Nanostructured disposable chips for electrochemiluminescence-based biosensing Ming Zhou∗a, Alexis Laforgueb, C. Geraldine Bazuin*b and Robert E. Prud'hommeb a Institute for Microstructural Sciences National Research Council Canada, Ottawa, ON, Canada, K1A 0R6 b Département de chimie, Université de Montréal C. P. 6128, succursale Centre-ville, Montréal, QC, Canada, H3C 3J7

ABSTRACT The electrochemiluminescence (ECL) of Ru(bpy)32+ (bpy = 2,2’-bipyridine and its derivatives) complexes has attracted interest from chem- and bio-analytical researchers. Particularly, by labeling bio-molecules with Ru(bpy)32+ derivatives, highly competitive ECL immunoassay and DNA probing have been employed in clinical and research laboratories and are now becoming standard methods. In the well-established commercial systems designed for bench-top applications, paramagnetic microbeads are used for capturing the analytes and separating the excess of labeled biomolecules from the flow cell. The large surface area of the beads provides a high capacity and efficiency for analyte capturing. However, the use of microbeads prevents the instrument from being miniaturized. Furthermore, only a tiny portion of species is enabled to generate luminescence because of the inaccessibility of the majority of the labels to the electrode surface. We propose to develop a handheld device with disposable chips based on the ECL signal modality. Central to this instrumentation is the fabrication of a nanostructured electrode with spatially selective bioimmobilization. The electrode surface is structured to reach the maximum capturing ability and, at the same time, maintain the effective electroactive region and the accessibility for the ruthenium label to be excited electrochemically. In this presentation, we present a manufacturable approach to the fabrication of such disposable nanostructured electrodes for ECL-based handheld devices. Key words: electrochemiluminescence, biosensor, nanostructure, bioimmobilization

1. INTRODUCTION The identification and quantification of chemical, biochemical and biological substances are often carried out by binding the substances of interest with labeled recognition partners through an affinitive interaction. The labels are molecules that generate detectable and highly characteristic signals. Enzymes, radioactive elements, fluorescent moieties, redox active moieties, chemiluminescent and electrochemiluminescent moieties are currently used as such signal generating moieties in clinical diagnostics, drug screening, genomics/proteomics and many other application areas. The performance of these immunoassays depends, often largely, on the signal detection techniques. Electrochemiluminescence (ECL)1-2 of ruthenium(II) tris(bipyridine) complexes (Ru(bpy)32+ , bpy = 2,2’-bipyridine or its derivative) has been developed into and proved to be a highly sensitive bioanalytical technology.1-11 There are several clinical diagnostic systems developed for the immunoassay and nucleic acid probe markets. The primary electrochemical and chemical reactions that lead to the luminescence are as follows: Ru(bpy)32+ - e- → Ru(bpy)33+

E0 = 1.06 V vs. Ag/AgCl

(1)

-

E0 = -1.48 V vs. Ag/AgCl

(2)

2+

+

Ru(bpy)3 + e → Ru(bpy)3 ∗

[email protected]; phone 613 993 2524; fax 613 993 0755. [email protected]; phone 514 340 5176; fax 514 340 5290. Nanosensing: Materials and Devices II, edited by M. Saif Islam, Achyut K. Dutta, Proc. of SPIE Vol. 6008, 60081A, (2005) · 0277-786X/05/$15 · doi: 10.1117/12.629735

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Ru(bpy)33+ + e- → Ru(bpy)32+* -

+

2+

Ru(bpy)3 - e → Ru(bpy)3 *

E0 = -1.06 V vs. Ag/AgCl

(3)

E0 = 0.64 V vs. Ag/AgCl

(4)

Ru(bpy)3 * → Ru(bpy)3 + hυ (620 nm) 2+

2+

(5)

The strong orange luminescence is generated from the decay of a triplet MLCT (Metal-to-Ligand Charge Transfer) state, i.e., Ru(bpy)32+*, to the ground state. According to the basic reactions (1)-(5), three different approaches have been used to produce the excited state Ru(bpy)32+*, namely, oxidative-reduction, reductive-oxidation and annihilation.1-2 The oxidative-reduction ECL is currently employed in commercial systems with tripropylamine3 as a coreactant. The proposed formation mechanism3 of the excited state is based on the study of anodic oxidation of aliphatic amines (Eqs. 6-8).12 N(C3H7)3 - e- → N(C3H7)3•+ or

(6)

3+

•+

2+

Ru(bpy)3 + N(C3H7)3 → Ru(bpy)3 + N(C3H7)3

(7)

N(C3H7)3•+ - H+ → CH3CH2C•HN(C3H7)2 •

3+

(8) 2+

CH3CH2C HN(C3H7)2 + Ru(bpy)3 → Ru(bpy)3 * + ?

(9)

The oxidation of tripropylamine on the electrode produces N(C3H7)3•+, which quickly deprotonates to form a free radical CH3CH2C•HN(C3H7)2 with strong reducing capability.3,12 The Ru(bpy)33+, simultaneously generated on the electrode, is then reduced by the deprotonated free radical to its luminescent excited state, Ru(bpy)32+*.

Incredibly Low Efficiency !!!

< 10 nm

Electrode

> 1000 nm

PMT

Electrical magnet

Figure 1: Schematic illustration of the ECL generation on magnetic microbeads with the analyte and the label in a sandwich format.

Figure 2: A small portion of a microbead in the electrochemical reaction region.

For bioanalyses, the luminophore Ru(bpy)32+ was made bioconjugatable,4,13,14 by introducing functional groups to one of the bipyridine rings, in order to label the biomolecules that recognize the analytes. The core of the ECL-based bioanalytical systems includes a platinum electrode in a flow cell and a photomultiplier tube (PMT) placed above the working electrode for light detection.7 The magnetic microbead capturing and separation techniques are used as a means to bring the analyte to the electrode surface and to separate the unbound reagents from the sample solution (see Fig. 1). Automated sample handling and fluidics delivery subsystems round out the analysis systems. Since the ECL signal is generated by a well-controlled low-voltage electrical stimulus, it is possible to reduce the system dimension even to a handheld size for medical settings where no large-volume and automated analysis is needed. However, for the following reasons, the current instrumentation doesn’t fit into the design concept for a miniaturized system demanded by the pointof-care diagnosis and other bioanalytical tasks.

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First, the flow cell and magnetic bead techniques, although commonly used, need a fluid handling system, including reagent reservoirs, pumps, valves and an electronic control unit. These contribute to a large part of the instrument size. Second, the platinum electrode mounted in the flow cell has to be regenerated after each measurement to allow multiple sequential analyses. An automated electrode regeneration procedure and additional reagents are needed. Third, the application of magnetic beads capturing/separation technique largely prevents the ECL technology from achieving higher detection sensitivity, because only a very small portion of the bead surface, which is in contact with the electrode, can be activated and thus generate luminescence (see Fig. 2). Furthermore, the luminescence could be attenuated by the opaque beads before it reaches the photomultiplier tube. It is understandable that the sensitivity will be generally reduced in a miniaturized system if the same technological principles are used. In order to miniaturize the current ECL instrumentation and adapt it to a handheld device for on-site test, we proposed a bead-free, disposable chip-based ECL device.

2. PROPOSED MINIATURIZATION Central to our proposed hand-held device is the elimination of magnetic microbeads and the related delivery system. Eliminating the beads from the system will require the electrochemical electrode to be a carrier for the bioactive species. However, if the species is directly captured on the whole electrode surface, the electrode will be passivated by surface modification and by immobilization of the bio-recognition elements. Our solution to this problem is to coat the electrode partially with the bio-recognition elements and leave the other part of the electrode electrochemically active. The electrode surface should be structured in a way that allows the maximum capturing ability and, at the same time, maintain the effective electroactive surface and the accessibility for the Ru-label to the electroactive region. An example is conceptually illustrated in Fig. 3. A patterned platinum layer functioning as an electrode is made on a substrate that can be biologically modified for capturing the bioactive species.

Pt Substrate Figure 3: The conceptual illustration of an ECL chip with an active electrode surface and bioimmobilization regions.

The chip will be mounted into a disposable cartridge with or without microfluidic channels. In the latter case (see, for example, Fig. 4), the liquid can be brought into the cartridge by using a syringe or external fluidic systems, and remains in contact with the surface modified chip for a sufficiently long period of time so that all the analyte can be captured by the recognition element immobilized on the chip. In addition to the patterned working electrode, the built-in counter and reference electrodes also need to be fabricated within the cartridge. The whole system includes a handheld reader, a disposable cartridge with individually addressed bioimmobilization on the nanostructured electrode chip, and a separate kit of specific reagents. This design concept removes the flow system, electrode regeneration reagents and the associated automation subsystem. Individually addressable cartridges for specific target analytes (such as cancer markers, cardiac marker, anthrax, etc.) can be made by immobilizing the target specific recognition elements on the chip surfaces.

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3 electrodes liquid in

liquid out

Figure 4: Schematic illustration of a disposable ECL cartridge with a nanostructure electrode (top layer) and biologically active regions (not in scale).

3. SPATIALLY SELECTIVE BIOIMMOBILIZATION ON NANOSTRUCTURED CHIPS Due to the size of the biomolecules (DNA, proteins, etc.) and the thickness of the electrochemical diffusion layer, it is obvious that a feature size at the micrometer level will not better address the efficiency issue suggested in Fig. 2. Thus, the fabrication of a nanostructured electrode with spatially selective bioimmobilization becomes a key to the realization of our idea. In contrast to the bead/electrode interface situation shown in Fig. 2, the nanostructured electrode surface (e.g., the top gold layer in Fig. 4) will allow a much bigger portion of the Ru-label molecule to be in the electrochemically active region, thus greatly increasing the signal level by more efficiently stimulating the label molecules, which are attached to the anti-analyte. A variety of methods have been developed to produce bio-functionalized surfaces with feature sizes down to 10 – 102 nanometers. In order to meet our design requirements, we employed the electrochemical template synthesis15-17 on nanostructured surfaces to produce the bioimmobilization regions. The nanostructured surface can be fabricated in two approaches. In one approach, a gold layer is deposited on a substrate followed by the deposition of an aluminum layer, which is then anodically oxidized to form nanopores.18,19 In another approach, the nanostructured surface is formed by diblock copolymers coated on the gold film of a substrate. The porous surface, either alumina or block copolymer, is then coated with a gold layer to form the electrode surface for stimulating ECL. The chip is then brought into an electrochemical cell for growing conducting polymer in the pores using the bottom gold layer as a working electrode. Specifically, a functionalized polymerizable monomer, 1-(2-carboxyethyl)pyrrole or its succinimidyl ester (NHS), is electrochemically polymerized in the nanopores to form nanometric polymer regions with bioconjugatable functional groups for immobilizing target specific biomolecules (Fig. 5).

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CN

O

CH2

HN NH

CH2 N

S (CH3)4 O C NH NH O C

(i ) K O H (i i ) H C l

CH2 CH2

O

COOH

N

Biotinylated PPy

O

O

CH2 ED C

N O

CH2

NHS

N

DCC

Carboxyl PPy

R N N

R N N R

N R

Succinimidyl PPy

Aluminium Gold Substrate Figure 5: Functionalized polypyrole (PPy) electrochemically grown in the nanopores to produce the bioimmobilization regions. A layer of antibody of the target analyte is formed only on the PPy surface. When used for detection, the target of interest will be captured on the PPy surface and the labeled antibody will be located in the vicinity of the ridge of the upper nanostructured gold layer.

3.1 Syntheses of 1-(2-carboxyethyl)pyrrole 1-(2-carboxyethyl)pyrrole was synthesized by following a reported procedure.20,21 25 g (0.20 mole) of 1-(2cyanoethyl)pyrrole was added to 60 mL of potassium hydroxide water solution (23.3 g KOH, 0.41 mole). The mixture was refluxed for six hours. After cooling down to room temperature, the amber solution was acidified with 8 M hydrochloric acid to pH = 5. The product was extracted with diethyl ether (5 × 100 mL) while maintaining the aqueous solution at pH = 5. The ether was roto-evaporated, leaving a brown residue. The crude product was recrystallized three times from boiling n-heptane. After vacuum drying, a colorless crystalline product was obtained (11.6 g, yield 40%). 1H NMR (400 MHz, CDCl3) δ 2.84 (t, J = 7.0 Hz, 2 H), 4.22 (t, J = 7.0 Hz, 2 H), 6.16 (t, J = 1.8 Hz, 2 H), 6.69 (t, J = 1.8 Hz, 2 H), 10.93 (b, 1 H). 3.2 Syntheses of 1-(2-carboxyethyl)pyrrole NHS ester (PyNHS) N,N-Dicyclohexylcarbodiimide (DCC, 99%, 4.15 g, 19.9 mmol) and N-hydroxysuccinimide (NHS, 97%, 2.37 g, 20.0 mmol) were mixed with 2.75 g of 1-(2-carboxyethyl)pyrrole (19.8 mmol) in 20 mL of DMF at 0°C. The solution was stirred at room temperature overnight. The precipitated urea was filtered out and the clear solution was roto-evaporated, giving a white product. The product was further dissolved in acetonitrile and some insoluble solid was removed using a

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0.2 µm syringe filter. 3.6 g (77% yield) of product was obtained. 1H NMR (400 MHz, CDCl3) δ 2.83 (s, 4 H), 3.06 (t, J = 7.0 Hz, 2 H), 4.29 (t, J = 7.0 Hz, 2 H), 6.15 (t, J = 2 Hz, 2 H), 6.69 (t, J = 2 Hz, 2 H). 3.3 Electropolymerization of functionalized pyrrole A preliminary study of the electropolymerization of the functionalized pyrrole was performed in acetonitrile with a platinum disc as the working electrode. The PPy conducting polymers can be either potentiodynamically22 or galvanostatically23 produced. Fig. 6 shows the electrochemical oxidation of PyNHS and the cyclic voltammogram of the produced polymer.

-0.2

0.0

0.2

0.4

0.6

0.8

1.0

1.2

E (V vs. Ag/AgCI)

Figure 6: Potentiodynamic polymerization of PyNHS in acetonitrile. Electrolyte: 0.1 M tetrabutylammonium hexafluorophosphate (TBAPF6). Electrode: 0.785 mm2 Pt.

3.4 Nanostructured surfaces formed by diblock copolymers An efficient method for obtaining regular surface patterns is by the deposition of block copolymer thin films.24-26 A new approach combining block copolymer deposition and supramolecular chemistry was recently developed to achieve nanoscopic patterns of holes on flat surfaces.27-28 This is illustrated by the system shown in Figure 7. An asymmetric polystyrene-b-poly-4-vinylpyridine (PS691-b-P4VP288) copolymer (obtained from Polymer Source) was dissolved in THF, along with 1,5-dihydroxynaphthalene (DHN). Hydrogen bonding takes place between the hydroxy groups of DHN and the pyridine groups of the P4VP, as demonstrated by infrared spectroscopy. The DHN thus selectively enriches the P4VP domains. The resulting thin films were deposited on the substrate by dip-coating. They present cylindrical nodules of P4VP-DHN domains embedded in a PS matrix (Fig. 7a), as confirmed by TEM measurements showing black dots of iodine-stained P4VP domains in a white PS matrix (not stained by iodine) (Fig 7a1). The fast Fourier transform of the image (inset in Fig 7a2) reveals an hexagonal order of the nanodots. The average center-to-center distance between nearest-neighbor cylinders is about 50 nm. The subsequent removal of DHN using methanol leads to the transformation of the nodules into holes (Fig. 7b) due to the decrease in the volume fraction of the P4VP domains. The PS matrix is not solvated by the methanol, so that the ordered structure is maintained, although there is a small decrease in film thickness. The depth of the holes (5-6 nm) compared to the film thickness suggests that they penetrate to the substrate surface, as desired. The light dots observed in the TEM image (Fig. 7b1) seem to corroborate this hypothesis, which is presently being tested by electrochemical methods.

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a1

•

4a

..• a** •• a'

a

•

a•

b1

a2

a3

a

~50

~11

~33

* a* • '

~8

*' *.•

* * a$

:: :.:

P4VP-DHN

b2

Substrate

PS

b3 ~18

~50 ~4

Substrate

~6 PS

P4VP

Figure 7: TEM images (x1), AFM height images (x2) and corresponding schematic representations (x3) of the block copolymer nanostructures after dip-coating (a) and after DHN removal with methanol (b). The nanostructures in the drawings have been laterally “aerated” for better viewing. All data in nm.

4. CONCLUSIONS The magnetic microbeads used in the current commercial systems play the dual role of capturing the analyte and separating the excess labeled antibody from the flow cell. The large surface area provides a high capacity and efficiency for analyte capturing. However, the method prevents the instrument from being miniaturized, and only a tiny portion of the luminescent labels is enabled to generate luminescence. It is proposed that a miniaturized system can be made without having to use the microbeads. The elimination of the magnetic beads and the disposability of the electrode can address some issues related to the miniaturization of bioanalytical devices based on the detection of electrochemiluminescence. A spatially selective ECL chip with nanometer feature sizes can be manufactured by different approaches. Anodic oxidation of aluminum thin film deposited on solid surfaces and the method involving selfassembled diblock copolymer systems represent two economical approaches to volume manufacture of the nanotemplates, on which the spatially selective bioimmobilization can be achieved by electropolymerization of the functionalized pyrrole in the nanopores.

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