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2007, 111, 327-330 Published on Web 12/22/2006
Early Aggregation in Prion Peptide Nanostructures Investigated by Nonlinear and Ultrafast Time-Resolved Fluorescence Spectroscopy Ying Wang and Theodore Goodson III* Department of Chemistry, UniVersity of Michigan, Ann Arbor, Michigan 48109 ReceiVed: October 29, 2006; In Final Form: December 4, 2006
We report the characterization of early aggregates in the self-assembly of prion peptides using nonlinear and ultrafast time-resolved fluorescence spectroscopy. The dye-labeled peptide and dye/peptide guest-host systems were used to demonstrate the feasibility of the new approach. By measuring the two-photon absorption crosssection, small aggregates of the dye labeled peptide were characterized. Ultrafast time-resolved fluorescence anisotropy spectroscopy reveals the packing state (microenvironment) of the probes to be tightly associated with aggregates and associated with aggregation progression of the peptides. Fluorescence intensity decay shows a correlation with growth of aggregates having a high level of structured β-sheet content. A new binding ligand Cascade Yellow shows promise for β-sheet recognition of prion peptide nanostructures. These findings may have implications for in vivo studies of neurotoxic aggregates targeting with fluorescence markers. Also, these results may provide insight into molecular design of peptide-based nanomaterials.
A broad range of scientific endeavors has established a growing interest in the naturally occurring self-assembly process of nanoscale objects by polypeptides. In this regard, the amino acids that constitute the peptide motifs have been used to fabricate novel supramolecular structures and advanced materials1-2 as drug delivery carriers,3 antibiotic agents,4 and molecular electronic components.5 The organization of these nanostructures via self-assembly starts from nucleation into ordered nanoscale fibrillar structures. Some particular examples are the prion protein and the amyloid peptide Aβ, which undertake a conformational transition from R-helices into β-sheets and further self-assembly of β-sheets into a nanofiber assembly ultimately leading to plaques.6-8 The self-assembly requires the equilibration between aggregated and nonaggregated states usually affected by environment and dominated by interactions of the proteins or peptides. Through this equilibration, various peptide-based nanostructures have been prepared in which their aggregation or conformers are critical for nanotechnological devices.9-11 Therefore, it is very important to characterize the dynamic intermediates prior to the ordered peptide nanostructures as this information may help the design and understanding of the critical features of novel biomimetic materials at the molecular level. The significance of this investigation also lies in substantial evidence on the toxicity of small aggregation intermediates of the proteins or peptides.12 The characterization of the early aggregates (oligomers) may help the finding of toxic components, in preventing its progression and targeting with therapeutic agents. However, it is very difficult to characterize these oligomeric intermediates due to the variable dynamical properties of the proteins/peptides. A number of biophysical tools have been developed to investigate soluble aggregates of the prion * To whom correspondence should be addressed. E-mail: tgoodson@ umich.edu.
10.1021/jp067098+ CCC: $37.00
protein13 and other amyloidogenic proteins/peptides.14 Among these techniques, the high sensitivity of fluorescence spectroscopy is particularly attractive in determining biomarkers of the neurodegenerative disease13 and analysis of residue-level dynamics of amyloid peptides15-17 as well as high throughput assays.18 Nonlinear spectroscopy, particularly two-photon absorption (TPA)19-20 has a number of remarkable features suitable for imaging/detection of amyloid peptide aggregates in highly scattered media.21 The excitation at specific electronic states reflects conformational changes of the peptide and aggregation of the polyelectrolytes.22 In this report, we probed two systems (i.e., a nonlinear chromophore labeled covalently to a prion peptide and the chromophore bound to the prion peptide in a guest-host arrangement) to demonstrate the feasibility of twophoton absorption and ultrafast time-resolved fluorescence measurements as new approaches to characterize early aggregation in prion peptide nanostructures. Specifically, the results of TPA measurements reveal that this technique possesses high sensitivity to small aggregates. Ultrafast fluorescence anisotropy measurements characterize the local motion of the probes, providing information on the packing state of the probes that are tightly associated with aggregates and the aggregation process in general. Fluorescence intensity decay measured in ultrafast time-scale shows a correlation with growth of aggregates having high structured β-sheet content. Moreover, a new ligand, Cascade Yellow, shows promise for recognition of β-structured aggregates in the prion peptide. The structures of nonlinear chromophore Cascade Yellow (pyridinium oxazole salt) ester, hydrolyzed Cascade Yellow (CY), and CY labeled PrP106-126 are shown in the Supporting Information. The use of this particular dye is advantageous as it is environmentally sensitive, hydrophilic,23 nonlinear-active, and chemically stable at neutral pH.24 Table 1 shows two-photon absorption cross-sections (δ, 1 GM ) 10-50 cm4 s/photon© 2007 American Chemical Society
328 J. Phys. Chem. B, Vol. 111, No. 2, 2007
Letters
TABLE 1: Comparison of Photophysical Properties and Two-Photon Absorption Cross-Sections of the CY Contained Systems samples small aggregates 23 µM CY labeled peptidea 2.13 µM CY 16.6 µM CY monomericb 10 µM peptide +2.13 µM CY β-structured aggregatesb 83 µM peptide+16.6 µM CY a
Aabs at 395 nm
quantum yield (η)
δ (GM)
0.70 0.034 0.18
0.104 0.26 0.19
990 ( 202 160 ( 30 195 ( 42
0.022
0.16
150 ( 30
0.28
0.27
129 ( 22
b
Measured after incubation for 72h. Measured after incubation for 41 days.
molecule) of several CY contained systems in PBS buffer (an aggregation inducing solvent). It was found that the CY-labeled peptide gives much larger TPA cross-section (∼1000 GM, approximately 330 GM per CY chromophore) than parent CY and CY bound peptide systems (smaller than 200 GM). The first concern for an interpretation of the amplified TPA effect is whether the labeling significantly contributes to this enhancement. For this concern, we first dissolved the dye labeled peptide in neat trifluoroethanol25 and dried and then dissolved into water without potential interferences from trace organic solvent or small aggregates (seeds) of the peptide. We then immediately measured and compared the UV-vis and fluorescence spectra of CY labeled peptide to that of parent CY in water. The UVvis (visible range) and fluorescence spectra were very similar, suggesting that the labeling does not increase the Stokes shift that is referred to the enhanced TPA effect via the increase in charge-transfer character, conjugation length and donoracceptor strength of the system.26 This result implies that the amplified TPA effect could not be explained by the effect of labeling, but may be explained by interactions of the labeled CY with the rest of amino acids presented in the labeled peptide. For this concern, we compared the UV-vis, fluorescence spectra (in water), and TPA cross-sections (in PBS buffer) of the CY in the guest-host systems to that of parent CY (Table 1). The similarity in TPA cross-sections and spectra of these systems implies that various interactions (electrostatic, hydrophobic, and H-bonding) between the dye and amino acids constituted in the PrP106-126 are too weak to enhance the TPA cross-section. Therefore, the amplified TPA effect is neither from the effects of labeling nor the interactions of the CY with peptide but from the intra and/or inter-chromophoric interactions arising from the peptide-peptide interactions. This interaction may lead to closer packing of the labeled CY to enhance the transition dipolar moment entirely and consequently to enhance the TPA effect.27 Not only can the formation of small aggregates be suggested from TPA measurements but also from ultrafast time-resolved fluorescence anisotropy measurements. Fluorescence anisotropy is an excellent technique to study the rotational motions of fluorophores in the excited states following the excitation using polarized light.28 Time-resolved anisotropy decay reveals multiple rotational motions experienced by the fluorophores in the excited states in terms of their rotational correlation times. Timecorrelated single photon counting (10 ns time window) has been used to measure rotational correlation times of all CY containing peptide systems. Only rotational correlation times within picosecond time scales were observed. The absence of any longer rotational correlation times (ns)16 may be a result of weak coupling or uncoupling of rotational characteristic of the probe with peptide motion.29 To resolve the picosecond dynamics of the probe, the good time-resolution (instrumental response
Figure 1. Ultrafast time-resolved fluorescence anisotropy decays of a 23 µM dye labeled peptide with incubation time up to 72 h.
function: ∼110 fs) of the fluorescence upconversion technique, rather than the time-correlated single photon counting (TCSPC) method, was utilized. Specifically, ultrafast fluorescence anisotropy decay provides useful information in the event that the depolarization of chromophore is faster than rotational diffusion within the aggregated assemblies.30 In our CY-labeled peptide system, global and local motions as well as ultrafast excitedstate energy transfer will affect the polarization of the fluorescence emission. The changes in global motions would give information on the aggregation states of the CY labeled peptide, whereas changes in local motions and appearance in tens of femtosecond anisotropy decay time would provide information on the packing state of the representative CY. Figure 1 shows ultrafast fluorescence anisotropy decay profiles of the 23 µM CY-labeled peptide taken at different incubation times. All anisotropy decays showed a oneexponential decay to the residue values (r∞) of 0.114, 0.139, and 0.165 for the three incubation times. There was also an increase in the rotational correlation time (φ, setting yo ) 0) from 215, 263, and 284 ps with increasing incubation time. These fast rotational correlation times were unlikely due to global or segmental motions (usually in ∼ns) of the peptides,17 and spatial assemblies of peptide aggregates signified by excitedstate energy transfer (usually tens of femtosecond).30 Indeed, we did not observe any ultrafast (fs) anisotropy decay for all of the systems investigated here. Therefore, the local motion of the attached CY chromophore is the main contributor to this decay. As incubation time progresses, the increases in the values (φ, r0, and r∞) suggest a more restricted movement of the attached CY due to the relative closer packing of the chromophores. Such packing is a result of small aggregate formation. These aggregates may be weakly bound or more exposed to solvent than the corresponding PrP106-126 aggregates formed at the same condition. It should be noted that these small aggregates may not be observed using electron microscopy as reported in literature31 or by CD spectra (Supporting Information) but can be observed from our ultrafast spectroscopic measurements. Thus, a combined technique (TPA and ultrafast time-resolved anisotropy) is a sensitive method to characterize small aggregates and follow their aggregating processes of the dye labeled peptide. In order to characterize oligomers of PrP106-126 or PrPSc isolated from biological specimens such as samples from patients, noncovalent recognition using extrinsic dyes is necessary. Several amyloid-staining compounds such as Congo Red, Thioflavin T, Pittsburgh Compound B, Chrysamine G, and
Letters recently developed NIAD-4 have been used for analysis of aggregated amyloid proteins.32 These studies relied on steadystate spectroscopy (UV and fluorescence) changes of the chromophores upon binding to amyloid plaques. Indeed, timeresolved spectroscopy provides more information on fluorescence properties associated with emission species and their correlation with the structures and dynamics of proteins and organic molecules in complex systems.33-34 Yet, there are very few reports16,35 where the binding of fluorescent compounds with peptide aggregates has been studied in detail using timeresolved fluorescence spectroscopy. Cascade Yellow is a new binding agent for recognition of β-structured PrP106-126 aggregates. This chromophore carries the negative charged sulfonate group to interact with the positively charged amino acid residues in β-structured PrP106-126 aggregates. The shape-specific recognition32 may also play an important role in binding of CY with the β-sheet aggregates due to the hydrophobic interactions between β-sheet and aromatic π-system of the CY. Table 1 also shows the comparison of absorbance and quantum yield of the mixed PrP106-126/ CY (molar ratio 5:1) to that of parent CY. As the CY chromophore does not bind to monomeric PrP106-126 (10 µM), the dye is accessible to water through H-bonding so that the absorbance of the CY chromophore decreases, and the emission is quenched. However, in a solution of β-structured PrP106-126 aggregates (83 µM, Supporting Information), an increased absorbance and quantum yield of the CY chromophores have been observed. This result is ascribed to the binding of CY to PrP106-126 aggregates where the conformational freedom of the CY is restricted to decrease radiationless decay rate and increase fluorescence quantum yield. It is noted that the absorbance and fluorescence maxima of the bound and unbound CY do not differ appreciably possibly due to the readily planarity of the hydrophobic core of the CY chromophore. Consequently, the CY chromophore may not undergo a geometry-induced bathochromic shift32 upon binding. The binding of CY to β-structured PrP106-126 aggregates is in the guest-host arrangement and can be illustrated by timeresolved fluorescence spectroscopy. The time-resolved fluorescence spectroscopic method provides more insight into the nature of the probe environments (free, bound to small aggregates, or bound to large aggregates) and thus can be used to follow the formation and growth of aggregates and consequently to address the mechanism of peptide aggregation. Similar to the dye labeled peptide system, the TCSPC measurements of the guest/host arrangement did not show any rotational correlation times within nanosecond time scales. Therefore, ultrafast fluorescence anisotropy decays of unbound and bound CY to PrP106-126 must be measured using fluorescence upconversion technique. Figure 2a shows the experimental results. The anisotropy of unbound CY displayed a one-exponential decrease to 0.130 with time, yielding a rotational correlation time (φ, setting yo ) 0) of 222 ps, whereas the anisotropy of bound CY showed one-exponential decay with rotational correlation time (φ, setting yo ) 0) of 249.4 ps and residue value (r∞) of 0.146. At a peptide concentration of 83 µM, the PrP106-126 formed the β-structured aggregates with approximately 25% β-sheet content. This aggregated PrP106-126 was more tightly packed than the monomeric PrP106-126 formed at 10 µM. From the discussion mentioned above, the binding mechanism of CY to β-structured aggregates is a combination of electrostatic interactions and hydrophobic interactions between β-sheet and the aromatic π system of the CY molecule. Therefore, the CY chromophores may incorporate within the β-sheet to tightly pack and are not
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Figure 2. (a) Ultrafast time-resolved fluorescence anisotropy decays of unbound and bound CY to PrP106-126 and (b) fluorescence intensity decays of the bound CY with the growth of β-structured PrP106-126 aggregates.
free to move, as observed by the increased rotational diffusion time and residual anisotropy value. To understand the detailed “rigidification” observed from ultrafast fluorescence anisotropy measurements, TCSPC was initially used to measure the fluorescence decays of the free CY and the CY bound peptide systems. It was found that the nanosecond decay component is insensitive to any environmental changes, but the picosecond decay component is very sensitive to aggregation process of the prion peptide. Thus, the dynamical processes of the CY chromophore accountable for the initial steps of prion peptide aggregation were resolved by the ultrafast fluorescence upconversion technique. Figure 2b shows the evolution of fluorescence intensity decays of the CY binding to the β-structured PrP106-126 aggregates with time. All fluorescence decays are fitted to a two-exponential function. The shortest decay component (∼2 ps) presented in all cases is assigned to a solvent relaxation process. The second decay component increases during an incubation time. Analysis of the CD spectra recorded at these incubation times and conditions show a clear change of β-sheet content from 14% to 25% (Supporting Information). An increase in fluorescence decay time suggests that the radiationless decays of the excited-state of the CY are less favorable only when the dye is in a hydrophobic environment and interacts with β-sheet aggregates. This result agrees with previous reports in the literature.31,35 It is clear that there is strong correlation between fluorescence decay times with aggregation progression and in turn with
330 J. Phys. Chem. B, Vol. 111, No. 2, 2007 population of β-structured aggregates. Thus, the changes of excited-state dynamics of CY as a function of prion peptide aggregation suggests that ultrafast time-resolved fluorescence spectroscopy is a promising technique to characterize β-structured peptide aggregates and their growth. Indeed, the combination of this technique with TPA properties of the CY probe could be capable of time-resolved multiphoton imaging of these prion peptide nanostructures, and this investigation is currently underway. In summary, we carried out the studies of CY-labeled (covalently) PrP106-126 and CY-bound PrP106-126 systems to demonstrate the sensitivity of nonlinear and ultrafast timeresolved spectroscopic methods to the characterization of oligomeric aggregates in prion peptides. The combined twophoton absorption and ultrafast fluorescence anisotropy technique has high sensitivity to the small aggregates of the dye labeled peptide. Ultrafast fluorescence decays (anisotropy and intensity) show the possibility for utilization of the fluorescence upconversion technique to characterize the β-sheet aggregates and their growth, as well as to understand the molecular mechanisms of the nucleation and growth of the peptide aggregates. The combination of these techniques may offer opportunities for defining protein aggregation pathways and characterizing amyloid aggregates, important for the synthesis of highly efficient peptide-based nanomaterials and for the treatment of neurodegenerative diseases. Acknowledgment. We thank the Army Research Office and the National Science Foundation for support. Supporting Information Available: Experimental details and steady-state spectra. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Carny, O.; Shalev, D. E.; Gazit, E. Nano Lett. 2006, 6, 15941597. (2) MacPhee, C. E.; Woolfson, D. N. Curr. Opin. Solid Sate Mater. Sci. 2004, 8, 141-149. (3) Davis, M. E.; Hsieh, P. C. H.; Takahashi, T.; Song, Q.; Zhang, S. G.; Kamm, R.; Grodzinsky, A. J.; Anversa, P.; Lee, R. T. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 8155-8160. (4) Fernandez-Lopez, S.; Kim, H. S.; Choi, E. C.; Delgado, M.; Granja, J. R.; Khasanov, A.; Kraehenbuehl, K.; Long, G.; Weinberger, D. A.; Wilcoxen, K. M.; Ghadiri, M. R. Nature 2001, 412, 452-455. (5) Scheibel, T.; Parthasarathy, R.; Sawicki, G.; Lin, X.-M.; Jaeger, H.; Lindquist, S. L. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 4527-4532. (6) Dobson, C. M. Nature 2003, 246, 884-890. (7) Prusiner, S. B. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 1336313383.
Letters (8) Hammarstro¨m, P.; Schneider, F.; Kelly, J. W. Science 2001, 293, 2459-2462. (9) Whitesides, G. M.; Grzybowski, B. Science 2002, 295, 2418-2421. (10) Reches, M.; Gazit, E. Curr. Nanosci. 2006, 2, 105-111. (11) Herland, A.; Bjo¨rk, P.; Nilsson, P. R.; Olsson, J. D. M.; Åsberg, P.; Konradsson, P.; Hammarstro¨m, P.; Ingana¨s, O. AdV. Mater. 2005, 17, 1466-1471. (12) Bucciantini, M.; Giannoni, E.; Chiti, F.; Baroni, F.; Formigli, L.; Zurdo, J.; Taddei, N.; Ramponi, G.; Dobson, C. M.; Stefani, M. Nature 2002, 416, 507-511. (13) Bieschke, J.; Giese, A.; Schulz-Schaeffer, W.; Zerr, I.; Poser, S.; Eigen, M.; Kretzschmar, H. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 54685473. (14) Bitan, G.; Fradinger, E. A.; Spring, S. M.; Teplow, D. B. Amyloid 2005, 12, 88-95. (15) Kaylor, J.; Bodner, N.; Edridge, S.; Yamin, G.; Hong, D.-P.; Fink, A. L. J. Mol. Biol. 2005, 353, 357-372. (16) Allsop, D.; Swanson, L.; Moore, S.; Davies, Y.; York, A.; El-Agnaf, O. M. A.; Soutar, I. Biochem. Biophys. Res. Comm. 2001, 285, 58-63. (17) Mukhopadhyay, S.; Nayak, P. K.; Udgaonkar, J. B.; Krishnamoorthy, G. J. Mol. Biol. 2006, 358, 935-942. (18) Liu, M.; Ni, J.; Kosik, K. S.; Yeh, L.-A. ASSAY Drug DeV. Technol. 2004, 2, 609-619. (19) Lakowicz, J., Ed.; Nonlinear and Two-Photon Induced Fluorescence. In Topics in Fluorescence Spectroscopy; Plenum Press: New York, 1997; Vol. 5. (20) Zipfel, W. R.; Williams, R. M.; Webb, W. W. Nat. Biotechnol. 2003, 21, 1368-1376. (21) Bacskai, B. J.; Skoch, J.; Hickey, G. A.; Allen, R.; Hyman, B. T. J. Biomed. Opt. 2003, 8, 368-375. (22) Nilsson, K. P. R.; Olsson, J. D. M.; Stabo-Eeg, F.; Lindgren, M.; Konradsson, P.; Inganas, O. Macromolecules 2005, 38, 6813-6821. (23) Anderson, M. T.; Baumgarth, N.; Haugland, R. P.; Gerstein, R. M.; Tjioe, T.; Herzenberg, L. A. Cytometry 1998, 33, 435-444. (24) Salafsky, J. S. Chem. Phys. Lett. 2003, 381, 705-709. (25) Buck, M. Q. ReV. Biophys. 1998, 31, 297-355. (26) Bhaskar, A.; Ramakrishna, G.; Lu, Z.; Twieg, R.; Hales, J. M.; Hagan, D. J.; Stryland, E. V.; Goodson, T., III J. Am. Chem. Soc. 2006, 128, 11840-11849. (27) Beljonne, D.; Wenseleers, W.; Zojer, E.; Shuai, Z.; Vogel, H.; Pond, S. J. K.; Perry, J. W.; Marder, S. R.; Bre´das, J.-L. AdV. Funct. Mater. 2002, 12, 631-641. (28) Lakowicz, J. R. Principles of Fluorescence Spectroscopy; Kluwer Academic/Plenum Press: New York, 1999. (29) Cohen, B. E.; Pralle, A.; Yao, X.; Swaminath, G.; Gandhi, C. S.; Jan, Y. N.; Kobilka, B. K.; Isacoff, E. Y.; Jan, L. Y. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 965-970. (30) Toele, P.; van Gorp, J. J.; Glasbeek, M. J. Phys. Chem. A 2005, 109, 10479-10487. (31) Jackson Huang, T. H.; Yang, D.-S.; Fraser, P. E.; Chakrabartty, A. J. Biol. Chem. 2000, 275, 36436-36440. (32) Nesterov, E. E.; Skoch, J.; Hyman, B. T.; Elunk, W. E.; Bacskai, B. J.; Swager, T. M. Angew. Chem., Int. Ed. 2005, 44, 5452-5456 and references cited therein. (33) Lindgren, M.; So¨rgjerd, K.; Hammarstro¨m, P. Biophys. J. 2005, 88, 4200-4212. (34) Varnavski, O. P.; Ranasinghe, M.; Yan, X.; Bauer, C. A.; Chung, S.-J.; Perry, J. W.; Marder, S. R.; Goodson, T., III J. Am. Chem. Soc. 2006, 128, 10988-10989. (35) De Rossi, U.; Hermel, H. Appl. Spec. 1999, 53, 505-509.
