Journal of Immunological Methods 295 (2004) 111 – 118 www.elsevier.com/locate/jim
Research paper
Silole nanocrystals as novel biolabels Cangel Pui-yee Chana, Matthias Haeusslerb, Ben Zhong Tangb, Yongqiang Dongb, King-keung Sina, Wing-cheung Makc, Dieter Traud, Matthias Seydacke, Reinhard Renneberga,* a
Department of Chemistry, Biosensors and Bioelectronics Laboratory, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong, China b Department of Chemistry, Institute of Nano Materials and Technology, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong, China c Bioengineering Graduate Program, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong, China d Division of Bioengineering and Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore e 8sens.biognostic AG, Robert-Roessle-Strasse 10, D-13125 Berlin, Germany Received 11 August 2004; accepted 22 September 2004 Available online 4 November 2004
Abstract A novel class of biofunctional silole nanocrystals with the potential to create highly sensitive immunoassay was firstly demonstrated. Biolabels were constructed by encapsulating nanocrystalline hexaphenylsilole [Ph2Si(CPh)4; HPS] within ultrathin polyelectrolyte layers via the layer-by-layer (LbL) technique that provided an binterfaceQ for the attachment of antibodies. A high ratio of fluorescent dyes to biomolecules (F/P ratio; 2.4103) was achieved without self-quenching problem. The aggregation-induced emission (AIE) feature offered silole biolabels the sensitivity 40- to 140-fold higher than that of a start-of-the-art immunoassay using directly fluorescent-labeled antibodies. D 2004 Elsevier B.V. All rights reserved. Keywords: Aggregation-induced emission (AIE); Biolabel; Fluorescence immunoassay (FIA); Layer-by-layer (LbL) technique; Silole
Abbreviations: AIE, aggregation-induced emission; BSA, bovine serum albumin; FITC, fluorescein isothiocyanate; FDA, fluorescein diacetate; FIA, fluorescence immunoassay; Gt a M IgG, goat antimouse IgG; [Ph2Si(CPh)4; HPS], hexaphenylsilole; HPC-SL, hydroxypropyl cellulose; LbL, layer-by-layer; LED, light-emitting diode; M IgG, mouse IgG; PAH, poly(allylamine hydrochloride); PBS, phosphate-buffered saline; PSS, poly(sodium 4-styrenesulfonate); F/P ratio, ratio of fluorescent dyes to biomolecules; SEM, scanning electron microscope; SDS, sodium dodecyl sulfate; THF, tetrahydrofuran. * Corresponding author. Tel.: +852 2358 7387; fax: +852 2705 9670. E-mail address:
[email protected] (R. Renneberg). 0022-1759/$ - see front matter D 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.jim.2004.09.016
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1. Introduction Ratio of fluorescent dyes to biomolecules (F/P ratio) is a key parameter that has an impact on sensitivity and is widely used for the evaluation of a fluorescent label in biochemical assay technologies— high number of labeling molecules per biomolecule is desired. Conventional fluorescence immunoassays (FIA) often encounter problems associated with selfquenching effects. If more than 10–15 fluorophores are attached to one antibody, their distance is close to or within their Foerster radiuses, resulting in significant losses in emission intensities or decreases in fluorescence quantum yields due to involved energy transfer. Aggregation of fluorophores often quenches light emission, which has been a thorny problem in the development of ultrasensitive FIA. In our previous studies (Chan et al., 2004; Trau et al., 2002), we reported on the preparation and utilization of a new class of particulate labels based on nanoencapsulated fluorogenic precursor fluorescein diacetate (FDA) with the potential to create highly amplified biochemical assays. A high molar ratio of fluorescent molecules present in the crystal core to biomolecules on the particle surface was achieved. Following the immunoreaction, the FDA core was dissolved by exposure to organic solvent/sodium hydroxide mixture, leading to the release of the fluorophores
(fluoresceins) into the surrounding medium and thus suppressing self-quenching effect. Our approach provides high sensitivity and low limits of detection without the need for long incubation times, making it an interesting alternative in biolabel technology. This is a pilot study to investigate the feasibility of utilizing siloles as fluorophores in immunoassays. Siloles are of considerable current interest because of their unusual electronic and optical properties, and because of their possible application as electrontransporting materials in light-emitting devices (Chen et al., 2003a,b,c; Luo et al., 2001; Tang et al., 2001; Yamaguchi and Tamao, 1998; Yamaguchi et al., 2000). Siloles exhibit high electron acceptability (Yamaguchi and Tamao, 1998; Sadimenko, 2001; Lee et al., 2000) and fast electron mobility (Murata et al., 2001), which make them efficient luminophores for a variety of optoelectronics applications. These properties arise from the unique low-lying LUMO level associated with the r*–p* conjugation arising from the interaction between the r* orbital of two exocyclic r-bonds on the silicon atom and the p* orbital of the butadiene moiety (Chen et al., 2003a; Tang et al., 2001). The siloles are a group of novel molecules that are highly photoluminescent in their aggregation state. This unique emission feature makes them promising candidate materials for light-emitting diode (LED) applications (Chen et al., 2003a; Tang et
Fig. 1. Schematic illustration of the preparation of biofunctional silole nanocrystals. HPS was synthesized according to published experimental procedure (Tang et al., 2001) (a) and was ball-milled into nanocrystals in an aqueous surfactant, HPC-SL (not shown in the figure) mixed with SDS (b), followed by encapsulation with polyelectrolyte multilayers of nanometer thickness (c), and the attachment of a specific immunoreagent (d).
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Fig. 2. Principle of a sandwich immunoassay using nanocrystalline silole biolabels. The analyte is first immobilized by the capture antibody preadsorbed on the solid phase (a) and then exposed to antibody-labeled nanocrystal detectors (b). Fluorescence intensity is proportional to the analyte concentration.
al., 2001; Yamaguchi et al., 2000; Murata et al., 2001, 2002; Ohshita et al., 2001; Hay et al., 2001) and as chemical sensors (Sohn et al., 2001, 2003). The silole biolabels were constructed by simply encapsulating hexaphenylsilole [Ph2Si(CPh)4; HPS] nanocrystals of 99 nm in average size within ultrathin polyelectrolyte layers of poly(allylamine hydrochloride) (PAH) and poly(sodium 4-styrenesulfonate) (PSS) via layer-by-layer (LbL) techniques (Trau et al., 2002). The polyelectrolyte coating was subsequently used as an interface for the attachment of antibodies through adsorption (Fig. 1). As the entire nanocrystal core is composed of silole molecules and the biomolecule forms a layer on the encapsulated crystal surface, the potentially reachable F/P ratio is exceptionally high. The general concept of the application of the nanocrystalline silole biolabels as fluorescent labels in FIAs is depicted in Fig. 2. The siloles are nonemissive (boffQ) when molecularly dissolved in organic solvents at room temperature, while the silole molecules in poor solvents cluster into nanoaggregates, which turn the emission bonQ and boost the photoluminescence quantum yields by up to two orders of magnitude (Chen et al., 2003a). This intriguing aggregation-induced emission (AIE) feature is an invaluable property for the development of ultrasensitive FIA.
2. Materials and methods 2.1. Synthesis of HPS This compound was prepared as shown in Fig. 1 according to published experimental procedure (Tang
et al., 2001). Briefly, clean lithium shavings (350 mg, 50 mmol) were added to a solution of diphenylacetylene (4.5 g, 25 mmol) in dry tetrahydrofuran (THF; 20 mL). The reaction mixture was stirred at room temperature for 2 h in a dry nitrogen atmosphere. The mixture was then diluted with 120 mL of THF, followed by the addition of 3.2 g (12.5 mmol) dichloro(diphenyl)silane. After refluxing for 5 h, the reaction mixture was cooled and filtered, and the filtrate was washed with water. The organic layer was extracted with diethyl ether and dried over magnesium sulfate. The solvent was removed, and the residue was purified by flash chromatography over silica gel using 10% diethyl ether in petroleum ether as the eluent. Recrystallization from ethanol gave a faintly greenishyellow crystal in 68% yield (4.6 g). 2.2. Preparation of nanocrystalline HPS biolabels Hexaphenylsilole nanocrystal suspension was prepared by ball milling using small glass beads (0.25–0.5 mm, Roth, Karlsruhe, Germany). A suspension of 20 mg of HPS in 2 mL of 1.0% (w/v) hydroxypropyl cellulose (HPC-SL, JE-1071; Nippon Soda, Japan) and 0.05% (w/v) sodium dodecyl sulfate (SDS) was mixed in a glass tube with 2 g of glass beads. The mixture was vortexed for 15 min at room temperature and then exposed to ultrasonic for 15 min. The milling process was repeated until desired particle size distribution was obtained. The colloidal suspension was centrifuged at 6000g (8000 rpm) for 10 min, and the collected supernatant was centrifuged at 16000g (13,100 rpm) for 15 min. The pellet was then resuspended in the surfactant solution. The morphology of the
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milled HPS nanocrystals was examined with a JEOL 6300F ultrahigh-resolution scanning electron microscope (SEM), operating at 10 kV. Particle size distribution was measured based on the Fraunhofer and Mie theories of light scattering using CoulterR LS230 (Beckman Coulter, USA) by polarization intensity differential scattering technology. Polyelectrolyte multilayers were assembled onto the nanocrystals by the sequential deposition of poly(allylamine hydrochloride) (PAH-polyelectrolyte; M w 15,000) and poly(sodium 4-styrenesulfonate) (PSS-polyelectrolyte; M w 70,000) as described by Trau et al. (2002); a small amount (0.1 mL) of the HPS nanocrystal suspension (1.0%, w/v) was added to 0.5 mL of PAH–polyelectrolyte solution (5 mg/mL, containing 0.5 M NaCl). The suspension was mixed at constant intervals for 15 min at room temperature. The excess polyelectrolyte was removed by three repeated centrifugation/washing with distilled water and redispersion cycles. For subsequent assembly of negatively charged PSS-polyelectrolyte, 0.5 mL of PSS–polyelectrolyte solution (5 mg/mL, containing 0.5 M NaCl) was added. The centrifugation/washing and polyelectrolyte incubation steps were repeated until the desired number of layers (typically four) was assembled. The surface charges of the milled and encapsulated HPS nanocrystals were examined by microelectrophoresis using a ZetaPlus potential analyzer (Brookhaven Instrument, Holtsville, NY) by taking the average of five measurements at the stationary level. The polyelectrolyte-coated HPS nanocrystals with an outermost layer of PSS-polyelectrolyte were conjugated to antibodies as described in our previous study (Chan et al., 2004). The particle suspension (0.0626%, w/v) was incubated with 200 Ag/mL of polyclonal goat antimouse IgG (Gt a M IgG, whole molecule; Arista Biologicals, USA) in 10 mM phosphate-buffered saline (PBS, pH 7.4) at 20 8C for 1 h. After centrifugation at 16,000g (13,100 rpm) for 10 min, the supernatant was removed, and its UV absorption was measured at 280 nm (Cary 50 Conc UV–Visible Spectrophotometer, Australia). The antibody surface coverage of nanoparticles was determined by the difference in absorption at 280 nm between supernatant and the original protein solution. The IgG-coated particles were then separated from soluble IgG by three centrifugation/washing cycles.
2.3. Solid-phase sandwich fluorescence immunoassay Two microgram per milliliter of Gt a M IgG (100 AL/well) was coated on Nunc Maxisorp 96-well microplates (Nunc International, Rochester, NY) in 0.1 mol/L carbonate buffer (pH 9.6) at 4 8C overnight. After rinsing three times with washing buffer [10 mM PBS, 0.1% (w/v) bovine serum albumin (BSA, fraction V), 0.5% (w/v) Tween-20], the wells were blocked with 300 AL/well of 1.0% BSA solution for half an hour at 37 8C. The plate was then washed four times and incubated with dilutions (100 AL/well) of mouse IgG (M IgG; Arista Biologicals) as an analyte at 37 8C for 1 h. After washing five times, antimousecoated nanocrystal suspensions (0.0125%, w/v) was dispensed into the wells (100 AL/well), and the microplate was incubated again at 37 8C for 1 h. Soluble fluorescein isothiocyanate (FITC)-labeled Gt a M IgG dilutions (100 AL/well) of 1:128 (Arista, protein concentration 1.1 mg/mL, F/P ratio 4.4) was used for comparison. After incubation, excess detector antibody conjugates were washed off by five washing cycles with buffer. The fluorescence intensity was measured using a FLUOstarOPTIMA multifunctional microplate reader (BMG Labtechnologies, Germany) with excitation/emission wavelengths of 380/500 and 485/520 nm for measurements of the nanocrystalline HPS-labeled and the FITC-labeled antibodies (gain setting of 1600).
3. Results and discussion This is a novel study to apply the aggregationinduced emission principle in immunoassay. Hexaphenylsilole was chosen as the fluorophore in this study due to the impressive performance of its LED device (Chen et al., 2003a). It was readily turned on at a low voltage (~4 V), emitted intensely at a moderate bias (55 880 cd/m2 at 16 V), and showed very high emission efficiencies (15 cd/A current efficiency, 10 1m/W power efficiency, and 7% external quantum efficiency). HPS was synthesized according to published procedures by ring-closing reaction of 1,4-dilithio-2,3,4,5-tetraphenyl-1,3-butadiene with dichlorodiphenylsilane (Tang et al., 2001). DMSO solution of HPS (22 AM) was virtually nonemissive. Only a noisy curve was
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Fig. 3. Photoluminescence spectra of 22 AM of HPS in DMSO (red) and water (blue); excitation wavelength: 380 nm.
obtained even in a 100 times magnified photoluminescent spectrum (Fig. 3). However, when the same concentration of HPS in a poor solvent (e.g., water), an intense signal was recorded under identical measurement conditions. This observation fits well to the data obtained from the previous investigation and confirms that HPS is AIE-active (Chen et al., 2003a).
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According to the previous study (Chen et al., 2003a), the AIE phenomenon is caused by the restricted intramolecular rotation. Rotational energy relaxation can nonradiatively deactivate excited species (Wise et al., 1998; Wong et al., 1998). In the solutions at room temperature, the active intramolecular rotations of the peripheral phenyl rings around the axes of the single bonds linked to the central silole core may effectively annihilate the excitons, thus making the silole molecules nonemissive. In the solid aggregates, the stacking forces involved in the crystal packing may restrict the intramolecular rotations, which may block the nonradiative channel and populate the radiative decay, thus making the silole molecules luminescent. The particle size distribution of the HPS nanocrystals determined by light-scattering measurements is shown in Fig. 4. A narrow disperse system was obtained with an average size of 99 nm. Approximately, 90% of the nanocrystals were found to be smaller than 185 nm, and 100% (or all of them) smaller than 500 nm. This result is in agreement with SEM analysis of the nanocrystals as shown in Fig. 4. SEM shows that the nanocrystals take different shapes. During the milling process, the hydrophobic surface of the stabilizer is most likely associated with the hydrophobic surface of the nanocrystal, and the hydrophilic portion of SDS is oriented towards the aqueous phase of the suspension (Caruso et al., 2000). The adsorbed HPC-SL layer makes the HPS nano-
Fig. 4. Particle size distribution and SEM micrograph of milled HPS nanocrystals.
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crystals dispersible in water and prevents their aggregation and crystal growth following extended storage periods, hence conferring colloidal stability (Liversidge et al., 1999). The adsorbed, negatively charged SDS layer introduces surface charges to the nanocrystals and also increases colloidal stability. This was verified by microelectrophoresis measurements. The nanocrystals exhibited a f-potential of 44.5 mV, indicating a negatively charged, highly stable particle suspension (Fig. 5). The subsequent alternate adsorptions of PAH- and PSS-polyelectrolyte layers onto the SDS-coated HPS nanocrystals yielded f-potentials of ca. +30 and 40 mV, respectively. The thickness of the polyelectrolyte coating, four layers total of PAH- and PSS-polyelectrolytes, assembled onto the amphiphile-modified HPS nanocrystals is only about 6–8 nm (Caruso et al., 1999; Caruso and Mo¨hwald, 1999). This thickness is negligible when compared to the total size of the HPS nanocrystals. The polyelectrolyte coatings did not cause any noticeable change in the morphology and size distribution of the HPS nanocrystals (data not shown). The polyelectrolyte coating on the HPS nanocrystals is to impart sufficient colloidal stability and to provide a suitable interface for the attachment of biomolecules to the nanocrystals. Gt a M IgG adsorption onto polyelectrolyte multilayers assembled onto polystyrene microparticles has been confirmed by monitoring the change in f-potentials as a function of particle suspension pH (Yang et al., 2001) and quantified by single particle light-scattering experiments (Caruso and Mo¨hwald, 1999). In this study, the amount of Gt a M IgG adsorbed onto the HPS nanocrystals was determined spectrophotometrically.
Fig. 5. Microelectrophoresis measurements of the HPS nanocrystals by taking the average of five measurements at the stationary level.
Fig. 6. Sandwich fluorescence immunoassay of M IgG using Gt a M IgG-HPS nanocrystals ( ) and Gt a M IgG-FITC ( ) as labels. The reported fluorescence intensities were obtained after the subtraction of the blank signal. Error bars correspond to standard deviations (FS.D., n=3).
The protein surface coverage was calculated by assuming an average size of 99 nm for the nanocrystals. Incubation of the nanocrystals with Gt a M IgG (200 Ag/mL) in 10 mM PBS buffer for 1 h at 20 8C resulted in the adsorption of 36.7% of the added protein, correlating to a surface coverage of 2.37 mg/m2. The theoretically calculated surface coverage value for a close-packed IgG monolayer is in the range of 2.0–5.5 mg/m2, depending on the different orientations of the adsorbed IgG molecules (Davalos-Pantoja et al., 2001; Caruso et al., 1998). To calculate the F/P ratio for the nanocrystal biolabel, a cubic crystal morphology with dimensions of 999999 nm was assumed. The calculated F/P value for the biolabel is 2.4103, which is much higher than the ratios of directly covalent labeled antibodies (carrying four to eight fluorophores in general). The F/P ratio of an immune detection system reflects its potential amplification rate. Fig. 6 shows calibration curves of the sandwich fluorescence immunoassay performed with Gt a M IgG adsorbed nanocrystal labels in comparison with a
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direct FITC-labeled antibody conjugate. The fluorescence signal is directly proportional to the M IgG concentration in the range of 0–100 Ag/L. A 40- to 140-fold higher sensitivity with the assays using the nanocrystal biolabels was observed compared with the direct FITC-labeled antibodies, depending on the analyte concentration. The higher sensitivity of the nanocrystal biolabels compared with standard FITC conjugates may be explained by the boosting effect of the higher ratio of dye molecules to binding molecules, and the suppressed self-quenching by the AIE feature could also have contributed to the improved signal. This is the first study to show the successful application of siloles as fluorophores in immunoassay. In conclusion, the applicability of a new class of biolabels for immunoassays was demonstrated. Chemo- and photostabilities make siloles a useful tool for labeling purposes. The preparation of nanocrystalline HPS biolabel is straightforward to perform and is controllable. By using a nanoscale polyelectrolyte coating as an interface for bioconjugation onto the nanocrystals, the siloles do not need to be watersoluble or possess groups for bioconjugation, as this functionality is provided by the polyelectrolytes. The quenching problem normally arising from F/P ratio labels can be prevented due to the AIE feature of siloles. Synthesis of water-soluble amphiphilic siloles through appropriate chemical modifications is now in progress. The soluble siloles may be used in homogenous assays, i.e., all reagents present in solution to achieve the measurement without any separation step. We believe the use of siloles provides us with a useful alternative to organic fluorophores after optimization. Acknowledgements The work described in this paper was partially supported by the Hong Kong Research Grant Council (HKUST6086/02M and HKUST6085/02P) and the University Grants Committee under an Area of Excellence (AoE) scheme (AoE/P-10/01-1). References Caruso, F., Mfhwald, H., 1999. Protein multilayer formation on colloids through a step-wise self-assembly technique. J. Am. Chem. Soc. 121, 6039.
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