INSTITUTE OF PHYSICS PUBLISHING NANOTECHNOLOGY

Nanotechnology 16 (2005) 1–8 doi:10.1088/0957-4484/16/0/000

Self-assembly of functionalized sphericalnanoparticles on chemically patternedmicrostructuresChristoph Huwiler1, Martin Halter1, Kurosch Rezwan2,Didier Falconnet1, Marcus Textor1 and Janos Voros1

1 BioInterfaceGroup, Laboratory for Surface Science and Technology, Department ofMaterials, Swiss Federal Institute of Technology (ETH) Zurich, CH-8093 Zurich, Switzerland2 Institute of Nonmetallic Inorganic Materials, Department of Materials, Swiss FederalInstitute of Technology (ETH) Zurich, CH-8093 Zurich, Switzerland

E-mail: janos.voeroes@mat.ethz.ch

Received 7 September 2005, in final form 7 October 2005PublishedOnline at stacks.iop.org/Nano/16/1

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AbstractThe production of hierarchical nanopatterns (using a top-downmicrofabrication approach combined with a subsequent bottom-upself-assembly process) will be an important tool in many research areas.We report the fabrication of silica nanoparticle arrays on lithographicallypre-patterned substrates suitable for applications in the field ofnanobiotechnology. Two different approaches to reach this goal arepresented and discussed: in the first approach, we use capillary forces toself-assemble silica nanoparticles on a wettability contrast pattern bycontrolled drying and evaporation. This allows the efficient patterning of avariety of nanoparticle systems and—under certain conditions—leads to theformation of novel branched structures of colloidal lines, that might help toelucidate the formation process of these nanoparticle arrays. The secondapproach uses a recently developed chemical patterning method that allowsfor the selective immobilization of functionalized sub-100 nm particles atdistinct locations on the surface. In addition, it is shown how thesenanocolloidal micro-arrays offer the potential to increase the sensitivity ofexisting biosensing devices. The well-defined surface chemistry (of particleand substrate) and the increased surface area at the microspots, where thenanoparticles self-assemble, make this patterning method an interestingcandidate for micro-array biosensing.

1. Introduction

Functional materials with topographically or chemicallycomplex (e.g. hierarchical) surfaces in the nanometre rangeare currently being developed in view of their promisingnew or improved optical or electronic properties that findpotential applications in biosensing or (opto-)electronicdevices. The still-emerging field of nanotechnology is inneed of reliable, fast (and therefore parallel), economicaland versatile methods for structuring surfaces with controlledsurface chemistry in the nanometre range. Nature hasbeen performing these tasks by self-assembling specific

biomolecules into complex supramolecular structures withgreat accuracy and high efficiency. This concept ofself-assembly can be adapted for the nanostructuring ofsurfaces. Nowadays, molecular self-assembly techniques(self-assembled monolayers (SAMs)) [1–3] are a well-known way to chemically functionalize surfaces. Complexmesoscale and millimetre-sized 3D objects [4–6] can be self-assembled into useful structures, and for a number of yearssubstantial efforts have been put into self-assembling colloidalparticles into 3D crystals [7–10], monolayers [11] or onpatterned substrates [12–20]. Spherical colloids have attractedconsiderable interest in such self-assembly experiments due

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to numerous advantageous properties (well-defined shape andsize distribution, controllable size and surface properties,variety of materials available) [20].

In colloidal self-assembly processes on patternedsubstrates, some general issues have to be addressed. Asa first step, the adsorption of the colloidal particles ontothe substrate pattern predetermines the final outcome ofthe colloidal nanostructures to a large extent. Relying onchemical binding to guide colloidal self-assembly, we need toprecisely structure the surface and control the properties of thenanocolloids in the suspension in order to achieve reproduciblenanocolloid patterns in the initial wet state. Of equal (or evengreater) importance, however, is the drying process followingthe adsorption step. If colloidal particles self-assemble onfunctionalized microstructures, drying of the suspension is aninherent part of the fabrication process. The colloid arraysmay be significantly altered by capillary forces during dryingif these exceed the colloid–substrate adhesion forces. For thisreason, capillary forces are a major concern in self-assemblyprocesses of colloidal particles [11, 21, 22]. However, theycan also be used intentionally to guide the self-assemblyprocess provided a suitable template is used (this template ismost commonly made up of relief structures patterned in thesubstrate) [20, 23–25].

In this work, we report the hierarchical fabrication ofcolloidal nanoarrays by two different approaches. Bothmethods rely on the photolithographic production of amicropattern as a first step, which is in a second step convertedinto a chemically patterned surface. These functionalizedmicropatterns act as the major structuring and guiding elementfor the assembly of sub-100 nm spherical silica colloids.

2. Experimental details

2.1. Selective molecular assembly patterning technique

In the selective molecular assembly patterning process(SMAP) [26], a photoresist film is spin-coated onto twotransparent sputter-coated metal oxide layers (a 100 nm SiO2

layer followed by a 12 nm TiO2 layer on top) (step 1 infigure 1). The desired geometrical features are transferredinto the photoresist using standard photolithography. Thispattern is then locally etched through the TiO2 by reactiveion etching (RIE) (step 2 in figure 1). Upon removal ofthe photoresist (and cleaning of the sample by a UV andO2 plasma treatment), the SiO2/TiO2 metal oxide pattern isthen immersed in an aqueous solution of 0.5 mM ammoniumdodecyl phosphate (CH3(CH2)11PO4(NH4)2) (DDPO4) [27]for 48 h and rinsed with high-purity water. This producesa well-defined hydrophobic self-assembled alkane phosphatemonolayer (SAM) specifically on the TiO2 patches whileleaving the SiO2 areas entirely uncovered. By this process,a wettability contrast pattern is achieved with the hydrophobicSAM covering the TiO2 regions (advancing contact angle= 110◦) and the hydrophilic silica areas (advancing contactangle <6◦).Q.1

2.2. Molecular assembly patterning by lift-off process

The recently developed molecular assembly patterning by lift-off (MAPL) [28] technique (figure 1(b)) is able to convert a

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Figure 1. Schematic illustration of the two patterning processesused in this work. (a) Selective molecular assembly patterningprocess (SMAP): (1) metal oxide pattern produced by standardphotolithography; (2) selective chemical modification of TiO2

surface area by an alkane phosphate SAM; (3) the hydrophobicalkane phosphate SAM is de-wetted during controlled drying;(4) capillary forces assemble the silica nanoparticles on thewettability contrast pattern. (b) Molecular assembly patterning bylift-off process (MAPL): (1) photolithographically producedphotoresist pattern coated with PLL-g-PEG-biotin polymer;(2) photoresist lift-off; (3) backfilling of open areas withPLL-g-PEG; (4) adsorption of PLL-g-PEG-biotin-coated silicacolloids using streptavidin as a linking molecule; (5) drying of thesample yields colloid arrays with functionalized nanoparticles(polymer brush will collapse). Note: sketches are not drawn toscale. For more information on the patterning processes see [26, 28].

(This figure is in colour only in the electronic version)

photolithographically prestructured photoresist film (Shipley1818) into a micropattern of biointeractive and non-interactiveregions. Negatively charged niobium oxide or titaniumoxide surfaces are used as substrates for this process. Ina first step, a biotinylated polyelectrolyte copolymer graftedwith PEG chains (cationic poly(L-lysine)-graft-poly(ethyleneglycol) (referred to as PLL-g-PEG-biotin)) is adsorbed bya simple dip and rinse step in an aqueous solution on thephotoresist patterned substrate (step 1 in figure 1(b)). Atneutral pH, the positively charged amino-terminated backboneof the PLL-g-PEG-biotin molecule adsorbs electrostatically tonegatively charged metal oxide surfaces, such as niobia, andto the photoresist as well. After lift-off of the photoresist,PLL-g-PEG-biotin and bare substrate regions are exposed(step 2 in figure 1(b)). Backfilling of the Nb2O5 substratewith non-biotinylated PLL-g-PEG is achieved by subsequentimmersion of the sample in a PLL-g-PEG solution. Allpolymer adsorption steps were performed at concentrationsof 0.1 mg ml−1 in 160 mM HEPES buffer. As a result ofthe MAPL process, a micropattern of biotinylated areas (PLL-g-PEG-biotin) in a non-adsorbing background (PLL-g-PEG)is achieved, for which the non-specific interactions of serumproteins is below 2 ng cm−2 (step 3 in figure 1(b)) [28].A PLL-g-PEG version with a molecular weight of the PLLbackbone of 20 kD, a PEG chain length of 2 kD molecularweight and a grafting ratio of 3.5 (e.g. a PEG chain is graftedto every 3.5 PLL monomer unit) was used in this work.The synthesis, structure and properties of the PLL-g-PEGcopolymer are described in more detail elsewhere [26, 29].The adsorbed amount of PLL-g-PEG on a silica surface was

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determined from optical waveguide light-mode spectroscopy(OWLS) measurements to be 150 ± 9 ng cm−2.

2.3. Colloidal particles

Colloidal silica suspensions (Klebosol, Clariant, France) oftwo different particle sizes were used. Particle sizes andsize distributions were measured by means of x-ray disccentrifuge (X-ray sedigraph, Brookhaven Instruments) andSEM image analysis (of at least 300 particles) (LEO 1530,Zeiss, Germany) and determined to be 41 ± 5 and 73 ±6 nm, respectively. Colloidal suspensions were received asaqueous suspensions and if not indicated otherwise bufferedto pH = 7.4 with HEPES (4-(2-hydroxyethyl) piperazine-1-ethanesulfonic acid) (Fluka Chemie, Switzerland) and6 M NaOH solution. Two different salt concentrations ofHEPES buffer were used: 10 mM HEPES buffer (HEPES 1)and 10 mM HEPES buffer with 150 mM NaCl (HEPES 2).

2.3.1. Coating of colloidal particles. Coating of colloidalsilica particles was achieved by mixing equal amounts ofcolloid suspension with the polymer solution, both of themthus containing twice the amount of colloidal particles orpolymer solution as required in the final suspension. Anaqueous solution of PLL-g-PEG-biotin (buffered at pH 7.4with HEPES) was used to coat the colloidal particles. PLL-g-PEG-biotin was added in a two-fold excess and the coatedsilica suspension was centrifuged and redispersed three timesto remove any free polymer molecules in solution. The specificsurface area of the 73 nm particles was 51 m2 g−1; thus 5 mlof a 1 wt% suspension was coated with 9.2 mg of polymer (atwo-fold excess assuming an adsorption density of the polymerof 150 ng cm−2 (as determined by OWLS)).

2.4. Optical waveguide light-mode spectroscopy (OWLS)technique

To determine the amount of adsorbed chemical species orcolloidal particles on a given substrate (see figures 4 and 6),optical waveguide lightmode spectroscopy (OWLS) was used.The OWLS technique is based on coupling a He–Ne laser intoa high-refractive-index waveguide via an optical diffractiongrating. The angle at which total internal reflection of thelaser occurs is measured. This incoupling angle depends onthe refractive index of the medium present at the surface ofthe waveguide. Therefore, molecules or particles adsorbing tothe waveguide surface induce a change in the incoupling angle,which can be detected and converted into adsorbed mass. Sinceonly species present in the evanescent field (extending a fewhundred nanometres into the fluid) have an influence on theincoupling angle, this method is very suitable for measuringadsorption processes in situ and with high sensitivity.

2.5. Colloid assembly experiments

Colloid adsorption experiments on 1 cm2 samples patternedwith the SMAP technique were carried out under controlledconditions to study the parameters that govern the colloidalself-assembly processes. Particles were either adsorbed bydrying a 50 µl drop of HEPES 1 buffered colloid suspensionof 0.1 wt% particle concentration under ambient conditions

(10 mM ionic strength and pH = 7.4) (drop drying) or byimmersing the sample into the suspension for 30 min, removingit from the suspension with a controlled speed, rinsing itwith Millipore water and blowing it dry under nitrogen flow(dip coating). Drop-dried samples were not rinsed beforescanning electron microscopy (SEM) investigations. All high-resolution SEM investigations were performed on samplessputter-coated with a Pt film of approximately 4 nm thickness.

Self-assembly experiments on MAPL samples wereperformed by first immersing a MAPL sample of 1 cm × 1 cm(exhibiting a PLL-g-PEG/PLL-g-PEG-biotin contrast) intoa streptavidin solution (25 µg ml−1 in HEPES 1) (Sigma-Aldrich, US) for 30 min. After rinsing and drying of thesample, it was immersed into a HEPES 1 suspension of PLL-g-PEG-biotin coated colloidal particles for 60 min. Sampleswere flooded with extensive amounts of millipore water toremove all non-adsorbed colloidal particles. They were thenrinsed and dried under nitrogen flow. In the case of serumadsorption experiments on colloid monolayers (figure 6), thesubstrate (SiO2 coating on an OWLS waveguide) was coatedwith a thin layer of poly(ethylene imine) (PEI). This coatingfirst renders the SiO2 surface positive at neutral pH and allowsthe rapid electrostatic adsorption of negatively charged silicacolloids. After adsorption of the colloid layer, another layer ofPEI was adsorbed to render the colloid sub-monolayer positiveagain so that serum adsorption on both samples (bare PEI-coated surface and colloid monolayer coated with PEI) couldbe compared.

3. Results and discussions

3.1. Nanocolloidal self-assembly by capillary forces

Our first patterning approach uses a hydrophobicity contrastcreated by the selective molecular assembly patterning process(SMAP) [26] depicted in figure 1(a). By this process, awettability contrast pattern is achieved with the hydrophobicSAM covering the TiO2 regions (advancing contact angle= 110◦) and the hydrophilic silica areas (advancing contactangle <6◦). This hydrophobicity contrast serves as a templatefor the self-assembly of nanocolloids by capillary forces incontrolled drying and dip-coating experiments (steps 3 and 4in figure 1(a)).

Large-scale (cm2) homogeneous colloidal patterns wereproduced by a dip-coating process using substrates with ahydrophobicity contrast pattern (see figure 2). A 1 cm2

sample patterned with the SMAP technique was immersedinto a 2 wt% suspension of 41 ± 5 nm silica colloids inHEPES 1 buffer for 60 min, removed from the suspensionat a speed of 2.5 µm s−1, rinsed with Millipore water anddried under nitrogen flow. The large difference in contactangle between the background and the pattern leads to de-wetting of the hydrophobic background during the removal ofthe sample from the suspension. However, if the suspensionwas diluted with large proportions of water before removalof the immersed chip, no colloidal particles were found onthe surface. This means that the colloidal particles did notinteract strongly with either the hydrophobicor the hydrophilic(silica) areas of the SMAP surface in suspension. We concludethat, in this case, capillary forces dominate the self-assemblyprocess: while the hydrophobic background is de-wetted just

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Figure 2. SEM micrographs of 41 nm silica colloidal particlesassembled on a hydrophobic–hydrophilic contrast produced bySMAP (see text). Dip-coating of the sample in a 2 wt% suspensionwas performed at a constant speed of 2.5 µm s−1. The backgroundconsists of TiO2 modified with a hydrophobic alkane phosphateself-assembled monolayer and is particle free due to thehydrophobic character of this surface and the de-wetting thattherefore occurs during drying. Well-ordered silica colloid arrayscan be obtained with this method by choosing suitable particleconcentration and dip-coating conditions (a). (b) Edge region of asingle square showing dense packing of the silica nanoparticles andgood confinement of the pattern. (c) The centre region of a squarerevealing depletion effects originating from the drying process.

outside the drying front, the hydrophilic squares are still filledwith suspension. Evaporation of the solvent on each of thesehydrophilic squares leads to the colloidal arrays observed infigure 2. The droplet left on such a square outside the dryingfront is pinned at the three-phase contact line around the edgeof the square, where therefore the highest rate of evaporationis observed. This results in a liquid flux dragging the particlestowards the three-phase contact line [30]. For this reason,edge regions of a hydrophilic square typically have very densemono- or multilayers while the centre region is depleted ofcolloidal particles (see insets of figure 2) or consists of a lowernumber of layers compared to the edge regions. We found thatdepletion effects could not be completely inhibited on suchhydrophobicity contrast patterns. While the number of colloidlayers formed on each individual hydrophilic square dependedmainly on the particle concentration and dip-coating speed andcould be controlled rather reproducibly (results not shown), thedepletion effects observed in the middle of a square were moredifficult to control systematically.

By performing drop-drying rather then dip-coatingexperiments (of a 50 µl drop of HEPES buffered colloidsuspension of 0.2 wt% particle concentration under ambientconditions), we observed the formation of typical colloidalcrystals in the edge of the drop that are often referred to as‘coffee’ rings [25]. However, we were also able to observethe formation of branch-like structures presented in figure 3.

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Figure 3. SEM micrographs of 73 nm silica colloidal particles on ahydrophobicity contrast pattern produced by SMAP (see text). Asmall 50 µl drop of a 0.2 wt% suspension was deposited and dried.The hydrophobic background SAM is particle free apart from theobserved colloid structures due to the de-wetting that occurs upondrying. Colloidal particles completely fill the hydrophilic SiO2 areas(60 µm × 60 µm squares). After filling of the pattern withnanoparticles during drying, remaining silica colloids line up andform branch-like structures on the hydrophobic background. Thethickness and range of these structures may show substantialvariations from area to area on a sample; the branched structure,however, is always observed. (b) and (c): insets revealing thedetailed structure of these drying structures.

These structures are only present in the central region of a drieddrop where the hydrophilic silica squares are completely filledwith a monolayer of the 73 nm silica particles (figure 3(a)) andthe branched structures of colloidal particles are observed onthe hydrophobic background. These branches consisting ofseveral layers of particles (inset of figure 3) have a fractal-type structure always originating at the edge of the silicasquares. While their width is typically in the range of 100 nmto 10 µm, their length varies between a few microns andseveral hundreds of microns depending on the experimentalconditions. At present, we have no clear understanding for thedescribed phenomenon. It seems plausible that these branchesare formed during the last stage of the drying process when thelarge drop of water breaks up and de-wetting of the backgroundcompetes with drying of the minute amount of liquid stillpresent at the surface. However, the presence of the branchesin these structures suggests that the branches grow out of thesilica squares. This might indicate that localized coagulation,related to the increased salt concentration upon drying, mightbe responsible for this effect.

Since in the SMAP system capillary forces are dominantover colloid–surface interactions, it is not surprising that we

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have not observed a significant influence of a variety ofparameters. Changing pH (3, 7.4 and 9.9) or ionic strength(10, 160 mM) did not alter the characteristics of the resultingcolloid nanoarrays. The influence of these parameters wasalso not observable by in situ OWLS measurements of theadsorption process without drying, i.e. in contact with thesuspension no interaction between the silica colloids and eitherthe hydrophobic alkane-phosphate SAM or the silica substratewas measured. This was also expected since ζ -potentialmeasurements of the silica colloids showed an isoelectric point(IEP) of around pH 2.3 (which is in agreement with literaturevalues) and the colloidal particles are therefore negativelycharged at all used pH values as is the silica substrate (figure 4).

3.2. Selective binding chemistry to form nanoparticle arrays

While in the case discussed above the nanocolloid self-assembly was driven predominantly by guided capillary forces,the colloidal particles are functionalized in the MAPL processto directly interact with the micropattern present on the surface.This leads to a reduced influence of capillary forces in thisapproach. Linking of the particles to the surface is achievedby binding biotinylated colloidal particles to biotinylatedsurface patches (produced by the MAPL [28] process) via astreptavidin molecule. The background of these pattern hasto be rendered non-interactive in order to avoid unspecificadsorption of colloidal particles to the background.

In a first step, the colloidal nanoparticles were coatedwith biotinylated PLL-g-PEG copolymer. This was achievedby mixing equal amounts of colloid suspension with a PLL-g-PEG-biotin solution (buffered at pH 7.4 with HEPES)as described in the experimental section. ζ -potentialmeasurements were conducted to monitor the adsorption ofthe copolymer to the colloid surface (figure 4). The isoelectricpoint (IEP) of the 73 nm silica colloids was determined to be atpH 2.3. Mixing of the colloidal suspension with the copolymersolution shifted the IEP to pH 6.5, indicating that the surfaceof the colloidal particles indeed became coated with a PLL-g-PEG-biotin film. The positively charged polyelectrolytecopolymer adsorbed to the negatively charged surface exposingthe uncharged, hydrophilic PEG chains. It is the exposedwater-like PEG chains that are believed to be responsible forthe change in the IEP to values close to pH 7. On the otherhand, coating the silica colloids with the positively chargedPLL (without the attached PEG chains) rendered the silicasurface positive (+35 mV at pH 7) and shifted the IEP to10.3. The PEG chains in the PLL-g-PEG molecule havean additional important function: they sterically stabilize thecolloid suspension preventing them from coagulating even atpHs close to the IEP.

OWLS measurements on homogeneous substrates furtherconfirmed that the coating process of the colloidal particleswas successful, as shown in figure 4. Uncoated silica particlesreadily adsorbed on a PLL-g-PEG coated surface withinminutes (depending on the concentration) and formed a stableand dense layer as observed by SEM and OWLS (figure 4(a)).If the colloidal particles were coated with a PLL-g-PEG-biotinlayer, the adsorption behaviour changed completely and thecoated particles no longer adsorbed to a PLL-g-PEG-coatedsurface (figure 4(b)). Thus, by coating the colloidal silica

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Figure 4. (a) ζ -potential measurements of uncoated 73 nm silicacolloids (��), the same colloids coated with the copolymerPLL-g-PEG-biotin (�) and with the polymer PLL (◦). The IEP ofthe silica colloids (2.3) shifts to 6.5 after coating withPLL-g-PEG-biotin and to 10.5 for the PLL-coated silica colloids,indicating that the colloid surface is indeed coated with thecopolymers and the IEP of the coated colloids shifts to thecorresponding pK value for the polymer coating. OWLSmeasurements (b) and SEM images ((c), (d)) illustrating theadsorption behaviour of PLL-g-PEG-biotin coated and uncoatedsilica colloids. Uncoated silica colloids show rapid adsorption on aPLL-g-PEG(-biotin) coated substrate as shown in the OWLSexperiment (b) and SEM image (c). However, if both particle andsubstrate are coated with a PLL-g-PEG(-biotin) copolymer layer,adsorption of colloidal particles is suppressed completely (d).

particles with a layer of PLL-g-PEG-biotin we can producenanoparticles that are not interacting with a PLL-g-PEG coatedsurface. Note that the addition of the biotin linker does notchange the adsorption properties significantly.

A PLL-g-PEG layer of the architecture used in this studyon a silica substrate extends about 8–10 nm [31] from thesurface forming a strongly hydrated and dense PEG brush. Ifparticle and surface are coated with a PLL-g-PEG adlayer,the interaction between particle and surface is efficientlyreduced. The reason why uncoated silica nanocolloids adsorbstrongly to a PLL-g-PEG coated surface is less obvious.

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Theoretical calculations show that the Debye-length—thelength at which the electric double layer is effective—is∼3 nm for a 10 mM electrolyte solution and <1 nm fora 160 mM solution. Therefore, the negative charges fromthe metal oxide surface should be completely shielded bythe adsorbed 8–10 nm thick PLL-g-PEG layer and indeeduncoated SiO2 microparticles did not interact with PLL-g-PEGcoated surfaces in colloidal probe AFM experiments [32]. Onthe other hand, according to van der Beek et al [33], protonatedsilica surfaces (and nanocolloids) may form hydrogen bondsbetween the protonated oxygen atoms of the silica surfaceand the oxygen atoms of the PEG layer and thus silicananocolloids can be immobilized on the surface. This has alsothe consequence that with increasing pH more and more Si–OH groups get deprotonated and fewer colloids will adsorbat higher pH values due to reduced hydrogen bonding assuggested by Gage et al [34]. Furthermore, this model alsoexplains why PLL-g-PEG coated surfaces with lower graftingratios (e.g. more EG-groups are present per surface area)exhibit increased colloid adsorption (results not shown).

Colloidal self-assembly experiments on MAPL sampleswere performed by first immersing a MAPL sample of 1 cm ×1 cm (exhibiting a PLL-g-PEG/PLL-g-PEG-biotin contrast)into a 25 µg ml−1 streptavidin HEPES solution for 30 min.After rinsing and drying, the sample was immersed into asuspension of PLL-g-PEG-biotin coated colloidal particles.Samples were flooded with water before removal, then rinsedwith Millipore water and dried under nitrogen flow. In figure 5,a PLL-g-PEG/PLL-g-PEG-biotin contrast of 30 µm squareswas produced and the PLL-g-PEG-biotin coated colloidalparticles were bound to these squares via the streptavidin linker.Since no unspecific adsorption occurs on the background(figures 4(a) and 5), the nanocolloid adsorption is restrictedto biotinylated areas of the substrate and formation of amonolayer is observed (insets in figure 5(b)). Regions inthe centre of the samples, however, typically exhibit dryingartefacts such as areas depleted of colloids (figure 5(d)). Theseartefacts originate from the capillary forces that act upon dryingand which are not negligible even if the colloids are boundto the surface by biotin–streptavidin interactions. However,we did not observe extensive multilayer formation on MAPLsamples as was the case for the SMAP samples if particleconcentration was increased and dip-coating speed reduced.This is attributed to the fact that on MAPL chips, there isno distinct hydrophobicity contrast pattern since there is nosignificant contact angle difference between a PLL-g-PEG-biotin coated and a PLL-g-PEG coated surface and thereforedifferent processes are responsible for the nanocolloid arrayformation in the two patterning approaches. However, in thelast stage of the drying process, when liquid bridges are stillpresent between individual particles and a capillary force isexerted, the particle arrays might still be deformed resulting inthe observed drying artefacts (figure 5(d)). If the particles arespaced far apart, no liquid bridge is formed during drying andcapillary forces will not influence the resulting pattern.

3.3. Using nanoparticle arrays to improve biosensing devices

Both patterning techniques presented here allow the productionof monolayers of colloidal particles at distinct places. These

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Figure 5. SEM micrographs showing colloidal nanoarrays obtainedby adsorption of PLL-g-PEG-biotin coated 73 nm silica colloids ona MAPL chip. On this chip, 30 µm squares are coated withPLL-g-PEG-biotin while the PLL-g-PEG coating on thebackground effectively resists particle adsorption. Streptavidin isadsorbed to the biotinylated regions of the substrate andsubsequently binds the biotinylated silica colloids. The achievedpatterns show good fidelity over a large area (b). Edge regions of thepattern are sharply confined (c). Depletion effects in the centreregions of an individual square are drying artefacts that are oftenseen in these colloidal nanoarrays (d).

colloidal arrays can be used as substrates for micro-array-type biosensing applications such as evanescent-field-excitedfluorescence readout providing an enhanced surface area forthe immobilization of biomolecules. The production ofcolloidal nanoarrays with a distinct chemistry, as obtainedby the MAPL technique, is of special interest for suchapplications. In the OWLS measurements presented infigure 6, serum adsorption (10% serum) was monitored on abare substrate and on a substrate coated with a sub-monolayerof colloidal particles, respectively. The substrate in thisexperiment (SiO2 layer on an OWLS waveguide) was coatedwith a thin PEI film (a positively charged polyelectrolyte) withan ellipsometric thickness of 1 nm. This coating renderedthe SiO2 surface positive at neutral pH and allowed the rapidelectrostatic adsorption of negatively charged silica colloids.After adsorption of the colloid layer, another layer of PEI wasadsorbed to render also the particles positive so that serumadsorption on both samples could be compared directly. Atwo-fold increase in serum-adsorption from 550 ng cm−2 tomore than 1000 ng cm−2 on the colloid coated surface wasobserved. This observed increase in serum adsorption on thecolloid-containing surface can be attributed to the increasein available surface area. This might offer the possibility to

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(b)

Figure 6. (a) OWLS measurements showing the influence of a colloid sub-monolayer on serum-adsorption on a substrate. More than500 ng cm−2 of a 10% serum solution adsorbs on a PEI-coated SiO2 surface after 40 min (solid line). The adsorption of a colloidsub-monolayer of 73 nm colloids leads to a two-fold increase in serum adsorption due to the increased surface area present at the surface(two set of experiments shown, dashed lines). In this case, the silica colloidal particles were adsorbed on a PEI-coated surface byelectrostatic interactions and the colloids were subsequently coated with a PEI layer as well to ensure comparable adsorption conditions.((b), (c)) SEM images representing the uncoated surface (b) and a surface with a sub-monolayer of 73 nm silica particles (c).

considerably increase the sensitivity of existing biosensingdevices that rely on the detection of small quantities ofanalyte. For biosensing applications, however, it is essentialto have precise control over the surface chemistry to preventunspecific adsorption on the background and have specificinteractions on the individual spots with for example selectedproteins or antibodies. Colloidal nanoarrays developed forbiosensing applications have to exhibit such non-foulingproperties and functionalization possibilities. The use ofpotentially non-fouling colloids due to a PLL-g-PEG coatingand the possibility of grafting diverse functionalities to thesecolloids [28] makes the approach presented here interestingfor such biosensing applications. However, a number ofunsolved problems have to be addressed beforehand. Theproblems range from economical issues (coating nanocolloidswith surface areas between 50–100 m2 g−1 with expensivechemicals), reproducibility and up-scaling difficulties tophysical limitations (kinetic and diffusion problems in such(multi-)colloid layer systems).

4. Conclusions

In summary, we presented and compared two different ap-proaches to produce colloidal nanoarrays on chemical mi-cropatterns combining a top-down method (photolithography)with a bottom-up colloidal self-assembly process. Both ap-proaches allow the efficient patterning of large areas (cm2)without geometrical constraints and have the potential to bedownscaled to the nanometre range (e.g. using nanoimprintlithography or x-ray interference lithography) [35].

The patterning approach based on the SMAP techniqueuses capillary forces to assemble nanoparticles on ahydrophilic/hydrophobic contrast pattern. While thehydrophilic patterns stay filled with colloid suspension, thehydrophobic areas de-wetted during drying, resulting inthe observed nanoparticle structures after drying. Undercertain conditions, the wettability contrast pattern induced theformation of novel fractal-like branched nanostructures with alength ranging from several microns up to millimetres.

The alternative approach is based on the MAPL processfor the production of nanoparticle micro-arrays with acomplete control over the surface chemistry of particles andsurface pattern. Furthermore, the coated nanoparticles arechemically bound to the surface which allows control of theappearance of the nanopatterns (density, number of layers).In contrast to the arrays formed by capillary forces, theMAPL nanopatterns have further benefits, for example in thearea of nanobiotechnology applications. There, non-foulingproperties (of the whole chip as well as the colloidal particlesassembled on it) and high selectivity are two key issues.

These nanoparticle arrays have specific surface functional-ization possibilities and non-fouling properties combined withthe necessary pattern fidelity. Colloidal arrays based on theMAPL technique, therefore, are highly attractive means to in-crease the sensitivity of existing micro-array biosensors. Wehave shown the feasibility of this concept to enhance the load-ing capacity of an optical waveguide technique.

In conclusion, both presented methods provide a versatileand flexible approach to produce geometrically definedpatterns of colloidal particles for applications in the areas ofbiotechnology, catalysis or for optical materials where large-scale nanoparticle micro-arrays with good homogeneity areneeded.

Acknowledgments

The authors thank Michael Horisberger for the metal oxidecoatings (PSI, Switzerland), Brandon Burgler for SEMimaging (ETH Zurich) and Stephanie Pasche for providing thepolymers. This work was financially supported by the SwissPriority Program on Nanotechnology, Top Nano 21 (Project5971.2), CTI (Project 7241.1),EPF Lausanne and ETH Zurich.

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