Reflective Interferometric Detection of Label-Free Oligonucleotides

Department of Chemical Engineering, Department of Biochemistry and Biophysics, ... Future Health, Department of Chemistry, and Department of Physics, ...
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Anal. Chem. 2004, 76, 4416-4420

Reflective Interferometric Detection of Label-Free Oligonucleotides Jinghui Lu,†,⊥ Christopher M. Strohsahl,‡,⊥ Benjamin L. Miller,‡,§,⊥ and Lewis J. Rothberg*,|,†,⊥

Department of Chemical Engineering, Department of Biochemistry and Biophysics, Department of Dermatology, Center for Future Health, Department of Chemistry, and Department of Physics, University of Rochester, Rochester, New York 14627

New chip-based methods for the detection of unmodified biomolecular targets have significant potential as enabling technology in fundamental biology and biomedical analysis. We report a method based on changes in reflectivity from specially fabricated substrates that is capable of detecting the binding of as little as an average of 0.2 nm (i.e., a fraction of a monolayer) of biomolecules. We demonstrate the method on detection of femtomole quantities of untagged oligonucleotides in an array format, showing that the amount of target bound can be determined quantitatively. The simplicity of the approach promises to make it broadly applicable for any biomolecule for which suitable molecular recognition chemistry is available. Sensitive and selective schemes to detect biomolecules are important enabling tools in medicine, environmental monitoring, and biological research.1 The vast majority of sensing instruments are based on fluorescent tagging of molecules in the sample under investigation,2-4 a time-consuming and expensive process. In addition, most of the fluorescent readout schemes currently available require expensive imaging systems and detectors. Simplification of the chemistry to avoid tagging of analytes and development of portable, inexpensive readout schemes continue to be central challenges. Surface plasmon resonance5,6 and ellipsometric7,8 and interferometric methods 9-12 have all proved to be viable ways to avoid tagging, but they still tend to require * To whom correspondence should be addressed. E-mail: rothberg@ chem.rochester.edu. † Department of Chemical Engineering. ‡ Department of Biochemistry and Biophysics. § Department of Dermatology. ⊥ Center for Future Health. | Department of Chemistry and Department of Physics. (1) Cooper, M. A. Nat. Rev. Drug Discovery 2002, 1, 515-528. (2) Iyer, V. R.; Eisen, M. B.; Ross, D. T.; Schuler, G.; Moore, T.; Lee, J. C. F.; Trent, J. M.; Staudt, L. M.; Hudson Jr., J.; Boguski, M. S.; Lashkari, D.; Shalon, D.; Botstein, D.; Brown, P. O. Science 1999, 283, 83-87. (3) Epstein, J. R.; Biran, I.; Walt, D. R. Anal. Chim. Acta 2002, 469, 3-36. (4) Chee, M.; Yang, R.; Hubbel, E.; Berno, A.; Huang, X. C.; Stern, D.; Winkler, J.; Lockhart, D. J.; Morris, M. S.; Fodor, S. P. A. Science 1996, 274, 610614. (5) Brockman, J. M.; Nelson, B. P.; Corn, R. M. Annu. Rev. Phys. Chem. 2000, 51, 41-63. (6) Knoll, W. Annu. Rev. Phys. Chem. 1998, 49, 569-638. (7) Jin, G.; Tengvall, P.; Lundstrom, I.; Arwin, H. Anal. Biochem. 1995, 232, 69-72. (8) Landry, J. P.; Zhu, X. D. Opt. Lett. 2004, 29, 581-583. (9) Jenison, R.; Yang, S.; Haeberli, A.; Polisky, B. Nat. Biotechnol. 2000, 19, 62-65.

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relatively complex detection systems, accompanying amplification chemistry, or both. In the present paper, we describe a method based on reflection that is sensitive and quantitative and has the potential to be implemented in an extremely simple format. We have demonstrated detection of oligonucleotides hybridizing with their surfacebound complements at the level of ∼0.02 monolayers. The method is suitable for highly parallel detection in microarray formats and can, in principle, be adapted to work under water. EXPERIMENTAL SECTION Surface Preparation. Nominally undoped silicon wafers with thermally grown oxides of ∼150 nm were purchased. Wafers were diced into pieces of 1.5 cm × 1.5 cm to be used as substrates. These were cleaned in boiling piranha etch solution, which was prepared by mixing 30% H2O2 and 98+% H2SO4 in volume ratio of 3:7, for 30 min and were rinsed several times with doubly deionized water (dd water). The cleaned substrates were then boiled in dd water for 30 min to have the oxide surface of the substrates fully hydrated and were stored in dd water at room temperature. Oligonucleotide Synthesis. DNA oligonucleotides (B1, B2, and C1) were synthesized by MWG Biotech Inc. (High Point, NC) and B3 (Variola complement), Variola target, and Monkeypox target by Invitrogen (Carlsbad, CA). The probe sequences were terminated with a biotin modifier at the 5′ end in the sequence. The target DNA sequences were unmodified. The sequences were as follows: Probe 1 (B1): biotin-5′-TTT TTT TTT GTT CTT CTC ATC ATC-3′ Probe 2 (B2): biotin-5′-TTT TTT TTT GAT GAT GAG AAG AAC-3′ Target (C1): 5′-GAT GAT GAG AAG AAC-3′ Probe 3 (B3,Variola): biotin-5′-AAG ATG CAA TAG TAA T-3′ Variola target: 5′-AAT ACT ATT GCA TCT T-3′ Monkeypox target: 5′-AGA CAA GCC TGT AA-3′ Hydrophobic Patterning. Photoresist (Shipley 1805) was micropipetted by hand in a pattern of two rows of four spots each, with spacing of 2 mm at the center of the substrates. Volumes of 0.2 µL were applied to each spot and spread to form 1-mm(10) Lin, V. S. Y.; Motesharei, K.; Dancil, K. P. S.; Sailor, M. J.; Ghadiri, M. R. Science. 1997, 278, 840-843. (11) Chan, S.; Li, Y.; Rothberg, L. J.; Miller, B. L.; Fauchet, P. M. Mater. Sci. Eng., C 2001, 15, 277-282. (12) Pan, S.; Rothberg, L. J. Nano Lett. 2003, 3, 811-814. 10.1021/ac0499165 CCC: $27.50

© 2004 American Chemical Society Published on Web 06/25/2004

Figure 1. Schematic of the substrate composition and measurement geometry (not to scale). The capital letters S, A, P, and D representing the optics are light source, aperture, polarizer, and detector, respectively. We used a 450-W Xe lamp filtered by a 0.3-m monochromator as S and a CCD camera as D. θ1 is the incident angle, and d is the aggregate thickness of the fabricated functional layers and thermally grown SiO2 layer. n1, n2, and n3, are the refractive indices of air, the oxide/biomolecular coating, and Si, respectively.

diameter dots, which were left 30 min to dry. The substrate was put into 50 mL of a solution of 30% carbon tetrachloride (Sigma Aldrich) and 70% hexadecane (Sigma Aldrich) containing 200 µL of OTS (octaldecyltrichlorosilane, Gelest) and ultrasonicated for 8 min at 18 °C. Subsequently, the substrate was washed in 20 mL of acetone three times to remove the photoresist and expose the patterned bare oxide surface. This was followed by 2 min of ultrasonication in dd water. Finally, the substrate was rinsed with dd water three times and dried under flowing high-purity nitrogen gas. Probe Immobilization. The patterned substrates were immersed in 10 mL of 5% (aminopropyl)triethoxysilane (APTES, Sigma Aldrich) solution in acetone for 1.5 h, rinsed with dd water, and baked in a drying oven at 100 °C for 1 h. The inner wells (designated 2, 3, 6, and 7 in Figure 2a) were then modified by applying 10 mM sulfo-NHS-biotin (Pierce EZ-Link) in phosphate buffer solution PBS (100 mM, pH ) 7.4) for 8 h. Excess sulfoNHS-biotin was removed with a dd water rinse, and nitrogen gas was used to dry the surface. Streptavidin (Sigma, 0.2 mg/mL in 10 mM PBS, pH ) 7.4) was applied to the biotinylated wells, and the biotin-modified oligonucleotides were attached to the streptavidin layer. Each of these reactions was allowed to proceed for 1 h in a covered Pyrex dish containing paper towel fragments saturated with dd water and was followed by dd water rinsing and nitrogen gas flow drying. For the probe attachment reaction, the B1, B2, and B3 probe molecules were dissolved in PBS (10

mM, pH ) 7.4) at 50 µM concentration, and 2 µL of the appropriate probe solution was micropipetted into the wells: B3 into 2 and 3, B1 into 6, and B2 into 7. Hybridization. Target solutions of single stranded DNA were made at a 10 µM concentration in PBS (10 mM, pH ) 7.4). In each case, a 2-µL droplet was pipetted into each probe well, left for 30 min in a humid Pyrex dish, and then rinsed briefly with ice-cold (∼4 °C) dd water (to minimize dehybridization) and dried under flowing nitrogen gas. Optical Measurement Apparatus and Data Analysis. The probe light was derived from a 450-W Xe lamp monochromatized to ∼1 nm bandwidth using a spectrometer. A polarizer was used to enforce s-polarization. The light was guided through two apertures ∼5 mm in diameter and separated by 60 cm to enforce collimation to better than 0.5°. The beam was incident on the substrate at 70.6°, and the reflected light was observed directly on a Roper Scientific CCD camera without imaging optics. We captured reflected images at 15 measurement wavelengths spaced by 2 nm in a range around the reflectivity minimum. The data were averaged to obtain manageable file sizes by binning regions of 5 × 5 pixels together. Curves of reflectivity versus wavelength were extracted for each position in the images, and these were fit to inverted parabolas to estimate the wavelength of minimum reflection. The resulting surface maps can be translated directly into topological maps of the surface using eq 2b. For the purposes of sensing, we compare topological maps of the surface before and after exposure to complementary DNA. RESULTS AND DISCUSSION Principle and Theory of the Measurement. A schematic of the substrate and measurement geometry is depicted in Figure 1. A substrate with a coating of thickness d is surface-functionalized with probe molecules for the desired target. Light is incident on the substrate at angle θ1 such that the reflection from the entire assembly vanishes by destructive interference for a particular wavelength λ. Upon binding with the target molecule, the wavelength for which destructive interference occurs shifts to the red or, alternatively, the reflectivity at the original wavelength increases. Although one could envision using several possible substrate-coating pairs, we have chosen to implement this method using silicon substrates coated with a thick thermal oxide. These

Figure 2. Surface topology of substrate with prearrayed bare wells. (a) 2D gray scale map of the patterned substrate surface. The x and y pixel numbers derive from those of the CCD array format (1317 (x) × 1035 (y)) after binning regions of 5 × 5 from raw data to reduce to an array 263 × 207. The actual distance scale is 8.98 mm × 7.04 mm overall, with each original pixel 6.8 × 6.8 µm. (b) Raw reflection versus wavelength derived at two different regions shown as open circles and closed circles corresponding to the rectangular regions outlined by white and black boxes labeled in (a). (c) Cross-section in the x direction of the topology of wells 5-8 in the OTS background for the substrate surface shown in (a) where ∼10 y positions have been averaged. d′ is the surface height in nanometers derived using eq 2b, where d′ ) d - 152 nm so that most of the oxide thickness is subtracted to properly utilize the dynamic range of the gray scale. Values for d′ correspond to the gray scale bar in (a). A height of 2.3 nm, the literature value for the thickness of a dense OTS monolayer, is shown at the right for reference.

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are easily obtained, very flat, and biomolecular attachment to SiO2 surfaces is well developed. In addition, backside reflections from the substrate are eliminated, since the Si is opaque in the visible region. The reflection R from the structure shown in Figure 1 is given by a simple analytic expression,13

R ) |(r12 + r23 exp(-2iβ))/(1 + r12r23 exp(-2iβ))|2

(1)

where rjk ) (nj cos θj - nk cos θk)/(nj cos θj + njk cos θk) are the Fresnel reflection coefficients for TE (s-polarized) light at the interface between layer j and layer k, nj are the complex refractive indices of the various layers, i ) x-1 and β ) (2π/λ)n2 d cos θ. The choice of s-polarized light is essential to the ability to minimize the reflection, an important distinction from other interferometric approaches.9 Using red probe wavelengths far below the direct band gap of silicon, the imaginary part of the reflection coefficient is negligible, and setting R ) 0 gives the conditions for the reflectivity minimum as 2β ≈ π and r12 ≈ r23. Physically, perfect destructive interference can be obtained when the reflections at the air/oxide interface and oxide/silicon interface have equal magnitude and travel distances that differ by a half wavelength. The conditions are easily solved to obtain the criteria for the incidence angle θ1min and oxide coating thickness dmin to make R ≈ 0:

θ1min ) sin-1 ({(n32 - n24/n12)/(n12 + n32 - 2n22)}1/2) (2a) dmin/λ ) 1/(4n2 cos θ2) ) (1/4)(n22 - n12 sin2 θ1)-1/2 (2b) For the air/SiO2/Si system under consideration and λ ) 660 nm, n1 ) 1, n2 ) 1.4563, and n3 ) 3.8251, where we ignore the imaginary part of the Si refractive index as assumed above. The theory predicts minimums for incidence angle θ1 ) 70.6° and dmin/λ ) 0.2253. In other words, for each 0.22 nm change in the coating thickness, there is a 1-nm shift in the reflectivity minimum, easily detectable using spectroscopic methods. Under ideal conditions where the surface is perfectly flat, the illumination is monochromatic and perfectly collimated, the reflectivity change at the minimum wavelength for a 0.22-nm thickness change is greater than a factor of 10. A more sophisticated transfer matrix model including the effects of nonidealities, and the fact that biomolecules have different refractive index than SiO2 has been developed and will be published shortly. Array Preparation and Concept Verification. Crystalline silicon substrates with thermally grown oxides provide a suitable substrate that can be functionalized with attachment chemistry developed for glass. Oxide thicknesses in the neighborhood of 100-150 nm were used to achieve reflection minimums in the visible wavelength range. We prepatterned the surface hydrophilicity to make reaction wells on the surface for attachment of various probe molecules. The areas between wells were separated by a compact and inert hydrophobic monolayer of octadecyl(13) Born, M.; Wolf, E. Principles of optics, 5th ed.; Pergamon Press: Oxford, New York, Toronto, Sydney & Braunschweig, 1975.

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trichlorosilane (OTS). The patterning confines droplets containing probe and target to the wells and also allows for “built-in” control experiments in which various steps of the treatment can be omitted. The topology of this surface was measured using reflective interferometry with incidence angle θ1 near 70.6°. Pictures of the spatial pattern of reflected probe light versus wavelength were used to identify the wavelength of minimum reflection at each location on the substrate. These wavelengths were determined by fitting raw data to a parabolic minimum and converted to height in nanometers using eq 2b. The result after deposition of patterned OTS to create eight hydrophilic wells of approximately millimeter dimension and separation is illustrated in Figure 2a. The rectangular field of view corresponds to the CCD camera dimensions of 8.98 mm × 7.04 mm (1317 × 1035 pixels), and the circular region mirrors the image of the reflected probe light. The oval features in the circular region correspond to circular hydrophilic wells on the substrate; their aspect ratio is distorted by tan(70.6°) due to the oblique incidence of the probe beam on the substrate. Figure 2b depicts the raw reflection versus wavelength data from which the topology is derived at two points on the surface, one inside a well and one between wells. Figure 2c presents a cross-section of the height of the substrate coating illustrating the definition and uniformity of the wells. Their height as derived using the theory is ∼2.3 nm, in excellent agreement with literature values for OTS monolayers14 and with what we measure using spectroscopic ellipsometry. These results confirm the quantitative nature of the reflective interferometric technique. Attachment of Oligonucleotide Probes. In each of the wells, the thermal oxide was silanized with (aminopropyl)triethoxysilane (APTES), leaving terminal amine groups that could be further reacted to attach probes. The four outer wells 1, 4, 5, and 8 were not treated further and were used as controls to verify that removal and replacement of the substrate in the apparatus did not modify the results or disturb the well registration. In the four inner wells, the amine groups were reacted with sulfo-NHS-biotin, followed by streptavidin. The streptavidin attachment site allows us to bind tightly to a wide variety of commercially available biotinylated probes. The large size of the streptavidin molecule also enforces sizable probe spacings that prevent hindrance of target binding by steric crowding. Again, we are able to use reflective interferometry to measure the efficacy of the layer attachments. In each case, the addition of material to the wells is not uniform, but is concentrated toward the center of the well. We assume this phenomenon results from the hydrophobicity of the walls deforming the droplet to bias binding to occur away from the edges. Measured in the center of the resulting bumps in the wells, the approximate thicknesses added by APTES, sulfo-NHS-biotin and streptavidin were 0.9, 1.3, and 1.5 nm, respectively. The first two are in good agreement with what is expected for a dense monolayer. Next, biotinylated DNA probes were attached to the streptavidin. In the top row of Figure 2a, the probe oligonucleotide in wells 2 and 3 is a 16-base fragment complementary to a segment of the crmB gene from the Poxviridae family, genus Orthopoxvirus. The gene encodes for the viral homologue of the tumor necrosis factor receptor (vTNFR). The sequence, originally described by (14) Silberzan, P.; Le´ger, L.; Ausserre´, D.; Benattar, J. J. Langmuir 1991, 7, 1647-1651.

Figure 3. Topology of the substrate surface in oligonucleotide sensing experiments. 2D gray scale maps of the substrate surface height in nanometers with bound probes are illustrated before (a) and after (b) exposure to target analytes. Height values are derived in the same manner as for Figure 2 (d′ ) d - 152 nm). (c) and (d) are cross-sections in x of the difference picture where (a) has been subtracted from (b) to give height changes ∆d following target exposure. Bands of 10 y values near the centers of the wells have been averaged together. For reference, the cross-sections of the wells before treatment (from Figure 1c) are shown to clarify the well positions, since these are no longer obvious in the difference image. x and y scales are identical to those in Figure 2.

Mikhailovich et al.,15 is unique to variola (smallpox) and can be used to specifically identify the smallpox virus. In the bottom row, two different probes with random base sequences of length 15 (B1 and B2) are used in wells 6 and 7, respectively. These two sequences also incorporated a 5′-TTTTTTTTT spacer to make the probe more easily accessible to the target sequence. After probe attachment, the resulting surface topologies are shown in Figure 3a. The concentration of material at the center of the wells, as noted above, is evident in the Figure. In each case, ∼1.5 to 2 nm of oligonucleotide probe is attached to the center of the wells, corresponding to roughly 20% of the thickness of a dense monolayer. Reflective Sensing of Oligonucleotides. In the top set of wells, the probes are the same, and different analytes were applied. Well 2 was exposed to an incorrect target sequence derived from monkeypox, whereas well 3 was exposed to an oligonucleotide containing the complementary sequence. In the bottom set of wells, the probes B1 and B2 are in wells 6 and 7, respectively, and identical analytes were applied containing an oligonucleotide C1 with sequence containing the complement of the probe in well 6. In each case, 2-µL droplets of analyte solution containing 10 µM target DNA (20 pmol) were applied to attempt hybridization. The surface topology after application of the target is illustrated (15) Lapa, S.; Mikheev, M.; Shchelkunov, S.; Mikhailovich, V.; Sobolev, A.; Blinov, V.; Babkin, I.; Guskov, A.; Sokunova, E.; Zasedatelev, A.; Sandakhchiev, L.; Mirzabekov, A. J. Clin. Microbiol. 2002, 40, 753-757.

in Figure 3b. Figure 3c and 3d depict cross-sections (3c for the top row, wells 1-4, and 3d for the bottom row, wells 5-8) through the well centers of a picture that is the difference in height between the pictures in 3b and 3a. For reference, the equivalent cross-sections of the bare wells are included to show where the differences are observed. Target binding in wells 3 and 6 is obvious. There appear to be negligible contributions of nonspecific target binding, as nothing is observed in the wells that are untreated or exposed to noncomplementary target to within our signal-to-noise ratio. Quantitative estimates of the amount of the bound DNA can be obtained from the data of Figure 3c and d. In the bottom set of wells, the height increase at the center of well 6 containing the complementary probe is ∼1.4 ( 0.2 nm, whereas it is 0.1 ( 0.2 nm for well 7, which has the noncomplementary probe, and also is within error of 0 for all of the control wells (1, 4, 5, and 8). From an integration of the topology, we can estimate the total amount of DNA deposition in the well. Averaged over the 1-mm diameter well size, there is ∼0.4 nm average addition of material, or ∼8% of a dense target monolayer. Assuming that a dense monolayer contains 2.5 × 1013/cm2, the amount of attached material corresponds to ∼40 femtomoles of oligonucleotide. An estimate based on a 0.4-nm layer 1 mm in diameter having the density of DNA and the molecular weight of the oligonucleotide yields nearly the same result.

CONCLUSIONS AND FUTURE WORK We have demonstrated a simple reflective interferometric biomolecular sensor capable of detecting submonolayer coverages of untagged DNA oligonucleotides. As are all reflection measurements, our approach is a special case of ellipsometry, but our method has the advantage that the most complex feature of ellipsometry (measurement of the phase of the reflected wave), is not required. The penalty for this is that one must work on specially coated substrates. Our approach also provides a large improvement in sensitivity over previous colorimetric methods, obviating the need for amplification chemistry and a sandwich assay format.9 It also provides quantitative data on the amount of target bound. The penalty for these is the need to use collimated and polarized probe light at a well-defined incidence angle. Reflective interferometry scales favorably to larger targets, since it is sensitive to changes in optical path length, which is a measurement of the total amount of material and not the thickness, per se. It is appropriate to any type of analyte in which moleculespecific recognition chemistry is available. Conversely, the method can be used to screen surface attached libraries to identify probe molecules for a desired target. We have realized a convenient implementation using silicon substrates and silicon oxide attachment chemistry. Although we have been able to detect binding of subpicomolar levels of short oligonucleotides, a number of modifications should increase the sensitivity substantially. These include scaling to smaller spot size, along with improved collimation and monochromaticity of the probe light. The ability to work under aqueous media is implicit in the method, and we are pursuing these applications. Perhaps even more important is the ability to envision inexpensive and portable sensing systems, in which a monochromatic probe such as a laser pointer is used with a split photodiode to detect Analytical Chemistry, Vol. 76, No. 15, August 1, 2004

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reflectivity differences between two spots after they are exposed to an analyte. We consider a variety of implementations of the method using a more rigorous and detailed theory in a separate paper to be published shortly. ACKNOWLEDGMENT We are grateful for support by a grant from Infotonics and funding from the Center for Future Health. We also wish to thank Dr. Al Raisanen from the RIT microelectronics fabrication laboratory for growth of the thermal oxides.

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SUPPORTING INFORMATION AVAILABLE Comparison of SPR and reflective interferometry using a transfer matrix formalism. This material is available free of charge via the Internet at http://pubs.acs.org.

Received for review January 14, 2004. Accepted May 18, 2004. AC0499165