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Phosphate biosensor based on polyelectrolyte-stabilized pyruvate oxidase
Analytica Chimica Acta 427 (2001) 271–277
Phosphate biosensor based on polyelectrolyte-stabilized pyruvate oxidase Vasilis G. Gavalas, Nikolas A. Chaniotakis∗ Laboratory of Analytical Chemistry, Department of Chemistry, University of Crete, 71409 Iraklion, Crete, Greece Received 27 June 2000; accepted 11 September 2000
Abstract In this work the development of a pyruvate oxidase-based phosphate biosensor is illustrated. The use of polyelectrolyte stabilized recombinant pyruvate oxidase in conjunction with a porous conductive carbon results in the development of a simple, reproducible and stable phosphate biosensor. The polyelectrolyte diethylaminoethyl-dextran or DNA was used as the enzyme stabilizer, and the resulting enzyme–polyelectrolyte complexes were physically adsorbed into the transducer, a highly porous and conductive carbon electrode, for the construction of the biosensor. The optimized biosensor exhibits high operational (67% remaining activity after 220 h) and storage (49% remaining activity after 24 weeks) stability, and very good sensor-to-sensor reproducibility. The optimized phosphate biosensor was used for the measurement of the phosphate ion activity in serum. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Recombinant pyruvate oxidase; Phosphate; Biosensor; Polyelectrolyte; Porous carbon electrode; Serum analysis
1. Introduction The simple and accurate measurement of the activity of the orthophosphate anion is very important in clinical, biological and environmental studies. A sensor system (ISE or biosensor) for the sensitive and selective measurement of this illusive anion is still a challenge [1]. The development of biosensors sensitive to phosphate has attracted many researches over the past 25 years [2]. Amperometric biosensors have been usually constructed based on multi-enzyme schemes: alkaline phosphatase and glucose oxidase [2,3] or polyphenol oxidase [4], nucleoside phosphorylase and xanthine oxidase [5–8], phosphorylase A, phospho∗ Corresponding author. Tel.: +30-81-393618; fax: +30-81-393601. E-mail address: [email protected] (N.A. Chaniotakis).
glucomutase and glucose-6-phosphate dehydrogenase [9,10], maltose phosphorylase and glucose oxidase [11], co-immobilized with phosphatase and muratorase [12] have been employed with various degrees of success. Camman et al. [12] have reported the lowest detection limit (0.01 M) up until now, while the best operational stability (70% remaining activity after 20 h) was reported by Coulet et al. [5] using nucleoside phosphorylase and xanthine oxidase. Additionally the best storage stability (66% remaining activity after 5 weeks dry storage at −20◦ C) was reported by Cosnier et al. [4] using alkaline phosphatase and polyphenol oxidase. The simplest and most efficient approach is achieved with the use of pyruvate oxidase (POD), an enzyme that catalyzes the following reaction Pyruvate + H2 PO4 − + O2 → Acetylphosphate + CO2 + H2 O2
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POD requires the presence of thiamine pyrophosphate (TPP), flavine adenine dinucleotide (FAD) and Mg(II) for the catalysis to take place. These cofactors make the system not very selective, while POD-based biosensors are shown to suffer form low operational and storage stability [13,14]. The absence of phosphate anions from the working and storage buffer that is crucial for the enzyme stability conduces to these problems [13]. In this work, the construction of stable phosphate biosensor based on recombinant pyruvate oxidase stabilized with polyelectrolyte is presented. Pyruvate oxidase from Lactobacillus plantarum contains the cofactors TPP, FAD and Mg(II) bound in the active center [15] and is genetically engineered to enhance its stability [16–18]. Bergmann et al. [19] have utilized this enzyme in conjunction with horseradish peroxidase for the construction of a bienzyme modified carbon paste biosensor for the measurement of pyruvate. In this work, the stabilized pyruvate oxidase is immobilized by physical adsorption into a highly porous and conductive carbon electrode. This material has been used with great success for the construction of highly stable and reproducible glucose and lactate biosensors [22–24]. The polyelectrolytes diethylaminoethyl-dextran and DNA are employed during the immobilization procedure to stabilize POD via a cage formation around the POD. The use of polyelectrolytes such as diethylaminoethyl-dextran or polyethylenimine (positively charged macromolecules) have been proved to stabilize enzymes [19–27] in various environments. It has been shown that these polyelectrolytes form a cage-like stable microenvironment around the enzyme via electrostatic interactions. Recently, DNA (a negatively charged macromolecule) was shown to enhance the stability of certain biosensors [28]. Here, the effects of the type and the amount of stabilizer used during the construction of the POD-based biosensor on the characteristics of the sensor are evaluated, and an optimized system is constructed. The optimized biosensor is then incorporated in a flow system for the measurement of phosphate levels in serum. The usually occurring serum interferences from the easily electro-oxidazible species are eliminated using an efficient preoxidizing cell [29], allowing for the fast and accurate measurements of phosphate in this complicated matrix.
2. Experimental 2.1. Materials-reagents Recombinant pyruvate oxidase (POD) from Lactobacillus plantarum (EC 1.2.3.3, 1.66 U/mg), available from Roche Diagnostics GmbH (Germany), was kindly provided by Dr. Spohn (University of Halle). Diethylaminoethyl-dextran (deae-dextran) (D-9885) and DNA sodium salt (D-1626) were obtained from Sigma. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), biochemica grade was purchased from Fluka. The porous carbon was obtained from BF HIRM (Austria). Vitreous carbon foam (bulk density 0.05 g/cm3 , porosity 96.5%), carbon fabric (weight/ m2 200 g, warp and weft yarn 200 Tex, plain weave) and platinum mesh (wire diameter 0.06 mm, wires/inch 82 × 82) were purchased from Goodfellow Cambridge Ltd. In all experiments nano-pure water (∼18 M, EASYpure model D7033, Barnstead) was used. All other reagents used were of analytical grade. 2.2. Biosensor construction A rod (2.1 mm diameter, 4.0 mm height) of the porous carbon was cleaned in a sonicated ethanol bath for 10 min and then in water bath for 10 min, followed by oven drying at 150◦ C for 30 min. Pyruvate oxidase was dissolved (to a final concentration 100 U/ml) in 10 mM HEPES buffer (pH 7.5) containing the desired amount of the stabilizer. The enzyme was allowed to react with the polyelectrolyte for 20 min at +4◦ C. After this, the cleaned and dried carbon rod was placed into the enzyme solution for 20 h at +4◦ C. The carbon rod was then removed from the solution, washed thoroughly with buffer, and placed in a testing holder. It is estimated, based on the amount of solution adsorbed by the carbon, that 1 unit per carbon rod is immobilized. The electrical contact was achieved from the backside through a platinum wire. When not used the biosensors were kept at +4◦ C in HEPES buffer. 2.3. Pre-oxidizing cell The pre-oxidizing cell used for the blood measurements was slightly modified from the one previously described [29]. The cell consists of two separate parts,
V.G. Gavalas, N.A. Chaniotakis / Analytica Chimica Acta 427 (2001) 271–277
the oxidation and the reduction part connected to a dc power supply. The constant potential method at an operational potential above that required to oxidize all interfering species, but bellow that for the oxidation of H2 O is chosen. Each part has separate inlet and outlet of the flow sample. The inlet of the oxidation part is connected to the injection valve so that the sample after the injection will pass only through this part of the cell. The outlet of the oxidation part (anode) of the cell is connected to the wall-jet flow cell where the biosensor is placed. The inlet of the reducing part (cathode) of the cell is connected to a reservoir containing 0.1 M KCl while the outlet is connected to the wastes. The electrical contact of the two half-cells is achieved through a cellulose acetate membrane (12– 16 kDa cut-off) placed between the upper side of the vitreous carbon foam and the lower side of the platinum mesh. The reactor where the oxidation takes place is the lower part of the cell, which contains the vitreous carbon foam (6.3 cm × 0.6 cm × 0.5 cm). It is composed of a polymethylmethacrylate (PMMA) plate (8.3 cm × 4.0 cm × 1.0 cm) where a hole in the dimensions of the foam is made. To assure homogeneous distribution of the potential throughout the carbon without any IR drop, the electrical contact to the power supply is achieved from the lower side of the carbon plate using carbon fabric. The cathode compartment (also made by PMMA 8.3 cm × 4.0 cm × 1.0 cm) contains the platinum mesh (3.5 × 0.6). The outlet of this part of the cell is designed so that the hydrogen bubbles generated during the experiment will be quickly removed from the platinum surface.
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The flow injection system consisted of a wall-jet flow cell, an injection valve with loop volume 400 l, while the solvent delivery was done using a peristaltic pump (No 7554-30, Cole-Palmer). The response is calculated as the peak height from the baseline.
3. Results and discussion A detailed experiment in order to obtain the optimum operational pH for the phosphate biosensor reveals that the pH profile of the sensor is bell shaped (data not shown), with optimum value at around pH 7.5. This is slightly shifted to basic pH compared with the POD pH profile that has been shown to be at pH 7.0 [30]. Further experiments were performed using 10 mM HEPES buffer adjusted to pH 7.5. 3.1. Effect of pyruvate concentration From Eq. (1) it is clear that pyruvate, a co-substrate in the enzymatic reaction, is required for the proper operation of the system, and the measurement of phosphate. The optimum amount of pyruvate was determined by varying its concentration in the carrier stream from 0.1–10 mM, while monitoring its effect on the sensitivity and detection limit as determined from the calibration curve obtained for each pyruvate concentration. The results shown in Fig. 1 indicate that an increase in the pyruvate concentration up to 1.0 mM results in an increase in the sensitivity (linear
2.4. Electrochemical measurements In all experiments a three electrode Metrohm 641 VA-Detector, a silver/silver chloride double junction reference electrode (Model 90-02, ORION Res. Inc.), and a platinum counter electrode (CORNING, cat. no. 476060) were used. The working potential was set at +800 mV in all experiments, except from the operational stability where the sensors were polarized at +600 mV versus Ag/AgCl. The signal was recorded via a personal computer equipped with a 16-bit A/D converter and controlled with software written in BASIC. Temperature control at 24.0±0.1◦ C was achieved with a circulating bath (Model 9105, PolyScience).
Fig. 1. Effect of pyruvate concentration, incorporated in the HEPES buffer, on the sensitivity (䊏) and the detection limit (䊊) of the phosphate biosensor.
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range 0.05–1.0 mM phosphate). Further increase in the pyruvate concentration results in lower sensitivities. Even though this decrease of the sensitivity was also observed by Kubo et al. [14], the large negative slope was not expected. Based on this observation, the detection limit of the sensor might also be influenced by the background pyruvate concentration. When the pyruvate concentration is increased from 0.1 up to 0.5 mM, there is no influence on the lower detection limit. When the concentration of pyruvate is increased from 0.5 to 10 mM the lower detection limit also increase linearly. The fact that the sensitivities obtained with 5 or 10 mM pyruvate were similar indicates that at these pyruvate concentrations the rate limiting substrate is oxygen. From these results the 1 mM pyruvate was chosen to be used for all further experiments. 3.2. Effect of polyelectrolyte The effect of the amount and type of polyelectrolyte on the phosphate biosensor was then investigated. Sensors were prepared from solutions containing various amounts of deae-dextran or DNA. Deae-dextran was varied from 0.1 to 0.5%, w/v, while two different amounts of DNA were tested (0.1 and 0.25%, w/v). Higher amount of DNA resulted in the formation of a suspension, preventing the construction of the sensor. To monitor the effect of the amount and type of polyelectrolyte, the sensors were periodically tested. Between the test times, all sensors were stored at 4◦ C in buffer solution (10 mM HEPES, pH 7.5). Table 1 summarizes the results of the sensitivity and the remaining activity of the sensors, 1 month after their construction. From this data it is clear that both polyelectrolytes increase the stability of the biosensor. It is interesting to note that when the amount of deae-dextran used is in-
creased beyond the level of 0.25%, w/v the stabilizing effect is eliminated and the sensor loses its sensitivity even faster than the control. The reduced biosensor’s stability at higher amounts of deae-dextran has been previously observed with other biosensors [22–24] and has been attributed to the reduction of enzyme activity at high polyelectrolyte concentrations. Additionally, it is shown that DNA has a smaller stabilizing effect on the sensor. When 0.1%, w/v DNA is used the initial sensor’s sensitivity is smaller than the control, and the stability of the sensor is slightly reduced. Increasing the concentration to 0.25%, w/v, the stability of the sensor is improved over that of the control, but to a lesser extent from the deae-dextran stabilized sensor. Based on these results sensors 1, 2 and 6 were chosen for further evaluation of their operational stability. For this experiment the sensors were incorporated in a flow system and were continuously polarized at +600 mV versus Ag/AgCl. The flow rate was 0.65 ml/min, the temperature was held constant at 24.0 ± 0.1◦ C and the working solution was 10 mM HEPES, 1 mM pyruvate and 1 mM phosphate adjusted to pH 7.5. Under these conditions (buffer stream containing the required substrates, pyruvate and phosphate), there is a continuous enzymatic catalysis. The system was calibrated daily, and the sensitivity was used as a measure to evaluate the activity of the sensor. For the calibration procedure, the phosphate was removed from the buffer stream and after equilibration different concentrations of phosphate were injected. Table 2 shows the remaining activity of the three sensors under the continuous operation test. It is interesting to note that the operational stability of all the sensors monitored is better than any such sensors presented so far in the literature (70% remaining activity after 20 h [5]). Moreover, the operational stability of
Table 1 Effect of polyelectrolyte type and concentration on the sensitivity and remain activity of the phosphate biosensor
1 2 3 4 5 6 a
Polyelectrolyte
Concentration (%, w/v)
Sensitivitya (A/mM)
– Deae-dextran Deae-dextran Deae-dextran DNA DNA
– 0.10 0.25 0.50 0.10 0.25
5.56 7.94 5.88 5.31 3.00 6.15
One month after construction.
± ± ± ± ± ±
0.06 0.12 0.14 0.08 0.03 0.12
Remaining activitya (%) 51.8 77.1 68.0 43.4 49.6 57.5
V.G. Gavalas, N.A. Chaniotakis / Analytica Chimica Acta 427 (2001) 271–277 Table 2 Operational stability of the 1, 2 and 6 phosphate biosensors Hours polarized
20 40 60 80 100 120 140 220
Remaining activity (%) 1
2
6
77 72 72 69 65 61 59 51
114 106 97 86 79 73 71 67
104 94 89 84 79 72 69 56
the sensor, 2 is exceptional, with 67% remaining activity after 220 h. DNA stabilized sensor, 6 has 56% and the control sensor has 50% remaining activity. 3.3. Characterization of the phosphate biosensor A batch of the most stable sensors (2) was then prepared. Three of these sensors were used to study their analytical characteristics, and the remaining were freeze dried and stored at 4◦ C for storage stability measurements. At the end of each month one sensor was rehydrated for 20 h and then its residual activity was calculated from the sensitivity obtained from the calibration curve. In Fig. 2, the initial average calibration curve of the three freshly prepared sensors is presented. The
Fig. 2. Calibration curve of the phosphate biosensor in 10 mM HEPES, 1 mM pyruvate buffer adjusted at pH 7.5. The insert presents the linear part of the curve. The working potential was +800 mV vs. Ag/AgCl.
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sensitivity is 10.3 ± 0.2 A/mM and the response is linear to phosphate concentrations from 50 to 1250 M. The limit of detection for phosphate based on a signal-to-noise ratio of three is 4.8 M. The response time varies between 20 and 40 s, depending on the concentration of the analyte. The sensor-to-sensor reproducibility based on the sensitivities is 2.2% RSD, while based on the response to 1 mM phosphate it is 7.6% RSD. These results indicate that the method developed enables the construction of very reproducible sensors. The storage stability of the phosphate biosensor was then examined, using the calibration curve of the fresh sensors as the reference curve. These studies indicate that there is a 84 ± 5% residual activity after 2 months of dry storage at 4◦ C, while after 6 months the residual activity is reduced to 49 ± 4%. 3.4. Application to serum analysis A key factor for the application of an amperometric sensor in real life samples is its selectivity towards the easily electro-oxidazible species. In the case of serum, ascorbic acid, uric acid, acetaminophen and many other compounds can interfere. The porous carbon used here for the construction of the phosphate sensor has shown to be sensitive to these species (between 10 and 13 A/mM), preventing its application to serum analysis. For this reason a pre-oxidizing cell [29] was incorporated in the FIA system, to oxidize these species before they reach the electrode surface. Initial experiments indicated the need for the buffer to be mixed with the sample after the pre-oxidation step. When the buffer was introduced prior to the cell the response of the sensor was not reproducible. The final experimental set-up is shown in Fig. 3. The concentration of pyruvate in blood varies considerably (70–150 M) [31]. Unless there is a high and constant concentration of pyruvate in the buffer stream, this fluctuation will introduce an error to the biosensor’s response. Therefore, the concentration of pyruvate in the buffer stream was raised from 1 to 10 mM. At the same time, the concentration of HEPES in the buffer stream was also raised in order to increase its buffer capacity. Thus, the buffer stream was 50 mM HEPES and 10 mM pyruvate adjusted to pH 7.5 at a flow rate of 0.65 ml/min and the flow rate of the sample was also 0.65 ml/min.
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Fig. 3. Diagram of the flow injection system. The buffer consists of 50 mM HEPES and 10 mM pyruvate adjust to pH 7.5. The sample passes through the pre-oxidizing cell where the interfering species are electrooxidized, then is mixed with the buffer and reach the biosensor to complete the analysis.
The analytical characteristics of the phosphate biosensor incorporated in the flow injection system are as follows. The sensitivity is 445 ± 9 nA/mM, the response is linear from 0.5 to 10 mM, while the detection limit is less than 0.3 mM. In Fig. 4, a typical recording of phosphate injections is presented. Prior to real sample analyses, the efficiency of the cell in the elimination of interferents was then investigated. With the cell current off, the phosphate biosensor exhibits sensitivities 1.72, 2.27 and 1.63 A/mM
for ascorbate, acetaminophen and urate, respectively. When the cell current is on, all sensitivities are reduced to 0.26, 0.30 and 0.51 A/mM, respectively. With these sensitivities the detection limit of the system for these species is 362, 853 and 816 M for ascorbate, acetaminophen and urate, respectively, values that are much higher than their normal concentration in serum. The above experimental set-up was then used to determine the phosphate level in serum. Three samples of serum were analyzed and the results were compared with those obtained with ion-chromatography. The samples were injected in the system without dilution and the calculated concentration was found to be 1.03 ± 0.06 mM. The results from the chromatographic analysis were 0.99 ± 0.02 mM, very close to the results obtained with the biosensor.
4. Conclusions
Fig. 4. Recording of the response of the biosensor incorporated in the flow injection system shown in Fig. 3 to phosphate injections (2.5, 5.0, 7.5 and 10 mM phosphate, respectively). The sensor is polarized at +800 mV vs. Ag/AgCl. The flow rate is 0.65 ml/min and the buffer consist of 50 mM HEPES and 10 mM pyruvate adjusted to pH 7.5.
The development and evaluation of a phosphate biosensor based on stabilized with polyelectrolytes recombinant pyruvate oxidase adsorbed into a porous carbon has been presented. Both electrolytes used were shown to have a positive effect on both the operational and storage stability of the resulting biosensor, with 0.1%, w/v deae-dextran being the optimum. The phosphate biosensor exhibits exceptional operational and storage stability while the simple construction
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procedure results in a very good sensor-to-sensor reproducibility. The use of the sensor in an FIA system equipped with a pre-oxidizing cell allows for the measurement of the activity of phosphate in difficult samples such as those of human serum with results in good agreement with those obtained with ion-chromatography.
Acknowledgements We would like to thank, Dr. U. Spohn (University of Hall) for supplying the enzyme, G. Kouvarakis (ECPL, University of Crete) for the ion-chromatography measurements and the Greek Secretary General of Research and Development for financial support through the YPER program (97 YPER-219). References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10]
S.O. Engblom, Biosens. Bioelectron. 13 (1998) 981. G.G. Guilbault, M. Nanjo, Anal. Chim. Acta 78 (1975) 69. Y. Su, M. Mascini, Anal. Lett. 28 (1995) 1359. S. Cosnier, C. Gondran, J.-C. Watelet, W. De Giovani, R.P.M. Furriel, F.A. Leone, Anal. Chem. 70 (1998) 3952. E.M. d’Urso, P.R. Coulet, Anal. Chim. Acta 239 (1990) 1. E.M. d’Urso, P.R. Coulet, Anal. Chim. Acta 281 (1993) 535. S.D. Haemmerli, A.A. Suleiman, G.G. Guilbault, Anal. Biochem. 191 (1990) 106. U. Wollenberger, F. Schubert, F.W. Scheller, Sens. Act. B 7 (1992) 412. J.J. Fernandez, J.R. Lopez, X. Correig, I. Katakis, Sens. Act. B 47 (1998) 13. J. Parellada, A. Narvaez, M.A. Lopez, E. Dominguez, J.J. Fernandez, V. Pavlov, I. Katakis, Anal. Chim. Acta 362 (1998) 47.
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[11] S. Huwel, L. Haalck, N. Conrath, F. Spener, Enz. Microb. Technol. 21 (1997) 413. [12] N. Conrath, B. Grundig, S. Huwel, K. Cammann, Anal. Chim. Acta 309 (1995) 47. [13] N. Gajovic, K. Habermuller, A. Warsinke, W. Schuhmann, F.W. Scheller, Electroanalysis 11 (1999) 1377. [14] I. Kubo, M. Inagawa, T. Sugawara, Y. Arikawa, I. Karube, Anal. Lett. 24 (1991) 1711. [15] Y. Muller, G. Schumacher, R. Rudolph, G. Schulz, J. Mol. Biol. 237 (1994) 315. [16] E. Kopetzki, K. Lehnert, P. Buckel, Clin. Chem. 40 (1994) 688. [17] B. Risse, G. Stempfer, R. Rudolph, H. Mollering, R. Jaenicke, Prot. Sci. 1 (1992) 1699. [18] B. Risse, G. Stempfer, R. Rudolph, G. Schumacher, R. Jaenicke, Prot. Sci. 1 (1992) 1710. [19] W. Bergmann, R. Rudolph, U. Spohn, Anal. Chim. Acta 394 (1999) 233. [20] A. Appleton, T.D. Gibson, J.R. Woodward, Sens. Act. B 43 (1997) 65. [21] T.D. Gibson, J.R. Woodward, in: P.G. Eldman, J. Wang (Eds.), Biosensors and Chemical Sensors, ACS Books, 1992, p. 40. [22] V.G. Gavalas, N.A. Chaniotakis, T.D. Gibson, Biosens. Bioelectron. 13 (1998) 1205. [23] V.G. Gavalas, N.A. Chaniotakis, Anal. Chim. Acta 404 (2000) 67. [24] V.G. Gavalas, N.A. Chaniotakis, Mikrochim. Acta, submitted for publication. [25] U. Spohn, D. Narasaiah, L. Gorton, Electroanalysis 8 (1996) 507. [26] H. Liden, J. Volc, G. Marko-Varga, L. Gorton, Electroanalysis 10 (1998) 223. [27] M. Tessema, E. Csoregi, T. Ruzgas, G. Kenausis, T. Solomon, L. Gorton, Anal. Chem. 69 (1997) 4039. [28] P. Dantoni, S.H.P. Serrano, A.M.O. Brett, I.G.R. Gutz, Anal. Chim. Acta 366 (1998) 137. [29] V.G. Gavalas, M.G. Fouskaki, N.A. Chaniotakis, Anal. Lett. 33 (2000) 2391. [30] Roche Molecular Biochemicals 1999/2000. [31] R.K. Murray, D.K. Granner, P.A. Mayes, V.W. Rodwell, Harper’s Biochemistry, Appleton & Lange, 1996, p. 829.
Phosphate biosensor based on polyelectrolyte-stabilized pyruvate oxidase Vasilis G. Gavalas, Nikolas A. Chaniotakis∗ Laboratory of Analytical Chemistry, Department of Chemistry, University of Crete, 71409 Iraklion, Crete, Greece Received 27 June 2000; accepted 11 September 2000
Abstract In this work the development of a pyruvate oxidase-based phosphate biosensor is illustrated. The use of polyelectrolyte stabilized recombinant pyruvate oxidase in conjunction with a porous conductive carbon results in the development of a simple, reproducible and stable phosphate biosensor. The polyelectrolyte diethylaminoethyl-dextran or DNA was used as the enzyme stabilizer, and the resulting enzyme–polyelectrolyte complexes were physically adsorbed into the transducer, a highly porous and conductive carbon electrode, for the construction of the biosensor. The optimized biosensor exhibits high operational (67% remaining activity after 220 h) and storage (49% remaining activity after 24 weeks) stability, and very good sensor-to-sensor reproducibility. The optimized phosphate biosensor was used for the measurement of the phosphate ion activity in serum. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Recombinant pyruvate oxidase; Phosphate; Biosensor; Polyelectrolyte; Porous carbon electrode; Serum analysis
1. Introduction The simple and accurate measurement of the activity of the orthophosphate anion is very important in clinical, biological and environmental studies. A sensor system (ISE or biosensor) for the sensitive and selective measurement of this illusive anion is still a challenge [1]. The development of biosensors sensitive to phosphate has attracted many researches over the past 25 years [2]. Amperometric biosensors have been usually constructed based on multi-enzyme schemes: alkaline phosphatase and glucose oxidase [2,3] or polyphenol oxidase [4], nucleoside phosphorylase and xanthine oxidase [5–8], phosphorylase A, phospho∗ Corresponding author. Tel.: +30-81-393618; fax: +30-81-393601. E-mail address: [email protected] (N.A. Chaniotakis).
glucomutase and glucose-6-phosphate dehydrogenase [9,10], maltose phosphorylase and glucose oxidase [11], co-immobilized with phosphatase and muratorase [12] have been employed with various degrees of success. Camman et al. [12] have reported the lowest detection limit (0.01 M) up until now, while the best operational stability (70% remaining activity after 20 h) was reported by Coulet et al. [5] using nucleoside phosphorylase and xanthine oxidase. Additionally the best storage stability (66% remaining activity after 5 weeks dry storage at −20◦ C) was reported by Cosnier et al. [4] using alkaline phosphatase and polyphenol oxidase. The simplest and most efficient approach is achieved with the use of pyruvate oxidase (POD), an enzyme that catalyzes the following reaction Pyruvate + H2 PO4 − + O2 → Acetylphosphate + CO2 + H2 O2
0003-2670/01/$ – see front matter © 2001 Elsevier Science B.V. All rights reserved. PII: S 0 0 0 3 - 2 6 7 0 ( 0 0 ) 0 1 2 0 4 - 6
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POD requires the presence of thiamine pyrophosphate (TPP), flavine adenine dinucleotide (FAD) and Mg(II) for the catalysis to take place. These cofactors make the system not very selective, while POD-based biosensors are shown to suffer form low operational and storage stability [13,14]. The absence of phosphate anions from the working and storage buffer that is crucial for the enzyme stability conduces to these problems [13]. In this work, the construction of stable phosphate biosensor based on recombinant pyruvate oxidase stabilized with polyelectrolyte is presented. Pyruvate oxidase from Lactobacillus plantarum contains the cofactors TPP, FAD and Mg(II) bound in the active center [15] and is genetically engineered to enhance its stability [16–18]. Bergmann et al. [19] have utilized this enzyme in conjunction with horseradish peroxidase for the construction of a bienzyme modified carbon paste biosensor for the measurement of pyruvate. In this work, the stabilized pyruvate oxidase is immobilized by physical adsorption into a highly porous and conductive carbon electrode. This material has been used with great success for the construction of highly stable and reproducible glucose and lactate biosensors [22–24]. The polyelectrolytes diethylaminoethyl-dextran and DNA are employed during the immobilization procedure to stabilize POD via a cage formation around the POD. The use of polyelectrolytes such as diethylaminoethyl-dextran or polyethylenimine (positively charged macromolecules) have been proved to stabilize enzymes [19–27] in various environments. It has been shown that these polyelectrolytes form a cage-like stable microenvironment around the enzyme via electrostatic interactions. Recently, DNA (a negatively charged macromolecule) was shown to enhance the stability of certain biosensors [28]. Here, the effects of the type and the amount of stabilizer used during the construction of the POD-based biosensor on the characteristics of the sensor are evaluated, and an optimized system is constructed. The optimized biosensor is then incorporated in a flow system for the measurement of phosphate levels in serum. The usually occurring serum interferences from the easily electro-oxidazible species are eliminated using an efficient preoxidizing cell [29], allowing for the fast and accurate measurements of phosphate in this complicated matrix.
2. Experimental 2.1. Materials-reagents Recombinant pyruvate oxidase (POD) from Lactobacillus plantarum (EC 1.2.3.3, 1.66 U/mg), available from Roche Diagnostics GmbH (Germany), was kindly provided by Dr. Spohn (University of Halle). Diethylaminoethyl-dextran (deae-dextran) (D-9885) and DNA sodium salt (D-1626) were obtained from Sigma. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), biochemica grade was purchased from Fluka. The porous carbon was obtained from BF HIRM (Austria). Vitreous carbon foam (bulk density 0.05 g/cm3 , porosity 96.5%), carbon fabric (weight/ m2 200 g, warp and weft yarn 200 Tex, plain weave) and platinum mesh (wire diameter 0.06 mm, wires/inch 82 × 82) were purchased from Goodfellow Cambridge Ltd. In all experiments nano-pure water (∼18 M, EASYpure model D7033, Barnstead) was used. All other reagents used were of analytical grade. 2.2. Biosensor construction A rod (2.1 mm diameter, 4.0 mm height) of the porous carbon was cleaned in a sonicated ethanol bath for 10 min and then in water bath for 10 min, followed by oven drying at 150◦ C for 30 min. Pyruvate oxidase was dissolved (to a final concentration 100 U/ml) in 10 mM HEPES buffer (pH 7.5) containing the desired amount of the stabilizer. The enzyme was allowed to react with the polyelectrolyte for 20 min at +4◦ C. After this, the cleaned and dried carbon rod was placed into the enzyme solution for 20 h at +4◦ C. The carbon rod was then removed from the solution, washed thoroughly with buffer, and placed in a testing holder. It is estimated, based on the amount of solution adsorbed by the carbon, that 1 unit per carbon rod is immobilized. The electrical contact was achieved from the backside through a platinum wire. When not used the biosensors were kept at +4◦ C in HEPES buffer. 2.3. Pre-oxidizing cell The pre-oxidizing cell used for the blood measurements was slightly modified from the one previously described [29]. The cell consists of two separate parts,
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the oxidation and the reduction part connected to a dc power supply. The constant potential method at an operational potential above that required to oxidize all interfering species, but bellow that for the oxidation of H2 O is chosen. Each part has separate inlet and outlet of the flow sample. The inlet of the oxidation part is connected to the injection valve so that the sample after the injection will pass only through this part of the cell. The outlet of the oxidation part (anode) of the cell is connected to the wall-jet flow cell where the biosensor is placed. The inlet of the reducing part (cathode) of the cell is connected to a reservoir containing 0.1 M KCl while the outlet is connected to the wastes. The electrical contact of the two half-cells is achieved through a cellulose acetate membrane (12– 16 kDa cut-off) placed between the upper side of the vitreous carbon foam and the lower side of the platinum mesh. The reactor where the oxidation takes place is the lower part of the cell, which contains the vitreous carbon foam (6.3 cm × 0.6 cm × 0.5 cm). It is composed of a polymethylmethacrylate (PMMA) plate (8.3 cm × 4.0 cm × 1.0 cm) where a hole in the dimensions of the foam is made. To assure homogeneous distribution of the potential throughout the carbon without any IR drop, the electrical contact to the power supply is achieved from the lower side of the carbon plate using carbon fabric. The cathode compartment (also made by PMMA 8.3 cm × 4.0 cm × 1.0 cm) contains the platinum mesh (3.5 × 0.6). The outlet of this part of the cell is designed so that the hydrogen bubbles generated during the experiment will be quickly removed from the platinum surface.
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The flow injection system consisted of a wall-jet flow cell, an injection valve with loop volume 400 l, while the solvent delivery was done using a peristaltic pump (No 7554-30, Cole-Palmer). The response is calculated as the peak height from the baseline.
3. Results and discussion A detailed experiment in order to obtain the optimum operational pH for the phosphate biosensor reveals that the pH profile of the sensor is bell shaped (data not shown), with optimum value at around pH 7.5. This is slightly shifted to basic pH compared with the POD pH profile that has been shown to be at pH 7.0 [30]. Further experiments were performed using 10 mM HEPES buffer adjusted to pH 7.5. 3.1. Effect of pyruvate concentration From Eq. (1) it is clear that pyruvate, a co-substrate in the enzymatic reaction, is required for the proper operation of the system, and the measurement of phosphate. The optimum amount of pyruvate was determined by varying its concentration in the carrier stream from 0.1–10 mM, while monitoring its effect on the sensitivity and detection limit as determined from the calibration curve obtained for each pyruvate concentration. The results shown in Fig. 1 indicate that an increase in the pyruvate concentration up to 1.0 mM results in an increase in the sensitivity (linear
2.4. Electrochemical measurements In all experiments a three electrode Metrohm 641 VA-Detector, a silver/silver chloride double junction reference electrode (Model 90-02, ORION Res. Inc.), and a platinum counter electrode (CORNING, cat. no. 476060) were used. The working potential was set at +800 mV in all experiments, except from the operational stability where the sensors were polarized at +600 mV versus Ag/AgCl. The signal was recorded via a personal computer equipped with a 16-bit A/D converter and controlled with software written in BASIC. Temperature control at 24.0±0.1◦ C was achieved with a circulating bath (Model 9105, PolyScience).
Fig. 1. Effect of pyruvate concentration, incorporated in the HEPES buffer, on the sensitivity (䊏) and the detection limit (䊊) of the phosphate biosensor.
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range 0.05–1.0 mM phosphate). Further increase in the pyruvate concentration results in lower sensitivities. Even though this decrease of the sensitivity was also observed by Kubo et al. [14], the large negative slope was not expected. Based on this observation, the detection limit of the sensor might also be influenced by the background pyruvate concentration. When the pyruvate concentration is increased from 0.1 up to 0.5 mM, there is no influence on the lower detection limit. When the concentration of pyruvate is increased from 0.5 to 10 mM the lower detection limit also increase linearly. The fact that the sensitivities obtained with 5 or 10 mM pyruvate were similar indicates that at these pyruvate concentrations the rate limiting substrate is oxygen. From these results the 1 mM pyruvate was chosen to be used for all further experiments. 3.2. Effect of polyelectrolyte The effect of the amount and type of polyelectrolyte on the phosphate biosensor was then investigated. Sensors were prepared from solutions containing various amounts of deae-dextran or DNA. Deae-dextran was varied from 0.1 to 0.5%, w/v, while two different amounts of DNA were tested (0.1 and 0.25%, w/v). Higher amount of DNA resulted in the formation of a suspension, preventing the construction of the sensor. To monitor the effect of the amount and type of polyelectrolyte, the sensors were periodically tested. Between the test times, all sensors were stored at 4◦ C in buffer solution (10 mM HEPES, pH 7.5). Table 1 summarizes the results of the sensitivity and the remaining activity of the sensors, 1 month after their construction. From this data it is clear that both polyelectrolytes increase the stability of the biosensor. It is interesting to note that when the amount of deae-dextran used is in-
creased beyond the level of 0.25%, w/v the stabilizing effect is eliminated and the sensor loses its sensitivity even faster than the control. The reduced biosensor’s stability at higher amounts of deae-dextran has been previously observed with other biosensors [22–24] and has been attributed to the reduction of enzyme activity at high polyelectrolyte concentrations. Additionally, it is shown that DNA has a smaller stabilizing effect on the sensor. When 0.1%, w/v DNA is used the initial sensor’s sensitivity is smaller than the control, and the stability of the sensor is slightly reduced. Increasing the concentration to 0.25%, w/v, the stability of the sensor is improved over that of the control, but to a lesser extent from the deae-dextran stabilized sensor. Based on these results sensors 1, 2 and 6 were chosen for further evaluation of their operational stability. For this experiment the sensors were incorporated in a flow system and were continuously polarized at +600 mV versus Ag/AgCl. The flow rate was 0.65 ml/min, the temperature was held constant at 24.0 ± 0.1◦ C and the working solution was 10 mM HEPES, 1 mM pyruvate and 1 mM phosphate adjusted to pH 7.5. Under these conditions (buffer stream containing the required substrates, pyruvate and phosphate), there is a continuous enzymatic catalysis. The system was calibrated daily, and the sensitivity was used as a measure to evaluate the activity of the sensor. For the calibration procedure, the phosphate was removed from the buffer stream and after equilibration different concentrations of phosphate were injected. Table 2 shows the remaining activity of the three sensors under the continuous operation test. It is interesting to note that the operational stability of all the sensors monitored is better than any such sensors presented so far in the literature (70% remaining activity after 20 h [5]). Moreover, the operational stability of
Table 1 Effect of polyelectrolyte type and concentration on the sensitivity and remain activity of the phosphate biosensor
1 2 3 4 5 6 a
Polyelectrolyte
Concentration (%, w/v)
Sensitivitya (A/mM)
– Deae-dextran Deae-dextran Deae-dextran DNA DNA
– 0.10 0.25 0.50 0.10 0.25
5.56 7.94 5.88 5.31 3.00 6.15
One month after construction.
± ± ± ± ± ±
0.06 0.12 0.14 0.08 0.03 0.12
Remaining activitya (%) 51.8 77.1 68.0 43.4 49.6 57.5
V.G. Gavalas, N.A. Chaniotakis / Analytica Chimica Acta 427 (2001) 271–277 Table 2 Operational stability of the 1, 2 and 6 phosphate biosensors Hours polarized
20 40 60 80 100 120 140 220
Remaining activity (%) 1
2
6
77 72 72 69 65 61 59 51
114 106 97 86 79 73 71 67
104 94 89 84 79 72 69 56
the sensor, 2 is exceptional, with 67% remaining activity after 220 h. DNA stabilized sensor, 6 has 56% and the control sensor has 50% remaining activity. 3.3. Characterization of the phosphate biosensor A batch of the most stable sensors (2) was then prepared. Three of these sensors were used to study their analytical characteristics, and the remaining were freeze dried and stored at 4◦ C for storage stability measurements. At the end of each month one sensor was rehydrated for 20 h and then its residual activity was calculated from the sensitivity obtained from the calibration curve. In Fig. 2, the initial average calibration curve of the three freshly prepared sensors is presented. The
Fig. 2. Calibration curve of the phosphate biosensor in 10 mM HEPES, 1 mM pyruvate buffer adjusted at pH 7.5. The insert presents the linear part of the curve. The working potential was +800 mV vs. Ag/AgCl.
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sensitivity is 10.3 ± 0.2 A/mM and the response is linear to phosphate concentrations from 50 to 1250 M. The limit of detection for phosphate based on a signal-to-noise ratio of three is 4.8 M. The response time varies between 20 and 40 s, depending on the concentration of the analyte. The sensor-to-sensor reproducibility based on the sensitivities is 2.2% RSD, while based on the response to 1 mM phosphate it is 7.6% RSD. These results indicate that the method developed enables the construction of very reproducible sensors. The storage stability of the phosphate biosensor was then examined, using the calibration curve of the fresh sensors as the reference curve. These studies indicate that there is a 84 ± 5% residual activity after 2 months of dry storage at 4◦ C, while after 6 months the residual activity is reduced to 49 ± 4%. 3.4. Application to serum analysis A key factor for the application of an amperometric sensor in real life samples is its selectivity towards the easily electro-oxidazible species. In the case of serum, ascorbic acid, uric acid, acetaminophen and many other compounds can interfere. The porous carbon used here for the construction of the phosphate sensor has shown to be sensitive to these species (between 10 and 13 A/mM), preventing its application to serum analysis. For this reason a pre-oxidizing cell [29] was incorporated in the FIA system, to oxidize these species before they reach the electrode surface. Initial experiments indicated the need for the buffer to be mixed with the sample after the pre-oxidation step. When the buffer was introduced prior to the cell the response of the sensor was not reproducible. The final experimental set-up is shown in Fig. 3. The concentration of pyruvate in blood varies considerably (70–150 M) [31]. Unless there is a high and constant concentration of pyruvate in the buffer stream, this fluctuation will introduce an error to the biosensor’s response. Therefore, the concentration of pyruvate in the buffer stream was raised from 1 to 10 mM. At the same time, the concentration of HEPES in the buffer stream was also raised in order to increase its buffer capacity. Thus, the buffer stream was 50 mM HEPES and 10 mM pyruvate adjusted to pH 7.5 at a flow rate of 0.65 ml/min and the flow rate of the sample was also 0.65 ml/min.
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Fig. 3. Diagram of the flow injection system. The buffer consists of 50 mM HEPES and 10 mM pyruvate adjust to pH 7.5. The sample passes through the pre-oxidizing cell where the interfering species are electrooxidized, then is mixed with the buffer and reach the biosensor to complete the analysis.
The analytical characteristics of the phosphate biosensor incorporated in the flow injection system are as follows. The sensitivity is 445 ± 9 nA/mM, the response is linear from 0.5 to 10 mM, while the detection limit is less than 0.3 mM. In Fig. 4, a typical recording of phosphate injections is presented. Prior to real sample analyses, the efficiency of the cell in the elimination of interferents was then investigated. With the cell current off, the phosphate biosensor exhibits sensitivities 1.72, 2.27 and 1.63 A/mM
for ascorbate, acetaminophen and urate, respectively. When the cell current is on, all sensitivities are reduced to 0.26, 0.30 and 0.51 A/mM, respectively. With these sensitivities the detection limit of the system for these species is 362, 853 and 816 M for ascorbate, acetaminophen and urate, respectively, values that are much higher than their normal concentration in serum. The above experimental set-up was then used to determine the phosphate level in serum. Three samples of serum were analyzed and the results were compared with those obtained with ion-chromatography. The samples were injected in the system without dilution and the calculated concentration was found to be 1.03 ± 0.06 mM. The results from the chromatographic analysis were 0.99 ± 0.02 mM, very close to the results obtained with the biosensor.
4. Conclusions
Fig. 4. Recording of the response of the biosensor incorporated in the flow injection system shown in Fig. 3 to phosphate injections (2.5, 5.0, 7.5 and 10 mM phosphate, respectively). The sensor is polarized at +800 mV vs. Ag/AgCl. The flow rate is 0.65 ml/min and the buffer consist of 50 mM HEPES and 10 mM pyruvate adjusted to pH 7.5.
The development and evaluation of a phosphate biosensor based on stabilized with polyelectrolytes recombinant pyruvate oxidase adsorbed into a porous carbon has been presented. Both electrolytes used were shown to have a positive effect on both the operational and storage stability of the resulting biosensor, with 0.1%, w/v deae-dextran being the optimum. The phosphate biosensor exhibits exceptional operational and storage stability while the simple construction
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procedure results in a very good sensor-to-sensor reproducibility. The use of the sensor in an FIA system equipped with a pre-oxidizing cell allows for the measurement of the activity of phosphate in difficult samples such as those of human serum with results in good agreement with those obtained with ion-chromatography.
Acknowledgements We would like to thank, Dr. U. Spohn (University of Hall) for supplying the enzyme, G. Kouvarakis (ECPL, University of Crete) for the ion-chromatography measurements and the Greek Secretary General of Research and Development for financial support through the YPER program (97 YPER-219). References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10]
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