Anal. Chem. 1996, 68, 3670-3678
Antifouling Membrane-Covered Voltammetric Microsensor for in Situ Measurements in Natural Waters M.-L. Tercier* and J. Buffle
Department of Inorganic, Analytical and Applied Chemistry, Sciences II, University of Geneva, 30 Quai E.-Ansermet, 1211 Geneva 4, Switzerland
A novel, effective, antifouling membrane-covered voltammetric microsensor has been developed. It combines the unique properties of microelectrodes and diffusion in gels. An agarose gel, with thickness varying in the range 0.40.9 mm, is used as the membrane and acts as a dialysis membrane, i.e. allows diffusion of small ions and molecules and retains colloidal materials. The voltammetric microelectrode measures the test compounds within the gel after equilibration. Diffusion of ions through various types of gels has been investigated by diffusion cell, square wave cathodic sweep voltammetry (SWCSV), and square wave anodic stripping voltammetry (SWASV) to characterize the diffusion properties of the agarose gel membranes. These studies revealed that (i) the agarose gel used is totally inert toward target compounds and (ii) the anticonvection properties of the agarose gel ensure purely diffusion-controlled transport within the membrane. Diffusion coefficient values of ions in the agarose membrane were found to be lower than those in free solution. Considering the inertness of the agarose gel used toward the ions measured, this discrepancy can be ascribed to the intrinsic physical properties of the agarose gel which influence the ion mobility. Reproducible values have been obtained for equilibration times and diffusion coefficients for gels prepared under controlled conditions with a given type and concentration of agarose. The agarose membranecovered, Hg-plated, Ir-based microelectrode (µ-AMMIE) was applied to lead and cadmium analysis by SWASV in the presence of 10-31 mg/L of fulvic and humic compounds and in raw river waters containing high concentrations of suspended matters (50-78 mg/L). Results of these measurements under such drastic conditions confirm the efficiency of the agarose gel membrane against adsorption of organic and inorganic matters on the Hg surface of the voltammetric microsensor. Thus, these results demonstate that direct measurements of analytes in complex media can be made with µ-AMMIE without physical and chemical interferences of the test solution, in particular due to the relatively large gel thickness compared to the diffusion layer thickness of the microelectrode and of the properties of the agarose gels used.
metal concentrations are required.1-5 Reliable speciation measurements at trace levels (10-11-10-8 M) and rigorous interpretation of the data can be achieved only if measurements can be performed with minimum analytical artifacts (contaminations, losses by adsorption, physicochemical modification of the samples, etc.). Hence, ideally, the chosen analytical method should allow in situ measurements, be highly sensitive and reliable, possess speciation capability, and be fairly rapid and readily automated with compact and moderate-cost instrumentation. All these requirements are met by voltammetric stripping techniques coupled with a reproducible and reliable Hg-plated, Ir-based microelectrode. It was shown previously6,7 that this electrode allows direct measurements of Pb(II) and Cd(II) at concentrations as low as 50 pM in low ionic strength freshwater, provided that dissolved organic matter and suspended particle contents are low. In natural waters containing significantly high concentrations of natural organic matter or inorganic colloids, voltammetric signals of trace metals may often be modified or may even be suppressed by adsorption of these organic and inorganic matters onto the mercury electrode surface (electrode fouling).8-10 Fouling problems are also the main limitation of direct voltammetric measurements in other complex matrices, i.e. biological or industrial samples. The most promising way to minimize such interferences is to prevent the diffusion of interfering compounds toward the voltammetric sensor surface. Several techniques have been proposed for this purpose; in particular, coating the mercury film electrode with a thin semipermeable protective membrane excludes the fouling material by size exclusion and/or electric charge exclusion. Specifically, cellulose acetate,11-13 Nafion,14-16 Nafion-cellulose acetate,17 polyaniline,18 poly(ester sulfonic acid),19
In order to understand the role and fate of toxic heavy metals in natural aquatic media, speciation studies in addition to total
(1) Fo ¨rstner, U.; Wittmann, G. Metal Pollution in the Aquatic Environment; Springer: New York, 1981. (2) Buffle, J., De Vitre, R. R., Eds. Chemical and Biological Regulation of Aquatic Systems; Lewis Publishers: Boca Raton, FL, 1994. (3) Stumm, W., Ed. Chemical Processes in Lakes; Wiley: New York, 1985. (4) Buffle, J. Complexation Reactions in Aquatic Systems. An Analytical Approach; Horwood: Chichester, U.K., 1988. (5) Ure, A. M., Davidson, C. M., Eds. Chemical Speciation in the Environment; Blackie Academic & Professional: Glasgow, U.K., 1995. (6) Tercier, M.-L.; Buffle, J. Electroanalysis 1993, 5, 187. (7) Tercier, M.-L.; Parthasarathy, N.; Buffle, J. Electroanalysis 1995, 7, 55. (8) Buffle, J.; Vuilleumier, J. J.; Tercier, M.-L.; Parthasarathy, N. Sci. Tot. Environ. 1987, 60, 75. (9) Brezonik, P. L.; Brauner, P. A.; Stumm, W. Water Res. 1976, 10, 605. (10) Sagberg, P.; Lund W. Talanta 1982, 29, 457. (11) Aldstadt, J. H.; Dewald, H. D. Anal. Chem. 1993, 65, 922. (12) Baldwin, R. P.; Thomsen, K. N. Talanta 1991, 35, 1. (13) Wang, J.; Bonakdar, M.; Pack, M. M. Anal. Chim. Acta 1987, 192, 215. (14) Vidal, J. C.; Vin ˜ao, R. B.; Castillo, J. R. Electroanalysis 1992, 4, 653. (15) Hoyer, B.; Florence, T. M. Anal. Chem. 1987, 59, 2839.
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© 1996 American Chemical Society
and poly(ethyl 3-thiopheneacetate)20 have been used. Unfortunately, these protective layers generally are neither effective enough for complete elimination of fouling nor completely inert vis a` vis analyte for most of them; moreover, their preparation modes are not reproducible enough. In the present study, the properties of microelectrodes are exploited to develop a novel form of electrode protection. Here, spherical diffusion occurs at the microelectrode surface, enabling trace metal measurements by stripping techniques in quiescent solution. This characteristic, together with low iR drop, makes it possible to cover the Hg-plated microsensor with a gel layer, comparatively thick (0.4-1 mm) with respect to the microelectrode size, in order to overcome the fouling problem. The low molecular weight test ions and molecules diffuse across the gel, whereas colloidal and/or macromolecular materials are retained by size exclusion, adsorption effects, or slow diffusion. The gel chosen should exhibit the following properties: (i) it should be inert toward the trace metal to be analyzed, i.e. neutral and noncomplexing, (ii) it should be easily prepared, and (iii) it should permit an easy renewal of the mercury layer without changing the gel membrane, i.e. deposition and reoxidation of the mercury film through the membrane. In addition, if the gel shows anticonvection properties and is not affected by temperature variation, then the analyte can be measured within the gel without chemical or physical influence from the test solution. On this basis, two different compounds seemed to be suitable for the preparation of a protective gel: polyacrylamide and agarose. Both these types of gels are widely used in chromatography and/or electrophoresis techniques. Agarose was chosen here since it has several advantages over polyacrylamide for in situ applications in natural systems: (i) agarose forms a more rigid gel (i.e. physically more resistant gel) than polyacrylamide at equal concentrations; (ii) high gel strength, even at low concentration, implies that agarose is an excellent anticonvection medium;21 (iii) agarose gels can be prepared at various concentrations more readily than polyacrylamide, which requires the use of mixtures of acrylamide (neurotoxin), bisacrylamide, and substances for initiating the polymerization (e.g., tetramethylethylenediamine (TEMED) and ammonium persulfate); and (iv) electrophoretic mobilities of ions in agarose gels have been shown to be unaffected22 by temperature variations in the range of 4-30 °C in contrast to polyacrylamide gels.23 In addition, agarose structure is strongly resistant to enzymatic hydrolysis.24 EXPERIMENTAL SECTION Apparatus and Operating Conditions. A specially designed diffusion cell was used for diffusion measurements of cations (NaI, TlI, PbII, CdII), anion (NO3-) and humic and fulvic acid compounds through agarose membrane. The diffusion cell consists of two plexiglass compartments, having 100 mL capacity, between which the agarose membrane of the desired thickness is inserted (see: (16) Hoyer, B.; Florence, T. M. Anal. Chem. 1987, 59, 1607. (17) Morrison, G. M. P.; Florence, T. M. Electroanalysis 1989, 1, 458. (18) Dam, M. E. R.; Thomsen, K. N.; Pickup, P. G.; Schrøder, K. H. Electroanalysis, 1995, 7, 70. (19) Wang, J.; Taha, Z. Electroanalysis 1990, 2, 383. (20) Wang, Z.; Galal, A.; Zimmer, H.; Mark, H. B. Electroanalysis 1992, 4, 77. (21) The agarose Monograph, 4th ed.; FMC Bioproducts: Rockland, ME, 1988. (22) Thomas, M.; Davis, R. W. J. Mol. Biol. 1975, 91, 315. (23) Allet, B.; Jeppesen, P. G. W.; Katagiri, K. J.; Delias, H. Nature 1973, 241, 120. (24) Armisen, R. Hydrobiologia 1991, 221, 157.
Agarose Membrane Preparation, below). Concentrations as a function of the diffusion time were measured in both compartments by atomic adsorption spectroscopy (AAS; Pye-Unicam SP9) for cations, UV/visible absorption (Perkin-Elmer 883 spectrophotometer) for anion, and either UV/visible absorption or TOC (Shimadzu TOC-5000) for humic/fulvic acids. Electrochemical measurements were performed by using a computer-controlled Amel 433A polarograph, a home-made plexiglass cell, and a three-electrode configuration. The reference electrode was a Metrohm Ag/AgCl/3 M KCl/0.1 M NaNO3 electrode and the counter electrode was a Metrohm platinum rod. The working electrode was an agarose membrane covered, Mercury-plated, Iridium-based microelectrode (µ-AMMIE) prepared as follows: a heat-shrinkable tubing of 2 mm diameter fixed at the tip of a home-made Ir-based microelectrode, described in detail in ref 7, was cut with a scalpel to protrude 0.4-0.9 mm from the surface of the microsensor and filled with the agarose gel (see: Agarose Membrane Preparation, below). The mercury layer was plated through the agarose onto the Ir substrate at -400 mV (vs Ag/AgCl/3 M KCl/0.1 M NaNO3) in a deoxygenated 5 mM Hg(CH3COO)2 and 10-2 M HClO4 solution. Reoxidation of mercury was carried out by scanning the potential linearly from -300 to + 300 mV, at 5 mV/s, in a degassed 1 M KSCN solution. Voltammetric measurements were performed in the direct reduction mode using square wave cathodic sweep voltammetry (SWCSV), as well as in the stripping mode using square wave anodic stripping voltammetry (SWASV). SWCSV conditions used were as follows: initial E, -100 mV; final E, -1100 mV; pulse amplitude, 25 mV; step amplitude, 4 or 8 mV; frequency, 50 Hz; precleaning E, -100 mV; precleaning time, 30 s. SWASV conditions used were as follows: deposition potential E, -1100 mV; deposition time, 5-15 min in quiescent solution; final E, -100 mV; pulse amplitude, 25 mV; step amplitude, 4 mV; frequency, 50 Hz; precleaning E, -100 mV; precleaning time, 60 s. A precleaning step was performed before each voltammetric measurements to ensure (i) uniform spreading of the mercury layer on the iridium surface and (ii) complete reoxidation of the elements preconcentrated in the mercury during the previous measurement. A home-made preamplifier (based on the system developed by Faulkner et al.25 with 1000× amplification was used. A Leitz Diavert optical inverted microscope and a Laborlux normal optical microscope were used respectively for looking perpendicularly at the Ir-based microelectrode surface to check that no impurities are present before covering it with the agarose gel and for looking tangentially at the agarose membrane-covered, Ir-based microelectrode to check the quality of the gel. Reagents. All reagents used were of suprapur grade, and all solutions were freshly prepared before use with Milli-Q water (Millipore). Two different kinds of humic and fulvic substances were used. The first one was a stock solution containing 5.28 g/L of purified peat humic acid (PPHA), supplied by the British Geological Survey Laboratory, Wallingford, U.K. This peat humic acid was prepared, purified and characterized in detail by Milne et al.26 The second one was a mixture of fulvic and humic substances purchased from Fluka (No. 53680; Mw ) 600-1000; Merck Index 11,4671). A stock solution of 700 mg/L was prepared as follows: 1.3 g of (25) Huang, H. J.; He, P.; Faulkner, L. R. Anal.Chem. 1986, 58, 2889. (26) Milne, C. J.; Kinniburgh, D. G.; De Wit, J. C. M.; Van Riemsdijk, W. H.; Koopal, L. K. Geochim.Cosmochim. Acta 1995, 59, 1101.
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product was added to 250 mL of Milli-Q water and left to stand for 24 h under stirring conditions. The solution was then filtered through a Schleicher & Schu¨ll membrane (No. 595), and the total organic carbon concentration in the filtrate was determined by TOC. Both stock solutions were stored in a refrigerator at 4 °C. Antifouling Agarose Membrane. Agarose Composition and Structure: Key Aspects. Agarose is obtained from agar,21 which itself is extracted from the cell walls of marine red algae. It can be separated into two major fractions: (i) a low molecular weight fraction (MW ≈16 000), containing a significant number of ionic groups (sulfate, uronic and pyruvic acids), called agaropectin and (ii) a higher molecular weight fraction (MW ≈120 000) considered as the neutral agar fraction, called agarose. The mechanism for agarose gelation was first proposed by Rees27 and later confirmed by Arnott et al.28 It consists of an exothermic, reversible process occurring in different stages. First the agarose dissolved by heating forms random coils. On cooling, the ramdom coils form a double helix (tertiary structure). The double helix has a pit of 1.9 nm, and each strand has three-fold helical symmetry, with an intercaterny distance of about 0.85 nm, forming a very compact structure.29 Finally, the macroreticular network is formed. The exclusion limits of the gel can, in principle, be changed merely by changing its concentration and/or cross-linking.30-33 In fact, the above schematic structure is ideal and is based on the assumption that agarose polysaccharide chains are composed of regular alternance of 1,3-linked β-D-galactopyranose and 2,4-linked 3,6-anhydro-R-L-galactopyranose (agarobiose unit). In reality, agarobiose units can be substituted with various functional groups, and it has been shown experimentally that breaks in stereoregularity of the agarose structure by the presence of another substituent modifies the agarose gelation processes; in particular, inhibition of double-helix formation and aggregation have been observed.34,35 With high-purity agarose, two effects cause changes in the gel structure upon increasing agarose concentration: the mean mesh size between fibers contracts and the fiber diameter increases. Visualization of agarose gels by electron microscopy36,37 revealed that macroreticular networks are more reproducible at concentrations >1%. At concentrations e1%, protuberances or culde-sacs in the network are observed and their number increases with decreasing concentration. The above observations clearly demonstrate that gels prepared from commercial agarose samples would show different properties linked not only to the origin and the type of the red algae but also to the extraction procedure, the degree of chemical purity of the agarose, and its concentration in the gel. Differences have been observed, e.g., in gelation temperature, gel strength (defined as the force, in g/cm2, that must be applied to fracture the gel), and electroendosmosis (EEO), which represents the flow of countercations hydration water when electric current is applied across an agarose gel.21 (27) Rees, D. A. Biochem. J. 1972, 126, 257. (28) Arnott, S.; Fulmer, A.; Scott, W. E., Dea, I. C. M.; Moorhouse, R.; Rees, D. A. J. Mol. Biol. 1974, 90, 269. (29) Corongiu, G.; Forlini, S. L.; Clementi, E. Int. J. Quant. Chem: Quant. Biol. Symp. 1983, 10, 227. (30) Griess, G. A.; Guiseley, K. B.; Server, P. Biophys. J. 1993, 65, 138. (31) Kirkpatrick, F. H. Curr. Commun. Cell. Mol. Biol. 1990, 1, 9. (32) Griess, G. A.; Moreno, E. T.; Eason, R. A.; Serwer, P. Biopolymers 1989, 28, 1475. (33) Waldmann-Meyer, H. J. Chromatogr. 1987, 410, 233. (34) Yaphe, W.; Duckworth, M. Int. Seaweed Symp. 1971, 7, 15. (35) Rees, D. A.; Steele, J. W.; Williamson, F. B. J. Polym. Sci. 1969, C28, 261. (36) Whytock, S.; Finch, J. Biopolymers 1991, 31, 1025. (37) Server, P.; Hayes, S. Y. Biochemistry 1989, 28, 5827.
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Therefore, it appears that the best reliability in preparation of agarose gel will be obtained by using high-purity agarose, in particular agarose with sulfate concentration and EEO as low as possible and gel concentration g1.5%. Nature of the Agarose Chosen. Two types of agarose were used: (i) Agarose A6013 type I from Sigma, having the following characteristics: EEO e 0.15; maximum sulfur, 0.35%; gel strength, 1000 g/cm2 minimum at 1%; gelling temperature, 36 ( 1.5 °C, and melting temperature, 86 ( 2 °C at 1.5%; maximum ash 0.25%. (ii) Agarose LGL from Biofinex (Switzerland), exhibiting the following characteristics: EEO e0.13; maximum sulfur 0.03%; gel strength, 2000 g/cm2 minimum at 1.5%; gelling temperature, 36 °C, and melting temperature, 98 °C at 2%; ash nondetectable. Agarose LGL exhibits two very interesting features for our specific application: high gel strength and very low sulfur content. The latter is frequently used as an indicator of agarose purity, as many undesirable properties of agarose are attributed to the ionic groups, of which sulfate is a major constituent. It must be noted that agarose with sulfur content
