14587
2009, 113, 14587–14590 Published on Web 07/09/2009
Noninvasive Real-time Monitoring of Amyloid-β Fibrillization via Simultaneous Label-free Dielectric Relaxation Spectroscopy and Dark-Field Imaging Yeonho Choi,† Soongweon Hong,† Taewook Kang,‡ and Luke P. Lee*,† Biomolecular Nanotechnology Center, Berkeley Sensor and Actuator Center, Department of Bioengineering, UniVersity of California at Berkeley, Berkeley, California 94720, and Department of Chemical and Biomolecular Engineering, Sogang UniVersity, Seoul, 121-742, Korea ReceiVed: May 4, 2009; ReVised Manuscript ReceiVed: May 26, 2009
Accumulated evidence suggests that amyloid-β has a critical role in chronic neurodegenerative disorders such as Alzheimer’s disease. Therefore, understanding of amyloid-β fibrillization is a focus of interest for the development of innovative therapeutic and diagnostic applications. The fibrillization of amyloid-β has similar growth characteristics of polymeric nanoparticles and current monitoring methods show only qualitative or static information. Here we describe a noninvasive real-time monitoring of nanoscale amyloid-β fibrillization by simultaneous Dielectric Relaxation Spectroscopy (DRS) and label-free dark-field imaging. First, the hydrodynamic radius is characterized by DRS, which can reflect the averaged radius of fibrilized amyloid-β, and we observe an increase from 19 to 21 nm during 48 h. Scattering intensity from dark-field imaging allowed us to visualize and quantify the fibrillization with respect to the incubation time of amyloid-β. Consequently, real-time observation and quantification of changes in both hydrodynamic radii and optical properties (i.e., scattering intensity) were performed simultaneously. Such a dual-mode technique may prove valuable for elucidating the mechanism of amyloid fibrillization and ultimately for designing possible diagnostic methods. Since the precise role of Amyloid-β (Aβ) aggregates in causing neurodegenerative diseases such as Alzheimer’s Disease (AD) is still unknown, systematic elucidation of oligomerization and aggregation of Aβ peptides could provide a clue to unveiling the conformation and dynamics of amyloid proteins as well as a definitive molecular basis for the clinical laboratory diagnosis of AD.1-11 In this regard, it is critical to develop characterization methods to track the structural evolution of Aβ oligomerization or fibrillation from soluble Aβ peptide. Direct visualization of Aβ fibrillation has been realized via transmission electron microscopy and atomic force microscopy, which have limitations in real-time monitoring and quantification of Aβ aggregation.10-12 Similarly, fluorescence and circular dichroism measurements have also been limited to qualitative analysis.13-16 Quartz crystal microbalance gives only quantitative information since it exploits a mass change of Aβ aggregate during aggregation.11,17 In the present work, we fabricated a transparent submicrometer gap electrode as a dual-response sensor platform exhibiting a dielectric and an optical response for combined optical dark-field imaging and dielectric relaxation spectroscopy (DRS) measurement of Aβ fibrillization noninvasively. In general, complementary measurement modalities provide more information and better cross checks for a precise characterization of biological and chemical events than either method alone can yield.18-20 In addition to this, our dual mode detection technology provides several unique advantages in the characterization of Aβ fibrillization. First, optical * To whom correspondence should be addressed. E-mail: lplee@ berkeley.edu. Phone: (510)642-5855. † University of California at Berkeley. ‡ Sogang University.
10.1021/jp904157j CCC: $40.75
and electrical measurements can be performed in aqueous solution where Aβ can sustain and progress its fibrillization under natural conditions. Second, in situ monitoring to characterize the fibrillization progress can be carried out via simple dual mode measurements. Third, based on the DRS result, the dynamic dipole moment’s change can be characterized during its fibrilization process. The calculated hydrodynamic radius offers quantified information for the fibrilized status of Aβ.21,22 Fourth, the optically measured scattering intensity change can be correlated with the hydrodynamic radius result. The underlying concept of our real-time dual-mode detection for Aβ fibrillization is illustrated schematically in Figure 1. Conformational changes of Aβ can be resolved by optical darkfield imaging and electrical DRS system. In the DRS measurement, an alternating voltage is applied across a sample and the sinusoidal current as a response is obtained as a function of frequency. On the basis of this measurement result, the various electrical properties such as complex impedance, inductance, permittivity, conductance, and phase angle can be calculated. Among these obtained values, the complex permittivity can be related with properties determined by the charge response in a system. Traditionally, a macroscale capacitor and relatively high frequency (in general, >1 MHz) are used in DRS.18 However, the dielectric properties at low frequency (smaller than 1 MHz) are highly related with the overall effective dipole moment change of detected molecules, because the results at low frequencies are primarily determined by the polarization of oscillating counterions over the entire structure of Aβ and can indicate the overall structural change (i.e., the progress of the fibrillization). Moreover, as Aβ peptides are aggregated and form bigger oligomers or plaques, the scattering intensity increases in 2009 American Chemical Society
14588
J. Phys. Chem. C, Vol. 113, No. 33, 2009
Letters
Figure 1. The schematics of the quantification for amyloid-β fibrilization by the optical and dielectric analysis system. (a) Amyloid-β polymerization steps in neurons. (b) Schematic diagram of experimental setup and the quantification of amyloid-β fibrillization by optical dark-field imaging and corresponding dielectric relaxation spectroscopy (DRS).
optical dark-field imaging measurements, and this optically resolved in situ dark-field imaging can generate additional characterization data of Aβ fibrillization in our system. As shown in Supporting Information Figure 2, a transparent submicrometer gap electrode for a dual-mode response is fabricated by using two polished indium-tin oxide (ITO)-coated glass slides (CG-51N-1115, Delta Technologies) as transparent electrodes and 50 nm polystyrene (PS) beads (Polyscience Inc.) as a dielectric spacer between the two glasses slides. In a typical experiment, 50 nm PS bead solutions in methanol (13 µL, 30 nM) is deposited onto two polydimethylsiloxane (PDMS)-confined areas (2.5 mm × 12.5 mm each) on bottom ITO-coated glass. After drying the solution, the PDMS block is removed, followed by placing another ITO-coated glass on top of the bottom ITO glass. In order to transport and confine a solution of interest through the space between the two ITO-coated glasses, 100 µm fluidic tubes are connected at both sides. For the prevention of leakage and the attachment between two glasses, the open structure of the flow channel is blocked by a dielectric material (Muscant, Acproducts Inc.). During this process, external force (∼30 N) is applied in order to prevent an increase in the gap size upon drying of the dielectric material. Finally, electrical wires are connected to both ITO-coated surfaces for DRS measurement. The gap size estimated by a simple capacitance equation (C ) εA/d, where C is capacitance [F], ε is permittivity [F/m], A is area [m2], and d is gap distance [m]) gives ca. 770 ( 63 nm. Aβ (Amyloid-β42 peptide, Sigma-Aldrich) is dissolved in deionized water at a concentration of 10 µM in order to minimize the ionic concentration of the sample solution. After loading the Aβ
solution into our submicrometer gap electrode, the scattering images and DRS signal are collected for 48 at 4 h intervals by using a dark-field microscopy system with a true-color imaging chargecoupled device camera and dielectric analyzer (Novocontrol) (Figure 1b). In DRS measurement, the resonance peak in the imaginary part of the permittivity is shifted from 1900 to 850 Hz and the peak intensity increased 67% (Figure 2a). On the other hand, in the case of the deionized water as a negative control experiment, the peak is not changed and the intensity is increased by only 7% (Supporting Information Figure 1). Assuming that the overall effective dipole moment of the aggregated Aβ is relatively small compared with soluble Aβ peptide due to the size, this change in dipole moment during the fibrilization can induce longer momentary delay with respect to the alternating electrical field, and the resonance peak shift to the lower frequency as well as intensity growth in permittivity are observed. Moreover, the effective hydrodynamic radius is calculated to further characterize the oligomerization and aggregation of Aβ based on the DRS result. First, the Cole-Cole equation as a dielectric relaxation model is applied to obtain the relaxation frequency24,25
ε* ) ε∞ +
where
∆ε 1 + i(ωτ0)1-R
(1)
Letters
J. Phys. Chem. C, Vol. 113, No. 33, 2009 14589
ε* ) ε′ - iε′′,
ω ) 2πf,
τ0 ) 2πf*,
∆ε ) ε0 - ε∞ (2)
ε* is the complex dielectric constant, f is the frequency, f* and τ0 are the relaxation frequency and time, respectively, ∆ε is the increment of the dielectric constant, ε0 and ε∞ are the dielectric constant at low and infinite frequency, respectively, and R is the empirical parameter that can be derived from the best fit to the experimental data. Since the Cole-Cole equation generates an arc of circle in which radius is r and center is (a, b) in the ε* complex plane, the DRS signal can be fitted into a form of
(ε′ - a)2 + (ε′′ - b)2 ) r2
(3)
Then, using eqs 1-3, ε0, ε∞, and R can be derived as
ε0 ) √r2 - b2 + a, b 2 sin-1 r R)π
ε∞ ) √r2 - b2 - a
()
(4)
Finally, based on the calculated result, the relaxation time can be obtained as
(
2 N 1 (ε0′ - ε∞′ ) + ε′′k 1 τ0 ) N k)1 2πfk (ε′ - ε′ )2 + ε′′ k ∞ k
∑
)
1/[2(1-R)]
(5)
where N is the number of the data points. From this relaxation frequency, the effective hydrodynamic radius r which represents the status of the fibrilized Aβ can be calculated using the equation under the assumption that the shape of Aβ is sphere24,25
r)
(
kBT
)
1/3
8π2ηf*
(6)
where η is the viscosity of the solvent. Since the calculated effective hydrodynamic radius is directly related with the motion of gyration, the fibrilized Aβ structure can be quantified with this radius. Figure 2b shows the calculated hydrodynamic radius change as a function of incubation time. It indicates systematic increase from 19.1 to 20.8 nm with increasing time. By stark contrast, the control shows that any change in hydrodynamic radius without Aβ except the increase in a standard deviation from 0.514 to 2.15% was not observed. The regression coefficient in the hydrodynamic radius versus the incubation time is higher than 0.97 for 4 days of measurements. In optical dark-field imaging, the scattered intensity by a single small particle from an unpolarized light input of wavelength λ and intensity I0 is given based on the equation of Rayleigh scattering
IScat ) I0
( )(
1 + cos2 θ 2π λ 2R2
4
n2 - 1 n2 + 2
)( 2
dAmyloid 2
)
6
(7)
where R is the distance to the particle, θ is the scattering angle, n is the refractive index of the particle, and d is the diameter of the amyloid-β particle. Therefore, even small changes in the size of fibrilized Aβ can induce huge different scattering intensity, because scattering intensity is proportional to the sixth order of the diameter of fibrilized Aβ. Upon fibrilization, soluble Aβ peptide gradually transforms into large-size insoluble oligomers or plaque. Therefore, as shown in Supporting Information Figure 3, bright dots after 16 h are clearly resolved in
Figure 3. Quantification of amyloid-β fibrilization by optical darkfield imaging. Total scattering intensity (IScat) changes as a function of incubation time (t).
Figure 2. DRS measurement of amyloid-β fibrillization. (a) The representative in situ complex permittivity (ε′′) and (b) calculated effective hydrodynamic radius (RH) change as a function of incubation time (t). The concentration of amyloid-β42 is 10 µM.
Figure 4. Hydrodynamic radius changes as a function of total Rayleigh scattering intensity.
14590
J. Phys. Chem. C, Vol. 113, No. 33, 2009
the dark-field image, and the number of dots increases with respect to the incubation time. Consistently, total scattering intensities in Figure 3 also correlate well with the dark-field images. These results suggest that the fibrilization process of Aβ can be resolved optically via the dark-field imaging. As shown in Figure 4, DRS results (i.e., the calculated hydrodynamic radii) are in a good agreement with the darkfield images showing a linear increase in the scattering intensities. In conclusion, noninvasive monitoring of nanoscale Aβ fibrilization by simultaneous label-free optical dark-field imaging and DRS is successfully performed with transparent electrodes. Oligomerization and aggregation of Aβ causes changes in hydrodynamic radius and scattering intensity, which are observed by DRS and dark-field imaging systems. Therefore our dual-mode electrode prepared via a simple and cost-effective method can give two independent sets of qualitative and quantitative information at the same time. We further expect our optofludic DRS and dark-field imagingbased dual-mode detection to impact on a wide range of research areas in neurodegenerative disease such as Alzheimer’s disease pharmacology and molecular diagnostics. Acknowledgment. This work was supported by the National Science Foundation (Award Number CBET-0239333) and the Center for Nanostructured Materials and Technology (CNMT) under the 21st Century Frontier Research Programs of the Korea government. Supporting Information Available: Control experiment results, fabrication steps for transparent submicrometer gap device, and representative time-resolved dark-field images. This material is available free of charge via the Internet at http://pubs.acs.org.
Letters References and Notes (1) Hardy, J.; Selkoe, D. J. Science 2002, 297, 353–356. (2) Dobson, C. M. Nature 2003, 426, 884–890. (3) Bossy-Wetzel, E.; Schwarzenbacher, R.; Lipton, S. A. Nat. Med. 2004, 10, S2-S9. (4) Quist, A.; et al. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 10427– 10432. (5) Lazo, N. D.; Downing, D. T. Biochemistry 1998, 37, 1731–1735. (6) Petkova, A. T.; et al. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 16742– 16747. (7) Ma, B.; Nussinov, R. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 14126– 14131. (8) Ma, B.; Nussinov, R. Curr. Opin. Struct. Biol. 2006, 10, 445–452. (9) Paravastu, A. K.; et al. Proc. Natl. Acad. Sci U.S.A. 2008, 105, 18349–18354. (10) Petkova, A. T.; et al. Science 2005, 307, 262–265. (11) Luhrs, T.; et al. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 17342– 17347. (12) Knowles, T. P. J.; et al. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 10016–10021. (13) Kammerer, R. A.; et al. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 4435–4440. (14) Li, Q.; et al. Angew. Chem., Int. Ed. 2004, 43, 6331–6335. (15) Ban, T.; et al. J. Biol. Chem. 2006, 281, 33677–33683. (16) Peter, K.; Nilsson, R.; Hammarstrom, P. AdV. Mater. 2008, 20, 2639–2645. (17) Hovgaard, M. B.; Dong, M.; Otzen, D. E.; Besenbacher, F. Biophys. J. 2007, 93, 2162–2169. (18) Tatsuma, T. Anal. Chem. 1997, 69, 887893. (19) Wang, J.; Pumera, M. Anal. Chem. 2002, 74, 5919–5923. (20) Cheesman, M. R.; et al. J. Am. Chem. Soc. 2004, 126, 4157–4166. (21) Yi, M.; Jeong, K.; Lee, L. P. Biosens. Bioelectron. 2005, 20, 1320– 1326. (22) Kamei, T.; Oobatake, M.; Suzuki, M. Biophys. J. 2002, 82, 418– 425. (23) Kremer, F.; Schonhals, A.; Luck, W. Broadband Dielectric Spectroscopy; Springer-Verlag: New York, 2002. (24) Bishay, S. T. Egypt. J. Sol. 2000, 23, 179–188. (25) Grant, E. H.; Sheppard, R. J.; South, G. P. Dielectric BehaVior of Biological Molecules in Solution, Clarendon, Oxford, 1978.
JP904157J
