Attoliter Chemistry for Nanoscale Functionalization of Graphene - ACS

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Attoliter chemistry for nano-scale functionalization of graphene Michael Hirtz, Sarah A. Varey, Harald Fuchs, and Aravind Vijayaraghavan ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b06065 • Publication Date (Web): 30 Nov 2016 Downloaded from http://pubs.acs.org on December 3, 2016

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ACS Applied Materials & Interfaces

Attoliter Chemistry for Nano-Scale Functionalization of Graphene Michael Hirtz,†,‡,* Sarah Varey,⊥,‡ Harald Fuchs, †,∥ and Aravind Vijayaraghavan⊥,* †

Institute of Nanotechnology (INT) & Karlsruhe Nano Micro Facility (KNMF), Karlsruhe

Institute of Technology (KIT), 76344 Eggenstein-Leopoldshafen, Germany. ⊥

School of Materials and National Graphene Institute, The University of Manchester,

Manchester M13 9PL, UK. ∥

Physical Institute and Center for Nanotechnology (CeNTech), University of Münster,

48149 Münster, Germany. KEYWORDS. graphene, click-chemistry, CuAAC, dip-pen nanolithography (DPN), microcontactspotting µCS

ABSTRACT

In order to realize graphene-based chemical and bio-sensors, the nano-scale, multiplexed functionalization of graphene in a device array is a critical step. We demonstrate that graphene can be functionalized with sub-micron resolution and in well-defined locations and patterns using reaction agents in attoliter quantities, utilizing dip-pen nanolithography or microchannel cantilever spotting. Specifically, we functionalize graphene with a biotin azide using clickchemistry and demonstrate the subsequent binding of fluorescently tagged streptavidin. The technique can be scaled up to multiplex functionalize graphene devices on a wafer-scale for sensor and biomedical applications.

ACS Paragon Plus Environment

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MAIN TEXT Among the many potential applications of graphene electronic devices, the ones likely to achieve real-world relevance is sensors.1–3 Graphene sensors have been demonstrated for a variety of gases, chemicals, and biological agents. In all these cases, the design of a graphene sensor follows a few underlying principles. Firstly, the graphene itself is quite a sensitive material, in part due to its atomic thinness which means that every atom of graphene is a surface atom which can respond to the environment. It is also sensitive because graphene is a zero band-gap semiconductor and its conductivity responds to changes in carrier concentration as a result of doping. However, graphene is not intrinsically selective, and the majority of research into graphene sensors has gone towards methods of functionalizing graphene with receptor molecules to impart selectivity to a desired analyte. Another advantage of a graphene sensor is that it can be fabricated on a wafer-scale and can be miniaturized to sub-micron dimensions with conventional photolithography techniques. Also the functionalization of graphene could be multiplexed, and an array of graphene sensors, each with a different functionality targeted at a different response to a mixture of analytes, in conjunction with pattern recognition routines can indeed result in a very powerful miniaturized electronic sensor system for complex mixtures of chemicals such as occurring in biological samples. However, while sub-micron scale physical patterning of graphene can be achieved rather readily by lithography, sub-micron scale chemical patterning of graphene or targeting functionalization of sub-micron scale physical graphene patterns has proved elusive.

Polymer pen lithography (PPL)4 and dip-pen nanolithography (DPN)5 offer high potential for localized

functionalization

of

surfaces.

Previously,


non-covalent

high-resolution

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functionalization of graphene was achieved by DPN with fluorophors6 and phospholipids.7 Covalent PPL printing via Diels-Alder reaction was also demonstrated.8 Both, PPL and DPN were employed in patterning modified silicon oxide surfaces by copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC).9–14 CuAAC15,16 being one of the most utilized click-chemistry reaction due to its orthogonal reactivity and taking place at room temperature. Here, we present the use of microchannel cantilever spotting (µCS) and DPN for highly localized and multiplexed functionalization of graphene by CuAAC (Figure 1). These functionalizations can be subsequently used for selective immobilization of proteins to the graphene, opening up routes to biosensor applications. Micromechanical cleavage of graphite yields graphene flakes of 100 s of µm lateral size, often containing a mixture of regions of monolayer, bi-layer and tri-layer graphene. Figure 2(a) shows an optical micrograph of a region of one such flake. In order to undertake the CuAAC functionalization on the graphene surface, the graphene needs to first be activated to contain exposed alkyne groups. This is achieved via an aryl diazonium salt reaction17 as shown schematically in Figure 1(a). It is well known that this type of diazonium coupling to graphene is an electron-transfer reaction wherein monolayer regions of graphene are significantly more reactive than bi-layer and tri-layer graphene regions, and edges and defects in graphene are even more reactive than monolayers.18,19 Hence, the extent of functionalization should be evident in the Raman spectrum of graphene. Figure 2(b) shows the Raman spectrum obtained at two representative locations on the graphene flake, one in the monolayer and one in the bilayer regions, covering the D (~1350 cm-1), G (~1580 cm-1) and 2D (~2700 cm-1) peak regions, before and after functionalization. The emergence of defect peaks is evidence of some perturbation to the pristine graphene structure,


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presumably from the functionalization by phenylacetylene moieties. An overtone of the D peak, the D´ (~1610 cm-1) peak, appears as a shoulder to the G peak. Raman peaks corresponding to the azo functional groups (indicated as Da at 1400 cm-1 and 1450 cm-1) also occur. Figures 2(c) and (d) compare the spatial maps of the ratio of the D peak integrated intensity (I[D]) to the G peak integrated intensity (I[G]), which is a measure of the extent of defect formation and in turn a measure of the extent of reaction on the surface. The individual maps of the G, D and 2D peak intensities and representative spectra from regions of 1 to 4 layers, edge and wrinkles are shown in supplementary Figure S1. It can be clearly seen that the monolayer graphene region shows a much higher I(D)/I(G) ratio compared to bi-layer and tri-layer regions. A streak of very high I(D)/I(G) is also seen across the monolayer region which we attribute to a wrinkle (line defect) in the graphene flake. This is attributed to a wrinkle rather than a tear because this region shows a very high 2D peak intensity and a very low D peak intensity before functionalization. Interestingly, the reactivity of the wrinkle in monolayer graphene is significantly high whereas reactivity of the wrinkle in bilayer graphene is indistinguishable from a flat bilayer region. This reactivity tendency is consistent with what has been reported for such diazonium coupling reactions on graphene. It has been reported that this type of reaction involving electron rich reagents can proceed beyond a simple monolayer of functional groups, and result in a polymerization to form poly-azo compounds on the graphene surface.20 A significant intensity of azo group Raman signature might be interpreted as evidence of such overreaction; however, this does not negatively affect the creation of reactive alkyl groups on the graphene surface. Furthermore, for diazonium salts having weakly electron withdrawing substituents in the para position such as the -C≡C-SiMe3, such secondary reactions are to be expected,20 but their impact is limited. The resulting graphene flakes contain a high density of reactive alkyne groups, which


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are protected by the trimethylsilyl (TMS) group. This protection group stabilizes the substrates for storage and is cleaved off just shortly before the patterning takes place to achieve best reactivity. This reaction, and the subsequent reaction with CuAAC, have been previous demonstrated in dispersions of graphene flakes.21 The use of microchannel cantilevers22,23 allows for the deposition of femtoliter sized droplets of ink that can act as reaction vessels for the CuAAC reaction to take place.24,25 We utilized a water based ink containing the azide compound to be immobilized, catalysts for the CuAAC and glycerol (to prevent drying out of the microdroplets). After loading of the microchannel cantilevers, spotting is carried out by bringing them into contact with the sample surface for controlled amount of time and allow the ink to flow down to the substrate by capillary forces. See Supporting Information Figure S2 for subsequent images of a spotting process. Assuming the droplets to be half spheres (as suggested by a contact angle about 90° of water on graphene),26 and taking a typical droplet diameter of ~8 µm, the typical reaction volumes would be estimated to be about 130 femtoliters. Typical results of this procedure are shown in Figure 3. To ensure, that the CuAAC was taking place as expected, first, biotin azide ink with all needed catalysts (‘+ Cu’) was spotted next to a negative control, with no catalysts in the ink (‘no Cu’), as seen in Figure 3a. After resting for 2 h to allow the reaction to take place, sample was washed, thus removing the reaction vessel droplets (Figure 3b). In order to reveal the surface bound biotin azide, selective binding of labelled streptavidin was performed.27 After protein binding, only the area previously covered with the catalyst containing ink lights up in fluorescence, while the area that was carrying the ‘no Cu’ ink remains dark (Figure 3c). This indicates, that – as expected – the CuAAC only took place in fast pace when catalysts were present, while for the areas without catalysts the washing steps could easily remove the not covalently bound biotin-azide.


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To validate the feasibility of this route for selective protein binding further, microdroplets of azide functionalized with dinitrophenol (DNP-azide) and biotin-azide were spotted (Figure 3d). After removing of the microdroplets by washing with PBS (Figure 3e), the sample was incubated with streptavidin as described above. Here, again exclusively the area functionalized with the biotin-azide lights up in fluorescence, while the DNP-azide functionalized area remains dark (Figure 3f). Additional images of another sample are provided in Supporting Information Figure S3. These results indicate, that the protein binding also remains selective in presence of other azides / binding motifs on the graphene flakes. To demonstrate high-resolution patterning, we also employed DPN for immobilization of biotinylated azide onto graphene (an in-situ image of a writing process is given in Supporting Information Figure S4). A typical result can be seen in Figure 4. After lithography has taken place, no fluorescence is visible at the patterned site (Figure 4a). After blocking and incubation of streptavidin as described above, the pattern emerges in fluorescence due to selective binding of the protein to the pattern (Figure 4b). The grid pattern has an overall size of 10 × 10 µm² with subunits of 2 × 2 µm². The linewidth as observed in the pattern are about 500 to 700 nm, bringing the correspondent reaction vessels into the attoliter regime. Individual droplets of this diameter corresponds to a reaction volume of 32 to 90 attoliters. In conclusion, we have successfully demonstrated that bioactive molecules can be grafted on to graphene in a multiplexed fashion, with the reaction restricted to sub-micron sized regions. The functionalized graphene was successfully used to selectively bind proteins. The µCS and DPN techniques can be extended from the single-cantilever writing used here to parallel functionalization using multi-cantilever arrays from tens to several ten thousand of individual cantilevers,28,29 each with potentially a different reaction agent delivering a different functional


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group on the graphene surface. Both graphene and graphene oxide can be functionalized in numerous ways, using the vast toolkit of organic chemistry and biochemistry.30,31 By utilizing this ultra-low volume and highly localized chemical delivery and immobilization, we can envisage a system where an array of micro-scale graphene sensor devices on a wafer scale can be functionalized with a variety of reactive groups each with a different reaction profile to a mixture of analytes, leading to scalable fabrication of a true miniaturized graphene based lab-on-a-chip system. ASSOCIATED CONTENT Supporting Information. Additional optical and Raman characterization of functionalized graphene flakes, Images of the lithography process. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Authors *E-mail: [email protected] *E-mail: [email protected] Notes The authors declare no competing financial interest. Author Contributions ‡These authors contributed equally. ACKNOWLEDGMENT


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This work was partly carried out with the support of the Karlsruhe Nano Micro Facility (KNMF, www.kmf.kit.edu), a Helmholtz Research Infrastructure at Karlsruhe Institute of Technology (KIT, www.kit.edu). SV was supported by the EPSRC Northwest Nanoscience Centre for Doctoral Training grant EP/L01548X/1 and AV acknowledges funding from EPSRC grants EP/K009451/1 and EP/G035954/1. REFERENCES (1)

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Figure 1. Scheme of localized functionalization showing (a) the preparation route for the graphene substrates, (b) writing process by µCS (left) performing chemistry in femtoliter volumes and DPN (right) in attoliter volumes. (c) CuAAC reaction scheme for immobilization of azide compound.


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Figure 2. Activation of graphene surface by diazonium functionalization. (a) optical micrograph of graphene flake, with the red square showing the region mapped by Raman spectroscopy, (b) Raman spectra including the D, G, D´ and 2D peaks of graphene and the azo group (Da) peaks for monolayer (1L) and bilayer (2L) regions of the graphene flake before and after diazonium functionalization. (c – d) Map of D to G peak intensity ratio of the graphene flake in the region of (a) marked by the red boarder, showing 1L, 2L, edge (E) and wrinkle (W) regions, before (c)


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and after (d) diazonium functionalization. Complete maps and sample spectra are shown in Figure S1.

Figure 3. Graphene flakes with spotting by microchannel cantilevers. (a) Microdroplets of catalyst containing biotin-azide ink (‘+ Cu’) and negative control (‘no Cu’) on a graphene flake. (b) After washing of the sample, droplets are removed, respective areas are marked. (c) After selective binding of fluorescently labelled streptavidin, only the area functionalized with the catalyst containing ink lights up in fluorescence. This indicates that CuAAC is taking place as expected. (d) Microdroplets of DNP- azide and biotin-azide on a graphene flake. (e) After washing both droplets are completely removed. (f) On incubation with streptavidin, selective binding to the biotin-containing area takes place, while the DNP-azide side remains dark. The microchannel cantilever used for spotting is visible out of focus in (a) and (d). All scale bars equal 50 µm.


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Figure 4. Fluorescence microscopy images of a graphene flake modified by DPN with biotinazide pattern. The position is marked with red circle. (a) The pattern is not visible after writing and washing of the flake but (b) lights up in fluorescence after incubation with Cy3 labelled streptavidin. The small squares in the grid pattern are of 2µm edge length, line widths range from approximately 500 to 800nm. Scale bars equal 20µm.


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Attoliter Chemistry for Nanoscale Functionalization of Graphene - ACS

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