Fabrication of Biological Nanostructures by Scanning Near-Field

NANO LETTERS

Fabrication of Biological Nanostructures by Scanning Near-Field Photolithography of Chloromethylphenylsiloxane Monolayers

2006 Vol. 6, No. 1 29-33

Shuqing Sun,† Matthew Montague,† Kevin Critchley,‡ Mu-San Chen,§ Walter J. Dressick,§ Stephen D. Evans,‡ and Graham J. Leggett*,† Department of Chemistry, UniVersity of Sheffield, Brook Hill, Sheffield S3 7HF, U.K., School of Physics and Astronomy, UniVersity of Leeds, Leeds, LS2 9JT, U.K., and Center for Bio/Molecular Science and Engineering (Code 6900), NaVal Research Laboratory, 4555 OVerlook AVenue, SW, Washington, D.C. 20375-5348 Received September 8, 2005; Revised Manuscript Received November 21, 2005

ABSTRACT We demonstrate the fabrication of sub-100-nm DNA surface patterns by scanning near-field optical lithography using a near-field scanning optical microscope coupled to a UV laser and a chloromethylphenylsiloxane (CMPS) self-assembled monolayer (SAM). The process involves 244-nm exposure of the CMPS SAM to create nanoscale patterns of surface carboxylic acid functional groups, followed by their conversion to the N-hydroxysuccinimidyl ester and reaction of the active ester with DNA to spatially control DNA grafting with high selectivity.

The organization of molecules into nanometer-scale patterns is a key challenge in nanoscale science and technology.1 A variety of approaches have been developed for the manipulation of adsorbate systems to fabricate templates that may be functionalized with molecular,2 biomolecular,3,4 and particulate species.5 For example, dip-pen nanolithography6 may be used to deposit a variety of inks onto solid substrates, and local oxidation7 via the application of an electrostatic potential to an AFM tip has been used to fabricate structures on self-assembled monolayers (SAM), semiconductors, and other surfaces. Photochemical techniques have proven to be powerful as a means of manipulating matter on larger-length scales because they offer extraordinary versatility and selectivity through a variety of means, including the selection of wavelength- and dose-dependent reaction pathways. For example, light-directed solid-phase synthesis8 is a powerful and highly efficient route to the fabrication of oligonucleotide arrays on solid substrates. However, photochemical methods have been explored little for the fabrication of molecular nanostructures. Recently we showed that the selective photochemical oxidation of alkanethiols could be achieved on length scales significantly beyond the conventional diffraction limit using a near-field scanning optical microscope (NSOM) coupled * Corresponding author. E-mail: [email protected]. † University of Sheffield. ‡ University of Leeds. § Naval Research Laboratory. 10.1021/nl051804l CCC: $33.50 Published on Web 12/09/2005

© 2006 American Chemical Society

to a UV laser.9,10 However, it was unclear to what extent this approach could be applied to other systems, and as yet there have been no demonstrations of its utility for the fabrication of functional molecular nanostructures. Here we present important new evidence that NSOM provides a versatile means for modification of surface structure (not restricted to alkanethiol systems) by demonstrating, for the first time, the fabrication of molecular nanostructures using carboxylic acids prepared by UV photolysis of chloromethylphenyl groups in siloxane SAMs. In particular, we show that it is possible to exploit the carboxylate photoproducts to fabricate arrays of nanospots of DNA. These data suggest that all of the flexibility and selectivity associated with photochemistry on larger-length scales may be directly transferable to nanofabrication, enabling a wide range of possibilities for the construction of complex molecular architectures. In our first reports of scanning near-field photolithography (SNP),9,10 a frequency-doubled argon ion laser coupled to a NSOM was used to selectively convert alkanethiolates to alkylsulfonates that could be displaced in a subsequent solution-phase step. Using this approach, it proved possible to match the resolution of electron beam lithography for the same materials.10 We hypothesized that the key requirement for achieving this level of resolution was the excitation of a specific photochemical reaction in a group distributed with monolayer coverage on a solid surface.

Figure 1. Reaction scheme for the oxidation of CMPS to first, an aldehyde, and subsequently on extended exposure, a carboxylic acid. [O] represents an adventitious oxygen source, such as O2 or H2O, at the SAM-air interface during exposure. The variation in contact angle as a function of a 244-nm exposure dose is shown in a, and the variation in the C1s region of the XPS spectrum as a function of exposure is shown in b. The C1s spectra have been normalized to facilitate comparison of the line shapes at different exposures. XPS N1s data for 2,4-dinitrophenylhydrazine-derivatized samples preexposed to varying amounts of irradiation are shown in c.

4-chloromethylphenylsiloxane (CMPS) SAMs, prepared by chemisorption on clean silicon oxide surfaces,11 provide a convenient means to test our hypothesis. Previous studies of chloromethylphenyl group photochemistry in SAMs12 and related thin films13 at 193 nm indicate that C-Cl bond photocleavage predominates, leading ultimately to the formation of surface aldehydes and carboxylic acids. Significant Si-C bond scission, often observed for simple aromatic siloxane SAMs14 at 193 nm, occurs only after C-Cl cleavage is complete, suggesting that similar film stability might be expected at the lower energy wavelength (244 nm) of interest to us here. Furthermore, excitation of chloromethylphenyl groups at ∼250 nm leads to well-characterized photochemical reactions involving radical intermediates in the bulk phase.15 The presence of similar reaction pathways in the SAM, if they occur, would provide an opportunity to generate aldehydes and/or carboxylates as useful surface photoproducts via radical abstractions of oxygen from ambient O2 and/ or H2O. CMPS SAMs were first exposed to light from a frequencydoubled argon ion laser (λ ) 244 nm)16 in the absence of a mask to determine the nature of the surface photoproducts formed during irradiation. The laser beam was passed through a divergent lens to expose a large enough area (ca. 1 cm2) for subsequent characterization by contact-angle measurement and surface spectroscopy. The sessile water drop contact angle fell from 70° to less than 20° during a period of 2 min exposure, corresponding to a dose of 4.8 J cm-2 (Figure 1a). The increase in surface energy was expected if polar functionalities such as carboxylic acid groups were created. After very long exposures (10 times as great) there was no further decrease in the contact angle. To identify the nature of the functional groups introduced to the CMPS monolayer, X-ray photoelectron spectroscopy 30

(XPS) was used. The data are shown in Figure 1. There was evidence of C-Si bond scission as the exposure increased. At a dose of 4.8 J cm-2, the C1s peak area had reduced to 48% of the value for an as-prepared monolayer. This declined to 36% after 9.6 J cm-2. However, in agreement with our initial hypothesis, the Cl signal declined substantially faster. At a dose of 4.8 J cm-2, Cl was undetectable by XPS. This confirms that Cl-C bonds are broken rapidly on exposure of the monolayer to UV light. Although some Si-C bonds are broken, substantial amounts of the adsorbate remain at the surface following exposure at doses in the range studied here. The C1s spectrum of the as-prepared monolayers exhibited a dominant peak centered at ∼285 eV (Figure 1b), with a shoulder at ∼287 eV arising from the carbon atom in the chloromethyl group. After a dose of 0.8 J cm-2, a new peak was evident at ∼289.3 eV, corresponding to the carboxylate carbon atom. Although the shoulder intensity at ∼287 eV also increased, consistent with formation of surface aldehyde groups (∼287.7 eV), a definitive CHO assignment could not be made due to the proximity of the C-Cl (∼287 eV) component of the C1s spectrum. Samples were therefore treated with 2,4-dinitrophenylhydrazine, which selectively converts any CHO groups present to hydrazones.17 The resulting N1s spectra are shown in Figure 1c. Two peaks are evident after an exposure of 0.8 J cm-2, corresponding to the two hydrazone bonding environments of N and confirming the presence of surface aldehyde groups. As exposure dose increased, the carboxylate carbon component of the C1s spectrum increased. These increases were paralleled by decreases in the areas of the N1s peaks of the hydrazone-derivatized samples, reflecting a decrease in the ratio of aldehyde to carboxylic acid groups. Quantitative analysis of the XPS spectra, based on the determination Nano Lett., Vol. 6, No. 1, 2006

Figure 2. Tapping-mode images (100 × 100 µm2) showing the results of immobilization to the active ester derivatives of carboxylic acid groups introduced to CMPS monolayers by photopatterning. (a and b): immobilization of amine-functionalized polymer nanoparticles; (c) control experiment identical to a and b, but without formation of the active ester; (d) immobilization of human plasma fibrinogen.

of the ratio of C(CHO)/C(all) and C(COOH)/C(all) from deconvoluted C1s spectra, revealed that after a dose of 0.8 J cm-2 the fraction of chloromethyl groups converted to aldehydes was 0.30, decreasing to 0.21 after 2.4 J cm-2 and 0.12 after 4.8 J cm-2. The fraction converted to carboxylate carbons was estimated to be 0.19, 0.36, and 0.84, respectively. The fraction of carboxylate carbons changed little after an exposure of 9.6 J cm-2, but the C1s peak area was reduced significantly and the peak was more noisy. It was concluded that at this dose, Si-C bond scission was leading to significant removal of adsorbates from the surface. Having determined the nature and relative abundance of the aldehyde and carboxylate surface photoproducts produced during 244-nm exposure of the CMPS SAMs, the feasibility of utilizing them for patterning was examined. To maximize the efficiency of selectively grafting materials to a patterned surface, it is desirable that a single surface photoproduct of high chemical reactivity be generated during exposure to maximize the surface density of reactive groups and minimize potential cross reactivities that may be associated with minor photoproducts. Although both aldehyde and carboxylic acid photoproducts are formed in our system, carboxylic acids can be made the predominant surface photoproduct by control of the exposure dose. Consequently, we elected to implement grafting chemistries utilizing carboxylic acid species in our work. Our grafting approach adapts methods from conventional peptide chemistry by exploiting the well-known reaction of carboxylic acid active esters18 with amines to form stable Nano Lett., Vol. 6, No. 1, 2006

amide linkages needed to spatially control tethering of materials to patterned surfaces. Similar approaches have been used previously by other workers to immobilize proteins.19,20 To demonstrate the method, we exposed CMPS SAMs at 244 nm (4.8 J cm-2) through a mask (an electron microscope grid) to create surface patterns of carboxylic acid groups, which were imaged using atomic force microscopy (AFM). The patterned samples were then treated under ambient conditions by immersion for 2 h in an ethanolic solution containing N-hydroxysuccinimide (25 mMol dm-3) and 1-ethyl-3, 3-dimethyl carbodiimide (20 mMol dm-3) to convert carboxylic acid groups in the irradiated regions of the SAM to active N-hydroxysuccinimidyl esters, which are reactive toward amine groups.19,20 In a subsequent step, the activated sample was derivatized by polymer nanoparticles by immersion for 1 h in a solution of 10 µL of 0.5% w/v amine-functionalized polymer latex nanoparticles (diameter 24 nm, Interfacial Dynamics Corporation, Portland, OR) in 1 mL of deionized water. Figure 2a and b shows AFM images of the results: nanoparticles have clearly been immobilized on the exposed areas (squares) but not on the masked areas (the bars). As a control, the experiment was repeated in the absence of the activation step, and the resulting AFM image shown in Figure 2c was featureless, confirming the specificity of the attachment process. Poor nanoparticle attachment was noted for samples exposed at doses 14.4 J cm-2 is consistent with scission of the Si-C bond and removal of the adsorbate from the surface. Figure 2d shows a sample that has been activated in the same way (via formation of an active ester) but subsequently derivatized with fibrinogen by immersion in a 10 µg mL-1 solution of the protein (Sigma, Poole, U.K.) in phosphatebuffered saline solution (Sigma) for 18 h. The contrast is weaker than that in Figure 2a and b because of the limited height of the adsorbed protein molecules (ca. 2 nm) and the large scan size, but the pattern is still sharply resolved. To fabricate nanostructures, we coupled an NSOM to the frequency-doubled argon ion laser as the light source (SNP). The NSOM probe was traced across the surface of the sample, leading to localized exposure, and the sample characterized by AFM. At a scan rate of 0.1 µm s-1, it was important not to use too powerful a UV intensity. With the laser output power set to its minimum value (1 mW), surface modification was mild, leading to frictional contrast but no topographical contrast. This indicated that the chemical structure of the surface had been modified without the ablation of adsorbate molecules. However, for higher powers topographical contrast was observed attributed to scission of the Si-C bond, leading to desorption of the organic part 32

of the adsorbate molecule and the localized growth of the oxide film. The immobilization of biomolecules on such heavily modified films was not successful. Figure 3 shows the results of oxidation carried out with a 244-nm laser power of 1 mW. The friction images show concentric rings, each with a line width of 85 nm (Figure 3a), and three parallel lines (Figure 3b), each with a width of only 45 nm. These feature dimensions are substantially beyond the diffraction limit at this wavelength (ca. λ/2 ) 122 nm). The frictional contrast arises because the carboxylic acid functionalized regions exhibit a higher surface free energy than the surrounding areas and adhere more strongly to the polar tip of the AFM as it scans the surface, leading to an increase in the friction force.21 The utility of these patterns for biomolecular patterning is demonstrated in the right-hand side of Figure 3. We followed an approach to DNA immobilization reported previously by Huang et al.22 in which amine groups of DNA molecules are attached to active ester groups. The carboxylic acid groups formed by UV exposure of the CMPTS monolayer have been converted to active esters using the same carbodiimide employed in Figure 2, and then exposed to a solution of calf thymus DNA (4 µg mL-1 in deionized water; Sigma, Poole, U.K.). The reaction of the NHS ester occurs Nano Lett., Vol. 6, No. 1, 2006

at a particularly basic N(7) nitrogen of the deoxyguanosine residue of the DNA strand, most efficiently where there is a pair of adjacent deoxyguanosine residues in the DNA strand.23 The arrays shown in Figure 3 consist of spots of DNA that are only 70 nm in diameter, placed at 500-nm intervals. It is clear that the SNP-patterned CMPS monolayers provide an excellent template for the immobilization of DNA. Given the simplicity and versatility of the carbodiimide coupling chemistry used here, we expect this approach to have widespread applicability. In summary, we have demonstrated for the first time that biomolecular nanostructures may be fabricated simply by exposing siloxane monolayers to UV light from a near-field scanning optical microscope. Controlled 244-nm exposure of 4-chloromethylphenylsiloxane monolayers chemisorbed on silicon oxide leads to the conversion of terminal chloromethyl groups to carboxylic acid groups. The resulting patterns are activated readily under ambient conditions using simple, widely applicable coupling chemistries to facilitate the formation of patterned nanoparticle, protein, and DNA structures. SNP promises to enable a wide variety of approaches for the photochemical manipulation of nanoscale molecular structure. Control of the exposure dose enables selection of one of three photochemical pathways (C-Cl scission leading to aldehyde or carboxylic acid photoproduct or Si-C scission leading to removal of the entire organofunctional group), emphasizing the versatility and control that photochemical nanofabrication methods should offer. Acknowledgment. S.S. and G.J.L. are grateful to the Engineering and Physical Sciences Research Council (EPSRC) (grant GR/N82197/01) and the Biotechnology and Biological Sciences research Council (BBSRC) (grant 50/EGM17711) for financial support. G.J.L. thanks EPSRC and the Royal Society of Chemistry Analytical Chemistry Trust Fund for their support. M.M. thanks the BBSRC for a Research Committee Studentship. References (1) Geissler, M.; Xia, Y. AdV. Mater. 2004, 16, 1249. (2) Kaholek, M.; Lee, W.-K.; LaMattina, B.; Caster, K. C.; Zauscher, S. Nano Lett. 2004, 4, 373. (b) Kaholek, M.; Lee, W.-K.; Ahn, S.-J.; Ma, H.; Caster, K. C.; LaMattina, B.; Zauscher, S. Chem. Mater. 2004, 16, 3688. (3) Hong, B. J.; Oh, S. J.; Youn, T. O.; Kwon, S. H.; Park, J. W. Langmuir 2005, 21, 4257. (b) Lee, K.-B.; Kim, E. Y.; Mirkin, C. A.; Wolinsky, S. M. Nano Lett. 2004, 4, 1869.

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(4) Leggett, G. J. Analyst 2005, 130, 259. (5) Cao, Y.-W. C.; Jin, R.; Mirkin, C. A. Science 2002, 297, 1536. (b) Park, S.-J.; Taton, A.; Mirkin, C. A. Science 2002, 295, 1503. (c) Nam, J.-M.; Thaxton, C. S.; Mirkin, C. A. Science 2003, 301, 1884. (d) Fresco, Z. M.; Frechet, J. M. J. Am. Chem. Soc. 2005, 127, 8302. (e) Brandow, S. L.; Dressick, W. J.; Dulcey, C. S.; Koloski, T. S.; Shirey, L. M.; Schmidt, J.; Calvert, J. M. J. Vac. Sci. Technol., B 1997, 15, 1818. (6) Ginger, D. S.; Zhang, H.; Mirkin, C. A. Angew. Chem., Int. Ed. 2004, 43, 30. (b) Piner, R. D.; Zhu, J.; Xu, F.; Hong, S.; Mirkin, C. A. Science 1999, 283, 661. (c) Lee, K.-B.; Park, S.-J.; Mirkin, C. A.; Smith, J. C.; Mrksich, M. Science 2002, 295, 1702. (7) Maoz, R.; Frydman, E.; Cohen, S. R.; Sagiv, J. AdV. Mater. 2000, 12, 725. (b) Maoz, R.; Frydman, E.; Cohen, S. R.; Sagiv, J. AdV. Mater. 2000, 12, 424. (c) Hoeppener, S.; Maoz, R.; Cohen, S. R.; Chi, L.; Fuchs, H.; Sagiv, J. AdV. Mater. 2002, 14, 1036. (d) Liu, S.; Maoz, R.; Schmid, G.; Sagiv, J. Nano Lett. 2002, 2, 1055. (8) Fodor, S. P. A.; Read, J. L.; Pirrung, M. C.; Stryer, L.; Liu, A. T.; Solas, D. Science 1991, 251, 767. (9) Sun, S.; Chong, K. S. L.; Leggett, G. J. J. Am. Chem. Soc. 2002, 124, 2414. (b) Sun, S.; Leggett, G. J. Nano Lett. 2002, 2, 1223. (10) Sun, S.; Leggett, G. J. Nano Lett. 2004, 4, 1381. (11) Dressick, W. J.; Dulcey, C. S.; Brandow, S. L.; Witschi, H.; Neeley, P. F. J. Vac. Sci. Technol., A 1999, 17, 1432. (12) Brandow, S. L.; Chen, M.-S.; Aggarwal, R.; Dulcey, C. S.; Calvert, J. M.; Dressick, W. J. Langmuir 1999, 15, 5429. (13) Brandow, S. L.; Chen, M.-S.; Fertig, S. J.; Chrisey, L. A.; Dulcey, C. S.; Dressick, W. J. Chem.sEur. J. 2001, 7, 4495. (14) Dressick, W. J.; Calvert. J. M. Jpn. J. Appl. Phys., Part 1 1993, 32, 5829. (15) Miranda, M. A.; Perez-Prieto, J.; Font-Sanchis, E.; Scaiano, J. C. Acc. Chem. Res. 2001, 34, 717. (16) Photopatterning was conducted using light from a frequency-doubled argon ion laser (Coherent FreD 300C, Coherent U.K., Ely) that emits at 244 nm. For scanning near-field photolithography, the laser was coupled to a ThermoMicroscopes Aurora III near-field scanning optical microscope (Veeco UK, Cambridge, U.K.) fitted with a fused silica fiber probe (Veeco, Cambridge, U.K.). The probe scan velocity was 0.1 µm s-1. Contact angles were measured using a Rame-Hart model 100-00 contact angle goniometer. XPS spectra were obtained for spin-cast samples of nanoparticles on an ESCALAB 250 instrument equipped with a monochromatic Al KR X-ray source, using a pass energy of 20 eV and a step size of 0.10 eV. (17) Vogel, A. I. In Textbook of Practical Organic Chemistry, 5th ed.; Furniss, B. S., Hannaford, A. J., Smith, P. W. G., Tatchell, A. R., Eds.; John Wiley: New York, 1989; p 1257. (18) Bodanszky, M. Principles of Peptide Synthesis; Springer-Verlag: New York, 1984; pp 28-49. (19) Corn, R. M.; Frey, B. L. Anal. Chem. 1996, 68, 3187. (20) Patel, N.; Davies, M. C.; Hatshorne, M.; Heaton, R. J.; Roberts, C. J.; Tendler, S. J. B.; Williams, P. M. Langmuir 1997, 13, 6485. (21) Leggett, G. J.; Brewer, N. J.; Chong, K. S. L. Phys. Chem. Chem. Phys. 2005, 7, 1107. (22) Huang, E.; Zhou, F.; Deng, L. Langmuir 2000, 16, 3272. (23) Millan, K. M.; Mikkelsen, S. R. Langmuir 1993, 9, 2317-2323.

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