Applied Physics Letters (In press 2014)
Inhibition of beta-amyloid aggregation by fluorescent dye labels
Mariana Amaro1, Thorben Wellbrock, David J.S. Birch, Olaf J. Rolinski2
Photophysics group, Centre for Molecular Nanometrology, Department of Physics, Scottish Universities Physics
Alliance, University of Strathclyde, 107 Rottenrow, Glasgow G4 0NG, UK.
Abstract
The fluorescence decay of beta-amyloid’s (A) intrinsic fluorophore tyrosine has been used for
sensing the oligomer formation of dye-labelled A monomers and the results compared with previously
studied oligomerization of the non-labelled A peptides. It has been demonstrated that two different
sized, covalently bound probes 7-diethylaminocoumarin-3-carbonyl (DEAC) and Hilyte Fluor 488
(HLF), alter the rate and character of oligomerization to different extents. The ability of HLF to inhibit
formation of highly ordered structures containing beta-sheets, was also shown. The implications of our
findings for using fluorescence methods in amyloidosis research are discussed and the advantages of this
auto-fluorescence approach highlighted.
1
Current address: Department of Biophysical Chemistry, J. Heyrovsky Institute of Physical Chemistry, Academy of Sciences of the Czech Republic,
Dolejškova 3, 182 23, Prague 8, Czech Republic.
2
Corresponding author: o.j.rolinski@strath.ac.uk, Fax: +44(0)1415522891 (Olaf J. Rolinski)
1
Misfolded proteins have a tendency to aggregate, leading to the diseases described as amyloidoses. This
group includes some of the most debilitating conditions in modern society such as Alzheimer's (AD),
type II diabetes, Creutzfeldt-Jakob disease and Parkinson's disease. AD is the most common of all
neurodegenerative disorders, making up 50 - 60% of all diagnosed dementias, according to Alzheimer's
Disease International. Until now there is still an open discussion about what is the actual cause of the
disease and the mechanisms that lead to neuron death.
Amyloidoses are characterised by the aggregation of particular proteins, specific to each disease,
that form stable and insoluble amyloid fibrils, which are not cleared from the body and deposit in the
tissue. In the case of AD, the amyloid fibrils are formed by aggregation of the beta-amyloid (Aβ)
protein.
Aβ is a cleavage product of the beta-amyloid precursor protein (APP). APP is a type I
transmembrane protein of up to 770 amino acids in its longer form with a large extracellular domain, a
single transmembrane region and a small intracellular part1. Aβ is a 38 to 42 amino acid long residue of
APP. The 40 and 42 amino acid species - Aβ40 and Aβ42 respectively – are the most common, with the
Aβ40 more abundant.2,3
Amyloid fibril formation appears to be a multi-step process during which a series of intermediate
aggregates are populated. The oligomeric prefibrillar aggregates are difficult to characterize because
they are short-lived, heterogeneous and may be present at low populations. Several different
mechanisms have been proposed to explain fibril formation yet all of them agree on the presence of
different intermediate structures: oligomers and protofibrils. The models of fibril formation, or
fibrillization, also seem to agree with the fact the fibrillization is a nucleation dependent
polymerization4-6.
The Aβ fibrils deposit in the brain as amyloid plaques, but these are not considered to be the
main pathogenic factor and, instead, are viewed as a reservoir of inactive species. It is believed that the
entities that spark neuronal dysfunction and cell death are the smaller soluble Aβ aggregates, termed
oligomers, which are formed in the initial stages of the aggregation process2,7. Therefore sensing of A
2
aggregation in its early stages is highly important for a full understanding of Alzheimer’s disease, and
finding the compounds that can prevent early aggregation may lead to the development of appropriate
therapeutics.
Spectroscopic techniques are widely used to study aggregation in real time, due to their high
sensitivity and the ability to detect low protein concentration. Both covalently bound and associating
probes have been used to study amyloid formation. Dyes such as 1-anilinonaphthalene-8-sulfonate
(ANS), 4-4-bis-1-phenylamino-8- naphthalene sulfonate (Bis-ANS), 4-(dicyanovinyl)-julolidine
(DCVJ)8, and the most commonly used Thioflavin T (ThT) are examples of associating probes.
However, the above dyes may not be particularly suitable to study the early stages of oligomers
formation. For example, the commonly used ThT assay seems to be fibril specific and therefore is not
sensitive to small oligomeric species formed.
On the other hand, covalently bound fluorescence labels are often preferred because their exact
location is known. A few recent examples of such labels used in the amyloid field are: pyrene 9, 5carboxyfluorescein10, Alexa-fluor 488, tetra-methyl rhodamine11, 5-carboxytetramethyl- rhodamine12
and Hilyte Fluor 48813.
Extrinsic fluorescence probe techniques currently used to study oligomerization, like anisotropy,
Förster resonance energy transfer and cross-correlation spectroscopy9,11,14 require labelling the
aggregating species with fluorescent tags.
It is commonly accepted, though rarely quantified, that the addition of an extrinsic fluorophore
tag can disturb to some extent the structure of the protein under study, and also the actual process of
oligomerization, which may lead to two kinds of consequences. Firstly, changes induced by any added
molecule can inhibit peptide aggregation, with clear implications for drug development. In fact, several
compounds have been attempted so far to inhibit beta-amyloid aggregation, including N-acetyl-Lcysteine capped quantum dots15, a 5-residue peptide “beta-sheet breaker”16, or co-polymeric
NiPAM:BAM nanoparticles17, but so far no extrinsic fluorophores were tested in this role.
3
On the other hand, an extrinsic fluorophore may produce results that are not an accurate
representation of the native behaviour of the protein, reducing the usefulness of the approach.
Using the intrinsic fluorescence of protein eliminates the disturbance due to the label completely
and gives a unique opportunity to study the unmodified protein and its natural aggregation process.
We have previously shown18,19 how the approach based on Aβ's intrinsic fluorescence of
tyrosine (Tyr) can be used for sensing oligomer formation noninvasively starting from the very early
onset of single peptide-to-peptide interactions. This approach takes advantage of A having only one
Tyr and no tryptophan in its structure, which allows its selective excitation and fluorescence
measurement. An AlGaN version of a pulsed light emitting diode is used in this work at 279 nm in order
to excite Tyr directly20. This highly specific, and environmentally sensitive response on the Angström
scale of the single Tyr fluorescence decay, has proved to be an efficient sensor of early peptide
aggregation, before the ThT (lag time) or other approaches detect measurable changes.
The other technique using intrinsic fluorescence is based on recently reported amyloid-specific
autofluorescence, which does not originate from fluorescent aminoacids, but appears during formation of
amyloid -sheets21,22 as a result of new electronic levels that become available during this process.
An assay exploring this effect23 has been used to quantify protein aggregation.
In this paper, we have tested the influence of dye-labelling on the oligomerization kinetics. For
this purpose, we have used again the approach based on Tyr fluorescence, and applied it to A40 peptides
labelled at the N-terminus with two dyes, which differ substantially in their structure and size. They are
(Fig.S1 in Supporting Information (SI) file): a small coumarin derivative 7-diethylaminocoumarin-3carbonyl (DEAC), and a larger dye Hilyte Fluor 488 (HLF). Both dyes are covalently bound through
their carbonyl group to the end amino acid residue of 40 N-terminal. The Tyr fluorescence decays
measured during aggregation in the samples containing labelled peptides were used as non-invasive
sensing observables and compared with those obtained in our previous studies for the unlabelled
peptides, to determine whether or not, and to what extent, the labelled dyes disturb the process of
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aggregation. The ratio of labelled A40 to unlabelled A40 was 1:26 for all samples. Further details of
sample preparation and fluorescence lifetime instrumentation are presented in SI.
The DAS6 data analysis package (Horiba Jobin Yvon IBH Ltd.) was used for multi-exponential
deconvolution analyses using a non-linear least squares method. All detected decays of Tyr exhibited a
three-exponential character
3
I  t     i exp  t /  i 
1
i 1
confirmed by means of the χ2 goodness of fit and the distribution of residuals criteria. This is consistent
with 3-rotamer model of Tyr kinetics, which, as we have shown before18,19, can be effectively used to
interpret main fluorescence properties of Tyr during the process of amyloid oligomerization. In other
words, the fractional contributions of the rotamers fi
fi 
 i i
3
 
k 1
 2
k k
and their lifetimes τi, indirectly report the stage of the A40 aggregation process.
Fig.1 presents the evolution of Tyr decay parameters with aggregation time reported in our
previous work19. These results will be used here for comparison with the new data obtained for dyelabelled A40 peptides. The parameters were retrieved from fitting Tyr decay to a three-exponential
model for an unlabelled sample of 50μM 40 in HEPES buffer at 37ºC. Our findings can be
summarised as follows: Three rotamers lifetimes, ~3.7 ns, ~1.1 ns and ~0.4 ns remain constant during
the whole process of oligomerization (Fig.1a), indicating, that there are no changes in the Tyr immediate
neighbourhood, which are significant enough to change the rates of the non-radiative transitions, like
quenching or charge transfer. At the same time, there is a decrease in fractional contribution f2 (Fig. 1b)
accompanied by increases in f1 and f3. Changes in the fi values suggest gradual evolution of the potential
wells the Tyr moieties remain in, and changes in the occupancy of the three minima corresponding to
three rotamers. Specifically, the ratio f1/f2 (Fig. 1c) increases quickly at the beginning, changing by
~100% during the first 15 hours and gradually slows down in the later stages. A unique sensitivity of
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this factor at the beginning of the process reflects alterations in the immediate surroundings of the Tyr10
in each individual peptide. The rate of the process can be arbitrarily characterized by fitting the f1/f2
versus time dependence to the exponential function
 f1 / f 2   t    A0  A  exp  t / taggr .   A
 3
with A0 and A∞ being technical parameters and taggr. a characteristic "aggregation time", which in this
experiment achieved a value of 33.3±3.1 hours.
The same lifetime measurements and the relevant analysis were performed for the DEAClabelled sample. DEAC is a relatively small dye and its size is comparable with the other amino acids
forming the A40 peptide. As the A40-DEAC absorption spectrum is well separated from the Tyr
emission (Fig.S1a), the Tyr→DEAC resonance energy transfer (FRET) is negligible. Direct emission
from DEAC, due to the possibility of exciting its S2 state, is discriminated against by the use of an
emission monochromator. Nevertheless, in order to minimize FRET or any other unforeseen influence of
the label on Tyr fluorescence, the samples were prepared so that only ~ 4% of the peptides were
labelled, meaning that Tyr fluorescence arises mainly from unlabelled peptides. This allows assumption
that the presence of DEAC affects Tyr decay not directly, but through its influence on the aggregation
performance.
The retrieved lifetimes τi and fractional contributions fi as functions of experiment time (Fig.2a
and b), were markedly different from those observed for the unlabelled A40 sample, and have shown a
more complex aggregation. Indeed, we have identified three stages of the evolution of the decay
parameters, thus apparently three stages of amyloid aggregation.
In the first stage, since the beginning until ~75 hours, the lifetimes are comparable to those in
the unlabelled A40 sample and remain constant, while the rotamer contributions fi evolve in a way
similar to the unlabelled sample. Some delay in the growth of the f1/f2 ratio (Fig.2c) is probably a result
of a slower initial coalescence of monomers into dimers, trimers, etc., caused by the DEAC moiety
reducing the probability of successful aggregation events. Nevertheless, the aggregation at this initial
stage occurs probably in a similar way as for the unlabelled peptides.
6
The essential difference appears in the second stage, between about 75 hours and 175 hours of
aggregation. During this period, contrary to the unlabelled sample, the lifetimes change: τ1 increases
gradually from ~4.0 ns to ~4.8 ns, and τ3 decreases from ~0.4 ns to ~0.2 ns. At the same time, the f1/f2
ratio remains constant. This change in the behaviour of Tyr parameters may indicate that the
oligomerization has changed its character. Indeed, our molecular dynamics studies24 suggest that the
initial oligomerization consists on formation of linear chains of peptides, bound end-to-end. As the
DEAC particles are bound to the N-terminal of the A40 peptide, its presence in an aggregating peptide
could terminate the formation of the peptide chain and different structures are formed. In the third stage,
after about 175 hours of aggregation, the lifetimes do not change any more and the f1/f2 ratio increases to
a level similar to the value typical for the unlabelled sample. This is probably caused by the oligomers
formed in the second stage joining together and possibly reorganizing into the familiar higher-order,
beta-sheet containing, aggregates. The spatial scale of this process is too big to be sensed by rotamers
lifetimes but still affects the parameters of the Tyr-peptide potential wells and thus the rotamer
preferences.
The absorption spectrum of HLF bound to A40 (Fig.S1b) is well separated from the Tyr’s
fluorescence spectrum, thus the Tyr→HLF FRET is unlikely. Nevertheless, the A40-HLF samples were
also prepared to yield only ~ 4% of the labelled peptides.
The lifetime τi and fractional contribution fi (Fig.3a and b) are this time substantially different
from the results obtained for the unlabelled and DEAC-labelled samples. The lifetimes display an initial
decrease followed by the recovery of their values, starting after approximately 50 hours. The changes in
the fractional contributions fi are also not monotonic and their initial values are different from those
observed in the previous samples. The initial decrease of the f1/f2 ratio (Fig.3c) is followed by an
increase after 50 hours of the process, supporting the existence of two stages in fluorescence responses.
In the first stage, the lifetimes decrease from the values typical for the monomeric solution to
lower values, which indicates increasing rates of the decay in all three rotamers. This effect may be due
to the presence of the additional non-radiative channels of depopulation of Tyr excited states through
7
quenching and/or charge transfer to other peptides and HLF molecules. The recovery of the lifetime
values in the second stage of the aggregation process could be caused by the consequent growth in size
and complexity of the oligomeric structures and relocation of the HLF moiety, which is connected to the
peptide by a relatively long linker. As the labelled peptides remain a part of the oligomeric structures,
they hinder the growth and slow the aggregation process. Such behaviour could explain the slower
characteristic aggregation time and also the lower asymptotic value recovered from fitting the f1/f2 ratio
to the eqn (3). The fact that the fi parameters do not reach their typical values could be an indication that
the final equilibrium is different from the one reached for unlabelled peptides, and that the aggregation
might be halted before the traditional higher-order oligomers/protofibrils and final fibrillar structures are
formed.
To further investigate the observed differences in the A40-DEAC and A40-HLF samples, we
have measured the DEAC and HLF fluorescence anisotropy decays at different stages of aggregation
(see SI). The rotational time r of the DEAC-containing sample has increased from 1.59 ns for a nonaggregated sample, to about 22.6 ns after 220 hours of aggregation. According to the Stokes-Einstein
dependence, the hydrodynamic diameter of the rotating structures, assuming their spherical shape, has
grown from 2.72 nm to 6.56 nm. Although the aggregating oligomers are not expected to be spherical,
the observed increase can be attributed to their oligomerisation. At the same time, the r factor has
increased from 0.11 to 0.20, indicating gradual loss of rotational freedom. The anisotropy responses of
the HLF sample were significantly different: the short rotational time of ~0.6 ns remained unchanged
during the aggregation, while the initial r was close to zero and has increased to 0.26 after 220 hours.
The stable r does not provide evidence of oligomerisation, and the increase in r suggests again limited
rotational freedom. As no growth of oligomers is observed in HLF-containing sample, its final structure
is likely to be different than that in the one containing DEAC.
To test the last hypothesis, we further investigated the A40-HLF sample aggregation by
measuring the fluorescence intensity of ThT molecules added to the sample in order to track the
formation of beta-sheets18 (see SI). We have found, that the ThT intensity growth in the ThT/A40-HLF
8
system ceased to increase and remained constant at a very low level, as compared to the A40 sample.
Approximately 100 hours after sample preparation, no further increase in ThT intensity was observed,
showing that the formation of -sheet structures is hindered by the presence of the labelled A40-HLF
peptides in the sample. This result confirms our hypothesis that HLF interferes significantly with the
process of oligomerisation from the very beginning, and its embedding hampers the peptide’s
aggregation to a significant extent leading to formation of a suspension of fragmented oligomers, rather
than the usual high-symmetry beta-sheet rich structures.
Clearly the size of the attached fluorescent molecule is a key factor disturbing the interactions
between A40 peptides leading to the formation of different oligomeric species. We have demonstrated
that the small, aminoacid-sized, DEAC molecule covalently bound to the A40 peptide modifies the
kinetics of oligomerization, but the final oligomers seem to be similar to those formed by the unlabelled
peptides. However, the substantially bigger HLF molecule disturbs and slows down the oligomerization
process to that extent, that it prevents formation of beta-sheet structures. In general, the use of labels
modifies the oligomerization process of A peptides even at low label/A ratios.
Our findings impact on current research in finding ways of sensing oligomerization and
preventing the formation of small Aoligomeric species. Awareness of the effect of dye-labelling and
its presence from the very beginning of the aggregation process is crucial to correctly interpreting such
data. We have demonstrated that the Aautofluorescence can be an effective alternative to the labellingbased fluorescence sensing approaches, carrying with it the significant advantage of non-invasiveness.
Discovering how to halt the formation of simple oligomers could be of fundamental importance
in the research on the mechanisms of neurodegenerative disease and even lead to more effective
therapeutics not only for Alzheimer’s but all amyloidoses.
The authors thank EPSRC and SFC for financial support including a Science and Innovation Award.
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Figure captions
1. Parameters obtained from fitting Tyr decay, in a 50μM A40 sample, to a 3 exponential
decay model21. a) Tyr fluorescence decay time; b) fluorescence intensity fractional
contributions. c) ratio between decay times fractional contributions f1/f2; data fitted
according to equation (3); All experimental errors are 3 standard deviations. (© IOP
Publishing. Reproduced by permission of IOP Publishing. All rights reserved).
2. Parameters obtained from fitting Tyr decay with A40-DEAC present at 1: 26 ratio to
A40. A 3 exponential decay model is used. a) Tyr fluorescence decay time; b)
fluorescence intensity fractional contributions. c) ratio between decay times fractional
contributions f1/f2; data fitted according to equation (3); All experimental errors are 3
standard deviations.
3. Parameters obtained from fitting Tyr decay in A40-HLF to a 3 exponential decay model.
a) Tyr fluorescence decay time; b) fluorescence intensity fractional contributions. c) ratio
between decay times fractional contributions f1/f2; data fitted according to equation (3);
All experimental errors are 3 standard deviations.
12