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Direct Patterning of Molecularly Imprinted Microdot Arrays for Sensors and Biochips Fanny Vandevelde,† Thierry Leı¨chle´,‡ Ce´dric Ayela,‡ Christian Bergaud,‡ Liviu Nicu,*,‡ and Karsten Haupt*,† Compie` gne UniVersity of Technology, CNRS UMR 6022, BP 20529, 60205 Compie` gne Cedex, France, and Laboratory of System Architecture and Analysis, CNRS UPR 8001, 31077 Toulouse, France ReceiVed February 5, 2007. In Final Form: March 23, 2007 We have used fountain pen microlithography to deposit arrays of molecularly imprinted polymer microdots on flat substrates. We visualize analyte binding to the dots by fluorescence microscopy with the aid of fluorescein as a model analyte. Elution and readsorption of the analyte to the MIP dots were possible if the porosity of the dots was improved by a sacrificial polymeric porogen. The imprinting effect was confirmed by using compounds structurally related to fluorescein. In addition, we show with another MIP specific to 2,4-D that, apart from the direct measurement of the binding of fluorescent compounds, a competitive immunoassay-type format can also be used to transduce the binding. We believe that this technique has a strong potential for the fabrication of biomimetic microchips and other types of integrated biosensors.
Introduction Biochips are arrays of small dots of biological molecules on a substrate used for the high-throughput detection and screening of target molecules, an area that is currently drawing considerable interest.1,2 A recent trend goes toward the miniaturization of these chips, which is expected to increase their portability tremendously, thus expanding the use of these arrayed biosensors to point-of-care clinical testing, environmental monitoring, security, and so forth.3 In biosensors, a chemical or physical signal is generated upon the binding of the target analyte to a recognition element. A transducer then translates this signal into a quantifiable output signal. The recognition elements in biosensors are in most cases biomacromolecules such as enzymes or antibodies. However, these molecules are far from perfect for this type of application: they are unstable out of their native environment, and moreover, a natural receptor for the particular target analyte of interest may not always exist. The design and synthesis of biomimetic receptor systems capable of binding a target molecule with similar affinity and specificity to their natural counterparts has long been a goal of bioorganic chemistry. One technique that is being increasingly adopted for the generation of artificial macromolecular receptors is molecular imprinting.4-6 A target molecule (or a derivative thereof) acting as a molecular template is utilized to direct the assembly of specific binding monomers, usually followed by a polymerization step. The thusobtained molecularly imprinted polymer (MIP) contains binding cavities that show selectivity for the target. These materials have a higher physical and chemical stability than biomacromolecules and can more easily be integrated in industrial fabrication processes. These properties make MIPs potentially very suitable as recognition elements for chemical sensors, biosensors, and biochips.7 * Corresponding authors. E-mail:
[email protected],
[email protected]. † CNRS UMR 6022. ‡ CNRS UPR 8001. (1) Espina, V.; Mehta, A. I.; Winters, M. E.; Calvert, V.; Wulfkuhle, J.; Petricoin, E. F., III; Liotta, L. A. Proteomics 2003, 3, 2091. (2) Pirrung, M. C. Ang. Chem., Int. Ed. 2002, 41, 1276. (3) Lynch, M.; Mosher, C.; Huff, J.; Nettikadan, S.; Johnson, J.; Henderson, E. Proteomics 2004, 4, 1695. (4) Arshady, R.; Mosbach., K. Makromol. Chem. 1981, 182, 687. (5) Wulff, G.; Sarhan, A. Ang. Chem., Int. Ed. 1972, 11, 341. (6) Haupt, K. Anal. Chem. 2003, 75, 376.
For multisensors and biochips, MIPs have to be patterned on surfaces and interfaced with a transducer. For this purpose, two different strategies can be considered. The prepolymerization mixture can be patterned in a precise configuration on the transducer surface and then polymerized in situ, or prepolymerized MIP nanoelements are deposited. Patterning methods that may be used with MIPs are soft lithography, microspotting techniques, and localized polymerization. There has been a first report on the use of soft lithography in combination with MIPs, using poly(dimethylsiloxane) (PDMS) stamps.8 However, one of the problems seems to be that the current imprinting recipes are not always compatible with the PDMS stamps used, which tend to swell in certain organic solvents. A recent report describes the use of microcontact printing of protein arrays into thin MIP films.9 To our knowledge, there is no report as yet of the use of localized in-situ polymerization for the preparation of MIP arrays. One possible approach could be an extension of continuous wave laser polymerization, which has been used by Shea and colleagues for microstereolithography of MIPs.10 Well adapted to microchip fabrication are standard microspotting techniques such as ink jetting11 and mechanical microspotting.12 For example, arrays of silicon microcantilevers have been used to deposit biomolecules onto glass slides.13 Depending on the solution to be deposited, the surface, and the cantilever, the diameter of the dots can vary but is normally in the higher micrometer range. In this letter, we describe the deposition of a prepolymerization solution for MIP synthesis with arrays of microcantilevers. Using a fluorescent model template, we show that these microdots are indeed molecularly imprinted and that the morphology of the dots has to be optimized for them to be useful as sensing elements. (7) Haupt, K. Chem. Commun. 2003, 171. (8) Yan, M.; Kapua, A. Anal. Chim. Acta 2001, 435, 163. (9) Lin, H. Y.; Hsu, C. Y.; Thomas, J. L.; Wang, S. E.; Chen, H. C.; Chou, T. C. Biosens. Bioelectron. 2006, 22, 534. (10) Conrad, P. G.; Nishimura, P. T.; Aherne, D.; Schwartz, B. J.; Wu, D.; Fang, N.; Zhang, X.; Roberts, M. J.; Shea, K. I. AdV. Mater. 2003, 15, 1541. (11) Blanchard, A. P.; Kaiser, R. J.; Hood, L. E. Biosens. Bioelectron. 1996, 11, 687. (12) Schena, M.; Schalon, D.; Davis, R. W.; Brown, P. O. Science 1995, 270, 467. (13) Belaubre, P.; Guirardel, M.; Garcia, G.; Leberre, V.; Dagkessamanskaia, A.; Tre´visiol, E.; Franc¸ ois, J. M.; Pourciel, J. B.; Bergaud, C. Appl. Phys. Lett. 2003, 82, 3122.
10.1021/la700320n CCC: $37.00 © 2007 American Chemical Society Published on Web 05/02/2007
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Figure 1. (a) Scanning electron microscope image of the tip of a silicon cantilever showing the integrated channel. (b) Photograph of the cantilevers during deposition on a glass slide.
Experimental Section Materials. Trimethylolpropane trimethacrylate (TRIM), 4-vinylpyridine (4-VP), poly(vinyl acetate) (PVAc, MW ) 140 000 g/mol), and 2,2-dimethoxy-2-phenylacetophenone (DPAP) were from Aldrich. Dimethylsulfoxid (DMSO), diethylene glycol dimethyl ether (diglyme), and fluorescein were from Fluka. All other chemicals were of analytical grade. 4-VP was vacuum distilled before use and kept in the dark at -20 °C. Molecularly Imprinted Polymers. MIPs were prepared using a 1:8:16 molar ratio between the template (fluorescein), the functional monomer (4-VP), and the cross linker (TRIM), respectively. The amount of polymerization initiator (DPAP) used was 1.2% relative to the number of moles of polymerizable groups. A 1:1 mixture of diglyme and DMSO was used as the porogen, which in some cases contained 1% PVAc as a co-porogen. The volume ratio between the porogen and the monomers was 3:1. Polymerization was initiated under UV light (360 nm, 6 W low-pressure lamp) in a nitrogen atmosphere. Microdot Deposition Tool and Procedure. MIP microdots were deposited on 10 × 10 mm2 glass slides. The slides were first cleaned and silanized with methacryloylamidopropyl trimethoxysilane, allowing for covalent bonding of the polymer layer. The deposition system consists of a silicon-based microcantilever array fixed on a three-stage automated spotter. The chip array, including 10 cantilevers, is fabricated using conventional microfabrication techniques.14 The cantilevers, shown in Figure 1, are 1500 µm long, 120 µm wide, and 5 µm thick. A fluidic channel is incorporated in the cantilever tip for liquid loading and deposition. The deposition protocol starts with loading the cantilevers. The cantilever tips are dipped for a few seconds into a reservoir containing the polymerization mixture, with the channel being filled by capillary forces. The cantilever array is then moved above the activated glass slide using an XYZ motion control system. Droplet deposition occurs by direct contact of the cantilever tip with the surface, allowing the liquid to be transferred onto the substrate. Matrices of dots are obtained by combining the deposition step with XY translation, without the (14) Leı¨chle´, T.; Saya, D.; Pourciel, J.-B.; Mathieu, F.; Nicu, L.; Bergaud, C. Sens. Actuators, A 2006, 132, 590.
Figure 2. Fluorescence microscope images of microdot matrices of the nonimprinted control polymer (left), positive control with the copolymerized template (middle), and imprinted polymer (right) synthesized with DMSO-diglyme as the porogen (a) directly after deposition, (b) after washing and elution of the template, and (c) after reincubation in a solution of 30 µM fluorescein. (Bottom row) white-light image of control dots. need to refill the cantilevers. The resulting droplets typically exhibit micrometric dimensions and picoliter volumes. After the deposition of the matrix, the slides were placed in a nitrogen atmosphere, and the matrix was irradiated for 10 min at room temperature (6 W, 365 nm, low-pressure mercury lamp). The microdot arrays on glass were then washed three times with a 1:4 mixture of acetic acid and ethanol, followed by three washing cycles with pure ethanol, and dried in a stream of nitrogen. Binding Experiments. MIPs were incubated in a 30 µM solution of fluorescein in DMSO for 2 h at room temperature. The surfaces were rinsed with acetone and dried in a stream of nitrogen. The fluorescence intensity of the microdots was observed using a fluorescent microscope (LEICA DMLB) with 480 nm EX and 527 nm EM filters. Surface Characterization of Microdots. Images of the microdot topography were acquired in tapping mode using a Digital Instruments Dimension 3100 AFM. The surfaces were also imaged with a Philips XL30 environmental SEM after vacuum deposition of a 5 nm gold film.
Results and Discussion Glass surfaces with attached MIP microdots were obtained on the basis of a copolymer of TRIM and 4-VP, with fluorescein as the imprinting template. As a positive control, microdot matrices were produced with a polymerizable fluorescein derivative, fluorescein methacrylate. This was done to rule out any matrix or solvent/pH effects on the fluorescence of the template. Moreover, non-imprinted microdot matrices were deposited without the addition of the template fluorescein to the monomer mixture. Figure 2a shows fluorescence microscope images of the dots directly after synthesis. As expected, only the MIP dots and the positive control are fluorescent. After washing and elution of the template (Figure 2b), one would expect to observe no fluorescence of the MIP dots but rather fluorescence of the positive control. However, the MIP dots are still fluorescent, indicating that the template could not be eluted. After rebinding experiments by incubation in a fluorescein solution in DMSO
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Figure 3. Fluorescence microscope images of microdot matrices of the nonimprinted control polymer (left), positive control with the copolymerized template (middle), and imprinted polymer (right) synthesized with DMSO-diglyme-1% PVAc as the porogen (a) directly after deposition, (b) after washing and elution of the template, and (c) after reincubation in a solution of 30 µM fluorescein. (Bottom row) white-light image of control dots.
(Figure 2c), no significant change in fluorescence is observed upon reincubation in fluorescein. (The intensity of the positive control dots increased by an average of 2%.) It should be noted that the nonimprinted control dots do not bind the fluorescein template. The impossibility of eluting the fluorescein template from the MIP dots might be due to either covalent attachment of the template to the polymer during radical polymerization or nonaccessibility of the binding sites created in the polymer. During earlier work on the generation of MIP films by spin coating monomers followed by conventional UV photopolymerization, we have found that a potential problem during the synthesis of thin films was their fast polymerization, which prevented pore formation by the nucleation-based phase-separation mechanism. The thus-obtained films were virtually nonporous and showed no binding capacity for the template. We were able to address this problem by using a sacrificial polymeric porogen,15,16 which accelerated phase separation and enhanced pore formation presumably by a spinodal decomposition mechanism. Because a similar situation might be at the origin of the above-mentioned problems with the microdots, we decided to test the same remedy, that is, the addition of a polymeric co-porogen, polyvinyl acetate. Figure 3 shows images of microdot arrays synthesized with a porogenic solvent containing 1% PVAc. The image directly after dot deposition (Figure 3a) is very similar to the one obtained from dots synthesized without PVAc except that the MIP dots appear to be larger. This could be due to a change in the surface wetting properties of the solution after addition of the PVAc. However, because this does not seem to be the case for the (15) Schmidt, R. H.; Mosbach, K.; Haupt, K. AdV. Mater. 2004, 16, 719. (16) Schmidt, R. H.; Haupt, K. Chem. Mater. 2005, 17, 1007.
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positive control, another possible explanation is that some porogen mixture containing template spreads onto the surface after polymerization. After washing and template elution (Figure 3b), we observe virtually no fluorescence of the MIP dots, whereas the positive control is still fluorescent. This confirms that the template has indeed been removed from the MIP dots and that the change in fluorescence is not simply due to a pH effect. (The fluorescence of fluorescein is pH-dependent, and acetic acid was used in some of the washing steps.) To confirm the effect of the PVAc co-porogen on the morphology of the microdots, we analyzed the surfaces of the MIP dots by atomic force microscopy. Representative 5 × 5 µm2 images of the surface topography are shown in Figure 4. Whereas the surface of dots synthesized without PVAc as co-porogen appears to be relatively featureless (Figure 4a), the addition of 1% of PVAc to the porogenic solvent results in the formation of clearly visible pores (Figure 4b). This indicates that the impossibility of template elution from the MIP microdots without PVAc was indeed due to the absence of porosity, despite a dot thickness of less than 1 µm (600 nm in the case of the nonporous dots, as determined by white-light interferometric microscopy). The presence of pores was confirmed by scanning electron microscopy of the surface. To further demonstrate that the MIP microdots are molecularly imprinted and show selective binding, we performed two additional control experiments. First, a competitive binding experiment was carried out using as the competitor 9-hydroxyxanthene, a smaller substructure of fluoresceine. Figure 5a shows that the presence of increasing concentrations of the inhibitor during incubation of the MIP and control microdots inhibits fluorescein binding to the MIP dots, whereas the (low) binding to the control dots does not change. We have also used the dye rose bengal, a structural analogue of fluorescein carrying several chlorine and iodine substituents, and have evaluated its binding to the dot arrays. With this molecule, only very weak binding was obtained, wereby no difference between the MIP and the control dots was observed (Figure 5b). This is not surprising and may be explained by the bulky substituents preventing the molecule from entering the imprinted sites. These two experiments confirm the existence of specific binding sites in the MIP. Finally, because fluorescein-MIP is only a model system, we have also deposited dots of another MIP specific to the 2,4dichlorophenoxyacetic acid (2,4-D) herbicide. This MIP is similar to the one described in one of our previous publications,17 but the polymer recipe used here was identical to that of the fluorescein MIP in order to adapt it for dot deposition. To evaluate the performance of the MIP, a competitive binding assay was performed with 7-carboxymethoxy-4-methylcoumarin (CMMC) as a structurally related fluorescent probe. As competitors, the original 2,4-D template molecule and the structurally related phenoxyacetic acid (POAc) were used. As can be seen in Figure 5c, the presence of 2,4-D at increasing concentrations competes for the binding of the probe, whereas POAc is much less effective as a competitor. This again confirms that our polymer microdots can be molecularly imprinted and at the same time indicates that a competitive pseudofluoroimmunoassay format can be used as the transduction method with these biomimetic microchips. In conclusion, we have used fountain pen microlithography to deposit arrays of molecularly imprinted polymer microdots on flat substrates. We were able to show analyte binding to the dots by fluorescence microscopy with the aid of a fluorescent model analyte. No binding to nonimprinted control dots was observed. We found that the elution and readsorption of the (17) Haupt, K.; Mayes, A.; Mosbach, K. Anal. Chem. 1998, 70, 3936.
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Figure 4. Atomic force microscope image (5 × 5 µm2, tapping mode) of the surfaces of microdots synthesized (a) without and (b) with the addition of polyvinylacetate (MW 140 000) to the porogenic solvent. (c) Scanning electron microscope image of pores on a surface of a microdot synthesized with the addition of polyvinylacetate.
Figure 5. Fluorescence microscopy studies of imprinted (gray bars) and nonimprinted control dots (white bars). The fluorescence intensity of the dots was quantified using ImageJ software (public domain, http://rsb.info.nih.gov/ij/). All measurements are mean values of three dots and were corrected for the background signal. (a) Template fluorescein: relative fluorescence after incubation in a 30 µM fluorescein solution in the presence of different concentrations of 9-hydroxyxanthene (480 nm EX-527 nm EM). (b) Template fluorescein: relative fluorescence after synthesis (1), after template elution (2), after incubation in 30 µM fluorescein (3) (all 480 nm EX - 527 nm EM), after washing (4), and after incubation in 30 µM of rose bengal (5) (all 546 nm EX-600 EM). (c) Template 2,4-D: relative fluorescence after incubation in 30 µM CMMC (340 nm EX-425 EM) in the presence of 2,4-D (1, 0 µM; 2, 30 µM; 3, 300 µM) and POAc (4, 0 µM; 5, 30 µM; 6, 300 µM).
analyte to the MIP dots was possible only if the dots were rendered porous by a technique based on a sacrificial polymeric porogen. The imprinting effect was confirmed by using molecules structurally related to fluorescein as competitors. In addition, we have shown, with another MIP, that apart from the direct measurement of the binding of fluorescent compounds a competitive immunoassay-type format can also be used to transduce the binding. Because of the ability to deposit dots of functional MIPs at precise and desired locations in a parallel manner, we believe that this technique has a strong potential for
the fabrication of highly integrated biomimetic microchips as well as other types of integrated biosensors. Acknowledgment. We gratefully acknowledge financial support by the CNRS Materials program. F.V. holds a Ph.D. fellowship from the French Ministry of Research and Higher Education. Partial financial support by the EC-funded project NaPa (contract no. NMP4-CT-2003-500120) is gratefully acknowledged. We thank Mr. Alexis Gautier for the interferometric microscopy measurements. LA700320N
