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 A40 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 4 aggregation. The ratio of labelled A40 to unlabelled A40 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 A40 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 A40 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 5 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 A40 peptide. As the A40-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 A40 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 A40 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 A40 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 A40 (Fig.S1b) is well separated from the Tyr’s fluorescence spectrum, thus the Tyr→HLF FRET is unlikely. Nevertheless, the A40-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 A40-DEAC and A40-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 A40-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/A40-HLF 8 system ceased to increase and remained constant at a very low level, as compared to the A40 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 A40-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 A40 peptides leading to the formation of different oligomeric species. We have demonstrated that the small, aminoacid-sized, DEAC molecule covalently bound to the A40 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 Aoligomeric 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 Aautofluorescence 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. 9 1 M. Gralle; and S.T. Ferreira, Progress in neurobiology 82,11 (2007). 2 D.M. Walsh; and D.J. Selkoe, Journal of neurochemistry 101,1172 (2007). 3 D.J. Selkoe; Nature Cell Biology 6, 1054 (2004). 4 S. Kumar, and J.B. Udgaonkar, Current Science 98, 639 (2010). 5 T.R. Serio, Science 289, 1317 (2000). 6 J.C. Rochet, and P.T. Lansbury, Current opinion in structural biology 10, 60 (2000). 7 K.N. Dahlgren, A.M. Manelli, W.B. Stine, et al., J.Biol.Chem. 277, 32046 (2002). 8 Lindgren, M. Sörgjerd, K.; Hammarström, P. Biophys.J. 88, 4200 (2005). 9 Thirunavukkuarasu, S.; Jares-Erijman, E. A.; Jovin, T.M. J.Mol.Biol. 378,1064 (2008). 10 Edwin, N.J.; Hammer, R.P.; McCarley, R.L.; Russo, P.S. Biomacromolecules 11, 341 (2010). 11 Nath, S.; Meuvis, J.; Hendrix, J.; Carl, S.A.; Engelborghs, Y.; Biophys.J. 98, 1302 (2010). 12 Wang, Y.; Clark, T.B.; Goodson, T. J.Phys.Chem. B. 114, 7112 (2010). 13 Schauerte, J.; Wong, P.T.; Wisser, K.C., et al. Biochemistry 49, 3031 (2010). 14 Kim. S.; Schwille, P. Current Opinion in Neurobiology 13, 583 (2003). 15 Xiao, L.; Zhao, D.; Chan, W.-H.; Choi, M.M.F.; Li, H.-W. Biomaterials 31, 91 (2010). 16 Soto, C.; Sigurdsson, E.M.; Modelli, I.; Kumar, R.A.; Castano, E.M.; Frangione, B. Nat.Med. 4, 822 (1998). 10 17 Cabaleiro-Lago, C.; Quinlan-Pluck, F.; Lynch, I.; Lindman, S.; Minogue, A.M.; Thulin, E., et al. J.Am.Chem.Soc. 130, 15437 (2008). 18 Rolinski, O.J.; Amaro, M.; Birch, D.J.S. Biosens&Bioelectr. 25, 2249 (2010). 19 Amaro, M.; Birch, D.J.S.; Rolinski, O.J. PCCP 13, 6434 (2011). 20 McGuinness, C.D.; Sagoo, K.; McLoskey, D.; Birch, D.J.S. Meas.Sci.Technol. 15, L19 (2004). 21 Chan, F.T.S; Kaminski Schierle, G.S.; Kumita, J.R.; Bertoncini, C.W.; Dobson, C.M.; Kaminski, C.F. Analyst 138, 2156 (2013). 22 Shukla, A.; Mukherjee, S.; Sharma, S.; Agrawal, V.; Radha Kishan, K.V.; Guptasarma, P. Arch.Biochem.Biophys. 428, 144 (2004). 23 Pinotsi, D.; Buell, A.K.; Dobson, C.M.; Schierle, G.S.K.; Kaminski, C.F. ChemBioChem 14, 846 (2013). 24 Amaro, M., Kubiak-Ossowska. K.; Birch, D.J.S.; Rolinski, O.J. Methods Appl. Fluoresc. 1, 015006 (2013). 11 Figure captions 1. Parameters obtained from fitting Tyr decay, in a 50μM A40 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 A40-DEAC present at 1: 26 ratio to A40. 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 A40-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