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Before they were amyloid: understanding the toxicity of disease-associated monomers and oligomers prior to their aggregation

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Before they were amyloid: understanding the toxicity of disease-associated monomers and oligomers prior to their aggregation

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Content BEFORE THEY WERE AMYLOID

UNDERSTANDING THE TOXICITY OF DISEASE-ASSOCIATED MONOMERS AND
OLIGOMERS PRIOR TO THEIR AGGREGATION

by

Natalie C. Kegulian

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETIC, MOLECULAR AND CELLULAR BIOLOGY)

August 2015
II

Acknowledgments
I would like to thank my kind and supportive mentor, Dr. Ralf Langen, for
believing in me and opening my world to the scientific and professional opportunities
that have informed my Ph.D. experience. I am also grateful to all my committee
members—Dr. Tobias Ulmer, Dr. Robert Farley, Dr. Jeannie Chen, and Dr. Ian
Haworth—for mentoring me as inspiringly as they have.
A big thank you to my fellow Langen lab members past and present! Your
spirited hard work is only surpassed by the precious friendships you have offered me.
Dr. J. Mario Isas, Alan Okada, Dr. Nitin Kumar Pandey, Kazuki Teranishi, Dr. Mark
Ambroso, Dr. Jobin Varkey, Dr. Balachandra Hegde, and Prabhavati Hegde: your
helpful feedback throughout our conversations and meetings as well as the practical
tools in lab that you have given me to succeed, all of which you gave with heart and
tireless energy, have been invaluable. Sean Chung, Fleur Lobo, and Meixin Tao: thank
you for being the vibrant new lifeblood of this beautiful laboratory and for teaching me
how to teach. Thank you to former Langen lab members Dr. Sajith Jayasinghe, Dr.
Melania Apostolidou, and Dr. Christine Jao, alongside whom I have not had the
pleasurable opportunity to work but who laid the foundations supporting my work.
Also major thanks to the members—and PIs!—of the Siemer and Ulmer labs for being
delightful and helpful coworkers and friends.
I would like to thank all our collaborators who have been instrumental to my
projects. I am very appreciative of all our colleagues across the pond at Istituto Ricerche
di Biologia Moleculaire Promidis Srl, including fearless leaders Dr. Andrea Caricasole
and Dr. Andreas Weiss. Grateful acknowledgment to Dr. Peter Butler at the University
of California, Los Angeles, for useful in vivo data and informative conversations. Thank
you to the Malmstadt lab here at USC, particularly to Shalene Sankhagowit for the
fascinating GUV videos and a cordial friendship.
III

I would like to extend my gratitude to our funding sources—the National
Institutes of Health (NIH), the Hereditary Disease Foundation (HDF), the Huntington’s
Disease Society of America (HDSA), and CHDI. In particular, I would like to
acknowledge the NIH Cellular, Biochemical, and Molecular (CBM) Training Program
for awarding me a predoctoral grant and thus supporting me for two of my Ph.D. years
as well as providing me with enriching opportunities for presenting my work, making
new contacts, and learning about their research. The director of this program, Dr.
Michael Stallcup, receives my deep thanks for his generosity as another mentor figure.
On that note, while all the professors I have encountered throughout this
extensive graduate program have made helpful contributions to my education, I would
particularly like to single out and thank Dr. Ite Laird-Offringa, Dr. Deborah Johnson,
Dr. Wei Li, Dr. Young-Kwon Hong, and Dr. Woojin An for their support and their
invaluable teachings.
Most of all, I send heartfelt thanks to my mother and my father, the two most
loving and generous cheerleaders that any human being could ever dare to hope for.
Mom and Dad, I could not have done any of this without your passionate, affectionate,
often creative and always steady support.

IV

Table of Contents

Acknowledgments

II
List of figures and tables

VI
Abstract

X
Chapter 1. Amyloid Proteins and Structural Studies 1
1.1 Proteins and amyloid formation: an introduction 2
1.1.1 α-Synuclein 3
1.1.2 IAPP 5
1.1.3 Huntingtin 5
1.2 Structural studies 7
1.2.1 α-Synuclein membrane-bound structure 9
1.2.2 IAPP membrane-bound structure 10
1.2.3 EPR with SDSL: a brief overview 11
1.3 Exertions by membrane-active proteins 12
1.4 Loss of membrane integrity in conjunction with membrane
remodeling
15
1.5 Huntingtin non-amyloid conformational behavior changes with Q-
length
19
1.6 Closing thoughts 22
1.7 References

23
Chapter 2. Measuring Exposure of Different α-Synuclein Residues to
Phospholipid Membranes Using a Fluorescent Probe
36
2.1 Introduction 37
2.2 Methods 38
2.3 Results and Discussion 40
2.4 References

44
Chapter 3. IAPP as a Membrane Curvature Inducer and Sensor 45
3.1 Abstract 46
3.2 Introduction 47
3.3 Methods 49
3.4 Results 54
3.5 Discussion 64
3.6 References

68
V

Chapter 4. Circular Dichroism Uncovers Temperature- and
Polyglutamine-Dependent Structural Changes in Huntingtin Exon 1
74
4.1 Introduction 75
4.2 Methods 77
4.3 Results 79
4.4 Discussion 85
4.5 References

89
Chapter 5. Locating Enhanced Structural Ordering in Polyglutamine-
Expanded Huntingtin Exon 1 by Electron Paramagnetic Resonance
91
5.1 Introduction 92
5.2 Methods 92
5.3 Results 94
5.4 Discussion 100
5.5 References

102
Chapter 6. Conclusions: On the Non-Amyloid Structures and Behaviors
of Amyloid-Forming Proteins
103
6.1 Membrane curvature induction: a possible common link to
amyloid toxicity (and beyond)
104
6.2 Jekyll-and-Hyde–like conformational switching in HDx1 109
6.3 References

112

VI

List of Figures and Tables

Fig. 1-1 Amino acid sequences of the three proteins of interest in this work 3
Fig. 1-2 Structural models of the α-synuclein helical structure 8
Fig. 1-3 Structural models of the hIAPP helical structure 9
Fig. 1-4 Spin label attaches to protein sulfhydryl groups and gives an EPR
signal that varies with mobility

11
Fig. 1-5 Schematics showing two major mechanisms by which proteins
induce membrane curvature

13
Fig. 1-6 Leakage of vesicles composed of negatively charged phospholipids
and loaded with the fluorophore 8-aminonaphthalene-1,3,6-
trisulfonic acid (ANTS) and its quencher p-xylene-bis(pyridinium
bromide) (DPX) in presence of α-synuclein and amphiphysin

15
Fig. 1-7 Leakage of POPS/POPC (~2:1 molar ratio) vesicles in the presence
of IAPP

16
Fig. 1-8 Clearance of negatively charged MLVs by α-synuclein 17
Fig. 1-9 α-Synuclein induces the formation of highly curved structures
from POPG membranes

18
Fig. 1-10 Expanded-polyQ HDx1 loses its ability to bring its N- and C-
termini in proximity

20
Fig. 1-11 Temperature-induced change in FRET efficiency is greater for
N548Htt-Q16 than for N548Htt-Q55

21
Fig. 2-1 Molecular structure of the environment-sensitive fluorophore
IANBD

37
Fig. 2-2 The four phospholipids utilized in this membrane study 38
VII

Fig. 2-3 Fluorescence spectroscopy indicates the formation of a continuous
α-synuclein helix bound to membranes

41
Fig. 2-4 Blue or red shifts in α-synuclein-IANBD fluorescence in the
presence of vesicles of the indicated lipid compositions

42
Fig. 3-1 Light scattering of MLVs 56
Table 3-1 Half-times to maximal ThT fluorescence for each protein-lipid
combination in which an increase was detected

57
Fig. 3-2 Dissolution of POPS membranes by IAPP visualized directly under
fluorescence microscopy

58
Fig. 3-3 Negative stain EM of the interaction of hIAPP or rIAPP with large
vesicles

59
Fig. 3-4 IAPP-induced vesicle leakage 60
Table 3-2 Time points at which light scattering reached its minimum (i.e., the
sample reached maximal clearance) and at which scattering began
to increase due to protein aggregation

61
Fig. 3-5 Membrane curvature sensitivity of rIAPP 62
Fig. 3-6 IAPP binding to mitochondria and vesicular structures in vivo 63
Fig. 3-7 Membrane curvature-inducing and -sensing behavior in α-helical
IAPP

66
Fig. 4-1 Typical CD spectra designating the indicated protein secondary
structures

75
Fig. 4-2 Temperature-dependent CD spectra for HDx1 fusion proteins of
four different Q-lengths

80
Fig. 4-3 CD difference spectra between temperature extremes 81
Fig. 4-4 Temperature-dependent CD spectra for HDx1 fusion proteins of
four different Q-lengths in salt-free phosphate buffer
82
VIII

Fig. 4-5 CD controls showing temperature effect is not concentration- or
THRX-dependent

83
Fig. 4-6 MRE values at 222 nm at each temperature tested for THRX-HDx1
of the indicated Q-lengths, with weighted THRX MRE values
subtracted

84
Fig. 4-7 Temperature-dependent α-helical variations are also observed in
untagged, larger N548Htt purified recombinant proteins

84
Fig. 4-8 Effect of the S16D phosphomimetic mutation on MRE values at 222
nm at each temperature tested for THRX-HDx1 of the indicated Q-
lengths, with weighted THRX MRE values subtracted

85
Fig. 4-9 Effect of the S13D phosphomimetic mutation on MRE values at 222
nm at each temperature tested for THRX-HDx1 of the indicated Q-
lengths, with weighted THRX MRE values subtracted

86
Fig. 4-10 Effect of the S13D/S16D double phosphomimetic mutation on MRE
values at 222 nm at each temperature tested for THRX-HDx1 of the
indicated Q-lengths, with weighted THRX MRE values subtracted

87
Fig. 5-1 Sample temperature-dependent EPR spectra for THRX-HDx1-Q46
spin labeled at the indicated sites on HDx1

93
Fig. 5-2 EPR spectra measured at the indicated temperatures for Q55-
containing THRX-HDx1 spin labeled at residue 35 or 75

94
Fig. 5-3 Ms plot for THRX-HDx1-Q46 spin labeled at the indicated positions
on HDx1 at –20 and 25°C

95
Fig. 5-4 Ms for the indicated spin labeled residues of HDx1 within THRX-
HDx1 protein constructs of the indicated Q-lengths

96
Fig. 5-5 Sample temperature-dependent EPR spectra for THRX-HDx1-Q25
spin labeled at the indicated sites on HDx1

97
Fig. 5-6 Overlay of EPR spectra for THRX-HDx1-Q25 spin labeled at the
indicated sites on HDx1 and spectra measured at 4–10°C higher
98
IX

temperatures for THRX-HDx1-Q46 spin labeled at the same sites

Fig. 5-7 Sample temperature-dependent EPR spectra for THRX-HDx1-Q7
and Q16 spin labeled at the indicated sites on HDx1

99
Fig. 5-8 Temperature-shifted overlays of EPR spectra for four THRX-HDx1
constructs of the indicated Q-lengths spin labeled at residue 21 of
HDx1

100
Fig. 6-1 Wedging of an amphipathic helix into a phospholipid bilayer could
lead to membrane curvature induction or toroidal pore formation

105
Fig. 6-2 Vesicle budding induced by amphipathic helix insertion could lead
to tubule or channel-like pore formation

106
Fig. 6-3 Proposed model for HTT/HDx1 conformational equilibrium 109
Fig. 6-4 Proposed models for folding of HDx1 of normal and expanded Q-
length

110

X

ABSTRACT
Many degenerative diseases are associated with proteins that misfold and aggregate
into amyloid fibrils that are deposited intra- or extracellularly at or near the site of
tissue degeneration. It has long been established that the protein species conferring
toxicity in each case is most likely an intermediate on the pathway to fibrillization or an
off-pathway oligomer rather than the fibrils themselves, which are inert and could even
exist as a protective mechanism. However, while the current understanding of the
toxicity of amyloid intermediates remains rudimentary, research upon the
conformational behavior and possible toxicity of amyloidogenic proteins in monomeric
or oligomeric (but not aggregated) form is still more severely lacking. Such research
should prove crucial to the development of protein-targeting pharmaceuticals for
treatment of these diseases; this speculation is based on evidence, part of which has
originated in my own research, that, depending on conditions, these proteins as
monomers or oligomers can severely damage membranes and/or even in solution form
unfavorable structures. My research focuses on the cases of α-synuclein, islet amyloid
polypeptide (IAPP), and huntingtin, amyloidogenic peptides associated with cell death
in the substantia nigra in Parkinson’s disease, pancreatic β-cell death in type II diabetes,
and spiny neuron degeneration in Huntington’s disease, respectively.
Structural studies have shown α-synuclein to assume multiple membrane-
binding modes when in contact with phospholipids, including an extended
amphipathic α-helix and a broken-helix conformation consisting of two shorter α-
helices interrupted by a short linker region. Other experiments have shown α-synuclein
to induce the formation of highly curved membranous structures upon binding to
negatively charged phospholipid bilayers. I reconciled these findings by probing the
helical structure of α-synuclein bound to membranes made up of compositions
corresponding to the latter findings. Using an environment-sensitive fluorophore, I
found the α-synuclein binding to these membranes to be in the extended α-helical form.
XI

My work with IAPP was more extensive. Using a variety of techniques, from
clearance assays (to measure light scattering loss by IAPP-affected vesicles) to leakage
assays, from fluorescence microscopy to electron microscopy, from circular dichroism
to thioflavin T fluorescence assays to ensure that the effects I was gauging were exerted
by non-fibrillized IAPP, I unprecedentedly uncovered a mechanism by which IAPP
damages membranes by remodeling their architecture, causing them to form more
highly curved structures much like the curved structure formation induced by α-
synuclein, suggesting that this could be a common mechanism among multiple protein
toxins. In addition, using circular dichroism on IAPP bound to vesicles of different
sizes, I discovered that this peptide is a membrane curvature sensor, not only an
inducer. In vivo data on immunogold-labeled IAPP in rat and human pancreatic cells
showing IAPP to bind mitochondrial cristae, which comprise membranes that are by
nature rich in high curvature, suggest physiological consequences of the peptide’s
membrane curvature-sensing ability.
My studies on huntingtin, an essential protein for a number of cellular processes
that is only known to cause Huntington’s disease when the polyglutamine (polyQ)
region in its first exon contains an above-threshold number of Q residues (Q-length), are
covered in Chapters 4 and 5, starting with circular dichroism studies showing
huntingtin exon 1 of abnormally high Q-length to adopt additional α-helical structure in
solution to the α-helical structure that is normally present in this protein. This structural
effect of increased Q-length can be exaggerated by decreasing the temperature of the
sample and diminished by raising the temperature; temperature effects are reversible
and are therefore typical of conformational transitions. Mimicking phosphorylation at
serine 16 with an aspartic acid mutation structurally alters huntingtin exon 1 protein of
higher Q-length such that its conformational behavior closely resembles that of its
normal Q-length counterpart. Chapter 5 focuses on my electron paramagnetic
resonance work, which clarifies the location of additional structure formation in
XII

huntingtin exon 1 of extended Q-length: it occurs in the 17-aa N-terminal region as well
as in residues of the polyQ region that are closer to the N-terminal end.
Altogether, these findings emphasize the importance of studying the structures,
functions, and mechanisms employed by amyloid-forming proteins under conditions
wherein they have not yet formed amyloid.
1

Chapter 1

Amyloid Proteins and Structural Studies

2

1.1 Proteins and amyloid formation: an introduction
Amyloid proteins are peptides or full-size proteins that misfold into insoluble β-sheet
fibrils that are then deposited in cells or in the extracellular matrix (1). Compelling
evidence continues to accumulate that, depending on conditions, any protein can form
amyloid (2,3), and some proteins in doing so are functional—curli fibers in certain
bacteria (4), the fungal HET-s prion that controls heterokaryon incompatibility (5), and
the long-term memory-stabilizing, prion-forming cytoplasmic polyadenylation element-
binding (CPEB) protein (6) are a few examples that come to mind. However, a subset of
proteins are notorious for their stubborn propensity to misfold into amyloid fibrils and
the association of their amyloid deposits with human disease. Among these proteins
and their associated diseases are α-synuclein in Parkinson’s disease (PD) (7), islet
amyloid polypeptide (IAPP) in diabetes (8-10), amyloid β-peptide and tau in
Alzheimer’s disease (11,12), the prion PrP
Sc
in Creutzfeldt-Jakob disease (13), and
huntingtin in Huntington’s disease (HD) (14).
Such amyloid proteins share a strong tendency to aggregate and form β-sheets
on their way to becoming fibrils (7,10,14,15). One of the most intriguing factors into
their aggregation and fibrillization kinetics has been found to be the presence of
membranes of physiologically relevant morphology and composition, which can
accelerate amyloid proteins’ misfolding kinetics (16-19). At least as intriguing is that
amyloid proteins affect membranes in turn; while the fibrils themselves are mostly
found to be inert, oligomers of these proteins on or off the pathway to fibril formation
damage cellular and cell-mimicking membranes and this could be the primary
mechanism behind amyloid disease-related cytotoxicity (20).
Since the three proteins of focus for my PhD studies have been α-synuclein,
IAPP, and huntingtin, I provide a brief introduction to each immediately below.
3

1.1.1 α-Synuclein
α-Synuclein is a 140-aa (sequence shown in Fig. 1-1A), brain-enriched protein
that is part of the synuclein family (21). α-, β-, and γ-synucleins share a lipid-binding
motif that resembles the amphipathic α-11/3 helix in certain apolipoproteins (22). α-
Synuclein is a component of cellular inclusions called Lewy bodies in PD as well as in
dementia with Lewy bodies and multiple system atrophy (23,24). While mutations in
the α-synuclein-encoding gene are not necessary for the development of disease, the
protein’s causative role in PD has been clarified upon a large body of evidence that
includes the discoveries of three point mutations (A53T, A30P, E46K) that cause familial
PD and of the fact that amplification of the α-synuclein gene locus leads to PD (25-30).
Aggregation and amyloid fibril formation by α-synuclein have been implicated as the
disease-causing mechanism (31), though intermediate structures formed by α-synuclein
on or off the pathway to fibrillization, rather than the fibrils themselves, are believed to
be the causative force (32). α-Synuclein associates intimately with membranes in vivo (as
reviewed below), while its fibrillization kinetics have been observed in vitro to be

Fig. 1-1. Amino acid sequences of the three proteins of interest in this work. A. α-synuclein
sequence. B. Human and rodent IAPP sequences, with the six amino acids in the latter that
are different shown in blue. C. Huntingtin exon 1, showing the N17 region in green, the
polyQ tract in red, and the polyproline-rich C-terminal region in gray.

4

directly affected by phospholipid membranes such that these membranes speed up or
slow down fibril formation depending upon their composition and the protein-to-lipid
ratio used (33). This observation is part of an overall pattern wherein membranes
stabilize secondary structure formation by intrinsically disordered proteins including α-
synuclein by allowing them to form helices along the membrane plane (20), yet also in
many cases accelerate their formation of fibrils, a very different conformation that is
chiefly composed of β-sheet structure (34).
Importantly, α-synuclein-membrane interactions also dramatically impact
membrane morphology. Physiological and disease-associated consequences of this
impact are abundantly evident, as a growing body of research points to the function of
α-synuclein as a regulator of normal presynaptic functions (35-37) and the impairment
of vesicle trafficking among neurons upon α-synuclein deregulation (38). α-Synuclein
promotes soluble N-ethylmaleimide sensitive fusion protein attachment protein
receptor (SNARE)-complex assembly via a non-classical chaperone mechanism that
involves its simultaneous binding of SNARE proteins and of membranes (37). α-
Synuclein regulates the size of pools of presynaptic vesicles in hippocampal neurons
(35). A recent study revealed the localization of this protein at the synaptic vesicle
apparatus of the enteric nervous system (39), showing that the physiological functions
of α-synuclein are not limited to the brain. On the other hand, the implications of α-
synuclein-synaptic vesicle machinery interactions for disease mechanisms must be paid
at least equal heed. α-Synuclein accumulation inhibits protein trafficking from the ER to
the Golgi, and the effect is worsened if α-synuclein contains the A53T PD mutation (40).
Neurons of transgenic mice overexpressing human α-synuclein have enlarged
presynaptic terminals at sites of overexpression, with extensive membranous structures
that display altered tubulovesicular architecture under the electron microscope (EM)
(41) Furthermore, extracellular accumulation of α-synuclein was found to lead to
leakage of hippocampal neuron membranes (42). These findings stress the importance
5

of probing into the mechanisms of alterations in membrane morphology and disruption
of membrane integrity caused by α-synuclein.
1.1.2 IAPP
IAPP, or amylin, is a 37-aa (sequences shown in Fig. 1-1B) member of the calcitonin
gene family of peptides (10) and is co-secreted with insulin upon the consumption of
food (43,44). It is believed to play a role in conjunction with insulin and other hormones
in promoting satiety (45). In humans, IAPP is the principal component of islet amyloid
commonly present in type II diabetes (46). Furthermore, numerous cell and animal
studies support a causative role of human IAPP (hIAPP) in type II diabetes (47-49). Rat
IAPP (rIAPP), as well as mouse IAPP, does not readily form amyloid fibrils, and
rodents do not spontaneously develop type II diabetes (50). Therefore, rIAPP is mostly
deemed to be nontoxic, but it has recently been found to cause the same cytotoxic
effects as hIAPP if present at much higher concentrations (51). The structural difference
between the peptides that has long been believed to be the reason that one is cytotoxic
and the other is (relatively) nontoxic is the three proline residues in rIAPP that hinder
the intermolecular interactions that would otherwise lead to fibril formation (52), but
more recent findings have also revealed the contribution of His18 in hIAPP (as opposed
to Arg18 in rIAPP; see Fig. 1-1B) to its ability to more effectively damage membranes
(53). Disruption of membrane integrity, as in the case of other amyloidogenic proteins
(54-56), is thought to be one of the mechanisms by which IAPP can cause toxicity
(47,51,57). The presence of synthetic vesicles of physiologically relevant membrane
compositions has been continually observed to accelerate hIAPP fibril formation (17,58);
conversely, IAPP-membrane interactions lead to a variety of membrane morphology
changes, including partial to total loss of membrane integrity (59). Studies on IAPP-
membrane interactions are evidently key to understanding this protein’s cytotoxicity.
1.1.3 Huntingtin
6

HD is a devastating neurodegenerative disorder of autosomal dominant inheritance
pattern caused by a CAG trinucleotide repeat expansion in the IT15 gene, which
encodes huntingtin, a ~350-kDa protein (60,61). As a result of the expansion, mutant
huntingtin has an abnormally long number (>35) of glutamines (Q-length) in a row in
its first exon (62). HD is one of nine polyglutamine (polyQ) expansion diseases, all of
which mainly cause neurological symptoms (63). In particular, HD-associated
neurodegeneration presents as loss of coordination, unsteady gait, chorea (involuntary
spasmodic movements of the trunk and limbs [64]), and psychiatric symptoms such as
depression and psychosis (62). HD also exerts a range of other physiological effects (65)
including reduced testosterone circulation concomitant with testicular degeneration in
males (66,67), marked, Q-length-dependent weight loss (68-70), and loss of muscle bulk
(71). The psychiatric symptoms eventually devolve into severe cognitive impairment
(72), and the HD patient dies typically within twenty years of initial symptom onset,
with the immediate cause of death often stemming from complications that result from
the loss of coordination (62).
Besides being the polyQ-containing segment, huntingtin exon 1 (HDx1) is
sufficient, when containing a polyQ expansion, to cause HD-like symptoms in
transgenic mice (73). Furthermore, it is the product of aberrant mRNA splicing in HD
knockin mouse models and HD patients (74). Huntingtin aggregate-containing cellular
inclusions in HD brains mainly consist of N-terminal huntingtin fragments, including
HDx1, which are the products of protease-induced cleavage processes that are
augmented in HD (64,75). HDx1 comprises a 17-aa N-terminal domain (N17) that is
thought to be largely helical, the polyQ tract, and a polyproline-rich C-terminal domain
(76) (sequence, with domains marked, shown in Fig. 1-1C). The N17 and C-terminal
domains immediately flanking the polyQ region affect its conformational behavior
(including amyloid fibril formation) and its toxicity (77-83). Therefore, in order to zone
7

in on residues that play direct structural roles in modulating polyQ-associated toxicity, I
have focused on HDx1 in my Ph.D. work.
The normal physiological functions of huntingtin appear to be manifold and
therefore have proven difficult to pinpoint. Huntingtin is necessary for development, as
huntingtin-null mice die as embryos (84). A growing body of research shows autophagy
and related vesicle trafficking events to be the main functions served by huntingtin (85-
87). In addition, huntingtin plays roles in ciliogenesis (88) and in mitotic spindle
establishment (89). Throughout its variety of functions, huntingtin interacts with a
plethora of binding partners, from the transport protein β-tubulin (90) to tumor
suppressor p53 (91), from protein kinase C and casein kinase 2 substrate in neurons
(PACSIN1) (92) to the nuclear export protein translocated promoter region (Tpr) (93), as
well as with membranes (94). PolyQ expansion can up- or downregulate many of these
interactions. Therefore, to better understand the normal and disease-associated roles of
huntingtin, it is important, as with other amyloid-forming proteins, to study it in its
monomeric/oligomeric forms.
1.2 Structural studies
Studies utilizing fluorescence-dependent techniques, X-ray diffraction, electron
paramagnetic resonance (EPR) with site-directed spin labeling (SDSL), and molecular
dynamics, including studies conducted in our laboratory, have generated visualization
of the structures of amyloid fibrils (95-101). It is also crucial, but at times proves more
elusive, to elucidate the structures of the monomers or oligomeric intermediates of these
proteins, since these are the structures that affect membranes, lend insight into the
proteins’ physiological functions, and serve as the starting structures for aggregation
and fibrillization. In particular, it is important to solve the membrane-bound structures
of these proteins, since their membrane interactions have such varied and complex
physiological and disease implications. The micelle-bound structure of the first 17
amino acids of huntingtin when probed by nuclear magnetic resonance (NMR) revealed
8

an amphipathic α-helix from residues 6 to 17 (102)—that is, an α-helix with one
hydrophobic face whose length is buried along the lipid and one hydrophilic face that is
in contact with solvent. α-Synuclein and IAPP, unlike huntingtin, are small enough to
have their structures studied whole rather than in fragments and their amino acids
mapped out individually. Structural studies on α-synuclein and IAPP are briefly
described immediately below; amphipathic helix formation on lipids has repeatedly
been observed by these proteins as by the huntingtin N-terminus.

Fig. 1-2. Structural models of the α-synuclein helical structure. (A) Cross-sectional view of α-
synuclein bound to membrane as an extended α-helix, model generated by simulated
annealing molecular dynamics using EPR data from our laboratory. From Jao et al., 2008. (B)
Cartoon representation of α-synuclein bound to an SDS micelle and to a vesicle, which as
opposed to the micelle can accommodate the length of its extended helical conformation.
From Jao et al., 2008. (C) α-Helical residues from the extended α-helix of α-synuclein plotted
on a helical wheel. Green designates residues found by EPR to face solvent, red found to face
lipid. Modified from Jao et al., 2004. (D) Helical wheels showing the two parts of the broken α-
helix formed by α-synuclein on micelles. From Bussell and Eliezer, 2003.
C
90
79
68
57
42
31
20
9
86
75
64
49
38
27
16
82
71
60
45
34
23
12
89 78 67 5241
30
19
85
74
63
48
37
26
15
81
70
59
44
33
22
11
88
77
66
51
40
29
18
84
73
62
47
36
25
14
80
10
21
32
43
58
69
17
28
39
50
65
76
87
13
24
35
46
61
72
83
A B
D
9

Fig. 1-3. Structural models of the hIAPP helical structure. (A) Native IAPP is unstructured in
solution. On membranes, it binds as a helix from residues 9 to 22. Residues 23 to 29 constitute the
amyloidogenic region. The proximity and alignment of IAPP molecules as membrane-bound
helices are likely the factors that lead to interactions between the regions between residues 23 and
29 that accelerate fibrillization. Modified from Apostolidou et al., 2008. (B) Helical wheel of hIAPP
helical residues, with blue designating solvent-exposed and red designating lipid-exposed
residues according to EPR data. From Apostolidou et al., 2008. (C) Schematic model of EPR data-
based structure of membrane-bound hIAPP. Residues 9-20, the central helical core, are shown as a
thick red ribbon; residues 21 and 22, which still appeared helical by EPR but showed elevated
mobility, as a thinner red ribbon; and residues 1 to 8 and 23 to 37, which showed no specific
structure, as a thin black dotted line. From Apostolidou et al., 2008.

C A B
1.2.1 α-Synuclein membrane-bound structure
α-Synuclein has long been observed by circular dichroism (CD) to transition
from its native unstructured conformation in solution to an α-helix upon addition of
lipid (103-105). NMR measurements of its SDS micelle-bound structure revealed a
broken helix, one from residues 1 to 41 and one from residues 45 to 94, bound parallel
to the micelle surface (106). In our laboratory, we used EPR with SDSL to solve the
structure of α-synuclein bound to negatively charged vesicles. We found it to form a
single extended helix (Fig. 1-2A) (107,108), in contrast to the broken helix previously
found on micelles. This difference can be explained by the much smaller size of a
micelle compared to a large unilamellar vesicle (LUV), such that the protein needs to
bend in order for more of it to bind a micelle, while an LUV provides a spacious, flat
(from the viewpoint of the protein) surface for binding without bending (Fig. 1-2B). EPR
10

power saturations showed the membrane-bound α-synuclein helix to be amphipathic
and inserted parallel to the membrane, with the more polar and in some cases charged
residues facing solvent and the less polar and in some cases nonpolar residues facing
lipid (α-helical wheel shown in Fig. 1-2C). Both the NMR study and our EPR study
showed α-synuclein α-helices to deviate from the commonly seen 3.6-amino acid-per-
turn structure of many α-helices and instead to have seven (or four-plus-three in the
NMR-solved broken helix) 11-aa repeats each taking up three helical turns, resulting in
α-11/3 helices (compare α-helical wheels in Fig. 1-2C and Fig. 1-2D).
1.2.2 IAPP membrane-bound structure
Our laboratory has also found IAPP to bind negatively charged membranes as an
amphipathic α-helix (109). SDSL and EPR led us to the discovery that IAPP, whose
native structure in solution like that of α-synuclein is largely a random coil, binds
negatively charged LUVs as an α-helix from residues 9 through 22 (Fig. 1-3A). EPR
power saturations showed this helix, like that of α-synuclein, to be amphipathic and
inserted parallel to the membrane (α-helical wheel shown in Fig. 1-3B). EPR spectra of
spin labels attached to regions defined by residues 1-8 and residues 23-37 showed high
mobility and no specific order, so by our model (Fig. 1-3C) the N- and C-terminal
regions of IAPP remain highly disordered, at least until the intramolecular proximity of
helices lined up on the membrane allow sufficient contact between the amyloidogenic
C-termini (residues 23-29) to lead to aggregation (Fig. 1-3A). While NMR data on
micelle-bound IAPP from other groups show a longer helix (110) or a broken helix
(111), these differences arise, as in the case of lipid-bound α-synuclein, from the
structural difference between micelles and bilayer membranes as binding surfaces, yet
their data and ours are still in agreement insofar as the α-helices (one of the helices, in
the case of the broken helix) in question are amphipathic and bound parallel to lipid.
The fact that α-synuclein and IAPP take on similar membrane-bound structures is one
more clue to a common thread in their cytotoxic mechanisms.
11

1.2.3 EPR with SDSL: a brief overview
EPR with SDSL is a technique for measuring protein structure at, in theory, any given
locus. SDSL is achieved by mutating a residue at the desired locus to cysteine; the
remainder of the protein must be non-cysteine-containing. Protein is labeled with spin
label (Fig. 1-4A), which is thiol-reactive and will bind only, out of all the canonical
amino acid side chains, to the thiol group intrinsic to cysteine (112,113).
Spin label contains an unpaired electron (Fig. 1-4A), which like other electrons
has two spin states normally of equal energy. EPR applies a magnetic field to the
sample, and the magnetic field separates the energies of the spin states. The resulting
readout shows an absorption peak at the field strength at which the energy difference
between spin states is equal to the energy of the electromagnetic radiation. EPR spectra
usually look like three peaks due to splitting of the absorption peak (Fig. 1-4B). This
splitting is due to hyperfine interactions between the large magnetic field generated by

Fig. 1-4. Spin label attaches to
protein sulfhydryl groups and
gives an EPR signal that varies
with mobility. (A) The
commonly used spin label [1-oxy-
2,2,5,5-tetramethyl-pyrroline-3-
methyl]-methanethiosulfonate
(MTSL), which we have used in
the studies described herein.
From Margittai and Langen, 2006.
(B) Spectra ranging from broad
(top) to sharp and narrow
(bottom) designate a range of
mobilities from rigid to highly
mobile, respectively, for the spin
label and therefore for the site at
which it is attached. From Feix
and Klug, 1998.

A B
12

EPR and the small magnetic field generated by the magnetic moment composed of the
unpaired electron of the spin label and the adjacent
14
N nucleus (Fig. 1-4A) (114).
EPR spectra tend to vary from broad (Fig. 1-4B, top) to sharp (Fig. 1-4B, bottom)
as the motion of the spin label varies from highly mobile to rigid depending on the
structure of its locus (112,113). In this way, EPR can determine how buried or exposed
the spin label, and therefore how structured or unstructured the site of labeling, is.
In addition, EPR can measure the accessibility of spin-labeled sites to
paramagnetic colliders (113). The manner in which our laboratory, for example,
identified areas of membrane-bound amphipathic α-helix formation in α-synuclein and
IAPP (107-109) (see above) involved bombarding spin-labeled derivatives of those
proteins with the polar paramagnetic collider nickel ethylenediamine diacetic acid
(NiEDDA) and O2, a nonpolar paramagnetic collider, and varying the power of the EPR
microwave to measure the accessibility of spin label at each individual site of the
protein to each collider. Sites facing the membrane (nonpolar) collided more with O2,
sites facing solvent (polar), with NiEDDA. The periodicity of the α-helix was
determined directly from the periodicity of accessibilities (113).
These are a few common applications of EPR, which can also, among other
things, be used to measure intra- and intermolecular distances by gauging spin-spin
coupling between nearby spin labels (115) and also to measure the polarity of the local
environment (116), another clue to the structure of the protein in that region. Different
EPR techniques can be combined to generate three-dimensional molecular structures
(107,109,117,118).
1.3 Exertions by membrane-active proteins
Structural models for membrane-bound forms of amyloid-forming proteins offer
a firm basis for understanding preamyloid toxicity by allowing us to visualize what is
most likely the starting structure of each preamyloid toxic agent. However, these
models do not directly answer the question of how proteins disrupt membranes in
13

Fig. 1-5. Schematics showing two
major mechanisms by which
proteins induce membrane
curvature. (A) The scaffold
mechanism. Modified from
Zimmerberg and Kozlov, 2006. (B)
The wedge mechanism. Modified
from Campelo et al., 2008.
A
B
Protein scaffolding
domain
Membrane bilayer
Membrane
bilayer
Protein amphipathic
helix
disease. A variety of mechanisms have been proposed whose time points if they occur
span from initial membrane binding to the point of fibril formation. Potentially, under
different conditions, all these mechanisms could apply. Fibril growth at the membrane
involves nascent fibrils tearing at and rupturing the
entangled membrane in the process (119-122).
Preamyloid oligomers have been observed to cause
permeabilization of membranes, in some cases by
an ion-channel-like mechanism as evidenced by
specific pore formation and discrete changes in ion
conductance (123-125), in others as nonspecific
thinning or deformation of membranes (126-128),
perhaps implicating mechanisms such as the
toroidal pore or carpet or “detergent-like” models
similar to those inferred for antimicrobial peptides
(129-131). These varied but not necessarily
conflicting reports have elicited much controversy
that has been both complicated and explicated by
the differing experimental conditions that
generated the seemingly contradictory outcomes.
Mostly overlooked has been the possibility
that amyloid and perhaps other membrane-active
proteins disrupt membrane integrity by inducing membrane curvature. Outside of the
amyloid disease field, membrane curvature induction is a widely studied but still
poorly understood necessity that enables a variety of crucial cellular processes, from cell
division to organelle formation, from protein transport by the ER-Golgi system to
synaptic vesicle transmission by neurons (132). Different proteins exert a variety of
mechanisms to conquer the energy barrier against high membrane curvature and
14

thereby permit these cellular functions. For instance, a Bin-Amphiphysin-Rvs (BAR)
domain protein will bind a membrane with the concave surface of its banana-like BAR
domain, forcing the membrane to take on the same curved shape (Fig. 1-5A) (132,133).
In contrast to this scaffolding mechanism, a protein can induce membrane curvature by
inserting amphipathic α-helices parallel to the bilayer between phospholipid
headgroups, creating an imbalance between the two leaflets of the bilayer and causing
the membrane to bend to accommodate this imbalance (Fig. 1-5B) (132,133). Epsin, a
brain-enriched protein involved in endocytosis, is a small globular protein, not a
scaffold (132), and drives membrane curvature by inserting epsin N-terminal homology
(ENTH) domains, which are amphipathic helices, into the outer leaflet of membranes
(134). Even some BAR domain proteins have been found to use this latter mechanism as
well, not purely the former scaffolding mechanism: in fact, the endocytic regulatory
BAR domain proteins amphiphysin and endophilin were found to require their N-
terminal amphipathic-helix domains to bind membranes and curve them into tubular
structures (135,136).
The ability of epsin to induce membrane curvature in the absence of a scaffolding
domain suggested that amphipathic helices alone were sufficient to remodel
membranes in this fashion. Given that amyloid-forming proteins such as α-synuclein
and IAPP were found to form amphipathic α-helices on anionic membranes (107-109),
did they also alter membrane architecture by this wedge mechanism? To answer this
question, we needed to view both membrane remodeling and loss of membrane
integrity while holding experimental conditions constant in the presence of these
proteins.

15

Fig. 1-6. Leakage of vesicles
composed of negatively charged
phospholipids and loaded with
the fluorophore 8-
aminonaphthalene-1,3,6-
trisulfonic acid (ANTS) and its
quencher p-xylene-bis(pyridinium
bromide) (DPX) in presence ofα-
synuclein and amphiphysin. Red,
α-synuclein with POPG vesicles
(1:20 protein/lipid molar ratio).
Broken blue, β-synuclein with
POPG vesicles (1:20 protein/lipid
molar ratio). Green, amphiphysin
with POPG/1-palmitoyl-2-oleoyl-
sn-glycero-3-phosphoethanolamine
(POPE) (2:1 molar ratio) vesicles
(1:100 protein/lipid molar ratio).
Black, control vesicles without α-
synuclein or β-synuclein. Blue,
control vesicles without
amphiphysin N-BAR domain.
Large extruded 1-μm vesicles were
used. From Varkey et al., 2010.
1.4 Loss of membrane integrity in conjunction with membrane remodeling
One widely utilized manner of evaluating loss of membrane integrity under test
conditions, such as treatment with antimicrobial peptides, is leakage measurements
(137). These experiments are performed by loading synthetic model vesicles with a
fluorophore and its quencher (or else a self-quenching fluorophore) and measuring the
fluorescence of the fluorophore over time upon
addition of the peptide in question, which if
leakage occurs should dramatically increase as
fluorophore and quencher are released as the
membrane loses its integrity. (A variation of this
method is to load vesicles instead with a
conditional fluorophore and the ion upon which
its fluorescence depends, then to include a
chelator in the outside solution. Leakage in this
case would manifest itself conversely as a drop in
fluorescence [see, for example, ref. 138].) Our
laboratory and others have measured leakage of
vesicle contents upon addition of α-synuclein or
IAPP. Both peptides, even if only data gleaned
during time frames within which the peptide
remained an α-helix on the membrane were
considered, have continually been found to induce
leakage of negatively charged membranes (56,139-
142). Our laboratory found α-synuclein to cause
leakage of POPG vesicles to an extent comparable
to that caused by the known curvature inducer
16

Fig. 1-7. Leakage of POPS/POPC (~2:1 molar ratio) vesicles in the presence of IAPP. (A)
Adding rIAPP at successively higher concentrations to these vesicles induces increasing
leakage, showing the ability of nonamyloidogenic IAPP to damage membranes. The detergent
Triton X is added at the end of the experiment to define the leakage reaction end point
(maximal fluorescence). From Melania Apostolidou. (B) hIAPP causes leakage to these vesicles
in a concentration-dependent manner. The concentration dependence is not linear; rather it is
sigmoidal, suggesting a cooperative mechanism for IAPP-induced membrane leakage. From
Melania Apostolidou’s Dissertation, 2009.

0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
0 1000 2000 3000 4000
Normalized fluorescence intensity
Time (seconds)
buffer
66:34
POPS/POPC
1 μM
rIAPP
1 μM
rIAPP
31 μM
rIAPP
16 μM
rIAPP
6 μM
rIAPP
Triton X
1 μM
rIAPP
6 μM
rIAPP
A B
amphiphysin and greater than that caused by the non-amyloid-forming β-synuclein
(Fig. 1-6). It should be noted that β-synuclein did induce considerable leakage (Fig. 1-6),
precluding the necessity of amyloid-forming ability and of possession of a BAR domain
for causing membranes to lose their integrity.
Our IAPP leakage studies (see also Chapter 3) similarly showed IAPP to induce
sufficient membrane disruption to lead to leakage without the need for a scaffold or
amyloid formation, as IAPP like α-synuclein contains no scaffolding domain and the
nonamyloidogenic rIAPP (Fig. 1-7A) caused leakage of partially negatively charged
vesicles nearly as much as did hIAPP (Fig. 1-7B). Note in Fig. 1-7B that IAPP
concentration seemed to affect the extent of leakage in a sigmoidal fashion. This
17

Fig. 1-8. Clearance of negatively charged
MLVs by α-synuclein. (A) Apparent
absorbance at 500 nm was monitored to
measure vesicle clearance. Control traces
for POPG vesicles (400 μM) in the absence
of α-synuclein are indicated by the broken
blue trace. POPG vesicles (400 μM)
incubated with 10 μM, 20 μM, and 40 μM
of α-synuclein are shown with the blue,
green, and red traces respectively. Control
POPC vesicles (400 μM) in the absence
and presence of 40 μM α-synuclein are
given by the black and broken red lines,
respectively. From Varkey et al., 2010.
(B) A glass tube containing POPG MLVs before and after α-synuclein treatment. Clearance can be seen
by the naked eye. From Varkey et al., 2010. (C) Clearance measured on other lipid compositions in a
similar fashion as in (A). 600 μM lipid was incubated with 15, 30, and 60 μM α-synuclein, as indicated by
the blue, red, and green traces, respectively. From Mizuno et al., 2012.

suggests a cooperative mechanism among α-helices causing or optimizing leakage, an
idea that was corroborated by studies from Dr. Andrew Miranker’s group (138,143).
In the case of α-synuclein, membrane damage was not limited to loss of
membrane integrity leading to leakage of vesicle contents, but also included a
mechanism not previously observed for amyloid proteins in the α-helical state: α-
synuclein-bound membranes could be remodeled into new morphological structures. In
our laboratory, multilamellar vesicles (MLVs) made up of the widely used negatively
charged phospholipid 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-RAC-(1-glycerol)]
(POPG) lost much of their light scattering after treatment with α-synuclein at 1:40, 1:20,
and 1:10 protein-to-single phospholipid molar ratios; this effect was measured by OD500
(Fig. 1-8A) but could even be qualitatively detected simply by holding the sample-
containing cuvette to the light before and after addition of α-synuclein (Fig. 1-8B).
Varying the acyl chains on the membrane phospholipids did not abrogate the effect of
vesicle clearance by α-synuclein (Fig. 1-8C).
18

Fig. 1-9. α-Synuclein induces the
formation of highly curved
structures from POPG membranes.
EM of (A-D) POPG alone (A) and in
the presence of α-synuclein at 1:40 (B),
1:20 (C), and 1:10 (D) protein-to-lipid
molar ratios. From Varkey et al., 2010.
(E) EM of supernatant obtained after
ultracentrifugation of wild-type
incubated with POPG vesicles at a
protein-to-lipid molar ratio of 1:10.
White arrowheads indicate
nanoparticles. From Varkey et al.,
2013. (F) Schematic of a helical
insertion inducing membrane
curvature by pushing phospholipid
headgroups apart. All scale bars, 100
nm. From Varkey et al., 2010.
Furthermore, electron microscopy (EM) studies in our laboratory have
uncovered the ability of α-synuclein to remodel negatively charged vesicles (intact
POPG multilamellar vesicles shown in Fig. 1-
9A). The nature of the remodeling observed was
dependent on the protein-to-phospholipid ratio.
At 1:40 as well as 1:20, the protein-to-
phospholipid ratio used in our synuclein-
induced leakage assays, α-synuclein induced the
formation of tubular structures by MLVs of
POPG (Fig. 1-9BC). Formation of such tubules
from vesicles is direct visual evidence of
curvature induction. At the higher ratio of 1:10,
the formation of smaller vesicular structures was
observed (Fig. 1-9D), suggesting that α-
synuclein induced smaller vesicles to pinch off
the large MLVs. At this higher ratio, EM
following ultracentrifugation to separate
different-sized particles within the sample
revealed the supernatant to contain yet smaller
lipidic structures (Fig. 1-9E) that further analysis
characterized as lipoprotein nanoparticles. Our
explanation, in view of these data, membrane-
bending mechanisms exerted by other proteins, as reviewed in (132), and the EPR-
solved structure of membrane-bound α-synuclein (107,108), was that α-helical α-
synuclein insertions wedged between phospholipid headgroups and pushed them
apart, forcing the membrane to adopt what would otherwise be an energetically
unfavorable highly curved shape (Fig. 1-9F). Even more severe remodeling into
19

nanoparticles resulted from α-synuclein helices at high local concentrations wrapping
around and further contorting the membrane. We correlate the observed membrane
remodeling with the membrane dissolution as viewed by clearance and leakage as
viewed by fluorescence that we observed under the same conditions. Another group
(144) independently observed tubulation induced by α-synuclein on membranes of
lower negative charge content.
Both the loss of light scattering by MLVs and the EM observations of highly
curved morphology formation demonstrate membrane remodeling by α-synuclein. At
the time, other amyloid proteins had yet to be tested for this phenomenon. Data
pointing to a membrane remodeling mechanism in other proteins would unveil a likely
source of membrane damage pervading a variety of amyloid diseases. In Chapter 3, I
tackle the question of whether IAPP displays a similar capacity to that of α-synuclein
for altering membrane architecture.
1.5 Huntingtin non-amyloid conformational behavior changes with Q-length
Abnormal polyQ expansion in huntingtin can have a variety of deleterious effects (as
reviewed in ref. 145), ranging from loss of function due to haploinsufficiency to a non-
amyloid-related gain of structure due to the Q-expansion, leading to a toxic gain of
function, to the possible formation of a β-sheet pocket that serves as a basis for
aggregation. Dr. Ray Truant and his group have hypothesized that polyQ expansion
forms a “rusty hinge” in the middle of exon 1 that abrogates switching between
conformations to carry out normal functions and this turns huntingtin from a functional
to a cytotoxic protein (145). They went on to perform F ӧ rster resonance energy transfer
(FRET) experiments in STHdh
Q7/Q7
cells (which are rat spiny neuron-like cells) by
attaching an mCerulean donor and an enhanced yellow fluorescent protein (eYFP)
acceptor to the N- and C-terminal ends, respectively, of HDx1 (146). The FRET effect
was enhanced (over intermolecular controls, in which each HDx1 molecule only had
either one of the fluorophores) in HDx1 constructs containing a normal number of
20

glutamines, but not in HDx1 constructs with above-disease threshold Q-length (Fig. 1-
10). It seems likely that the polyQ region becomes rigid above that threshold,
preventing the joining of the mCerulean- and eYFP-laden ends (Fig. 1-10, the authors’
model). Furthermore, this effect was modulated by phosphorylation and
phosphomimicry. Mutating Ser13 and Ser16 to glutamic acid (which mimics
phosphorylation) or
treating cells with an
IKK inhibitor (which
leads to increased
phosphorylation)
caused a loss of FRET
efficiency, while
treatment with CK2
inhibitors improved
FRET efficiency in
the expanded-polyQ
construct (146). This
suggests that
tinkering with
phosphorylation in
the N17 region can
remedy the structural
behavior of HDx1 that confers cytotoxicity in HD.
Lower temperature can unmask a tendency in protein to form extraneous
structures. Dr. Andrea Caricasole’s group incubated a purified huntingtin N-terminal
construct consisting of the first 548 amino acids of huntingtin (N548Htt) with the N17-
region-binding and polyQ-tract-binding antibodies 2B7 and MW1, respectively; 2B7
Fig. 1-10. Expanded-polyQ
HDx1 loses its ability to
bring its N- and C-termini
in proximity. From top:
Sample live-cell
fluorescence and
fluorescence lifetime
imaging microscopy images
for HDx1-Q17 (normal-level
Q-length) tagged with
mCerulean at the N-
terminus and eYFP at the C-
terminus, HDx1-Q17 tagged
with either one as an
intermolecular control,
HDx1-Q138 (disease-level
Q-length) tagged with the
two fluorophores, and
HDx1-Q138 tagged with
either one. Authors’ models
of normal (left) and
expanded (right) HDx1
that, respectively, is and is
not able to join its N- and C-
terminal ends. From Caron
et al., 2013.

21

Fig. 1-11. Temperature-induced change in FRET efficiency is
greater for N548Htt-Q16 than for N548Htt-Q55, with the room-
temperature baseline for the former being much lower than for
the latter. Shown: TR-FRET fluorescence signals from a dilution
curve of N548Htt proteins of the indicated Q-lengths, obtained
using the indicated antibody combination at the indicated
temperatures. Modified from Fodale et al., 2014.
ng protein
was conjugated to the FRET donor terbium, MW1 to its acceptor D2 (147). They
measured FRET efficiency between the N548Htt-bound antibodies at room temperature
and at 4°C and found a temperature-dependent increase in FRET efficiency from
samples of normal- and of expanded-Q-length N548Htt (Fig. 1-11); however, expanded-
Q-length N548Htt showed a relatively high FRET enhancement at both temperatures
with only a small increase between them, while normal-Q-length N548Htt showed a
dramatic increase in FRET efficiency with lowered temperature from a much lower
FRET signal reading at room temperature (Fig. 1-11). Similar results were found in full-
length huntingtin and HDx1. While this might seem like a contradiction to the Truant
group’s results (ref. 146
and Fig. 1-10), the Truant
group had examined
FRET between the N17
and C-terminal regions,
not the N17 and polyQ
regions (when the
Caricasole group
performed another
version of their
experiment using 2B7
and the HDx1 C-
terminal-binding
antibody 4C9,
conjugating them with
terbium and the terbium fluorescence acceptor A647, they found no significant
temperature effect on FRET efficiency [147]); also, just as the results from the Truant
group appear to be caused by increased rigidity resulting from a polyQ expansion, the
22

increased FRET efficiency observed by the Caricasole group arises from lowered
temperature, which generally increases structure and herein appears to be modifying
the conformational behavior of normal-Q-length protein to appear approximately as
expanded-Q-length protein would at room temperature.
These findings are exciting but still require clarification. They reveal the effect of
polyQ expansion on huntingtin intramolecular interactions, but they do not define the
exact structural changes that come with these altered interactions, or the locations of
these structural changes. Furthermore, the use of antibodies precludes absolute
certainty that the changes seen are structural in nature because they could result from
differential temperature effects on antibody/epitope recognition. In Chapters 4 and 5, I
tackle the questions of actual temperature-dependent conformational changes in HDx1
and the location on the protein of these changes.
1.6 Closing thoughts
Research into amyloid diseases and their proteins has blossomed into a wide set of
findings of possible physiological and disease mechanisms for those proteins in their
varied shape-shifting forms. However, these findings are incomplete and, to continue to
grow, require a deeper molecular understanding of what intramolecular,
intermolecular, and membrane interactions amyloid-forming proteins can have in the
cell. In my PhD studies, I have used a variety of techniques, from CD to EPR to
microscopy, to add to the stepping-stones of known information about the
conformational behaviors that α-synuclein, IAPP, and huntingtin (specifically, HDx1)
exhibit before they form amyloid fibrils.

23

1.7 References
1. Bolshette, N. B., Thakur, K. K., Bidkar, A. P., Trandafir, C., Kumar, P., and Gogoi,
R. (2014) Protein folding and misfolding in the neurodegenerative disorders: a
review. Rev. Neurol. (Paris) 170, 151-161
2. Dobson, C. M. (1999) Protein misfolding, evolution and disease. Trends Biochem.
Sci. 24, 329-332
3. Schmittschmitt, J. P., and Scholtz, J. M. (2003) The role of protein stability,
solubility, and net charge in amyloid fibril formation. Protein Sci. 12, 2374-2378
4. Wang, X., Zhou, Y., Ren, J. J., Hammer, N. D., and Chapman, M. R. (2010)
Gatekeeper residues in the major curlin subunit modulate bacterial amyloid fiber
biogenesis. Proc. Natl. Acad. Sci. U. S. A. 107, 163-168
5. Ritter, C., Maddelein, M. L., Siemer, A. B., Luhrs, T., Ernst, M., Meier, B. H.,
Saupe, S. J., and Riek, R. (2005) Correlation of structural elements and infectivity
of the HET-s prion. Nature 435, 844-848
6. Raveendra, B. L., Siemer, A. B., Puthanveettil, S. V., Hendrickson, W. A., Kandel,
E. R., and McDermott, A. E. (2013) Characterization of prion-like conformational
changes of the neuronal isoform of Aplysia CPEB. Nat. Struct. Mol. Biol. 20, 495-
501
7. Conway, K. A., Harper, J. D., and Lansbury, P. T., Jr. (2000) Fibrils formed in
vitro from alpha-synuclein and two mutant forms linked to Parkinson's disease
are typical amyloid. Biochemistry 39, 2552-2563
8. Westermark, P., Wilander, E., and Johnson, K. H. (1987) Islet amyloid
polypeptide. Lancet 2, 623
9. Clark, A., Cooper, G. J., Lewis, C. E., Morris, J. F., Willis, A. C., Reid, K. B., and
Turner, R. C. (1987) Islet amyloid formed from diabetes-associated peptide may
be pathogenic in type-2 diabetes. Lancet 2, 231-234
10. Westermark, P., Wernstedt, C., Wilander, E., and Sletten, K. (1986) A novel
peptide in the calcitonin gene related peptide family as an amyloid fibril protein
in the endocrine pancreas. Biochem. Biophys. Res. Commun. 140, 827-831
11. Glenner, G. G., and Wong, C. W. (1984) Alzheimer's disease: initial report of the
purification and characterization of a novel cerebrovascular amyloid protein.
Biochem. Biophys. Res. Commun. 120, 885-890
12. Grundke-Iqbal, I., Iqbal, K., Quinlan, M., Tung, Y. C., Zaidi, M. S., and
Wisniewski, H. M. (1986) Microtubule-associated protein tau. A component of
Alzheimer paired helical filaments. J. Biol. Chem. 261, 6084-6089
13. Wadsworth, J. D., Joiner, S., Hill, A. F., Campbell, T. A., Desbruslais, M., Luthert,
P. J., and Collinge, J. (2001) Tissue distribution of protease resistant prion protein
in variant Creutzfeldt-Jakob disease using a highly sensitive immunoblotting
assay. Lancet 358, 171-180
24

14. Huang, C. C., Faber, P. W., Persichetti, F., Mittal, V., Vonsattel, J. P., MacDonald,
M. E., and Gusella, J. F. (1998) Amyloid formation by mutant huntingtin:
threshold, progressivity and recruitment of normal polyglutamine proteins.
Somat. Cell Mol. Genet. 24, 217-233
15. Zhou, Z., Yan, X., Pan, K., Chen, J., Xie, Z. S., Xiao, G. F., Yang, F. Q., and Liang,
Y. (2011) Fibril formation of the rabbit/human/bovine prion proteins. Biophys. J.
101, 1483-1492
16. Zhu, M., and Fink, A. L. (2003) Lipid binding inhibits alpha-synuclein fibril
formation. J. Biol. Chem. 278, 16873-16877
17. Jayasinghe, S. A., and Langen, R. (2005) Lipid membranes modulate the structure
of islet amyloid polypeptide. Biochemistry 44, 12113-12119
18. Terakawa, M. S., Yagi, H., Adachi, M., Lee, Y. H., and Goto, Y. (2015) Small
liposomes accelerate the fibrillation of amyloid beta (1-40). J. Biol. Chem. 290, 815-
826
19. Suopanki, J., Gotz, C., Lutsch, G., Schiller, J., Harjes, P., Herrmann, A., and
Wanker, E. E. (2006) Interaction of huntingtin fragments with brain membranes--
clues to early dysfunction in Huntington's disease. J. Neurochem. 96, 870-884
20. Relini, A., Marano, N., and Gliozzi, A. (2014) Probing the interplay between
amyloidogenic proteins and membranes using lipid monolayers and bilayers.
Adv. Colloid Interface Sci. 207, 81-92
21. George, J. M. (2002) The synucleins. Genome Biol. 3, REVIEWS3002
22. Clayton, D. F., and George, J. M. (1998) The synucleins: a family of proteins
involved in synaptic function, plasticity, neurodegeneration and disease. Trends
Neurosci. 21, 249-254
23. Baba, M., Nakajo, S., Tu, P. H., Tomita, T., Nakaya, K., Lee, V. M., Trojanowski, J.
Q., and Iwatsubo, T. (1998) Aggregation of alpha-synuclein in Lewy bodies of
sporadic Parkinson's disease and dementia with Lewy bodies. Am. J. Pathol. 152,
879-884
24. Spillantini, M. G., Crowther, R. A., Jakes, R., Cairns, N. J., Lantos, P. L., and
Goedert, M. (1998) Filamentous alpha-synuclein inclusions link multiple system
atrophy with Parkinson's disease and dementia with Lewy bodies. Neurosci. Lett.
251, 205-208
25. Polymeropoulos, M. H., Lavedan, C., Leroy, E., Ide, S. E., Dehejia, A., Dutra, A.,
Pike, B., Root, H., Rubenstein, J., Boyer, R., Stenroos, E. S., Chandrasekharappa,
S., Athanassiadou, A., Papapetropoulos, T., Johnson, W. G., Lazzarini, A. M.,
Duvoisin, R. C., Di Iorio, G., Golbe, L. I., and Nussbaum, R. L. (1997) Mutation in
the alpha-synuclein gene identified in families with Parkinson's disease. Science
276, 2045-2047
25

26. Kruger, R., Kuhn, W., Muller, T., Woitalla, D., Graeber, M., Kosel, S., Przuntek,
H., Epplen, J. T., Schols, L., and Riess, O. (1998) Ala30Pro mutation in the gene
encoding alpha-synuclein in Parkinson's disease. Nat. Genet. 18, 106-108
27. Zarranz, J. J., Alegre, J., Gomez-Esteban, J. C., Lezcano, E., Ros, R., Ampuero, I.,
Vidal, L., Hoenicka, J., Rodriguez, O., Atares, B., Llorens, V., Gomez Tortosa, E.,
del Ser, T., Munoz, D. G., and de Yebenes, J. G. (2004) The new mutation, E46K,
of alpha-synuclein causes Parkinson and Lewy body dementia. Ann. Neurol. 55,
164-173
28. Singleton, A. B., Farrer, M., Johnson, J., Singleton, A., Hague, S., Kachergus, J.,
Hulihan, M., Peuralinna, T., Dutra, A., Nussbaum, R., Lincoln, S., Crawley, A.,
Hanson, M., Maraganore, D., Adler, C., Cookson, M. R., Muenter, M., Baptista,
M., Miller, D., Blancato, J., Hardy, J., and Gwinn-Hardy, K. (2003) alpha-
Synuclein locus triplication causes Parkinson's disease. Science 302, 841
29. Chartier-Harlin, M. C., Kachergus, J., Roumier, C., Mouroux, V., Douay, X.,
Lincoln, S., Levecque, C., Larvor, L., Andrieux, J., Hulihan, M., Waucquier, N.,
Defebvre, L., Amouyel, P., Farrer, M., and Destee, A. (2004) Alpha-synuclein
locus duplication as a cause of familial Parkinson's disease. Lancet 364, 1167-1169
30. Ibanez, P., Bonnet, A. M., Debarges, B., Lohmann, E., Tison, F., Pollak, P., Agid,
Y., Durr, A., and Brice, A. (2004) Causal relation between alpha-synuclein gene
duplication and familial Parkinson's disease. Lancet 364, 1169-1171
31. Irvine, G. B., El-Agnaf, O. M., Shankar, G. M., and Walsh, D. M. (2008) Protein
aggregation in the brain: the molecular basis for Alzheimer's and Parkinson's
diseases. Mol. Med. 14, 451-464
32. Volles, M. J., and Lansbury, P. T., Jr. (2003) Zeroing in on the pathogenic form of
alpha-synuclein and its mechanism of neurotoxicity in Parkinson's disease.
Biochemistry 42, 7871-7878
33. Zhu, M., Li, J., and Fink, A. L. (2003) The association of alpha-synuclein with
membranes affects bilayer structure, stability, and fibril formation. J. Biol. Chem.
278, 40186-40197
34. Nelson, R., Sawaya, M. R., Balbirnie, M., Madsen, A. O., Riekel, C., Grothe, R.,
and Eisenberg, D. (2005) Structure of the cross-beta spine of amyloid-like fibrils.
Nature 435, 773-778
35. Murphy, D. D., Rueter, S. M., Trojanowski, J. Q., and Lee, V. M. (2000) Synucleins
are developmentally expressed, and alpha-synuclein regulates the size of the
presynaptic vesicular pool in primary hippocampal neurons. J. Neurosci. 20, 3214-
3220
36. Chandra, S., Fornai, F., Kwon, H. B., Yazdani, U., Atasoy, D., Liu, X., Hammer, R.
E., Battaglia, G., German, D. C., Castillo, P. E., and Sudhof, T. C. (2004) Double-
knockout mice for alpha- and beta-synucleins: effect on synaptic functions. Proc.
Natl. Acad. Sci. U. S. A. 101, 14966-14971
26

37. Burre, J., Sharma, M., Tsetsenis, T., Buchman, V., Etherton, M. R., and Sudhof, T.
C. (2010) Alpha-synuclein promotes SNARE-complex assembly in vivo and in
vitro. Science 329, 1663-1667
38. Eisbach, S. E., and Outeiro, T. F. (2013) Alpha-synuclein and intracellular
trafficking: impact on the spreading of Parkinson's disease pathology. J. Mol.
Med. (Berl.) 91, 693-703
39. Bottner, M., Fricke, T., Muller, M., Barrenschee, M., Deuschl, G., Schneider, S. A.,
Egberts, J. H., Becker, T., Fritscher-Ravens, A., Ellrichmann, M., Schulz-Schaeffer,
W. J., and Wedel, T. (2015) Alpha-synuclein is associated with the synaptic
vesicle apparatus in the human and rat enteric nervous system. Brain Res.
40. Cooper, A. A., Gitler, A. D., Cashikar, A., Haynes, C. M., Hill, K. J., Bhullar, B.,
Liu, K., Xu, K., Strathearn, K. E., Liu, F., Cao, S., Caldwell, K. A., Caldwell, G. A.,
Marsischky, G., Kolodner, R. D., Labaer, J., Rochet, J. C., Bonini, N. M., and
Lindquist, S. (2006) Alpha-synuclein blocks ER-Golgi traffic and Rab1 rescues
neuron loss in Parkinson's models. Science 313, 324-328
41. Boassa, D., Berlanga, M. L., Yang, M. A., Terada, M., Hu, J., Bushong, E. A.,
Hwang, M., Masliah, E., George, J. M., and Ellisman, M. H. (2013) Mapping the
subcellular distribution of alpha-synuclein in neurons using genetically encoded
probes for correlated light and electron microscopy: implications for Parkinson's
disease pathogenesis. J. Neurosci. 33, 2605-2615
42. Pacheco, C. R., Morales, C. N., Ramirez, A. E., Munoz, F. J., Gallegos, S. S.,
Caviedes, P. A., Aguayo, L. G., and Opazo, C. M. (2015) Extracellular alpha-
synuclein alters synaptic transmission in brain neurons by perforating the
neuronal plasma membrane. J. Neurochem. 132, 731-741
43. Kahn, S. E., D'Alessio, D. A., Schwartz, M. W., Fujimoto, W. Y., Ensinck, J. W.,
Taborsky, G. J., Jr., and Porte, D., Jr. (1990) Evidence of cosecretion of islet
amyloid polypeptide and insulin by beta-cells. Diabetes 39, 634-638
44. Butler, P. C., Chou, J., Carter, W. B., Wang, Y. N., Bu, B. H., Chang, D., Chang, J.
K., and Rizza, R. A. (1990) Effects of meal ingestion on plasma amylin
concentration in NIDDM and nondiabetic humans. Diabetes 39, 752-756
45. Lutz, T. A. (2013) The interaction of amylin with other hormones in the control of
eating. Diabetes Obes. Metab. 15, 99-111
46. Cooper, G. J., Willis, A. C., Clark, A., Turner, R. C., Sim, R. B., and Reid, K. B.
(1987) Purification and characterization of a peptide from amyloid-rich
pancreases of type 2 diabetic patients. Proc. Natl. Acad. Sci. U. S. A. 84, 8628-8632
47. Janson, J., Soeller, W. C., Roche, P. C., Nelson, R. T., Torchia, A. J., Kreutter, D. K.,
and Butler, P. C. (1996) Spontaneous diabetes mellitus in transgenic mice
expressing human islet amyloid polypeptide. Proc. Natl. Acad. Sci. U. S. A. 93,
7283-7288
27

48. Gurlo, T., Ryazantsev, S., Huang, C. J., Yeh, M. W., Reber, H. A., Hines, O. J.,
O'Brien, T. D., Glabe, C. G., and Butler, P. C. (2010) Evidence for proteotoxicity in
beta cells in type 2 diabetes: toxic islet amyloid polypeptide oligomers form
intracellularly in the secretory pathway. Am. J. Pathol. 176, 861-869
49. Zhang, S., Liu, H., Chuang, C. L., Li, X., Au, M., Zhang, L., Phillips, A. R., Scott,
D. W., and Cooper, G. J. (2014) The pathogenic mechanism of diabetes varies
with the degree of overexpression and oligomerization of human amylin in the
pancreatic islet beta cells. FASEB J. 28, 5083-5096
50. Matveyenko, A. V., and Butler, P. C. (2006) Islet Amyloid Polypeptide (IAPP)
Transgenic Rodents as Models for Type 2 Diabetes. ILAR J. 47, 225-233
51. Magzoub, M., and Miranker, A. D. (2012) Concentration-dependent transitions
govern the subcellular localization of islet amyloid polypeptide. FASEB J. 26,
1228-1238
52. Westermark, P., Engstrom, U., Johnson, K. H., Westermark, G. T., and Betsholtz,
C. (1990) Islet amyloid polypeptide: pinpointing amino acid residues linked to
amyloid fibril formation. Proc. Natl. Acad. Sci. U. S. A. 87, 5036-5040
53. Brender, J. R., Hartman, K., Reid, K. R., Kennedy, R. T., and Ramamoorthy, A.
(2008) A single mutation in the nonamyloidogenic region of islet amyloid
polypeptide greatly reduces toxicity. Biochemistry 47, 12680-12688
54. Burke, K. A., Hensal, K. M., Umbaugh, C. S., Chaibva, M., and Legleiter, J. (2013)
Huntingtin disrupts lipid bilayers in a polyQ-length dependent manner. Biochim.
Biophys. Acta 1828, 1953-1961
55. Thapa, A., Vernon, B. C., De la Pena, K., Soliz, G., Moreno, H. A., Lopez, G. P.,
and Chi, E. Y. (2013) Membrane-mediated neuroprotection by curcumin from
amyloid-beta-peptide-induced toxicity. Langmuir 29, 11713-11723
56. Stefanovic, A. N., Stockl, M. T., Claessens, M. M., and Subramaniam, V. (2014)
alpha-Synuclein oligomers distinctively permeabilize complex model
membranes. FEBS J 281, 2838-2850
57. Janson, J., Ashley, R. H., Harrison, D., McIntyre, S., and Butler, P. C. (1999) The
mechanism of islet amyloid polypeptide toxicity is membrane disruption by
intermediate-sized toxic amyloid particles. Diabetes 48, 491-498
58. Knight, J. D., and Miranker, A. D. (2004) Phospholipid catalysis of diabetic
amyloid assembly. J. Mol. Biol. 341, 1175-1187
59. Butterfield, S. M., and Lashuel, H. A. (2010) Amyloidogenic protein-membrane
interactions: mechanistic insight from model systems. Angew. Chem. Int. Ed. Engl.
49, 5628-5654
60. (1993) A novel gene containing a trinucleotide repeat that is expanded and
unstable on Huntington's disease chromosomes. The Huntington's Disease
Collaborative Research Group. Cell 72, 971-983
28

61. Gusella, J. F., and MacDonald, M. E. (1995) Huntington's disease: CAG genetics
expands neurobiology. Curr. Opin. Neurobiol. 5, 656-662
62. Walker, F. O. (2007) Huntington's disease. Lancet 369, 218-228
63. Cortes, C. J., and La Spada, A. R. (2015) Autophagy in polyglutamine disease:
Imposing order on disorder or contributing to the chaos? Mol. Cell. Neurosci.
64. Imarisio, S., Carmichael, J., Korolchuk, V., Chen, C. W., Saiki, S., Rose, C.,
Krishna, G., Davies, J. E., Ttofi, E., Underwood, B. R., and Rubinsztein, D. C.
(2008) Huntington's disease: from pathology and genetics to potential therapies.
Biochem. J. 412, 191-209
65. van der Burg, J. M., Bjorkqvist, M., and Brundin, P. (2009) Beyond the brain:
widespread pathology in Huntington's disease. Lancet Neurol. 8, 765-774
66. Markianos, M., Panas, M., Kalfakis, N., and Vassilopoulos, D. (2005) Plasma
testosterone in male patients with Huntington's disease: relations to severity of
illness and dementia. Ann. Neurol. 57, 520-525
67. Van Raamsdonk, J. M., Murphy, Z., Selva, D. M., Hamidizadeh, R., Pearson, J.,
Petersen, A., Bjorkqvist, M., Muir, C., Mackenzie, I. R., Hammond, G. L., Vogl, A.
W., Hayden, M. R., and Leavitt, B. R. (2007) Testicular degeneration in
Huntington disease. Neurobiol. Dis. 26, 512-520
68. Djousse, L., Knowlton, B., Cupples, L. A., Marder, K., Shoulson, I., and Myers, R.
H. (2002) Weight loss in early stage of Huntington's disease. Neurology 59, 1325-
1330
69. Hamilton, J. M., Wolfson, T., Peavy, G. M., Jacobson, M. W., Corey-Bloom, J., and
Huntington Study, G. (2004) Rate and correlates of weight change in
Huntington's disease. J. Neurol. Neurosurg. Psychiatry 75, 209-212
70. Aziz, N. A., van der Burg, J. M., Landwehrmeyer, G. B., Brundin, P., Stijnen, T.,
Group, E. S., and Roos, R. A. (2008) Weight loss in Huntington disease increases
with higher CAG repeat number. Neurology 71, 1506-1513
71. Goodman, A. O., Murgatroyd, P. R., Medina-Gomez, G., Wood, N. I., Finer, N.,
Vidal-Puig, A. J., Morton, A. J., and Barker, R. A. (2008) The metabolic profile of
early Huntington's disease--a combined human and transgenic mouse study.
Exp. Neurol. 210, 691-698
72. Simpson, S. A. (2007) Late stage care in Huntington's disease. Brain Res. Bull. 72,
179-181
73. Mangiarini, L., Sathasivam, K., Seller, M., Cozens, B., Harper, A., Hetherington,
C., Lawton, M., Trottier, Y., Lehrach, H., Davies, S. W., and Bates, G. P. (1996)
Exon 1 of the HD gene with an expanded CAG repeat is sufficient to cause a
progressive neurological phenotype in transgenic mice. Cell 87, 493-506
74. Sathasivam, K., Neueder, A., Gipson, T. A., Landles, C., Benjamin, A. C.,
Bondulich, M. K., Smith, D. L., Faull, R. L., Roos, R. A., Howland, D., Detloff, P.
J., Housman, D. E., and Bates, G. P. (2013) Aberrant splicing of HTT generates the
29

pathogenic exon 1 protein in Huntington disease. Proc. Natl. Acad. Sci. U. S. A.
110, 2366-2370
75. Lunkes, A., Lindenberg, K. S., Ben-Haiem, L., Weber, C., Devys, D.,
Landwehrmeyer, G. B., Mandel, J. L., and Trottier, Y. (2002) Proteases acting on
mutant huntingtin generate cleaved products that differentially build up
cytoplasmic and nuclear inclusions. Mol. Cell 10, 259-269
76. Kim, M. W., Chelliah, Y., Kim, S. W., Otwinowski, Z., and Bezprozvanny, I.
(2009) Secondary structure of Huntingtin amino-terminal region. Structure 17,
1205-1212
77. Arndt, J. R., Brown, R. J., Burke, K. A., Legleiter, J., and Valentine, S. J. (2015)
Lysine residues in the N-terminal huntingtin amphipathic alpha-helix play a key
role in peptide aggregation. J. Mass Spectrom. 50, 117-126
78. Kokona, B., Rosenthal, Z. P., and Fairman, R. (2014) Role of the coiled-coil
structural motif in polyglutamine aggregation. Biochemistry 53, 6738-6746
79. Wetzel, R. (2012) Physical chemistry of polyglutamine: intriguing tales of a
monotonous sequence. J. Mol. Biol. 421, 466-490
80. Jayaraman, M., Kodali, R., Sahoo, B., Thakur, A. K., Mayasundari, A., Mishra, R.,
Peterson, C. B., and Wetzel, R. (2012) Slow amyloid nucleation via alpha-helix-
rich oligomeric intermediates in short polyglutamine-containing huntingtin
fragments. J. Mol. Biol. 415, 881-899
81. Jayaraman, M., Mishra, R., Kodali, R., Thakur, A. K., Koharudin, L. M.,
Gronenborn, A. M., and Wetzel, R. (2012) Kinetically competing huntingtin
aggregation pathways control amyloid polymorphism and properties.
Biochemistry 51, 2706-2716
82. Thakur, A. K., Jayaraman, M., Mishra, R., Thakur, M., Chellgren, V. M., Byeon, I.
J., Anjum, D. H., Kodali, R., Creamer, T. P., Conway, J. F., Gronenborn, A. M.,
and Wetzel, R. (2009) Polyglutamine disruption of the huntingtin exon 1 N
terminus triggers a complex aggregation mechanism. Nat. Struct. Mol. Biol. 16,
380-389
83. Crick, S. L., Ruff, K. M., Garai, K., Frieden, C., and Pappu, R. V. (2013)
Unmasking the roles of N- and C-terminal flanking sequences from exon 1 of
huntingtin as modulators of polyglutamine aggregation. Proc. Natl. Acad. Sci. U.
S. A. 110, 20075-20080
84. Zeitlin, S., Liu, J. P., Chapman, D. L., Papaioannou, V. E., and Efstratiadis, A.
(1995) Increased apoptosis and early embryonic lethality in mice nullizygous for
the Huntington's disease gene homologue. Nat. Genet. 11, 155-163
85. Steffan, J. S. (2010) Does Huntingtin play a role in selective macroautophagy? Cell
cycle 9, 3401-3413
86. Martin, D. D., Ladha, S., Ehrnhoefer, D. E., and Hayden, M. R. (2015) Autophagy
in Huntington disease and huntingtin in autophagy. Trends Neurosci. 38, 26-35
30

87. Ochaba, J., Lukacsovich, T., Csikos, G., Zheng, S., Margulis, J., Salazar, L., Mao,
K., Lau, A. L., Yeung, S. Y., Humbert, S., Saudou, F., Klionsky, D. J., Finkbeiner,
S., Zeitlin, S. O., Marsh, J. L., Housman, D. E., Thompson, L. M., and Steffan, J. S.
(2015) Potential function for the Huntingtin protein as a scaffold for selective
autophagy. Proc. Natl. Acad. Sci. U. S. A. 111, 16889-16894
88. Keryer, G., Pineda, J. R., Liot, G., Kim, J., Dietrich, P., Benstaali, C., Smith, K.,
Cordelieres, F. P., Spassky, N., Ferrante, R. J., Dragatsis, I., and Saudou, F. (2011)
Ciliogenesis is regulated by a huntingtin-HAP1-PCM1 pathway and is altered in
Huntington disease. J. Clin. Invest. 121, 4372-4382
89. Godin, J. D., Colombo, K., Molina-Calavita, M., Keryer, G., Zala, D., Charrin, B.
C., Dietrich, P., Volvert, M. L., Guillemot, F., Dragatsis, I., Bellaiche, Y., Saudou,
F., Nguyen, L., and Humbert, S. (2010) Huntingtin is required for mitotic spindle
orientation and mammalian neurogenesis. Neuron 67, 392-406
90. Smith, R., Bacos, K., Fedele, V., Soulet, D., Walz, H. A., Obermuller, S., Lindqvist,
A., Bjorkqvist, M., Klein, P., Onnerfjord, P., Brundin, P., Mulder, H., and Li, J. Y.
(2009) Mutant huntingtin interacts with {beta}-tubulin and disrupts vesicular
transport and insulin secretion. Hum. Mol. Genet. 18, 3942-3954
91. Steffan, J. S., Kazantsev, A., Spasic-Boskovic, O., Greenwald, M., Zhu, Y. Z.,
Gohler, H., Wanker, E. E., Bates, G. P., Housman, D. E., and Thompson, L. M.
(2000) The Huntington's disease protein interacts with p53 and CREB-binding
protein and represses transcription. Proc. Natl. Acad. Sci. U. S. A. 97, 6763-6768
92. Plomann, M., Lange, R., Vopper, G., Cremer, H., Heinlein, U. A., Scheff, S.,
Baldwin, S. A., Leitges, M., Cramer, M., Paulsson, M., and Barthels, D. (1998)
PACSIN, a brain protein that is upregulated upon differentiation into neuronal
cells. Eur. J. Biochem. 256, 201-211
93. Cornett, J., Cao, F., Wang, C. E., Ross, C. A., Bates, G. P., Li, S. H., and Li, X. J.
(2005) Polyglutamine expansion of huntingtin impairs its nuclear export. Nat.
Genet. 37, 198-204
94. Atwal, R. S., Xia, J., Pinchev, D., Taylor, J., Epand, R. M., and Truant, R. (2007)
Huntingtin has a membrane association signal that can modulate huntingtin
aggregation, nuclear entry and toxicity. Hum. Mol. Genet. 16, 2600-2615
95. Der-Sarkissian, A., Jao, C. C., Chen, J., and Langen, R. (2003) Structural
organization of alpha-synuclein fibrils studied by site-directed spin labeling. J.
Biol. Chem. 278, 37530-37535
96. Schneider, R., Schumacher, M. C., Mueller, H., Nand, D., Klaukien, V., Heise, H.,
Riedel, D., Wolf, G., Behrmann, E., Raunser, S., Seidel, R., Engelhard, M., and
Baldus, M. (2011) Structural characterization of polyglutamine fibrils by solid-
state NMR spectroscopy. J. Mol. Biol. 412, 121-136
31

97. Bedrood, S., Li, Y., Isas, J. M., Hegde, B. G., Baxa, U., Haworth, I. S., and Langen,
R. (2012) Fibril structure of human islet amyloid polypeptide. J. Biol. Chem. 287,
5235-5241
98. Bugg, C. W., Isas, J. M., Fischer, T., Patterson, P. H., and Langen, R. (2012)
Structural features and domain organization of huntingtin fibrils. J. Biol. Chem.
287, 31739-31746
99. Xu, W., Su, H., Zhang, J. Z., and Mu, Y. (2012) Molecular dynamics simulation
study on the molecular structures of the amylin fibril models. J. Phys. Chem. B
116, 13991-13999
100. Rothlein, C., Miettinen, M. S., Borwankar, T., Burger, J., Mielke, T., Kumke, M.
U., and Ignatova, Z. (2014) Architecture of polyglutamine-containing fibrils from
time-resolved fluorescence decay. J. Biol. Chem. 289, 26817-26828
101. Langkilde, A. E., Morris, K. L., Serpell, L. C., Svergun, D. I., and Vestergaard, B.
(2015) The architecture of amyloid-like peptide fibrils revealed by X-ray
scattering, diffraction and electron microscopy. Acta Crystallogr. D Biol.
Crystallogr. 71, 882-895
102. Michalek, M., Salnikov, E. S., and Bechinger, B. (2013) Structure and topology of
the huntingtin 1-17 membrane anchor by a combined solution and solid-state
NMR approach. Biophys. J. 105, 699-710
103. Davidson, W. S., Jonas, A., Clayton, D. F., and George, J. M. (1998) Stabilization
of alpha-synuclein secondary structure upon binding to synthetic membranes. J.
Biol. Chem. 273, 9443-9449
104. Weinreb, P. H., Zhen, W., Poon, A. W., Conway, K. A., and Lansbury, P. T., Jr.
(1996) NACP, a protein implicated in Alzheimer's disease and learning, is
natively unfolded. Biochemistry 35, 13709-13715
105. Eliezer, D., Kutluay, E., Bussell, R., Jr., and Browne, G. (2001) Conformational
properties of alpha-synuclein in its free and lipid-associated states. J. Mol. Biol.
307, 1061-1073
106. Bussell, R., Jr., and Eliezer, D. (2003) A structural and functional role for 11-mer
repeats in alpha-synuclein and other exchangeable lipid binding proteins. J. Mol.
Biol. 329, 763-778
107. Jao, C. C., Hegde, B. G., Chen, J., Haworth, I. S., and Langen, R. (2008) Structure
of membrane-bound alpha-synuclein from site-directed spin labeling and
computational refinement. Proc. Natl. Acad. Sci. U. S. A. 105, 19666-19671
108. Jao, C. C., Der-Sarkissian, A., Chen, J., and Langen, R. (2004) Structure of
membrane-bound alpha-synuclein studied by site-directed spin labeling. Proc.
Natl. Acad. Sci. U. S. A. 101, 8331-8336
109. Apostolidou, M., Jayasinghe, S. A., and Langen, R. (2008) Structure of alpha-
helical membrane-bound human islet amyloid polypeptide and its implications
for membrane-mediated misfolding. J. Biol. Chem. 283, 17205-17210
32

110. Patil, S. M., Xu, S., Sheftic, S. R., and Alexandrescu, A. T. (2009) Dynamic alpha-
helix structure of micelle-bound human amylin. J. Biol. Chem. 284, 11982-11991
111. Nanga, R. P., Brender, J. R., Vivekanandan, S., and Ramamoorthy, A. (2011)
Structure and membrane orientation of IAPP in its natively amidated form at
physiological pH in a membrane environment. Biochim. Biophys. Acta 1808, 2337-
2342
112. Feix, J. B., and Klug, C. S. (1998) Site-Directed Spin Labeling of Membrane
Proteins and Peptide-Membrane Interactions. in Biological Magnetic Resonance,
Volume 14: Spin Labeling: The Next Millennium (Berliner, L. J. ed.), Plenum Press,
New York. pp 251-281
113. Margittai, M., and Langen, R. (2006) Spin labeling analysis of amyloids and other
protein aggregates. Methods Enzymol. 413, 122-139
114. Weber, R. T., Jiang, J., and Barr, D. P. (1998) EMX User's Manual, Bruker
Instruments, EPR Division, Billerica, MA
115. Persson, M., Harbridge, J. R., Hammarstrom, P., Mitri, R., Martensson, L. G.,
Carlsson, U., Eaton, G. R., and Eaton, S. S. (2001) Comparison of electron
paramagnetic resonance methods to determine distances between spin labels on
human carbonic anhydrase II. Biophys. J. 80, 2886-2897
116. Marsh, D. (2010) Spin-Label EPR for Determining Polarity and Proticity in
Biomolecular Assemblies: Transmembrane Profiles. Applied magnetic resonance 37,
435-454
117. Furlan, S., La Penna, G., Appelhans, D., Cangiotti, M., Ottaviani, M. F., and
Danani, A. (2014) Combined EPR and molecular modeling study of PPI
dendrimers interacting with copper ions: effect of generation and maltose
decoration. J. Phys. Chem. B 118, 12098-12111
118. Mustafi, D., Telser, J., and Makinen, M. W. (1992) Molecular Geometry of
Vanadyl-Adenine Nucleotide Complexes Determined by EPR, ENDOR, and
Molecular Modeling. J. Am. Chem. Soc. 114, 6219-6226
119. Sparr, E., Engel, M. F., Sakharov, D. V., Sprong, M., Jacobs, J., de Kruijff, B.,
Hoppener, J. W., and Killian, J. A. (2004) Islet amyloid polypeptide-induced
membrane leakage involves uptake of lipids by forming amyloid fibers. FEBS
Lett. 577, 117-120
120. Zhao, H., Tuominen, E. K., and Kinnunen, P. K. (2004) Formation of amyloid
fibers triggered by phosphatidylserine-containing membranes. Biochemistry 43,
10302-10307
121. Engel, M. F., Khemtemourian, L., Kleijer, C. C., Meeldijk, H. J., Jacobs, J., Verkleij,
A. J., de Kruijff, B., Killian, J. A., and Hoppener, J. W. (2008) Membrane damage
by human islet amyloid polypeptide through fibril growth at the membrane.
Proc. Natl. Acad. Sci. U. S. A. 105, 6033-6038
33

122. Milanesi, L., Sheynis, T., Xue, W. F., Orlova, E. V., Hellewell, A. L., Jelinek, R.,
Hewitt, E. W., Radford, S. E., and Saibil, H. R. (2012) Direct three-dimensional
visualization of membrane disruption by amyloid fibrils. Proc. Natl. Acad. Sci. U.
S. A. 109, 20455-20460
123. Lashuel, H. A., Petre, B. M., Wall, J., Simon, M., Nowak, R. J., Walz, T., and
Lansbury, P. T., Jr. (2002) Alpha-synuclein, especially the Parkinson's disease-
associated mutants, forms pore-like annular and tubular protofibrils. J. Mol. Biol.
322, 1089-1102
124. de Planque, M. R., Raussens, V., Contera, S. A., Rijkers, D. T., Liskamp, R. M.,
Ruysschaert, J. M., Ryan, J. F., Separovic, F., and Watts, A. (2007) beta-Sheet
structured beta-amyloid(1-40) perturbs phosphatidylcholine model membranes.
J. Mol. Biol. 368, 982-997
125. Quist, A., Doudevski, I., Lin, H., Azimova, R., Ng, D., Frangione, B., Kagan, B.,
Ghiso, J., and Lal, R. (2005) Amyloid ion channels: a common structural link for
protein-misfolding disease. Proc. Natl. Acad. Sci. U. S. A. 102, 10427-10432
126. Kayed, R., Sokolov, Y., Edmonds, B., McIntire, T. M., Milton, S. C., Hall, J. E., and
Glabe, C. G. (2004) Permeabilization of lipid bilayers is a common conformation-
dependent activity of soluble amyloid oligomers in protein misfolding diseases.
J. Biol. Chem. 279, 46363-46366
127. Sokolov, Y., Kozak, J. A., Kayed, R., Chanturiya, A., Glabe, C., and Hall, J. E.
(2006) Soluble amyloid oligomers increase bilayer conductance by altering
dielectric structure. J. Gen. Physiol. 128, 637-647
128. Haataja, L., Gurlo, T., Huang, C. J., and Butler, P. C. (2008) Islet amyloid in type 2
diabetes, and the toxic oligomer hypothesis. Endocr. Rev. 29, 303-316
129. Jayasinghe, S. A., and Langen, R. (2007) Membrane interaction of islet amyloid
polypeptide. Biochim. Biophys. Acta 1768, 2002-2009
130. Bechinger, B., and Lohner, K. (2006) Detergent-like actions of linear amphipathic
cationic antimicrobial peptides. Biochim. Biophys. Acta 1758, 1529-1539
131. Rausch, J. M., Marks, J. R., Rathinakumar, R., and Wimley, W. C. (2007) Beta-
sheet pore-forming peptides selected from a rational combinatorial library:
mechanism of pore formation in lipid vesicles and activity in biological
membranes. Biochemistry 46, 12124-12139
132. Zimmerberg, J., and Kozlov, M. M. (2006) How proteins produce cellular
membrane curvature. Nat. Rev. Mol. Cell Biol. 7, 9-19
133. Campelo, F., McMahon, H. T., and Kozlov, M. M. (2008) The hydrophobic
insertion mechanism of membrane curvature generation by proteins. Biophys. J.
95, 2325-2339
134. Ford, M. G., Mills, I. G., Peter, B. J., Vallis, Y., Praefcke, G. J., Evans, P. R., and
McMahon, H. T. (2002) Curvature of clathrin-coated pits driven by epsin. Nature
419, 361-366
34

135. Takei, K., Slepnev, V. I., Haucke, V., and De Camilli, P. (1999) Functional
partnership between amphiphysin and dynamin in clathrin-mediated
endocytosis. Nat. Cell Biol. 1, 33-39
136. Farsad, K., Ringstad, N., Takei, K., Floyd, S. R., Rose, K., and De Camilli, P. (2001)
Generation of high curvature membranes mediated by direct endophilin bilayer
interactions. J. Cell Biol. 155, 193-200
137. Weinstein, J. N., Ralston, E., Leserman, L. D., Klausner, R. D., Dragsten, P.,
Henkart, P., and Blumenthal, R. (1984) Self-Quenching of Carboxyfluorescein
Fluorescence: Uses in Studying Liposome Stability and Liposome-Cell
Interaction. in Liposome Technology, Volume III: Targeted Drug Delivery and
Biological Interaction (Gregoriandis, G. ed.), CRC Press, Inc., Boca Raton, FL. pp
183-204
138. Last, N. B., Rhoades, E., and Miranker, A. D. (2011) Islet amyloid polypeptide
demonstrates a persistent capacity to disrupt membrane integrity. Proc. Natl.
Acad. Sci. U. S. A. 108, 9460-9465
139. Varkey, J., Isas, J. M., Mizuno, N., Jensen, M. B., Bhatia, V. K., Jao, C. C., Petrlova,
J., Voss, J. C., Stamou, D. G., Steven, A. C., and Langen, R. (2010) Membrane
curvature induction and tubulation are common features of synucleins and
apolipoproteins. J. Biol. Chem. 285, 32486-32493
140. Brender, J. R., Lee, E. L., Cavitt, M. A., Gafni, A., Steel, D. G., and Ramamoorthy,
A. (2008) Amyloid fiber formation and membrane disruption are separate
processes localized in two distinct regions of IAPP, the type-2-diabetes-related
peptide. J. Am. Chem. Soc. 130, 6424-6429
141. Brender, J. R., Lee, E. L., Hartman, K., Wong, P. T., Ramamoorthy, A., Steel, D.
G., and Gafni, A. (2011) Biphasic effects of insulin on islet amyloid polypeptide
membrane disruption. Biophys. J. 100, 685-692
142. Heyl, D. L., Osborne, J. M., Pamarthy, S., Samisetti, S., Gray, A. W., Jayaprakash,
A., Konda, S., Brown, D. J., Miller, S. R., Eizadkhah, R., and Milletti, M. C. (2010)
Liposome damage and modeling of fragments of human islet amyloid
polypeptide (IAPP) support a two-step model of membrane destruction. Int. J.
Pept. Res. Ther. 16, 43-54
143. Knight, J. D., Hebda, J. A., and Miranker, A. D. (2006) Conserved and cooperative
assembly of membrane-bound alpha-helical states of islet amyloid polypeptide.
Biochemistry 45, 9496-9508
144. Pandey, A. P., Haque, F., Rochet, J. C., and Hovis, J. S. (2011) alpha-Synuclein-
induced tubule formation in lipid bilayers. J. Phys. Chem. B 115, 5886-5893
145. Truant, R., Atwal, R. S., Desmond, C., Munsie, L., and Tran, T. (2008)
Huntington's disease: revisiting the aggregation hypothesis in polyglutamine
neurodegenerative diseases. FEBS J 275, 4252-4262
35

146. Caron, N. S., Desmond, C. R., Xia, J., and Truant, R. (2013) Polyglutamine
domain flexibility mediates the proximity between flanking sequences in
huntingtin. Proc. Natl. Acad. Sci. U. S. A. 110, 14610-14615
147. Fodale, V., Kegulian, N. C., Verani, M., Cariulo, C., Azzollini, L., Petricca, L.,
Daldin, M., Boggio, R., Padova, A., Kuhn, R., Pacifici, R., Macdonald, D.,
Schoenfeld, R. C., Park, H., Isas, J. M., Langen, R., Weiss, A., and Caricasole, A.
(2014) Polyglutamine- and temperature-dependent conformational rigidity in
mutant huntingtin revealed by immunoassays and circular dichroism
spectroscopy. PLoS One 9, e112262

36

Chapter 2

Measuring Exposure of Different α-Synuclein Residues to Phospholipid Membranes
Using a Fluorescent Probe

Partly extracted from Mizuno, N., Varkey, J., Kegulian, N. C., Hegde, B. G., Cheng, N.,
Langen, R., and Steven, A. C. (2012) Remodeling of lipid vesicles into cylindrical
micelles by α-synuclein in an extended α-helical conformation. J. Biol. Chem. 287, 29301-
29311.
37

2.1 Introduction
As a plethora of studies implicate α-synuclein in several physiological and cytotoxic
roles that involve direct interaction with cellular membranes (as summarized in
Chapter 1), studies focusing on α-synuclein-
membrane interactions are crucial. In view of prior
and concurrent experiments by my colleagues
showing α-synuclein to induce loss of light
scattering by multilamellar vesicles (MLVs) made
up of phosphatidylglycerol with varying acyl
chains, I used an environment-sensitive probe to
identify the α-synuclein residues that bound the
vesicles under the same conditions that the protein
caused them to clear. Different studies have shown α-synuclein to adopt a broken
helical structure on micelles (1), an extended helical structure on large unilamellar
vesicles (LUVs) (2,3), and a combination of broken and extended helices on small
unilamellar vesicles (SUVs) (4). (Interestingly, since my study, our group has also
uncovered the ability of α-synuclein in the extended conformation to remodel vesicles
into small, lipid-containing nanoparticles on which α-synuclein binds in the broken
helical conformation [5]. This and the SUV study showcase the high propensity of this
protein to interconvert between helical structures according to the shape of the lipid to
which it is bound.) I wanted to probe different residues on α-synuclein in order to
identify which of these helical structures it takes on when inducing the clearance we
observed.
The probe I utilized was N,N'-dimethyl-N-(iodoacetyl)-N'-(7-nitrobenz-2-oxa-1,3-
diazol-4-yl)ethylenediamine (IANBD) (Fig. 2-1), a fluorophore whose fluorescence
emission spectrum is enhanced when in a lipidic or other hydrophobic environment
(6,7), such that fluorescence signal is both higher and blue-shifted. Fluorophores usually

Fig. 2-1. Molecular structure of
the environment-sensitive
fluorophore IANBD.

38

lose energy after excitation and emit at a lower energy than their energy of excitation, in
a phenomenon called the Stokes shift (8). A blue shift under one condition as compared
to another signifies that the fluorophore in question has lost less energy under the
former condition and is therefore emitting at a higher energy than under the latter
condition, and for polarity-sensitive fluorophores, a hydrophobic environment such as
that provided by a lipid membrane is a common cause of blue fluorescence shifting
relative to fluorescence of the same molecule in a hydrophilic environment (9,10).
Since IANBD is thiol-reactive (11), it selective makes a covalent bond at the thiol
group on each cysteine residue of a protein. Taking advantage of the fact that human α-
synuclein is natively cysteine-free (Fig. 1-1A), I labeled single cysteine mutants of α-
synuclein with this fluorophore and measured the resulting fluorescence spectra in the
presence and absence
of MLVs of some of
the same compositions
utilized in the
experiments showing
loss of light scatter.
2.2 Methods
Materials
IANBD amide was
purchased as a
powder from
Molecular Probes
(Eugene, OR), kept at –
20°C in the dark, and
dissolved in dimethyl sulfoxide prior to use. 1-palmitoyl-2-oleoyl-sn-glycero-3-
[phospho-RAC-(1-glycerol)] (POPG), 1,2-dioleoyl-sn-glycero-3-[phospho-RAC-(1-

Fig. 2-2. The four phospholipids utilized in this membrane
study. Membranes were composed of lipids of negatively
charged compositions due to their common phosphatidylglycerol
headgroup. Acyl chains varied in length and saturation.

39

glycerol)] (DOPG), 1,2-dimyristoyl-sn-glycero-3-[phospho-RAC-(1-glycerol)] (DMPG),
and 1,2-diarachidonoyl-sn-glycero-3-[phospho-RAC-(1-glycerol)] (DAPG) (Fig. 2-2)
purchased from Avanti Polar Lipids Inc. (Alabaster, AL) as solutions in chloroform
were transferred to glass tubes and the films dried under N2 gas and then stored 6
hours to overnight under vacuum in a desiccator. MLVs were generated from lipid
films by adding 20 mM 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid
(HEPES), pH 7.4, 100 mM NaCl, and vortexing for ≥ 1 minute to resuspend the lipids in
this buffer.
Preparation and labeling of α-synuclein
α-Synuclein was expressed and purified as previously described (12). Briefly, lysate
from α-synuclein-expressing Escherichia coli BL21(DE3)pLys-S cells was boiled and the
supernatant subsequently acid-precipitated. The resulting supernatant was passed
through anion exchange columns and eluted with a 0–1.0 M NaCl gradient. All buffers
used contained 5 mM dithiothreitol (DTT) to prevent dimerization via disulfide bridge
formation between cysteine residues. Different single cysteine mutants were buffer-
exchanged through PD-10 columns (GE Healthcare) into 20 mM HEPES, pH 7.4, 100
mM NaCl, in order to separate them from the DTT. IANBD was immediately added at a
ten-fold molar excess, and the protein sample was rotated at ~40 rpm either for 2 hours
at room temperature or overnight at 4°C. To separate labeled α-synuclein from free
label, the sample was again passed through a PD-10 column. Samples were
concentrated by centrifugation at 5000 rcf in Millipore filter units (3000 MWCO).
Absorbance spectra were measured from 250 to 500 nm in a Jasco V-550
spectrophotometer with a 1-cm-path-length quartz cuvette. Labeling efficiency was
assessed with the following calculation:

(Eq. 2-1)

C
f
A
maxf
ε
280p

C
p
A
280
ε
maxf
– A
maxf
ε
280f

= × 100%
40

where Amaxf is maximal absorbance of the fluorophore (478 nm for IANBD), ε280p is the
molar extinction coefficient of the protein (number of tyrosine residues × 1280 M
-1
cm
-1
),
and εmaxf and ε280f are the molar extinction coefficients of the fluorophore at maximal
absorbance and at 280 nm (18,492 and 1250 M
-1
cm
-1
for IANBD, respectively).
Fluorescence measurements
Emission spectra were taken from 500 to 600 nm in a Jasco FP-6500 spectrofluorometer
with excitation wavelength set at 478 nm and excitation and emission slit widths of 3
nm and 5 nm, respectively. First, a spectrum was taken for 15 µM protein alone. MLVs
were promptly added and mixed in by pipetting up and down to a single phospholipid
concentration of 300 µM. After approximately 1 hour to allow for maximal binding, the
protein-with-lipid spectrum was taken.
Spectra were normalized by dividing every fluorescence value for both spectra of
each α-synuclein mutant by the maximal fluorescence of the protein-alone spectrum for
that particular mutant. This method set maximal fluorescence of protein alone to 1 and
allowed for direct assessment of fluorescence change upon addition of vesicles. Blue
shifts were measured as the difference between the wavelength at maximal fluorescence
intensity in the absence of lipid (~543 nm on average) and the wavelength at maximal
fluorescence intensity in the presence of lipid.
2.3 Results and Discussion
MLVs composed of POPG, which contains one cis-double bond in one acyl chain and
whose other acyl chain is saturated (Fig. 2-2), were added to α-synuclein labeled with
IANBD at solvent-exposed (residues 31 and 76) and membrane-exposed (residues 26,
48, 52, 63, and 70) sites in the α-helix, as well as sites further in the N-terminus (residues
5 and 8) and sites in the non-membrane-binding C-terminus (residues 100, 124, and 136)
(refer to Fig. 1-2C for residue positions on the helical wheel). The resulting spectra (Fig.
2-3A) showed markedly enhanced fluorescence for residues located in the N-terminus
(even those below position 9, which according to our EPR study marks the start of the
41

helical region [3]) compared to the C-terminal residues, which only fluoresced close to
baseline. All residues within the α-helix, regardless of the direction each is known to
face, showed significant fluorescence increases, suggesting that IANBD is not a
sufficiently specific probe for studying α-synuclein residue orientation. However, there
was a trend toward less of a fluorescence increase for residues in the C-terminal part of
the α-helix (Fig. 2-3A), corresponding to a recent NMR study showing that, at low
protein-to-lipid ratios such as that used here, the population of α-synuclein molecules
includes pools of
α-synuclein
helices only far-
N-terminal parts
of which are
membrane-bound
(13). In addition,
while residues
124 and 136
showed spectra
as low as
baseline, the C-
terminal residue
100 showed a
mild increase
(Fig. 2-3A),
suggesting some
interplay between
the part of the C-

Fig. 2-3. Fluorescence spectroscopy indicates the formation of a
continuous α-synuclein helix bound to membranes. Normalized
fluorescence spectra for 15 µM IANBD labeled α-synuclein and 300 µM (A)
POPG, (B) DOPG, (C) DAPG, and (D) DMPG MLVs. Black, green, and gray
broken lines represent α-synuclein that was IANBD-labeled at position 26,
48, or 11, respectively, before addition of lipid. All other lines represent α-
synuclein-IANBD ~1 hour after addition of the indicated lipid, where
labeling was at the following positions: turquoise dash-dot line, 136; red
dotted line, 124; green broken line, 100; turquoise solid line, 86; yellow solid line,
80; orange broken line, 76; purple broken line, 70; blue solid line, 63; purple solid
line, 52; green solid line, 48; red solid line, 31; black solid line, 26; magenta solid
line, 22; gray solid line, 11; orange solid line, 8; and jute solid line, 5. Some
traces from part (A) were published in Mizuno et al., 2012.

42

terminus close to the α-helix and POPG. Lipid compositions containing greater
numbers of acyl double bonds, namely DOPG and DAPG (Fig. 2-2), yielded spectra
showing similar trends but more apparent difficulty in binding by residues in the C-
terminal part of the α-helix and by C-terminal residues close to the α-helix (Fig. 2-3BC).

Fig. 2-4. Blue or red shifts in α-synuclein-IANBD fluorescence in the presence of vesicles of
the indicated lipid compositions.
43

In contrast, spectra from α-synuclein and DMPG, a completely saturated lipid (Fig. 2-2),
showed no discrimination between N- and C-terminal parts of the helix (Fig. 2-3D).
Blue shifts in environment-sensitive fluorescence are another way to quantify
residue-by-residue membrane binding by a probe-labeled protein. Using blue shift
length as the criterion for binding, we no longer observe a trend toward lower binding
by C-terminal residues within the α-helix (Fig. 2-4). However, residue 100 did show a
slight blue shift in the presence of POPG (Fig. 2-4, left), again mildly suggesting
interactions with the membrane, and no blue shift (even a 2-nm red shift) in the
presence of DOPG (Fig. 2-4, top right), suggesting greater abrogation by unsaturated
acyl chains of interactions between C-terminal residues and the membrane.
These experiments have lent a piece of insight into the structures taken on by α-
synuclein while it induces remodeling of negatively charged membranes.

44

2.4 References
1. Bussell, R., Jr., and Eliezer, D. (2003) A structural and functional role for 11-mer
repeats in alpha-synuclein and other exchangeable lipid binding proteins. J. Mol.
Biol. 329, 763-778
2. Jao, C. C., Der-Sarkissian, A., Chen, J., and Langen, R. (2004) Structure of
membrane-bound alpha-synuclein studied by site-directed spin labeling. Proc.
Natl. Acad. Sci. U. S. A. 101, 8331-8336
3. Jao, C. C., Hegde, B. G., Chen, J., Haworth, I. S., and Langen, R. (2008) Structure
of membrane-bound alpha-synuclein from site-directed spin labeling and
computational refinement. Proc. Natl. Acad. Sci. U. S. A. 105, 19666-19671
4. Lokappa, S. B., and Ulmer, T. S. (2011) Alpha-synuclein populates both elongated
and broken helix states on small unilamellar vesicles. J. Biol. Chem. 286, 21450-
21457
5. Varkey, J., Mizuno, N., Hegde, B. G., Cheng, N., Steven, A. C., and Langen, R.
(2013) alpha-Synuclein oligomers with broken helical conformation form
lipoprotein nanoparticles. J. Biol. Chem. 288, 17620-17630
6. Shore, J. D., Day, D. E., Francis-Chmura, A. M., Verhamme, I., Kvassman, J.,
Lawrence, D. A., and Ginsburg, D. (1995) A fluorescent probe study of
plasminogen activator inhibitor-1. Evidence for reactive center loop insertion and
its role in the inhibitory mechanism. J. Biol. Chem. 270, 5395-5398
7. Salvucci, M. E. (2004) Potential for interactions between the carboxy- and amino-
termini of Rubisco activase subunits. FEBS Lett. 560, 205-209
8. Lakowicz, J. R. (1999) Principles of fluorescence spectroscopy, 2nd ed., Kluwer
Academic/Plenum, New York
9. Gaus, K., Zech, T., and Harder, T. (2006) Visualizing membrane microdomains
by Laurdan 2-photon microscopy. Mol. Membr. Biol. 23, 41-48
10. Parasassi, T., Krasnowska, E. K., Bagatolli, L., and Gratton, E. (1998) Laurdan and
Prodan as Polarity-Sensitive Fluorescent Membrane Probes. Journal of
Fluorescence 8, 365-373
11. Jordanova, R., Radoslavov, G., Fischer, P., Liebau, E., Walter, R. D., Bankov, I.,
and Boteva, R. (2005) Conformational and functional analysis of the lipid binding
protein Ag-NPA-1 from the parasitic nematode Ascaridia galli. FEBS J 272, 180-
189
12. Der-Sarkissian, A., Jao, C. C., Chen, J., and Langen, R. (2003) Structural
organization of alpha-synuclein fibrils studied by site-directed spin labeling. J.
Biol. Chem. 278, 37530-37535
13. Bodner, C. R., Dobson, C. M., and Bax, A. (2009) Multiple tight phospholipid-
binding modes of alpha-synuclein revealed by solution NMR spectroscopy. J.
Mol. Biol. 390, 775-790
45

Chapter 3

IAPP as a Membrane Curvature Inducer and Sensor

Natalie C. Kegulian, Shalene Sankhagowit, Melania Apostolidou, Sajith A. Jayasinghe,
Noah Malmstadt, Peter C. Butler, and Ralf Langen

Adapted from Kegulian, N. C., Sankhagowit, S., Apostolidou, M., Jayasinghe, S. A.,
Malmstadt, N., Butler, P. C., and Langen, R. (2015) Membrane curvature-sensing and -
inducing activity of islet amyloid polypeptide and its implications for membrane
disruption. J. Biol. Chem. Under revision.
46

3.1 Abstract
Islet amyloid polypeptide (IAPP) is a 37-amino-acid amyloid protein intimately
associated with pancreatic islet β-cell dysfunction and death in type II diabetes. In this
study, we combine spectroscopic methods and microscopy to investigate α-helical
IAPP-membrane interactions. Using light scattering and fluorescence microscopy we
observe that larger vesicles become smaller upon treatment with human or rat IAPP.
Electron microscopy shows the formation of various highly curved structures such as
tubules or smaller vesicles in a membrane-remodeling process, and spectrofluorometric
detection of vesicle leakage shows disruption of membrane integrity. This effect is
stronger for hIAPP than for the less toxic rIAPP. From CD spectra in the presence of
different-sized vesicles, we also uncover the membrane curvature-sensing ability of
IAPP and find that it transitions from inducing to sensing membrane curvature when
lipid negative charge is decreased. Our in vivo EM images of immunogold-labeled
rIAPP and hIAPP show both forms to localize to mitochondrial cristae, which contain
not only locally curved membranes but also cardiolipin, a lipid with high spontaneous
negative curvature. Disruption of membrane integrity by induction of membrane
curvature could apply more broadly to other amyloid proteins and be responsible for
membrane damage observed in other amyloid diseases as well.

47

3.2 Introduction
Following my study of α-synuclein-membrane interactions, I examined the different
outcomes emerging from adding IAPP to membranes. As described in Chapter 1, IAPP
is also associated with cellular membrane damage, in this case in pancreatic islet β-cells
in type II diabetes. Mitochondrial and ER membranes in human IAPP (hIAPP)-
transgenic mice and in hIAPP-expressing human insulinoma cells were found to lose
their integrity (1). However, the exact mechanism behind IAPP-induced membrane
damage remains under debate.
Different mechanisms have been proposed for IAPP-dependent membrane
damage, with some of the more recent studies (2-4) illustrating IAPP-mediated
membrane disruption to occur in two or possibly more (5) steps. In one model, hIAPP
fibrils growing at the membrane extract lipid in the process, tearing at the membrane
and fragmenting it in a nonspecific manner (6,7). However, rat IAPP (rIAPP), which
does not readily form fibrils (8), has also been seen to cause membrane leakage in vitro
(5) and confer some (albeit strongly reduced) cytotoxicity in vivo (9). Also, the
nonamyloidogenic hIAPP1-19 fragment as well as a number of nonamyloidogenic full-
length variants of IAPP have been observed to cause leakage of synthetic membranes
(2,4,10), and hIAPP1-19 and rIAPP1-19 can permeabilize pancreatic islet cell membranes
(11). Therefore, fibril growth cannot be the sole mechanism involved. Other
mechanisms have been suggested that are analogous to those utilized by antimicrobial
peptides (12-14). Peptides that employ the detergent-like carpet mechanism cause
widespread defects throughout the membrane surface in a nonspecific manner (15-18).
Pore models, in contrast, entail monomers assembling to form a discrete oligomer in the
membrane that acts as a pore with ion channel-like properties (19,20) or, alternatively,
peptides inducing the membrane to form a toroidal pore lined by its own phospholipids
(21-23). Different studies have proposed a variety of mechanisms such as these for
IAPP, including pore mechanisms (5,24-28). Given the increasingly established bi- or
48

multiphasicity of IAPP-mediated membrane damage (2-5), one or more of the above
mechanisms could be part of an early step, with the later or latest step involving β-sheet
fibril formation.
One possibility that remains unexplored is that IAPP induces membrane
curvature as an α-helical wedge. Prior to forming β-sheet fibrils, IAPP is known to bind
negatively charged lipids and detergents using an amphipathic α-helical structure (29-
33). Here we hypothesize that the insertion of such a helical structure into negatively
charged membranes might induce and/or sense membrane curvature. Other proteins
that form membrane-bound amphipathic α-helices are known to induce membrane
curvature by pushing phospholipid headgroups apart, thereby effectively acting as
wedges. Examples include the H0 α-helices of N-BAR domain proteins such as
amphiphysin and endophilin (34-41). In addition, α-synuclein binds to negatively
charged membranes as an amphipathic α-helix that alone is sufficient to induce
membrane curvature—that is, to remodel liposomes into tubular structures, small,
highly curved vesicles, or protein-lipid nanoparticles (42-45). Importantly, such
membrane remodeling and induction of membrane curvature can cause disruption of
membrane integrity (42). Furthermore, at presynaptic termini in transgenic mice,
overexpressed human α-synuclein was associated with remodeling of organelle
membranes into highly curved structures (46). We therefore tested whether some of the
previously observed membrane-disrupting effects of IAPP might have been caused by
induction of membrane curvature. Moreover, prior studies on membrane curvature-
inducing proteins have shown that such proteins can also exhibit curvature-sensitive
membrane binding under some conditions (34,47-51). Inasmuch as sensing of
membrane curvature could have implications for physiological and pathogenic
recruitment of IAPP to membranes, we also investigated the curvature sensitivity of
IAPP-membrane interaction.
49

Using circular dichroism (CD), spectrophotometry, and fluorescence microscopy
of giant unilamellar vesicles (GUVs), we find that α-helical IAPP converts large,
negatively charged vesicles into much smaller structures. Dye leakage experiments
indicate that this membrane remodeling coincides with significant disruption of
membrane integrity, and electron microscopy (EM) reveals the IAPP-dependent
formation of lipid tubules and smaller vesicles. By uncovering IAPP as a curvature
inducer, we can correlate the previously discovered early phase of membrane leakage
with the modulation of membrane architecture into highly curved structures. When
using only weakly negatively charged membranes, we find that IAPP transitions from
an inducer of membrane curvature to a sensor of membrane curvature. The induction or
sensing of membrane curvature may impact IAPP’s physiological and pathological
functions and govern its membrane localization in vivo. In support of this notion, we
show that IAPP preferentially localizes to mitochondria, where it interacts with cristae.
Cristae are curved membrane structures rich in cardiolipin, a lipid with pronounced
negative spontaneous curvature.
3.3 Methods
Materials
Synthetic hIAPP was obtained from Bachem (Torrance, CA), synthetic rIAPP from
BiomerTech (Pleasanton, CA). 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine
(POPC), 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-L-serine] (POPS), 1-palmitoyl-2-
oleoyl-sn-glycero-3-[phospho-RAC-(1-glycerol)] (POPG), 1,2-dioleoyl-sn-glycero-3-
[phospho-RAC-(1-glycerol)] (DOPG), 1-palmitoyl-2-{12-[(7-nitro-2-1,3-benzoxadiazol-4-
yl)amino]dodecanoyl}-sn-glycero-3-phosphoserine (NBD-PS), 1,2-dioleoyl-sn-glycero-3-
phosphoethanolamine-N-[lissamine rhodamine B sulfonyl] (Rh-PE), and 1,2-
dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl) (biotin-PE) were
obtained as solutions in chloroform from Avanti Polar Lipids, Inc. (Alabaster, AL).
Hexafluoroisopropanol (HFIP), 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid
50

(HEPES), thioflavin T (ThT), Triton-X 100, and asolectin were obtained from Sigma-
Aldrich (St. Louis, MO), ANTS, DPX, and avidin from Invitrogen (Life Technologies,
Grand Island, NY), guanidine hydrochloride from Thermo Scientific (USA), and uranyl
acetate from Electron Microscopy Sciences (EMS, Hatfield, PA). Polydimethylsiloxane
(PDMS) was purchased from Dow Corning (Midland, MI).
Preparation of peptides
hIAPP or rIAPP powder was initially dissolved in HFIP. To quantify peptide
concentrations, a small amount of peptide was isolated, the HFIP evaporated under a
stream of nitrogen gas and replaced with 8 M guanidine HCl, and absorbance at 280 nm
measured. hIAPP in HFIP was then aliquotted, flash-frozen in liquid N2, and
lyophilized overnight. Immediately prior to each experiment, hIAPP was redissolved in
10 µl deionized water containing 0.5% acetic acid, to which 40 µl 110 mM HEPES, pH
7.4, was added. The resulting peptide solution was added to vesicles to a final peptide
concentration of 25 µM (unless otherwise indicated). For rIAPP, HFIP was evaporated
under a nitrogen stream and the peptide redissolved in 10 mM HEPES, pH 7.0, 50 mM
KCl, or 10 mM sodium phosphate, pH 7.0, no salt. Immediately before each experiment,
peptide solution was centrifuged in a tabletop Eppendorf 5415D centrifuge at 13,200
rpm for 10 minutes, the supernatant taken, and the peptide concentration determined
using A280. To perform the experiment, the appropriate amount of peptide was then
added to vesicles to reach a final peptide concentration of 25 μM (unless otherwise
indicated).
α-Synuclein was prepared as described in Chapter 2.
Preparation of phospholipid vesicles
Multilamellar vesicles (MLVs) composed of POPC, POPS, POPG, DOPG, or
POPS/POPC mixtures were prepared as described in Chapter 2, rehydrating in 10 mM
HEPES, 50 mM KCl, pH 7.0. For leakage assays, large unilamellar vesicles (LUVs) with
a diameter of 100 nm were prepared. First, lipids were rehydrated in 10 mM HEPES, pH
51

7.0, 2.4 mM KCl, 1 mM EDTA, 3 mM sodium azide, 9 mM ANTS, and 25 mM DPX. This
resuspension was subjected to 10 cycles of freeze/thaw, followed by at least twenty-one
extrusions through 2 100-nm-cutoff polycarbonate membranes (Avanti).
Unencapsulated ANTS and DPX were removed by gel filtration using a PD-10 column,
eluting the LUVs in 10 mM HEPES, pH 7.0, 50 mM KCl, 1 mM EDTA, and 3 mM
sodium azide.
For in vitro curvature sensitivity measurements, lipids were rehydrated in 10 mM
sodium phosphate, pH 7.0, no salt. 100-nm LUVs were prepared by freeze/thawing and
extruding the vesicle suspension as described above. Small unilamellar vesicles (SUVs)
were prepared by bath-sonicating the suspension, then centrifuging it in an Optima
TLX Ultracentrifuge (Beckman Coulter, Inc.) at 60,000 rpm for 1 hour using an Optima
TLA-100.3 fixed-angle rotor (Beckman Coulter) and collecting the supernatant.
Preparation of giant unilamellar vesicles (GUVs) was modified from a previous
method (52). Briefly, the lipid mixture dissolved in chloroform was dried on an indium-
tin oxide (ITO)-coated glass slide (Delta Technologies, Loveland, CO) and stored under
vacuum overnight. The resulting lipid film was hydrated with the addition of 10 mM
HEPES, pH 7.0, 100 mM sucrose to the volume enclosed by two ITO-coated slides and a
2.5-mm-thick silicon spacer. The GUVs were formed at room temperature (~23°C) by
applying a 1.5-V AC field with a frequency of 10 Hz for 1 hour.
Vesicle clearance assay
Clearance of MLVs was monitored by measuring change in light scattering as a function
of time at a wavelength of 500 nm and a slit width of 2 nm in a Jasco V-550
spectrophotometer. hIAPP, rIAPP, α-synuclein, or buffer as a control was added to
vesicles in a quartz cuvette. IAPP was added to a final protein/phospholipid molar ratio
of 1:20. α-Synuclein was added at a lower molar concentration to a final protein/lipid
molar ratio of 1:74, to match the mass to lipid ratio used in the IAPP experiments. Each
experiment was performed at least in triplicate. Data were normalized by setting initial
52

values to 1 and thereby plotting the fraction of remaining light scattering as OD500 at
each time point.
CD
Measurements were taken using a Jasco J-810 spectropolarimeter in a 1-mm quartz cell
every 0.5 nm at a 50-nm/minute scan rate and a response time of 1 second. For each
spectral measurement performed in parallel to the clearance assay, eight scans were
averaged and the appropriate background subtracted; then, the spectrum was
smoothed by the Savitsky-Golay method. For CD spectra measured in order to
determine curvature sensitivity, sixteen scans were averaged and the appropriate
background subtracted, and experiments were performed in 10 mM sodium phosphate,
pH 7.0, no salt. In all cases, values were normalized in order to obtain mean residue
ellipticity (MRE).
ThT assays
To monitor possible fibril formation from peptides, ThT fluorescence assays were also
performed in parallel using 25 µM ThT in the sample. Emission intensities at 482 nm
were measured as a function of time with excitation at 450 nm, excitation and emission
slit widths of 1 and 10 nm, respectively, and a response time of 1 second. Measurements
were taken using a Jasco FP-6500 spectrofluorometer and normalized to the initial
fluorescence values.
Fluorescence microscopy
The NBD-PS and Rh-PE labeled GUVs were observed with confocal fluorescence
microscopy on a Nikon TI-E inverted microscope, using illumination provided by 50
mW solid-state 491- and 561-nm lasers for the dyes and capturing fluorescent emissions
at 525 and 595 nm, respectively. To minimize volumes used during observations of
GUVs, a ~4.6-mm-thick PDMS sheet with a 6-mm-diameter cylindrical hole was
attached to the coverslip to form a ~130-µL observation well. The 0.3% biotin-PE GUVs
were immobilized with avidin-biotin tethers to a supported lipid bilayer on the glass
53

coverslip surface. The supported bilayer was prepared by rupturing 85:15
asolectin/biotin-PE LUVs onto glass. These LUVs were formed by 30-minute sonication
at 30°C of overnight vacuum-dried lipid film hydrated to a 2 mg/mL concentration with
10 mM HEPES, pH 7.0, 50 mM KCl. The LUVs were then filtered through the 0.45-µm
cellulose membrane of a syringe filter (VWR International, Radnor, PA). This LUV
solution was deposited onto the glass surface, left for 30 minutes, and rinsed via
pipetting with 10 mM HEPES buffer, pH 7.0, no salt. Next, a 1 mg/mL avidin solution in
water was added and left for 15 minutes and was followed by the same rinsing
procedure but with the 10 mM HEPES, pH 7.0, 50 mM KCl. The GUVs were added to
the observation well and allowed 10 minutes to sediment to the coverslip surface, after
which excess vesicles were removed to prevent the peptides from acting on floating,
unobservable vesicles prior to reaching those in focus. The resulting GUV lipid molar
concentration was estimated to be on the order of 2.5 to 5 times the final concentration
of peptide used in each experiment.
Leakage assay
Release of ANTS and its quencher DPX was gauged as an increase in ANTS
fluorescence intensity as a function of time. One hundred percent leakage was attained
using a final concentration of 0.04% Triton-X 100, and all data were normalized to
fluorescence intensity at this amount. Excitation and emission wavelengths were set at
380 and 520 nm with slits of 1 and 12 nm, respectively, and a response time of 1 second.
EM of vesicles
Samples were deposited as 10-μl droplets on parafilm and carbon-coated formvar films
mounted on copper grids (EMS) floated atop for 5 minutes. Then, after blotting the
excess liquid with filter paper, grids were floated on 10-μl droplets of 1% uranyl acetate
for 1 minute for negative staining. Microscopy was performed using a JEOL 1400
transmission electron microscope accelerated to 100 kV. To build size distribution
histograms, MLVs composed of POPS were initially extruded to 1 μm in diameter and
54

an arbitrarily chosen quadrant was sampled for each of a series of images taken of
POPS alone, POPS with hIAPP, and POPS with rIAPP; quadrants chosen throughout
each series were those at the same placement as the first arbitrarily chosen quadrant for
that series. Subsequently, the vesicles within the chosen quadrant for each series were
counted for every image in that series, then categorized by size.
EM of pancreatic cells
With Animal Use and Care Committee approval whole pancreas was removed from a
wild type female FVB mouse after euthanasia and tissue was minced in Trump's
solution at 4°C, fixed for 3 hours, and then transferred to 0.1 mol/1 phosphate buffer.
With IRB approval, human insulinoma tissue previously obtained at surgery as
described (53) was also minced immediately in Trump’s solution and then postfixed in
1% osmium tetroxide. Samples from both the mouse pancreas and human insulinoma
were then dehydrated in increasing concentrations of acetone, embedded in Quetol
resin, and polymerized at 40°C, and 80-nm sections were placed on nickel grids for
immunogold labeling of IAPP as described previously (53).
3.4 Results
IAPP transforms negatively charged vesicles into smaller structures
In order to test whether IAPP can remodel membranes, we added human or rat IAPP to
MLVs made up of POPS, a negatively charged phospholipid, and monitored the change
in light scattering by measuring OD500 over time, as previously described for α-
synuclein (42). Fig. 3-1A shows that hIAPP and, to a lesser extent, rIAPP cleared
scattering by POPS MLVs, indicating their ability to transmute large vesicles into
smaller entities. We performed the same assay using buffer alone as a negative control
or α-synuclein, which is known to cause clearance of phosphatidylglycerol MLVs
(42,44). Buffer induced no difference in light scattering, while α-synuclein caused some
POPS clearance, though less so than either type of IAPP (Fig. 3-1A). Vesicles made up
entirely of the zwitterionic phospholipid POPC showed no clearance upon addition of
55

any peptide or buffer control (Fig. 3-1BC). These results suggest that the electrostatic
interactions between IAPP and the membrane lead to loss of scattering by vesicles. In
order to more systematically investigate how the change in scattering depends on the
presence of negatively charged POPS, we reduced the percentage of POPS in the vesicle
composition by replacing it with POPC. We found that the less POPS the vesicle
membranes contained, the less clearance was observed (Fig. 3-1C). hIAPP was generally
more potent than rIAPP at reducing the scattering and this difference was particularly
pronounced at 33% POPS. Readings shown were collected before the onset of
aggregation, as illustrated by the example of CD spectra measured in the presence of
100% POPS (Fig. 3-1D), which after 1 hour of clearance contained negative peaks at 208
and 222 nm, indicating α-helical, as opposed to β-sheet, structure. ThT fluorescence
measurements taken in parallel showed no increase in ThT fluorescence (Fig. 3-1E;
compare to the sigmoidal increase in ThT fluorescence of hIAPP over a longer
timecourse in figure in Table 3-1), also indicating no misfolding within the time frame
of the experiment.
In addition, we tested IAPP clearance of POPG and DOPG vesicles, since
phosphatidylglycerol is a membrane component that is widely used in the literature
(27,54,55). Both hIAPP and, albeit less so, rIAPP moderately cleared light scattering
from POPG more than that from DOPG, though not to the extent that they cleared
POPS light scattering (as summarized in Fig. 3-1C). Both types of IAPP showed a less
pronounced effect on DOPG than did α-synuclein (Fig. 3-1C). For all the data shown, α-
synuclein, a 140-aa protein, was used at a mass per volume equal to that of IAPP,
resulting in a 1:74 rather than a 1:20 protein/lipid molar ratio for α-synuclein and
therefore a smaller clearance effect by α-synuclein on POPG in this experiment than in
our previously published study, which utilized a 1:20 α-synuclein/POPG molar ratio
(42). We also measured ThT fluorescence for our phosphatidylglycerol-containing
samples to ensure that the agent inducing vesicle clearance was not a β-sheet. As with
56

100% POPS, for all peptides, ThT fluorescence showed no aggregation within the time
frame required to reach vesicle clearance (Table 3-1).

Fig. 3-1. Light scattering of MLVs. Changes in light scattering upon addition of hIAPP
(magenta), rIAPP (green), α-synuclein (blue), or buffer (black) determined from the optical
density at 500 nm. Clearance curves of 100% POPS (A) or 100% POPC (B) in the presence of each
protein or buffer control. (C) Lowest light scattering reading within the first hour of clearance for
MLVs of the indicated phospholipid compositions. Error bars represent standard error; N ≥ 3.
CD spectra of hIAPP, rIAPP, and α-synuclein mixed with 100% POPS (D). ThT fluorescence
timecourses for the three proteins mixed with 100% POPS (E). Phospholipid concentration is 500
μM, IAPP concentration is 25 μM, and α-synuclein concentration is matched to the IAPP
concentration by mass, at 6.74 μM. Clearance and ThT fluorescence readings are normalized by
setting their initial readings to 1. CD readings are normalized to MRE.

57

IAPP reduces negatively charged giant vesicles into smaller lipidic structures as
visualized by fluorescence microscopy
Our next step was to view the consequences of IAPP-membrane interactions directly.
We immobilized GUVs and observed them under fluorescence confocal microscopy.
Within 6 seconds of adding hIAPP, POPS vesicles dissolved into small structures (Fig.
3-2A, green), while control POPC vesicles present in the same preparation remained
intact (Fig. 3-2A, blue). POPS GUVs treated with rIAPP also disbanded into smaller
lipidic structures, with POPC remaining intact (Fig. 3-2B). Samples were observed for at

Table 3-1. Half-times to maximal ThT fluorescence for each protein-lipid
combination in which an increase was detected. Times to half-maximal
fluorescence intensity (t1/2) were measured as the time points at which ThT
fluorescence was at half its value when it reached its plateau, for protein-lipid
combinations that yielded sigmoidal curves (example of hIAPP and 100% POPS
shown in figure).
Lipid composition
t1/2 (hours)
hIAPP rIAPP α-synuclein
100% POPS 1.9 NA NA
66:34 POPS/POPC 0.7 NA NA
33:67 POPS/POPC 1.8 NA NA
100% POPC NA NA NA
100% POPG 0.7 NA NA
100% DOPG NA NA NA
NA, not applicable (ThT fluorescence did not significantly increase within 2 hours).

58

least 10 minutes after addition of either variety of IAPP, at which point still no
dissolution of POPC vesicles was observed (Fig. 3-2AB, iv).

IAPP modulates membrane architecture to form structures of high curvature as
visualized by EM
Thus far, we have observed that, when added to negatively charged vesicles, α-helical
IAPP leads to a population of smaller lipidic structures. To glean more information on
the structures formed from affected vesicles, we used negative stain EM to visualize
large vesicles before (example of POPS shown in Fig. 3-3A) and after (Fig. 3-3BC)
treatment with IAPP. Both hIAPP (Fig. 3-3B) and rIAPP (Fig. 3-3C) induced formation
of smaller vesicles from large ones made of POPS. The size distribution histogram (Fig.
3-3D) for treated and untreated POPS vesicles shows a greater proportion of small
vesicles in the presence of either IAPP compared to POPS alone; in particular, <20% of
i ii iii iv
i ii iii iv
A
B
Fig. 3-2. Dissolution of POPS membranes by IAPP visualized directly under fluorescence
microscopy. GUVs composed of 99.2:0.5:0.3 POPS/NBD-PS/biotin-PE (green) or of 99.2:0.5:0.3
POPC/Rh-PE/biotin-PE (blue) before (i), immediately after (ii), 6 seconds after (iii), and 10
minutes after (iv) treatment with hIAPP (A) or rIAPP (B). Scale bar, 15 μm. Images taken by
Shalene Sankhagowit.
59

the vesicles
counted that
were untreated
measured
under 150 nm
in diameter,
whereas ~70%
of those treated
with hIAPP and
~60% of those
treated with
rIAPP
measured
under 150 nm.
In
addition, we
observed tubule
formation,
another way in
which
curvature
induction can
manifest itself, in some IAPP-lipid combinations (Fig. 3-3E-G). However, the result that
we detected was not a consistent generation of tubules upon mixing lipids with IAPP
but was instead a heterogeneous mix of images across individual EM grids, such that
even combinations that led to tubulation showed no tubes in other places on their
respective grids (as illustrated in Fig. 3-3HI). This heterogeneity is in contrast to the
Fig. 3-3. Negative stain EM of the interaction of hIAPP or rIAPP with
large vesicles. POPS in the absence of peptide (A), with hIAPP (B), and
with rIAPP (C). (D) Protein/single phospholipid molar ratios are 1:20.
Size distributions of 100% POPS vesicles alone (black), with hIAPP
(magenta), or with rIAPP (green). A negligible percentage of vesicles
measured over 1200 nm in diameter. DOPG (E) in the presence of hIAPP.
Protein/single phospholipid molar ratio is 1:20. Vesicles composed of
20:80 POPS/POPC (F) and of 5:95 POPS/POPC (G) in the presence of
rIAPP. Protein/single phospholipid molar ratios are 1:180. (H) Same
20:80 POPS/POPC sample as in part (F), and (I) same 5:95 POPS/POPC
sample as in part (G), areas of respective grids showing little tubulation.
(J) 5:95 POPS/POPC in the absence of peptide, showing some baseline
tubulation (inset).
60

stable, consistent tubulation generated by α-synuclein (42,44). Furthermore, in the 5:95
POPS/POPC MLVs in the absence of peptide (Fig. 3-3J), we did observe a small amount
of tubulation (Fig. 3-3J, inset), which indicates a baseline of spontaneous tubulation in
high-POPC content vesicles. Thus, in contrast to higher negative charge conditions, it is
more difficult to cleanly conclude that
membrane curvature was induced by
IAPP at 5% PS content. Also
observable in some IAPP-lipid
combinations (see, for example, Fig. 3-
3C) was the formation of irregularly
shaped amorphous structures that
resemble neither tubules nor vesicles.
These structures might have been a
consequence of membrane
fragmentation (56).
IAPP induces vesicle leakage in a
charge-dependent manner
In order to assess whether the
reduction in vesicle size correlated with loss of membrane integrity, we next measured
vesicle leakage. IAPP was added to LUVs encapsulating the fluorophore ANTS and its
quencher DPX, and ANTS fluorescence was monitored as a measure of membrane
leakage. LUVs made up of 66:34 POPS/POPC leaked up to their respective plateaus
within 10 minutes of addition of hIAPP or rIAPP with the latter being slightly less
potent under these conditions (Fig. 3-4). In both cases, the time frame was faster than
that required for clearance of IAPP-treated vesicles of various amounts of POPS to reach
its plateau (Fig. 3-1A and Table 3-2). This timecourse indicates that vesicle integrity is
already breached at the onset of membrane remodeling. Overall, the timecourse

Fig. 3-4. IAPP-induced vesicle leakage. Leakage
observed for LUVs encapsulating ANTS and its
quencher DPX with membranes made of 66:34
POPS/POPC in the presence of hIAPP (green,
double line) or rIAPP (blue, long dashes); 10:90
POPS/POPC in the presence of hIAPP (red,
solid) or rIAPP (orange, short dashes); or 100%
POPC in the presence of hIAPP (black, large
dots) or rIAPP (gray, small dots). Data collected
in collaboration with Sajith Jayasinghe and
Melania Apostolidou.
61

corresponds to those previously observed for α-synuclein and amphiphysin (42). While
neither hIAPP nor rIAPP appreciably causes leakage of 100% POPC-containing LUVs
(Fig. 3-4), 10:90 POPS/POPC-containing LUVs leaked in the presence of hIAPP whereas
rIAPP only caused minor leakage (Fig. 3-4). Thus, as in the case of the clearance
experiments, hIAPP had a stronger effect on membranes and the difference between the
peptides was most pronounced at intermediate lipid charge densities.
IAPP senses membrane curvature under conditions that only moderately favor
binding
Depending on conditions, some membrane curvature-inducing proteins have been
shown to act alternately as sensors (47,50). To delve into the possible membrane
curvature-sensing properties of IAPP, we used CD to quantify the helicity of rIAPP on
LUVs and SUVs, which are about 100 and 40 nm in diameter, respectively, and the
latter of which is by nature more highly curved. We restricted this part of our study to
rIAPP, which does not misfold readily and can thus be more stably investigated by CD
over a wide range of conditions. We measured the CD spectra of rIAPP on LUVs and
SUVs consisting of 10:90, 20:80, 33:67, and 66:34 POPS/POPC, using only three-quarters
Table 3-2. Time points at which light scattering reached its minimum (i.e., the
sample reached maximal clearance) and at which scattering began to increase due
to protein aggregation.

Lipid composition
Time (in hours) at light scattering minimum/light scattering
increase
hIAPP rIAPP α-synuclein
100% POPS 0.3/NA 0.3/NA 0.8/NA
66:34 POPS/POPC 0.2/0.4 0.3/NA 1/NA
33:67 POPS/POPC 0.1/0.3 NA/NA NA/NA
100% POPC NA/NA NA/NA NA/NA
100% POPG 0.5/0.9 1/NA 1/NA
100% DOPG 0.2/0.4 0.9/NA 1/NA
NA, not applicable (light scattering did not significantly decrease/increase within 2
hours).
62

of the lipid molar concentration for SUVs as for LUVs in order to offset the fact that
approximately two-thirds of the phospholipids in SUV membranes, as opposed to
about half of the phospholipids in LUV membranes, compose the outer leaflet (57). As
an example, spectra from 20:80 POPS/POPC are shown in Fig. 3-5A. rIAPP in the
absence of vesicles yields a spectrum that peaks with negative ellipticity at ~200 nm,
indicating a mostly random coil structure with only slight helicity. In contrast, the CD
spectrum of rIAPP in the presence of 20:80 POPS/POPC SUVs displays negative peaks
at 208 and 222 nm, signifying an α-helical structure. rIAPP with LUVs presents a CD
spectrum that is intermediate between the other two. The three spectra intersect at a
wavelength of 204 nm, an isosbestic point typical of spectra of a peptide transitioning
between random coil and helical states. We conclude that, at this composition, the more
highly curved vesicles induce more rIAPP to enter the α-helical, and therefore
membrane-bound, state. To quantify this difference, we plotted MRE values at 222 nm
as a measure of α-helicity of rIAPP in the presence of LUVs or SUVs of varying POPS

Fig. 3-5. Membrane curvature sensitivity of rIAPP. (A) Representative MRE spectra for rat
rIAPP alone (black) or in the presence of 20:80 POPS/POPC LUVs (red) or of SUVs of the
same composition (turquoise). Spectra intersect at the isosbestic point of 204 nm. (B) MRE
values at 222 nm plotted against POPS content for rIAPP on LUVs (circles) and SUVs
(squares) of different compositions. Each point represents the average of three experiments.
Error bars represent standard error; N = 3. Where error bars appear as horizontal lines,
standard error is too small to be seen as a vertical line. Single phospholipid concentration is
2 mM for LUVs, 1.5 mM for SUVs. rIAPP concentration is 25 μM in each case.
63

content (Fig. 3-5B). MRE values at 222 nm differed substantially between SUVs and
LUVs in each case except for 66:34 POPS/POPC, wherein values were similar. We
conclude that, at low negative charge, α-helicity of rIAPP is dependent upon membrane
curvature. Thus, it appears that IAPP acts as a membrane curvature sensor under
conditions that limit binding and as a membrane curvature inducer under conditions
that strongly favor binding (see Discussion).
Having established that IAPP has the ability to sense and/or induce membrane
curvature, we next wanted to investigate whether membrane curvature may be a factor
in the cellular localization of IAPP. To probe this possibility, we viewed immunogold-
labeled endogenous IAPP
under EM in mouse
pancreatic islet β-cells (Fig. 3-
6A) and in cells from an
insulinoma removed from a
human (Fig. 3-6BC) and
observed it to bind to the
cristae of the inner
mitochondrial membrane, as
well as other intra- and
extracellular vesicular
structures. Although our EM
images lack the three-
dimensional resolution to
show that IAPP exclusively
interacts with curved membranes, it should be noted that mitochondrial cristae as well
as crista-crista junctions are predominantly highly curved, containing extensive tubular
as well as bent membranes (58-60). Aside from the physical curvature, mitochondrial

Fig. 3-6. IAPP binding to mitochondria and vesicular
structures in vivo. EM images showing immunogold-
labeled endogenous IAPP bound chiefly to mitochondria
in (A) female mouse islet cells and (B,C) human
insulinoma cells. Images taken by Peter Butler.

64

cristae are rich in cardiolipin, a lipid known to harbor significant amounts of negative
spontaneous curvature (61,62). Such spontaneous curvature is the consequence of a low
headgroup to acyl chain ratio causing low packing densities in the headgroup region,
thereby mimicking the effects of physical curvature. Both this physical curvature and
the negative spontaneous curvature of cardiolipin are likely to contribute to IAPP’s
binding affinity for cristae.
3.5 Discussion
The present study sought to uncover the effects of α-helical IAPP on membranes. We
detected charge-dependent induction of membrane curvature as a result of IAPP-
membrane interactions, leading to the formation of a range of highly curved structures
accompanied by disruption of membrane integrity as well as leakage. This effect was
strongest for hIAPP, although rIAPP was also capable of inducing membrane curvature.
In addition, we observed IAPP to act as a membrane curvature sensor under conditions
that limit membrane binding. Our data show IAPP to be α-helical for the duration of
this membrane curvature-inducing and -sensing activity.
α-Synuclein has previously been observed to induce membrane curvature in a
similar manner (42-45) to the behavior of IAPP in the current study. We propose that
membrane curvature, whether induced by α-synuclein or by IAPP, is mediated by the
wedging of membrane-inserting amphipathic helices. While both IAPP and α-synuclein
induce tubulation and vesiculation of membranes, α-synuclein has a tendency to
generate more stable tubules, while tubulation exerted by IAPP is far weaker. This
difference between the proteins in ensuing lipid tubule stability is likely due to the
unusually extended helical structure of α-synuclein that is ~140 Å in length (63) and
that we have observed in Chapter 2 to be capable of binding membranes from the N-
terminus to as far as residue 100. We previously noted that the long helical structure of
α-synuclein generates a highly anisotropic curvature strain that is ideally suited for
stabilizing the anisotropic curvature of tubes (42). In agreement with this notion, we
65

find that the shorter IAPP helix is less capable of stabilizing tubular structures.
Regardless of this difference, our data from the current study link the membrane
curvature-inducing abilities of both proteins to a loss of membrane integrity.
Furthermore, we observed amorphous structures that were neither tubular nor
vesicular. Those structures could be similar to the protein-lipid complexes previously
observed for α-synuclein (45) and could explain the previous finding of membrane
fragmentation reported for IAPP (56).
Considering the general prevalence of disruption of membrane integrity by
amyloid proteins (64), could the inferred membrane curvature induction mechanism be
more general? Amyloid β-peptide and huntingtin have been observed to bind to
membranes using the insertion of amphipathic helices (65,66). This manner of
membrane binding is similar to that of IAPP (29-33), and one might expect potential
induction of membrane curvature via helical wedges in those proteins as well. Such
membrane remodeling could further be facilitated by oligomeric structures that act as
delivery vehicles, creating a high local protein density. The membrane curvature
induction mechanism could also conceivably apply to membrane damage by
antimicrobial peptides, as they seem to remodel membranes and disrupt their integrity
along the same energetic landscape as amyloid proteins (67). However, additional
structural studies are needed to assess utilization of this membrane curvature
mechanism by proteins other than IAPP and α-synuclein.
Membrane binding by IAPP depends upon membrane curvature (Fig. 3-5) and
charge density (Fig. 3-1C), where a higher density of negative charge in the membrane
leads to a stronger driving force for membrane interaction and therefore curvature
induction (Fig. 3-1 and Fig. 3-3). At lower negative charge densities, some membrane
remodeling is observed (Fig. 3-3FG), but under such conditions membrane binding
becomes strongly sensitive to membrane curvature (Fig. 3-5). Thus, with the decrease in
driving force for IAPP-membrane interactions due to lower negative charge density
66

comes an increasingly
pronounced positive effect on
membrane binding
originating from the
additional spacing between
headgroups in more highly
curved membranes (Fig. 3-
7A), whereas high negative
charge densities cause IAPP
to insert into membranes
without the need for existing
membrane curvature, and
this insertion induces new
membrane curvature (Fig. 3-7B). Therefore, membrane curvature sensing and inducing
are likely to be coupled phenomena that depend upon the affinity of IAPP for the
membrane, as governed by negative charge density. Our in vitro data suggest that
hIAPP and rIAPP disrupt membrane integrity by induction of membrane curvature, but
that hIAPP does so more efficiently, especially in the presence of membranes composed
of more modest, physiologically relevant levels of negative charge. This quantitative
rather than qualitative difference between hIAPP and rIAPP is consistent with the
recent finding that rIAPP can become toxic, but that much higher concentrations are
required for this peptide (9).
Curiously, in β-cells of hIAPP-expressing rodent models of type II diabetes as
well as those of humans with type II diabetes, mitochondrial membranes, which are
partially negatively charged, are particularly prone to disruption (1). Our in vivo EM
data (Fig. 3-6) show human and rat IAPP to localize to mitochondria. Considering that
hIAPP is more potent at remodeling membranes, one might expect that hIAPP becomes

Fig. 3-7. Membrane curvature-inducing and -sensing
behavior in α-helical IAPP. (A) IAPP easily binds, even at
low negative charge density, to highly curved membranes
by fitting between the spaced headgroups. (B) At high
membrane negative charge, IAPP binds membranes
regardless of local curvature, wedging headgroups apart
in planar membranes to force curvature. Cartoon by Ralf
Langen.

67

more disruptive to these membranes than rIAPP. The driving force for such membrane
disruption could further be enhanced by increased negative charge density of the
mitochondrial lipids or increased local concentrations of hIAPP, possibly by
oligomerization. The notion that α-helical hIAPP senses membrane curvature and
disturbs the architecture of crista membranes is further supported by the discovery that
α-helical IAPP abrogates mitochondrial function (9). Furthermore, other studies showed
that α-helical hIAPP oligomers permeabilize pancreatic β-cell membranes and cause
those cells to lose their normal morphology and viability (68) and that targeting the α-
helical form of hIAPP can be protective to cells and membranes (69). The finding that
induction of membrane curvature by amyloidogenic proteins results in disruption of
membrane integrity and presumably toxicity might ultimately lead to new therapeutic
strategies.
68

3.6 References
1. Gurlo, T., Ryazantsev, S., Huang, C. J., Yeh, M. W., Reber, H. A., Hines, O. J.,
O'Brien, T. D., Glabe, C. G., and Butler, P. C. (2010) Evidence for proteotoxicity in
beta cells in type 2 diabetes: toxic islet amyloid polypeptide oligomers form
intracellularly in the secretory pathway. Am. J. Pathol. 176, 861-869
2. Brender, J. R., Lee, E. L., Cavitt, M. A., Gafni, A., Steel, D. G., and Ramamoorthy,
A. (2008) Amyloid fiber formation and membrane disruption are separate
processes localized in two distinct regions of IAPP, the type-2-diabetes-related
peptide. J. Am. Chem. Soc. 130, 6424-6429
3. Brender, J. R., Lee, E. L., Hartman, K., Wong, P. T., Ramamoorthy, A., Steel, D.
G., and Gafni, A. (2011) Biphasic effects of insulin on islet amyloid polypeptide
membrane disruption. Biophys. J. 100, 685-692
4. Heyl, D. L., Osborne, J. M., Pamarthy, S., Samisetti, S., Gray, A. W., Jayaprakash,
A., Konda, S., Brown, D. J., Miller, S. R., Eizadkhah, R., and Milletti, M. C. (2010)
Liposome damage and modeling of fragments of human islet amyloid
polypeptide (IAPP) support a two-step model of membrane destruction. Int. J.
Pept. Res. Ther. 16, 43-54
5. Last, N. B., Rhoades, E., and Miranker, A. D. (2011) Islet amyloid polypeptide
demonstrates a persistent capacity to disrupt membrane integrity. Proc. Natl.
Acad. Sci. U. S. A. 108, 9460-9465
6. Sparr, E., Engel, M. F., Sakharov, D. V., Sprong, M., Jacobs, J., de Kruijff, B.,
Hoppener, J. W., and Killian, J. A. (2004) Islet amyloid polypeptide-induced
membrane leakage involves uptake of lipids by forming amyloid fibers. FEBS
Lett. 577, 117-120
7. Engel, M. F., Khemtemourian, L., Kleijer, C. C., Meeldijk, H. J., Jacobs, J., Verkleij,
A. J., de Kruijff, B., Killian, J. A., and Hoppener, J. W. (2008) Membrane damage
by human islet amyloid polypeptide through fibril growth at the membrane.
Proc. Natl. Acad. Sci. U. S. A. 105, 6033-6038
8. Westermark, P., Engstrom, U., Johnson, K. H., Westermark, G. T., and Betsholtz,
C. (1990) Islet amyloid polypeptide: pinpointing amino acid residues linked to
amyloid fibril formation. Proc. Natl. Acad. Sci. U. S. A. 87, 5036-5040
9. Magzoub, M., and Miranker, A. D. (2012) Concentration-dependent transitions
govern the subcellular localization of islet amyloid polypeptide. FASEB J. 26,
1228-1238
10. Cao, P., Abedini, A., Wang, H., Tu, L. H., Zhang, X., Schmidt, A. M., and Raleigh,
D. P. (2013) Islet amyloid polypeptide toxicity and membrane interactions. Proc.
Natl. Acad. Sci. U. S. A. 110, 19279-19284
69

11. Brender, J. R., Hartman, K., Reid, K. R., Kennedy, R. T., and Ramamoorthy, A.
(2008) A single mutation in the nonamyloidogenic region of islet amyloid
polypeptide greatly reduces toxicity. Biochemistry 47, 12680-12688
12. Shai, Y. (1999) Mechanism of the binding, insertion and destabilization of
phospholipid bilayer membranes by alpha-helical antimicrobial and cell non-
selective membrane-lytic peptides. Biochim. Biophys. Acta 1462, 55-70
13. Jayasinghe, S. A., and Langen, R. (2007) Membrane interaction of islet amyloid
polypeptide. Biochim. Biophys. Acta 1768, 2002-2009
14. Almeida, P. F., and Pokorny, A. (2009) Mechanisms of antimicrobial, cytolytic,
and cell-penetrating peptides: from kinetics to thermodynamics. Biochemistry 48,
8083-8093
15. Gazit, E., Miller, I. R., Biggin, P. C., Sansom, M. S., and Shai, Y. (1996) Structure
and orientation of the mammalian antibacterial peptide cecropin P1 within
phospholipid membranes. J. Mol. Biol. 258, 860-870
16. Oren, Z., and Shai, Y. (1998) Mode of action of linear amphipathic alpha-helical
antimicrobial peptides. Biopolymers 47, 451-463
17. Sato, H., and Feix, J. B. (2006) Peptide-membrane interactions and mechanisms of
membrane destruction by amphipathic alpha-helical antimicrobial peptides.
Biochim. Biophys. Acta 1758, 1245-1256
18. Chang, W. K., Wimley, W. C., Searson, P. C., Hristova, K., and Merzlyakov, M.
(2008) Characterization of antimicrobial peptide activity by electrochemical
impedance spectroscopy. Biochim. Biophys. Acta 1778, 2430-2436
19. Matsuzaki, K., Murase, O., Fujii, N., and Miyajima, K. (1995) Translocation of a
channel-forming antimicrobial peptide, magainin 2, across lipid bilayers by
forming a pore. Biochemistry 34, 6521-6526
20. Langham, A. A., Ahmad, A. S., and Kaznessis, Y. N. (2008) On the nature of
antimicrobial activity: a model for protegrin-1 pores. J. Am. Chem. Soc. 130, 4338-
4346
21. Allende, D., Simon, S. A., and McIntosh, T. J. (2005) Melittin-induced bilayer
leakage depends on lipid material properties: evidence for toroidal pores.
Biophys. J. 88, 1828-1837
22. Sengupta, D., Leontiadou, H., Mark, A. E., and Marrink, S. J. (2008) Toroidal
pores formed by antimicrobial peptides show significant disorder. Biochim.
Biophys. Acta 1778, 2308-2317
23. Bertelsen, K., Dorosz, J., Hansen, S. K., Nielsen, N. C., and Vosegaard, T. (2012)
Mechanisms of peptide-induced pore formation in lipid bilayers investigated by
oriented 31P solid-state NMR spectroscopy. PLoS One 7, e47745
24. Green, J. D., Kreplak, L., Goldsbury, C., Li Blatter, X., Stolz, M., Cooper, G. S.,
Seelig, A., Kistler, J., and Aebi, U. (2004) Atomic force microscopy reveals defects
70

within mica supported lipid bilayers induced by the amyloidogenic human
amylin peptide. J. Mol. Biol. 342, 877-887
25. Smith, P. E., Brender, J. R., and Ramamoorthy, A. (2009) Induction of negative
curvature as a mechanism of cell toxicity by amyloidogenic peptides: the case of
islet amyloid polypeptide. J. Am. Chem. Soc. 131, 4470-4478
26. Xu, W., Wei, G., Su, H., Nordenskiold, L., and Mu, Y. (2011) Effects of cholesterol
on pore formation in lipid bilayers induced by human islet amyloid polypeptide
fragments: a coarse-grained molecular dynamics study. Phys. Rev. E Stat. Nonlin.
Soft Matter Phys. 84, 051922
27. Last, N. B., and Miranker, A. D. (2013) Common mechanism unites membrane
poration by amyloid and antimicrobial peptides. Proc. Natl. Acad. Sci. U. S. A.
110, 6382-6387
28. Pannuzzo, M., Raudino, A., Milardi, D., La Rosa, C., and Karttunen, M. (2013)
alpha-helical structures drive early stages of self-assembly of amyloidogenic
amyloid polypeptide aggregate formation in membranes. Sci. Rep. 3, 2781
29. Jayasinghe, S. A., and Langen, R. (2005) Lipid membranes modulate the structure
of islet amyloid polypeptide. Biochemistry 44, 12113-12119
30. Knight, J. D., Hebda, J. A., and Miranker, A. D. (2006) Conserved and cooperative
assembly of membrane-bound alpha-helical states of islet amyloid polypeptide.
Biochemistry 45, 9496-9508
31. Apostolidou, M., Jayasinghe, S. A., and Langen, R. (2008) Structure of alpha-
helical membrane-bound human islet amyloid polypeptide and its implications
for membrane-mediated misfolding. J. Biol. Chem. 283, 17205-17210
32. Patil, S. M., Xu, S., Sheftic, S. R., and Alexandrescu, A. T. (2009) Dynamic alpha-
helix structure of micelle-bound human amylin. J. Biol. Chem. 284, 11982-11991
33. Nanga, R. P., Brender, J. R., Vivekanandan, S., and Ramamoorthy, A. (2011)
Structure and membrane orientation of IAPP in its natively amidated form at
physiological pH in a membrane environment. Biochim. Biophys. Acta 1808, 2337-
2342
34. Peter, B. J., Kent, H. M., Mills, I. G., Vallis, Y., Butler, P. J., Evans, P. R., and
McMahon, H. T. (2004) BAR domains as sensors of membrane curvature: the
amphiphysin BAR structure. Science 303, 495-499
35. Zimmerberg, J., and Kozlov, M. M. (2006) How proteins produce cellular
membrane curvature. Nat. Rev. Mol. Cell Biol. 7, 9-19
36. Campelo, F., McMahon, H. T., and Kozlov, M. M. (2008) The hydrophobic
insertion mechanism of membrane curvature generation by proteins. Biophys. J.
95, 2325-2339
37. Low, C., Weininger, U., Lee, H., Schweimer, K., Neundorf, I., Beck-Sickinger, A.
G., Pastor, R. W., and Balbach, J. (2008) Structure and dynamics of helix-0 of the
N-BAR domain in lipid micelles and bilayers. Biophys. J. 95, 4315-4323
71

38. Gallop, J. L., Jao, C. C., Kent, H. M., Butler, P. J., Evans, P. R., Langen, R., and
McMahon, H. T. (2006) Mechanism of endophilin N-BAR domain-mediated
membrane curvature. EMBO J. 25, 2898-2910
39. Jao, C. C., Hegde, B. G., Gallop, J. L., Hegde, P. B., McMahon, H. T., Haworth, I.
S., and Langen, R. (2010) Roles of amphipathic helices and the
bin/amphiphysin/rvs (BAR) domain of endophilin in membrane curvature
generation. J. Biol. Chem. 285, 20164-20170
40. Cui, H., Mim, C., Vazquez, F. X., Lyman, E., Unger, V. M., and Voth, G. A. (2013)
Understanding the role of amphipathic helices in N-BAR domain driven
membrane remodeling. Biophys. J. 104, 404-411
41. Ambroso, M. R., Hegde, B. G., and Langen, R. (2014) Endophilin A1 induces
different membrane shapes using a conformational switch that is regulated by
phosphorylation. Proc. Natl. Acad. Sci. U. S. A. 111, 6982-6987
42. Varkey, J., Isas, J. M., Mizuno, N., Jensen, M. B., Bhatia, V. K., Jao, C. C., Petrlova,
J., Voss, J. C., Stamou, D. G., Steven, A. C., and Langen, R. (2010) Membrane
curvature induction and tubulation are common features of synucleins and
apolipoproteins. J. Biol. Chem. 285, 32486-32493
43. Pandey, A. P., Haque, F., Rochet, J. C., and Hovis, J. S. (2011) alpha-Synuclein-
induced tubule formation in lipid bilayers. J. Phys. Chem. B 115, 5886-5893
44. Mizuno, N., Varkey, J., Kegulian, N. C., Hegde, B. G., Cheng, N., Langen, R., and
Steven, A. C. (2012) Remodeling of lipid vesicles into cylindrical micelles by
alpha-synuclein in an extended alpha-helical conformation. J. Biol. Chem. 287,
29301-29311
45. Varkey, J., Mizuno, N., Hegde, B. G., Cheng, N., Steven, A. C., and Langen, R.
(2013) alpha-Synuclein oligomers with broken helical conformation form
lipoprotein nanoparticles. J. Biol. Chem. 288, 17620-17630
46. Boassa, D., Berlanga, M. L., Yang, M. A., Terada, M., Hu, J., Bushong, E. A.,
Hwang, M., Masliah, E., George, J. M., and Ellisman, M. H. (2013) Mapping the
subcellular distribution of alpha-synuclein in neurons using genetically encoded
probes for correlated light and electron microscopy: implications for Parkinson's
disease pathogenesis. J. Neurosci. 33, 2605-2615
47. Bhatia, V. K., Madsen, K. L., Bolinger, P. Y., Kunding, A., Hedegard, P., Gether,
U., and Stamou, D. (2009) Amphipathic motifs in BAR domains are essential for
membrane curvature sensing. EMBO J. 28, 3303-3314
48. Madsen, K. L., Bhatia, V. K., Gether, U., and Stamou, D. (2010) BAR domains,
amphipathic helices and membrane-anchored proteins use the same mechanism
to sense membrane curvature. FEBS Lett. 584, 1848-1855
49. Cui, H., Lyman, E., and Voth, G. A. (2011) Mechanism of membrane curvature
sensing by amphipathic helix containing proteins. Biophys. J. 100, 1271-1279
72

50. Pranke, I. M., Morello, V., Bigay, J., Gibson, K., Verbavatz, J. M., Antonny, B., and
Jackson, C. L. (2011) alpha-Synuclein and ALPS motifs are membrane curvature
sensors whose contrasting chemistry mediates selective vesicle binding. J. Cell
Biol. 194, 89-103
51. Ouberai, M. M., Wang, J., Swann, M. J., Galvagnion, C., Guilliams, T., Dobson, C.
M., and Welland, M. E. (2013) alpha-Synuclein senses lipid packing defects and
induces lateral expansion of lipids leading to membrane remodeling. J. Biol.
Chem. 288, 20883-20895
52. Angelova, M. I., Soléau, S., Méléard, P., Faucon, J. F., and Bothorel, P. (1992)
Preparation of giant vesicles by external AC electric fields. Kinetics and
applications. Progress in Colloid & Polymer Science 89, 127-131
53. O'Brien, T. D., Butler, A. E., Roche, P. C., Johnson, K. H., and Butler, P. C. (1994)
Islet amyloid polypeptide in human insulinomas. Evidence for intracellular
amyloidogenesis. Diabetes 43, 329-336
54. Brender, J. R., Heyl, D. L., Samisetti, S., Kotler, S. A., Osborne, J. M., Pesaru, R. R.,
and Ramamoorthy, A. (2013) Membrane disordering is not sufficient for
membrane permeabilization by islet amyloid polypeptide: studies of IAPP(20-29)
fragments. Phys. Chem. Chem. Phys. 15, 8908-8915
55. Caillon, L., Lequin, O., and Khemtemourian, L. (2013) Evaluation of membrane
models and their composition for islet amyloid polypeptide-membrane
aggregation. Biochim. Biophys. Acta 1828, 2091-2098
56. Brender, J. R., Durr, U. H., Heyl, D., Budarapu, M. B., and Ramamoorthy, A.
(2007) Membrane fragmentation by an amyloidogenic fragment of human Islet
Amyloid Polypeptide detected by solid-state NMR spectroscopy of membrane
nanotubes. Biochim. Biophys. Acta 1768, 2026-2029
57. Gennis, R. B. (1989) Biomembranes : molecular structure and function, Springer-
Verlag, New York
58. Perkins, G., Renken, C., Martone, M. E., Young, S. J., Ellisman, M., and Frey, T.
(1997) Electron tomography of neuronal mitochondria: three-dimensional
structure and organization of cristae and membrane contacts. J. Struct. Biol. 119,
260-272
59. Frey, T. G., Renken, C. W., and Perkins, G. A. (2002) Insight into mitochondrial
structure and function from electron tomography. Biochim. Biophys. Acta 1555,
196-203
60. Zerbes, R. M., van der Klei, I. J., Veenhuis, M., Pfanner, N., van der Laan, M., and
Bohnert, M. (2012) Mitofilin complexes: conserved organizers of mitochondrial
membrane architecture. Biol. Chem. 393, 1247-1261
61. Rustenbeck, I., Matthies, A., and Lenzen, S. (1994) Lipid composition of glucose-
stimulated pancreatic islets and insulin-secreting tumor cells. Lipids 29, 685-692
73

62. Horvath, S. E., and Daum, G. (2013) Lipids of mitochondria. Prog. Lipid Res. 52,
590-614
63. Jao, C. C., Hegde, B. G., Chen, J., Haworth, I. S., and Langen, R. (2008) Structure
of membrane-bound alpha-synuclein from site-directed spin labeling and
computational refinement. Proc. Natl. Acad. Sci. U. S. A. 105, 19666-19671
64. Relini, A., Marano, N., and Gliozzi, A. (2014) Probing the interplay between
amyloidogenic proteins and membranes using lipid monolayers and bilayers.
Adv. Colloid Interface Sci. 207, 81-92
65. Terzi, E., Holzemann, G., and Seelig, J. (1997) Interaction of Alzheimer beta-
amyloid peptide(1-40) with lipid membranes. Biochemistry 36, 14845-14852
66. Atwal, R. S., Xia, J., Pinchev, D., Taylor, J., Epand, R. M., and Truant, R. (2007)
Huntingtin has a membrane association signal that can modulate huntingtin
aggregation, nuclear entry and toxicity. Hum. Mol. Genet. 16, 2600-2615
67. Last, N. B., Schlamadinger, D. E., and Miranker, A. D. (2013) A common
landscape for membrane-active peptides. Protein Sci. 22, 870-882
68. Bram, Y., Frydman-Marom, A., Yanai, I., Gilead, S., Shaltiel-Karyo, R.,
Amdursky, N., and Gazit, E. (2014) Apoptosis induced by islet amyloid
polypeptide soluble oligomers is neutralized by diabetes-associated specific
antibodies. Sci. Rep. 4, 4267
69. Kumar, S., Schlamadinger, D. E., Brown, M. A., Dunn, J. M., Mercado, B., Hebda,
J. A., Saraogi, I., Rhoades, E., Hamilton, A. D., and Miranker, A. D. (2015) Islet
Amyloid-Induced Cell Death and Bilayer Integrity Loss Share a Molecular Origin
Targetable with Oligopyridylamide-Based alpha-Helical Mimetics. Chem. Biol. 22,
369-378

74

Chapter 4

Circular Dichroism Uncovers Temperature- and Polyglutamine-Dependent Structural
Changes in Huntingtin Exon 1

Partly extracted from Fodale, V., Kegulian, N. C., Verani, M., Cariulo, C., Azzollini, L.,
Petricca, L., Daldin, M., Boggio, R., Padova, A., Kuhn, R., Pacifici, R., Macdonald, D.,
Schoenfeld, R. C., Park, H., Isas, J. M., Langen, R., Weiss, A., and Caricasole, A. (2014)
Polyglutamine- and temperature-dependent conformational rigidity in mutant
huntingtin revealed by immunoassays and circular dichroism spectroscopy. PLoS One 9,
e112262.

75

4.1 Introduction
Huntingtin (HTT) is a multifunctional, polyglutamine (polyQ)-containing protein in
which a trinucleotide repeat-encoded expansion of the polyQ tract leads to the
occurrence of Huntington’s disease (HD), a devastating, fatal neurodegenerative
disorder (see Chapter 1 for full review and citations). Recent findings have suggested
that HTT with an expanded polyQ
region has an elevated conformational
rigidity as compared to HTT containing
a normal number of glutamine residues
(Q-length) (1,2) (also summarized in
Chapter 1). However, those fluorescence
lifetime- and immunoassay-based
discoveries did not yield a full readout
delineating the nature of the
conformation in the longer polyQ
constructs that appeared to be “stuck
on.” For this reason, we chose to analyze
HTT exon 1 (HDx1), the segment of HTT
comprising the polyQ region flanked by a largely α-helical, 17-aa N-terminal (N17)
domain and a proline-rich C-terminal region (as reviewed in Chapter 1), by circular
dichroism (CD), a spectroscopic method that directly measures secondary structure.
To gauge secondary structure of proteins and other molecules, CD relies upon
the fact that linearly polarized light is composed of left-handed and right-handed
circularly polarized light of equal amplitudes to one another (3). When linearly
polarized light passes through an optically active medium (such as proteins or nucleic
acids folded into structures that themselves possess “handedness”), its left-handed and
right-handed circularly polarized components are absorbed to different extents (4). The

Fig. 4-1. Typical CD spectra designating the
indicated protein secondary structures.
Source: Ohio State University Department of
Chemistry.
https://chemistry.osu.edu/~grenkes/STATEME
NT%202/CD/ExampleCDSpectra.gif

76

result is elliptically polarized light, which yields a measure of ellipticity as the readout
at different wavelengths (4). Consequently, CD spectra showing different secondary
structures are distinct for one another (3). For proteins, random coil, α-helix, β-sheet,
type I β-turn, and polyproline II (PPII) helix structures have individual signature CD
spectra (Fig. 4-1). In practice, proteins usually contain two or more of these secondary
structures and will manifest spectra comprising components of different signature
spectra, which can then be quantified (3). In our study, we thus utilized CD to quantify
the different secondary structure components that changed with respect to temperature
within each of the HDx1 proteins that we used, which bore a thioredoxin (THRX)
moiety fused to the N-terminal end to improve their stability and solubility. We found
lowering the temperature to unmask an increased propensity in polyQ-expanded HDx1
to adopt an α-helical structure that could not solely be accounted for by the N17
residues and most likely includes part of the polyQ tract. We also tested the
temperature-dependent CD of N-terminal fragments comprising the first 548 residues
of wild-type HTT (N548 HTT) and found the conformational effects of Q-length and
temperature to follow a similar trend. In addition, we tested the effects of mimicking
phosphorylation at serine residues 13 and 16 on our protein constructs’ temperature-
dependent CD profiles and found phosphomimicry at position 16 to have a potentially
therapeutic effect. The rationale behind mimicking phosphorylation at this site and at
position 13 is that not only do post-translational modifications exert a wide range of
effects on the functionality and toxicity of HTT and other polyQ-containing proteins (5),
but in particular, phosphorylation of HTT at serine residues 13 and 16 has been
implicated in changes in its intracellular localization, cytoskeletal organization, and
interactions with binding partners (6-9); phosphomimetic mutations at those sites
resulted in decelerated fibrillization kinetics (10) and amelioration of HD-like
symptoms in transgenic mice (11). Our results could have profound implications in the
development of pharmaceuticals to treat HD.
77

4.2 Methods
Materials
Purified N548 HTT proteins with Q19, Q22, Q25, Q42, and Q52 were kindly provided
by CHDI. Purified N548 HTT proteins with Q55 were kindly provided by IRBM
Promidis.
THRX-HDx1 protein expression and purification
C-terminal His tag-containing THRX-HDx1-Q7, Q25, Q46, and Q55 were expressed by
the previously used pET32a parent constructs (12,13). To generate phosphomimetic
mutants, aspartic acid mutations were introduced by site-directed mutagenesis at
positions 13 and 16 of the N17 region, two of the known sites of HDx1 phosphorylation
(6). Expression and subsequent lysis of cell pellets were performed as previously
described (12). Then, similarly to the previously described protocol, proteins were
purified by centrifuging the lysates at 19,000 × g for 30 minutes; incubating with nickel-
nitrilotriacetic acid-agarose beads (Qiagen) on a rocker at 4°C for 1 hour; washing with
several column volumes of 20 mM Tris-HCl, pH 8.0, 300 mM NaCl, 50 mM imidazole;
and eluting with 20 mM Tris-HCl, pH 7.4, 300 mM NaCl, 250 mM imidazole. Eluted
proteins were diluted in 10 mM Tris, pH 7.4, then purified on a HiTrap Q XL column
(GE Healthcare) with an AKTA FPLC system (Amersham Pharmacia Biotech), taking 1-
mL fractions and using a phosphate-buffered saline gradient from 20 mM sodium
phosphate, pH 7.4, to 20 mM sodium phosphate, pH 7.4, 1 M NaCl. For isolation of the
THRX fusion partner, the fusion protein was cleaved with EKmax (Life Technologies,
Grand Island, NY) at room temperature for 30 minutes. Then, 1 M urea was added to
the resulting fragments, which were subsequently applied to nickel-nitrilotriacetic acid-
agarose beads and briefly centrifuged in a tabletop Eppendorf 5415D centrifuge at
13,200 rpm. The supernatant was diluted 1:10 in 20 mM Tris, pH 7.4, and further
purified by FPLC as described for THRX-HDx1 above. Prior to each experiment, the
protein concentration was measured using the A280 of the sample.
78

Temperature-controlled CD
CD was performed using a Jasco 815 spectropolarimeter (Jasco Inc., Easton, MD).
Temperature was regulated by a Jasco PFD-425S Peltier type FDCD attachment
connected to a PolyScience recirculator (PolyScience, Niles, IL). For spectra at each
temperature, measurements were taken every 1 nm from 200 to 260 nm, scanning at 50
nm/minute and an averaging time of 1 second. For spectra spanning from 190 to 260
nm, protein was eluted into 20 mM sodium phosphate, pH 7.4, no salt, via a PD-10
column (GE Healthcare) and CD measurements taken with the parameters denoted
above. Ten scans were averaged for each sample spectrum; background spectra were
obtained by averaging twenty scans and the appropriate ones were subtracted from the
respective sample spectra. Spectra were smoothed by the Savitsky-Golay algorithm.
Mean residue ellipticity (MRE) was determined for each reading by dividing the
observed CD reading by the protein concentration, path length, and number of
residues. Single-wavelength readings at 222 nm were obtained at –10°C, 0°C, 4°C, 10°C,
20°C, 30°C, and 37°C. For each THRX-HDx1 sample, ellipticity was measured every 1
second for 300 seconds; the 301 readings for each sample at each temperature were
averaged. The same was done for the cleaved THRX control and the weighted
contribution of the THRX moiety subtracted from the averaged single-wavelength
reading for each THRX-HDx1 sample at each temperature as follows:

(4.1)
where obsMRE222, fus.prot. is the MRE observed at 222 nm at a given temperature for each
sample of THRX-HDx1 fusion protein, obsMRE222, THRX is the MRE observed at 222 nm at
that same temperature for THRX alone, the number of amino acids in THRX is equal to
131, and the number of amino acids in the fusion protein varies by Q-length.
Analysis of CD data
79

The number of amino acids in each HDx1 sample to experience a change in helicity
from –10°C to 37°C was estimated using a previously developed helix-coil transition
model, which gives the change in fraction of helicity (ΔfH) by the following equation
(14,15):

(4.2)
where fH at each temperature is given as follows:

(4.3)
where

(4.4)
and

(4.5)
given that T is the temperature in °C and Nr is the number of residues.
4.3 Results
Temperature-dependent α-helix formation in THRX-HDx1 is dependent upon Q-
length
CD data on our HTT protein constructs were collected at a wide range of temperatures
(–10 to 37°C) that encompassed the conditions employed in the temperature-controlled
Förster resonance energy transfer (FRET) assays described in Chapter 1. THRX-HDx1-
Q7, Q25, Q46, and Q55 had spectra shown in Fig. 4-2A-D displaying significant,
reversible changes with temperature and an isosbestic point near 203 nm, which is
commonly observed for conversions between random coil and α-helical structure. The
difference spectra for all four proteins exhibit minima at 208 and 222 nm (Fig. 4-3A),
which are characteristic of α-helical structure (as illustrated in Fig. 4-1) and which in
this case, since these are difference spectra, indicate increased formation of α-helical
structure at lower temperatures.
80

Another key feature of α-helical CD spectra is a maximum near 190 nm (as
illustrated in Fig. 4-1), which was not possible to view at high salt concentrations. We
therefore recorded CD spectra for our four constructs in salt-free phosphate buffer.
These spectra (Fig. 4-4A-D) also show isosbestic points near 203 nm and reversibility of
the temperature effect on conformation, and the difference spectra show the peaks near
190, 208, and 222 nm that are indicative of an α-helix (Fig. 4-3B).
Two controls were in order: whether the observed temperature-dependent
conformational change was intramolecular or relied on intermolecular interactions

Fig. 4-2. Temperature-dependent CD spectra for HDx1 fusion proteins of four different Q-
lengths. CD spectra for THRX-HDx1 Q7 (A), Q25 (B), Q46 (C), and Q55 (D) at all the
temperatures tested showing an isosbestic point around 203 nm (arrows) and showing the
spectra for each construct at –10˚C after measurements had been taken at 37˚C to be nearly
identical to those viewed beforehand, indicating the reversibility of the temperature effect.
Parts B and C from Fodale et al., 2014.
81

needed to be
tested, and it had
to be verified that
the residues
showing the
conformational
change were part
of HDx1 and not
the THRX moiety.
Evaluating the
MRE values at 222
nm of two of our
constructs at three
temperatures over
a range of
concentrations showed that concentrations did not affect the temperature-dependent
conformational change (Fig. 4-5A), ruling out intermolecular interactions as the cause
for the temperature-dependent effect. Cleaving off the HDx1 moiety left THRX to yield
virtually identical spectra across temperatures (Fig. 4-5B), indicating that HDx1 was the
segment of our construct that changed conformations in each case.
The increased formation of α-helical structure at lower temperatures signified by
the difference spectra in Fig. 4-3A and B appears more pronounced for each successive
Q-length in both buffer compositions. To better view this differential effect, we plotted
the MRE values obtained at 222 nm for all our THRX-HDx1 constructs minus the
weighted contribution of the THRX moiety (see Methods for explanation) versus
temperature. Using this measure, we can clearly see that cooling increases helicity more
strongly for constructs with higher Q-lengths (Fig. 4-6). To quantify each increase in

Fig. 4-3. CD difference spectra between temperature extremes.
Spectra from THRX-HDx1 of the indicated Q-lengths gleaned from
subtracting spectra measured at 37°C from spectra measured at –10°C
in phosphate buffered saline (A) and in salt-free phosphate buffer (B).
Q25 and Q46 traces from Fodale et al., 2014.
82

helicity, we applied a previously developed helix-coil model as one means to calculate
the number of residues that experienced a change in helicity (14,16), since the consistent
isosbestic point across temperatures (Fig. 4-2A-D and 4-4A-D) and the typically α-
helical difference spectra (Fig. 4-3A and B) suggest that the conformational changes are
dominated by a helix-coil transition, justifying this method of calculation. Using this
model, we estimate that on the order of 11, 23, 31, and 35 residues become helical upon
cooling from 37 to –10°C for the Q7, Q25, Q46, and Q55 constructs, respectively. This
suggests that the observed change in helicity in the constructs of longer polyQ region

Fig. 4-4. Temperature-dependent CD spectra for HDx1 fusion proteins of four different Q-
lengths in salt-free phosphate buffer. CD spectra measured in salt-free phosphate buffer for
THRX-HDx1 Q7 (A), Q25 (B), Q46 (C), and Q55 (D) at all the temperatures tested showing an
isosbestic point around 203 nm (arrows) and showing the spectra for each construct at –10˚C
after measurements had been taken at 37˚C to be nearly identical to those viewed beforehand,
indicating the reversibility of the temperature effect. Parts B and C from Fodale et al., 2014.
83

cannot exclusively stem from structural changes in the N17 region and likely involves
the polyQ region as well. The proline-rich C-terminal region can adopt a PPII structure
(17-19), a conformation whose stability may also increase with decreasing temperature
(12,20). However, the CD spectrum of a PPII structure is entirely different from that of
an α-helix (Fig. 4-1). While we cannot exclude that more subtle changes might also have
occurred in the polyproline region, the structural changes observed by CD are
dominated by α-helical structure (Fig. 4-3A and B). Although the temperature-
dependent change in helicity increases with Q-length, it is important to note that the
overall helicity still remains higher at higher Q-lengths at 37°C (Fig. 4-6). Thus, polyQ-
expanded HDx1 has enhanced structural ordering at all temperatures, including
physiologically relevant temperatures.
Temperature-dependent α-helix formation in N548 HTT is dependent upon Q-length
The temperature-controlled FRET experiments (2) described in Chapter 1 utilized N548
HTT fragments. To ascertain that such fragments would display CD profiles compatible
with our findings on HDx1, we performed CD analysis on N548 HTT with a number of
different Q-lengths and found a similar trend as in THRX-HDx1, although the impact of
the polyQ region on CD measurements of the larger fragments is significantly diluted

Fig. 4-5. CD controls showing temperature effect is not concentration- or THRX-dependent.
(A) MRE values at 222 nm for THRX-HDx1-Q25 and Q46 at indicated concentrations. (B) CD
spectra for the THRX fusion partner alone at indicated temperatures. From Fodale et al., 2014.
-14
-12
-10
-8
-6
-4
-20 0 20 40
[ θ]
MRE
10
-3
deg cm
2
dmol
-1
222 nm
Temperature ( C)
THRX-HDx1
Q25 1.9 μM
Q25 3.8 μM
Q25 7.6 μM
Q25 19 μM
Q46 1.6 μM
Q46 3.7 μM
Q46 20 μM
-8
-6
-4
-2
0
2
4
6
8
10
200 220 240 260
[ θ]
MRE
10
-3
deg cm
2
dmol
-1
Wavelength (nm)
THRX Tag
-10 deg
0 deg
4 deg
10 deg
20 deg
37 deg
A B
84

out. For example, N548 Q25 HTT spectra at
–10°C and at 37°C show increased helicity
with decreased temperature (Fig. 4-7A).
Also, we again found MRE values to
become more negative, indicating
increased α-helical structure, with
decreasing temperature. In fact, we
observed a similar influence by Q-length
on N548 HTT α-helical content (Fig. 4-7B)
as on THRX-HDx1.
A phosphomimetic mutation at position
16 could result in reduced temperature-
dependent α-helix formation
In order to mimic phosphorylation at
positions 13 and 16, we mutated the serine
residues at those positions to aspartic acid

Fig. 4-6. MRE values at 222 nm at each
temperature tested for THRX-HDx1 of the
indicated Q-lengths, with weighted
THRX MRE values subtracted. All data
are represented as mean ± S.D. of three
independent experiments. Q25 and Q46
traces adapted from Fodale et al., 2014.

Fig. 4-7. Temperature-dependent α-helical variations are also observed in untagged, larger
N548Htt purified recombinant proteins. (A) MRE spectra for the Q25 N548Htt protein at –
10°C and 37°C, showing that the temperature effect can still be seen in this longer HTT
fragment. (B) MRE values at 222 nm at each indicated temperature relative to MRE values at
37°C for N548Htt of increasing Q-length (Q19, Q22, Q25, Q42, Q52, Q55). From Fodale et al.,
2014.
-10
-8
-6
-4
-2
0
2
200 220 240 260
[ θ]
MRE
10
-3
deg cm
2
dmol
-1
Wavelength (nm)
N548Htt Q25
37 deg
-10 deg
0.9
0.95
1
1.05
1.1
1.15
1.2
Q19 Q22 Q25 Q33 Q42 Q52 Q55
222-nm CD signal relative to
that at 37 C
N548Htt
-10 deg
0 deg
4 deg
10 deg
20 deg
A B
85

by site-directed
mutagenesis (see
Methods). The S16D
mutation in our THRX-
HDx1 constructs resulted
in the Q46 construct
having a distribution of
THRX-weighted MRE
values at 222 nm
resembling that of THRX-
HDx1-Q25 that does not
bear such a mutation, and
the Q25 construct having a
CD profile resembling that
of unmutated Q7 (Fig. 4-
8). In other words, mimicking phosphorylation at this site reduces the enhanced
structural ordering conferred by polyQ expansion. In contrast, mimicking
phosphorylation in THRX-HDx1-Q25 and THRX-HDx1-Q46 at serine 13 (Fig. 4-9) or
simultaneously at serine 13 and serine 16 (Fig. 4-10) did not appear to result in a
significant change in CD profile from the CD profiles of the non-phosphomimetic
counterparts of those constructs.
4.4 Discussion
In this study, we utilized CD to identify the temperature-sensitive secondary structure
taken by HDx1 to be α-helical and to observe that such structure forms to a greater
extent in polyQ-expanded HDx1. These experiments succeed findings using FRET that
polyQ-expanded HDx1 experiences reduced intramolecular interaction between its N-
and C-termini in vivo (1) and that its N17 and polyQ regions in contrast have elevated

Fig. 4-8. Effect of the S16D phosphomimetic mutation on
MRE values at 222 nm at each temperature tested for THRX-
HDx1 of the indicated Q-lengths, with weighted THRX MRE
values subtracted.
86

interactions that
increased temperature
does not abrogate in vitro
(2). Taken together, these
results suggest reduced
flexibility in polyQ-
expanded HDx1, as
HDx1 of normal Q-length
showed relatively high
levels of N-terminus-C-
terminus intramolecular
interaction in vivo (1), as
well as the ability to lose
its N17-polyQ interaction
in vitro in response to
elevated temperature (2), indicating a propensity in HDx1 of normal Q-length to adopt
a range of conformations rather than remaining “stuck” in one. Our CD data have
fleshed out the abstract suggestions by the FRET data of increased structure in the
polyQ-expanded proteins, by clarifying that this increased structure indeed exists and is
α-helical in nature. Since the N17 region is mostly known to be α-helical and the
number of amino acids in α-helical form significantly exceeds 17 in the polyQ-expanded
constructs, we can easily hypothesize that the helix in the N17 region in these cases
expands into the polyQ tract. CD spectra do not show the polyproline region in the C-
terminus to be affected, leaving the polyQ tract as the prime suspect for containing the
additional α-helical structure. Studies that zone in on specific residues are needed to
confirm this hypothesis and this need is addressed using electron paramagnetic

Fig. 4-9. Effect of the S13D phosphomimetic mutation on
MRE values at 222 nm at each temperature tested for THRX-
HDx1 of the indicated Q-lengths, with weighted THRX MRE
values subtracted.
87

resonance (EPR) with site-
directed spin labeling
(SDSL) in the experiments
described in Chapter 5.
The data showing
an S16D mutation to
attenuate the expanded
polyQ region-associated
α-helical structure are
preliminary and should be
applied to more
constructs. However, one
can already envision the
reason that the effects of
post-translational
modifications or of mimics thereof on HTT have been so convoluted and contradictory,
such that while mimicking phosphorylation at residues 13 and 16 ameliorated the HD-
like syndrome exhibited by transgenic mice (11), the presumably “healthy” FRET-
observed interaction between the N- and C-termini of HDx1 of normal Q-length was
abrogated by phosphorylation at these same sites as well as by mimicry thereof (1). The
reason might be opposing effects by each individual mutation or phosphorylation.
While S16D appears to have a therapeutic effect on conformation (Fig. 4-8), not only
does S13D appear to have no effect (Fig. 4-9), but also the double mutation S13D/S16D
does not correct the polyQ expansion-associated structural enhancement (Fig. 4-10).
Phosphorylation or phosphomimicry at position 13 could have a detrimental effect on
the seemingly therapeutic effect of phosphorylation or phosphomimicry at position 16.
Such cross-talk should be examined further.

Fig. 4-10. Effect of the S13D/S16D double phosphomimetic
mutation on MRE values at 222 nm at each temperature
tested for THRX-HDx1 of the indicated Q-lengths, with
weighted THRX MRE values subtracted.
88

Altogether, this study, by showing dramatic effects of temperature on HDx1
conformation, demonstrates a great propensity in HDx1 conformation for modulation
by external factors and thereby suggests tremendous potential for pharmaceuticals that
act as conformational modulators to emerge as HD therapeutics.
89

4.5 References
1. Caron, N. S., Desmond, C. R., Xia, J., and Truant, R. (2013) Polyglutamine
domain flexibility mediates the proximity between flanking sequences in
huntingtin. Proc. Natl. Acad. Sci. U. S. A. 110, 14610-14615
2. Fodale, V., Kegulian, N. C., Verani, M., Cariulo, C., Azzollini, L., Petricca, L.,
Daldin, M., Boggio, R., Padova, A., Kuhn, R., Pacifici, R., Macdonald, D.,
Schoenfeld, R. C., Park, H., Isas, J. M., Langen, R., Weiss, A., and Caricasole, A.
(2014) Polyglutamine- and temperature-dependent conformational rigidity in
mutant huntingtin revealed by immunoassays and circular dichroism
spectroscopy. PLoS One 9, e112262
3. Holde, K. E. v., Johnson, W. C., and Ho, P. S. (2006) Principles of Physical
Biochemistry, 2nd ed., Pearson Prentice Hall, Upper Saddle River, NJ
4. Tinoco, I., Sauer, K., Wang, J. C., and Puglisi, J. D. (2002) Physical chemistry :
principles and applications in biological sciences, 4th ed., Prentice Hall, Upper Saddle
River, N.J.
5. Ehrnhoefer, D. E., Sutton, L., and Hayden, M. R. (2011) Small changes, big
impact: posttranslational modifications and function of huntingtin in Huntington
disease. Neuroscientist 17, 475-492
6. Maiuri, T., Woloshansky, T., Xia, J., and Truant, R. (2013) The huntingtin N17
domain is a multifunctional CRM1 and Ran-dependent nuclear and cilial export
signal. Hum. Mol. Genet. 22, 1383-1394
7. Caron, N. S., Hung, C. L., Atwal, R. S., and Truant, R. (2014) Live cell imaging
and biophotonic methods reveal two types of mutant huntingtin inclusions.
Hum. Mol. Genet. 23, 2324-2338
8. Khoshnan, A., and Patterson, P. H. (2011) The role of IkappaB kinase complex in
the neurobiology of Huntington's disease. Neurobiol. Dis. 43, 305-311
9. Atwal, R. S., Desmond, C. R., Caron, N., Maiuri, T., Xia, J., Sipione, S., and
Truant, R. (2011) Kinase inhibitors modulate huntingtin cell localization and
toxicity. Nat. Chem. Biol. 7, 453-460
10. Mishra, R., Hoop, C. L., Kodali, R., Sahoo, B., van der Wel, P. C., and Wetzel, R.
(2012) Serine phosphorylation suppresses huntingtin amyloid accumulation by
altering protein aggregation properties. J. Mol. Biol. 424, 1-14
11. Gu, X., Greiner, E. R., Mishra, R., Kodali, R., Osmand, A., Finkbeiner, S., Steffan,
J. S., Thompson, L. M., Wetzel, R., and Yang, X. W. (2009) Serines 13 and 16 are
critical determinants of full-length human mutant huntingtin induced disease
pathogenesis in HD mice. Neuron 64, 828-840
12. Bugg, C. W., Isas, J. M., Fischer, T., Patterson, P. H., and Langen, R. (2012)
Structural features and domain organization of huntingtin fibrils. J. Biol. Chem.
287, 31739-31746
90

13. Bennett, M. J., Huey-Tubman, K. E., Herr, A. B., West, A. P., Jr., Ross, S. A., and
Bjorkman, P. J. (2002) A linear lattice model for polyglutamine in CAG-expansion
diseases. Proc. Natl. Acad. Sci. U. S. A. 99, 11634-11639
14. Luo, P., and Baldwin, R. L. (1997) Mechanism of helix induction by
trifluoroethanol: a framework for extrapolating the helix-forming properties of
peptides from trifluoroethanol/water mixtures back to water. Biochemistry 36,
8413-8421
15. Jayasinghe, S. A., and Langen, R. (2004) Identifying structural features of fibrillar
islet amyloid polypeptide using site-directed spin labeling. J. Biol. Chem. 279,
48420-48425
16. Rohl, C. A., and Baldwin, R. L. (1997) Comparison of NH exchange and circular
dichroism as techniques for measuring the parameters of the helix-coil transition
in peptides. Biochemistry 36, 8435-8442
17. Darnell, G., Orgel, J. P., Pahl, R., and Meredith, S. C. (2007) Flanking polyproline
sequences inhibit beta-sheet structure in polyglutamine segments by inducing
PPII-like helix structure. J. Mol. Biol. 374, 688-704
18. Kim, M. W., Chelliah, Y., Kim, S. W., Otwinowski, Z., and Bezprozvanny, I.
(2009) Secondary structure of Huntingtin amino-terminal region. Structure 17,
1205-1212
19. Darnell, G. D., Derryberry, J., Kurutz, J. W., and Meredith, S. C. (2009)
Mechanism of cis-inhibition of polyQ fibrillation by polyP: PPII oligomers and
the hydrophobic effect. Biophys. J. 97, 2295-2305
20. Ma, K., and Wang, K. (2003) Malleable conformation of the elastic PEVK segment
of titin: non-co-operative interconversion of polyproline II helix, beta-turn and
unordered structures. Biochem. J. 374, 687-695

91

Chapter 5

Locating Enhanced Structural Ordering in Polyglutamine-Expanded Huntingtin Exon 1
by Electron Paramagnetic Resonance

Work done in collaboration with Dr. J. Mario Isas.

92

5.1 Introduction
Chapter 4 delineates the discovery via circular dichroism (CD) of increased
temperature-dependent α-helix formation in polyglutamine (polyQ)-expanded
huntingtin (HTT) exon 1 (HDx1). Using electron paramagnetic resonance (EPR) with
site-directed spin labeling (SDSL), we have now located the regions in this protein
where additional glutamine residues result in conformational rigidity. To this end, we
generated a variety of single cysteine mutant derivatives of the N-terminal thioredoxin
(THRX)-fused constructs of HDx1 containing 7, 16, 25, 46, and 55 glutamine residues in
their respective polyQ regions and thus had a variety of sites at which we could attach
spin label and obtain an EPR readout. As a result, I find that enhanced structural
ordering in response to lowered temperature occurs in the 17-aa N-terminal (N17)
region and the part of the polyQ tract that is closer to the N17 region, and this
heightened rigidity is more severe with greater Q-length.
5.2 Methods
Protein expression, labeling and purification
C-terminal His tag-containing THRX-HDx1-Q7, Q16, Q25, Q46, and Q55 carrying single
cysteine mutations generated by site-directed mutagenesis were expressed in the same
pET32a parent constructs and using the same protocol as indicated in Chapter 4.
Purification from protein-expressing bacterial cell pellets entailed including 1 mM
dithiothreitol (DTT) in the lysis buffer; washing nickel-nitrilotriacetic acid-agarose
beads containing bound protein with 20 mM Tris-HCl, pH 7.4, 300 mM NaCl, 20 mM
imidazole; and spin labeling protein by adding S-(2,2,5,5-tetramethyl-2,5-dihydro-1H-
pyrrol-3-yl)methyl methanesulfonothioate (MTSL) at a threefold molar excess
immediately upon elution of protein from nickel-nitrilotriacetic acid-agarose beads into
20 mM Tris-HCl, pH 7.4, 300 mM NaCl, 250 mM imidazole. Further purification using a
phosphate-buffered saline gradient via an AKTA FPLC system (Amersham Pharmacia
93

Biotech) with a HiTrap Q XL column (GE Healthcare) was performed as described in
Chapter 4.

Fig. 5-1. Sample temperature-
dependent EPR spectra for
THRX-HDx1-Q46 spin labeled
at the indicated sites on HDx1.
Mobile (M) and immobile (I)
components are labeled where
applicable. Temperatures
tested were, from top to
bottom, –20, –10, 0, 4, 10, 15, 20,
25, 30, and 37°C.

94

Continuous-wave EPR spectra
Protein concentrations were adjusted to between 15 and 20 μM to maximize EPR signal
while minimizing the risk of aggregation. Proteins were subsequently loaded into
borosilicate capillaries (0.6-mm inner diameter × 0.84-mm outer diameter, VitroCom,
Mt. Lakes, NJ); EPR spectra were recorded on an X-band Bruker EMX spectrometer
(Bruker Biospin Corporation) with a scan width of 100 gauss and an incident
microwave power of 12.60 milliwatts. Spectral recordings were performed at –20, –10, 0,
4, 10, 15, 20, 25, 30, and 37°C. Temperature was controlled using a Bruker ER 4131VT
variable temperature accessory. Spectra were processed for analysis using the Bruker
suite of software programs.
5.3 Results
In THRX-HDx1 of expanded polyQ,
residual structure forms in N17 and
the N-terminal region of the polyQ
tract
To locate the temperature-dependent
structural changes in HDx1, we
generated a number of spin-labeled
derivatives of our previously used
THRX-HDx1-Q46 construct (see
Chapter 4) and viewed their EPR
spectra at temperatures ranging from
–20 to 37°C. Fig. 5-1 shows sample
spectra from spin labels resonating at
representative loci. Spectra from
mutants with N17 residues labeled

Fig. 5-2. EPR spectra measured at the indicated
temperatures for Q55-containing THRX-HDx1
spin labeled at residue 35 (within the polyQ
stretch) or 75 (within the polyproline region).
Mobile (M) and immobile (I) components are
indicated where applicable.

95

Fig. 5-3. Ms plot for THRX-HDx1-Q46 spin labeled at the
indicated positions on HDx1 at –20 and 25°C.
showed broad lines at low temperature that grew sharp, as well as an immobile
component (I) that diminished and gave way to the mobile component (M), as
temperature was increased. Spectra from mutants with “early” (closer to the N-
terminus) polyQ residues labeled showed a similar pattern in response to temperature.
In contrast, spectra closer to the C-terminus, whether in the “late” polyQ tract or in the
C-terminal proline-rich region, showed a general line-sharpening trend with increasing
temperature but no immobile component even at the lowest temperature tested. While
increased rigidity with
decreasing temperature
can always be expected,
a peak designating an
immobile component
signifies inaccessibility
of the spin label due to a
rigid structure in or
around the labeled
residue. Therefore,
increased structuring
unmasked by lowered
temperature exists in N17 and the N-terminal side of the polyQ stretch of HDx1. The
separation between mobile and immobile components can be seen as dramatically in
THRX-HDx1-Q55, the other expanded polyQ construct that we analyzed, such that, for
instance, spin label at position 35, which is situated “early” in the polyQ region, shows
a strong elevation of the immobile component at low temperatures (Fig. 5-2, left panel);
in contrast, this same construct spin labeled at position 75 gives EPR spectra with sharp
lines that simply dull with lowered temperature but do not show significant separation
of components (Fig. 5-2, right panel).
96

Fig. 5-4. Ms for the indicated spin labeled residues of HDx1 within THRX-HDx1 protein
constructs of the indicated Q-lengths.
Another manner of gauging mobility of spin-labeled sites analyzed by EPR is by
plotting the linewidths of their central resonances. Broader central linewidths indicate
lower mobility of the site; higher values for inverse central linewidth indicate higher
mobility. Normalizing the inverse central linewidths (δ
-1
) to scaled mobility (Ms) using
the equation Ms = (δ
-1
– δi
-1
) / (δm
-1
– δi
-1
) (1), where δi
-1
and δm
-1
are the inverse central
linewidths of the spin label at the most immobile and mobile sites, respectively, of the
97

THRX-HDx1-Q46 construct at each temperature, we obtain the scaled mobility plot
shown in Fig. 5-3. The mobilities of different residues at 25°C show that residues above
a point between positions 21 and 30, a part of the polyQ stretch close to the N-terminus,
possess a higher level of mobility than those below position 21, with the exception of 1.
In contrast, at –20°C, the “leap” to that higher mobility does not occur until position 62,
at the C-terminal end of the polyQ stretch. This difference between temperatures in
position of the “leap” could account for the increased α-helicity observed by CD at
lower temperatures for constructs of this Q-length (see Chapter 4), such that the stretch
of polyQ residues that remain ordered at low temperature as viewed by EPR are the
ones that account for the typically α-helical difference spectra between high and low
temperatures as measured by CD (as in Fig. 4-3). Comparing Ms across different

Fig. 5-5. Sample temperature-dependent EPR spectra for THRX-HDx1-Q25 spin labeled at
the indicated sites on HDx1. Mobile (M) and immobile (I) components are labeled where
applicable. Temperatures tested were, from top to bottom, –20, –10, 0, 4, 10, 15, 20, 25, 30, and
37°C. Spectra were measured using scan widths of 150 G.
98

Fig. 5-6. Overlay of EPR spectra for THRX-HDx1-Q25 spin labeled at
the indicated sites on HDx1 and spectra measured at 4–10°C higher
temperatures for THRX-HDx1-Q46 spin labeled at the same sites.

Q46 Q25 position 11 Q46 Q25 position 35 Q46 Q25 position 5
temperatures (Fig. 5-4), we assess that intermediately positioned polyQ residues
respond to temperature by increasing gradually in mobility from –20 to 37°C, while N17
and polyproline region residues as well as residues on either end of the polyQ tract
change relatively little across temperatures, with the N-terminal residues having the
most mobility overall and the C-terminal residues having the least.
THRX-HDx1 constructs with shorter polyQ stretches also show structuring at their N-
terminal ends but show less temperature sensitivity
Sample spectra
from spin-labeled
THRX-HDx1-Q25
(Fig. 5-5) show an
immobility
component (I) at
low temperatures
for positions 5 and
11, which are in the
N17 region, and for
position 35, which
is near the middle
of the polyQ tract,
but no immobility component in the polyproline-rich C-terminus, as exemplified by the
spectra at position 50. While the apparent gain in structure in N-terminal residues at
decreased temperature (and lack thereof in C-terminal residues, as exemplified by
residue 50) may at first glance seem equivalent to the profile of the Q46 construct, there
is a temperature shift between the two profiles. Specifically, upon superimposing
spectra from THRX-HDx1-Q46 spectra over spectra from THRX-HDx1-Q25 where the
latter were measured at 10°C-lower (or 4–7°C-lower at higher temperatures, at which
99

spectra grew more similar) temperatures, the spectra from the different constructs
overlap almost perfectly (Fig. 5-6). This observation suggests that the addition 21 Q
residues rigidify HDx1 at those sites to an extent close to that at which lowering the
temperature by 4–10°C (depending on the temperature range) would.

Fig. 5-7. Sample
temperature-
dependent EPR
spectra for
THRX-HDx1-Q7
and Q16 spin
labeled at the
indicated sites on
HDx1. Mobile
(M) and immobile
(I) components
are indicated
where applicable.

100

Spectra from THRX-HDx1
constructs of still shorter Q-
lengths (Fig. 5-7) show
temperature-dependent
rigidifying of the N17 and
“early” polyQ regions in a
similar pattern to that observed
for longer polyQ constructs, yet
they generally show more
mobility across temperatures
and loci. Superimposing spectra
from an N-terminal spin labeled
residue from constructs of four
different Q-lengths shows each
construct to require a
temperature shift of approximately 5°C higher in order to obtain similar spectra to those
from the construct that has approximately 10 fewer Q residues in the polyQ tract
(example of position 21 shown in Fig. 5-8).
5.4 Discussion
This EPR study shows increased temperature-dependent conformational rigidity in the
N17 region and on the N-terminal side of the polyQ region of HDx1 of expanded
polyQ. This finding confirms the suspicion brought up in Chapter 4 that increased α-
helical structure observed by CD stems from α-helicity in the N17 region extending into
the polyQ tract in cases of polyQ expansion. THRX-HDx1 constructs of five different Q-
lengths were examined for this study, and temperature-dependent rigidity correlated
positively with Q-length across all five, even those of normal Q-length, in keeping with
CD-obtained findings described in Chapter 4 showing the Q7 construct to form much

Fig. 5-8. Temperature-shifted overlays of EPR spectra
for four THRX-HDx1 constructs of the indicated Q-
lengths spin labeled at residue 21 of HDx1.
101

less α-helical structure than the Q25 construct. This suggests that the rigidity associated
with Q-length lies on a continuum rather than on an on/off basis, even though in
practice a disease threshold exists. Apparently the rigidity of HTT at disease threshold
reaches a point at which it can no longer fulfill its normal functions and (in cases of
heterozygosity) interferes with the functionality of normal HTT. The EPR-based
discovery that the polyQ region carries this residual rigidity further supports the “rusty
hinge” hypothesis (2,3). The “hinge” that becomes “rusty” in cases of polyQ expansion
and thereby blocks intramolecular interactions and, conceivably, intermolecular
interactions between HTT and some its various binding partners is contained in the
polyQ region and formed by the additional glutamine residues that make this protein
an unhealthy version of itself.
Further EPR studies, via spin labeling of more sites of polyQ-expanded HDx1
including double labeling to measure intramolecular distances, will be needed to map
the full three-dimensional fold of this unwanted, disease-associated structure. Post-
translational modifications that may modulate this expanded polyQ-associated rigidity
should also be studied by EPR—for instance, by spin labeling and measuring the EPR
spectra of constructs containing the S16D mutation, which has appeared to ameliorate
the effect of polyQ expansion on temperature-dependent conformational rigidity
(Chapter 4)—in order to lend insight into how this rigidity could be altered.

102

5.5 References
1. Hubbell, W. L., Cafiso, D. S., and Altenbach, C. (2000) Identifying conformational
changes with site-directed spin labeling. Nat. Struct. Biol. 7, 735-739
2. Caron, N. S., Desmond, C. R., Xia, J., and Truant, R. (2013) Polyglutamine
domain flexibility mediates the proximity between flanking sequences in
huntingtin. Proc. Natl. Acad. Sci. U. S. A. 110, 14610-14615
3. Truant, R., Atwal, R. S., Desmond, C., Munsie, L., and Tran, T. (2008)
Huntington's disease: revisiting the aggregation hypothesis in polyglutamine
neurodegenerative diseases. FEBS J 275, 4252-4262

103

Chapter 6

Conclusions: On the Non-Amyloid Structures and Behaviors of Amyloid-Forming
Proteins

104

Equally important to studying amyloid fibril formation by proteins is delving into their
conformations, functions, and toxicity occurring separately from or prior to their
amyloidogenesis. α-Synuclein (1-4), islet amyloid polypeptide (IAPP) (5,6), and
huntingtin (HTT) (7) all interact with physiological membranes independently (though
not exclusively independently) of their propensity to form fibrils. In addition, HTT exon
1 (HDx1) in solution (not aggregated) apparently adopts levels of α-helical
conformation that vary according to the number of glutamine residues (Q-length) in its
polyglutamine (polyQ) region, which is the variable that determines whether or not the
form of HTT in question is toxic (8). Therefore, the structural enhancement observed in
HDx1 of expanded polyQ (explicated in Chapters 4 and 5) could be at the root of its
toxicity, or at least its loss of function, in Huntington’s disease. The cellular damage
conferred by α-synuclein and IAPP in Parkinson’s disease (9) and type II diabetes (10),
respectively, could also stem from their structures and behaviors, most notably their
membrane interactions, as monomers/oligomers rather than as aggregated β-sheets. The
mechanisms informing non-amyloid or preamyloid toxicity of these proteins could also
apply to the preamyloid forms of other amyloid-forming proteins.
6.1 Membrane curvature induction: a possible common link to amyloid toxicity (and
beyond)
My IAPP research (as detailed in Chapter 3) couples membrane damage to a novel
membrane curvature induction mechanism. IAPP in amphipathic α-helical form causes
a large part of the disruption of membrane integrity observed upon adding IAPP to
synthetic liposomes in vitro or viewed in in vivo samples of affected cellular membranes
(11,12). This new model for the mechanism of membrane disruption by IAPP and
possibly other similar proteins does not preclude the validity of previously proposed
models, such as the stable, oligomeric pore (13) or the carpet model (14). For one thing,
different studies, including studies on a single membrane-active peptide, have utilized
different membrane compositions; a protein could preferentially employ one membrane
105

interaction mechanism on membranes of a certain composition and another on
membranes of another composition. For another, a number of proposed models could
all stand on a continuum of protein-membrane interactions. This continuum could be
multidimensional, encompassing membrane energetics, protein-to-lipid ratio, and
insertion size. In their review about membrane-active peptides (13), Drs. Last,
Schlamadinger, and Miranker point out that protein-membrane interactions involve a
measure of give-and-take between the protein and the membrane, rather than, as
discussions within the literature at large may often imply, exertions by proteins on
completely passive membranes. They explain that membranes can thin and distort
themselves spontaneously and bound peptides—curvature-sensitive ones such as IAPP
(Chapter 3) and others (15-18), i.e., proteins that have the tendency to bind more highly
curved membranous structures than those of low curvature—could then bind to and
stabilize these distortions (13). Therefore, it is important to consider that membrane
curvature induction may not simply be the creation of new structures from an
otherwise unperturbed structure, but an energy shift toward more distorted states in

Fig. 6-1. Wedging of an amphipathic helix (red dot) into a phospholipid bilayer could lead
to (A) membrane curvature induction or (B) toroidal pore formation. Building blocks
extracted from Last et al., 2013.
A B
106

the membrane that is mutually initiated by membrane and protein. Membrane
curvature induction that at some phospholipid compositions could result in the
formation of highly curved structures due to a helical insertion pushing phospholipid
headgroups apart and pinching part of the membrane outward (Fig. 6-1A) could
instead, in cases of a membrane phospholipid composition with a higher energetic
propensity for adopting transient toroidal distortions (Fig. 6-1B), result in the energetic
stabilization of the resulting toroidal pore by the insertion. Thus, the toroidal pore
mechanism (14) and curvature induction by α-helical insertion could be related.
Furthermore, toroidal pores are linked to consequent activation of sinking-raft or
carpet/detergent mechanisms that ultimately result in dissolution of the membrane (14);
this strand of the protein-membrane interaction continuum could therefore encompass
several of these mechanisms.
Protein-to-phospholipid ratio could inform another dimension of the continuum
of protein-membrane interactions. The disruption of membrane integrity by IAPP has
been found to rely on a cooperative mechanism (19) (also Fig. 1-7B). Therefore, the
mechanism depends on, or is at least substantially accelerated by, different IAPP

Fig. 6-2. Vesicle budding induced by amphipathic helix insertion could lead to (top) tubule
or (bottom) channel-like pore formation. Adapted from Varkey et al., 2010.
107

monomers coming together on the membrane in some way rather than interacting with
the membrane in isolation. This coming together would predictably occur increasingly
at higher protein concentrations and could include oligomer formation. It is not difficult
to envision that several helices bound to a membrane would more efficiently, say, cause
formation of a tubule and stabilize it than would one. Thus, it would not be surprising
to find membrane curvature induction to be a cooperative process (Fig. 6-2), as is
membrane damage, which is coupled to this process. At sufficiently high protein
concentrations relative to available membrane area, the membrane curvature induction
process could cross over into stable pore formation (Fig. 6-2), as the localization of
several monomers could shift their association equilibrium in favor of forming an
oligomer that in this case would act as a membrane pore.
Finally, the size, or more specifically the length, of the helix could determine the
type of membrane curvature induction that occurs. The examples of α-synuclein and
IAPP have served to illustrate that tubulation and vesiculation occur at varying degrees
that depend upon helix length. Since a single helix can only bend a membrane in one
direction, a longer helix induces more anisotropic curvature strain on a membrane,
leading to a greater degree of tubulation, than several shorter helices with the same
total number of residues, leading to increased vesiculation (1). This could be a general
rule for helical wedges in membranes, such that it would take an additional factor
supporting tubule stabilization, such as a protein scaffold working in conjunction with
helical insertions, a non-membrane-binding domain adjacent to the membrane-bound
helical domain keeping helical wedges sufficiently apart to compensate for their
isotropicity, or a membrane composition that prohibits a great extent of binding by the
peptide in question, to make a short helix form stable tubules to the same extent as a
long helix. Because the α-synuclein α-helix is much longer than the IAPP helix and
possibly also because the former is adjacent to a longer non-membrane-binding C-
terminus, it generates tubules to a greater extent and more stably. Furthermore, lipid-
108

bound α-synuclein has been found to include populations of different helix lengths (20)
(also suggested in Chapter 2), including populations of broken helices (2,21), which are
correlated with formation of lipoprotein nanoparticles (2), which appear to be stabilized
by the α-synuclein broken helix conformation while the elongated helix conformation
stabilizes lipid tubules. Membrane interactions by amyloid proteins apart from α-
synuclein and IAPP should be analyzed for similar membrane architecture modulation
patterns, as should antimicrobial peptides, since melittin, for example, has been found
to induce both poration and budding of vesicles, leading to their shrinkage (22),
possibly pointing to the likely generality of membrane disruption by curvature
induction.
Apart from the continuums between membrane curvature induction and pore
formation and between tubulation and vesiculation, membrane-bending proteins also
have a tendency to sense membrane curvature and therefore may lie on an inducing-
sensing continuum, as illustrated by Fig. 3-7. Like IAPP (Chapter 3), helical α-synuclein
and amyloid β-peptide sense membrane curvature (18,23), as do similarly amphipathic
α-helical motifs in Bin/Amphiphysin/Rvs (BAR) domain proteins (15). This has broad
implications for where these proteins will bind cellular membranes in vivo and could
also be part of a mechanism that could be generalizable to other amyloid proteins and
possibly other membrane-active peptides such as antimicrobial peptides. Furthermore,
membrane curvature sensing plays an essential role in the energetic give-and-take
between proteins and membranes (see above), since it enables proteins to bind
selectively to membrane distortions, stabilizing curved structures formed by the
membrane that would be transient and energetically unfavorable otherwise. It is likely
that membrane curvature sensing most likely leads to proteins remaining bound to the
highly curved membrane structures generated by curvature induction exercised by
those same proteins; therefore, the equilibrium between curvature sensing and
109

Fig. 6-3. Proposed model for HTT/HDx1 conformational equilibrium.
Increasing the temperature or decreasing Q-length promotes disorder and
flexibility in HTT/HDx1 conformation, while decreasing the temperature or
increasing the number of Q repeats promotes a collapsed, more highly
helical state.
curvature induction could be shifted toward the former over time, with the highly
curved structures acting as a protein-trapping sink.
6.2 Jekyll-and-Hyde–like conformational switching in HDx1
My CD and
EPR data
regarding
HDx1
(Chapters 4
and 5) show
enhanced
structuring in
this protein in the presence of an expanded polyQ region. The fact that the enhanced
structure proved temperature-labile signifies a conformational difference across Q-
lengths, as opposed to an irreversible reaction (e.g., forming and/or breaking covalent
bonds). The additional Q residues appear to shift the protein to a more ordered energy
state, in much the same way as does lowering the temperature, as dramatically
demonstrated by superposition of EPR spectra from HDx1 of Q-lengths 9–18 residues
apart collected 4–12°C apart (Fig. 5-8). There appear to exist two conformational states
of HDx1—an ordered one, whose additional structure consists of α-helical residues
according to CD data (Chapter 4), and a disordered one—and the equilibrium between
these two states can be modulated by varying temperature or varying Q-length (Fig. 6-
3). The interchangeability between varying temperature and varying Q-length as the
means to produce a conformational state suggests that pharmaceutical means could be
devised to promote adoption of the disordered state, or at least a less ordered state, in
HTT of expanded polyQ.
EPR data (Chapter 5) have enabled the start to visualization of what the two
conformational states of HDx1 might look like. We hypothesize that the normal
110

structure of HDx1 allows for a conformation in which it is loosely folded in on itself
(Fig. 6-4A). PolyQ expansion stiffens the polyQ region, as per the “rusty hinge”
hypothesis (24,25), reducing the protein’s ability to form this fold fully and replacing it
with a more rigid, collapsed fold (Fig. 6-4B). This rigidifying extraneous structure in
HDx1 of expanded polyQ is likely to alter intramolecular interactions between this HTT
segment and regions on its C-terminal side (24) and to enhance some intramolecular
interactions within HDx1 and abrogate others (Fig. 6-4), all of which could structurally
account for observed polyQ expansion-associated changes in interactions with binding
partners. For instance, interactions with protein kinase C and casein kinase 2 substrate
in neurons (PACSIN1), which binds directly to the proline-rich C-terminus of HDx1
(26), are enhanced in HTT with expanded polyQ (27). This could be due to increased
exposure of the C-terminus because it interacts less with the N17 region, as Förster
resonance energy transfer (FRET) data show (25) and our hypothesis would imply (Fig.
6-4B).

Fig. 6-4. Proposed models for folding of HDx1 of normal and expanded Q-length. (A) HDx1
of normal Q-length has a low level of α-helix formation in the polyQ tract and will fold
loosely, bringing the N17 and C-terminal regions in proximity. (B) HDx1 of expanded Q-
length has more α-helicity in the polyQ region and will fold into a rigid, collapsed structure
that instead brings the N17 and polyQ regions in proximity.
111

Conversely, interactions between HTT fragments and the nuclear export protein
known as translocated promoter region (Tpr) are significantly reduced in the presence
of a polyQ expansion, resulting in the nuclear accumulation of polyQ-expanded HTT
fragments (28). Unlike PACSIN1, Tpr binds to the N17 region of HDx1 (28). The
decreased flexibility that we found in the “early” (N-terminal) polyQ region could be
generating the enhanced interaction between this region and the N17 domain seen by
FRET (29) and decreasing the exposure of the N17 region to binding partners (Fig. 6-4B).
The effect of expanded polyQ-associated rigidity on the N17 region, which has
membrane-associating properties (7), most probably affects its membrane interactions
as well. HDx1 membrane interaction studies have been embarked upon in our
laboratory and are important for gauging the effect of the structural enhancement and
for assessing possible mechanisms in HDx1 toxicity to membranes, which could be
related to the above-discussed mechanisms uncovered for α-synuclein and IAPP.

112

6.3 References
1. Varkey, J., Isas, J. M., Mizuno, N., Jensen, M. B., Bhatia, V. K., Jao, C. C., Petrlova,
J., Voss, J. C., Stamou, D. G., Steven, A. C., and Langen, R. (2010) Membrane
curvature induction and tubulation are common features of synucleins and
apolipoproteins. J. Biol. Chem. 285, 32486-32493
2. Varkey, J., Mizuno, N., Hegde, B. G., Cheng, N., Steven, A. C., and Langen, R.
(2013) alpha-Synuclein oligomers with broken helical conformation form
lipoprotein nanoparticles. J. Biol. Chem. 288, 17620-17630
3. Mizuno, N., Varkey, J., Kegulian, N. C., Hegde, B. G., Cheng, N., Langen, R., and
Steven, A. C. (2012) Remodeling of lipid vesicles into cylindrical micelles by
alpha-synuclein in an extended alpha-helical conformation. J. Biol. Chem. 287,
29301-29311
4. Pandey, A. P., Haque, F., Rochet, J. C., and Hovis, J. S. (2011) alpha-Synuclein-
induced tubule formation in lipid bilayers. J. Phys. Chem. B 115, 5886-5893
5. Jayasinghe, S. A., and Langen, R. (2007) Membrane interaction of islet amyloid
polypeptide. Biochim. Biophys. Acta 1768, 2002-2009
6. Hebda, J. A., and Miranker, A. D. (2009) The interplay of catalysis and toxicity by
amyloid intermediates on lipid bilayers: insights from type II diabetes. Annual
review of biophysics 38, 125-152
7. Atwal, R. S., Xia, J., Pinchev, D., Taylor, J., Epand, R. M., and Truant, R. (2007)
Huntingtin has a membrane association signal that can modulate huntingtin
aggregation, nuclear entry and toxicity. Hum. Mol. Genet. 16, 2600-2615
8. Gusella, J. F., and MacDonald, M. E. (1995) Huntington's disease: CAG genetics
expands neurobiology. Curr. Opin. Neurobiol. 5, 656-662
9. Volles, M. J., and Lansbury, P. T., Jr. (2003) Zeroing in on the pathogenic form of
alpha-synuclein and its mechanism of neurotoxicity in Parkinson's disease.
Biochemistry 42, 7871-7878
10. Gurlo, T., Ryazantsev, S., Huang, C. J., Yeh, M. W., Reber, H. A., Hines, O. J.,
O'Brien, T. D., Glabe, C. G., and Butler, P. C. (2010) Evidence for proteotoxicity in
beta cells in type 2 diabetes: toxic islet amyloid polypeptide oligomers form
intracellularly in the secretory pathway. Am. J. Pathol. 176, 861-869
11. Last, N. B., Rhoades, E., and Miranker, A. D. (2011) Islet amyloid polypeptide
demonstrates a persistent capacity to disrupt membrane integrity. Proc. Natl.
Acad. Sci. U. S. A. 108, 9460-9465
12. Magzoub, M., and Miranker, A. D. (2012) Concentration-dependent transitions
govern the subcellular localization of islet amyloid polypeptide. FASEB J. 26,
1228-1238
13. Last, N. B., Schlamadinger, D. E., and Miranker, A. D. (2013) A common
landscape for membrane-active peptides. Protein Sci. 22, 870-882
113

14. Butterfield, S. M., and Lashuel, H. A. (2010) Amyloidogenic protein-membrane
interactions: mechanistic insight from model systems. Angew. Chem. Int. Ed. Engl.
49, 5628-5654
15. Bhatia, V. K., Madsen, K. L., Bolinger, P. Y., Kunding, A., Hedegard, P., Gether,
U., and Stamou, D. (2009) Amphipathic motifs in BAR domains are essential for
membrane curvature sensing. EMBO J. 28, 3303-3314
16. Cui, H., Lyman, E., and Voth, G. A. (2011) Mechanism of membrane curvature
sensing by amphipathic helix containing proteins. Biophys. J. 100, 1271-1279
17. Ouberai, M. M., Wang, J., Swann, M. J., Galvagnion, C., Guilliams, T., Dobson, C.
M., and Welland, M. E. (2013) alpha-Synuclein senses lipid packing defects and
induces lateral expansion of lipids leading to membrane remodeling. J. Biol.
Chem. 288, 20883-20895
18. Pranke, I. M., Morello, V., Bigay, J., Gibson, K., Verbavatz, J. M., Antonny, B., and
Jackson, C. L. (2011) alpha-Synuclein and ALPS motifs are membrane curvature
sensors whose contrasting chemistry mediates selective vesicle binding. J. Cell
Biol. 194, 89-103
19. Knight, J. D., Hebda, J. A., and Miranker, A. D. (2006) Conserved and cooperative
assembly of membrane-bound alpha-helical states of islet amyloid polypeptide.
Biochemistry 45, 9496-9508
20. Bodner, C. R., Dobson, C. M., and Bax, A. (2009) Multiple tight phospholipid-
binding modes of alpha-synuclein revealed by solution NMR spectroscopy. J.
Mol. Biol. 390, 775-790
21. Lokappa, S. B., and Ulmer, T. S. (2011) Alpha-synuclein populates both elongated
and broken helix states on small unilamellar vesicles. J. Biol. Chem. 286, 21450-
21457
22. Yu, Y., Vroman, J. A., Bae, S. C., and Granick, S. (2010) Vesicle budding induced
by a pore-forming peptide. J. Am. Chem. Soc. 132, 195-201
23. Matsuzaki, K., and Horikiri, C. (1999) Interactions of amyloid beta-peptide (1-40)
with ganglioside-containing membranes. Biochemistry 38, 4137-4142
24. Truant, R., Atwal, R. S., Desmond, C., Munsie, L., and Tran, T. (2008)
Huntington's disease: revisiting the aggregation hypothesis in polyglutamine
neurodegenerative diseases. FEBS J 275, 4252-4262
25. Caron, N. S., Desmond, C. R., Xia, J., and Truant, R. (2013) Polyglutamine
domain flexibility mediates the proximity between flanking sequences in
huntingtin. Proc. Natl. Acad. Sci. U. S. A. 110, 14610-14615
26. Plomann, M., Lange, R., Vopper, G., Cremer, H., Heinlein, U. A., Scheff, S.,
Baldwin, S. A., Leitges, M., Cramer, M., Paulsson, M., and Barthels, D. (1998)
PACSIN, a brain protein that is upregulated upon differentiation into neuronal
cells. Eur. J. Biochem. 256, 201-211
114

27. Modregger, J., DiProspero, N. A., Charles, V., Tagle, D. A., and Plomann, M.
(2002) PACSIN 1 interacts with huntingtin and is absent from synaptic
varicosities in presymptomatic Huntington's disease brains. Hum. Mol. Genet. 11,
2547-2558
28. Cornett, J., Cao, F., Wang, C. E., Ross, C. A., Bates, G. P., Li, S. H., and Li, X. J.
(2005) Polyglutamine expansion of huntingtin impairs its nuclear export. Nat.
Genet. 37, 198-204
29. Fodale, V., Kegulian, N. C., Verani, M., Cariulo, C., Azzollini, L., Petricca, L.,
Daldin, M., Boggio, R., Padova, A., Kuhn, R., Pacifici, R., Macdonald, D.,
Schoenfeld, R. C., Park, H., Isas, J. M., Langen, R., Weiss, A., and Caricasole, A.
(2014) Polyglutamine- and temperature-dependent conformational rigidity in
mutant huntingtin revealed by immunoassays and circular dichroism
spectroscopy. PLoS One 9, e112262

Abstract (if available)

Abstract Many degenerative diseases are associated with proteins that misfold and aggregate into amyloid fibrils that are deposited intra‐ or extracellularly at or near the site of tissue degeneration. It has long been established that the protein species conferring toxicity in each case is most likely an intermediate on the pathway to fibrillization or an off‐pathway oligomer rather than the fibrils themselves, which are inert and could even exist as a protective mechanism. However, while the current understanding of the toxicity of amyloid intermediates remains rudimentary, research upon the conformational behavior and possible toxicity of amyloidogenic proteins in monomeric or oligomeric (but not aggregated) form is still more severely lacking. Such research should prove crucial to the development of protein-targeting pharmaceuticals for treatment of these diseases

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Asset Metadata

Creator Kegulian, Natalie C. (author)

Core Title Before they were amyloid: understanding the toxicity of disease-associated monomers and oligomers prior to their aggregation

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 07/10/2015

Defense Date 06/15/2015

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag alpha-synuclein,Amyloid,biochemistry,circular dichroism,Diabetes,electron microscopy,electron paramagnetic resonance,fluorescence microscopy,fluorescence spectroscopy,huntingtin,Huntington's disease,islet amyloid polypeptide,membrane,OAI-PMH Harvest,Parkinson's disease,protein,protein structure,protein-lipid interaction

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Ulmer, Tobias S. (committee chair), Chen, Jeannie (committee member), Haworth, Ian S. (committee member), Langen, Ralf (committee member)

Creator Email kegulian@usc.edu,NatalieCKegulian@aol.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-590175

Unique identifier UC11301253

Identifier etd-KegulianNa-3581.pdf (filename),usctheses-c3-590175 (legacy record id)

Legacy Identifier etd-KegulianNa-3581.pdf

Dmrecord 590175

Document Type Dissertation

Format application/pdf (imt)

Rights Kegulian, Natalie C.

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA