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DNA Immobilization on GaP(100) Investigated by Kelvin Probe Force Microscopy David N. Richards,† Dmitry Y. Zemlyanov,‡ Rafay M. Asrar,† Yena Y. Chokshi,† Emily M. Cook,† Thomas J. Hinton,† Xinran Lu,† Viet Q. Nguyen,† Neil K. Patel,† Jonathan R. Usher,† Sriram Vaidyanathan,† David A. Yeung,† and Albena Ivanisevic*,†,§ Weldon School of Biomedical Engineering, Birck Nanotechnology Center, and Department of Chemistry, Purdue UniVersity, West Lafayette, Indiana 47907 ReceiVed: June 27, 2010; ReVised Manuscript ReceiVed: August 4, 2010
Understanding changes in the properties of semiconductor materials after immobilization of biomolecules on the surface is essential for the fabrication of well-tuned and programmable devices. The work examines changes in the properties of gallium phosphide (GaP) after modification with an organic linker, a single stranded DNA, and its complementary strand. We investigated changes in surface potential with Kelvin probe force microscopy (KPFM). Analysis revealed that a more ordered adlayer of ssDNA was present when a lower concentration of linker molecule was used. KPFM data combined with coverage data obtained from X-ray photoelectron spectroscopy (XPS) further confirmed this result. Successful hybridization with the complementary strand was confirmed by both KPFM and Raman spectroscopy. The results indicate that one can control the amount of DNA on the surface by changing the initial concentration of the organic linker, and thus modulate the surface potential of the semiconductor material. Introduction The ability to interface organic and inorganic material is a topic that has garnered much research interest. Forming a stable organic monolayer on a surface has become attractive due to its potential applications in biosensors, proteomics, and photovoltaics. Even more important in this field is the potential to use these organic monolayers as an anchor for biological molecules such as DNA, proteins, and peptides.1 The choice of substrate material is an important parameter. Materials such as silicon nitride,2 gold,3,4 indium-tin-oxide,5 and carbon nanotubes6 have been used for biomolecule immobilization for a variety of purposes. Semiconductors are also highly researched substrates because of their technological maturity. Silicon is more studied than others due to its domination of the semiconductor industry. Indeed, there have been many examples of biomolecule immobilization on silicon, especially for the generation of biosensor devices.7-10 However, due to its relatively small bandgap energy, it is not an ideal material for future devices.1 III-V semiconductors are promising materials due to their high electron mobility and larger bandgap energies. We have chosen to use gallium phosphide (GaP), a III-V semiconductor that has demonstrated favorable biocompatibility in the past.11 In a previous study, we demonstrated the ability of an alkene linker to functionalize a surface of GaP more efficiently compared to an alkanethiol.12 In this study, we use the alkene, undecenoic acid (UDA), to pattern a GaP surface. Using this alkene’s carboxylic acid moiety, we immobilize single stranded DNA (ssDNA) onto the GaP surface and use it to hybridize a cDNA sequence. A number of biosensor applications require an understanding of changes in the surface properties of material after biomolecule immobilization. Therefore, we chose to use Kelvin probe * To whom correspondence should be addressed. E-mail: albena@ purdue.edu. Phone: 765-496-3676. Fax: 765-496-1459. † Weldon School of Biomedical Engineering. ‡ Birck Nanotechnology Center. § Department of Chemistry.
force microscopy (KPFM) to map the surface potential before and after DNA immobilization. Only a few reports analyze the organic monolayer on III-V semiconductors with KPFM.13,14 Most KPFM is performed on III-V semiconductors to investigate the quality of the semiconductor film.15-17 We utilized KPFM to investigate the progressive functionalization of GaP with UDA, ssDNA, and its cDNA strand. In order to contrast the difference between a nonfunctionalized surface and a functionalized surface, we used PDMS stamping techniques to generate a pattern of organic linker molecules on the GaP surface with which to bind singlestranded DNA. Additional surface characterization and spectroscopic techniques are used to confirm and further understand the data obtained from KPFM. Experimental Section Sample Preparation. Wafers of S-doped GaP(100) were purchased from University Wafer (South Boston, MA). Undeceonic acid (UDA, 98%) was purchased from Sigma-Aldrich. Amine-modified DNA with a six-carbon spacer (5′-amine-C6TTAAGGTCTGGACTGGCCTG-3′) and its Cy3-labeled complement (5′-Cy3-CAGGCCAGTCCAGACCTTAA-3′) were purchased from Integrated DNA Technologies (Coralville, IA). Ultrapure water was added to the lyophilized DNA and stored at -20 °C when not in use. The GaP wafers were cut into 1 × 1 cm2 pieces and ultrasonically degreased with water and ethanol for 10 min. The samples were subsequently dried with N2 and immediately exposed to NH4OH for 15 s. The samples were rinsed with water and dried with N2 before being exposed to a 40% solution of NH4F for 15 min. 10, 20, and 30% solutions of UDA in toluene were used for microcontact printing. A drop of the respective solution was placed on a PDMS stamp, fully covering the pattern, and left there for 1 min. The stamp was blown dry with N2 and left alone for 5 min. The cleaned GaP samples were patterned by stamping for 1 min. The stamped samples were immediately
10.1021/jp105927t 2010 American Chemical Society Published on Web 08/20/2010
Kelvin Probe Microscopy on GaP exposed to UV light (302 nm) for 30 min followed by a toluene rinse and drying with N2. Stamped GaP surfaces were exposed to a 10 µM solution of amine-modified DNA for 2 h and subsequently rinsed with water and dried with N2. The DNA-immobilized surfaces were then exposed to a 2 nM complementary strand solution of 1 M phosphate-buffered saline (PBS), 0.35 M sodium chloride (NaCl), and 0.025% sodium dodecyl sulfate (SDS) for 2 h. The samples were then immediately washed in Arrayit (Sunnyvale, CA) wash buffers 1, 2, and 3 for 5 min, 5 min, and 1 s, respectively, per the provided instructions. Samples were finally blown dry with N2. Surface Characterization. Kelvin probe force microscopy was accomplished with a Veeco Multi-Mode Nanoscope IIIa atomic force microscope using the surface potential mapping mode. Conducting tips (model AC240TM Electrilevers, resonant frequency 45-95 kHz, Al coated) were purchased from Asylum Research (Santa Barbara, CA). All KPFM experiments were performed in ambient conditions using a drive frequency of 10 V and a lift height ranging from 2 to 10 nm. XPS data were obtained using a Kratos Ultra DLD spectrometer using monochromatic Al KR radiation (hν ) 1486.58 eV). Survey and high resolution spectra were collected at a fixed analyzer pass energy of 160 and 20 eV, respectively, and acquisition was performed at photoemission angles of 0 and 60° measured with respect to the surface normal. Binding energy values were referenced to the Fermi edge, and charge corrections were done using the C 1s peak set at 284.80 eV. Curve fitting was performed after linear or Shirley type background subtraction assuming a Gaussian/ Lorentzian peak shape. Raman spectra were obtained using a LabRam HR800 Raman microscope (Horiba Jobin Yvon, Edison, NJ) with laser excitation of 661 nm. The slit width and confocal aperture were set to 200 and 1000 µm, respectively. Results and Discussion Atomic force microscopy (AFM) is a very powerful tool for investigating the functionalization of surfaces. In this report, we use a derivative of AFM known as Kelvin probe force microscopy (KPFM), which is capable of mapping the surface potential. The surface potential map is generated by recording the magnitude of the voltage applied to a conducting tip in order to nullify the electrostatic force between the sample and tip.18 This technique allows for the visualization of charged molecules on a surface.19 KPFM has been used to evaluate the change in electronic properties of a surface after modification with alkyl monolayers20 as well as to detect the binding events of various biomolecules.5,21 After functionalization, we hybridize the ssDNA present on the GaP surface with its complementary strand. Figure 1 shows a schematic representation of two possible ways the DNA can be arranged on the surface. It has been demonstrated that the most efficient hybridization occurs when the ssDNA is in an upright conformation as opposed to mostly lying parallel to the surface.22,23 Since DNA is a charged molecule, an investigation of the electrical properties of the functionalized surface is deemed very important for characterization and optimization for future inorganic/biomolecular interfaces. In order to confirm the presence of the Cy3-labeled cDNA on the surface, Raman spectroscopy was performed after amine-DNA and Cy3-labeled cDNA incubation (Figure 2). The bottom spectrum corresponds to a physisorbed Cy3-labeled DNA on GaP, while the three spectra above it were taken at
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Figure 1. Two possible molecular arrangements of the DNA molecules on the GaP surface: (A) an ordered layer of DNA; (B) a more disordered layer of DNA.
Figure 2. Raman spectra of physisorbed Cy3-labeled DNA on GaP and three random spots on a patterned GaP sample after amine-DNA and Cy3-labeled cDNA incubation. The dashed boxes are used to guide the eye to the most intense peaks.
three random spots on the patterned sample where the molecules are expected to be. The Raman data indicates that labeled cDNA is on the surface based on characteristic peaks in the regions indicated by the dotted lines. Topographical Characterization. Due to its nature, KPFM also records a topographical image with every surface potential image. Topographical characterization revealed successful patterning and an average step-height between the bare GaP surface and patterned surface of 0.7 nm. This value correlates well with our previous study in which we calculated an adlayer thickness of about 0.8 nm.12 After incubation in the amine-modified DNA solution, the step height increased by roughly 1 nm when the 10% UDA solution was used compared to about 0.7 nm after the 20 and 30% UDA solutions were used (Figure 3). These results indicate a more ordered adlayer for which the DNA can bind. Consequentially, the DNA ordered itself more perpendicular to the GaP surface. Figure 3 also depicts a representative topographical image before and after DNA immobilization including a crosssectional profile. KPFM Pre- and Post-DNA Immobilization. Using KPFM, various patterned samples of 10, 20, and 30% UDA were analyzed to determine the average surface potential difference between the bare and patterned areas, before and after DNA immobilization. These findings are presented in Figure 4. Before DNA immobilization, the potential difference for the 10% samples is about 4 mV. An increase in UDA concentration to 20 and 30% increases the potential difference to about 9 mV for each, within error. Again, this can be attributed to the exposure of the more positive carbon backbone relative to the GaP surface. Indeed, previous studies have demonstrated that one -CH2- is capable of altering the surface potential by roughly 924 to 14 mV.25 After DNA immobilization, the 10% UDA patterns show a large increase in the average contact potential difference. This phenomenon can be attributed to the large dipole moment that is associated with single-stranded DNA.26 As previously mentioned, the DNA is arranged in a more perpendicular fashion relative to
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Figure 3. (left) The average step height increase after DNA immobilization using various UDA concentrations. (right) Representative topographical images before DNA immobilization (A) and after DNA immobilization (B) using a 30% UDA solution. (bottom) The crosssectional profiles corresponding to the dashed lines (scale bar corresponds to 1 µm).
Richards et al.
Figure 5. Contact potential difference as a function of UDA linker coverage before DNA immobilization (A) and after DNA immobilization (B).
Coverage was calculated using the following equation:
dσk GaP sadlayer Nadlayer(θ) dΩ × Λe (Es) cos θ coverage ≡ ) ssubstrate NGaP(θ) dσl ×d dΩ
Figure 4. Average contact potential difference (mV) for various samples using 10, 20, and 30% UDA concentration before and after DNA immobilization.
the surface when the 10% UDA solution is used, owing to the increase in surface potential after immobilization. However, for the 20 and 30% UDA patterns, we see a large decrease in surface potential after DNA immobilization which can be explained by the exposure of the negatively charged DNA backbone. In other words, the orientation of the UDA molecules is most likely parallel to the surface which forces the DNA to arrange in a conformation that is parallel to the surface. KPFM vs XPS. The nonpatterned (flat) portion of a PDMS stamp was used to create an interface in which one side was functionalized and one side was clean GaP. The interface was analyzed by KPFM and then sent for XPS analysis to evaluate coverage. The samples were then incubated in amine-modified DNA, and the interface was again analyzed by KPFM. A summary of the results can be found in Figure 5.
where sadlayer and ssubstrate are the mean surface densities of atoms in the adlayer and substrate, respectively. Nadlayer(θ) and NGaP(θ) are the peak intensities of the adlayer and Ga 3d or P 2p, respectively; dσk/dΩ and dσl/dΩ are differential cross sections; ΛGaP e (Es) is the Ga 3d or P 2p electron attenuation length; d is the distance between adjacent gallium (phosphorus) planes for GaP(100); and θ is the electron emission angle. Atmospheric contaminants were accounted for by extracting values for Nadlayer(θ) corresponding to the carboxyl carbon atom (HOs CdO).12 Figure 5A indicates that, as the coverage increases, the average surface potential difference between the bare surface and the functionalized surface also increases. The bare GaP surface is at a more negative potential than the UDA linker molecule’s carbon backbone; therefore, this finding suggests that the UDA layer is becoming more disordered and not exposing its negative carboxylic acid moiety at higher coverage. At lower coverage values, a few data points suggest that the functionalized surface is at a lower potential than the bare GaP, indicated by the negative value of the contact potential difference. This most likely indicates a more ordered UDA layer that is exposing its carboxylic acid. However, other points in this same coverage region indicate a positive potential difference suggesting a disordered UDA layer. This unusual spread in values is most likely a result of the error associated with the coverage value
Kelvin Probe Microscopy on GaP
J. Phys. Chem. C, Vol. 114, No. 36, 2010 15489 cross-sectional profile of the 10% UDA progression from parts G-I corresponding to the dashed lines. The increase in surface potential after DNA immobilization and the subsequent decrease after hybridization is clearly seen. Conclusions
Figure 6. KPFM progression of UDA pattern, amine-DNA immobilization, and Cy3-labeled cDNA hybridization for 30% UDA (A-C), 20% UDA (D-F), and 10% UDA (G-I). (bottom) The crosssectional profiles corresponding to the dashed lines in parts G-I (scale bar corresponds to 1 µm).
calculations. The trend after amine-DNA modification is apparent and more applicable to the current discussion (Figure 5B). The surface potential difference is high at low coverage and decreases as UDA coverage increases. This is similar to the results summarized in Figure 4. The bulkiness of the DNA molecule requires space for it to be able to stand in an upright conformation. With an increased coverage, the DNA molecules are forced to lie more parallel to the GaP surface, exposing their negatively charged phosphate backbone. This results in a more negative surface potential. KPFM Progression with Complementary DNA. The 10, 20, and 30% UDA patterned samples were exposed to aminemodified DNA and subsequently incubated in a solution of a Cy3-labeled complementary strand. A KPFM progression is shown in Figure 6. The 30% progression (Figure 6A-C) shows an expected decrease in contact potential difference after amine-DNA addition. However, after the cDNA strand is added, there is a notable increase in surface potential along the patterned portion. These spots of increased surface potential can be attributed to the salt counterions binding to the hybridized DNA in order to screen the negative charge of the DNA backbone. Figure 6D-F shows the progression of the 20% UDA concentration. As expected, there is a decrease in potential difference after amine-DNA modification followed by a further decrease after hybridization with the complementary strand. Figure 6G-I shows the progression with a 10% UDA concentration. An obvious increase in surface potential is evident after the amine-DNA modification which was noted earlier. After hybridization, a decrease in potential down to the original UDA potential is observed. This phenomenon can be attributed to compensation of the DNA dipole moment by the complementary strand.27 A few high potential spots on Figure 6I indicate areas where hybridization did not occur, most likely caused by an agglomeration of DNA which limited its exposure to the complementary strand. Figure 6 also depicts a representative
We have characterized changes in surface potential on GaP associated with functionalization with DNA molecules. AFM topographical images suggest a more ordered adlayer of DNA when using a solution of 10% UDA compared to a 20 and 30% solution. KPFM further confirms this finding by demonstrating an increase in surface potential after DNA immobilization when using a 10% solution due to the dipole moment inherent in single stranded DNA. A decrease in surface potential when using 20 and 30% solutions indicates exposure of the negatively charged DNA backbone and, thus, a more disordered adlayer. Taken together, XPS data and KPFM indicate that a higher coverage of UDA results in decreased surface potential after DNA immobilization which is explained by the exposure of the negatively charged DNA backbone. Therefore, a lower concentration of linker molecules seems to allow for a more ordered layer of DNA, most likely due to the bulkiness of the DNA molecule. Further KPFM imaging indicates the binding of the cDNA strand to the surface, and Raman spectroscopy confirms it. The findings in this paper can be applied to a variety of fields, including the development of biosensor devices. Acknowledgment. Instructional materials used to engage undergraduate students in this research were made possible by National Science Foundation Undergraduate Research Center award (CHE-0418902) that supports the Center for Authentic Science Practice in Education (CASPiE). References and Notes (1) Stutzmann, M.; Garrido, J. A.; Eickhoff, M.; Brandt, M. S. Phys. Status Solidi A 2006, 203, 3424–3437. (2) Uno, T.; Tabata, H.; Kawai, T. Anal. Chem. 2007, 79, 52–59. (3) Sato, S.; Hirano, A.; Takenaka, S. Anal. Chim. Acta 2010, 665, 91–97. (4) Steichen, M.; Brouette, N.; Buess-Herman, C.; Fragneto, G.; Sferrazza, M. Langmuir 2009, 25, 4162–4167. (5) Saoud, M.; Blaszykowski, C.; Ballantyne, S. M.; Thompson, M. Analyst 2009, 134, 835–837. (6) Chen, R. J.; Bangsaruntip, S.; Drouvalakis, K. A.; Kam, N. W. S.; Shim, M.; Li, Y. M.; Kim, W.; Utz, P. J.; Dai, H. J. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 4984–4989. (7) Bunimovich, Y. L.; Shin, Y. S.; Yeo, W. S.; Amori, M.; Kwong, G.; Heath, J. R. J. Am. Chem. Soc. 2006, 128, 16323–16331. (8) Cui, Y.; Wei, Q. Q.; Park, H. K.; Lieber, C. M. Science 2001, 293, 1289–1292. (9) Gao, Z. Q.; Agarwal, A.; Trigg, A. D.; Singh, N.; Fang, C.; Tung, C. H.; Fan, Y.; Buddharaju, K. D.; Kong, J. M. Anal. Chem. 2007, 79, 3291–3297. (10) Wei, F.; Qu, P.; Zhai, L.; Chen, C. L.; Wang, H. F.; Zhao, X. S. Langmuir 2006, 22, 6280–6285. (11) Hallstrom, W.; Martensson, T.; Prinz, C.; Gustavsson, P.; Montelius, L.; Samuelson, L.; Kanje, M. Nano Lett. 2007, 7, 2960–2965. (12) Richards, D.; Zemlyanov, D.; Ivanisevic, A. Langmuir 2010, 26, 8141–8146. (13) Frolov, L.; Rosenwaks, Y.; Richter, S.; Carmeli, C.; Carmeli, I. J. Phys. Chem. C 2008, 112, 13426–13430. (14) Martz, J.; Zuppiroli, L.; Nuesch, F. Langmuir 2004, 20, 11428– 11432. (15) Barbet, S.; Aubry, R.; di Forte-Poisson, M. A.; Jacquet, J. C.; Deresmes, D.; Melin, T.; Theron, D. Appl. Phys. Lett. 2008, 93, 3. (16) Glatzel, T.; Sadewasser, S.; Shikler, R.; Rosenwaks, Y.; Lux-Steiner, M. C. Mater. Sci. Eng., B 2003, 102, 138–142. (17) Rosenwaks, Y.; Shikler, R.; Glatzel, T.; Sadewasser, S. Phys. ReV. B 2004, 70, 7. (18) Nonnenmacher, M.; Oboyle, M. P.; Wickramasinghe, H. K. Appl. Phys. Lett. 1991, 58, 2921–2923.
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(19) Leung, C.; Kinns, H.; Hoogenboom, B. W.; Howorka, S.; Mesquida, P. Nano Lett. 2009, 9, 2769–2773. (20) Magid, I.; Burstein, L.; Seitz, O.; Segev, L.; Kronik, L.; Rosenwaks, Y. J. Phys. Chem. C 2008, 112, 7145–7150. (21) Gao, P.; Cai, Y. G. Anal. Bioanal. Chem. 2009, 394, 207–214. (22) Opdahl, A.; Petrovykh, D. Y.; Kimura-Suda, H.; Tarlov, M. J.; Whitman, L. J. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 9–14. (23) Schreiner, S. M.; Shudy, D. F.; Hatch, A. L.; Opdahl, A.; Whitman, L. J.; Petrovykh, D. Y. Anal. Chem. 2010, 82, 2803–2810.
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JP105927T
