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Published online 9 January 2007
Published in J Environ Qual 36:233-238 (2007)
DOI: 10.2134/jeq2006.0216
© 2007 American Society of Agronomy, Crop Science Society of America, and Soil Science Society of America
677 S. Segoe Rd., Madison, WI 53711 USA
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Biosensing Paraoxon in Simulated Environmental Samples by Immobilized Organophosphorus Hydrolase in Functionalized Mesoporous Silica

Chenghong Lei, Michelle M. Valenta, K. Prasad Saripalli and Eric J. Ackerman*

Pacific Northwest National Laboratory, P.O. Box 999, Richland, WA 99352

* Corresponding author (Eric.Ackerman{at}pnl.gov)

Received for publication May 31, 2006.

   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL SECTION
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
There is a critical need for highly sensitive, cost-effective sensors to conduct ecological analyses for environmental and homeland security-related applications. Enzyme biosensors, which are currently gaining acceptance for environmental monitoring applications, need improvements to deliver faster measurements with stabilized sensing elements, e.g., enzymes. We report here on a method which significantly overcomes this difficulty, and demonstrate its application in a biosensor for aquatic environmental applications. A fast-responding and stable biosensor was developed via immobilization of organophosphorus hydrolase (OPH) in functionalized mesoporous silica (FMS) with pore sizes in tens of nanometers. The OPH-FMS composite was held on glassy carbon electrode by a dried Nafion gel and FMS protected OPH from Nafion-resulted activity loss. The resulting enzyme biosensor, when integrated with an electrochemical instrument, responded rapidly to low paraoxon concentration and achieved steady-state current in less than 10 s, with a detection limit of 4.0 x 10–7 M paraoxon. The biosensor was tested for detection of paraoxon in simulated environmental samples, under wide-ranging physicochemical conditions. Results clearly indicate high recovery efficiencies in aqueous solutions (96 to 101%) at different pH, total organic carbon, total dissolved solids, and total suspended solids, and demonstrate the ability of the biosensor unit to continuously monitor paraoxon in aqueous conditions similar to those found in river and lake systems.

Abbreviations: OPH, organophosphorus hydrolase • FMS, functionalized mesoporous silica • GDAH, glutaric dialdehyde • GCE, glassy carbon electrode • TDS, total dissolved solids • TOC, total organic carbon • BTC, simulated breakthrough curve


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL SECTION
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
ELECTROCHEMICAL enzyme biosensors are portable, even implantable, and suitable for on-line monitoring in the field (Graham, 1998). A vexing problem with conventional enzyme immobilization technology is its inability to produce fast-responding and stabilized enzymes. We report here a fast-responding and stable biosensor for aquatic environmental applications based on active and stabilized enzymes immobilized in functionalized mesoporous silica (FMS).

Mesoporous silica has been extensively investigated for industrial and environmental processes such as catalysis, adsorption, ion exchange, and sensing due to its open pore structure, well-defined pore size (diameter) and pore shape. Significant progress in this area occurred with the introduction of FMS (Bagshaw et al., 1995; Feng et al., 1997). The combined effects of monolayer versatility and controllable mesopore sizes would result in high selectivity and binding capacity of FMS. Mesoporous silica has been made in pore sizes from 20 to 500 Å, a size range suitable for many enzymes, proteins, and their complexes. With rigid and open pore structures and controllable pore sizes, mesoporous silica can facilitate mass transportation of the enzyme substrate and product, resulting in fast enzymatic reaction and fast sensor response. Mesoporous silica can also protect the enzyme because the pore size is sufficiently small to eliminate any invading bacteria (at least 2 µm in size). It has also been reported that confinement from molecular crowding in biological cells can both stabilize and induce order-of-magnitude enhancements in catalytic reaction rates compared with enzymes in solution (Minton, 2001; Zhou and Dill, 2001). Thus, enzyme immobilization in FMS may both stabilize and enhance catalytic activity.

Recently, we reported that organophosphorus hydrolase (OPH) spontaneously entrapped in controllably-designed FMS with pore size of 30 nm could exhibit dramatically increased enzyme activity compared with normal porous materials (Lei et al., 2002). As a result of the unique nanoscaled mesoporous structure and surface chemistry, the carboxylic acid (HOOC)-FMS provided both high affinity for OPH and a favored microenvironment that resulted in exceptionally high immobilization efficiency and enzyme stability, in contrast to conventional enzyme immobilization approaches such as encapsulation by sol-gel silica yielding low specific activity due to low surface area and nonopen porous structure (Glad et al., 1985; Braun et al., 1990; Schmidtsteffen and Staude, 1992; Bhatia and Brinker, 2000; Wei et al., 2000; Lei et al., 2002; Reetz et al., 2003). Amino acid residues of proteins can also be readily covalently linked with NH2– or HOOC-functionalized FMS using bioconjugate techniques to avoid enzyme release from FMS due to electrostatic interaction (Hermanson, 1996). In this work, NH2–FMS were first reacted with the bifunctional cross-linking agent, glutaric dialdehyde (GDAH). This resulted in covalently linking one aldehyde end with the internal wall of FMS by forming a Schiff base, leaving the other aldehyde end available for covalent linkage with the target protein, OPH.

Immobilized OPH has been investigated for detection, decontamination, and destruction of poisonous agents and pesticides (Caldwell and Raushel, 1991; Havens and Rase, 1993; LeJeune and Russell, 1996; LeJeune et al., 1998; Flounders et al., 1999; Mulchandani et al., 1999; Singh et al., 1999; Gaberlein et al., 2000; Richins et al., 2000; Sacks et al., 2000; Mullchandani et al., 2001; Simonian et al., 2001; Shimazu et al., 2003; Cao et al., 2004; Ji et al., 2005). In the case of OPH biosensors, for example, direct detection of organophosphates has been achieved based on immobilized OPH in sol-gel silica (Flounders et al., 1999; Singh et al., 1999). Amperometric thick-film electrodes were constructed earlier to monitor organophosphorus nerve agents (Mulchandani et al., 1999). Disposable potentiometric enzyme biosensor has been used for determination of organophosphorus insecticides (Gaberlein et al., 2000). Ji et al. demonstrated the use of quantum dots-OPH bioconjugate for the detection of paraoxon (Ji et al., 2005). In the present study, the immobilized, active, and stable OPH in FMS was loaded onto glassy carbon electrode (GCE) surfaces to construct a fast-responding and stable electrochemical biosensor for organophosphorus compounds. The OPH-FMS composite was held on GCE by a dried Nafion gel and FMS protected OPH from Nafion-resulted activity loss. Paraoxon, used in many toxicological studies as a surrogate for chemical warfare agents, was selected as the model compound. This work focused on testing the sensor performance under rugged environmental conditions: simulated and actual river water, soapy water, water with pulverized biomass (leaves, roots), high concentration of clay colloids, water with petroleum compounds, water with organic interferences (e.g., hexane, naphthalene, pyrene), and water with inorganic interferences (e.g., Fe, Na, K, Cs, etc.).


   EXPERIMENTAL SECTION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL SECTION
RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Materials
Organophosphorus hydrolase (OPH) in pH 7.5, 0.1 M HEPES was prepared and purified as reported earlier (Lei et al., 2002). Mesoporous silica (pore size, 30 nm) and 20% NH2–FMS was prepared as described previously (Lei et al., 2002). Paraoxon, Nafion aqueous solution, and other chemicals were obtained from Sigma Chemical Co. (St. Louis, MO). Deionized (DI) water was used to prepare the solutions used in this work.

Preparation of Organophosphorus Hydrolase Functionalized Mesoporous Silica Composite
To react NH2–FMS with the bifunctional cross-linking reagent GDAH, an aliquot of 2 to 8 mg of NH2–FMS in a 1.8-mL tube was added with 200 to 400 µL of 5% GDAH in the same buffer solution as used for the enzyme stock, and shaken for 10 min at room temperature (21 ± 1°C). Subsequently, it was centrifuged at 1200 rpm for 6 min and the supernatant containing the excess covalent linker was decanted. The resulting deposit was washed with 3 x 400 µL of the same buffer and then immediately incubated with 100 to 1200 µL of the enzyme stock, and shaken at 1400 min–1 on an Eppendorf Thermomixer 5436 for 1 to 3 h at 25°C. The enzyme incubation solution was separated by centrifugation, and the resulting deposit was washed exhaustively at room temperature and centrifuged to remove any nonfirmly immobilized enzyme. Finally, the washed deposit was resuspended in pH 7.5, 20 mM HEPES by 300 µL of the buffer per mg of original FMS.

Preparation of the Biosensor
One hundred µL of the OPH-FMS suspension was mixed with the same volume of 0.2% Nafion aqueous solution. Aliquots (5 uL) of OPH-FMS/Nafion mixture were deposited on a GCE (3 mm in diameter) and allowed to dry at 21°C to form OPH-FMS/GCE. The resulting biosensor was rinsed with H2O before use.

Apparatus
All electrochemical experiments were performed with the Autolab PSTAT12 (Eco Chemie, Netherlands). A working buffer of pH 8.5, 0.1 M HEPES/0.1 M NaCl was used throughout this work. The Autolab was connected with a YSI 2700 (YSI Instruments, Ohio, USA) electrochemical reactor (0.5 mL) equipped with a Ag/AgCl (3.0 M NaCl) reference electrode and a platinum wire auxiliary electrode. Sample volume for this work was set at 25 µL. Sample injection, stirring, and flushing of the reactor can be controlled by the YSI control panel. Before measuring, the electrochemical reactor and three electrodes in the reactor were completely flushed with fresh working buffer. The stirring speed was set to be the minimum level fixed by the YSI instrument.

Measurement Procedure
The YSI electrochemical reactor was specially designed such that a predetermined volume of sample replaces the equivalent volume of the previous working solution in the reactor (Fig. 1). The equation for calculating the final concentration of a single sample is:

Formula 1[1]
where C is the final concentration in the reactor; Vs is the sample volume; Cs is the sample concentration and Vr is the reactor volume. Final concentration Cn during a continuous injection of the same volume of standard sample for a calibration plot is:

Formula 2[2]


Figure 1
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  		Fig. 1. Schematic of the biosensor testing unit. (a) the modified YSI electrochemical chamber (reactor); (b) YSI sipper tube; (c) glassy carbon electrode (GCE); (d) Ag/AgCl reference electrode; (e) Pt wire auxiliary electrode is separately drawn, whose position on the chamber is marked. The body of the auxiliary electrode was vertical to the plane of the drawing paper and the Pt wire spring was immersed in the working solution; (f) the flowing direction for the flushed solution; (g) the stirring bar; (h) the organophosphorus hydrolase functionalized mesoporous silica (OPH-FMS) layer held on GCE by a Nafion gel. The three electrodes are connected with Autolab PSTAT12. The sipper tube is driven by a YSI mechanism system so that the sipper tube can descend into or ascend from the chamber, or rotate above the chamber. Thus, the samples can be injected into the chamber and the tested solution can be flushed by refilling the working buffer solution via the sipper tube.

 
Preparation of Simulated Aqueous Samples
Simulated samples representing conditions found in lake and river water were prepared with the following variants: pH, ionic strength, turbidity, total dissolved solids (TDS), total organic carbon (TOC), clay colloids, soap content, biomass, petroleum, an inorganic interferent, and an organic interferent. This preparation was designed to simulate water quality parameter ranges found in natural waters for pH, ionic strength, TDS, and TOC (U.S. Geological Survey, 1985).

pH was adjusted in a range of 6.5 to 8.5 by the addition of 0.01 M NaOH to DI water. Ionic strength was varied between 10–2 and 10–5 M by adding NaNO3 to DI water. Sodium chloride was dissolved in water ranging from 15.0 to 500.0 mg L–1 for TDS and as a representative inorganic interferent. Humic acid was dissolved in water ranging from 3.0 to 6.0 mg L–1 for TOC. The soapy water was prepared by dissolving 11.4 mg of handsoap (Backdown antimicrobial handsoap manufactured by Decon Laboratories) in 100 mL of H2O and subsequently diluting this for a range of soap concentrations between 1.0 and 10.0 mg L–1. Kaolin was used to prepare water containing clay colloids in concentrations ranging from 1.0 to 100.0 mg L–1, by mixing and suspending in DI water. For the preparation of biomass-containing water samples, 106.1 mg of ground-up spinach was vigorously shaken for 5 h in 100 mL of DI water, the suspension was filtered, and the filtrate was diluted from 1 to 40 times as the biomass samples. Similarly, for the samples containing petroleum compounds, 572.7 mg of crude oil from the Alaskan North Slope was vigorously shaken for 5 h in 100 mL of DI water, the suspension was filtered, and the filtrate was diluted from 1 to 40 times as the petroleum samples. Hexane was used as a representative organic interferent, dispersed in DI water in the range of 1.0 to 100.0 mg L–1.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL SECTION
 RESULTS AND DISCUSSION
CONCLUSIONS
 REFERENCES
 
Cyclic Voltammetry of 4-Nitrophenol at Glassy Carbon Electrode
Organophosphorus hydrolase catalyzes the hydrolysis of paraoxon and produces the equimolar product 4-nitrophenol. Fig. 2 shows the cyclic voltammetry of 4-nitrophenol at GCE. Nitrophenol can be easily oxidized at the GCE at a potential window of 0.70 to 1.20 V vs. Ag/AgCl. In contrast, paraoxon itself did not show any oxidation peak at the same potential window. Therefore, by means of immobilization of OPH on GCE, the resulting biosensor is poised at a fixed potential to generate an oxidation current of the hydrolyzed product proportional to the paraoxon concentration and accordingly paraoxon can be monitored.


Figure 2
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  		Fig. 2. Cyclic voltammetry of 0.25 mM 4-nitrophenol (solid line) and paraoxon (dotted line) in pH 8.5, 0.1 M HEPES/0.1 M NaCl at glassy carbon electrode (GCE). Scan rate: 0.1 V s–1.

 
Experimental Conditions and Calibration
The OPH-FMS composite could not be firmly attached on GCE by itself. Therefore, the Nafion polymer was tested for holding OPH-FMS on GCE. The preliminary experiments indicated that the free OPH in 1% Nafion aqueous solution only maintained 13% of its initial activity in the absence of Nafion. This result shows that Nafion could be poisonous to some enzymes although it is extensively used as an immobilization matrix (Mulchandani et al., 1999; Karyakin et al., 2002; Jia et al., 2003). In contrast, OPH-FMS in the same concentration of Nafion solution kept 89% of its initial activity, indicating FMS could prevent OPH from being denatured by the strong ion exchange polymer Nafion. In practice, a minimum concentration of 0.1% Nafion was used to prepare OPH-FMS suspension before being deposited on the electrode surface.

pH can influence the paraoxon biosensor response in several ways. The catalyzed hydrolysis of paraoxon involves participation of hydroxyl anions. A lower pH (less than 7.0) would decrease OPH activity. The hydroxyl anion also promotes electrochemical oxidation of 4-nitrophenol, so these considerations favor using a weakly basic working buffer (higher than 8.0). However, if pH is too high (higher than 10.0) the structure of mesoporous silica would be damaged. Therefore we chose pH 8.5 for this work.

Electrochemical oxidation of 4-nitrophenol was observed at approximately +0.70 V (Fig. 2), and the steady-state current increased with the applied potential increasing up to +1.0 V. However, it was hard to reach a steady-state current when the applied potential was higher than +0.95 V. We chose +0.90 V as the working potential. At this potential, molecular oxygen was not involved in the electrode reaction. Hence, the working buffer was not deaerated. Extensive testing and calibration in batch mode shows excellent sensitivity and accuracy of the biosensor. Dynamic response of the paraoxon biosensor and a typical calibration plot are shown in Fig. 3A and 3B, respectively. At the end of each batch measurement, the biosensor and the electrochemical reactor could be rapidly flushed to the baseline for the next measurement (Fig. 3A). The biosensor integrated with the electrochemical instrument responded rapidly to low paraoxon concentration, achieved 95% of the steady-state current within 5 s, and the full steady-state current in less than 10 s, with a detection limit of 4.0 x 10–7 M paraoxon. The fast response of the biosensor to paraoxon was ascribable to the large, rigid, and open mesoporous structure. The prepared paraoxon biosensor was stored in pH 7.5, 0.1 M HEPES in a refrigerator when not in use. The stability of the biosensor was checked twice a week. After 5 wk, the decrease of the response sensitivity was less than 5%. The high stability of the biosensor can be attributed to the unique enzyme-binding and immobilization properties of FMS.


Figure 3
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  		Fig. 3. (A) Dynamic response of the paraoxon biosensor. Step = 10 µM; (B) Calibration plot of the paraoxon biosensor.

 
Recovery of Paraoxon in Simulated Samples with Single Variants
Aqueous simulated samples were prepared for each single variant of pH; ionic strength (NaNO3: 10 mM to 10 µM); inorganic salt (NaCl); clay colloid (kaolin clay); humic acid; soapy water; biomass (filtered and diluted spinach juice); petroleum (Alaskan crude oil saturated); and organic interferent (suspension of hexane). These variants of the simulated samples correspond to real ecological conditions (U.S. Geological Survey, 1985). A sample of river water (Columbia River, Richland, WA) was also tested. Shown in Table 1 are paraoxon recoveries from the river water and these simulated aqueous samples determined by the biosensor unit. The recovery of paraoxon in all the samples tested was nearly 100%. Thus, the results demonstrate that these environmentally important variants did not significantly affect the biosensor performance. Fig. 4 shows the continuous measurement of four simulated samples at different pH from 6.5 to 8.5. Each sample (the same volume) was added twice to obtain two steady responding steps. Then the electrochemical reactor was flushed with the fresh working buffer, resulting in the amperometric response decreasing rapidly to the baseline, so that the biosensor was again ready for the same measurement of next sample (Fig. 4). Although the instrumental baseline drift was observed, the amperometric response did not vary significantly for different pH values of samples due to the small sample volume and the buffering ability of the working solution. The biosensor unit displayed convenient operation, fast response, excellent measurement flexibility, sensitivity, and accuracy.


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  		Table 1. Recoveries of paraoxon in simulated aqueous samples with water quality variants measured by the biosensor.

 

Figure 4
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  		Fig. 4. Amperometric response of the paraoxon biosensor in simulated aqueous samples at different pH. The concentration of paraoxon in the testing solutions was 15.0 µM.

 
Determination of Paraoxon in Simulated Samples with Multiple Variants
Sensors deployed for field monitoring usually are required to function in a continuous detection mode. After the biosensor construction and extensive testing in the batch mode for its performance under various single environmental conditions, we also tested the biosensor in continuous detection mode for simulated samples with multiple variants. The objective of this test was to investigate the ability of biosensor to continuously detect samples in a flow mode. A simulated breakthrough curve (BTC) for up to 0.4 mM concentration in aqueous solutions was prepared as a set of discrete batch samples. This batch of BTC samples were continuously analyzed using the biosensor (Fig. 5). Results from this analysis clearly indicate excellent agreement of the measured concentrations with the expected concentrations, and demonstrate the ability of the biosensor unit to continuously monitor paraoxon in aqueous conditions similar to river and lake systems.


Figure 5
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  		Fig. 5. Continuous mode testing of the paraoxon biosensor on a simulated breakthrough curve. The samples contained 1 mM NaOH, 10 mM NaNO3, 50 mg L–1 clay colloid, 500 mg L–1 NaCl, and 5 mg L–1 humic acid.

 

   CONCLUSIONS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL SECTION
 RESULTS AND DISCUSSION
 CONCLUSIONS
REFERENCES
 
A fast-responding and stable electrochemical biosensor based on the immobilization of OPH in FMS was constructed to monitor paraoxon concentrations with high accuracy under a variety of simulated aqueous environmental conditions. The OPH-FMS composite was held on the electrode surface by a dried Nafion gel. Functionalized mesoporous silica protected OPH from the activity losses occurring in conventional immobilization approaches, where Nafion is used as the immobilization matrix alone. We selected OPH for the recognition of organic molecules and demonstrated its use for biosensing paraoxon as a model compound of environmental concern. The resulting paraoxon biosensor showed good stability, sensitivity, reproducibility, and fast response to paraoxon under simulated rugged environmental conditions. Results from this analysis clearly indicate excellent agreement of the measured concentrations with the expected concentrations, and demonstrate the ability of the biosensor unit to continuously monitor paraoxon in aqueous conditions similar to river and lake systems. Functionalized mesoporous silica was found to be a suitable material for enzyme immobilization and for protecting enzymes from the harsh matrix-resulted activity loss for enzyme biosensors. It will be worthwhile to further develop the biosensor for field testing and eventual ecological applications.


   ACKNOWLEDGMENTS
 
We gratefully acknowledge funding of this work by Laboratory Directed Research and Development funds within Pacific Northwest National Lab.'s Environmental Technology Division and the U.S. Dep. of Energy Office of Biological and Environmental Research.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL SECTION
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 





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