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TECHNICAL NOTES Carbon-Fiber Ultramicroelectrodes Modified with Conductive Polymeric Tetrakis(3-methoxy-4-hydroxyphenyl)porphyrin for Determination of Nickel in Single Biological Cells Frederick Bailey and T a d e u s z Malinski* Department of Chemistry, Oakland University, Rochester, Michigan 48039-4401 Frederick Kiechle Department of Clinical Pathology, William Beaumont Hospital, Royal Oak, Michigan 48072 INTRODUCTION Chemically modified microelectrodes add the dimension of selectivity to the inherent sensitivity of ultramicroelectrodes (1, 2). The modification of conventional scale electrodes (1-mm diameter) with conductive nickel poly(tetrakis(3methoxy-4-hydroxypheny1)porphyrin)has been described previously ( 3 ) . In this work, this modification is applied to ultramicroelectrodes for the detection of nickel in a single biological cell. Although no specific function for nickel has been established, evidence exists that supports the hypothesis that Ni(I1) is an essential metal that occurs a t trace concentrations in physiological and environmental systems. At higher concentrations, it has been found to cause contact dermatitis, fibrosis, emphysema, and pulmonary lesions, and of most concern, it was found to be carcinogenic (4,5). Subsequently, it is desirable to monitor the levels of nickel in organisms and the environment. Inhalation is the predominant route of exposure in man. Studies have shown that following inhalation, nickel is distributed in the following way, listed from high to low concentrations: lung, heart, kidney = brain, liver (6). Total nickel concentrations in cells have been determined by using radiochemical or liquid scintillation counting techniques with 63Ni. However, autoradiographic techniques cannot differentiate between nickel bound by proteins or adsorbed onto cellular membranes and unbound nickel in intracellular fluids. Ultramicroelectrodes can be used as an invasive technique to determine unbound nickel directly in intracellular fluids. Ultramicroelectrodes have been designed for the detection of small biological molecules and drugs in physiological fluids (7). They can be fabricated with appropriate dimensions to be inserted in a single cell or tissue. Microelectrodes have increased mass transport, decreased double-layer capacitance, and decreased ohmic loss with the general effect of an increased signal to noise ratio, a time independent response, and higher sensitivity ( I , 8). Selectivity of ultramicroelectrodes can be achieved by modification of the electrode surface (9). This paper presents a method for determination of the unbound nickel in single biological cells, through the use of ultramicroelectrodes modified with a conductive polymeric porphyrin film. Nickel tetrakis(3-methoxy-4-hydroxypheny1)porphyrin (TMHPPNi) is used as an agent to modify the carbon-fiber microelectrodes. Initial oxidation of the monomeric TMHPPNi leads to polymerization and formation of highly conductive polymer film on the electrode surface. The polymer undergoes facile demetalation in acid solution leaving an intact, adherent, conductive film on the electrode
* To whom correspondence should be addressed.
surface that then selectively chemically incorporates Ni(I1) cations from analyte solutions. The electrode can then be transfered to a blank electrolysis solution where current, due to either the Ni(II)/Ni(III) oxidation or the catalytic oxidation of water, can be used as an analytical signal observed by differential-pulse voltammetry (DPV) (3). EXPERIMENTAL S E C T I O N Reagents and Materials. Nickel tetrakis(3-methoxy-4hydroxypheny1)porphyrin was synthesized according to a procedure described previously (IO). Celion G50-300 carbon fibers with a diameter of 6 pm were obtained from BASF. All acids and bases used were Suprapure (Ultrex, J. T. Baker). Standard solutions of cations were prepared from lo00 wg mL1 ICP reference stock standards (Spex Industries) for Ni(II), Fe(III), Cd(II), Pb(II), Cu(II), and Co(I1). NBS 1643b Trace Elements in Water, used as an analytical standard, was obtained from the U.S. National Bureau of Standards. A culture of H4-11423 rat hepatoma cells were grown on glass plates with Dalbecco's modified Eagle medium (Gibco Laboratories), which contained L-glutamine at a concentration of 4 mM and 15% controlled-processserum replacement (Sigma). Nickel accumulations and analytical measurements were performed with the cells in Trizma buffer. This solution was held at 37 "C. Apparatus. A classical, three-electrode system in a quartz cell fitted with a Teflon cap was used for modification of carbon fibers with poly-TMHPP and for Ni(I1) determinations. The working electrode was a poly-TMHPP ultramicroelectrode prepared as described below. Platinum wire was used as the auxiliary electrode, and a saturated calomel electrode (SCE), as the reference electrode. The SCE was separated from the analytical solution by a bridge, fitted with a Vycor disk (IBM), containing saturated KCl. All potentials are reported vs SCE. A PAR Model 174A polarographic analyzer with a PAR Model 181 current-sensitive preamplifier were used for DPV, and the resulting current-voltage curves were recorded with a PAR Model 9002A X-Y recorder. Ultramicroelectrode Fabrication. Ultramicroelectrodeswere produced by threading an individual carbon fiber (Celion G50-300) through the pulled end of a capillary tube with approximately 1 cm left protruding. Nonconductive 5-min epoxy (Devcon)was put a t the glass/fiber interface. When the epoxy that is drawn into the tip of the capillary dried, the carbon fiber was sealed in place. A key factor in these studies is that the length and area of the electroactive surface must be appropriate for the dimensions of the cells under investigation. Submerging the end of the electrode in melted wax coats the carbon fiber. Carefully burning the coated fiber in a controlled temperature gradient obtained with a microburner sharpens the tip (11). The length and shape of the electroactive area obtained is controlled by the thickness of the wax coating, which is controlled by the temperature of the melted wax. In this work, the sharpened tip (0.5-1 pm) of the fiber was covered electrochemically with 10-90 monolayers of polymeric porphyrin, demetalated, implanted, and used for Ni(I1) accu-
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Figure 1. Microscopic photograph of a carbon-fiber ultramicroelectrode. 1
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mulations and current determinations. Figure 1 is a photograph of a sharpened electrode observed under a microscope. Waxing prior to the thermal sharpening step produces a short “spearhead” that is the only electrically conductive area that can be plated with polymeric TMHPPNi film. Electron micrographs of the carbon fiber with and without the film are shown in Figure 2. In the final step of electrode fabrication, the other end of the carbon fiber was attached to a copper wire lead with the use of silver epoxy (A. I. Technology). Poly-TMHPP Electrode Modification. The electrode modification process begins with deposition of conductive, poly-TMHPPNi film on the electrode surface and verification of film deposition. Deposition is followed by demetalation and verification of demet alat ion. Polymeric film is deposited from a solution of 0.1 M NaOH M TMHPPNi by constant-potential eleccontaining 5 X trolysis at 0.7 V. A peak attributable to the Ni(II)/Ni(III) couple appears at E l i 2 = 0.50 V in a DPV scan. The number of monolayers deposited is dependent upon the initial concentration of TMHPPNi and the time of electrolysis. After film formation, the electrode is removed from the deposition solution, rinsed, and immersed in 0.1 M NaOH. The presence of poly-TMHPPNi film is confirmed by the Ni(II)/Ni(III) couple, which appears at EIi2 = 0.50 V. The poly-TMHPPNi film is then demetalated by placing the electrode in a stirred 0.1 M HCl solution for 120 s. The electrode then is removed from the solution, rinsed, and immersed in 0.1 M NaOH. The absence of peaks associated with the Ni(II)/Ni(III) couple when scanning between 0.0 and 0.70 V confirms the absence of Ni in the demetalated poly-TMHPP film.. Electrodes have been stored with intermittent use for periods as long as 1 month without deterioration in response. Chemical
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preconcentration of nickel was performed in 5-mL solutions contained in a quartz cell or from a single biological cell. Implantation of the ultramicroelectrodes into individual cells was effected by using a Nikon TMS inverted microscope and a Stoelting HS 6 micromanipulator. Preconcentration times studied varied from 1.5-30 min. Solutions were not stirred. The poly-TMHPP film electrode with its incorporated nickel was removed from the analyte solution, rinsed with H20, and then transferred to 0.1 or 1.0 M NaOH, and nickel was determined by DPV. Liquid Scintillation Counting of “Ni. A tissue culture of H4-II-C3 rat hepatoma cells was solubilized in organic quaternary amines (PCS, New England Nuclear). Typically, 0.2-0.3 g of the wet cell sample was vortexed with 1-2 mL of the amines, incubated overnight at 50 “C,and then mixed with liquid scintillator (Aquasol-Beckman). A 1-mL aliquot of solubilized sample was added to 15-20 mL of liquid scintillator. The counting efficiency, using the tritium channel (Eo-max, 60 keV), was about 50%. Samples were standardized internally with no more than 25 p L of‘ “Ni/20 mL of counting mixture. RESULTS AND DISCUSSION A detailed characterization of the TMHPPNi film formation on coiiventional electrodes has been the subject of previous works (IO). Figure 3 shows the growth patterns for TMHPPNi in continuous-scan cyclic voltammetry from 0.00 to 1.00 V a t a carbon-fiber electrode (20-pm2 surface area). Peak Ia corresponds to the oxidation of the porphyrin ring on an ultramicroelectrode and peaks IIa and IIc, observed a t = 0.50 V, correspond to the Ni(II)/Ni(III) redox couple. Peak IIIa corresponds to the catalytic oxidation of water to molecular oxygen. Deposition of polymeric TMHPPNi on ultramicroelectrodes during continuous-scan cyclic voltammetry is relatively slow and requires about 90-100 scans from to obtain 30 equivalent 0.00 to 0.70 V (scan rate 0.10 V monolayers on a 78-pm2electrode. This is less efficient than a t larger electrodes (area of 0.07 cm2) on which polymerization and film formation is observed after a few scans using cyclic voltammetry. In order to obtain an electrode capable of preconcentrating Ni(I1) from the analyte solution, the original Ni in the polymeric TMHPPNi film has to be removed. The extent of the demetalation process was observed by placing the polyTMHPPNi/carbon-fiber ultramicroelectrode electrode that
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potential, V Figure 5. Differential-pulse voltammograms of nickel obtained with polymeric TMHPPNi film in 0.1 M NaOH (a) background and of preconcentrated nickel from 1 X lo4 M (b) and 5 X M (c) solutions at pH 7.4 for 15 min.
had been previously immersed in acid, which leaches the Ni(I1) from the porphyrin, in a 0.1 M NaOH solution and monitoring the current due to oxidation of Ni(I1) to Ni(II1) (peak IIa) or the catalytic oxidation of water (peak IIIa). A demetalation time of 2 min in a 0.1 M HC1 solution was found to be sufficient to remove the Ni(I1) to a concentration level below that detectable by differential-pulse voltammetry. Demetalation of the polymeric porphyrin film strongly depends on the pH of the solution. Figure 4 shows the relationship between pH of the solution and demetalation of the film on an ultramicroelectrode. This figure gives an indication of the useable pH range. Once demetalated, the poly-TMHPP film can be expected to readily reincorporate nickel from the analytical system ( 3 ) . Figure 5 shows the anodic Ni(I1) oxidation wave obtained with the poly-TMHPP ultramicroelectrode for three concentrations of standard nickel solutions. The reincorporation of nickel occurs with high efficiency from solutions having pH values of 7.0-9.0. In solutions with pH values lower than 6, a significant decrease in the rate of reincorporation of Ni(I1) is observed. On the other hand, if the pH is greater than 10.0, the Ni(I1) forms insoluble Ni(OH), (k!, = 1.6 X 10-14),decreasing the concentration of Ni and inhibiting the metalation process ( 3 ) . Both preconcentration time and initial nickel concentration should affect chemical preconcentration (12). Preconcentration yield was studied for each of these parameters. Solution concentrations of 5-10000 WMNi(I1) were investigated. For 1.5-3.0-min preconcentration times, a linear relationship
Flgure 6. Differential-pulse voltammogram of catalytic oxidation of water obtained with polymeric TMHPPNi film in 0.1 M NaOH. Nickel was preconcentrated from 1 X M solutions at pH 7.4 for 15 min; the electrode area was 20 pm2.
between preconcentration time and peak current was observed. In general, with longer preconcentration time (greater than 3 min for 1 x M), a steady decrease of A i / A t will be observed due to the decrease of the nickel concentration gradient, between the solution and film. A linear relationship between the current of the Ni(II)/ Ni(II1) reaction in the film and nickel concentration in the bulk solution was observed for the concentration range between l x and l x lo-'- M. A detection limit of 5 X M was obtained by using this redox couple as the analytical signal. However, the high current observed for catalytic oxidation of water on poly-TMHPPNi (wave 111) using ultramicroelectrodes creates a means of lowering detection limits. A good linear relationship between concentration of nickel in solution and catalytic current was observed with ultramicroelectrodes. Thus, in addition to voltammetric determination of nickel based on wave IIa, a convenient method based on monitoring a catalytic current due to oxidation of water to oxygen (wave 111) can be used. The catalytic current is several orders of magnitude higher than current due to Ni(II)/Ni(III) process. Therefore, a decrease of the detection limit by this same order magnitude would be expected. The relationship between Ni concentration and catalytic current in 0.1 M NaOH is linear between 5 x and 5 X lo+ M Ni(I1) with a detection limit approaching lo+ M. The analytical signal due to the catalytic process can be enhanced by 1 order of magnitude if 1.0 M NaOH is used for the Ni determination step instead of 0.1 M NaOH. This is consistent with the mechanism of the oxidation of water on polymeric TMHPPNi surfaces (IO). Figure 6 shows the differential-pulse voltammogram of catalytic oxidation of water by nickel accumulated in the porphyrinic film from a 2 X M solution of Ni(I1) at a pH of 7.4 for 15 min. No interferences were observed for solutions M Ni(I1) that contained 50-fold excesses of Cu(II), of 1 X Pb(II), Cd(II), or Fe(II1). The interference of the Co(II), when the catalytic peak was observed, was less significant (about 40% reduction of current) compared to that seen in earlier studies (80% for 50-fold excess) when the Ni(II)/Ni(III) peak was followed under similar conditions. In order to determine whether the presence of protein molecules in biological fluid would block the electrode surface, thereby interfering with the uptake of Ni(II), solutions mimetic of intercellular fluids, were prepared containing 50 mg of albumin in 5 mL of HzO. This concentration should be representative of total protein concentrations in cells. Determination of Ni(I1) was then successfully performed in this solution. This experiment showed that Ni(I1) uptake did occur. Therefore, the electrode surface is not significantly blocked by the high cellular protein content.
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The use of ultramicroelectrodes, now proven for the measurement of Ni(I1) concentrations in solution, was applied to the measurement of Ni(I1) in an individual cell of a cell culture of H4-II-C3 rat hepatoma cells. Cells in this particular cell line have an approximately spherical shape with a diameter nearing 10-15 pm. Thus the length of the sharpened/electroactive tip, which is plated with TMHPP, should be somewhat less than this dimension. The cell culture was grown directly on glass plates in Dalbecco‘s modified Eagle medium at 37 “C in a 10% C 0 2 atmosphere. The media was drawn off, and Tris buffer M Ni) was (containing 3% BSA, 2 mM glucose, and 1 X added. The cells were soaked in this Tris buffer solution for 45 min in order to allow Ni uptake. Following the uptake time, this Tris buffer was withdrawn and a fresh Tris buffer solution that did not contain Ni(I1) was added. Cells grown on glass slides in a Petri dish were placed on the inverted microscope, and the electrode was implanted by using the micromanipulator. When the electrode pierced the cell wall, the timing was initiated. The electrode was left in place for the desired preconcentration time. At the end of this time period the electrode was extracted, rinsed with deionized water, and placed in a 0.1 M NaOH solution for the determination of Ni(I1). The current from this analysis relates to the concentration of unbound Ni(I1) in the cell. For a 2-min preconcentration time, if the measurement was taken after the M Ni(I1) cell culture was subjected to a buffer with 1.0 x for 45 min, the resulting current represented a (1.0 f 0.1) X M Ni(I1) concentration in the cell. Total nickel accumulation, for conditions which were identical with those employed with the microelectrode studies, was determined by using liquid scintillation counting of 63Ni. The calculated nickel accumulation was 10 pg/million cells. Assuming the cells have a spherical shape with a radius of 7 pm, the total concentration of accumulated nickel, bound and unbound, in each cell would be 1 X M. Therefore, in a single rat hepatoma cell after 45 min, the total nickel accumulation was 0.01 pg and only 6% (0.6 fg) of this amount was unbound, as determined by using the ultramicroelectrode and differential-pulse voltammetry. After 5 h of allowed accumulation, the total amount of nickel doubled to 0.02 pg/cell. However, no significant change in the amount of unbound nickel was observed. A similar total accumulation of nickel (0.015 pg/cell) for a 4 h incubation period from 3 X M nickel solution was observed for FM3H cells from C3H mammary tissue (13). These data indicate that relatively high amounts of nickel accumulate in the cells studied. This effect has been noted
previously and attributed to the fact that the cell membrane is negatively charged, enabling it to compete for nickel binding with other ligands. Nickel internalization by the cell is possible by either active transport, most likely through calcium and/or magnesium channels, or passive diffusion of neutral, lipophilic complexes (14). While the exact nature of the nickel-binding sites in the cell is unknown, studies have pointed to the involvement of phosphate groups of both RNA and to a lesser extent DNA along with proteins, phospholipids, amino acids, mononucleotides, and other ligands. These extensive binding capabilities within the cell for nickel account for the relatively small proportion of total nickel concentration left unbound. ACKNOWLEDGMENT We thank Drs. J. R. Fish and K. Kasprzak for helpful comments and suggestions. Registry No. Ni, 7440-02-0; nickel poly(tetrakis(3-methoxy4-hydroxyphenyl)porphyrin),126752-50-9.
LITERATURE CITED Fieischmann, M.; Pons, S.; Roiison, D. R.; Schmidt, P. P. Ukramicroelectrodes; Datatech Systems Inc.: Morganton, NC, 1987. Guadalupe, A. R.; Abruna, H. D. Anal. Chem. 1985, 5 7 , 142-149. Malinski, T.; Ciszewski, A,; Fish, J. R.; Czuchajowski, L. Anal. Chem. 1990, 62, 909-914. Fishbein, L.; Mehiman, M. A. Advances in Modern Environmental Toxicology; CRC Press: Boca Raton, FL, 1987; pp 145-183. Costa, M. Env. Heanh. Perspect. 1989, 8 1 , 73-76. Coogan, T. P.; Latta, D. M.; Snow, E. T.; Costa, M. Crit. Rev. Toxicol. 1989, 19, 341-384. Ponchon, J. L.;Cespugiio, R.; Gonon, F.; Jouvet, M.; Pujol. J. F. Anal. Chem. 1979, 51, 1483-86. Tanaka, K.; KashiwaQi, N. J. Electroanal. Chem. Interfacial Eiectrochem. 1989, 275, 95-98. Turner, A. P. F.; Karube, I.; Wilson, G. S. Biosensors Fundamentals and Applications : Oxford University Press: Oxford, New York, Tokyo, 1987. Malinski, T.; Ciszewski, A,; Bennett, J. E.; Czuchajowski, L. Proceedings of the Symposium on Nickel Hydroxide Nectrodes; The Electrochemical Society Meeting, Hollywood, FL, October 1989; Corrigan. D. A., Zimmerman, A. H. Eds.; The Electrochemical Society: Pennington, NJ. 1990; pp 177-193. Josowicz, M.; Potje, K.; Liess, H.-D.; Besenhard, J. 0.; Kurtze, A. Electrochemistry, Sensors and Analysis; Elsevier: Amsterdam, 1987: pp 87-104. Whiteley, L. D.; Martin, C. R. Anal. Chem. 1987, 59, 1746-1751. Nishimura, M.; Umeda, M. Mutat. Res. 1979, 6 8 , 337-349. Nieboer, E.; Stafford, A. E.; Evans, S. L.; Dolovich, J. Nickel in the Human Environment; Sunderman, F. W., Ed.: IARC Science Publication No. 53; International Agency for Research on Cancer: Lyon, 1984; pp 321-331.
RECEIVED for review June 20, 1990. Accepted October 26, 1990. This work was supported by a grant from William Beaumont Hospital Research Institute.
Tissue Bioelectrode for Eliminating Protein Interferences Joseph Wang,* Li Huey Wu, Sandra Martinez, and Juanita Sanchez Department of Chemistry, New Mexico State University, Las Cruces, New Mexico 88003 One of the major difficulties in clinical applications of voltammetry stems from the adsorption of proteins onto the electrode surface. A degraded and irreproducible response characterizes these deactivation processes. Such loss of electrode activity is commonly referred to as “fouling” or “poisoning”. Various approaches have been used to alleviate the protein fouling problem. Permselective coatings represent one useful avenue. Size-selective cellulose acetate films have been particularly attractive for this task (1,2). Reactivation
* To whom correspondence should be addressed.
schemes, based on electrochemical or laser pulses, have also proved useful for the prevention of surface passivation effects (3,4).
This paper describes a novel and highly useful approach, based on tissue-modified electrodes, for the prevention of biofouling. The rich biocatalytic activity of tissue-containing surfaces is exploited for in situ enzymatic digestion of interfering proteins. The use of plant or animal tissues in place of isolated enzymes can offer attractive properties for biosensing probes, including extended lifetimes, high enzymatic activity, and low cost ( 5 , 6). These advantages have been
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