ASSESSING THE ROLE OF STABILITY IN FIBRIL FORMATION IN
APOLIPOPROTEIN A-I VARIANT L178H USING UREA AND GUANIDINE
DENATURATION
A Thesis
Presented to the faculty of the Department of Chemistry
California State University, Sacramento
Submitted in partial satisfaction of
the requirements for the degree of
MASTER OF SCIENCE
in
Chemistry
(Biochemistry)
by
Trang Thanh Thi Duong
SUMMER
2012
© 2012
Trang Thanh Thi Duong
ALL RIGHTS RESERVED
ii
ASSESSING THE ROLE OF STABILITY IN FIBRIL FORMATION IN
APOLIPOPROTEIN A-I VARIANT L178H USING UREA AND GUANIDINE
DENATURATION
A Thesis
by
Trang Thanh Thi Duong
Approved by:
__________________________________, Committee Chair
Linda Roberts, Ph.D.
__________________________________, Second Reader
Mary McCarthy Hintz, Ph.D.
__________________________________, Third Reader
Jeffrey Mack, Ph.D.
____________________________
Date
iii
Student: Trang Thanh Thi Duong
I certify that this student has met the requirements for format contained in the
University format manual, and that this thesis is suitable for shelving in the Library
and credit is to be awarded for the thesis.
__________________________, Department Chair
Linda Roberts, Ph.D.
Department of Chemistry
iv
___________________
Date
Abstract
of
ASSESSING THE ROLE OF STABILITY IN FIBRIL FORMATION IN
APOLIPOPROTEIN A-I VARIANT L178H USING UREA AND GUANIDINE
DENATURATION
by
Trang Thanh Thi Duong
Apolipoprotein (apo) A-I is a major component of high density lipoprotein
(HDL) which removes and transports cholesterol from peripheral cells to the liver to
be excreted in a process known as reverse cholesterol transport (RCT). Plasma apo A-I
levels are correlated with reduced cardiovascular disease (CVD) risk (1,2),
presumably due to the role of apo A-I in RCT, as well as apo A-I’s demonstrated antiinflammatory and anti-oxidant properties (3). However, mutations in apo A-I result in
fibril formation, which can lead to severe diseases including atherosclerosis,
neuropathy, nephropathy, and peptic ulcer disease (4,5). These mutations cluster in
two regions, within the first 100 amino acid residues of the protein and in a small
cluster spanning residues 173-178. The two clusters appear to be associated with tissue
specific deposition of fibrils (6). Two full-length fibril-forming proteins with a
mutation in either the N-terminal region (G26R) or the C-terminal region (L178H)
have been studied. The G26R mutation leads to the formation of amyloid fibrils with
increased β-strand structure (7) that causes hereditary systemic amyloidosis, resulting
in neuropathy, nephropathy and visceral amyloidosis. The L178H mutation results in
helical fibril formation leading to fibril deposits in the larynx and in cardiac tissue (6).
A first step in understanding differences between the two mutation clusters and how
v
they are manifested in vivo is to explore the properties of variants from each cluster.
To that end, we are exploring the structure and stability of L178H. A common feature
of fibril-forming proteins is that they are generally less stable, and therefore more
prone to misfolding, than their wild-type (WT) counterparts (8-10). The goal of this
thesis project is to analyze the stability of L178H in comparison to that of wild-type
apo A-I using equilibrium solvent denaturation.
In this work, the protein stability and cooperativity of unfolding were
measured through the intrinsic fluorescence of tryptophan residues. From intrinsic
fluorescence measurements, the Gibbs free energy of protein unfolding (ΔG),
effectiveness of unfolding (m), and the midpoint of denaturation (D1/2) were obtained.
The results indicate L178H is less stable and unfolds less cooperatively than WT
protein. In addition, the stability of L178H under all conditions tested is dominated
more by hydrophobic interactions than electrostatic interactions compared to WT apo
A-I. These factors are likely responsible for the propensity of this mutant to form
fibrillar structures in vivo.
___________________________________, Committee Chair
Linda Roberts, Ph.D.
___________________________________
Date
vi
ACKNOWLEDGMENTS
I would like to thank the people who have helped me complete this thesis.
First, I thank my parents who have raised me and made me the person I am today.
Next, thanks to my committee members Dr. Roberts, Dr. McCarthy, and Dr. Mack,
who helped me through the whole thesis process. Thanks to the faculty and staff of the
chemistry department who helped me with various questions and problems: Dr. Dixon,
Dr. Baker, Dr. Gherman, Dr. Crawford, Ted, John, Michelle, Kathleen. Thanks to my
research group members and friends for their help and support: Aurelia, Himali,
Roman, Soraya, Megan, Tanya, Jessica, Jason, Kris, Yad, Holly, Sonya, Lauren.
vii
TABLE OF CONTENTS
Page
Acknowledgements ...................................................................................................... vii
List of Tables.................................................................................................................. x
List of Figures ............................................................................................................... xi
Chapter
1. INTRODUCTION ................................................................................................... 1
1.1 Atherosclerosis ............................................................................................. 1
1.2 Role of apo A-I and HDL Function ............................................................. 2
1.3 Apo A-I Structure......................................................................................... 3
1.4 Amyloid ..................................................................................................... 18
1.5 Apolipoproteins and Amyloid .................................................................... 23
1.6 Apo A-I Amyloid ....................................................................................... 25
1.7 The Current Study ...................................................................................... 27
2. EXPERIMENTAL ................................................................................................. 28
2.1 Materials ..................................................................................................... 28
2.2 Methods ...................................................................................................... 29
3. RESULTS AND DISCUSSION ............................................................................ 38
3.1 Protein Purification .................................................................................... 38
3.2 Equilibrium Solvent Denaturation ............................................................. 40
3.3 Intrinsic Fluorescence Measurements ........................................................ 42
viii
3.4 Comparison of Experimental Data to Literature Values ............................ 44
3.5 Stability of Initial Structure ........................................................................ 45
3.6 Stability Over 14 Day Period ..................................................................... 52
4. CONCLUSIONS AND FUTURE WORK ............................................................ 58
Appendix A: List of Abbreviations .............................................................................. 60
Appendix B: Experimental Data .................................................................................. 64
References .................................................................................................................... 66
ix
LIST OF TABLES
Tables
Page
Table 1.1. The eight classes of steric zippers ............................................................... 21
Table 1.2. Apo A-I mutations associated with amyloidosis......................................... 26
Table 3.1 Comparison of experimental values to literature values for WT protein. .... 45
Table 3.2. Comparison of L178H to WT at 4⁰C ......................................................... 47
Table 3.3. Comparison of L178H to WT after incubation at 37⁰C .............................. 48
x
LIST OF FIGURES
Figures
.........................Page
Figure 1.1. Atherosclerosis ............................................................................................ 1
Figure 1.2. Putative amphipathic helices in apo A-I protein.......................................... 5
Figure 1.3. Secondary structure of apo A-I based on different methods ....................... 8
Figure 1.4. Lipid-free apo A-I tertiary structure .......................................................... 10
Figure 1.5. Stabilizing interactions of the lipid-free ∆(185-243) apo A-I dimer ......... 12
Figure 1.6. Twisting of double belt structure ............................................................... 14
Figure 1.7. Monomer, dimer, and tetramer form of ∆(1-43) apo A-I .......................... 16
Figure 1.8. Proposed mechanism for lipid binding by apo A-I................................... 17
Figure 1.9. Overview of amyloid assembly mechanism .............................................. 19
Figure 1.10. Underlying structure of amyloid .............................................................. 20
Figure 1.11. β-sheet stacking interfaces ....................................................................... 20
Figure 1.12. Clustering of single amino acid mutations in apo A-I. ............................ 27
Figure 3.1. SDS-PAGE analysis of the integrity of WT and L178H protein............... 39
Figure 3.2. Structures of urea and guanidine hydrochloride. ....................................... 40
Figure 3.3. Emission spectra of WT apo A-I in varying denaturant concentrations .... 43
Figure 3.4. A) Denaturation curve for WT apo A-I monitored by intrinsic fluorescence
xxxxxxxxxB) Determination of ∆GH2O of WT apo A-I by linear extrapolation ........ 44
Figure 3.5. Urea induced unfolding of WT and L178H apo A-I at 4⁰C ...................... 46
Figure 3.6. Denaturation of WT and L178H apo A-I after incubation at 37⁰C ........... 48
xi
Figure 3.7. Location of the L178H and R173C mutations ........................................... 51
Figure 3.8. Percent of reduction in thermodynamic value of L178H and R173C
dddddddddcompared to WT at 4⁰C ............................................................................. 52
Figure 3.9. Change in free energy after incubation at 4⁰C ......................................... 54
Figure 3.10. Change in free energy after incubation at 37⁰C ..................................... 55
xii
1
Chapter 1
INTRODUCTION
1.1 Atherosclerosis
Atherosclerosis is a condition in which fat, cholesterol, calcium, and other
materials form deposits inside artery walls that can harden to form plaques (Figure 1.1)
(11). Atherosclerosis can affect many different organs such as the lungs, heart, kidneys,
brain, intestines and limbs. The plaques building up in these arteries can lead to
myocardial infarction (heart attack), stroke, coronary heart disease (CHD), abdominal
aortic aneurysm, kidney disease, mesenteric artery ischemia, peripheral artery disease,
renal artery stenosis, hypertension and thoracic aortic aneurysm. CHD is the number
one killer of men and women in the United States each year (12). In 2007, CHD killed
406,351 people. In 2008, it was estimated that 16.3 million Americans 20 years or older
had CHD (13). In addition, in 2007, 132,968 Americans died of heart attack and
135,952 died of stroke. These numbers indicate that atherosclerosis is a significant
health concern for Americans.
Figure 1.1. Atherosclerosis. Artery narrowed by plaque buildup, leading to heart attack
or stroke (14).
2
The risk factors associated with atherosclerosis, in particular CHD, are high
triglyceride levels, increased low density lipoprotein (LDL) cholesterol levels, and
decreased high density lipoprotein (HDL) cholesterol levels (15). HDL, commonly
referred to as the “good cholesterol,” has been shown to inversely correlate with CHD
risk, independent of triglyceride and LDL levels (16). That is, the more HDL present in
the plasma, the lower the risk for CHD.
1.2 Role of apo A-I and HDL Function
Apolipoprotein (apo) A-I is present in all HDL particles, and constitutes
approximately 70-100% of HDL proteins (17,18). The intestines and liver synthesize
and secrete apo A-I into the plasma as a lipid-poor protein. Immediately after secretion,
apo A-I acquires phospholipids and unesterified cholesterol from macrophages with the
help of ATP-binding cassette transporter A1 (ABCA1) to form discoidal HDL particles.
This nascent HDL acquires more phospholipids and unesterified cholesterol from other
peripheral cells such as skeletal myocytes, adipocytes, and skin fibroblasts. In addition,
HDL receives phospholipids from other lipoproteins by activity of the phospholipid
transfer protein (PLTP). Lecithin-cholesterol acyltransferase (LCAT) converts the
cholesterol into a cholesteryl ester by transferring a fatty acid from a phospholipid to
free cholesterol. The cholesteryl ester forms the hydrophobic core of HDL, which turns
the nascent HDL into spherical (mature) HDL. Cholesteryl ester transport protein
(CETP) transports the mature HDL to the liver where scavenger receptor class BI (SR-
3
BI) promotes uptake of cholesterol from HDL to eventually be excreted in bile and
feces. This process is known as reverse cholesterol transport (RCT).
As a primary factor in RCT, apo A-I has important anti-atherosclerotic
properties. Furthermore, apo A-I possesses both anti-microbial and anti-inflammatory
properties (19). Studies comparing apo A-I, HDL and LDL cholesterol level as markers
for cardiovascular mortality show apo A-I can be a powerful predictor of CHD
compared to HDL level (1, 2). In fact, there is a stronger correlation between CHD and
apo A-I level than between CHD and HDL level.
1.3 Apo A-I Structure
Since apo A-I is the main component of HDL, even compared to lipids, and the
primary factor in RCT, it is important to understand its structure at various points along
the RCT pathway. In human plasma, apo A-I exists in multiple lipid-free and lipidbound states. Approximately 5-10% of apo A-I in circulating plasma exists in a lipidfree state, while the rest is distributed among various lipid-bound states (20). Depending
on solution conditions, lipid-free apo A-I may exist in oligomeric forms of dimers,
trimers, tetramers and higher order oligomers. The physiological distribution of the
oligomer forms is unknown and may include amyloid or other fibrillar forms that
contribute to disease. Due to its importance in RCT and human health, the multiple
forms of apo A-I have received intensive study over the past 30 years. Although much
headway has been made in the past 10 years, significant gaps in understanding lipid-free
and lipid-bound structures remain. With the increased attention on apo A-I amyloid
4
formation, there is an increasing awareness of the importance of apo A-I selfassociation in solution and in binding to lipid. To fully understand how apo A-I forms
fibrils, a complete understanding of apo A-I structure in all forms is needed.
1.3.1 Apo A-I
The human apo A-I gene is located on chromosome 11 (6). The gene encodes a
pre-propeptide containing 267 amino acids (residues), which is cleaved intracellularly
to a 249 amino acid propeptide (21). After secretion into the plasma, the propeptide is
cleaved to yield a 243 amino acid, 28.1 kilodalton (kDa) polypeptide protein that lacks
glycosylation and disulfide linkages (20); the latter is due to the absence of cysteine
residues. The primary structure of apo A-I has been extensively analyzed using
computer programs developed to identify and characterize the signature motif of all
apolipoproteins, the amphiphatic helix (22). The computer models led to the
identification of amphipathic helix classes found predominantly in apolipoproteins: G*,
A, and Y (Figure 1.2). The G* helix, which is characterized by random radial
arrangement of positively and negatively charged residues on the polar face of the helix
(22), lies in residues 1-43 of the N-terminus of apo A-I. Residues 44-243 contain ten
amphipathic α-helices containing either 11 or 22 residues belonging to class A or class
Y (Figure 1.2). The class A amphipathic helix is associated with strong lipid-binding
properties and is defined by positively charged residues at the polar-non polar interface
and negatively charged residues at the center of the polar face. Class Y helices contain
two negative residue clusters on the polar face and three positive residue clusters
5
forming a Y motif. In general, the lipid affinity for the three amphipathic helix classes is
A>Y>G*.
Figure 1.2. Putative amphipathic helices in apo A-I protein (22).
Based on the proposed placement of amphipathic helices in apo A-I by these
computer programs, Mishra et al. synthesized the peptides for each putative helix to
probe their lipid-associating properties (23). The order of lipid affinity was determined
to be helix 9-10 >1-2>4-5 (Figure 1.2); the other helices do not possess much affinity.
Researchers subjected the full-length protein to point mutations and or deletion
mutations to analyze apo A-I based on its lipid binding affinity and ability to efflux
cholesterol. The resulting data suggests helices 1, 9, and 10 function to initiate the
binding of lipid, helix 4 and 5 promote HDL formation, and helix 6 and 7 activate
LCAT (Figure 1.2) (24, 25).
1.3.2 Apo A-I Secondary Structure
Extensive analysis of the primary sequence suggests the type and number of
amphipathic helices in apo A-I. A number of experimental approaches have been used
to determine the actual helix content. Collectively, these approaches indicate lipid-free
apo A-I contains 50-60% helix (26-30), whereas lipid-bound contains approximately
6
70-80% helix (31-34). The placement of helices using different methods has given
conflicting results, especially for the lipid-free protein as discussed below.
The full-length apo A-I in solution has a flexible and adaptable structure, which
makes it difficult to determine the lipid-free as well as lipid-bound secondary structure
using direct methods such as nuclear magnetic resonance (NMR) and X-ray
crystallography (X-RC). Thus, secondary structures were determined using both
indirect methods on full-length and direct methods on the less flexible truncated apo A-I
mutants. The location of helices in lipid-free apo A-I was first suggested by a mixture of
indirect methods (MIM) which include: circular dichroism (CD), sedimentation
velocity, ANS binding, fluorescence quenching and limited proteolysis of full-length
and truncated protein (35). The data from MIM were used to create a structural model
of lipid-free apo A-I that contains six helices near the N-terminus (residues 1-189) and
no helices near the C-terminus (residues 190-243) (Figure 1.3). The structure contains
60% helix, generally consistent with CD analysis of lipid-free apo A-I. More than a
decade after the MIM model, a second model was determined by hydrogen exchange
and mass spectrometry (HX-MS; 26). This second model has five helices instead of six,
and the placement of the helices is somewhat different from that of the MIM model.
However, the HX-MS model has 51% helix content, similar to that of the MIM model
and also consistent with CD analysis. A third model, generated from the structure of a
truncated apo A-I (∆(185-243) apo A-I) that was determined by X-RC (36), has five
helices near the N-terminus, but the placement of helices is different from both the MIM
7
and HX-MS model. The three secondary structural models are similar in that they all
have helices that span residues 8-33 and residues 44-54 and no helices for residues 135141. A fourth model was proposed using data from electron paramagnetic resonance
(EPR; 37). This structure has 53% helix content, consistent with CD analysis, but it is
not in good agreement with the previous three models, having helices placed in the Cterminal region and containing four β-strands (12% β-strand content). The β-strand
content from the EPR model is consistent with CD analysis and Fourier transform
infrared spectroscopy (FTIR) data which indicate apo A-I contains 8-16% β-strand
content (27-30). A small amount of β-strand structure could have important
implications for fibril formation, and the EPR data has been used to propose a model of
lipid-free apo A-I that contains β-strand stabilizing regions. Until a high resolution
structure of the full-length protein is determined, the existence and role of β-strands, as
well as absolute placement of helices, remains unclear.
There are two secondary structural models proposed for lipid-bound apo A-I.
The first model was based on X-RC data (38) of a truncated lipid-free protein (∆(1-43)
apo A-I), but it is believed to represent the lipid-bound structure for three reasons: It has
a different secondary structure (increased helical content) compared to full-length apo
A-I, its secondary structure does not change when it binds to lipid, and it has a
proteolytic cleavage pattern similar to that of full-length lipid-bound apo A-I. The
second model was based on NMR data for the full-length protein in the presence of
sodium dodecyl sulfate (SDS), a lipid-mimicking solution (39). Both structural models
8
differ in the exact placement and amount of α-helices (Figure 1.3), but they both place
helices throughout the length of apo A-I. They also have helices in the C-terminus,
absent in most of the proposed lipid-free structures. The full length NMR structure has
75% helical content, consistent with CD analysis of lipid-bound apo A-I. The truncated
∆(1-43) apo A-I has 93% helical content.
Figure 1.3. Secondary structure of apo A-I based on different methods. The boxes
denote α-helix, and the arrows denote β-sheet. The structural models are derived from
A) combination of methods including CD, sedimentation velocity, ANS binding,
fluorescence quenching, and limited proteolysis of truncated mutant (35), B) hydrogen
exchange and mass spectrometry (26), C) X-ray crystallography of ∆(185-243) apo A-I
(36), D) EPR of full-length apo A-I (37), E) X-ray crystallography of ∆(1-43) apo A-I
(38), and F) NMR of apo A-I in SDS (39)
In summary, due to apo A-I's inherent flexibility (and therefore intractability for
analysis by X-RC and NMR), and despite decades of research there is still no single
9
unified model that researchers can agree upon for the secondary structure of apo A-I.
The same can be said of the tertiary structure, although, as discussed below, there is
some agreement on its general features.
1.3.3 Apo A-I Tertiary Structures
In humans, the total serum apo A-I concentration is approximately 1 mg/ml,
with the lipid-free concentration being less than 0.1 mg/ml (40). At concentrations <0.1
mg/ml, apo A-I self-association is disrupted and apo A-I is assumed to be monomeric.
At ≥ 0.1 mg/ml, apo A-I is found to self-associate due to hydrophobic interaction
between amphipathic helices, forming oligomers (41). Apo A-I self-associates in both
lipid-free and lipid-bound states, but the mechanism of self-association is unknown.
Tertiary structure models of the protein in the presence and absence of lipid must take
into account both monomeric and oligomeric states especially in light of the importance
of these states in amyloid formation. While the self-associated structure of lipid-bound
apo A-I is fairly well understood, that of lipid-free apo A-I is not.
Several models of lipid-free apo A-I tertiary structures have been proposed.
Early studies using MIM (refer to section 1.3.2) led to one of the first models of lipidfree monomeric apo A-I (Figure 1.4A). The MIM data suggests monomeric apo A-I has
two structural domains. The N-terminal domain (residues 1-189) forms a helix bundle
and the C-terminal domain (residues 190-243) is mostly unstructured (35). Two separate
subsequent studies, deletion mutagenesis (42) and Förster Resonance Energy Transfer
(FRET; 43), also support the two domain structure. The deletion mutagenesis data
10
indicate that, when a part or the entire N-terminal domain is deleted, the number of
residues that form helices decreases significantly, indicating the N-terminal domain
forms some kind of helical structure. When the C-terminal domain is deleted, there is no
change in the number of residues that form helices, indicating the C-terminus is
unstructured. The FRET study measured the distance between W50-R83 and W50-R173
in apo A-I, which proved to be consistent with a compact N-terminal structure such as a
helical bundle.
Figure 1.4. Lipid-free apo A-I tertiary structure. A) Structure of monomer of full length
apo A-I determined by MIM (35). B) Structure of dimer of ∆(185-243) apo A-I
determined by X-RC (36). C) Structure of proposed monomer and dimer of full length
Apo A-I determined by EPR (37)
The X-RC study of the truncated ∆(185-243) apo A-I protein led to a model of
the lipid-free dimeric protein (Figure 1.4B) (36). The X-RC model indicates the
monomers in the dimer lie in an antiparallel position to each other and form a
semicircular structure with a 17 Å height and 110 Å diameter (Figure 1.5). The dimer
contains two N-terminal helix bundles. Each helix bundle consists of three helices
11
(residues 7-41, 55-65, 69-121) of one apo A-I molecule, bundling together with one
helix (residues 143-185) of the second apo A-I molecule. The N-terminal helix bundle
is held together by four stabilizing interactions: two by the N-terminal and C-terminal
aromatic clusters and two by the N-terminal and C-terminal π-cation interactions. The
aromatic clusters are composed of aromatic residues at each terminus, forming a
hydrophobic interaction. The π-cation interactions at each terminus contain an aromatic
residue and a cation residue, forming a π-cation bond. The N-terminal aromatic cluster
involves nonpolar interactions between W8, F71 and W72 of one monomer and nearby
leucines (14, 60, 64, 75 from the same monomer and 170, 174, 178, 181 from the
second monomer), holding residues 44-66 and 67-77 close together (44). The Caromatic cluster consists of F33, F104, and W108 of one monomer and nearby
aliphathic residues (V30, L38, L42, L44, L46 from the same monomer and V156, L159
from the second monomer), holding the N-terminal helix close to residues 99-121. The
N-terminal π-cation interaction consists of W8 and R61 in each monomer positioning
residues 55-66 close to residues 1-43 and covering the N-aromatic cluster (W8, F71,
and W72) of the same monomer. The C-terminal π-cation interaction consists of K23
and W50 in each monomer, holding residues 55-66 toward residues 1-43. In addition to
interactions stabilizing the helix bundles, the central hinge segment (residues 121-143)
connects the two monomers via a leucine zipper composed of L122, 126, 137, and 141
from each monomer. Additional stabilization is provided by intramolecular salt bridges
(E5-R9, R11-E15-R18, E16-R20, E4-R7, K88-D89, E120-R123) and intermolecular
salt bridges (K96-E69, D89-R173-E92, E85-R177-D89).
12
Figure 1.5. Stabilizing interactions of the lipid-free ∆(185-243) apo A-I dimer. One apo
A-I molecule is gray and the other is black (36).
The EPR study of full-length apo A-I plus spin coupling data led to a different
model of the lipid-free dimeric protein (Figure 1.4C) (37). The spin coupling data
indicate that, for each monomer in the dimer, residues 24, 44, 64, 167, 217, and 226
align to form a helical bundle. The four β-strands serve as stabilizing structures. The
two center β-strands (residues 102-115 and 130-148) form a hairpin with a 15 residue
loop to stabilize the center of the protein. The N-terminal β-strand (residues 20-25)
stabilizes the N-terminus to the rest of the protein. The C-terminal β-stand (residues
214-220) stabilizes the C-terminus to prevent it from unraveling into random coils in
13
the absence of lipids, through hydrophobic association with the rest of the protein. An
important feature of this model is a loop spanning residues 130-139 that initiates lipid
binding. The dimer is formed from the anti-parallel association of two monomers
aligned at residues 24, 44, 64, 167, 217, and 226. The authors suggest the two center βstrands, which flank the loop in each monomer, convert to α-helices as the protein binds
lipids, so that the protein can re-arrange itself into the lipid-bound conformation.
The X-RC and EPR tertiary structural model of lipid-free dimeric apo A-I are
substantially different. The X-RC model is of a truncated apo A-I instead of full-length,
but X-RC has the advantage of being a high resolution method. The EPR method has
lower resolution, but it has the advantage of being conducted on the full-length protein.
The EPR model also accounts for the small amount of β-structure measured in lipid-free
apo A-I by CD (29,30) and FTIR (28). Until the full-length structure is resolved by high
resolution techniques such as X-RC and NMR, it is difficult to determine which of the
two models is more accurate.
Lipid-bound apo A-I exists in multiple forms of discoidal and spherical HDL.
These HDL have various lipid compositions and proteins in addition to apo A-I, such as
apo A-II and apo C, resulting in different HDL sizes and shapes (45). Lipid-bound apo
A-I has been studied most often in reconstituted HDL (rHDL), which are created by
combining
lipid-free
apo
dimyristoylphosphatidylcholine
A-I
(DMPC)
with
or
synthetic
lipids,
usually
palmitoyloleoylphosphatidylcholine
(POPC). The rHDLs range from 7-12 nm in diameter and contain 2-5 molecules of apo
A-I (45). Currently, there is a lack of substantial data on spherical rHDL and rHDL
14
containing more than two apo A-I molecules; thus, the focus here will mainly be on
discoidal rHDL containing two molecules of apo A-I. Indirect methods including crosslinking studies (45), FRET (46) fluorescence quenching (47), and computational
experiments combined with NMR spectroscopy and electron microscopy (48) all
support a double belt model for lipid-bound protein (Figure 1.6). In this model, two
anti-parallel apo A-I molecules wrap around a lipid bilayer to form discoidal HDL. The
structure is stabilized by extensive intermolecular salt bridges between the two apo A-I
molecules, in which residue 121-143 of one molecule directly overlaps residues 121143 of the second molecule (49). Computational analysis suggest the size variation in
discoidal HDL that contains only two apo A-I molecules is due to the incremental
twisting or untwisting of the double belt structure (Figure 1.5;48). When the double belt
structure is twisted, it resembles the shape of a saddle (“saddle-shaped” structure), but
the salt-bridges linkage remains the same and this results in smaller HDL particles.
Figure 1.6. Twisting of double belt structure. A) Conversion of saddle-shaped structure
to double belt structure, the black arrows denote the N-terminus. B) Same as in A, but
shown with lipid molecules in black (48)
15
The X-RC structure of ∆(1-43) apo A-I (Figure 1.7) was the first crystal
structure of apo A-I to be reported, and it supports the double belt model (38). The ∆(143) apo A-I tertiary structure was obtained in lipid-free conditions. However, for the
reasons discussed above, it is believed to represent lipid-bound structure (as discussed
above). The X-RC data indicates that ∆(1-43) apo A-I protein assumes a tetrameric
structure, formed from two dimers. It has an elliptical shape with dimensions 125 x 80 x
40 Å, suitable for wrapping around a lipid disc to form HDL. The monomers within the
dimer lie antiparallel to each other, and the dimers in the tetramer are also in an
antiparallel position. When the ∆(1-43) apo A-I monomers form a dimer (Figure 1.7),
21% of the surface of each monomer is no longer solvent accessible. Half of this 21% is
due to hydrophobic interactions and 30% is due to intermolecular salt bridges formed
between Arg/Lys and Asp/Glu residues. The dimer possesses both a hydrophobic and
hydrophilic face. The structure of ∆(1-43) apo A-I strongly supports the double belt
model. The hydrophobic interaction and intermolecular salt bridge found in the ∆(1-43)
apo A-I dimer is also comparable to those discussed in the ∆(185-243) apo A-I dimer.
When the dimers combine to form the tetramer, the buried surface area of each
monomer increases to 42% and the lipid-binding regions of the protein are buried
within the tetramer (Figure 1.7). It is suggested that because the X-RC structure of ∆(143) apo A-I was acquired in a lipid-free condition, the dimers combined to form
tetramers to shield the hydrophobic lipid-binding region from water. Due to the buried
lipid-binding regions, the authors believe the dimeric rather than the tetrameric structure
of ∆(1-43) apo A-I represents the true lipid-bound structure.
16
Figure 1.7. Monomer, dimer, and tetramer form of ∆(1-43) apo A-I determined by XRC (38).
1.3.4 Lipid-free to Lipid-bound Mechanism
Structural models of lipid-free and lipid-bound apo A-I can be used to propose a
mechanism for how lipid-free apo A-I binds to lipid and forms HDL. Based mainly on
the X-RC data of ∆(185-243) apo A-I, Mei and Atkinson proposed a mechanism in
which lipid-free apo A-I converts to lipid-bound apo A-I (Figure 1.8). In plasma, apo AI is represented as monomers. As the monomers interact with lipids, they come in close
contact to each other and form a dimer with a semicircular structure. The π-cation
interactions that hold residues 44-66 in place serve as a gate for lipids to enter into the
hydrophobic core of the protein. As lipids enter the hydrophobic core, the helices in the
N-terminal helix bundle move away from each other to open up the bundle. This is due
to the disruption of the four stabilizing interactions (Figure 1.6). The hydrophobic core
is exposed, and the dimer is inserted into the lipid membrane surface. This in turn
unhinges the N-terminal helix bundle and causes residues 77-99 to extend out, forming
a double belt structure, resulting in discoidal HDL formation. The hydrophobic surface
of the dimer faces the inside of the belt structure and the hydrophilic surface faces the
17
outside. This double belt structure is strongly supported by the X-RC structure of ∆(143) apo A-I.
Figure 1.8. Proposed mechanism for lipid binding by apo A-I (36).
Apo A-I is quite flexible because it needs to unfold and wrap itself around lipids
to form HDL. However, this may leave the protein prone to misfolding, which can have
several consequences, including lack of stable HDL formation or increased abnormal
self-association, leading to the formation of insoluble aggregates or fibrils. A number of
fibril-forming or amyloid variants of apo A-I have been identified, and because WT apo
A-I is found in arterial plaque, it is important to understand the factors that can cause or
accelerate improper folding in apo A-I. The propensity of protein in general to misfold
and the research conducted to identify misfolding mechanisms is described in the next
section.
18
1.4 Amyloid
In 1854, Schleiden and Virchow coined the term “amyloid” to describe pale and
waxy looking macroscopic tissue that produced a positive iodine staining similar to
starch, found during organ autopsies (50). Thus, amyloids were initially thought to be
carbohydrates. In 1859, amyloids were discovered to be proteins when their high
nitrogen content was determined (50). Research conducted in 1927 found amyloids
specifically bind congo red and produce an apple-green birefringence under polarized
light suggesting their fibrillar structure, and the fibrillar structure was confirmed by
electron microscopy in 1959. X-ray diffraction analysis in 1968 found the fibrils contain
β-sheets running perpendicular to the fibril axis (50). Years of subsequent research
indicated amyloids form from specific proteins, and these proteins become
amyloidogenic upon denaturation of tertiary and quaternary structures in vitro. X-ray
crystallography analysis of short amyloidogenic peptides led to a new definition for
amyloids, which are protein fibers of varying length, consisting of repeating β-sheets
running perpendicular to the fibril axis (cross-β-sheet) (51). Two or more stacked βsheets form a filament, two or more filaments form a protofibril, and two or more
protofibrils form a fibril (Figure 1.9). The cross-β-sheet can either be parallel or antiparallel and has a characteristic X-ray fiber diffraction pattern with 4.7 Å meridional
reflection and 6-11 Å equatorial reflection (Figure 1.10 C). The meridional reflection
corresponds to the inter-β-strand spacing and the equatorial reflection corresponds to
the distance between stacked β-sheets (Figure 1.10B). All fibrils have helical symmetry
as well as translational symmetry parallel to the fibril axis. The β-sheet stacking
19
contains two types of interfaces: wet and dry. The wet interface has hydrogen bonds
between the side chains and water molecules. The dry interface has complementary
side chains interlocking to form a steric zipper and is empty of water molecules (Figure
1.11). The interactions that stabilize the fibril are both polar (alternating charges and
hydrogen-bond ladders) and nonpolar (Van der Waals dispersion and aromatic
stacking). Despite all these same characteristics in many amyloids, there are also
important differences among the amyloids.
Figure 1.9. Overview of amyloid assembly mechanism (52).
20
Figure 1.10. Underlying structure of amyloid. A) Electron micrograph of amyloid
fibrils. B) Schematic diagram of the cross-β-sheets in a fibril. C) Diffraction pattern of
amyloid fiber showing meridianal reflection (black dashed box) and equatorial
reflection (white dashed box;51)
Figure 1.11. β-sheet stacking interfaces. Pictured are six rows of β-sheets that run
horizontally. The + signs represent water molecules. The big dashed box indicates one
row of β-sheet. The small dashed box highlights the steric zipper formed by interlocking
side chains (53).
Amyloid structure can vary in terms of the numbers of wet and dry interfaces
making up a fibril. The dry interface contains steric zippers and there are eight possible
classes of steric zipper structures, five of which have been observed experimentally.
These steric zippers differ by whether the β-strands are parallel or anti-parallel, are
packed face-to-face or face-to-back, and are packed up-up (both sheet have the same
21
edge up) or up-down (the sheets have one edge up and one down) (Table 1;54).
Amyloid fibrils can also vary by the number of protofibrils. There are four types of
polymorph (multiple structure states) amyloids can adopt: packing, segmental, sidechain, and assembly. The polymorphs differ in which segment of the protein forms the
amyloid fibril and the orientation of the side chains in the fibril. Multiple polymorphs
can exist in a fibril.
Table 1.1. The eight classes of steric zippers
Example
protein
Class
β-strand
1
parallel
up-up
face-to-face
tau, sup35, RNase
2
parallel
up-up
face-to-back
prion
3
parallel
up-down
face-to-face
not yet observed
4
parallel
up-down
face-to-back
amyloid-β, sup35
5
anti-parallel up=down
face-to-face
not yet observed
6
anti-parallel up=down face-to-back
not yet observed
7
anti-parallel
up-up
face=back
insulin
8
anti-parallel
up-down
face=back
amyloid-β
Packing
Structure
22
1.4.1 Amyloid Properties
There are functional as well as toxic amyloids. Disease caused by toxic
amyloids is described by the term amyloidosis. For the purpose of this discussion, the
focus will be on toxic amyloids. Since the cross-β fold in amyloid can be as simple as
having four residues of β-strand that assemble repetitively, any protein that can form βstrands can eventually form amyloids. In fact, many proteins aggregate into amyloids or
amyloid-like states under abnormal conditions such as high protein concentration, heat
denaturation, extreme pH and non-aqueous solvents. However, even under normal
conditions, some properties of the polypeptide itself can lead to amyloid formation. The
primary mechanism for amyloid formation is destabilization of the folded (presursor
amyloid protein) structure, leading to misfolding and aggregation (55). Studies of single
point mutations in amyloidogenic proteins, such as human lysozyme, transthyretin and
human immunoglobulin light chain, indicate that the mutants that form amyloid are
significantly less stable than the wild-type counterparts (8-10). Hierarchical analysis of
2351 point mutations in 44 globular proteins using empirical tools indicates that the
magnitude of destabilization in a mutant protein may correlate with amyloid formation
(56). It has been found that, in general, mutants that are more stable than the wild-type
counterparts do not form amyloids (10).
Besides the mutations that result in a globally unstable structure, amyloids can
form from mutations that change a specific physical property of the protein. Analysis of
protein sequences using the Tango algorithm, a computer algorithm that predicts
23
aggregating regions in unfolded polypeptide chains, found proline (P), glycine (G),
lysine (K), arginine (R), glutamate (E), and aspartate (D) residues play a protective role
in flanking aggregating stretches within proteins (55). Mutations that reduce the
number of P, G, K, R, E, and D residues increase the propensity for aggregation.
Proline’s structural constraints make it difficult to form β-sheets; whereas glycine is
small, has no β-carbon atom, and is highly flexible, requiring a high entropic cost to
form it into β-sheets (55). Meanwhile, K, R, E, and D residues have the lowest
hydrophobicity and are charged, also making it hard to form β-sheets. Properties that
increase a protein’s propensity to aggregate into amyloids are long stretches of five or
more hydrophobic residues, increased hydrophobicity in the protein core, decreased
overall net charge, increased clustering of hydrophobic residues in a region, and high
sequence identity between adjacent domains. These properties all make β-sheet
formation in proteins easier.
1.5 Apolipoproteins and Amyloid
Currently, over 25 proteins have been shown to form amyloids, contributing to
diseases such as Alzheimer’s, Parkinson’s, the prion diseases and adult onset diabetes
(57). Proteins in the apolipoprotein family also form amyloids that lead to disease.
Understanding how apolipoproteins as a family form amyloids will aid in understanding
how apo A-I forms amyloid because of the shared characteristics among these proteins.
Apolipoproteins contain a high proportion of class A and Y amphipathic α-helices (57).
Their function involves binding to lipids to form lipoprotein particles. Despite their
24
generally high helical content, which results in stable structures (57), they are prone to
amyloid formation. For example, one of the three apo E isoforms is associated with
enhancing amyloid formation of Aβ peptide (Alzheimer’s disease), while apo C-II
spontaneously forms amyloids in lipid-free conditions and was found in atherosclerotic
plaques. Mutations in apo A-II are responsible for familial systemic amyloidosis, and
mutations in apo A-I lead to atherosclerosis and tissue specific amyloid deposition
throughout the body.
It is not clear why apolipoproteins form amyloids, but studies of apo E isoforms
and apo C-II indicate that low conformational stability may be the main factor
contributing to amyloid formation. Studies of apo E revealed that apo E fragments form
helical fibrils and full-length apo E forms amyloid-like fibrils. The rate of apo E
forming amyloid-like fibrils is related to the conformational stability of the N-terminal
domain. Further investigation indicates that mutations that lower the conformational
stability of the N-terminal portion of the protein increase the rate of apo E fibrillation
(58). Apo C-II, unlike apo E, has no ordered structure (helix or β-strands) in lipid-free
conditions and spontaneously forms amyloid (59). In a high micellar lipid environment,
apo C-II forms structures with stable conformations that do not form amyloid (57).
The disordered structure of apo C-II in lipid-free condition is less stable than its ordered
structure in the lipid environment. These studies with both apo E and apo C-II indicate a
decrease in conformational stability leads to fibril formation. Thus, low conformational
stability may also be a factor in amyloid formation in apo A-I.
25
1.6 Apo A-I Amyloid
There are 19 mutations in apo A-I that lead to amyloid formation (Table 2). Of
the 19 mutations, 12 are single amino acid mutations that cluster in two regions,
spanning residues 26-90 and 170-178 (Figure 1.12). Both clusters are within the Nterminal helix bundle (residues 1-189). Mutations near the N-terminus (“inside”
mutations) lead to hereditary systemic amyloidosis, while mutations nearer to the Cterminus (“outside” mutations) lead to specific amyloid deposits in the larynx, cardiac
tissue, kidneys, and cutaneous tissues (60). The mechanism of tissue-specific deposition
of these amyloid variants is unknown. Mutations in both clusters result in fibrils in vivo
consisting mainly of an N-terminal peptide of 80-100 residues in length, commonly 193 (61).
Two full-length fibril-forming proteins, each with a mutation in either the Nterminal region (G26R) or the C-terminal region (L178H), have been studied by our
laboratory in conjunction with Dr. John Voss at University of California, Davis, and Dr.
Jens Lagerstedt at Lund University in Sweden. These are the only studies to date on
full-length fibril-forming mutants of apo A-I. We found that the G26R mutation, which
causes neuropathy, nephropathy and visceral amyloidosis (5), leads to the formation of
amyloid fibrils with increased β-strand structure (62). In contrast, the L178H mutation,
which deposits fibrils in the larynx and in cardiac tissue (5), results in helical fibril
formation (63). We hypothesize that while lowered stability may underlie fibril
formation in mutants from both clusters, fibril type may be determined by mutation
26
Table 1.2. Apo A-I mutations associated with amyloidosis
Apo A-I
Variant
Clinical Features
Renal impairment, Hepatomegaly,
Peripheral neuropathy, Nephropathy,
Gly26Arg
Peptic ulcer disease, Visceral
amyloidosis,
Glu34Lys
Renal impairment
Renal impairment, Hepatomegaly,
Trp50Arg
Nephropathy, Visceral Amyloid
Renal impairment, Hepatomegaly,
Leu60Arg
Cardiomyopathy, Nephropathy,
Visceral Amyloid
Leu64Pro
Renal impairment
Phe71Tyr
Palatal mass, (normal organ function)
Leu60Hepatomegaly, Visceral Amyloid
Phe71delins6
particularly in liver
Glu70_Trp72del Renal impairment
Renal impairment, Gastrointestinal
Asn74LysfsX10
amyloid, amyloid detected in uterus,
6
ovaries, pelvic lymph nodes
Hepatomegaly, Renal impairment,
Leu75Pro
hepatic amyloid, gastrointestinal
amyloid
Skin lesions, cardiomyopathy,
dysphonia, nephropathy, visceral
Leu90Pro
amyloid, dermal amyloid, laryngeal
amyloid,
Angina associated with amyloid in
Lys107del
aortic intima
Ala154GlyfsX48 Renal impairment, polyneuropathy
His155MetfsX46 Renal impairment, polyneuropathy
Leu170Pro
laryngeal amyloid
Dermal amyloid, vocal cord amyloid,
Arg173Pro
visceral amyloid, cardiomyopathy,
Infertility, cardiomyopathy, bilateral
Leu174Ser
carpal tunnel syndrome,
polyneuropathy, systemic amyloid
Laryngeal amyloid, Infertility, visceral
Ala175Pro
amyloid
Dermal amyloid, laryngeal amyloid,
Leu178His
neuropathy, cardiomyopathy
Cause of Death
Refs
Renal Failure
5, 6,
64,
Renal Failure
Renal Failure
Renal Failure
6
5, 6,
64
5, 6,
64
Renal Failure
Renal Failure
6, 64
6
5, 6,
64
6, 64
6, 64
Renal Failure
6, 64
Liver Failure
Cardiomyopathy 5, 6,
64
6, 64
6, 64
6
6, 64
Cardiomyopathy 5, 6,
64
Cardiomyopathy 5,
6,64
Cardiomyopathy 5,
6,64
Cardiomyopathy 5, 6,
64
27
Figure 1.12. Clustering of single amino acid mutations in apo A-I.
position. The first step in understanding the fibril-forming properties of these and
eventually other mutants of apo A-I is to characterize their stabilities.
1.7 The Current Study
The aim of this study was to compare the stability of apo A-I L178H, which has
an “outside” mutation, to wild-type protein using equilibrium solvent denaturation.
Denaturation of the protein was measured through changes in the intrinsic fluorescence
of the protein’s four tryptophan residues. From intrinsic fluorescence measurements, the
Gibbs free energy of the protein (ΔG⁰), the effectiveness of unfolding (m), and the
midpoint of denaturation (D1/2) were obtained. These values tell us how stable L178H is
compared to WT protein. Understanding the difference in stability caused by the
mutation is key to understanding the mechanism of fibril formation in the protein.
28
Chapter 2
EXPERIMENTAL
2.1 Materials
Ethanol (95%, ACS/USP grade) was obtained from Pharmco Products, Inc.
Nanopure water was obtained by filtration through an EASYpure LF Compact
Ultrapure Water System (Barnstead). His-bind (nickel chelating) resin was purchased
from Novagen®. Ampicilin sodium salt (AMP), Triton-X-100 (electrophoresis grade),
guanidine hydrochloride, ethylenediaminetetraacetic acid (EDTA) disodium salt
(electrophoresis grade), sodium chloride, disodium hydrogen phosphate (Na2HPO4),
monosodium phosphate (NaH2PO4), Luria broth components (LB; 10 g/L tryptone, 5
g/L yeast extract, 10 g/L sodium chloride), sodium dodecyl sulfate (SDS), glacial acetic
acid, glycerol, Coomassie brilliant blue R-250, and methanol were purchased from
Fisher Scientific®. Protease inhibitor cocktail P2714 (inhibitory components: AEBSF,
Aprotinin, Bestatin, EDTA, E-64, Leupeptin) and P8849 (inhibitory components:
AEBSF, Bestatin, E-64, Pepstatin A, Phosphoramidon) were purchased from SigmaAldrich Chemical Company®. Isopropyl-β-D-thiogalactopyranoside (IPTG) analytical
grade was purchased from Affymetrix, Inc. Ultra pure urea and guanidine hydrochloride
(Gn-HCl) were purchased from MP Biomedical, LLC. Imidazole was purchased from
Janssen Chimica. Glycine was purchased from BDH Chemicals. Sodium dodecylsulfate
polyacrylamide gel electrophoresis (SDS-PAGE) gels were purchased from Jule, Inc.
Acetone was purchased from Pharmco-Aaper. NiSO4•6H2O and 2-mercaptoethanol
were purchased from Aldrich Chemical Co. Tris(hydroxymethyl)aminomethane
29
hydrochloride (Tris-HCl) was purchased from EMD Chemicals. Bromophenol blue was
purchased from Sigma Chemical Co. Tris base was purchased from J.T.Baker®.
2.2 Methods
2.2.1 Protein Production
2.2.1.1 Clones and Cell Lines
Wild type and mutant (L178H) apo A-I were donated from the laboratory of Dr.
John Voss (UC Davis, CA) as inserts in pNFXex plasmid in Escherichia coli (E. coli)
strain BL21(DE3) pLysS cells (Invitrogen) (65). The original pNFXex plasmid was
constructed from pBluescript KS(+) vector (Stratagene, La Jolla, CA) by Ryan and coworkers to encode the human apo A-I gene (66). The (pNFXex) plasmid, constructed by
Lagerstedt and co-workers, has increased ultility with the addition of 13 restriction
endonuclease sites and improved expression due to optimizing codon for use in E. coli,
compared to the original plasmid (65).
2.2.1.2 Induction of Expression and Cell Pellet Storage
Fifity ml LB supplemented with 50 μl of 500 mg/ml AMP in a 125 ml
Erlenmeyer flask was inoculated with E. coli containing the apo A-I plasmid either with
one colony from an agar plate or from scraping the surface of a frozen (-80˚C) glycerol
stock solution. The flask was then incubated in a New Brunswick Scientific Excella E24
Incubator Shaker Series at 37˚C with shaking (225 rmp) overnight (12-16 hours). Two 1
L flasks, each containing 500 ml LB broth and 500 μl of 50 mg/ml AMP, were
30
inoculated with 25 ml of the overnight culture until the OD at 600 nm reached 0.8 ±
0.05. IPTG was then added to a concentration of 0.5 mM, inducing expression of the
target protein. After 3-5 hours, the cells were pelleted by centrifugation in a SuperLite® SLA-1500 rotor in a Sorvall® RC 5B PLUS centrifuge at 12,500 rpm for 15
minutes at 4˚C. The cell pellets were quickly frozen in a bath of dry ice and acetone and
then stored at -80˚C.
2.2.1.3 Cell Lysis
Ten ml of lysis buffer (phosphate saline buffer (PBS; pH = 7.4, 1.5 M sodium
chloride, 0.2 M Na2HPO4, 0.2 M NaH2PO4 in nanopure water), 3 M Guanidine HCl,
0.1% Triton-X-100, 500 μl P2774 Sigma® protease inhibitor cocktail) was added to
each frozen cell pellet and mixed by first inverting and then incubating on an InnOva
2300 Platform Shaker (30 rpm) for 20 minutes at room temperature. The solution then
was kept in an ice bath and sonicated 3-5 times using a Fisher Sonic Dismembrator
Model 150 from Artek Systems Corporation at 45 % burst power for 30 seconds with a
30 second cooling period. The solution was centrifuged in a Super-Lite® SLA-1500
rotor in a Sorvall® RC 5B PLUS centrifuge at 12,500 rpm for 15 minutes at 4˚C. The
pellet was discarded and the supernatant was filtered through a syringe with a 0.45 µm
Millex® Syringe Driven Filter Unit filter.
2.2.1.4 Protein Purification
An econo chromatography column (Bio-Rad®) was loaded with 5 ml of Hisbind resin (binding capacity 5-10 mg protein/ml resin). The column was then washed
31
with 20 ml deionized water, charged with 25 ml charge buffer (400 mM NiSO4), and
equilibrated with 25 ml binding buffer (PBS containing 40 mM imidazole). The cell
extract was loaded onto the column with a flow rate of 0.5-1 ml/min. A 15 µl sample of
the flow through was collected for SDS-PAGE analysis for the presence of protein (See
section 2.2.1.5). The column was then washed with 70 ml binding buffer and the first
seven 1.5 ml fractions were collected for SDS-PAGE analysis for the presence of
protein. The target protein was then eluted with elution buffer (PBS containing 500 mM
imidazole), and the first twelve 1.5 ml fractions were collected for SDS-PAGE analysis
for the presence of protein. The column was then stripped with 30 ml strip buffer (0.5 M
NaCl, 100 mM EDTA, 20 mM Tris-HCl, pH 8.0), to be re-used in the next round of
protein purification.
2.2.1.5 Protein Analysis
Sodium dodecylsulphate polyacrylamide gel electrophoresis (SDS-PAGE) was
carried out with 15% Tris-Glycine gels (Jule, Inc.) to identify the protein-containing
fractions collected from the column. Gels were loaded with 15 μl of column extract
mixed with 5 μl of 4X sample buffer (240 mM Tris-HCl pH 6.8, 40% glycerol, 8%
SDS, 0.04% bromophenol blue, 5% 2-mercaptoethanol), and then run at 4ºC for 1.5
hours at 120 V in running buffer (25 mM Tris base, 192 mM glycine, 0.1% SDS, pH
8.3). Gels were then stained with Coomassie blue (0.1% w/v Coomassie brilliant blue
R-250, 10% glacial acetic acid, 45% methanol) for at least one hour, then de-stained
overnight in 10% glacial acetic acid, 10% methanol. The fractions containing protein
32
were pooled and dialyzed in 1 L PBS for at least 9 hours, with a change of 1 liter of
fresh PBS every 3 hours, to remove imidazole.
2.2.2 Determination of Protein Concentration
The concentration of the pooled protein fractions were measured
spectrophotometrically with an Ocean Optics, Inc. Chem 2000-UV-VIS
spectrophotometer. Protein concentration was calculated from the OD at 280 nm using ε
= 1.13 (cm2/mg).
2.2.3 Equilibrium Solvent Denaturation
Apo A-I protein was diluted in PBS to a final concentration of 0.10 mg/ml and
incubated at either 4˚C or 37˚C without stirring for 0 to 14 days. For 4˚C samples,
protease inhibitor (P8849) was added at a 1:10,000 (v/v) ratio at day 3 of incubation.
For 37˚C samples, P8849 was added at day 3 and again at day 10, both at a 1:10,000
(v/v) ratio, to prevent protein degradation. Aliquots of protein were removed
periodically and combined with a denaturant (urea or Gn-HCl) in equilibrium
denaturation experiments. Stock solutions of 8 M urea and Gn-HCl were prepared from
their respective ultra pure solid crystals. Samples containing 0.050 mg/ml ApoA-I and
varying concentrations of urea or Gn-HCl (0.0-4.0 M) were prepared. The final volume
of each sample was 2.0 ml. The samples were incubated with denaturant for 24 hrs at
4°C prior to fluorescence measurements.
2.2.4. Intrinsic Fluorescence
33
Fluorescence measurements were conducted with a Shimadzu RF-5301
spectrofluorophotometer equipped with thermostat-controlled cell holders set to
maintain the temperature constant at 25˚C using a circulating water bath. The
experiments were performed using a 1 cm cell. The excitation wavelength for
tryptophan was set at 295 nm in order to exclude the contribution of tyrosine residues
(max absorbance at 280 nm) to the overall fluorescence emission. Emitted light was
measured at 310-400 nm with a 5 nm excitation and 3 nm emission slit width, and
corrected for background signal. The unfolding transition was monitored by recording
the wavelength at maximum fluorescence intensity (WMF). When tryptophan is
excited, the emission spectrum produces a curve with a WMF between 308-355 nm
depending on the degree of solvent polarity around the tryptophan (67). During
unfolding, the WMF increases and the fluorescence intensity decreases as the solvent
polarity around the tryptophan increases (68).
2.2.5. Data Analysis
Calculating D1/2, ΔGDº, and m
Based on the unfolding of the protein monitored through the WMF, the stability
of the protein can be calculated. A background scan of PBS was subtracted from each
sample scan. The WMF of triplicate dilutions of each sample were averaged and plotted
against the denaturant concentration. The resulting curves exhibited characteristics of a
sigmoidal function and were fitted with the Hill equation (69),
𝑦 = 𝑦0 +
𝑎𝑥 𝑏
𝑐 𝑏+ 𝑥𝑏
[1]
34
where y is the WMF, y0 the minimum WMF (when the protein is folded), a is the
difference between the minimum WMF and maximum WMF (when the protein is
completely unfolded), b is the Hill coefficient, x is the denaturant concentration, and c is
the midpoint of denaturation (D1/2). The midpoint occurs at a denaturant concentration
where half of the protein population is folded and half is unfolded, which is determined
by the spectroscopic change. Assuming a two-state model (folded and unfolded), the
fraction of unfolded protein (fD) can be calculated using:
𝑓𝐷 =
𝑦− 𝑦0
𝑦𝑚𝑎𝑥 − 𝑦0
[2]
where ymax is the maximum WMF. The equilibrium constant (KD) for the process can be
calculated as:
𝐾𝐷 =
𝑓𝐷
1−𝑓𝐷
[3]
and the free energy of denaturation (ΔGD) can be calculated as
∆GD = -RTlnKD
[4]
Plotting ΔG versus denaturant concentration results a linear plot with the equation:
∆GD = ∆GD0 – m [denaturant]
[5]
where ΔGDº is the free energy of protein folding in water (0 M denaturant), and m the
effectiveness of denaturation. To obtain the D1/2, fD is set to 0.5 (in which half of the
protein population is unfolded), resulting in KD = 1 and ΔGD = 0. The linear equation
becomes 0 = ∆GD0 – m(D1/2). Thus,
35
𝐷1/2 =
°
∆𝐺𝐷
𝑚
[6]
The goodness of fit for the sigmoidal function was determined by the correlation
coefficient (R2) value.
2.2.6 Statistical Analysis
2.2.6.1 One Sample T-Test
One sample t-tests are used to compare a mean value to a population mean. The
variation in data (standard deviation) obtained in this project was due to instrument
model, cell line, protein batch, reagents, and pipetting techniques, and one could assume
the same sources of variation in other labs. In comparing this project’s data to a
literature value, the standard deviations of both data are comparable using an unpaired ttest. However, this project’s data is compared to multiple literature values (five to six
literature sources); and this project’s standard deviation is not comparable to the 5-6
standard deviations of the combined literature sources. Thus, the one sample t-test
compares this project’s data to the literature data regardless of each literature data’s
standard deviation. A one sample t-test uses the equation:
𝑡=
|𝑥− 𝜇|
𝑠/√𝑛
[7]
where x is the mean of the literature value, μ is the experimental mean value (from this
study), s is the standard deviation of the literature values from their mean, and n is the
number of literature values. The calculated t value is then used to obtain the p-value.
36
The data is tested at the 95% confidence level in which p > 0.05 indicates there is no
difference between the two means, and p < 0.05 indicates the two means are different.
2.2.6.2 Unpaired T-Test
Unpaired t-tests are used to compare two population means. Here, the unpaired
t-test was used to compare the mean thermodynamic value between the two types of
proteins (WT and L178H) in the same conditions. Due to the small sample sizes the
variance may differ between each set of data, but if the sample size were to be infinite
the true variance is assumed to be equal, due to the fact that the data were all obtained
in the same way. Thus, this test assumes the two sets of data have equal variances. The
test uses the equation:
𝑡=
|𝑥̅ 1 −𝑥̅2 |
𝑠𝑝𝑜𝑜𝑙𝑒𝑑
𝑛1 𝑛2
[8]
√𝑛
1 +𝑛2
where 𝑠𝑝𝑜𝑜𝑙𝑒𝑑 = √
𝑠12 (𝑛1 −1)+𝑠22 (𝑛2 −1)
𝑛1 +𝑛2 −2
[9]
s is the standard deviation, 𝑥 is the mean, and n is the number of samples.
The calculated t value is then used to obtain the p-value. The data is tested at the 95%
confidence level, in which p > 0.05 indicates there is no difference between the two
means, and p < 0.05 indicates the two means are different.
2.2.6.3 Propagation of Error
37
Propagation of error is used to find the uncertainty associated with a function
based on the uncertainty associated with the individual values. When comparing the
average thermodynamic values of L178H to the average values of WT (meta-average),
propagation of error is needed to find the uncertainty (standard deviation) of the metaaverage. To statistically compare the reduction in stability of L178H in urea to that in
Gn-HCl at both 4⁰C and 37⁰C using an unpaired t-test, the standard deviation must be
calculated for the reduced value. The equation for propagation of error was used to
calculate the standard deviation.
For a multiplying or dividing function (e.g. 𝑞 = 𝑥1 𝑥2 or 𝑞 = 𝑥1 ⁄𝑥2 ), the
resulting standard deviation of q (stotal) can be calculated as:
𝑠𝑡𝑜𝑡𝑎𝑙 =
𝑥1
𝑥2
2
2
√(s1 ) + (s2 )
x
x
1
2
where x is the mean and s is the standard deviation.
[10]
38
Chapter 3
RESULTS AND DISCUSSION
3.1 Protein Purification
The stability of L178H and WT apo A-I was determined through equilibrium
solvent denaturation monitored by intrinsic fluorescence at two temperatures (4⁰C and
37⁰C) and in two denaturants (urea and Gn-HCl) over a 14 day period. This requires a
large amount of protein (3.5-4.0 mg) for each experiment, so a good protein yield is
important.
In order to obtain protein, E. coli cells containing the apo A-I gene (WT or
L178H in the pNFXex plasmid) (66) were first inoculated overnight in 50 ml of LB
media. The next day, the cells were transferred to a new flask containing 500 ml of
media and were induced to express the proteins. After induction, the cells were lysed,
the protein was purified using nickel column chromatography, and the protein purity
was confirmed using SDS-PAGE. The day the protein was extracted from the cell is
considered day zero. The protein was then dialyzed for at least nine hours to eliminate
imidazole salts. Finally, the protein’s concentration was determined and used in
denaturation experiments beginning at day 1. The protein yield on average was five
milligrams per 500 ml of culture, enough for one denaturation experiment (one
expression of one protein at one temperature with one denaturant run as triplicate
samples). In order to complete all the experiments, protein production was performed
numerous times to obtain the required amount of protein. As the experiments were run
39
over a 14 day period, SDS-PAGE was used to periodically check for degradation
because apo A-I is highly sensitive to proteolytic degradation (70); only intact protein
was used in denaturation experiments. SDS-PAGE analysis confirmed both WT and
L178H protein remained intact during the 14 days incubation at 4⁰C, as the band
corresponding to intact protein persists through day 14 (Figure 3.1A). At 37ºC, both
proteins were degraded by day 7, as the intact apo A-I band for day 7 is non-existent
(Figure 3.1B). To prevent degradation, protease inhibitor cocktail (P8849 Sigma®) was
added to the 37⁰C incubated protein at a 1:10,000 v/v dilution every seven days. With
the addition of protease inhibitors, the proteins remained intact during the 14 days of
incubation (Figure 3.1C).
Figure 3.1. SDS-PAGE analysis of the integrity of WT and L178H protein at A) 4⁰C,
B) 37⁰C, C) 37⁰C with protease inhibitor cocktail P8849. D1, D7, D14 corresponds to
day 1, 7, and 14 respectively. The expected molecular weight of apo A-I is 28.1 kDa.
40
3.2 Equilibrium Solvent Denaturation
Mechanism of Solvent Denaturation
Equilibrium solvent denaturation is the unfolding of protein induced by
increasing the denaturant (urea or guanidine hydrochloride) concentration. Urea is an
uncharged molecule, whereas guanidine (a hydrochloride salt) is charged (Figure 3.2).
The mechanism of denaturation for both denaturants is unclear, but research suggests
that denaturation may be due to direct binding of denaturant to the protein, or to the
ability of denaturants to change the properties of the water solvent (71), or to both.
Urea has a carbonyl group and guanidine hydrochloride (Gn-HCl) has a charged
guanidinium group that can interact with the protein backbone or side chains. The
polarity of urea and Gn-HCl is different from that of the water solvent, which means
their presence could change the bulk solvent properties in ways that promote protein
unfolding.
Figure 3.2. Structures of urea and guanidine hydrochloride.
Because urea and Gn-HCl differ in their charges, they have different denaturing
effects and yield different thermodynamic values (72). This was demonstrated by
examination of the stability of a series of 4-helix coiled-coil analogs in which residues
41
involved in polar or electrostatic helix-helix interactions were systematically varied
from attractive to repulsive charges. The residues contributing to nonpolar helix-helix
interactions were the same in all four analogs. The stability of the four analogs was
determined and yielded different thermodynamic values for each protein in urea but
nearly identical values in Gn-HCl. This indicates that urea denaturation measures the
contribution of both hydrophobic and electrostatic interactions that stabilize the protein,
whereas Gn-HCl denaturation only measures the contribution of hydrophobic
interaction (72). This is due to the fact that Gn-HCl is an ionic compound that masks
electrostatic interactions. Thus, comparing proteins using two denaturants allows for a
relative determination of stabilizing forces, electrostatic versus hydrophobic, between
the proteins.
To examine the stability of WT and L178H apoA-I, equilibrium solvent
denaturation was conducted on both proteins using Gn-HCl and urea. Protein was
diluted from a stock solution to 0.10 mg/ml and incubated at 4⁰C or 37⁰C for 24 hours
to ensure the protein is in its monomeric form. The protein was then combined with
denaturants (urea or Gn-HCl) in triplicate samples containing 0.05 mg/ml protein in
varied concentrations of denaturants (0.0-4.0 M). The samples were incubated at 4⁰C
for another 24 hours to allow the protein and denaturant mixture to reach equilibrium.
Finally, the samples were measured using intrinsic fluorescence.
42
3.3 Intrinsic Fluorescence Measurements
When tryptophan is excited, the emission spectrum produces a curve with a
wavelength at maximum fluorescence intensity (WMF) between 308-355 nm depending
upon the degree of solvent polarity around the tryptophan (67). During unfolding, the
WMF increases and the fluorescence intensity decreases as the solvent polarity around
the tryptophan increases (68). Typically, a WMF around 335 nm is characteristic of
tryptophan residues buried inside the hydrophobic core of the protein, while a WMF of
around 350 nm is characteristic of tryptophan residues exposed to aqueous solvent (67).
Emission spectra typical for lipid-free WT apo A-I are shown in figure 3.3. Apo A-I
contains four tryptophan residues (W8, 50, 72, 108) which are located in the N-terminal
helix bundle (residues 1-189). Fluorescence studies indicate that all four of the
tryptophans in lipid-free apo A-I exist in a non-polar environment and are protected
from solvent (73). When the folded apo A-I is excited at 295 nm the buried tryptophan
gives an emission spectrum with a wavelength at maximum fluorescence intensity
(WMF) of around 338 nm (Curve A). As apo A-I unfolds due to denaturant, the
tryptophan residues become exposed to solvent, and the solvent polarity around the
tryptophan increases. This leads to an increase in WMF due to solvent relaxation of the
excited fluorophore and a decrease in the fluorescence intensity due to increased
quenching by the solvent. The WMF for fully unfolded apo A-I is approximately 352
nm (Curve D).
43
Figure 3.3. Emission spectra of WT apo A-I in varying denaturant concentrations. A) 0
M Gn-HCl, B) 1.5 M, C) 2.0 M, and D) 3.0 M.
To determine stability of the protein, the WMF obtained from intrinsic
fluorescence was plotted against denaturant concentration. This generated sigmoidal
plots for both proteins (Figure 3.4A). The curves were then fitted using the Hill
equation as described in Chapter 2. The sigmoidal plot shows the unfolding of apo A-I
is a two state mechanism, transitioning from a native (N) state to denatured (D) state,
without a substantially populated intermediate state. The cooperativity of the protein
unfolding is dependent on the steepness of the transition from the N to D state.
44
Figure 3.4. A) Denaturation curve for WT apo A-I monitored by intrinsic fluorescence.
B) Determination of ∆GH2O of WT apo A-I by linear extrapolation; ∆G is plotted as a
function of denaturant concentration. Each point on both plots is an average of three
samples from the same batch of protein.
Parameters used to determine the stability of the protein are the midpoint of
denaturation (D1/2), Gibbs free energy (∆G⁰) of unfolding, and protein stability in
response to denaturant concentration (m). The m value has been shown to correlate with
the change of accessible surface area (∆ASA) of the protein going from the native (N)
to the denatured (D) state (74). The values were determined by plotting ∆G versus
denaturant concentration (Figure 3.4B). The resulting linear function was analyzed
using the equation ∆G = ∆G⁰ - m[denaturant] to obtain ∆G⁰, D1/2, and m.
3.4 Comparison of Experimental Data to Literature Values
In the course of obtaining the stability data, there are many factors that could
influence the data such as instruments, cell line, protein batch, reagents, pipetting
technique, and fitting method. In order to prove the validity of our technique, data
obtained for wild-type (WT) protein in urea and guanidine hydrochloride for day 1 at
4⁰C was compared to literature values, also obtained at 4⁰C (Table 3.1). There is no
45
literature value for WT at 37⁰C and no literature value for L178H at either temperature.
The p-values from one-sample t-tests indicate that, statistically, there is no significant
difference between the project values and literature values for either urea or Gn-HCl.
This validates our technique and, by extension, the data obtained for WT and L178H
protein at both temperatures.
Table 3.1 Comparison of experimental values to literature values for WT protein.
Urea*
Gn-HCl**
D1/2
D1/2
∆G⁰
∆G⁰
WT
m
m
This Project
2.81
6.16
2.19
1.23
4.74
3.85
Literature 2.80±0.19 6.95±1.42 2.46±0.65 1.11±0.14 4.18±0.85 3.88±0.76
n=5
n=5
n=5
n=6
n=6
n=6
p-value
p>0.05
p>0.05
p>0.05
p>0.05
p>0.05
p>0.05
(Diff?)
(no)
(no)
(no)
(no)
(no)
(no)
* Literature value for urea is an average of data taken from ref. 75-79.
**Literature value for Gn-HCl is an average of data taken from ref.78-83.
Units for the thermodynamic parameters are as follow: D1/2 (M), ∆G⁰ (kcal•mol-1),m
(kcal•M-1mol-1)
3.5 Stability of Initial Structure (Day 1)
The stability of the initial structure (at day 1) of WT and L178H apo A-I was
determined by incubating the proteins at 4⁰C and 37⁰C in both urea and Gn-HCl for one
day. Protein incubated at 4⁰C was evaluated in order to compare the data to literature
values, whereas the 37⁰C temperature was used to reflect relevant physiological (body)
temperature. The data was collected from three batches of protein; each batch of protein
resulted in a sigmoidal plot, as discussed above. Thus, the thermodynamic parameters
are reported as an average from three batches of protein (three plots).
46
3.5.1 Stability at 4⁰C
The urea denaturation curve at 4⁰C indicates L178H has reduced stability
compared to WT, as shown by the shift of the unfolding curve towards lower urea
concentration (Figure 3.5A). The denaturation curve for L178H is also less steep than
that of WT indicating reduced cooperativity. The midpoint of denaturation (D1/2),
conformational stability (∆G⁰), and the dependence of ∆G value on denaturant
concentration (m) determined from linear plots of ∆G as a function of denaturant
concentration are listed in table 3.2. Compared to WT, the stability of L178H is reduced
by approximately 38%, 59%, and 33% corresponding to the D1/2, ∆G⁰, and m values,
respectively. Similar to the results for urea denaturation, the values obtained for Gn-HCl
denaturation indicate L178H is less stable and less cooperative than WT (Figure 3.5B).
Compared to WT, the stability of L178H is reduced by approximately 12%, 59%, and
54% corresponding to the D1/2, ∆G⁰, and m values, respectively (Table 3.2).
Figure 3.5. Urea induced unfolding of WT and L178H apo A-I at 4⁰C in A) urea and B)
Gn-HCl. The solid line represents WT and the dashed line represents L178H.
47
Table 3.2. Comparison of L178H to WT at 4⁰C
Urea
Gn-HCl
D1/2
D1/2
∆G⁰
∆G⁰
m
m
WT
2.81±0.05
6.16±0.19 2.19±0.11 1.23±0.02 4.74±0.38 3.85±0.34
L178H
1.74±0.03
2.55±0.07 1.46±0.07 1.08±0.07 1.93±0.14 1.76±0.06
Reduction of
38.2±1.1% 58.7±1.2% 33.3±3.1% 12.5±5.4% 59.2±3.0%, 54.2±1.5%
L178H vs. WT
p-value
p<0.05
p<0.05
p<0.05
p<0.05
p<0.05
p<0.05
(Diff?)
(yes)
(yes)
(yes)
(yes)
(yes)
(yes)
-1
Units for the thermodynamic parameters are as follow: D1/2 (M), ∆G⁰ (kcal•mol ),
m (kcal•M-1mol-1)
3.5.2 Stability at 37⁰C
Denaturation was then performed on protein incubated for one day at 37⁰C
because it is the relevant body temperature and the temperature at which apo A-I
normally functions. Similar to the results at 4⁰C, L178H incubated at 37⁰C has reduced
stability and cooperativity (Figure 3.6). Urea denaturation shows the stability of L178H
was reduced by approximately 37%, 59%, and 30% corresponding to the D1/2, ∆G⁰, and
m values, respectively, compared to WT (Table 3.3). Gn-HCl denaturation at 37⁰C
indicates L178H has reduced stability by approximately 25%, 59%, and 45%
corresponding to the D1/2, ∆G⁰, and m values, respectively (Table 3.3).
48
Figure 3.6. Denaturation of WT and L178H apo A-I after incubation at 37⁰C in A) urea
and B) Gn-HCl. The solid line represents WT and the dashed line represents L178H.
Table 3.3. Comparison of L178H to WT after incubation at 37⁰C
D1/2
Urea
∆G⁰
5.39±0.33
2.34±0.12
m
1.91±0.08
1.34±0.05
D1/2
Gn-HCl
∆G⁰
3.78±0.08
1.55±0.27
m
3.14±0.02
1.74±0.31
WT
2.82±0.07
1.20±0.03
L178H
1.78±0.12
0.90±0.02
Reduction of
L178H vs.
36.9±4.1% 55.8±2.3% 29.7±2.5% 25.5±1.3% 58.8±7.5%, 44.6±9.8%
WT
p-value
p<0.05
p<0.05
p<0.05
p<0.05
p<0.05
p<0.05
(Diff?)
(yes)
(yes)
(yes)
(yes)
(yes)
(yes)
Units for the thermodynamic parameters are as follow: D1/2 (M), ∆G⁰ (kcal•mol-1),
m (kcal•M-1mol-1)
The results indicate that L178H is significantly less stable than WT protein, as
indicated by the approximately 60% reduction in ∆G⁰ in both denaturants after one day
incubation at either 4⁰C or 37⁰C. The D1/2 values for L178H in guanidine and urea are
decreased at both temperatures compared to WT but the decrease is greater for urea than
for guanidine. Since urea measures both electrostatic and hydrophobic contributions
and Gn-HCl measures mainly hydrophobic contributions to stability (72), this indicates
49
L178H (relative to WT) has a larger reduction in electrostatic contributions than
hydrophobic contributions. In other words, L178H has a larger proportion of
hydrophobic interactions contributing to its stability. The m value of L178H is less than
that of WT in both denaturants at both temperatures, but the reduction in m value in
urea is less than that in Gn-HCl. The m value represents the dependence of ∆G⁰ on
denaturant concentration (m = ∆G⁰/D1/2; Chapter 2, section 2.2.5, eq. 6). In general, the
greater the m value, the more effective the denaturant because less denaturant is
required to achieve a given ∆G⁰. Since the reduction in m value of L178H compared to
WT is less in guanidine, this indicates guanidine is less effective than urea in denaturing
L178H compared to WT. This result is consistent with the D1/2 results, since the more
the protein is resistant to denaturation the less effective the denaturant. The m value is
also proportional to the change in solvent accessible surface area (∆ASA) of the protein
going from the native (N) to the denatured (D) state (74). The reduction in m values of
L178H compared to WT indicates the change in surface area going from the N to D
state for L178H is less than that of WT. WT is more stable and therefore its native state
may be more ordered, so going from the N to D state would result in a large ∆ASA and
thus a large m value. In contrast, L178H is less stable and therefore its native state may
be more disordered, so the ∆ASA between the N and D state is smaller. Hence, the
smaller m values for L178H. The more disordered N state of L178H is consistent with
results obtained using ANS binding. ANS binds to accessible hydrophobic surfaces in
proteins. The ANS binding of L178H is nearly 4 times that of WT indicating L178H
has increased exposed hydrophobic surface area (63).
50
The lower stability of L178H compared to WT apo A-I is consistent with fibrilforming proteins being generally less stable than their WT counterparts. Recent analysis
of the ∆(185-243) apo A-I crystal structure model by Gursky et al. reveals how the
L178H mutation may destabilize the protein (36). In the crystal structure, all the
amyloidogenic mutations in apo A-I are packed in the N-terminal helix bundle (Figure
3.7A; 44). The authors hypothesize that the mutations destabilize the bundle causing
increased solvent exposure and leading to fibril formation. The L178H mutation is
located in the hydrophobic cluster near the bottom of the helix bundle (Figure 3.7A).
The mutation replaces a non-polar alkyl residue (L) with a polar imidazole ring (H),
therefore disrupting the hydrophobic cluster and destabilizing the bundle. The
disruption likely also changes salt bridge interactions involved in stabilizing the helix
bundle. This could explain the reduced electrostatic contributions as indicated by the
larger reduction of D1/2 in urea versus Gn-HCl for L178H at both temperatures,
compared to WT. Since the crystal structure is that of a truncated apo A-I and not fulllength apo A-I, the position of L178H is also examined in the EPR model of full-length
apo A-I. The EPR model of lipid-free apo A-I (37) also places position 178 in a helix
bundle (Figure 3.7B) which would likely be destabilized by the L178H mutation. Thus,
the lower stability of L178H can be explained by either structural model as being due to
destabilization of the helix bundle.
51
Figure 3.7. Location of the L178H and R173C mutations in A) ∆(185-243) lipid-free
apo A-I crystal structure and B) EPR model of lipid-free apo A-I.
Whether lowered stability leads to fibril formation may depend on more subtle
features of alterations caused by mutation. The apo A-I variant R173C is significantly
less stable than WT protein; its conformational stability (∆G⁰) is reduced by
approximately 50% in Gn-HCl and 70% in urea (Figure 3.8;78). Yet, the R173C
mutation does not result in fibril formation. The location of R173C in both the X-RC
and EPR models is also in the helix bundle, close to the location of L178H (Figure 3.7).
The mutation replaces a charged residue (R) with a non-polar residue (C), potentially
disrupting helix-helix interactions within the bundle (44). Therefore, since L178H and
R173C destabilize the helix bundle, leading to similar reductions in global stability
(Figure 3.8) while only L178H forms fibrils, it is unlikely that decreased stability per se
is the only factor in fibril formation. An additional factor may be the balance of
52
electrostatic versus hydrophobic interactions that stabilize the mutant protein. R173C is
less resistant to urea denaturation and slightly less resistant to guanidine denaturation
compared to L178H, indicating that, relative to each other, R173C stability is
dominated more by electrostatic forces while L178H stability is dominated more by
hydrophobic forces. This suggests that the ability to maintain salt bridge interactions, a
hallmark feature of helix-helix association in apolipoproteins (20,45), may help prevent
fibril formation.
Figure 3.8. Percent of reduction in thermodynamic value of L178H and R173C
compared to WT at 4⁰C in A) urea and B) Gn-HCl. The white bar represents L178H
and the gray bar represents R173C (78).
3.6 Stability Over 14 Day Period
The magnitude of the initial stability difference of WT and its fibril forming
mutants may be less important in fibril formation than the stability change over time. To
examine this parameter, the thermodynamic values for WT and L178H were obtained
on proteins incubated over a period of 14 days. One set of thermodynamic values for
day 1-14 was determined from one batch of protein and the entire experiment was
53
repeated twice (i.e. three batches of each protein used). During the course of the thesis
research, our research collaborators found that L178H incubated at 37⁰C converts to
helical fibrillar structures over time with a t1/2 of 12 days, while WT fails to form fibrils
(although small spherical aggregates were observed) (63). Since the distribution of
monomer, aggregates, and fibrillar species for L178H and WT in solution over time is
unknown, only the change in ∆G⁰ values over time were examined because ∆G⁰
measures the change in free energy of the whole system (all the protein species). The
contribution of hydrophobic and electrostatic interactions of the system is much more
complicated than that of a monomeric protein and will therefore not be discussed.
3.6.1 Stability changes at 4oC
To compare the stability change over time, the ∆G⁰ values of day 7 and 14 were
calculated as fractions of day 1. Both proteins experienced a decrease in stability when
incubated at 4oC (Figure 3.9). The decrease in stability at day 7 is similar for the two
proteins in both denaturants with the fraction of day 1 ∆G0 ranging from 0.79 to 0.84.
However, the stability of WT protein to guanidine denaturation at day 14 is much lower
than in urea, indicating a greater loss of hydrophobic interactions after two weeks.
L178H also exhibits a bigger drop in stability in the second week of incubation but the
change in the ∆G0 value is similar for the two denaturants for this protein (0.60 and 0.67
for urea and guanidine, respectively). These results indicate that both proteins undergo
structural changes when incubated at 4oC and that the change is greater in the second
week of incubation. The interactions involved in these changes are different for the two
54
proteins as WT experiences a greater loss of hydrophobic interactions than L178H
(fractional decrease to 0.44 for WT versus 0.60 for L178H). Thus, even though both
proteins are destabilized by incubation at 4oC, the resulting L178H structure is more
stabilized by hydrophobic interactions than the WT structure.
Figure 3.9. Change in free energy after incubation at 4⁰C. The ∆G⁰ values of day 7
and 14 were calculated as fractions of day 1 ∆G⁰ values. The standard deviations for the
fractions were calculated using propagation of error. The white bar represents urea and
the gray bar represents Gn-HCl.
3.6.2 Stability changes at 37oC
Both proteins exhibited an overall increased stability when incubated at 37oC,
but to different extents in the two denaturants (Figure 3.10). The WT stability to urea is
similar to that of guanidine at both day 7 and day 14, as shown by the error bar
(standard deviation), suggesting the increase in the stability of WT is due to an increase
in both electrostatic and hydrophobic interactions. The stability at day 7 is similar to
that of day 1 having a fraction of approximately 1, whereas at day 14 the stability
increases 130% (fraction = 1.3) indicating a large change in stability occurs after day 7.
55
In contrast, L178H exhibits only a minor increase in stability to urea at both Day 7 and
Day 14, whereas there is a large increase in stability of L178H to guanidine at Day 7
and again at Day 14. This indicates the increased stability of L178H is due to a
substantial gain in hydrophobic interactions and a minor gain in electrostatic
interactions.
Figure 3.10. Change in free energy after incubation at 37⁰C. The ∆G⁰ values of day 7
and 14 were calculated as fractions of day 1 ∆G⁰ values. The standard deviations for the
fractions were calculated using propagation of error. The white bar represents urea and
the gray bar represents Gn-HCl.
3.6.3 Structural Changes in L178H versus WT
It is clear from the denaturation data that while both proteins became less stable
over time when incubated at 4oC and more stable over time when incubated at 37oC, the
interactions involved in the stability changes are different for the two proteins. While at
4oC, both proteins experienced the greatest decrease in ∆G0 in guanidine at Day 14, the
decrease was larger for WT, implying that L178H remains more stabilized by
56
hydrophobic interactions. The same trend toward greater hydrophobic interactions in
L178H is given by the 37oC incubation data which shows L178H becomes much more
stable to guanidine denaturation than WT.
The basis for the decrease in stability at 4oC is unclear, but it may be due to cold
denaturation. At low temperatures, the Gibbs free energy of hydration of nonpolar
residues increases, leading to proteins unfolding and exposing nonpolar amino acids to
water (84). Although this normally occurs in proteins at temperatures much below
freezing, it may occur at 4oC in apolipoproteins due to their increased flexibility and
increased hydrophobic surface exposure compared to globular proteins.
The increase in stability at 37oC is most likely due to increased aggregation of
the protein, with L178H experiencing greater aggregation. An increase in aggregation
is supported by two types of evidence. First, as previously noted, L178H forms fibrils
after one month of incubation at 37oC whereas WT forms smaller, spherical aggregates
(63). Second, we have observed that light scattering in both WT and L178H is
increased upon incubation for two weeks at 37oC, but the increase is much higher in
L178H (Baal, Y., Nguyen, H., Roberts, L., unpublished observations). While the
stability of aggregate protein species versus the native protein is unknown in this
mixture, it has been shown that amyloid fibrils generally have higher stabilities than
their normally folded counterparts (85,86). Thus, the increased stability of WT over
time at 37⁰C is due to some form of aggregation. Presumably, the effect of the L178H
57
mutation is to both lower the stability of the protein and promote its rate of aggregation
over time through increased hydrophobic interactions.
58
Chapter 4
CONCLUSIONS AND FUTURE WORK
Research indicates that fibril-forming proteins are generally less stable than their
wild-type counterparts, and that less stable proteins are more prone to misfolding and
forming fibrils with increased stability (8-10). The goal of this thesis was to determine
whether stability is a factor in fibril formation in apo A-I by comparing the stability of
the fibrillogenic apo A-I mutant L178H to wild-type protein using equilibrium solvent
denaturation. The stability data obtained in this work indicates L178H is significantly
less stable than WT protein and that its stability is dominated more by hydrophobic than
electrostatic interactions. Furthermore, the greater influence of hydrophobic
interactions in L178H compared to WT persists through two weeks of incubation at
either 4oC or 37oC. The increased stability of both proteins upon incubation at 37⁰C is
most likely due to a greater tendency to form aggregated structure, with L178H
experiencing more aggregation than WT protein.
The formation of fibrils or other aggregated structures does not depend solely on
conformational destabilization, as other apo A-I mutants, most notably R173C, have
lower stability without forming fibrils (78). There are two clusters of single site
mutations in apo A-I that lead to tissue-specific fibrillar deposits (6). L178H, from the
173-178 cluster, is the first full-length fibril-forming apoA-I mutant to be characterized
in terms of its stability. Therefore, it is not yet known whether mutations from both
clusters lead to lowered protein stability. Furthermore, the relationship between
59
stability and direction of fibril formation has yet to be established. The G26R protein
forms amyloid (β-sheet) fibrils (7) whereas the L178H forms helical fibrils (63). Thus,
even if it is found that G26R and other amyloidogenic apoA-I mutants have lowered
stability, additional factors must dictate the type of fibril that forms. To understand the
relationships between stability, fibril type, and mutation position, it will be necessary to
characterize both the structures and stability of more mutants from both clusters.
Fibril formation further depends on the protein's change in hydrophobic versus
electrostatic interactions over time. There is almost no research at present on timedependent changes in stability and structure of apoA-I mutants, either amyloidogenic or
non-amyloidogenic. To fully understand the development of fibrillar protein in disease
will require these kinds of analyses for both types of mutants.
60
Appendix A
List of Abbreviations
∆ASA: change of accessible surface area (∆ASA) of the protein going from the native
(N) to the denatured (D) state
∆G: free energy of denaturation (protein unfolding)
∆G°: Gibbs free energy of protein. It represents the conformational stability of the
protein, and is the energy required to unfold the protein in water (0 M denaturant).
ABCA1: ATP-binding cassette transporter A1
Amino acid abbreviations:
Symbol
3-Letter 1-Letter
Alanine
Ala
A
Arginine
Arg
R
Asparagine
Asn
N
Aspartic acid
Asp
D
Cysteine
Cys
C
Glutamic Acid
Glu
E
Glutamine
Gln
Q
Glycine
Gly
G
Histidine
His
H
Isoleucine
Ile
I
Name
Symbol
3-Letter 1-Letter
Leucine
Leu
L
Lysine
Lys
K
Methionine
Met
M
Phenylalanine
Phe
F
Proline
Pro
P
Serine
Ser
S
Threonine
Thr
T
Tryptophan
Trp
W
Tyrosine
Tyr
Y
Valine
Val
V
Name
ANS: 8-Anilinonaphthalene-1-sulfonate, ANS binds to accessible hydrophobic surfaces
in proteins
Apo A-I: apolipoprotein A-I
61
ATP: Adenosine-5'-triphosphate
CD: circular dichroism
CETP: Cholesteryl ester transport protein
CHD: coronary heart disease
CVD: cardiovascular disease
D1/2: the midpoint of denaturation. It represent the stability against denaturant and is
the concentration in which half the population of protein is unfolded.
DMPC: dimyristoylphosphatidylcholine
Electrostatic interaction: the interaction of charged amino acids along a helix. For
example, a positively charge amino acid nearing a negatively charge amino acid results
in electrostatic attraction and amino acids with the same charge (negative or positive)
nearing each other result in electrostatic repulsion.
EPR: electron paramagnetic resonance
Equilibrium solvent denaturation: the unfolding of protein induced by increasing the
denaturant (urea or guanidine hydrochloride) concentration.
FRET: Förster Resonance Energy Transfer
FTIR: Fourier transform infrared spectroscopy
62
G26R: a mutation at position 26 (in apo A-I), in which a glycine (G) residue mutates to
an arginine (R)
Gn-HCl: guanidine hydrochloride
HDL: high density lipoprotein
HX-MS: hydrogen exchange and mass spectrometry
Hydrophobic interaction: the contribution of hydrophobic interaction depends on the
number of buried hydrophobic amino acid side chain in the folded protein (87). These
hydrophobic side chains are shielded from solvent. The more shielded hydrophobic side
chains in a protein generally results in greater stability.
L178H: a mutation at position 178 (in apo A-I), in which a leucine (L) residue mutates
to a histidine (H)
LB: luria broth
LCAT: lecithin-cholesterol acyltransferase
LDL: low density lipoprotein
m: effectiveness of denaturation
MIM: mixture of indirect method, which includes circular dichroism (CD),
sedimentation velocity, ANS binding, fluorescence quenching and limited proteolysis of
full-length and truncated protein (35)
63
NMR: nuclear magnetic resonance
PBS: phosphate buffered saline
PLTP: phospholipid transfer protein
POPC: palmitoyloleoylphosphatidylcholine
R173C: a mutation at position 173 (in apo A-I), in which an arginine (R) residue
mutates to a cysteine (C)
RCT: reverse cholesterol transport
rHDL: reconstituted HDL
SDS: sodium dodecyl sulfate
SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis
SR-BI: scavenger receptor class BI
WMF: wavelength at maximum fluorescence intensity
WT: Wild-type
X-RC: X-ray crystallography
64
Appendix B
Experimental Data
For all the data listed, units for the thermodynamic parameters are as follow:
D1/2 (M), ∆G⁰ (kcal•mol-1), m (kcal•M-1mol-1).
Initial stability data (day 1) for WT and L178H.
Urea
D1/2
∆G⁰
Gn-HCl
m
D1/2
∆G⁰
m
WT 4°C
2.81±0.05
6.16±0.19
2.19±0.11
1.23±0.02
4.74±0.38
3.85±0.34
L178H 4°C
1.08±0.07
1.93±0.14
1.76±0.06
1.74±0.03
2.55±0.07
1.46±0.07
WT 37°C
2.82±0.07
5.39±0.33
1.91±0.08
1.20± 0.03
3.78 ±0.08 3.14± 0.02
L178H 37°C 1.78±0.12
2.34±0.12
1.34±0.05
0.90±0.02
1.55±0.27
1.74±0.31
* All thermodynamic values are an average of values from three batches of protein (n=3); except for
L178H at 37°C in urea, which the values are an average of four batches of protein (n=4)
Time course data at 4°C
Urea
Gn-HCl
D1/2
D1/2
Day
∆G⁰
∆G⁰
m
m
1
2.76±0.04 6.46±0.72
2.34±0.23
1.21±0.02
4.95±0.15 4.11±0.16
7
WT
2.82±0.06 5.09±0.20
1.81±0.11
1.18±0.03
4.14±0.12 3.51±0.15
14
2.68±0.02 4.80±0.12
1.79±0.03
1.13±0.03
2.20±0.10 1.94±0.13
1
1.70±0.04 2.63±0.10
1.55±0.05
1.02±0.05
1.85±0.09 1.83±0.06
7
L178H
1.57±0.15 2.18±0.25
1.39±0.07
0.96±0.03
1.56±0.08 1.63±0.08
14
1.51±0.04 1.75±0.13
1.17±0.12
0.87±0.04
1.11±0.08 1.27±0.04
*All thermodynamic values are an average of three values (n=3) from triplicate samples. The data
were obtained from the same batch of protein for each conditions (e.g. for WT in urea, the same batch
of protein is used for day 1, 7, and 14 ).
65
Time course data at 37°C
Urea
Gn-HCl
D1/2
D
Day
∆G⁰
m
∆G⁰
m
1/2
1
2.86±0.08 5.52±0.73 1.92±0.20
1.22±0.03 3.85±0.19 3.16±0.09
7
WT
2.81±0.08 5.39±0.55 1.93±0.25
1.30±0.09 4.27±0.55 3.30±0.52
14
2.56±0.08 7.38±1.36 2.88±0.46
1.22±0.02 5.09±0.23 4.17±0.13
1
1.85±0.01 2.49±0.21 1.34±0.11
0.90±0.11 1.34±0.27 1.48±0.17
7
L178H
1.81±0.17 2.58±0.36 1.44±0.29
0.82±0.06 1.75±0.12 2.15±0.14
14
1.63±0.10 2.74±0.27 1.68±0.07
0.77±0.05 2.13±0.13 2.76±0.22
*All thermodynamic values are an average of three values (n=3) from triplicate samples. The data
were obtained from the same batch of protein for each conditions (e.g. for WT in urea, the same batch
of protein is used for day 1, 7, and 14 ).
66
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