indicated by a much higher than average heavy
atom root mean square deviation (r.m.s.d.) of LR3IGF-I and an increase in T 2 values for
several residues (Figure 3.3), suggesting that solvent was accessible to this region.

Surprisingly, our MS data from analyses of peptic fragments 40 —- 60 and 38 - 55
indicate that both of the above-described scenarios may occur simultaneously. The mass
shiﬁ of the segment 40 — 60 is smaller than that of segment 38 — 55 as listed in Table 3.1;
this disparity seems strange at ﬁrst glance because the latter segment is almost
completely included in the former. However, a reasonable explanation could be that
these two peptic fragments are actually ﬁom two different conformational states of a
given redox state and that pepsin can distinguish between them. This dual-conforrnation
explanation is supported by NMR results [7, 8], where two distinct conformations of
Gly55 were detected, which indicate that the backbone between Lys4o and Cyséo has two
different conformations in native LR3IGF-I, which might be induced by isomerization of
Pro.“ and/or Prosz. Therefore, pepsin may be able to attack cleavage sites in the
Lys4o/Cy560 segment in one conformational state of LR3IGF-I, but not in the other. The

backbone, including the Phe3g/G1Y55 segment, is probably much less structured in one of

113

the two conformational states. In contrast, at least some of the Phe3g/GIYS5 segment is
more structured in the other conformational state, a condition precluding access of
pepsin, and the resulting missed cleavage gives rise to the peptic fragment 40 — 60. This
sort of evidence for the putative dynamic conformational equilibrium observed in the
NMR studies [7, 8] and here in our HDX-MS study of LR3IGF-I has also been reported
in an NMR study of highly mobile regions of an ATPase protein [34].
D. Some Figures of Merit for the Integrated Methodologies

Some ﬁgures of merit for our HDX methodology can be gleaned from a detailed
examination of our results for two of the ten peptic fragments of LR3IGF-I listed in Table
3.2. Analyses of one of these peptic fragments, a 6-mer (LRRLEM), from triplicate HDX
experiments shows considerable slow amide hydrogen exchange in one region of various
redox states of LR3IGF-I, while results from analyses of an 18-mer
(FNKPTGYGSSSRRAPQTG) indicate fast amide hydrogen exchange in a different
region; these results are in agreement with an NMR study of the conformation of native
IGF—I and LR3IGF-I [8-10]. The extent of deuterium incorporation into the two peptic
fragments of interest (the 6-mer and the 18-mer described earlier) is summarized in Table
3.2. For each peptic fragment, the listed constituent residues are identiﬁed according to
their residue position in the sequence of LR3IGF-I. The number of amide hydrogens in
the fragment (in the longer peptic fragment, the number of amide hydrogens is less than
the number of residues because of constituent prolines, which do not have an amide
hydrogen) together with the experimentally determined number of deuterium atoms

incorporated (not adjusted for back-exchange) is also listed.

114

Table 3. 2 Summary of mass shifts for two peptic fragments from different red-ox
species after 10-sec HDX. (Results are the average from three independent experiments;

plus/minus terms indicate the 95% conﬁdence interval).

(A) 6-mer (Sequence = LRRLEM; number of amide hydrogens = 5)

 

 

 

 

 

 

 

 

 

 

Disulﬁde Species R 1S 2S N
(31-74) (31-74, 19-61) (31-74, 19-61,
60-65)
Experimental mass shift (Da) 2.3 :t 0.05 1.7 i 0.1 0.77 i 0.05 0.59 :t 0.05
Adjusted 4.5 3.3 1.5 1.2
mass shift (Da)
Percent deuteration 90% 66% 30% 24%
4.5/5.0 3.3/5.0 1.5/5.0 1.2/5.0
Degree of “protection” 10% 34% 70% 76%
(5.0-4.5)/5.0 (5.0-3.3)/5.0 (5.0-1.5)/5.0 (5.0-1.2)/5.0

 

 

(B) 18-mer (Sequence = FNKPTGYGSSSRRAPQTG; number of amide hydrogens = 15)

 

 

 

 

 

 

 

 

 

 

Disulﬁde Species R 15 2S N
(31-74) (31-74, 19-61) (31-74, 19-61,
60-65)
Experimental mass shift 8.5 :t 0.2 8.8 i 0.2 8.5 i 0.7 8.5 :t 0.1
(Da)
Adjusted 14.8 15.2 15 14.8
mass shift (Da)
Percent deuteration 98% 100.7% 99.3% 98.7%
14.8/15.1 15.2/15.1 15/15.l 14.8/15.1
Degree of “protection” 2% -0.7% 0.7% 2%
(15.1-14.8)/15.1 (15.1-15.2)/15.1 (15.1-15)/15.1 (15.1-14.8)/15.1

 

115

 

In particular, examine the HDX results for the peptic fragment 67 — 72. In
consideration of back-exchange, the listed 2.3-deuterium atom incorporation (or 4.5 Da
after adjustment for back-exchange) in Table 3.2 (A) means that essentially all ﬁve amide
hydrogens were exchanged for deuterium atoms during the HDX experiment; this result
indicates that little protection against exposure to D20 is afforded these ﬁve amide
hydrogens in the reduced (R) form of LR3IGF-I. In Table 3.2 (A), the listed 1.7-
deuterium atom incorporation means that 3.3, on average, of the ﬁve amide hydrogens
were exchanged for deuterium atoms during the HDX experiment with l-S. This reduced
level of deuteration of 66% (3.3/5.0) indicates that some degree of secondary structure
appears in the l-S intermediate, which partially protects the ﬁve amide hydrogens from
complete exposure to the D20 during HDX. We numerically indicate this degree of
protection as the difference between the maximum possible shift and the observed mass
shift relative to the maximum possible shift (in this case, (5 - 3.3)/5 x 100% = 34%).
The results for the 2-S intermediate show a substantially smaller mass shift of 0.77 Da,
which gives an adjusted mass shift of 1.5 Da; this corresponds to 30% deuteration or 70%
protection, meaning that there is signiﬁcantly more secondary structure in this segment
after the second disulﬁde bond has formed. Finally, the HDX results for the native
structure show the smallest mass shift of 0.59 Da, which gives an adjusted mass shift of
1.2 Da; this corresponds to 24% deuteration or 76% protection, which means that the
native structure contains the greatest amount of secondary structure in this segment, as
expected. The results for HDX of residues 67 — 72 in the native structure of LR3IGF-I
are in good agreement with HDX assessment of the conformation of these structures

using NMR [7, 9, 10].

116

Abbreviated segments of the MALDI mass spectra of the peptic fragment 67 - 72
after HDX of the reduced, l-S intermediate, 2-S intermediate, and native forms of
LR3IGF-I are shown in Figure 3.4 (A). The mass spectrum of the normal (no HDX)
peptic fragment (6-mer) of LR3IGF-I shown in the top panel of Figure 3.4 (A) serves as a
reference point from which to compare the mass shift associated with this peptic
fragment after HDX of the polypeptide in the various redox states indicated in the lower
four panels of Figure 3.4 (A). The mass shift is assessed by determining the centroid of
the multiplet of isotope peak intensities representing the peptic fragment (monoisotopic
mass with no HDX = 816.4640 Da). The centroid of the multiplet of peaks in the third
panel of Figure 3.4 (A) indicates a mass shift of 2.3 Da from the reference point in the top
panel of Figure 3.4 (A); this result, when adjusted for back-exchange, gives the shift of
4.5 Da for the reduced (R) protein reported in the second column, second row of Table
3.2 (A). The centroid of the multiplet of peaks in the fourth panel of Figure 3.4 (A)
indicates a mass shift of 1.7 Da from the reference point in the top panel of Figure 3.4
(A); this result, when adjusted for back-exchange, gives the shift of 3.3 Da for the l-S
intermediate reported in Table 3.2 (A). Similarly, the centroid in the ﬁfth panel of Figure
3.4 (A) indicates a mass shift of 0.77 Da, which corresponds to the adjusted value of 1.5
Da in Table 3.2 (A) for the 2-8 intermediate. Finally, the centroid in the sixth panel of
Figure 3.4 (A) indicates a mass shift of 0.59 D3, which corresponds to the adjusted value
of 1.2 Da in Table 3.2 (A) for the native form of LR3IGF-I. The mass spectrum in the
second panel of Figure 3.4 (A) (corresponding to the ﬁrlly deuterated species) served as
the basis for determining the degree of back-exchange under the sample-handling

conditions described above.

117

Figure 3. 4 Mass spectra of (A) the segment 67 - 72 (LRRLEM, 6-mer) from different
redox species and (B) the segment 38 — 55 (FNKPTGYGSSSRRAPQTG, l8-mer) from
different redox species. The ﬁrst row shows the mass spectra of the peptides before
deuteration; the second row shows the mass spectra after sustained HDX to achieve a
high degree of deuteration of the peptides. The remaining four rows are mass spectra of
indicated redox species (R: reduced; 1-S: one-disulﬁde intermediate; 2-S: two-disulﬁde

intermediate, N: native) after a 10-s exposure to D20 at pD 6.8.

118

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For comparison, examine the HDX results in Table 3.2 (B) for the peptic
fragment 38 — 55. In consideration of back-exchange, the 8.5-deuterium atom
incorporation listed in Table 3.2 (B) means that essentially all ﬁfteen available amide
hydrogens were exchanged for deuterium atoms during the HDX experiment. This result
indicates that no protection against exposure to D20 is afforded these ﬁfteen amide
hydrogens in the reduced (R) form of LR3IGFI. Similarly, the HDX results for the ﬁfteen
hydrogens in this eighteen-residue peptic fragment from all other red-ox states indicate
that no protection is ever apparent, and thus little protection from solvent is available to
this segment of the sequence in any of the redox states of LR3IGF-I. The abbreviated
mass spectra of the peptic fragment 38 — 55 (lS-mer) from which the data in Table 3.2
(B) were gathered for different redox states of LR3IGF-I are shown in Figure 3.4 (B).

As is apparent from a cursory examination of the mass spectra, a substantial and
nearly constant mass shift was detected in all cases, indicating that none of the amide
hydrogens is afforded protection from exposure to DZO. These results indicate that there
is no signiﬁcant conformational structure among residues 38 — 55 in any of the indicated
redox states of LR3IGF-I. Further examination of the mass spectra in Figure 3.4 (B)
might suggest a lesser mass shift in the third panel than in the lower three; however, the
mass shift in all ﬁve panels is essentially the same as indicated by the centroid calculation
on the cluster of peaks in each panel. The major factor in the lower three panels is
apparently the substantial leading edge to the most intense peaks; because the magnitude
of the ion current in the lower three panels is two to three times that in the third panel, the
leading edge in the spectra is real with regard to ion counting statistics, and it causes the

centroid to be approximately the same as that in the third panel.

120

The thiocyanocyanate group on the modiﬁed cysteines is not expected to
inﬂuence the conformation of the intermediates in any signiﬁcant way, as reasoned by
analogy to NMR work by Talluri et a1. [35], who used the (2-aminoethyl)
methanethiosulfonate derivative of cysteine, a blocking group much larger than the

thiocyanate group.

IV. Conclusion

These results demonstrate that cyanylation-based disulﬁde mapping methodology
can be combined successfully with HDX methodology for a more comprehensive
assessment of the conformation of cystinyl proteins as a function of cysteine status. The
cyanylation-based methodology is effective in determining and controlling the cysteine
status in cystinyl proteins during assessment of its conformation by HDX. The results
reported here show that these integrated methodologies can monitor a clear and distinct
dependence of the conformation of a cystinyl protein on the progressive formation of its
constituent disulﬁde bonds during oxidative refolding.

Our results of HDX on LR3IGF-I and its 2-disulﬁde folding intermediate as
monitored by MALDI-MS provide evidence for two dynamic conformations of a
segment of the polypeptide backbone; this ﬁnding is consistent with results from several
NMR studies of LR3IGF-I and IGF-1 [7-10]. In this way, the integrated methodologies
will help elucidate the detailed molecular mechanisms of protein dynamics and protein

folding.

121

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[21] R. King, J. R. Wells, P. Krieg, M. Snoswell, J. Brazier, C. J. Bagley, J. C. Wallace,
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recombinant insulin-like grth factor-1(IGF-I) and potent analogues of IGF-I,
with Gly or Arg substituted for Glu3, following their expression in Escherichia
coli as fusion proteins, J Mol Endocrinol 8 (1992) 29-41.

[22] G. L. Francis, M. Ross, F. J. Ballard, 8. J. Milner, C. Senn, K. A. McNeil, J. C.

Wallace, R. King, J. R. Wells, Novel recombinant fusion protein analogues of
insulin-like grth factor (IGF)-I indicate the relative importance of IGF-binding

123

protein and receptor binding for enhanced biological potency, J Mol Endocrinol 8
(1992) 213-23.

[23] J. G. Mandel], A. M. Falick, E. A. Komives, Measurement of amide hydrogen
exchange by MALDI-TOF mass spectrometry, Anal Chem 70 (1998) 3987-95.

[24] P. K. Glasoe, F. Long, Use of glass electrodes to measure acidities in deuterium
oxide, Notes 64 (1960) 188-90.

[25] Y. Bai, J. S. Milne, L. Mayne, S. W. Englander, Primary structure effects on peptide
group hydrogen exchange, Proteins 17 (1993) 75-86.

[26] Z. Zhang, D. L. Smith, Thennal-induced unfolding domains in aldolase identiﬁed by
amide hydrogen exchange and mass spectrometry, Protein Sci 5 (1996) 1282-9.

[27] D. L. Smith, Z. R. Zhou, Strategies for locating disulﬁde bonds in proteins, Methods
Enzymol 193 (1990) 374-89.

[28] J. Qi, J. Wu, G. A. Somkuti, J. T. Watson, Determination of the disulﬁde structure of
sillucin, a highly knotted, cysteine-rich peptide, by cyanylation/cleavage mass
mapping, Biochemistry 40 (2001) 4531-8.

[29] J. R. Engen, B. E. Morton, X. Chen, Using Stable-Isotope-Labeled Proteins for
Hydrogen Exchange Studies in Complex Mixtures, Analytical Chemistry 74
(2002) 1680-6.

[30] Y. Deng, H. Pan, D. L. Smith, That Hydrogen Exchange at Individual Peptide
Amide Linkages Can Be Determined by Collision-Induced Dissociation Mass
Spectrometry, Journal of the American Chemical Society 121 (1999) 1966-7.

[31] G. S. Anand, C. A. Hughes, J. M. Jones, S. S. Taylor, E. A. Komives, Amide H/2H
exchange reveals communication between the CAMP and catalytic subunit-
binding sites in the R(Dalpha subunit of protein kinase A, J Mol Biol 323 (2002)
377-86.

[32] D. L. Smith, Y. Deng, Z. Zhang, Probing the non-covalent structure of proteins by
amide hydrogen exchange and mass spectrometry, J Mass Spectrom 32 (1997)
135-46.

[33] R. A. Sayle, E. J. Milner-White, RASMOL: biomolecular graphics for all, Trends
Biochem Sci 20 (1995) 374.

[34] Y. T. Chou, J. F. Swain, L. M. Gierasch, Functionally signiﬁcant mobile regions of

Escherichia coli SecA ATPase identiﬁed by NMR, J Biol Chem 277 (2002)
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124

[35] S. Talluri, D. M. Rothwarf, H. A. Scheraga, Structural characterization of a three-
disulﬁde intermediate of ribonuclease A involved in both the folding and
unfolding pathways, Biochemistry 33 (1994) 10437-49.

125

CHAPTER 4
CHARACTERIZATION OF PEPTIDE FOLDING NUCLEI
BY HYDROGEN/DEUTERIUM EXCHANGE-MASS SPECTROMETRY AND

CIRCULAR DICHROISM

I. Introduction

Protein folding is a central problem of molecular biophysics. An early model of
folding hypothesized that proteins adopt “ﬂickering” native structure in high-propensity
segments, and that protein folding proceeds by the gradual association and mutual
stabilization of such marginally stable segments [1, 2]. Consistent with this hypothesis,
mixing synthetic peptide fragments of a protein in sufﬁciently high concentrations can
produce folded species analogous to those in the corresponding segments of the intact
protein [3]. On the other hand, numerous experiments show that most peptides
corresponding to secondary-structure elements (e.g., tit-helices, B-hairpins, B-meanders,
etc.) from naturally occurring proteins exhibit little secondary structure in aqueous
solution [4], probably owing to competition between water-amide and amide-amide
hydrogen bonds (H-bonds) [5, 6]. Therefore, long-range interactions are usually invoked
to account for the stabilization of short-range structure during protein folding. However,
the mechanisms by which interactions between peptides stabilize the structure within
peptides have not been studied systematically. It would be useful, for example, to assess
the correlations between secondary, side-chain, and tertiary structure in pairs of
covalently linked peptides and to characterize quantitatively the stabilization of local

structure conferred by various factors, e.g., amino-acid sequence, speciﬁc side-chain

126

packing, steric, and/or hydrophobic exclusion of competing water molecules, etc.

For clarity, we distinguish three types of structure. The degree of secondary
structure is deﬁned by the average number of speciﬁc H-bonds between amide groups of
the protein backbone, which imposes local (i.e., short-range in the polymer sense)
conformational order on the backbone. The average is taken over the equilibrium
conformational ensemble under the experimental conditions (e.g., the thermodynamic
average). A polypeptide system has stable secondary structure if, in the thermodynamic
average, the majority of its amide groups are involved in H-bonds with speciﬁc amide
groups. The degree of tertiary structure is deﬁned by global conformational order, i.e.,
the ordering of the protein backbone in space as measured by the mean CQRMSD within
the conformational ensemble. Thus, a polypeptide system has stable tertiary structure if
the majority of conformations in its equilibrium ensemble are within 6 A CaRMSD of a
single, physically plausible conformation. Finally, the degree of side-chain structure is
deﬁned by the ordering in the dihedral angles of the protein side chains. Such structure is
often associated with tertiary structure (side chains are constrained because of dense
packing), but may also arise from local secondary structure. For example, a speciﬁc side-
chain rotarner may predominate because of steric interactions or H-bonding to the
backbone (e. g., a-helix capping) or other side-chains interactions (e. g., salt bridges or or-
helices). Thus, a polypeptide system has stable side-chain structure if, on the average, its
side-chain x dihedral angles are within 30° of a speciﬁed rotarner.

HDX, as monitored by MS and NMR spectroscopy, has been used widely to study
conformations and conformational dynamics of proteins in various conditions ranging

from isolated states to non-covalent interactions [7, 8]. The cru'rent understanding of

127

protein folding suggests three connected hypotheses that can be tested by HDX on pairs
of covalently linked peptides. (H1) The association of any two peptides should lower the
rate of HDX (even in the absence of signiﬁcantly increased secondary structure) because
steric hindrance alone will inhibit water from encountering the amide groups. (HZ) The
association of any two peptides should increase the secondary structure of both peptides
because amide-amide H-bonds will have less competition from amide-water H-bonds.
(H3) The association of any two sufﬁciently nonpolar peptides may provoke stable
secondary, side-chain, and/or tertiary structure.

In this chapter, the three hypotheses (Hl—H3) are tested by comparing the
structure in monomeric and dimeric peptides with HDX-MS and CD. HDX has been
rarely applied to study oligopeptide systems [9], nor has HDX-MS been widely applied to
the analysis of polypeptides containing disulﬁde bonds [10]. We chose to measure HDX
using MS because it requires signiﬁcantly less material than NMR and tolerates poorly
soluble peptides. The major advantage of HDX-NMR, as compared to HDX-MS, is its
resolution at the single amino acid residue level; while the resolution of HDX-MS
analysis is conventionally limited by the size of the peptic fragments of deuterated
polypeptides (usually 5 — 12 resiudes). However, in recent years it was demonstrated that
collision-induced dissociation tandem mass spectrometry (CID-MS/MS) can assess the
deuterium incorporation of individual amides from the b-series of CID fragment ions [11-
13]. In 2004, HDX-CID-MS/MS results were reported in a study of two oligopeptides
and their homodimers formed by non-covalent interactions using ESI with an ion-trap
mass spectrometer [14].

The main purpose of this chapter is: (1) To test the three hypotheses of protein

128

folding; (2) To investigate the feasibility of characterizing protein folding nuclei by the

combined approaches of HDX-MS and CD.

II. Materials and Methods
Materials

The model peptides (PO. and PB, see deﬁnition in Section HI. A.) were synthesized
by the Genomics Technology Support Facility (GTSF) at Michigan State University and
puriﬁed by rp-HPLC using a Vydac C13 analytical column (catalog #218TP54). All other
reagents were of the highest purity commercially available and used without further
puriﬁcation. Dibasic sodium phosphate (NaZI-IP04) and sodium sulfate (Na2804) were
purchased ﬁ'om Spectrum Chemical Mfg. Corp. (Gardena, CA) and Columbus Chemical
Industries, Inc. (Columbus, WI), respectively. Gdn-HCl was purchased from Invitrogen
(Carlsbad, CA). ACN (UV grade) and ACN (HPLC grade) were purchased from
Honeywell International, Inc. (Muskegon, MI) and EMD Chemicals (Gibbstown, NJ),
respectively. The TCEP solutions were freshly prepared before use. All other chemicals
were purchased from Sigma.

All pH/pD values were measured using a Beckman (D 40 pH meter or ColorpHast
pH indicator strips (pH 0 — 14; EM Science, Gibbstown, NJ). pD values were computed
by adding 0.4 to the corresponding pH readings [15]. The standard buffer in this study
contains 200 mM Na2804, 10 mM NaHzPOa, adjusted to pH 6.0. For the monomeric Pa
and PB peptides, the buffer also contained 1 mM reduced dithiothreitol to prevent
dimerization of the monomers. No noticeable oxidation was detected after the

experiment, as checked by rp-HPLC.

129

Preparation of the Peptide Dimers

The P.m and PB]; homodimers and the Pap heterodimer were prepared as described
previously [16]. Brieﬂy, roughly equal amounts of the Pa and PB monomers were mixed
in a solution containing 5 M Gdn-HCl and 200 mM Tris-HCl at pH 8.0 and were air-
oxidized for 48 hr at 25°C. The dimers were separated and puriﬁed on a Vydac C13
analytical column using a water/ACN gradient in the presence of 0.1% TFA. Each dimer
was collected manually and dried under reduced pressure for further use.

CD
CD spectra were obtained at 0 °C on a Jasco J-810 CD spectropolarimeter (Jasco,

Inc., Easton, MD) using a therrnostatted l-mm pathlength cell. The samples were
dissolved in the standard buffer. The concentrations were determined by UV absorbance
at 205 nm [17] and were typically 0.15 mg/ml.
HDX

The HDX experiments were carried out in a cold room, typically at 4 °C. Each
peptide sample was equilibrated with the standard buffer (1 nmol/uL) for about half an
hour before each experiment. HDX was initiated by diluting 1 uL of aqueous protein
solution with 19 uL of standard buffer made with D20 at pD 6.4. After 15 s, 18 11L of
aqueous TCEP solution (500 mM in 2 M GdnHCl, pH 2.5, 4 °C) were added to quench
the exchange by decreasing the pH to 2.5 and to reduce the disulﬁde bond. The pH was
maintained at 2.5 in all subsequent steps. Incubation of the sample with D20 for 15 5
should be long enough to exchange exposed amide hydrogens with deuteriums [18], but

short enough to avoid signiﬁcant exchange with H-bonded/protected amide hydrogens.

130

Analysis by MALDI-TOF Mass Spectrometry

Mass spectra were acquired on a PerSpective Biosystems Voyager DE STR
instrument (Applied Biosystems Inc., Framingham, MA) in the positive ion reﬂectron
mode of operation. The accelerating voltage was 20 kV; the grid voltage and guide wire
voltage were 76% and 0.04% of the accelerating voltage, respectively. The mirror
voltage ratio was 1.12, whereas the extraction delay time was 150 ns. The matrix was on-
CHCA in a 5 mg/mL solution containing ACN, ethanol and H20 (1:1:1, v:v:v); the ﬁnal
pH was adjusted to 2.5 with TFA. The measurement was done using next-spot external
calibration with the Pa and Pp monomers as the calibrants, for which the calculated
monoisotopic [M + H]+ masses are 1671.7219 Da and 1681.8160 Da, respectively. The
average mass of each peptide was measured experimentally by determining the centroid
of the distribution of its isotope peak intensities using Data Explorer TM 4.0. Ion current
from 256 laser shots was accumulated for each spectrum.

Immediately after quenching the HDX process, the sample was desalted with cold
water containing 0.1% TFA (pH 2.5) using a C13 ZipTip® (Millipore, Bedford, MA)
pipette tip, eluted with matrix solution, and spotted on a chilled MALDI target. The
target was immediately placed in a desiccator under moderate vacuum to dry the spot
within 1 — 2 min. The target was immediately transferred to the MALDI instrument. A
highly deuterated standard was prepared by dissolving the sample in D2O at pH 7.0 with
8 M d4-urea and incubating at room temperature for over 48 hr. Treating the highly
deuterated standard by the same protocol indicated that the extent of back-exchange

during the experimental procedures was 35%.

131

Analysis by CID Tandem Mass Spectrometry

Mass spectra were acquired on a Finnigan LCQDecal

ion-trap mass spectrometer
(ThermoQuest, San Jose, CA) with an ion-spray voltage of 3 kV, a capillary voltage of 24
V, a tube lens offset of 55 V, and a capillary temperature of 150 °C. Data-dependent
MS/MS conditions were set with a default collision energy of 35%, a default charge state
of 2, and an isolation width of 5 (m/z). Scans were taken over the range m/z 235 to 1500.
Typically, 100 scans were accumulated per spectrum.

Immediately after quenching the HDX process, the sample was desalted with cold
water containing 0.1% formic acid (FA) (pH 2.5) using a C13 ZipTip® pipette tip and
eluted with 100 11L of a water/ACN solution (1:1, v:v) pre-adjusted to pH 2.5 with FA.
The processed sample was infused into the LCQDeca immediately with a pre-chilled (ice)
lOO-uL syringe at a ﬂow rate of 20 uL/min; the syringe was kept buried in ice throughout
the experiment. The extent of back-exchange during the infusion-CID-MS/MS
procedures was 25%, as determined from analysis of the highly deuterated standard
prepared as described above. Mass spectra were obtained with the operating software

Xcalibur, and the centroid values were calculated with the Magtran software [19].

III. Results and Discussion
A. Criteria for Selection of Peptide Models

To validate our approach for studying protein folding nuclei, the selected model
peptides should adopt distinct conformations in different states (e.g., signiﬁcantly folded
in one state and essentially unstructured in another state), and these conformations should
have been well characterized using conventional techniques (i.e., NMR); to test the three

hypotheses of protein folding, the peptide models should be from a protein that is

132

important and interesting to scientists in the area of protein structure and folding. In
addition, to enforce the association of designated peptides, we restrict our study here to
( 1) individual monocysteinyl peptides that are linked by a disulﬁde bond in the native
structure of a folded protein and (2) all possible covalent dimers of these peptides.

BPTI is a small single-domain protein that consists of 58 residues including 6 cysteines in
the form of three intra-molecular disulﬁde bonds (Figure 4.1). The native structure of
BPTI consists of an amino-terminal 310-helix, a highly twisted anti-parallel B-sheet, and
an a-helix near the C-terminus (Figure 4.2). The native structure is stabilized by the
three disulﬁde bonds. This protein is one of the most extensively studied systems in
protein sciences. It has served as a model system for the examination of almost all
aspects of protein structure including X-ray crystallography and neutron diffraction [20],
the development of two-dimensional solution nuclear magnetic resonance (NMR)
techniques for protein structure determination [21], measurement of protein dynamics
[22, 23], studies of the position and dynamics of water molecules [24]. BPTI is also a
popular model in protein folding. It has been used for development of computational
approaches to protein folding [25-27], and its oxidative folding pathway has been
thoroughly studied in terms of the disulﬁde-bonded intermediates that accumulate during

folding [28-34].

133

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134

 

Figure 4. 2 Ribbon diagram of BPTI [35], with peptides Pa and PB depicted in dark and
light gray, respectively. The three native disulﬁde bonds are also shown: Cyss-Cysss
(left), Cysm-Cysn (right), and Cysm-Cyssl (middle). The ﬁgure was drawn using the

MOLMOL software [36].

135

We illustrate the application of our method using two cysteinyl peptides and their
three possible disulﬁde-linked pairs, denoted P0,, P3, Pm, Pm}, and Pap. The model
peptides mimic selected segments of the sequence of BPTI; speciﬁcally, the monomers,
Pa and PB, correspond to residues 43-58 (NNFKSAEDCMRTAGGA) and 20-33
(RYFYNAKAGLCQTF), respectively, which form the disulﬁde-linked C-terrninal 0t-
helix and central B-hairpin, respectively, in native BPTI (Figure 4.2). Because Pa and
PB each have a cysteine, they can be disulﬁde-bonded to form the P.m and PM,
homodimers and the P04; heterodimer. The Pap heterodimer corresponds to the
(Cys3o—Cys51) disulﬁde species of intact BPTI, which accumulates in the one-disulﬁde
ensemble and seems to be partially folded [33, 37]. Oas and Kim have shown that the
Pug heterodimer has stable secondary structure by CD and lD-NMR, whereas the Pa and
Pp monomers do not [16]. The Pa... and P35 dimers have not been studied structurally.
However, it was reported recently that the homodimer of another peptide model for the
BPT I B-sheet folds into a four-stranded antiparallel B-sheet when the two monomers are
connected by an “optimized” synthetic linker, although homodimers formed using
“unoptimized” linkers and the monomer peptide do not form stable structure [9].
Throughout this chapter, the residues are numbered according to their position in native

BPTI (e.g., Cys30, Ser47, etc).

136

B. Conformational Study of Selected Peptide Models by CD Spectroscopy
Circular dichroism spectroscopy is a convenient tool to study the secondary
structure of a polypeptide. In this study, we ﬁrst used CD spectroscopy to analyze each
peptide model. The CD spectra of Pap (Figure 4.3 (A)), PC. and P.m (Figure 4.3 (B)), and
PB and P88 (Figure 4.3 (C)) suggest that the presence of the disulﬁde bond increases the
secondary structure content of all three dimers (PM, Ppp, and Pap) relative to that in their
component monomers (Pa and PB)- Although the absolute magnitude of the CD signal
shown in Figure 4.3 is smaller than expected ideally, the spectral shape is clearly
consistent with or-helical and B-sheet structure [38]. The clearest evidence of secondary
structure is visible in Pug, which has been shown previously to contain stable tertiary
structure [16]. At 0 °C, the CD spectrum of Pap exhibits two minima near those expected
for helical proteins (208 nm and 222 nm) [38]; at 60 oC, these troughs are replaced by one
trough at 204 nm characteristic of random-coil structure [3 8]. The summed CD spectrum
of the Pa and PB peptides resembles the spectrum of Pap at 60 °C, but shifted to lower
wavelengths; this contrasts with results from the earlier study [16], which found little
deviation. A marked nonlinearity in the temperature dependence is observed for
Pug (Figure 4.3 (D)), which is consistent with our interpretation that Pap is at least
partially structured; however, the absence of a fully sigrnoidal transition suggests that the
folded Pug structure could be further stabilized, even at 0°C. Comparison of Pa and Pa...
shows that Pa... at 0 °C has more (it-helical structure than P.m at 60 °C, which has more or-
helical structure than P... at 0 °C (judging by ellipticity at 222 nm). Similarly, Pm; at 0 °C
has signiﬁcantly more secondary structure than its corresponding monomer PB at 0 °C

(judging by ellipticity at 217 nm) [3 8]. These results are consistent with hypothesis (HZ)

137

that local secondary structure content is stabilized upon the association of two peptides,
presumably by the exclusion of water from attacking the amide-arnide H-bonds.
However, the increase in secondary structure in PM seems small and local, not global; the

PM CD spectrum is not fully ct-helical even at 0 °C, and the difference spectrum of

PM and Pa

138

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142

does not exhibit the double minima at 208 nm and 222 nm expected for a-helical
structure (Figure 4.3 (8)). Hence, the enhanced protection of P,m against HDX (which
rivals that of the native-like Pap) likely stems from the association of the two PCl
monomers connected by the disulﬁde bond, rather than from enhanced secondary
structure within each, which supports hypothesis (H1). By contrast, the CD spectrum of
the PM; peptide at O 0C (but not at 60 °C) suggests the presence of side-chain structure;
speciﬁcally, the positive peak at 230 nm likely corresponds to the CD signal of one or
more ordered-tyrosine sidechains [39]. This suggests that Ppp may also have stable
tertiary structure, similar to that in Pug; further evidence is cited below.

C. Study of the Conformational Features of Selected Peptide Models by HDX-

MALDI-MS

Results from HDX and matrix-assisted laser-desorption/ionization time-of-ﬂight
mass spectrometry (MALDI-TOF-MS) of the ﬁve peptides are reported as total
deuterium incorporation determined by the observed mass shiﬁ (Figure 4.4 and Table
4.1). The monomeric PCl and PB peptides showed average mass shifts of 10.2 Da and 8.6
Da, respectively, from their 15 and 13 exchangeable hydrogens. Hence, Pa and PB show
some protection against HDX under our experimental conditions. This protection may
reﬂect “ﬂickering” local structure, as has been observed in other experiments [9, 40] and
in our CD and CID-MS/MS experiments reported below. Much smaller mass shifts of
6.8 Da and 3.5 Da were measured for the subunits of the P,m and PM; dimers, respectively;
the native-like Pap heterodimer showed slightly smaller mass shifts of 5.5 Da (on-subunit)
and 2.9 Da (B-subunit). Hence, all three dimers showed signiﬁcantly more protection

than their component monomers, consistent with hypothesis (H1).

143

Table 4. 1 HDX results for a- and B-subunits as assessed by MALDI-TOF-MS.

 

Peptide model

Pa

PB

P0,,

P00.

Pm:

 

Mass shift'

 

 

10.2

 

8.6

 

5.5 (or-chain)
2.9 (IS-chain)

 

6.8
(each oc-chain)

 

3.5
(each B-chain)

 

'Mass shift (adjusted for back-exchange) is the average from triplicate runs. The uncertainty of the

reported values is $0.1Da expressed as standard deviation.

144

 

Figure 4. 4 Mass spectra of (A) PM before HDX, (B) PM aﬁer HDX, (C) Pa after HDX,
(D) PB after HDX, (E) PM after HDX, (F) Pm, after HDX. The disulﬁde bond in each
dimer was present during HDX, but was reduced before the mass measurement. The
cluster of peaks corresponding to the Pa-chain is on the left, while that for the Pp-chain is
on the right in (A) and (B). Likewise, PM and P59 were exposed to HDX as dimers, but

reduced to the monomer prior to analysis by MALDI-MS.

145

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146

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148

In this study, CD spectroscopy was used for a preliminary investigation of the
conformation of each peptide model. Although CD results have been qualitatively
related to HDX results in several studies [41, 42] (the more structure detected by CD, the
more protection against HDX observed), more research will be necessary to determine
whether estimation of secondary structure content based on CD can be quantitatively
correlated to protection against HDX. For example, amide hydrogens along a
polypeptide chain that adopts 100% speciﬁc secondary structure (i.e., oc-helix) may be
replaced by deuteriums during HDX [10] depending on various factors and conditions
including breathing of the polypeptide chain, the HDX conditions (e.g., pD and labeling
time), and the degree of involvement of the amide hydrogens in intra-molecular H-bonds.
Protection against HDX, on the other hand, may result from factors other than those
associated with well-deﬁned secondary structures that can be detected by CD
spectroscopy (e.g., lack of exposure to the deuterated solvent due to tertiary structure).
Therefore, it seems that one should hesitate before quantitatively correlating HDX results
with secondary structural content of a polypeptide.

In addition, a quantitative study of the CD results does not help signiﬁcantly to
validate our strategy of studying small peptides using HDX-MS. The resolution of CD
analysis is low, and the accuracy of quantitating secondary structure by CD has been
disappointing [40, 43, 44]. To further characterize the 3-D structure of each peptide
model (including determining the secondary structural content), NMR or X-ray
crystallography should be used. On the other hand, as demonstrated in this study, when
the deuterium content of each amide linkage can be deduced from HDX-CID-MS/MS, a

qualitative correlation of the extent of deuterium incorporation and intra-molecular H-

149

bonding (or other structural features of the polypeptide chain) is reasonable to a ﬁrst
approximation and can provide a means to monitor changes in local structure of peptides
as a ﬁmction of cysteinyl/cystinyl status, for example.

D. Assess Deuterium Content at the Single Amino Acid Residue Level by HDX-
CID-MS/MS

The anti—parallel B-sheet segment is the core element of BPTI [9]. To identify the
position(s) of the enhanced HDX protection induced by the dimerization in the B-unit, we

used CID-MS/MS to probe the HDX of individual amide groups in P13: P33, and PM.

Deca

These measurements were done using direct infusion ESI on an LCQ ion trap mass

spectrometer. HDX-CID of b-ions has been reported to be reliable for assessing
deuterium content at the single amino-acid level, whereas that of y-ions is unreliable [11-
13]. A series of b-ions (b 3 - b1o) was formed during the CID process for the PB subunit
(Figure 4.5); however, only y-ions were produced for the Pa subunit (Figure 4.6). The
masses of the b 3 — b ,0 ions were used to assess deuterium incorporation in residues Tyr23
— LCUzg of the PB subunit in PB, PBS, and PM (Table 4.2). The results indicate that the
amide hydrogens of Ty1'23, Asn24, Ly326, and Alan exchange readily in PB (0.6 - 1.1 Da),
but are protected in Pap (0.1 — 0.3 Da) and even more so in PM (0.0 Da for Lys26, 01 Da
for Alan). By contrast, the Ala25 amide hydrogen exchanges moderately and equally well
(0.4 Da) in the monomer (Pp) and both dimers (PW, and PM). Finally, the GIY28 and
LCUzg amide hydrogens exchange weakly in PB (0.2 Da) and less so in PM; and Pap (0.1
and 0.0 Da, respectively). Due to the missing fragment in the series of b1, b2, bu-bn

ions, deuterium incorporations for Tyr21, Phezz, Cys3o — Phe33 are not available.

150

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153

However, the 2.0-Da mass shift of the b3 ion of PB indicates that Tyrzi and Phezz amide
hydrogens exchange readily in the monomer. In addition, the l.O—Da decrease in the
mass shift of the b3 ion in P53 and P043 suggests that one of these two amide hydrogens is
protected in the dimers. Similarly, segment Cys3o — Phe33 of the [3 sequence shows less
protection in PB (2.3 Da) than in Pm; and Pap (0.9 Da and 1.0 Da, respectively).

These site-speciﬁc HDX results for the PB subunit are generally consistent with
the solvent exposure and H-bonding of the native structure (See Figure 4.7). If the [3-
chain adopts its native structure, the amide hydrogens of Phezz, Asn24, Glyzg, Gln31, and
Phe33 should be well protected because they are all involved in strong backbone (amide-
amide) H-bonds; however, the Phe33-Arg20 H—bond is unlikely to be maintained in the
peptide model, owing to the well-known ﬂexibility of chain termini. Consistent with
these predictions, signiﬁcant protection for Asn24 and Glyzg is observed in PM; and Pap
(Table 4.2). Similarly, the mass shifts of the b3 and b” ions of PBB and P04; are consistent
with signiﬁcant protection of Phezz and Gln31 against HDX, which are mutually H-
bonded in the native structure. The amide hydrogen of Leuzg is well protected in PM; and
Pap (Table 4.2); in the native structure, the amide hydrogen of Leuzg is protected from
exchange by the surrounding residues and is weakly H-bonded to the backbone oxygen of
Asn24 (although it competes with the amide hydrogen of Glyzg). Interestingly, the amide
hydrogens of Gl)’23 and LCUzg exchange relatively slowly in monomeric PB, suggesting
that Pp adopts local structure in its B-hairpin (residues 5 —10). Finally, the Lyszo and
Alan amide hydrogens form a bifurcated H-bond with the side-chain oxygen of Asn24 in
the native structure of BPTI. These hydrogens are well protected in PM, and even more

so in Pap, consistent with a structured Asn24 side-chain. These two hydrogens are chieﬂy

154

responsible for the differences in the protection of the Pp subunit against HDX in Pap and
PM Combined with the CD results, these data suggest signiﬁcant native secondary and

side-chain structure in the P53 homodimer, as well as the Pug heterodimer [16].

 

Figure 4. 7 Ribbon diagram of the natively structured PB peptide; Asnz4 is indicated in
light gray and relevant hydrogen bonds are shown in dark gray. The mainchain-
mainchain H—bond donor/acceptor pairs from bottom: Phe33/Arg20, Phe22/Gln31,
Gln31/Phe22, Asn24/Leu29, Leu29/Asn24, and Glyzg/Asn24. The H-bonds between the amide
hydrogens of Lys7 and Alas to the sidechain atom 051 of Asns are also shown. The ﬁgure

was drawn using the MOLMOL software [36].

155

The absence of protection against HDX also can be a cross-check of native
structure; the amide hydrogens of Alazs, Cysm, and My should exchange readily

because they are solvent-exposed and not H-bonded in the native structure. The
increased structure in Pm; and Pug does not provide enhanced protection to Alazs (Table
4.2). Notably, Alazs does not fully exchange even in PB; this effect remains to be
explained, although slow exchange at this position has been noted in BPTI variants [45],
and may reflect a stable B-turn at residues Asn24-Leu29. The mass shifts of b], and blo
from PM; and Pug suggest that at least one residue in the segment Cys3o-Phe33 is
unprotected. Finally, the amide hydrogens of Tyr21 and Tyr23 should be protected in the
Pap dimer owing to intermolecular H-bonds with Phezz and Asnzo from the Pa subunit.
Consistent with this prediction, no protection is observed for Tyrz: and Wm in P5, but
modest and roughly equal protection of Tyr23 is observed in both P33 and Pap (Table 4.2).
This protection of Tyr23 in the P39 dimer suggests stable intermolecular H-bonding (and,
thus, tertiary structure) in Ppp, in addition to the secondary and side-chain structure cited

above.

156

IV. Conclusions

We have demonstrated the usefulness of HDX-MS, when combined with CD, for
studying conformational features of model peptide folding systems. Our protocol uses
MALDI-MS to assess overall deuterium incorporation and CID-MS/MS to obtain
residue-level HDX data. These data coupled with CD spectra (with thermal transitions)
allow us to assay secondary and other structural changes that occur upon dimer formation
in model peptides. Our method is general and should be useful in studying the physics of

protein folding and disorder.

Applied to peptide models from BPTI, our results provide evidence for the three
hypotheses outlined in the introduction. First, Pa shows some protection against HDX,
but the P,m dimer has almost as much protection as the native-like Pug heterodimer,
despite having signiﬁcantly less secondary structure than Pug (as assessed by CD). This
supports hypothesis (H1) that the association of two peptides can inhibit HDX by
lowering the local activity of water independently of secondary structure. Second, the
P,m dimer exhibits more secondary structure than the Pa monomer, even at 60 °C;
analogous results pertain to P3 and Pm}. This supports hypothesis (H2) that the decreased
local activity of water stabilizes local secondary structure. Finally, the Pm dimer does
not appear to be fully structured, judging from its non-helical CD spectrum; by contrast,
the data suggest that the Pap and P33 dimers have stable secondary, side-chain, and even
tertiary structure. Thus, our results are consistent with hypothesis (H3) that stable
secondary, side-chain, and even nonnative tertiary structure may develop when two
sufﬁciently nonpolar peptides are linked covalently. The difference between the PM and

Pap dimers may result from the hydrophobicity of the two peptides; PC, has only two large

157

hydrophobic residues (Phe45 and Metsz) of 16 residues, whereas PB has four (Tyr21, Phezz,
Tyrzg, and LCU29) of 14 residues. Therefore, the PM; dimer is better able to form a
hydrophobic core around its disulﬁde bond, further shielding its amide-amide H-bonds
from attack by water molecules. This is consistent with the global HDX results; PM;

exchanges 27% (3.5/ 13) of its amide hydrogens, whereas P,m exchanges 45% (6.8/ 15).

158

V. References

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[2] H. A. Scheraga, Theoretical and experimental studies of conformations of
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[3] H. Taniuchi, C. B. Anﬁnsen, Simultaneous formation of two alternative enzyrnology
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[4] G. T. Montelione, H. A. Scheraga, Formation of local structures in protein folding,
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[6] I. M. Klotz, J. S. Franzen, Hydrogen bonds between model peptide groups in solution,
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exchange and cyanylation-based methodology for conformational studies of
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[11] M. Y. Kim, C. S. Maier, D. J. Reed, M. L. Deinzer, Site-speciﬁc amide
hydrogen/deuterium exchange in E. coli thioredoxins measured by electrospray
ionization mass spectrometry, J Am Chem Soc 123 (2001) 9860-6.

[12] Y. Deng, H. Pan, D. L. Smith, Selective Isotope Labeling Demonstrates That
Hydrogen Exchange at Individual Peptide Amide Linkages Can Be Determined
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[14] X. Cai, C. Dass, Structural characterization of methionine and leucine enkephalins
by hydrogen/deuterium exchange and electrospray ionization tandem mass
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[15] P. K. Glasoe, F. Long, Use of glass electrodes to measure acidities in deuterium
oxide, Notes 64 (1960) 188-90.

[16] T. G. Oas, P. S. Kim, A peptide model of a protein folding intermediate, Nature 336
( 1988) 42-8.

[1 7] R. K. Scopes, Measurement of protein by spectrophotometry at 205 nm, Anal
Biochem 59 (1974) 277-82.

[1 8] Y. Bai, J. S. Milne, L. Mayne, S. W. Englander, Primary structure effects on peptide
group hydrogen exchange, Proteins 17 (1993) 75-86.

[1 9] Z. Zhang, A. G. Marshall, A universal algorithm for fast and automated charge state

deconvolution of electrospray mass-to-charge ratio spectra, J Am Soc Mass
Spectrom 9 (1998) 225-33.

[20] A. Wlodawer, J. Nachman, G. L. Gilliland, W. Gallagher, C. Woodward, Structure

of form III crystals of bovine pancreatic trypsin inhibitor, J Mol Biol 198 (1987)
469-80.

[21] K. Wuthrich, G. Wider, G. Wagner, W. Braun, Sequential resonance assignments as

a basis for determination of spatial protein structures by high resolution proton
nuclear magnetic resonance, J Mol Biol 155 (1982) 311-9.

[22] G. Wagner, K. Wuthrich, Amide protein exchange and surface conformation of the
basic pancreatic trypsin inhibitor in solution. Studies with two-dimensional
nuclear magnetic resonance, J Mol Biol 160 (1982) 343-61.

[23] K. S. Kim, C. Woodward, Protein internal ﬂexibility and global stability: effect of

urea on hydrogen exchange rates of bovine pancreatic trypsin inhibitor,
Biochemistry 32 (1993) 9609-13.

[24] G. Otting, E. Liepinsh, K. Wuthrich, Protein hydration in aqueous solution, Science
254 (1991) 974-80.

[25] M. Levitt, Effects of proline residues on protein folding, J. Mol. Biol. 145 (1981)
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[26] S. Cusack, J. Smith, J. Finney, B. Tidor, M. Karplus, Inelastic neutron scattering
analysis of picosecond internal protein dynamics. Comparison of harmonic theory
with experiment, J Mol Biol 202 (1988) 903-8.

[27] V. Daggett, M. Levitt, A model of the molten globule state from molecular dynamics
simulations, Proc Natl Acad Sci U S A 89 (1992) 5142-6.

[28] T. E. Creighton, Experimental studies of protein folding and unfolding, Prog
Biophys Mol Biol 33 (1978) 231-97.

[29] T. E. Creighton, D. P. Goldenberg, Kinetic role of a meta-stable native-like two-
disulphide species in the folding transition of bovine pancreatic trypsin inhibitor,
J Mol Biol 179 (1984) 497-526.

[30] J. S. Weissman, P. S. Kim, Kinetic role of normative species in the folding of bovine
pancreatic trypsin inhibitor, Proc Natl Acad Sci U S A 89 (1992) 9900-4.

[31] J. S. Weissman, P. S. Kim, The pro region of BPTI facilitates folding, Cell 71 (1992)
841 -5 1 .

[32] J. S. Weissman, P. S. Kim, Efﬁcient catalysis of disulphide bond rearrangements by
protein disulphide isomerase, Nature 365 (1993) 185-8.

[33] J. S. Weissman, P. S. Kim, Reexamination of the folding of BPTI: predominance of
native intermediates, Science 253 (1991) 1386-93.

[34] J. S. Weissman, P. S. Kim, A kinetic explanation for the rearrangement pathway of
BPTI folding, Nat Struct Biol 2 (1995) 1123-30.

[35] S. Parkin, B. Rupp, H. Hope, Structure of bovine pancreatic trypsin inhibitor at 125
K deﬁnition of carboxyl-terminal residues Gly57 and Ala58, Acta Crystallogr D
Biol Crystallogr 52 (1996) 18-29.

[36] R. Koradi, M. Billeter, K. Wuthrich, MOLMOL: a program for display and analysis
of macromolecular structures, J Mol Graph 14 (1996) 51-5, 29-32.

[37] J. P. Staley, P. S. Kim, Formation of a native-like subdomain in a partially folded
intermediate of bovine pancreatic trypsin inhibitor, Protein Sci 3 (1994) 1822-32.

[38] R. W. Woody, H. Sugeta, T. S. Kodama, [Circular dichroism of proteins: recent
developments in analysis and prediction], T anpakushitsu Kakusan Koso 41 (1996)
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[39] N. Sreerama, M. C. Manning, M. E. Powers, J. X. Zhang, D. P. Goldenberg, R. W.

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161

pancreatic trypsin inhibitor, Biochemistry 38 (1999) 10814-22.

[40] E. M. Goodman, P. S. Kim, Folding of a peptide corresponding to the alpha-helix in
bovine pancreatic trypsin inhibitor, Biochemistry 28 (1989) 4343-7.

[41] C. S. Maier, M. I. Schimerlik, M. L. Deinzer, Thermal denaturation of Escherichia
coli thioredoxin studied by hydrogen/deuterium exchange and electrospray

ionization mass spectrometry: monitoring a two-state protein unfolding transition,
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[42] M. Y. Kim, C. S. Maier, D. J. Reed, M. L. Deinzer, Conformational changes in
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[43] S. W. Provencher, J. Glockner, Estimation of globular protein secondary structure
from circular dichroism, Biochemistry 20 (1981) 33-7.

[44] S. Y. Tetin, F. G. Prendergast, S. Y. Venyaminov, Accuracy of protein secondary
structure determination from circular dichroism spectra based on immunoglobulin
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[45] B. A. Schulman, P. S. Kim, Hydrogen exchange in BPTI variants that do not share a
common disulﬁde bond, Protein Sci 3 (1994) 2226-32.

162

CHAPTER 5
IDENTIFICATION AND CHARACTERIZATION OF AN AUTONOMOUS

FOLDING UNIT FOR LRJIGF-I

I. Introduction

To clarify the mechanism of protein folding, great effort has been made to
characterize the 3-D structure of intermediates at various stages along the folding
pathway [1-4]. However, the cooperativity of protein folding and the limited solubility of
many folding intermediates often hinder researchers’ efforts [5, 6]. One way to
circumvent these problems is to design and characterize peptide models that can fold into
a native-like conformation. The protein subdomains that correspond to these peptide
models are called AFU [7]. AFUs from various proteins have been described and
characterized in recent years [8-10]. In this chapter, a new AFU for recombinant human
LR3IGF-I is described and characterized by HDX-MS combined with CD spectroscopy.

Recombinant human long R3 insulin-like grth factor-I (LR3IGF-I) is a highly
potent growth factor [11, 12]. Its amino acid sequence is shown in Figure 3.1.
Cysteinyl/cystinyl intermediates involved in progressive disulﬁde-bond formation during
the refolding of LR3IGF-I have been trapped and characterized by our
cyanylation/cleavage methodology [13]. During this study, it was interesting to observe
that only one of the ﬁfteen theoretically possible l-S intermediates, (namely, that
containing the disulﬁde bond Cy531—Cy874) accumulates to a signiﬁcant degree during the

oxidative refolding process (Figure 5.1). One explanation for this phenomenon involves

163

 

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Figure 5. 1 HPLC chromatogram showing the time-dependent distribution of CDAP-

trapped intermediates during the refolding of LR3IGF-I [13]. The disulﬁde structure of

each species is: I’ (Cys31 — Cys74), II’ (Cys31 — Cy574, CYS19 - Cysm), III’ (Cys31 —

Cy819 - Cysoo, Cysor -' Cysos), N (CYS31 - CyS74, Cy819 - CySm, Cysoo - CySos), R

(completely reduced). (experimental details on page 99 in Chapter 3)

164

protective tertiary structure formed via long-range interactions between the residues
ﬂanking this particular disulﬁde bond. According to this hypothesis, the disulﬁde bond
stabilizes the local structure, while the structure protects the disulﬁde bond, in turn, from
attack by reducing reagents in the system (such as the other free sulfhydryl groups in the
protein).

To test this hypothesis and investigate the relationship between disulﬁde bond
formation and the stabilization of local folded structure, peptide models were designed
and characterized that mimic the sequence of LR3IGF-I in the vicinity of Cys31 and Cysu.
Speciﬁcally, synthetic monomer P1 corresponds to residues 17 — 39
(RLSGAELVDALQFVCGDRGFYFS) in LR3IGF-I, and synthetic monomer Pn
corresponds to residues 54 — 75 (RGIVDEASFRSVDLRRLEMYCQ). Segment 17 — 39
corresponds to helix 1 in LR3IGF-I, and segment 54 — 75 corresponds to helix 2 and helix
3. As shown in Figure 3.3, helix 1 and helix 3 are connected by the disulﬁde bond
Cys31—Cys74. Because P1 and Pu each contain one cysteine residue, they can be disulﬁde—
bonded to form the PIP! or the PuPu homodimers or the PIP” heterodimer. The P1P"
heterodimer corresponds to the Cys31—Cys74 l-S intermediate that accumulates early
during the refolding of LR3IGF-I [13].

The two peptides, P1 and P", were synthesized and puriﬁed, and then used as
reagents to form the three dimers (PIPI, PnPn, and P1P") by air oxidation. The structure of
each of the ﬁve peptide models (PI, Pu, P1P], PnPn, and P1P") was probed using CD
spectroscopy and HDX-MS. CD reports on the secondary structure (i.e., or-helix and [3-
sheet structure) of each peptide model [14], whereas HDX-CID-MS monitors the degree

of protection of amide hydrogens against exposure to solvent [15-17]. The experimental

165

results demonstrate that signiﬁcant 3-D structure is associated with the formation of the
disulﬁde bond in each dimer. This observation supports our hypothesis that P; and P"

correspond to an AFU subdomain.

11. Materials and Methods
Materials

The P. and P" peptides were synthesized by the Genomics Technology Support
Facility (GTSF) at Michigan State University. The target peptides (approximately 10%
yield in the synthetic mixture) were then puriﬁed by rp-HPLC using a Vydac C13
analytical column (catalog #218TP54) in our laboratory. All other reagents were of the
highest purity commercially available and used without further puriﬁcation. Dibasic
sodium phosphate (NazHPO4) was purchased from Spectrum Chemical Mfg. Corp.
(Gardena, CA). ACN (UV grade) and ACN (HPLC grade) were purchased from
Honeywell International, Inc. (Muskegon, MI) and EMD Chemicals (Gibbstown, NJ),
respectively. Gdn-HCl was purchased from Invitrogen (Carlsbad, CA). All other
chemicals were purchased from Sigma.

All pH/pD values were measured using a Beckman (D 40 pH meter or ColorpHast
pH indicator strips (pH 0-14; EM Science, Gibbstown, NJ). pD values were computed by
adding 0.4 to the corresponding pH readings [18]. The standard buffer in this study
contains 0.1 M NaCl and 10 mM NaHzPO4, adjusted to pH 7.0. For the monomeric PI
and P" peptides, the buffer also contained 1 mM reduced dithiothreitol to prevent
dimerization of the monomers. No noticeable oxidation during the experiment was

detected as checked by rp-HPLC.

166

Preparation of the Peptide Dimers

The P1P; and PnPu homodimers and the P1P" heterodimer were prepared as
described previously [6]. Brieﬂy, roughly equal amounts of the P1 and P" monomers
were mixed in a solution containing 5 M Gdn-HCl and 200 mM Tris-HCl at pH 8.0 and
were air-oxidized for 48 hr at 25 °C. The dimers were separated and puriﬁed on a Vydac
C13 analytical column using a water/ACN gradient in the presence of 0.1% TFA. Each
dimer was collected manually and dried under reduced pressure for further use.
CD Spectroscopy

CD spectra were obtained at 0 °C on a J asco J-810 CD spectropolarimeter (J asco,
Inc., Easton, MD) using a therrnostatted 1-mm pathlength cell. The samples were
dissolved in the standard buffer. The concentrations were determined by UV absorbance
at 205 nm [19] and were typically 0.11 mg/ml.
HDX

The HDX experiments were carried out on ice. Each peptide sample was
equilibrated with the standard buffer (1 nmol/uL) for about half an hour before each
experiment. HDX was initiated by diluting 1 uL of aqueous protein solution with 19 uL
of standard buffer made with D20 at pD 7.4. After 60 s, 80 uL of buffer A solution
(containing 0.1% FA and 2.5% ACN) was added to quench the exchange by decreasing
the pH to 2.5 and to reduce the disulﬁde bond. The pH was maintained at 2.5 in all
subsequent steps. Incubation of the sample with D20 for 60 3 should be long enough to
exchange exposed amide hydrogens with deuteriums [20], but short enough to avoid

signiﬁcant exchange with H-bonded/protected amide hydrogens.

167

Analysis by CID Tandem Mass Spectrometry

De“ XP ion—trap mass

Mass spectra were acquired on a Finnigan LCQ
spectrometer (ThermoQuest, San Jose, CA) with an ion-spray voltage of 2.2 kV, a
capillary voltage of 43 V, a tube lens offset of 0 V, and a capillary temperature of 225 0C.
Data-dependent MS/MS conditions were set with a default collision energy of 40%, a
default charge state of 1, and an isolation width of 10 (m/z units). Scans were taken over
the range m/z 235 to 2000. Typically, 10 scans were accumulated per spectrum.

Immediately after quenching the HDX process, the sample was loaded onto a C13
trap (Upchurch, C281) using a pre—chilled syringe and ﬂushed with 100 [4L of buffer A
(pH 2.5) for 60 °C in for desalting. The processed sample was eluted into the LCQDeca
XP by adjusting the gradient to 50% buffer B (97.5% ACN, 0.1% FA) at a ﬂow rate of
100 uL/min; the injector and all the loops were buried in ice. The extent of back-
exchange during the procedures was 20%, as determined from analysis of the highly
deuterated standard prepared as described above. Mass spectra were obtained with the

operating software Xcalibur, and the centroid values were calculated with the Magtran

soﬁware[21].

168

III. Results and Discussion
A. Design of the peptide models
In native LR3IGF-I, the disulﬁde bond between Cys31 and Cysu connects two of
the three (it-helices (helix 1 and helix 3 in Figure 3.3). The core region of the designed
peptide model therefore includes residues that correspond to the three helices described
earlier. Residues outside this core region also were included if they satisﬁed either of the
following criteria:
(1) A residue that is conserved in the IGF family;
(2) A residue in one peptide interacts with residues in the second peptide according to
the NMR structure of LR3IGF-I [22, 23].
Our goal was to include all residues that satisfy these criteria, while keeping the peptides
as short and soluble as possible.
P1P" represents approximately half of LR3IGF-I (45 out of 83 residues) and
includes most of its hydrophobic core. The amino-acid sequences of P1 and P" are:
P; (17-39): RLSGAELVDALQFVCGDRGFYFS
Pu (54-75): RGIVDEASFRSVDLRRLEMYCQ
Pr contains 23 residues and corresponds to residues 17 — 39; this includes helix 1
(21 — 31) of LR3IGF-I. Pu contains 22 residues corresponding to residues 54 — 75; this
includes both helix 2 (56 - 63) and helix 3 (67 — 73). Eight residues of LR3IGF-I were
replaced to Optimize the solubility and stability of our peptides, and to encourage the
formation of b-ions during CID-fragmentation: in P1, Thrn was replaced by Arg, CYS19 by
Ser, and Asn39 by Ser; in P", Thr54 was substituted by Arg, CYSéO by Ala, Cys61 by Ser,

CYSﬁS by Val, and Ala75 by Gln. To improve solubility, we replaced the terminal residues

169

of our peptides with charged or more polar/ﬂexible residues: Thrn by Arg, Asn39 by Ser,
Thr54 by Arg, and Alan by Gln. We also eliminated four of the six cysteines, generally
by substituting them with Ala (when the cysteine was buried in the hydrophobic core) or
the isosteric residue serine (when the cysteine was exposed). Cys(,5 was mutated to Val
(instead of Ala) for two reasons: (1) to help ﬁll the void in the hydrophobic core left by
the elimination of the cyS60-‘Cy865 disulﬁde bond; and (2) Val forces the extended
conformation (ct = —80°, w = +139°) of the backbone at position 65. By contrast, the
Cysés to Ala substitution might have encouraged the formation of one long a-helix
(because Ala is helix-favoring), instead of the two broken helices observed in the native
structure (Figure 5.2). Throughout this chapter, the residues are numbered according to

their position in native LR3IGF-I (e.g., Cys3r, Va165, etc).

170

 

Figure 5. 2 The natively structured Pr and P11 [24]. P1 is depicted in dark gray, with its
initial residue, Argn, shown at the far left and its ﬁnal residue, Ser39, shown in the back
on the right. Similarly, P" is depicted in light gray, with its initial residue, Arg54, shown
at the bottom and its ﬁnal residue, Gln75, shown at the far right. The crossing-linking
disulﬁde bond, Cys3r—Cys74, is shown in the upper right. The side chains of the
substituted residues are also shown in dark gray (in P1) and light gray (in P“). The ﬁgure
was drawn using the MOLMOL software [25]. The side-chains were drawn using

SCWRL software [26].

171

B. CD results

To investigate the structural changes occurring upon disulﬁde bond formation, the
ﬁve peptides (P1, P", P1P", P1P], PUP") were ﬁrst studied using CD spectroscopy.
Although the absolute magnitude of the CD signal shown in Figure 4.3 is smaller than
expected ideally, the spectral shape is clearly consistent with a-helical structure [14]. As
shown in Figure 5.3 (A), the CD spectrum of P1P" at 0 0C shows minima near 208 nm and
222 nm, which are characteristic of helical proteins [14]; at 60 oC and 80 °C, these
troughs are replaced by one trough near 204 nm as expected for random-coil structure
[14]. The summed CD spectrum of P1 and P11 at 0°C is dominated by contribution from
random structure; the minimum near 222 nm suggests that a small population of the
monomers probably takes or-helical structure, although the majority of the population
does not. The summed spectrum resembles the spectra of P1P" at 60 °C and at 80 °C.
These results suggest that signiﬁcant oc-helical structure is formed in the PIP"
heterodimer, consistent with its native structure.

At 0 °C, the PIP; homodimer also gives CD signals characteristic of or-helical
structure; the two minima disappear as the temperature increases to 60 °C and 80 0C
(Figure 5.3 (B)). The CD spectrum of P; at 0 oC resembles that of PP at 60 °C and at 80
°C. The CD spectrum of PuPu at 0 °C is signiﬁcantly different from that of Pu, and the
minimum near 217 nm disappears gradually as the temperature increases (Figure 3 (C)).
This observation indicates that, although the majority of the P" monomers adopt a
random coil structure, a novel folded structure is formed in the PUP“ homodimer. The

changes in 3-D structure are probably caused by long-range interactions induced by the

172

formation of the disulﬁde bond. Interestingly, the CD spectrum of PuPn resembles that of

a pure B-sheet protein (a negative signal around 217 nm [14]), raising the possibility that

173

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PUP" adopts a different secondary structure than observed in the native structure of
LR3IGF-I.

In summary, the comparison between the CD spectrum of each dimer and its
corresponding monomer(s) suggests that formation of the disulﬁde bond induces the
formation of a more folded structure in all three dimers.

C. HDX results

During the last decade, HDX-MS has become more and more popular in the study
of protein conformation and conformational dynamics [15, 27, 28]. Here, HDX was used
to further characterize the structural features of each peptide model.

HDX results of the ﬁve intact peptides are reported as total deuterium
incorporation determined by the observed mass shift (Table 5.1). For monomeric P1 and
P", 76% (16.7 out of 22) and 86% (18.1 out of 21), respectively, of the total
exchangeable amide hydrogens were replaced by deuteriums from the solvent. Under the
experimental conditions used in this study, P1 and P11 show some protection against HDX,
which may reﬂect “ﬂickering” local structure in each monomer as indicated by our CD
results and which has been observed in other experiments [29, 30]. The three dimers
showed much less deuterium incorporation (9% for PIP", 14% for P1P], and 15% for
PuPu). In summary, all the dimers exhibit much more protection than their corresponding
free monomers. The HDX results are consistent with the observation in our CD
experiments that signiﬁcantly more folded structure is present in each dimer than in the
individual constituent monomers.

In a few studies, resolution of the loci of incorporated deuteriums has been

improved by analyzing the peptides by HDX-CID-MS [16, 17, 31, 32]. When a series of

177

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178

b-ions is produced during fragmentation by CID, the degree of deuterium incorporation
of individual amides can be assessed. Here, we applied CID-MS/MS to the ﬁve peptides
(Pl, Pu, P1P", P1P], and PUP"); the spectrum of each peptide is shown in Figure 5.4.

The fragmentation pattern of each dimer is similar to that of its constituent
monomer(s). During CID-MS/MS of the P1 monomer, major fragmentation takes place at
the C-tcrminal side of Phe36 and Try37, giving rise to b202+ and b212+ (Figure 5.4 (A)). For
the PIP] homodimer, major fragmentation takes place at the same sites, but in only one of
the two F; chains during the CID-MS/MS process, giving rise to analogous b-ions that are
more massive (shiﬁed by the mass of the P1 monomer); these ions are quadruply charged
as annotated in Figure 5.4 (B). As shown in Figure 5.4 (B), the most abundant ions
produced from PIPI are [(bzo of P1) + P1]4+ and [(1)21 of P1) + Pd“; When the P11 monomer
was subjected to CID-MS/MS fragmentation, y92+ and b132+ were the major ions produced
as a result of fragmentation at the C-terminal side of Asp66 (Figure 5.4 (C)). For the PUP"
homodimer, the same b132+ and bmz+ ions from one of the component P" chains are the
major products (but mass-shifted by the mass of P") of the CID-MS/MS process (Figure
5.4 (D)). In the case of the PIP" heterodimer, major CID-MS/MS fragmentation takes
place at the C-terminal side of Asp“ of the component Pu chain; b- and y-ions are
produced that are analogous to those produced from the Pu monomer, but mass shifted by
the mass of P; (Figure 5.4 (E)). In the heterodimer, the P1 chain and the P11 chain are held
together covalently by the disulﬁde bond between Cys3l (in P1) and Cys74 (in Pu). Thus,
the P1 chain is connected with the yg fragment of the P11 chain through this disulﬁde bond.
As shown in Figure 5.4 (E), the most abundant ions produced from P1P” are b132+ of P"

and [09 of P11) + P11“.

179

 

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Because each of the ﬁve peptides showed essentially only two or three channels
of fragmentation during CID-MS/MS, the ion current ﬁ'om the most populated channels
of b-ions was subjected to CID-MS/MS/MS in an effort to promote further
fragmentation. Representative spectra are shown in Figure 5.5 and Figure 5.6. Ions with
reasonable abundance were identiﬁed and their corresponding peaks were labeled in the
ﬁgures. None of the ﬁve peptides produces a series of b-ions or y-ions. Both P1 and P”
contain more than one basic residue (e.g., Arg) in each sequence, and, thus, the
fragmentation pattern of these peptides is complicated and unpredictable [33]. In this
study, only b-ions with reasonable abundance were used to assess the level of deuterium
incorporation in different sections of each peptide because b-ions of very low abundance,
as well as y-ions, are not reliable for assessment of deuterium incorporation [31, 32].

The HDX-CID-MS/MS and HDX-CID-MS/MS/MS results are summarized in
Table 5.2. Although single amino-acid residue resolution was not achieved due to the
absence of a series of b-ions for each sequence, the b-ions detected for each peptide
enable some insight into the deuterium incorporation of a few sections along each
polypeptide chain. All the sections along the polypeptide chain of the monomers are
signiﬁcantly deuterated. However, the deuterium incorporation level of each section
along the backbone of the dimers is limited. For instance, Argn-Aspzs of the P1 sequence
has 8 eligible amide hydrogens; 7 of these 8 amide hydrogens were replaced by
deuterium in the P1 monomer (Table 5.2 (A)); on the other hand, only 1.4 of them were
deuterated in the PIP; homodimer (Table 5.2 (B)). This observation is consistent with the

fact that the dimers adopt a signiﬁcantly folded structure, while the monomers do not.

185

 

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These results support our hypothesis that long-range interactions induced by the

formation of a disulﬁde bond can stabilize surrounding local folded structure.

IV. Conclusion

The peptide models (P1 and P") were designed to represent the amino acid
sequence in the vicinity of the Cy331-Cys74 disulﬁde bond in the l-S folding intermediate
of LR3IGF-I. It was assumed that they would mimic the conformational features
promoted by the corresponding environment in the intact protein. Our experimental
results (CD and HDX—MS) show that the native-like heterodimer (PxPn) and the two
homodimers (P1P; and PUP") adopt significant folded structure as compared with that in
the component monomers. During the folding of LR3IGF-I, long-range interactions
induced by the formation of the Cy831-Cy574 disulﬁde bond may stabilize the local folded
structure around it. The local folded structure, in turn, protects this disulﬁde bond against
attack from reducing reagents in the solution such as free sulfhydryl groups. This
provides a structural explanation for the observation that the Cys31-Cys74 l-S

intermediate accumulates at high levels in early folding intermediates of LR3IGF-I.

199

 

 

V. Future Work

Further experiments (i.e., size-exclusion chromatography) will be carried out to
investigate if any of the dimers aggregate to form non-covalently associated multimers
under our experimental conditions. If so, then the stacking of the peptide chains in the
multimer probably also contributes to the protection against HDX as observed in our
experiments. In addition, further interpretation of our experimental results will be
correlated with and rationalized by computer simulation of the folding process of each
peptide model. NMR and/or X-ray crystallography can be used to obtain detailed

conformational information of the peptide models.

200

 

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