Anal. Chem. 2008, 80, 573-582
Articles
Poly(styrene-block-vinylpyrrolidone) Beads as a Versatile Material for Simple Fabrication of Optical Nanosensors Sergey M. Borisov,* Torsten Mayr, and Ingo Klimant
Institute of Analytical Chemistry and Radiochemistry, University of Technology Graz, Stremayrgasse 16, 8010 Graz, Austria
A versatile platform for designing optical nanosensors is proposed. The “sensing chemistries” are entrapped into the poly(styrene-block-vinylpyrrolidone) nanobeads having the average size of 245 nm in aqueous media. Addressable staining into the core or the shell of the beads results in nanosensors for essential analytes such as dissolved oxygen, temperature, pH, chloride, and copper ions. Two immobilization procedures are developed: staining in the polystyrene core is performed from a tetrahydrofuran/water mixture (50:50 v/v) and staining in the poly(vinylpyrrolidone) shell is achieved by using the ethanol/water mixture (70:30 v/v). The oxygen and temperature indicators should be preferably immobilized into the core, whereas nanosensors for ions are manufactured by staining into the shell. In the case of the lipophilic pH indicators both procedures result in similar pKa values. The unique properties of the beads make them promising for sensing and imaging even in very complex media, multianalyte sensing, and monitoring of very fast processes. Optical chemical sensing has become very widespread in the last 2 decades. It is mostly based on the use of smart materials with optical properties that respond to a chemical parameter.1,2 Most sensor materials consist of an indicator dye dissolved in a polymeric matrix which also acts as a solid support and as a permeation-selective membrane. Such materials are used in numerous formats including planar sensor spots,3,4 fiber-optic systems,5 two-dimensional foils,6,7 and pressure-sensitive paints.8,9 * Corresponding author. Phone: +43 316 873 4326. Fax: +43 316 873 4329. E-mail:
[email protected]. (1) Wolfbeis, O. S. Anal. Chem. 2006, 78, 3859-3874. (2) Wolfbeis, O. S. J. Mater. Chem. 2005, 15, 2657-2669. (3) Hartmann, P.; Trettnak, W. Anal. Chem. 1996, 68, 2615-2620. (4) Mills, A.; Thomas, M. Analyst 1997, 122, 63-68. (5) Rosenzweig, Z.; Kopelman, R. Anal. Chem. 1995, 67, 2650-2654. (6) Koese, M. E.; Carrol, B. F.; Schanze, K. S. Langmuir 2005, 21, 91219129. (7) Liebsch, G.; Klimant, I.; Frank, B.; Holst, G.; Wolfbeis, O. S. Appl. Spectrosc. 2000, 54, 548-559. (8) Zelelow, B.; Khalil, G.; Phelan, G.; Carlson, B.; Gouterman, M.; Callis, J. B.; Dalton, L. R. Sens. Actuators, B 2003, 96, 304-314. (9) Hradil, J.; Davis, C.; Mongey, K.; McDonagh, C.; MacCraith, B. D. Meas. Sci. Technol. 2002, 13, 1552-1557. 10.1021/ac071374e CCC: $40.75 Published on Web 01/04/2008
© 2008 American Chemical Society
Imaging, in particular, becomes increasingly popular due to the possibility of real time analyte monitoring over a large area or in volume. Imaging of oxygen,10,11 pH,12,13 CO2,14,15 ions,16 and biomolecules such as glucose17 is of wide interest in various fields of science and technology including marine research,10,13,14,18 clinical medicine,19 and biotechnology15 to mention only a few. Imaging of the total pressure of air (via oxygen pressure) on the surface of aircrafts and cars20,21 is performed by means of pressuresensitive paints. Planar sensor foils and pressure-sensitive paints which have been mostly used so far for imaging purposes are, however, not universal tools for several reasons. These include (a) inability of monitoring of very fast processes such as enzymatic reactions since significant thickness of the sensor layer (usually several micrometers) and the presence of a thick virtually gasimpermeable support results in relatively long response times. This can be handled by using (sub)micrometer-sized fiber-optic sensors5,22 which respond virtually in real time; however, mechanical stability of those is often poor; (b) unsuitability for imaging in a variety of flow-through cells and bioreactors, where only small spots of the planar material can be positioned; (c) certain difficulties for multianalyte sensing, which can be compromised by inhomogeneities of a sensor layer and reduced photostability.23 The above limitations can be overcome by making use of analyte(10) Koenig, B.; Kohls, O.; Holst, G.; Glud, R. N.; Kuehl, M. Mar. Chem. 2005, 97, 262-276. (11) Schroeder, C. R.; Polerecky, L.; Klimant, I. Anal. Chem. 2007, 79, 60-70. (12) Liebsch, G.; Klimant, I.; Krause, Ch.; Wolfbeis, O. S. Anal. Chem. 2001, 73, 4354-4363. (13) Stahl, H.; Glud, A.; Schroder, C. R.; Klimant, I.; Tengberg, A.; Glud, R. N. Limnol. Oceanogr.: Methods 2006, 4, 336-345. (14) Zhu, Q.; Aller, R. C.; Fan, Y. Mar. Chem. 2006, 101, 40-53. (15) Borisov, S. M.; Krause, Ch.; Arain, S.; Wolfbeis, O. S. Adv. Mater. 2006, 18, 1511-1516. (16) Mayr, T.; Liebsch, G.; Klimant, I.; Wolfbeis, O. S. Analyst 2002, 127, 201203. (17) Schaeferling, M.; Wu, M.; Wolfbeis, O. S. J. Fluoresc. 2004, 14, 561-568. (18) Holst, G.; Grunwald, B. Sens. Actuators, B 2001, 74, 78-90. (19) Babilas, P.; Liebsch, G.; Schacht, V.; Klimant, I.; Wolfbeis, O. S.; Szeimies, R. M.; Abels, C. Microcirculation 2005, 12, 477-487. (20) Demas, J. N.; DeGraff, B. A.; Coleman, P. B. Anal. Chem. 1999, 71, 793A800A. (21) Gouterman, M.; Callis, J.; Dalton, L.; Khalil, G.; Mebarski, Y.; Cooper, K. R.; Greiner, M. Meas. Sci. Technol. 2004, 15, 1986-1994. (22) Tan, W.; Shi, Z.-Y.; Kopelman, R. Anal. Chem. 1992, 64, 2985-2990. (23) Borisov, S. M.; Neurauter, G.; Schroeder, C.; Klimant, I.; Wolfbeis, O. S. Appl. Spectrosc. 2006, 60, 1167-1173.
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Figure 1. Chemical structures of the indicators used in nanosensors.
sensitive nanobeads which will also allow for three-dimensional (3D) imaging. Moreover, since nanobeads can be dispersed in small volume, very low absolute limits of detection are achieved. Dissolved indicators can also be used, but such systems are prone to various interferences, such as water, ions, and other species, and they can interact with other components of the medium such as proteins. Moreover, many of the indicators possess extremely low solubility in aqueous solutions. Dendrimeric indicators are promising,24 but sophisticated synthesis is necessary. The group of Kopelman has developed a variety of analytesensitive nanobeads, so-called “PEBBLEs”,25 which are mostly based on polyacrylamide and poly(decyl methacrylate) and have typical size from 50 to 300 µm. Nanobeads for sensing and imaging oxygen,26,27 pH,28 Ca2+,28,29 Mg2+,30 K+,31 Fe3+,32 Zn2+,33 and Cl- 34 were reported. The group also developed nanobiosensors for (24) Wilson, D. F.; Lee, W. M. F.; Makonnen, S.; Finikova, O.; Apreleva, S.; Vinogradov, S. A. J. Appl. Physiol. 2006, 101, 1648-1656. (25) Clark, H. A.; Hoyer, M.; Philbert, M. A.; Kopelman, R. Anal. Chem. 1999, 71, 4831-4836. (26) Xu, H.; Aylott, J. W.; Kopelman, R.; Miller, T. J.; Philbert, M. A. Anal. Chem. 2001, 73, 4124-4133. (27) Cao, Y.; Lee Koo, Y.-E.; Kopelman, R. Analyst, 2004, 129, 745-750. (28) Clark, H. A.; Hoyer, M.; Parus, S.; Philbert, M. A.; Kopelman, R. Mikrochim. Acta 1999, 131, 121-128. (29) Clark, H. A.; Kopelman, R.; Tjalkens, R.; Philbert, M. A. Anal. Chem. 1999, 71, 4837-4843. (30) Park, E. J.; Brasuel, M.; Behrend, C.; Philbert, M. A.; Kopelman, R. Anal. Chem. 2003, 75, 3784-3791. (31) Brasuel, M.; Kopelman, R.; Miller, T. J.; Tjalkens, R.; Philbert, M. A. Anal. Chem. 2001, 73, 2221-2228. (32) Sumner, J. P.; Kopelman, R. Analyst 2005, 130, 528-533. (33) Sumner, J. P.; Aylott, J. W.; Monson, E.; Kopelman, R. Analyst 2002, 127, 11-16. (34) Brasuel, M. G.; Miller, T. J.; Kopelman, R.; Philbert, M. A. Analyst 2003, 128, 1262-1267.
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Cu2+ 35 and glucose.36 Other groups reported dyed polystyrene nanoparticles for sensing oxygen,37 quantum dots for temperature sensing (however, suitable for measurements only in some nonaqueous solvents),38,39 and beads based on plasticized poly(vinyl chloride) for sensing chloride ion.40 Oxygen-sensitive liposomes (∼70 nm in diameter) were shown to degrade constantly with time.41 In this work we report poly(styrene-block-vinylpyrrolidone) as a novel matrix for designing optical nanosensors. This versatile matrix allows for “sensing chemistry” to be incorporated either into the core or the shell of a bead which for some nanosensors can slightly tune the sensitivity and helps to minimize interferences. Examples include nanosensors for oxygen, temperature, pH, chloride, and copper ions. EXPERIMENTAL SECTION Materials and Suppliers. Platinum(II) 5,10,15,20-tetrakis(2,3,4,5,6-pentafluorophenyl)-porphyrin (PtTFPP) was obtained from Frontier Scientific (www.frontiersci.com), poly(styrene-blockvinylpyrrolidone) emulsion in water (38% w/w emulsion in water) was purchased from Aldrich (www.sigmaaldrich.com). Nile red, lucigenin, lucifer yellow-CH, sodium dodecyl sulfate (SDS), hexadecyltrimethylammonium chloride (HDTMA), sodium hy(35) Sumner, J. P.; Westerberg, N. M.; Stoddard, A. K.; Fierke, C. A.; Kopelman, R. Sens. Actuators, B 2006, 113, 760-767. (36) Xu, H.; Aylott, J. W.; Kopelman, R. Analyst 2002, 127, 1471-1477. (37) Im, S. H.; Khalil, G. E.; Callis, J.; Ahnb, B. H.; Gouterman, M.; Xia, Y. Talanta 2005, 67, 492-497. (38) Jorge, P. A.; Mayeh, M.; Benrashid, R.; Caldas, P.; Santos, J. L.; Farahi, F. Meas. Sci. Technol. 2006, 17, 1032-1038. (39) Wang, S.; Westcott, S.; Chen, W. J. Phys. Chem. B 2002, 106, 11203-11209. (40) Ceresa, A.; Qin, Y.; Peper, S.; Bakker, E. Anal. Chem. 2003, 75, 133-140. (41) McNamara, K. P.; Rosenzweig, Z. Anal. Chem. 1998, 70, 4853-4859.
Table 1. Composition of the Nanosensors code
analyte
dye
content, % w/w
procedure
code
analyte
dye
content, % w/w
procedure
N1 N2 N3 N4
oxygen oxygen T pH
PtTFPP PtTFPP Eu(tta)3L CHFOE
1.5 1.5 1.5 0.25
EtOH THF THF EtOH
N6 N7 N8 N9
pH pH ClCu2+
HPTS(OA)3 HPTS(OA)3 lucigenin lucifer yellow-CH
1.5 1.5 0.23 0.175
EtOH THF EtOH EtOH
N5
pH
CHFOE
0.25
THF
drogen phosphate, sodium dihydrogen phosphate, and sodium chloride were from Fluka (www.sigmaaldrich.com). Tetrahydrofuran (THF) and ethanol were obtained from Roth (www.carlroth.de). Nitrogen and synthetic air (all of 99.999% purity) were obtained from Air Liquide (www.airliquide.at). All the components of the cultivation media were obtained from Becton, Dickinson and Company (www.bd.com) and Roth. Synthesis of 2′-chloro-7′-hexylfluorescein octadecylester (CHFOE) is reported elsewhere.42 The pH indicator 1-hydroxypyrene3,6,8-tris-octylsulfonamide (HPTS(OA)3) was prepared according to the procedure of Mohr et al.43 The temperature probe europium(III) tris(thenoyltrifluoroacetonate) dipyrazoltriazine complex (Eu(tta)3L) was prepared according to the literature procedure.44 The chemical structures of the indicators are presented in Figure 1. Preparation of the Oxygen-Sensitive Nanobeads N1. Five hundred twenty-six milligrams of the polymer emulsion (containing 200 mg of the polymer beads) was diluted with the mixture of 80 mg of ethanol and 40 mg of water. Then, 3 mg of PtTFPP was dissolved in 20 mL of ethanol, and the solution was added dropwise under vigorous stirring into emulsion of the polymer. The emulsion was concentrated under reduced pressure until all ethanol was removed. It was then diluted with water up to 20 mL overall volume. Preparation of the Oxygen-Sensitive Nanobeads N2. Five hundred twenty-six milligrams of the polymer emulsion was diluted with the mixture of 50 mL of water and 30 mL of THF. Three milligrams of PtTFPP was dissolved in 20 mL of THF, and the solution was added dropwise under vigorous stirring into emulsion of the polymer. THF was removed under reduced pressure, and the suspension was diluted with water up to 20 mL overall volume. Preparation of the Temperature-Sensitive Nanobeads N3. It was performed according to the previous protocol; however, 3 mg of Eu(tta)3L was used instead of the same amount of PtTFPP. Preparation of the pH-Sensitive Nanobeads N4. One thousand and fifty-two milligrams of the polymer emulsion was diluted with the mixture of 80 mg of ethanol and 40 mg of water. Then, 1 mg of CHFOE was dissolved in 20 mL of ethanol, and the solution was added dropwise under vigorous stirring into the emulsion of the polymer. The emulsion was concentrated under reduced pressure until all ethanol was removed. It was then diluted with water up to 20 mL overall volume. Preparation of the pHsensitive nanobeads N6 was performed in the similar manner; however, 6 mg of the dye (HPTS(OA)3) was used. Other (42) Weidgans, B. M.; Krause, Ch.; Klimant, I.; Wolfbeis, O. S. Analyst 2004, 129, 645-650. (43) Mohr, G. J.; Werner, T.; Wolfbeis, O. S. J. Fluoresc. 1995, 5, 135-138. (44) Borisov, S. M.; Wolfbeis, O. S. Anal. Chem. 2006, 78, 5094-5101.
pH-sensitive nanobeads (N5 and N7) were prepared according to the procedure for the oxygen-sensitive nanobeads N1, and 0.5 mg of CHFOE and 1.5 mg of HPTS(OA)3 were used for 526 mg of polymer emulsion, for preparation of N5 and N7, respectively. Preparation of the Chloride-Sensitive Nanobeads N8. Three hundred milligrams of polymer emulsion was diluted with 2.7 mL of water. Then, 200 µL of an aqueous lucigenin stock solution (1.3 mg/mL) and 200 µL of an aqueous SDS stock solution (30 mg/mL) were added to 2 mL of ethanol. This solution was added dropwise under vigorous stirring into 1 mL of the polymer emulsion. To this emulsion 9 mL of water was added. The emulsion was dialyzed against water. The reservoir was changed four times every 3 days until it was colorless. Preparation of the Copper(II)-Sensitive Nanobeads N9. Three hundred milligrams of polymer emulsion was diluted with 2.7 mL of water. Then, 200 µL of aqueous lucifer yellow-CH stock solution (1 mg/mL) and 64 µL of aqueous HDTMA stock solution (38 mg/mL) were added to 2 mL of ethanol. This solution was added dropwise under vigorous stirring into 1 mL of the polymer emulsion. To this emulsion 9 mL of water was added. The emulsion was dialyzed against water. The reservoir was changed four times every 3 days until it was colorless. The composition of the sensor materials is summarized in Table 1. Measurements. The size of the beads was determined with a particle size analyzer Zetasizer Nano ZS (www.malvern.de). Luminescence phase shifts were measured with a two-phase lock-in amplifier (SR830, Stanford Research Inc., www.thinksrs.com). The emulsion of the beads in water (∼1 mg/mL) contained in a glass vial was excited with the light of a violet LED (λmax 405 nm, www.roithner-laser.com) which was sinusoidally modulated at 5 kHz or at 700 Hz, respectively, in the case of the oxygen-sensitive and temperature-sensitive beads. A bifurcated fiber bundle was used to guide the excitation light to the vial and to guide back the luminescence after passing through the OG 590 (Schott) glass filter. The luminescence was detected with a photomultiplier tube (H5701-02, Hamamatsu, www.sales.hamamatsu.com). Temperature was controlled by a cryostat ThermoHaake DC50. In the case of the oxygen-sensitive beads the temperature was kept constant at 1, 25, and 50 °C. Gas calibration mixtures were obtained using a gas mixing device (MKS, www.mksinst.com). Three independent measurements were performed to obtain a calibration curve. Fluorescence measurements for pH-sensitive beads N4 and N5 were performed on a Hitachi F-7000 fluorescence spectrometer (www.inula.at). The pH was adjusted to the desired value using phosphate buffer. The pH of the buffer solutions was controlled Analytical Chemistry, Vol. 80, No. 3, February 1, 2008
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by a digital pH meter (InoLab pH/ion, WTW GmbH & Co. KG, www.wtw.com) calibrated at 20 ( 2 °C with standard buffers of pH 7.0 and 4.0 (WTW GmbH & Co. KG). The buffers were adjusted to constant ionic strength using sodium chloride as the background electrolyte. Three independent measurements were performed to obtain a calibration curve. Fluorescence measurements of the nanobeads of types N6 and N7 were performed in 96-well microplates using a BMG Labtech Fluorstar Optima microplate reader (www.bmg-labtechnologies.com) equipped with 485/20 and 560/20 band-pass filters for excitation and emission, respectively. Eight independent measurements were performed in parallel to obtain a calibration curve. Fluorescence measurements of ion-sensitive nanobeads were performed in 96-well microplates using a BMG Labtech Fluorstar Optima microplate reader with 420/20 and 540/20 band-pass filters for excitation and emission, respectively. The pH of the copper(II)-containing solutions was adjusted to pH 5 using a 10 mM acetate buffer. The pH of the chloride sample solutions was adjusted to 7.1 with a phosphate buffer (C(H2PO4- + HPO42-) ) 13 mM) and to constant ionic strength of 230 mM using sodium fluoride as the background electrolyte. Eight independent measurements were performed in parallel to obtain a calibration curve. Leaching Tests. Two milliliters of the emulsion in water (10 mg beads/mL) was placed in a syringe and pressed through a fine filter (220 nm pore size, Rotilabor, Roth, www.carl-roth.de). The first 0.5 mL of the slightly turbid filtrate was discarded, and the main transparent fraction was collected and investigated spectroscopically. Leaching in the standard Luria broth () LB) medium (10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl) was investigated in the same manner. Prior to filtration, the mixture of 2 mL of the bead emulsion and 2 mL of LB medium was stirred at room temperature for 24 h. Toxicity and Penetration Tests. A preculture of Pichia pastoris (strain: X-33 pGAPZB-PaHNL5R (BT2869)) was grown in 50 mL YPD 1% (w/v) medium (1% yeast extract, 2% peptone, 1% dextrose) in 500 mL baffled shake flasks to OD600 ) 2.0. The main culture was incubated to OD600 ) 0.26 (7.5 mL of the preculture was added to 50 mL of the main culture) in BMD 1% (w/v) medium (1.34% yeast nitrogen base w/o amino acids, 1% dextrose, 0.02% D-biotin, 2% 10× PPB (30.13 g/L K2HPO4‚3H2O, 118.13 g/L KH2PO4)) in 500 mL buffeled shake flasks. Growth conditions for both cultures were 28 °C and 120 rpm (incubator: Certomat BS-1, www.sartorius-stedim.com). Two milliliters of the emulsion of the nanobeads (N3, 10 mg/mL) was added to the main culture, and the cells were monitored microscopically after 6, 12, and 24 h of cultivation. A preculture of Escherichia coli (strain: BL 21 gold) was grown in 25 mL of LB medium to OD600 ) 4.0 in 100 mL shake flasks. The main culture was incubated to OD600 ) 0.52 (7.5 mL of the preculture was added to 50 mL of the main culture) in LB medium in 600 mL shake flasks. Growth conditions for both cultures were 30 °C and 160 rpm (incubator: Certomat BS-1). Two milliliters of the emulsion of the nanobeads (N3, 10 mg/mL) was added to the main culture, and the cells were monitored microscopically after 3 and 6 h of cultivation. The images were acquired on an Axiolab microscope (Zeiss, www.zeiss.de) equipped with a ProgRes C14 digital camera (Zeiss). 576 Analytical Chemistry, Vol. 80, No. 3, February 1, 2008
Figure 2. Size and distribution of the beads in different solvents and solvent mixtures.
Fitting was performed using Origin version 7.5 (www.originlab.com) software. RESULTS AND DISCUSSION Properties and Staining of the Beads. As specified by the manufacturer, the poly(styrene-block-vinylpyrrolidone) nanobeads consist of 64% w/w of styrene and 36% w/w of vinylpyrrolidone and are 0.9997). Since f1 values are rather uniform for all the temperatures investigated, medium values were used for calculation of the Stern-Volmer constants. For the beads obtained from ethanol the fitting gives K1SV of 0.201, 0.329, and 0.500 kPa-1 for 1, 25, and 50 °C, respectively. The K2SV was found to be 0, and f1 was 0.836 ( 0.01. For the beads stained from THF the fitting gave K1SV of 0.151, 0.271, and (49) Lee, S.; Okura, I. Anal. Commun. 1997, 34, 185-188. (50) Amao, Y.; Miyashita, T.; Okura, I. J. Fluorine Chem. 2001, 107, 101-106. (51) Spellane, P. J.; Gouterman, M.; Antipas, A.; Kim, S.; Liu, Y. C. Inorg. Chem. 1980, 19, 386-391. (52) Borisov, S. M.; Klimant, I. Anal. Chem. 2007, 79, 7501-7509. (53) Sacksteder, L.; Demas, J. N.; DeGraff, B. A.; Bacon, J. R. Anal. Chem. 1993, 65, 3480-3483.
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Figure 4. Calibration curves for the oxygen-sensitive beads based on PtTFPP: (a and b) beads stained from EtOH/H2O (N1); (c and d) beads stained from THF/H2O (N2). Fitting of the Stern-Volmer plots (b and d) is performed using eq 1.
0.357 kPa-1, K2SV of 0.022, 0.012, and 0.017 kPa-1 for 1, 25, and 50 °C, respectively (f1 was 0.767 ( 0.14). It is evident that staining from ethanol results in more sensitive nanobeads. Fine-tuning of sensitivity to oxygen is therefore possible. Notably, quenching efficiency in the case of the beads stained in the core is slightly higher than that for the PS film (τ0/τ at 21.3 kPa was 2.9, 3.4, and 3.9, respectively, for 1, 25, and 50 °C in the case of the beads, and τ0/τ was 2.9, 3.2, and 3.3, respectively, for 1, 25, and 50 °C in the case of the sensor films). This can be explained by the fact that the decay times in the absence of oxygen are higher in the beads (65.1, 61.1, and 56.9 µs at 1, 25, and 50 °C, respectively) than in the film (59.2, 55.4, and 51.0 µs at 1, 25, and 50 °C, respectively). As expected, the nanosensors exhibit higher sensitivity to oxygen at elevated temperatures (Figure 4). Such behavior is common for all optical oxygen sensors, and the temperature effects need to be compensated for if measurements are performed at varying temperatures. Evidently, the sensitivity of the nanosensors can be varied significantly if other oxygen indicators are used instead of the platinum(II) porphyrin. For example, the palladium(II) complex of 5,10,15,20-tetrakis-(2,3,4,5,6-pentafluoro-phenyl)-porphyrin (PdTFPP) is an ideal candidate if measurements of trace oxygen in practically anaerobic solutions are performed, since its decay time of ∼1 ms will enable high oxygen sensitivity of the beads. On the other side, if measurements need to be carried out under pO2 exceeding 21 kPa, a widely used ruthenium(II)-tris-4,7-diphenyl1,10-phenanthroline probe (Ru-dpp, τ0 ∼ 6 µs) can be immobilized. In fact, we have found that PdTFPP and Ru-dpp, as well as phosphorescent iridium(III) coumarin complexes52 are easily immobilized into the beads using the same protocols.54 For example, in the case of Ru-dpp τ0/τ was found to be 1.45 at 25 °C and 21.3 kPa.54 (54) Borisov, S. M.; Klimant, I. Unpublished results, 2007.
578 Analytical Chemistry, Vol. 80, No. 3, February 1, 2008
Figure 5. Calibration curves for the temperature-sensitive beads of type N3. Fitting is performed using eq 2.
Nanobeads for Sensing Temperature (N3). We have previously shown44 that the europium(III) tris(thenoyltrifluoroacetonate) complex with coordinated antenna chromophore (“Eu(tta)3L”) can serve as an excellent temperature probe for luminescence sensing and imaging. This complex is excitable by visible light (λmax 402 nm in toluene) and benefits from very strong brightness (Bs, defined as the product of quantum yield and the molar absorbance at the excitation wavelength) which is ∼28 000 at 25 °C and ∼48 000 at 1 °C.44 In this work, the complex was immobilized into the core of the nanobeads using the THF/water procedure. It should be noted that it is not possible to immobilize the europium(III) complex via the ethanol procedure due to extremely poor stability of the complex in ethanol. When immobilized from THF, the dye in the beads was found to remain stable even at pH ∼1, while the indicator in solution is already destroyed at higher pH. Elevating the temperature from 1 to 51 °C results in ∼2-fold decrease of the luminescence decay time (Figure 5). A fit (with a correlation coefficient r2 > 0.9998) is performed using an Arrhenius-type equation:55,56 (55) Coyle, L. M.; Gouterman, M. Sens. Actuators, B 1999, 61, 92-99.
(
(
τ ) k0 + k1 exp -
∆E R(T + 273)
))
-1
(2)
where k0 is the temperature-independent decay rate for the excited-state deactivation, k1 is the pre-exponential factor, ∆E is the energy gap between emitting level and higher excited-state level, and R is the gas constant. The temperature dependence of the decay time (in the absence of oxygen) can be fitted with the following parameters: k0 ) 1491 s-1, k1 ) 1.98 × 1010 s-1, and ∆E ) 43.7 kJ‚mol-1. Since cross-sensitivity to oxygen is rather pronounced, the temperature measurements can be compromised when the oxygen concentration differs significantly. In this case, oxygen concentration should be known, which can be achieved by dispersing oxygen-sensitive nanobeads along with the temperature-sensitive ones. Both can be excited, for example, with a bright 405 nm LED, with emissions separated spectrally or by decay time. As was previously demonstrated for planar sensor foils,44 several iterations are enough to obtain true values of pO2 and temperature. Nanobeads for Sensing pH. The nanosensors for pH make use of the lipophilic indicators CHFOE and HPTS(OA)3. CHFOE incorporated into a polyurethane hydrogel membrane was shown previously42 to be suitable for optical pH sensing at physiological conditions (pKa 6.96). Moreover, ratiometric measurements are possible, since both protonated and deprotonated forms of the pH indicator emit light at different wavelengths (Figure 6). Figure 7a shows dependence of the ratio of fluorescence intensities at 522 and 566 nm (λexc ) 475 nm) on pH at different ionic strengths (IS). Minor dependence is observed at low IS (pKa is 6.2 and 5.9 at IS 0.02 and 0.05 M, respectively); however, there is practically no dependence at higher IS (pKa 5.9 at IS 0.4 M) which is relevant for physiological applications (IS ∼ 0.1 M) and marine systems (IS < 0.6 M). In contrast to many charged polymer matrixes, the PS-PVP is highly promising for designing pH nanosensors, since it makes it possible to significantly reduce cross-sensitivity to IS. It should be mentioned that, unlike other indicator loaded beads, both immobilization methods result in virtually identical results. In fact, when using the THF/H2O procedure, pKa of the beads was measured to be 6.0 at IS 0.05 M, compared to the value of 5.9 for the EtOH/H2O procedure. It is likely that the amphiphilic nature of the indicator (which is composed of a hydrophilic chromophore and a long lipophilic tail) favors its localization in the region of intermediate polarity between the core and shell. It should be considered that the pKa values obtained by ratiometric measurements are only apparent ones because of the fluorescence energy resonance transfer (FRET) which complicates the situation. In fact, the emission spectrum of the acidic form overlaps with the excitation spectrum of the basic form so that FRET becomes increasingly efficient with increasing pH and the fluorescence intensity of the acidic form drops faster. The problem can be of course tackled by preparing beads with much lower amount of the dye (C , 0.6% in the PVP shell), but this will result in significantly lower brightness. More accurate determination of the acidity constant is, however, possible by measuring the fluorescence intensity in the emission maximum of the basic form. Alternatively, the part of the emission from the basic form is (56) Liebsch, G.; Klimant, I.; Wolfbeis, O. S. Adv. Mater. 1999, 11, 1296-1299.
Figure 6. Emission spectra of the pH-sensitive nanobeads N4 (CHFOE from EtOH) at different pH (IS 0.1 M, λexc ) 475 nm).
Figure 7. Calibration curves for the pH-sensitive beads: (a) CHFOE from EtOH (N4); (b) HPTS(OA)3 from THF (N7).
collected using a 560/20 band-pass filter (e.g., in the microplate reader). The pKa values obtained in both methods are essentially the same and are ∼0.4 units higher than those obtained by ratiometric measurements. The calibration curves for the sensor beads N7 (HPTS(OA)3 immobilized via the THF procedure) are presented in Figure 7b. In contrast to the widely used 1-hydroxypyrene-3,6,8-trisulfonate, the sulfonamide derivative bears little negative charge so that the sensor beads exhibit virtually no cross-sensitivity to IS. In fact, the pKa values were found to be 6.53, 6.51, 6.37, and 6.37 for IS 0.02, 0.05, 0.2, and 0.5 M, respectively. The same trend is observed for the sensor beads N6, which are stained from EtOH, and the pKa values are 6.69, 6.57, 6.59, and 6.48 for the same IS. Notably, staining from EtOH/H2O results in brighter beads than staining from THF/H2O. Nevertheless, as in the case of CHFOE both procedures result in rather similar materials in respect to pKa values. Although the sensing materials based on CHFOE and HPTS(OA)3 are suitable for measurements in physiological condition and application in biotechnology, they are not fully adequate for some applications (e.g., for those of marine science). Different substituents were shown to shift the pKa of fluorescein derivatives to higher or lower values.42 These lipophilic indicators can be incorporated into the PS-PVP beads in the same manner as was shown for CHFOE.54 Such beads can be used for pH monitoring in neutral condition, slightly acidic, or slightly basic media. For example, at IS ) 0.2 M the pKa values for the beads stained with Analytical Chemistry, Vol. 80, No. 3, February 1, 2008
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Figure 8. Calibration curves for the nanosensors for Cl- (N8) and Cu2+ (N9).
2′,7′-didodecylfluorescein (from THF) and 2′,7′-dichlorofluorescein octadecylester (from THF) were found to be 7.7 and 5.7, respectively.54 Being based on different fluorescein derivatives, all the beads exhibit virtually identical spectral properties (excitation and emission) but different pKa values. Thus, a homogeneous mixture of the beads can be prepared, which will possibly enable pH monitoring in a wide range, compared to the dynamic range of the pH electrode. Nanobeads For Sensing Ions. Sensing schemes for ionsensitive nanobeads (Cl- and Cu2+) are based on the fluorescence quenching of a fluorescent indicator dye immobilized in the hydrophilic shell of the beads. Lucigenin and lucifer yellow-CH are known to be efficiently quenched by chloride57 and copper(II),58 respectively. The dyes show good solubility in water due to their positively charged amino (lucigenin) or negatively charged sulfo groups (lucifer yellow-CH). The dyes were immobilized by exchanging the counterions with lipophilic counterions (dodecyl sulfate or hexadecyltrimethylamine). This yields ion pairs that are localized in the hydrophilic shell of the beads and do not leach into the aqueous medium. In fact, the reservoir from dialysis was colorless after four cycles indicating that all indicator dye is located in the beads, not in the solvent. Response of the chloride nanosensor (N8) is shown in Figure 8. The titration plot exhibits a typical sigmoidal shape. The dynamic range of the nanosensors is 0.5-100 mM of chloride. The nanosensors respond to a 100 mM chloride solution by a decrease in fluorescence intensity of -80%, and the concentration at which 50% of the fluorescence is quenched (c1/2) is 6.3 ( 0.3 mM. The apparent c(1/2) is ∼3-fold higher than the value found in solution (c(1/2) ) 2.3 mM)59 and ∼4-fold lower than to the value reported for lucigenin immobilized to a hydrogel (c(1/2) ) 25 mM) composed of poly(acrylamide) and poly(acrylonitrile). The more efficient quenching in solution can be attributed to the better accessibility of the quenchers to the fluorophores, whereas the lower c(1/2) compared to that of the hydrogel can be explained by the more hydrophilic environment. The response of copper(II)-sensitive nanospheres (N9) is also shown in Figure 8. The nanosensors respond to a 0.1 and 100 (57) Huber, Ch.; Klimant, I.; Krause, Ch.; Werner, T.; Mayr, T.; Wolfbeis, O. S. Fresenius’ J. Anal. Chem. 2000, 368, 196-202. (58) Mayr, T.; Wencel, D.; Werner, T. Fresenius’ J. Anal. Chem. 2001, 371, 4448. (59) Huber, Ch.; Krause, Ch.; Werner, T.; Wolfbeis, O. S. Microchim. Acta 2003, 142, 245-253.
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Figure 9. Course of the enzymatic oxidation of glucose monitored with help of nanobeads N1 (2 mg/mL) and of a standard sensor foil (PtTFPP in PS, 1.5% w/w, ∼6 µm thick). The concentration of glucose is 0.25 M. An amount of 100 µL of glucose oxidase solution (C ) 10 mg/mL) is added to the test solution (V ) 5 mL). Excitation source: 405 nm LED. The excitation and emission filters are BG 12 and RG 630, respectively. The modulation frequency is 5 kHz. The experiment was performed at room temperature.
mM copper(II) solution by a decrease of fluorescence intensity of -15.1% and -76.6%. The dynamic range is from 0.01 to 100 mM. Similar dynamic ranges were found for nanosensors prepared with various molar ratios of anionic dye and added lipophilic counterion. The dynamic range is 1000-fold higher than that found for dye solutions and sensing membranes based on lucifer yellow immobilized on anion exchanger cellulose. This is attributed to a repulsion of the analyte by an excess of immobilized lipophilic cation. It was also found that if the “sensing chemistries” were incorporated into the core of the beads by using the THF/water procedure, the nanobeads did not exhibit any sensitivity for ions. Response. A very important feature of the nanosensors which makes them advantageous over planar sensor foils and even microsensors includes very short response times. Since the nanobeads introduce virtually no diffusional barrier for the diffusion-limited quenching processes, real time monitoring of fast processes becomes possible. Figure 9 shows the course of enzymatic reaction of glucose oxidation monitored with help of a standard planar sensor foil (PtTFPP in PS, ∼6 µm thick) and using the oxygen-sensitive beads N1. The apparent response of the nanobeads is ∼4 s, which represents, however, the overall time needed for mixing the enzyme and the enzymatic reaction itself. In fact, if a concentrated emulsion of beads is added under vigorous stirring to the anaerobe aqueous solution, the whole signal change occurs in less than 1 s. The response of the pH and ion nanosensors is also very fast and occurs in less than 1 s, which is evidently the time needed for stirring. The nanosensor for temperature responds virtually instantaneously, and so are temperature-sensitive foils, since no diffusion of the analytes is required. Dye Leaching. As was demonstrated above the nanobeads can very easily be stained with the indicators. Since only covalent coupling insures that no leaching occurs, it can be possible for the stained poly(styrene-block-vinylpyrrolidone) nanobeads. The emulsion of the beads (10 mg/mL) was filtered through a fine filter (220 nm pore size), and absorption and emission spectra of the filtrate were recorded. It was found that no leaching was
observed for any of the nanosensors even after 2 months of storage in aqueous solution. We also investigated leaching in the complex (LB) medium, which contains proteins and amino acids and an inorganic salt (NaCl). After 24 h of continuous stirring of the emulsion in the LB medium (1:1 v/v) the filtration was performed. In the case of the oxygen, temperature, and pH nanosensors (N1-N7, stained both from EtOH/H2O and THF/ H2O) no leaching was detected. On the other side, lucifer yellowCH was leached completely out of the Cu2+ nanosensors (N9). It is likely that leaching is promoted by the exchange of hexadecyltrimethylammonium cation in the ion pair by sodium cations. The Cl- nanosensors can evidently not be used in LB medium, and leaching there was therefore not investigated. Interference Tests. Water-soluble indicators are rarely used for sensing purposes in complex media since they tend to absorb on the proteins and other components of the media which can dramatically alter the sensing properties. Apart from those interactions, quenching of luminescence by other components of the medium can occur. These interferences are generally significantly reduced, but not always removed completely, upon immobilization. The nanosensors for oxygen, temperature, and pH were thus investigated in LB medium. In the case of all the nanosensors immobilization in the core via the THF/H2O procedure results in more robust sensors. In fact, at 25 °C τ0 for the N2 nanosensor (PtTFPP from THF/H2O) is 5% lower in the LB medium compared to aqueous emulsion and τ0/τ is 2.7, which is slightly lower than the value obtained in water (τ0/τ is 2.88). Some decrease in decay time can be, however, due to the background fluorescence of the LB medium which is not fully discarded using the emission filters. The LB medium definitely has pronounced effect on the decay time of PtTFPP when the beads are stained from EtOH/H2O. In fact, the decay time drops by 28% in LB medium. The sensitivity also was much lower (τ0/τ was 2.5 and 3.72 for the beads in LB medium and in water, respectively). The data indicate that PtTFPP located in the shell is likely to interact with the components of the medium. In the case of the temperature-sensitive beads N3, the decay times in LB medium are 6% and 11% lower (at 1 and 50 °C, respectively) than in water. Since brightness of the beads increases at lower temperatures, the influence of the LB medium can again be partly cause by background fluorescence. From the pH-sensitive nanobeads only those of type N5 (CHFOE from THF) show identical behavior in aqueous solution and in LB medium. As in the case of the oxygen indicator, immobilization of CHFOE from THF/H2O helps to minimize the interferences. Notable changes occur in the calibration curve for the beads stained from EtOH/H2O (N4), and the pKa value was found to increase by 0.5 units. The calibration plots for the beads based on HPTS(OA)3 were both altered in the LB medium and lost their typical sigmoidal form so that determination of acidity constants became unreliable. Toxicity and Uptake by Cells. We have demonstrated above that the nanosensors for oxygen, temperature, and pH which are obtained by staining from THF/H2O are rather robust to perform adequately even in complex media such as those used in bioreactors. It is, however, also important that the materials exhibit
Figure 10. Microscopic image of the cultivation media containing E. coli (left) and P. pastoris (right) and the beads N3 stained with the temperature indicator (0.5 mg/mL).
no cell toxicity and no uptake by the cell occurs over prolonged time. Therefore, cultivation of E. coli and P. pastoris was performed in the presence of the nanobeads (N3). We have found that the beads in a concentration of 0.5 mg/mL do not influence the growth of the cells. We assume that the nanobeads are not toxic for these robust cells; however, no experiment was performed with more sensitive mammal cells. In addition, we have investigated the possibility of the uptake of the beads by the cells. Figure 10 shows the photographic images obtained with a fluorescence microscope of the cultivation media after 3 and 6 h (in the case of E. coli and P. pastoris, respectively). Evidently, no interaction is visible between the cells and the beads which remain outside of the cell membranes. Similar results were obtained after longer cultivation times (6 and 24 h for E. coli and P. pastoris, respectively). The results confirm that the beads can be successfully used for sensing and imaging in cell cultures. Storage Stability. All the beads can be stored as an aqueous emulsion for at least 3 months (at 4 °C) without alteration of their properties. Freeze-drying can enhance the storage durability. The effect of freeze-drying was investigated for oxygen, pH, and temperature nanosensors. The freeze-dried beads are dispersible in water within 2 min under ultrasonication. The calibration curves for the temperature nanosensor and the oxygen nanosensor N2 (PtTFPP stained from THF/H2O) are not affected by freeze-drying. In the case of the oxygen-sensitive beads obtained from EtOH/ H2O an ∼25% decrease in the decay times (at 0 kPa O2 and at air saturation) was observed after freeze-drying which can be caused by aggregation of the indicator in the dry shell. Freeze-drying has only a minor effect on the properties of the pH-sensitive nanobeads except those based on HPTS(OA)3 stained from THF/ H2O (N7), the brightness of which was reduced significantly after freeze-drying. For the other pH nanosensors, freeze-drying results in the increase of pKa values. In fact, for the nanobeads N4, N5, and N6 the pKa values increased by 0.1, 0.21, and 0.23 units, respectively. The data indicate that all the beads can be freezedried and redispersed without significant alteration of the sensing properties, especially those stained from THF/H2O. Multianalyte Sensing. Above we have shown that fluorescence intensity, fluorescence intensity ratio, and luminescence Analytical Chemistry, Vol. 80, No. 3, February 1, 2008
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decay time (via phase fluorometry) can serve as analytical parameters for sensing with the help of the nanobeads. Ratiometric measurements were demonstrated for the dually emissive pHsensitive probe. In general, ratiometric measurements can be performed using a second (analyte-insensitive) dye; in this case the reference dye and the indicator should possess overlapping absorption spectra to enable their simultaneous excitation but significantly different emission spectra. Note that ratiometric measurements are quite common,25,30 especially when measurements by means of confocal microscopy are performed. The flexibility of the presented material can enable immobilization of the indicator and the reference in different parts of the beads, i.e., into the core and the shell. Alternatively, two types of beads (i.e., sensor beads and reference beads) can be dispersed together. In this case, the ratio of the components can be optimized easily. Another exciting possibility includes multianalyte sensing and imaging using the modified dual lifetime referencing method (mDLR).11,23,60 Particularly, nanosensors with long luminescence decay time (for oxygen or temperature) can be mixed together with fluorescent nanobeads (such as those for pH or ions). Multianalyte sensing with mDLR becomes possible if the two types of beads are excitable at a single wavelength and the emission of both is detected simultaneously at different modulation frequencies.23,60 Because of their small size, the beads result in truly homogeneous emulsions. Since spectral properties of the nanosensors are very different and so are the optical windows required for excitation of the beads and reading of their emission, multianalyte measurements can be performed in another way. A typical example is a mixture of the oxygen-sensitive beads and the temperature-sensitive beads which is suitable for simultaneous determination of both parameters (Figure 11). The emissions can be isolated with two sets of filters, and the luminescence decay times can be detected independently for each indicator. On the other side, no spectral separation of the signals is required using imaging in the time domain since the decay times of both indicators differ significantly and can be resolved. In summary, we have described a novel and versatile platform for designing luminescent nanosensors. The sensing nanobeads can easily be prepared using commercially available poly(styreneblock-vinylpyrrolidone) aqueous emulsion. The “sensor chemistries” can be immobilized either into the core or into the shell of (60) Vasilevska, A. S.; Borisov, S. M.; Krause, Ch.; Wolfbeis, O. S. Chem. Mater. 2006, 18, 4609-4616.
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Figure 11. Emission spectra of the mixture of the oxygen-sensitive beads N2 (C ) 0.66 mg/mL) and the temperature-sensitive beads N3 (C ) 0.167 mg/mL) in aqueous solution (λexc ) 405 nm). All the solutions contained 0.1 M glucose. The deoxygenated solutions additionally contained 0.2 mg/mL glucose oxidase.
the beads by using different staining protocols. The practically uncharged PVP shell allows for the unique stability of the nanosensors which show no tendency to aggregate even when used in high concentration in complex media. It is demonstrated that the obtained materials can be used for optical sensing and imaging of important analytes, such as O2, Cl-, and Cu2+ but also for determination of temperature and pH. Simultaneous sensing of two analytes is also possible. It is shown that the nanosensors for oxygen, temperature, and pH can be used in complex LB medium. The nanosensors can be used for sensing and imaging of even very fast processes in biotechnology, marine science, environmental monitoring, etc. ACKNOWLEDGMENT This work was partly supported by the European Commission Fifth and Sixth Framework programs MAST (N 1645000012) and CLINICIP (EU506965). We thank Dr. Jochen Gerlach from the Research Center of Applied Biocatalysis (Petersgasse 14, 8010 Graz, Austria) for the help in performing the cell penetration tests.
Received for review June 28, 2007. Accepted October 25, 2007. AC071374E
