Influence of Surface Characteristics on the Stability of Cryptosporidium parvum Oocysts

Microelectrophoresis is a common technique for probing the surfacechemistry of the Cryptosporidium parvum oocyst. Results of previousstudies of the electrophoretic mobility of C. parvum oocystsin which microelectrophoresis was used are incongruent. In thiswork we demonstrated that capillary electrophoresis may alsobe used to probe the surface characteristics of C. parvum oocysts,and we related the surface chemistry of C. parvum oocysts totheir stability in water. Capillary electrophoresis resultsindicated that oocysts which were washed in a phosphate buffersolution had neutrally charged surfaces. Inactivation of oocystswith formalin did not influence their electrophoretic mobility,while oocyst populations that were washed in distilled waterconsisted of cells with both neutral and negative surface charges.These results indicate that washing oocysts in low-ionic-strengthdistilled water can impart a negative charge to a fraction ofthe oocysts in the sample. Rapid coagulation experiments indicatedthat oocysts did not aggregate in a 0.5 M NaCl solution; oocyststability in the salt solution may have been the result of Lewisacid-base forces, steric stabilization, or some other factor.The presence of sucrose and Percoll could not be readily identifiedon the surface of C. parvum oocysts by attenuated total reflectance-Fouriertransform infrared spectroscopy, suggesting that these purificationreagents may not be responsible for the stability of the unchargedoocysts. These findings imply that precipitate enmeshment maybe the optimal mechanism of coagulation for removal of oocystsin water treatment systems. The results of this work may helpelucidate the causes of variation in oocyst surface characteristics,may ultimately lead to improved removal efficiencies in full-scalewater treatment systems, and may improve fate and transportpredictions for oocysts in natural systems.

INTRODUCTION

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Introduction

Cryptosporidium parvum is a parasitic protozoan that causesthe diarrheal disease cryptosporidiosis. It has been associatedwith many outbreaks of disease in recent years (7, 12, 16).There was an outbreak in Milwaukee, Wis., in which C. parvumwas linked to over 400,000 illnesses and 104 deaths (27). Whenoutside the host, C. parvum exists as an oocyst, which is highlyresistant to a myriad of environmental conditions (12).

Efficient removal of C. parvum oocysts from drinking water suppliesby conventional treatment processes has been reported previously(31). However, C. parvum oocysts continue to break through watertreatment plants and enter drinking water distribution systems(7, 19, 25, 32, 35). Repeated breakthroughs of C. parvum oocystsmay be the result of an incomplete understanding of how oocystsinteract chemically and physically under conditions encounteredin water treatment systems (5, 7). Knowledge of the oocyst'ssurface chemistry and subsequent surface interactions duringthe water treatment process may be used to prevent such breakthroughs.

In order to study the surface chemistry of C. parvum oocysts,oocysts must be purified from infected animals since there iscurrently no cost-effective method to grow oocysts in vitro.There are a number of methods for isolating oocysts from animalfeces (37). In addition, oocysts are typically washed and storedin a variety of solvents, such as distilled water, prior touse. Knowledge of the influence of purification methods on oocystsurface chemistry is limited (2), yet oocysts are still studiedwithout taking into account this potentially crucial factor.Current guidelines for carrying out studies with oocysts donot take into account the potential effects of altered surfacecharges due to storage media (11). Use of oocysts with disparatesurface characteristics in laboratory and pilot plant studiescan lead to results which may not be extrapolated to full-scaleoperations. Indeed, changes in particle surface chemistry canhave a significant influence on particle aggregation and adhesion(1, 4, 15, 24, 26). An improved understanding of the influenceof oocyst purification and handling methods on oocyst surfacechemistry and subsequent stability (vide infra) in water maylead to more effective oocyst removal in engineered processes.

Microelectrophoresis has commonly been used to quantify theelectrophoretic mobility of oocysts (electrophoretic mobilityprovides an estimate of surface charge). Unfortunately, theresults reported previously for electrophoretic mobility havebeen incongruent. For example, some studies have shown thatC. parvum oocysts have an isoelectric point near pH 2.5 (10,20, 22, 23, 29, 33), while other reports (2) have demonstratedthat the oocyst surface remains neutral at pH values between2 and 10. Thomas and coworkers (36) recently reported that oocystssuspended in CaCl₂ solutions had near-zero electrophoretic mobility.Brush et al. (2) have reported that variations in observed electrophoreticmobilities were caused by dissimilarities in purification techniques.

Capillary electrophoresis has been used to quantify the electrophoreticmobilities of biological samples whose sizes are on the orderof 0.5 µm (13). Glynn et al. (13) reported that if similarnumbers of replicate measurements were used, the standard deviationsof electrophoretic mobility measurements obtained by capillaryelectrophoresis were typically lower than the standard deviationsof measurements obtained by microelectrophoresis. They attributedthe lower standard deviations to the larger sample size andlonger time required for separation with the capillary electrophoresistechnique. These investigators further reported variations incharge within monoclonal populations for several different bacterialstrains, a significant advantage of capillary electrophoresisover the traditional microelectrophoresis technique. This ispossible because the results of capillary electrophoresis arereported on an electropherogram on which retention time is afunction of electrophoretic mobility. On the other hand, theresults of microelectrophoresis are commonly reported as a singlevalue. Capillary electrophoresis was used in this study to evaluatethe influence of selected solvents that are often used to washor store oocysts on the electrophoretic mobility of oocysts.In capillary electrophoresis, particles and molecules in anelectrolyte solution are separated based on charge in a longcapillary tube, which has an electric potential applied acrossits length. The electric potential induces an electroosmoticflow due to a double layer that is formed at the capillary-solventinterface, which causes the bulk electrolyte solution to movefrom the positive electrode, located at the column influent,to the negative electrode, located at the column effluent. Thefar end of the column contains a detector (typically a UV detectorfor biological applications) that measures the presence of moleculesand particles at that point in the column. Neutrally chargedparticles and molecules move through the column with the bulksolvent via electroosmotic flow. A neutrally charged dopant,such as mesityl oxide, is used to measure the bulk solvent flow.Negatively charged molecules and particles move through thecolumn at a rate that is less than that of the dopant. The electrophoreticmobility of a particle can be quantified by the following relationship(13, 21, 34):

where µ is the electrophoreticmobility (in meters squared per volt·second), L_t is thetotal length of the capillary (in meters), L_d is the lengthof the capillary from the sample inlet to the detector (in meters),V is the voltage applied across the length of the capillary(in volts), t_r is the retention time of the neutrally chargeddopant (in seconds), and t_p is the retention time of the particle(in seconds) (see references 13, 21, and 34 for a more detailedexplanation of capillary electrophoresis theory).

The term stability is used to describe the tendency of particlesto resist aggregation. The stability of particles is a functionof repulsive and attractive forces between the particles (9,41). Factors that affect particle stability in water includepH, ionic strength, surface functional groups, and adsorptionof inorganic and organic matter from solution. During the productionof drinking water, colloidal particles in raw surface waterare typically destabilized, a process which promotes the formationof larger aggregates that can be removed by subsequent physicochemicalprocesses, such as sedimentation, filtration, or flotation.Particle destabilization is achieved by adjusting solution chemistrysuch that the net force between particles is attractive. Mechanismsof particle destabilization and aggregation include compressionof the double layer, charge neutralization, precipitate enmeshment,and interparticle bridging (1). In this study we examined theeffects of the double-layer-compression mechanism of coagulationon the stability of C. parvum oocysts. In double-layer compression,electrostatic repulsive forces, caused by charged moieties onparticle surfaces, are screened out by increasing the ionicstrength with an indifferent electrolyte, such as NaCl. If theelectrostatic force is the only repulsive interaction betweentwo particles, then the particles should aggregate via the attractivevan der Waals force. However, if other repulsive forces (e.g.,Lewis acid-base forces or hydration forces) are present in asystem or if the particles are sterically stabilized, then theymay not aggregate when the electrostatic force is screened viadouble-layer compression (14).

The objective of this project was to determine if the stabilityof C. parvum oocysts, used as received from commercial sources,was due to their surface charge. In this study we examined theelectrophoretic mobility and surface characteristics of C. parvumoocysts as affected by typical isolation and handling procedures.Capillary electrophoresis has not been used previously to determinethe electrophoretic mobility of this organism. Knowledge ofthe electrophoretic mobility and surface chemistry of oocystsmay elucidate causes of variation in oocyst surface characteristicsreported previously, may ultimately lead to improved removalefficiencies in full-scale water treatment systems, and mayimprove fate and transport predictions for oocysts in naturalsystems.

MATERIALS AND METHODS

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Materials and Methods

C. parvum oocysts.
C. parvum oocysts were obtained from Waterborne, Inc. (New Orleans,La.) and Pleasant Hill Farm (Troy, Idaho). Oocysts were isolatedfrom feces from experimental infected calves by sucrose andPercoll density gradient centrifugation after initial extractionof the feces with diethyl ether. The oocysts were stored at4°C in phosphate buffer amended with 1,000 U of penicillinper ml and 125 µg of streptomycin sulfate per ml to preventbacterial growth. Prior to use, the C. parvum oocysts were examinedmicroscopically for purity, and samples that had bacterial contaminationwere not used. The oocysts were collected by centrifugation(16,060 x g for 5 min) and suspended in distilled water or aphosphate buffer solution (20 mM, pH 7). Oocysts used in diffusereflectance infrared spectroscopy were obtained from the Departmentof Microbiology and Immunology, Cornell University (Ithaca,N.Y). The purification and storage procedures used for theseoocysts were the procedures described by Brush et al. (2). Theseoocysts were washed as described above. Unless otherwise stated,all oocysts were used within 2 months of the time that theywere received. The actual age of the stocks may have been slightlygreater because the time between purification and delivery wasnot known.

Microspheres.
Use of capillary electrophoresis to quantify the electrophoreticmobility of particles whose size was comparable to that of C.parvum oocysts (3 to 5 µm) was validated with latex microspheresthat were approximately the same size. Surfactant-free, 5-µm,carboxylate-modified polystyrene latex spheres (CML) were obtainedfrom Interfacial Dynamics Corp. (Portland, Oreg.). Stock suspensionsof microspheres were washed in deionized water until monodisperse(nonaggregated) suspensions were produced, and the microsphereswere suspended in a 1 mM NaHCO₃ electrolyte solution. Monodispersesuspensions were verified by light microscopy and particle-countingtechniques.

Capillary electrophoresis.
Capillary electrophoresis experiments were performed by usinga Dionex capillary electrophoresis system (Dionex, Sunnyvale,Calif.). The fused silica capillary, supplied by Phenomenex(Torrance, Calif.), was approximately 57 cm long and 75 µmin diameter. As presented by Dionex, a window for the detectorwas placed (the polyamide was burned off) 52 cm from the sampleinjection reservoir. Because more than one capillary was usedduring this study, the times required for the sample to reachthe detector window varied slightly. To ensure consistency,detection times were quantified for all experiments with a neutraldopant. A 1 mM NaHCO₃ electrolyte solution (pH 8.4) was usedfor all experiments. Sodium bicarbonate was selected becauseit buffers the pH in natural systems in which C. parvum oocystsmay be encountered. Samples were injected by gravity and wereeluted through the column at 30 kV. The detector wavelengthwas set at 210 nm by using the method of Glynn et al. (13).Mesityl oxide was used as a neutral dopant to quantify the bulkfluid movement through the column. Experiments were conductedboth in the presence and in the absence of the dopant to determinethe influence of the dopant on oocyst mobility. The capillarywas cleaned between samples by injection of 0.1 M H₃PO₄, followedby 0.5 M NaOH. The capillary was then rinsed with the 1 mM NaHCO₃electrolyte solution until a stable baseline was obtained. Allexperiments were performed in triplicate. Proper operation ofthe instrument was verified by sampling the neutral dopant andexamining the migrating medium. For example, the presence ofoocysts corresponding to a particular peak was confirmed by(i) repeating the experiment, (ii) stopping the flow of electrolytejust prior to the retention time observed for oocysts in theprevious experiment by removing the applied voltage, (iii) immediatelyremoving the capillary from the instrument and separating itjust upstream of the window, (iv) placing the migrating mediumon a microscope slide, and (v) verifying the presence of C.parvum oocysts with a light microscope at a magnification ofx1,000.

Microscopy.
A light microscope and a Petroff-Hauser counting chamber wereused to evaluate the tendency of C. parvum oocysts and CML microspheresto aggregate in distilled water and 0.5 M NaCl. The percentageof aggregates was estimated by dividing the total number ofaggregates by the total number of particles counted in the chamberfor each treatment. For purposes of comparison, the concentrationsof oocysts and CML microspheres were maintained at the sameorder of magnitude for all experiments.

Infrared spectroscopy.
Attenuated total reflectance (ATR)-Fourier transform infrared(FTIR) spectra were obtained by placing a suspension of oocystson a ZnSe ATR crystal. Oocysts were allowed to settle for 20min to obtain the best spectra. The sample chamber was flushedwith N_2(g) to minimize the effects of gaseous CO₂ and H₂O. Thespectra were then corrected for peaks resulting from residualCO₂ and H₂O. Diffuse reflectance spectra were obtained by usingfreeze-dried oocysts, which were mixed with dry KBr. Spectrawere corrected for small impurities (absorbed H₂O) in the KBr.

RESULTS

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Results

Electrophoresis.
Initially, the capillary electrophoresis technique was validatedwith CML microspheres that were approximately the same diameteras C. parvum oocysts. Microspheres with carboxyl surface functionalgroups were used because C. parvum oocysts have been reportedto have similar surface functional groups (22). Figure 1a showsa typical electropherogram of CML microspheres and a mesityloxide dopant. Peaks from 0 to ca. 30 s were the result of noisecaused by application of the 30-kV energy potential followingsample injection. This noise did not influence the parametersrequired to calculate the electrophoretic mobility (see equationabove). The peak on the electropherogram at 1.8 min correspondsto the dopant, and the peaks at ca. 3 min correspond to theCML microspheres. The electropherogram in Fig. 1a shows thatthere was a distribution of CML microspheres with various surfacecharges. Similar findings have been reported for latex spheres(13) and single-strain microbial populations (38). Figure 1ashows that the electrophoretic mobility of the CML microsphereswas ca. -3.6E-08 m²/V·min. This value is comparable tothe value expected by the manufacturer (James Goodwyn, InterfacialDynamics Corp., personal communication).

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FIG. 1. (a) Representative electropherogram of CML microspheres and mesityl oxide dopant. Peaks at the start of the electropherogram (from 0 s to ca. 30 s) result from noise caused by application of the 30-kV energy potential following sample injection. (b) Representative electropherogram of C. parvum oocysts, which were analyzed as received. The single peak at 1.9 min corresponds to the oocysts. The peak indicated by the dashed line represents the retention time for both the mesityl oxide dopant and C. parvum oocysts, showing that the two have the same value. (c) Representative electropherogram for oocysts washed in distilled water. The peak at ca. 2 min represents both oocysts and the mesityl oxide dopant.

Figure 1b is a typical electropherogram of C. parvum oocystssupplied by Waterborne Inc., and the sample was analyzed asreceived. The single peak at 1.9 min corresponds to the oocysts.The presence of C. parvum oocysts corresponding to the peakin Fig. 1b was verified by microscopic examination of the migratingmedium. The retention time for the dopant was also 1.9 min.The results indicate that the oocysts migrated through the columnvia electroosmosis with the carrier electrolyte, which impliesthat they were not charged. Similar results were obtained withadditional batches of C. parvum oocysts that were supplied byWaterborne Inc. and Pleasant Hill Farm.

Figure 1c shows a typical electropherogram for oocysts (fromthe first batch of Waterborne Inc. oocysts) that were washedin distilled water. The peak at ca. 2 min represents oocystsand the dopant. Additional peaks represent charged oocysts.Similar results were obtained with additional batches of C.parvum oocysts that were supplied by Waterborne Inc. and washedin distilled water. These results indicate that a fraction ofthe oocysts became negatively charged after they were washedwith low-ionic-strength distilled water. Washing oocysts inthe phosphate buffer solution did not impart a negative chargeto the oocysts (the electropherograms for these oocysts werecomparable to those shown in Fig. 1b). A third batch of oocysts,which were washed in distilled water by Waterborne Inc., containedboth neutral and negatively charged oocysts.

Microscopy.
A Petroff-Hauser counting chamber was used to determine if theeffects of double-layer compression could be observed with C.parvum oocysts. As when capillary electrophoresis was used,the experimental approach was validated with CML microspheres.The aggregation of CML microspheres suspended in either distilledwater or 0.5 M NaCl is shown in Fig. 2a. The results of a similarexperiment performed with C. parvum oocysts are shown in Fig.2b. Unlike the CML microspheres, the oocysts did not aggregatein the 0.5 M NaCl solution. Figure 3 shows a photomicrographof oocysts and CML microspheres in the Petroff-Hauser countingchamber suspended in the 0.5 M NaCl solution. Aggregates ofCML microspheres are plainly visible.

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FIG. 2. (a) Aggregation of CML microspheres suspended in distilled water and 0.5 M NaCl. (b) Aggregation of C. parvum oocysts suspended in distilled water and 0.5 M NaCl. Error bars represent 1 standard deviation.

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FIG. 3. Photomicrographs of CML microspheres (a) and oocysts (b) in a Petroff-Hauser counting chamber. The microspheres and oocysts were suspended in a 0.5 M NaCl solution.

Infrared spectroscopy.
We hypothesized that sucrose, which was used in the primarypurification of C. parvum oocysts from feces, may have coatedthe oocyst surface, possibly resulting in polar characteristicsdue to hydrogen bonding, without adding a surface charge. ATR-FTIRspectroscopy was used to determine if selected reagents usedin the purification process could be found on the oocyst surface.ATR spectra for Waterborne Inc. oocysts, sucrose, and Percollare shown in Fig. 4. The presence of sucrose and Percoll couldnot be identified in the spectrum for the C. parvum oocysts.Figure 5 shows a spectrum for C. parvum oocysts that were treatedwith formalin and washed. This spectrum shows that formalinwas present on the surface of the oocysts, although formalindid not impart a surface charge. Diffuse reflectance spectrawere obtained with freeze-dried oocysts mixed with KBr. Theresulting spectra indicated that there were two identifiablefunctional groups: alkyl C-H stretches and carbonyl groups (Fig.6). Based on oocyst composition, Karaman et al. (22) also reportedthe presence of carbonyl groups on the oocyst surface. The resultsindicate that the oocyst surfaces observed in this study werecapable of forming hydrogen bonds, which allowed an oocyst witha neutral surface to interact with water and/or polar domainsvia Lewis acid-base forces (14). The spectra also show thatthere were similarities between oocysts obtained from two differentsources. It has been observed that freeze-drying may alter thecomposition of the cell surface (30).

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FIG. 4. ATR spectra of C. parvum oocysts and chemicals commonly used in the oocyst purification process.

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FIG. 5. ATR spectra of formalin-treated oocysts (spectrum a) and oocysts not exposed to formalin (spectrum b). Note the absorption bands identified in spectrum a by asterisks that correspond to formaldehyde. These bands may arise from either formaldehyde adsorbed onto the surface of the oocyst or formaldehyde absorbed into the oocyst. Also note the absence of these bands in spectrum b.

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FIG. 6. Diffuse reflectance FTIR spectra of C. parvum oocysts from Cornell and Idaho.

DISCUSSION

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Discussion

The findings shown in Fig. 1b and c may help explain previousconflicting reports concerning surface charge (3, 10, 20, 23,22, 29, 33, 36). For example, using microelectrophoresis, Brushet al. (2) reported finding neutrally charged C. parvum oocysts.In contrast, other workers have reported that the surface ofthe C. parvum oocyst is exclusively negatively charged at circumneutralpH values (10, 20, 23, 22, 29, 33). It should be noted thatwashing procedures were not clearly outlined in some of thesestudies. It is possible that ionized functional groups on theoocyst are coated with up to 20 to 30 nm of a carbohydrate-richglycocalyx (3, 28). Washing oocysts in low-ionic-strength solventsmay result in removal of a glycocalyx and a concomitant increasein the electrophoretic mobility. The observed increase in C.parvum oocyst electrophoretic mobility as a result of washingin low-ionic-strength solvents is consistent with results reportedby Pembrey et al. (30) for other microorganisms. Formalin, whichis used to inactivate C. parvum oocysts, has also been reportedto increase the electrophoretic mobility of C. parvum oocysts(2). Our results suggest that formalin did not impart a significantcharge to the oocysts surface. (The electropherograms for formalin-treatedoocysts were comparable to those shown in Fig. 1b.)

The lack of surface charge indicated by the capillary electrophoresisdata was supported by stability (aggregation) experiments. Comparedto the data obtained with distilled water, the larger percentageof aggregates in the electrolyte suspension suggests that double-layercompression may have resulted in a reduction in electrostaticrepulsion between the CML microspheres. Unlike the microspheres,the oocysts did not aggregate in the 0.5 M NaCl solution. Thisobservation was consistent for oocysts received from severalsuppliers. These results support the electrophoresis data presentedabove and suggest that C. parvum oocysts may not have a significantsurface charge. The stability of oocysts in the salt solutionmay have been the result of Lewis acid-base forces, steric stabilization,or other factors (14). The polar nature of the oocyst surface(i.e., the ability to hydrogen bond) was observed in the ATRspectra, which may implicate Lewis acid-base interactions (14).A rationale for oocyst stability has been described previously.For example, Walker and Montemagno (42) reported that sorptionof oocysts to Al₂O₃ was observed only for oocysts that had theirwalls disrupted by some environmental stress, such as freeze-thawcycles. Using laterally resolved force microscopy, Considineet al. (6) found that proteinaceous materials on the oocystsurface may be responsible for steric interactions. Furthermore,Harter et al. (18) reported reversible adhesion of oocysts tosandy soils and aquifer sediments and reported only a very weakelectrostatic energy of interaction. Recently, Dai and Boll(8) reported that oocysts entrained in overland flow travelfreely in the water column and do not attach to particulatematter in the sediment load as a result of electrostatic interactions.Although the reason for the stability of oocysts remains unclear,our aggregation results are consistent with the capillary electrophoresisdata shown in Fig. 1b. The upshot is that the capillary electrophoresisdata, aggregation data, and ATR spectra obtained in this workindicate that oocysts remain stable in a water column withouta significant net surface charge.

Although ATR spectroscopy can penetrate up to several micronsbeyond the surface of a sample (17), it has been shown thatinfrared spectroscopy is useful for probing the molecular propertiesof other microorganisms (39, 40), and it was therefore considereda relevant technique for this work. The purpose of using thistechnique was to determine the presence of residual solventsused in the purification processes, whether on the surface ofthe oocysts or inside the oocysts. Because sucrose and Percollcould not be identified in the spectra for the C. parvum oocyststhat were examined, we presumed that these compounds did notbind to the oocyst surface during the purification procedureand consequently did not influence the stability of the oocysts.The spectra for formalin-treated oocysts shown in Fig. 5 didindicate the presence of formalin residue in the oocysts studied.It is conceivable that formalin penetrated into the cell wallfollowing this treatment step, but it is clear that the formalincould not be removed after several water washes. Penetrationof formalin into the interior of the cell was expected sinceformalin is used to inactivate oocysts. This may explain whyformalin-treated oocysts did not have a significant surfacecharge as a result of formalin.

The results reported here indicate that C. parvum oocysts donot always have a surface charge and that the surface chargemay be altered by washing oocysts in distilled water. This findingis noteworthy because it suggests that the stability of C. parvumoocysts in the water column is related to properties other thansurface charge, such as hydrogen bonding or steric stabilization.The observed stability without a surface charge may hinder removalof C. parvum oocysts by typical water treatment techniques becausethe charge neutralization mechanism of coagulation is not effectivefor particles with this surface characteristic. This may helpexplain why other workers (3, 31) have reported that optimalremoval of C. parvum oocysts requires the precipitate enmeshmentmechanism of coagulation. Additional research is required toidentify whether naturally occurring oocysts are charged andto evaluate the influence of natural waters on oocyst surfacechemistry. Such information might improve the detection of oocystsin environmental systems and improve the efficacy of treatmentprocesses designed to remove oocysts from water streams. Thecapillary electrophoresis technique might be useful in thisregard.

Finally, the results suggest that storage and preparation ofoocysts may affect the outcomes of bench-scale treatment studies.Current guidelines related to oocyst stocks are very specificabout many aspects of oocyst suspensions used for such experiments,but the results of this work suggest that storage and preparationaspects should be considered carefully as well.

ACKNOWLEDGMENTS

We thank Dwight Bowman, Department of Microbiology and Immunology,Cornell University, for supplying oocyst samples. We gratefullyacknowledge the guidance of J. C. Baygents, Department of Chemicaland Environmental Engineering, The University of Arizona, concerningthe use of capillary electrophoresis for biological samples.We also acknowledge the technical assistance of Michael Kelley,Department of Geography and Environmental Engineering, UnitedStates Military Academy, and Russell Forney, David Loehle, andDawn Riegner, Department of Chemistry and Life Sciences, UnitedStates Military Academy.

This project was supported by grants from the Army ResearchLaboratory and the Army Research Office.

FOOTNOTES

* Corresponding author. Mailing address: Environmental Engineering Program, Department of Geography and Environmental Engineering, United States Military Academy, West Point, NY 10996. Phone: (845) 938-2820. Fax: (845) 938-3339. E-mail: bm8375{at}exmail.usma.edu.

REFERENCES

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