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Trace Organic Analysis

Revised 4/17/97

Applications of Capillary Electrophoresis/Laser-Induced Fluorescence Detection to Groundwater Migration Studies

W. C. Brumley,* P. L. Fergusona and A. H. Grangeb

* U.S. Environmental Protection Agency, National Exposure Research Laboratory, Environmental Sciences Division, P.O. Box 93478, Las Vegas, NV 89193-3478

J. R. Donnelly, Lockheed-Martin Environmental Services Group, 980 Kelly Johnson Drive, Las Vegas, NV 89119

John Farley, Department of Physics, University of Nevada-Las Vegas, Las Vegas, NV 89154

a This work was performed while P.L.F. held an internship with the UNLV Environmental Studies Program.

b This work was performed while A. H. G. held a National Research Council/NERL (CRD-LV) Senior Research Associateship.

Original citation:
W. C. Brumley, P. L. Ferguson, A. H. Grange, J. L. Donnelly, and J. W. Farley, "Applications of Capillary Electrophoresis/Laser-Induced Fluorescence Detection to Groundwater Migration Studies," J. Cap. Electrophoresis, 3, 295-299 (1996)

Indexing terms:

Capillary electrophoresis, fluorescent dyes, groundwater tracers, laser-induced fluorescence, solid-phase extraction, TinopalTM

Abbreviations:
CE, capillary electrophoresis; CZE, capillary zone electrophoresis; HPLC, high performance liquid chromatography; LIF, laser-induced fluorescence; SPE, solid-phase extraction; UV, ultraviolet.

ABSTRACT

Capillary electrophoresis (CE) has been applied to the determination of groundwater migration based on laser-induced fluorescence (LIF) detection and traditional spectrofluorimetry. Detection limits of injected dye/fluorescent whitening agent (tinopal) in the low ppt ranges have been accomplished with both CE/LIF based on the HeCd laser and with a spectrofluorometer. This approach was used for a real world problem in determining groundwater migration between adjacent RCRA and Superfund sites. Fluorescent dyes were injected into wells and were discovered in monitoring wells by using extracting pads that adsorbed the dye. The approaches based on CE/LIF exhibits increased specificity over existing approaches due to the separation and unique migration time of the dye. Additional studies were aimed at achieving sub-ppt levels in the water using solid-phase extraction and field-amplified injection techniques.

INTRODUCTION

Groundwater migration is an important parameter in determining the distribution and fate of environmental pollutants originating from various waste sites or in tracking plumes that result from specific sources.1-3 The analytical problem can be approached by using various types of tracer compounds that can be determined very sensitively. Examples of such methodology include the use of fluorescent dyes or whiteners, radioactive tracers, and, historically, the use of microscopic organisms or solid particles such as chaff.2

Fluorescent dyes are a convenient choice because of the ease of sensitive detection. Spectrofluorimetry, high performance liquid chromatography (HPLC)/UV or fluorescence detection, and capillary LC/fluorescence detection have been used.2-6 For HPLC or capillary LC, the retention of dye analytes is enhanced using ion-pairing techniques. Logically, applications of capillary electrophoresis/laser-induced fluorescence (CE/LIF) detection should be ideal for the determination of anionic (or cationic) dyes. In free-zone CE, the dyes are separated very simply on the basis of their mobilities in aqueous buffers. Among several reports on CE/LIF detection techniques are two recent papers based on detecting fluorescent dyes very sensitively.7,8 Environmental applications of CE have been reviewed9 and techniques of sample preparation for environmental analysis using CE have also been reviewed.10

In this work, we applied CE/LIF detection to the determination of TinopalTM (a fluorescent whitener) as a tracer compound for groundwater migration, and compared the performance of CE/LIF with the more traditional spectrofluorimetry. Feasibility of determining dyes directly in the water through solid-phase extraction (SPE) preconcentration is also shown. To our knowledge, this is the first application of CE/LIF to groundwater migration using fluorescent dyes or whiteners.

EXPERIMENTAL SECTION

Chemicals

All organic compounds were obtained from Aldrich Chemical Company, Inc. (Milwaukee, WI, USA) and Molecular Probes (Eugene, OR) unless otherwise specified. Other chemicals were from standard sources of supply, and all were used as received. Deionized water (ASTM Type II) was used for all aqueous solutions. Buffer solutions were freshly prepared at least weekly. Solutions of dye standards were prepared from solid dye and serially diluted.

Capillary Electrophoresis

An optical bench with components and light-tight enclosure was constructed and used for all LIF experiments. The overall design was based on that of Nie et al.,11 but modified in certain respects. A special cylindrically symmetric capillary holder was machined to insure optical alignment with the laser beam, lenses, slit, and the newly installed capillary with window made by removing the polyamide coating. The entrance lens was an L-50X and the exit lens was an M-60X (Newport, Irvine, CA). A mass spectrometer slit (VG 7070EQ source slit, Micromass, Beverly, MA) was mounted in a special holder to exclude wall fluorescence transmitted to the detector as previously described.3 The bench was fitted with a fused-silica capillary (Polymicro Technologies, Phoenix, AZ) 57 cm X 75-m I.D., 50 cm to the detector, with LIF detection using the 354-nm line of the HeCd ion laser, 3 mW, model 7203N (Liconix, Santa Clara, CA ) and two 450DF100 (i.e., 50% transmission at 400 nm and 500 nm) emission filters (Omega Optical, Battleboro, VT) in series. The power supply for CE was a Series EL (Glassman, White House Station, NJ). The temperature of the capillary was 25 C, and electrophoretic runs were about 10 minutes at 20 kV using a 40 mM borate buffer at pH 9.1, which was prepared by weighing 0.381 g of sodium tetraborate decahydrate and dissolving in 100 mL of deionized (DI) water. The capillary was equilibrated with running buffer at the start of each experiment, and washed extensively (minimum 2 min each) with 0.1 M sodium hydroxide, DI water, and running buffer between analyses. Rinsing was accomplished by using capillary rinse reservoirs (SGE, Austin, TX) at about 20 psi of nitrogen pressure by fitting the injection end of the capillary through septum seals on the reservoirs. Migration times, peak widths, and detection limits were either read directly from the monitor or from printouts of the data system (Austin P-90 computer, Austin, TX loaded with Beckman System Gold, Ver. 8.1, Fullerton, CA) with data acquisition using a Beckman 406 analog interface (2 V full scale output). The photomultiplier tube (PMT) was model R928 (185-900 nm) fitted with socket E0719-21 (Hamamatsu Photonics Systems, Bridgewater, NJ) and was operated at 900 V with power supply model 230-03R (Bertan, supplied by Hamamatsu). The current amplifier for the optical signal was a model 428 (Keithley Instruments, Cleveland, OH) and used the auto-current suppression facility of the amplifier to zero the background signal and maintain full amplifier dynamic range (0 to 10 V output). Corrected peak areas, as computed by using a spreadsheet (peak area multiplied by the velocity of the ion [length to the detector divided by time]), were normalized to the corrected peak area of the internal standard (7-hydroxycoumarin-4-acetic acid) as a control for the variations in the nominal volumes of the gravity injections (10 sec to 40 sec at 30 cm height corresponding to about 40 to 170 nL). A microampere electrometer with 0 to 1 V output (1 V = 200 A) was constructed for measuring current through the capillary and was also interfaced to the Beckman 406 to provide a record of the electrophoretic current.

Groundwater Injection Studies

Four different dyes (Tinopal CBS-X, fluorescein (acid yellow 73), rhodamine WT, and eosin Y) were injected into four wells at a RCRA site and were monitored at three wells at a nearby Superfund site. Each dye (10 - 30 lbs) was injected with 2000 L of water resulting in a 10 mM concentration level for each dye. Thereafter, 8000 L of water was used to flush the dyes into the surrounding groundwater. Samples were taken before injection and for about two months afterward resulting in about 22 samples. Here, we report results only for the Tinopal tracer in a selection of 6 samples. Samples consisted of vial samples of water, "receptors," and 1-L water samples at the monitoring wells. Results are reported here for the extracts of the receptors. The "receptors" consisted of fiberglass mesh filled with coconut charcoal and weighted to remain near the bottom of the well. The standard protocol called for 1 g of charcoal from the receptor to be extracted with 10 mL of a solution consisting of 5:3:2 (propanol:water:concentrated ammonium hydroxide). Results for Tinopal are reported as ppt-levels in the 10-mL extractant.

SPE Sample Handling

Tinopal was isolated from spiked DI water samples using SPE with styrene-divinylbenzene (SDVB) extraction disks. The disks were prepared following the manufacturer's directions by soaking in 10 mL acetone and then pulling the solvent through the disk. The process was repeated with10 mL methanol and then water without letting the disk become dry. Samples were then added, adjusted to pH 1.0, and pulled through at 25 mm Hg vacuum. The disks were dried for 2 min and then eluted twice with 6 mL of methanol. The methanol eluant was concentrated as necessary with a gentle stream of nitrogen with gentle warming to achieve a recovered concentration within the detection limits of the CE/LIF technique.

Spectrofluorimetry

Fluorescence emission spectra were recorded using a Spex Fluorolog 2, which is a double monochrometer instrument (excitation and emission) with 450 W Xe lamp (Instruments S. A., Edison, NJ). Groundwater studies used maximum sensitivity (1-cm cells) with 5.0-mm slits (8.5-nm bandpass) and a synchronous scan (1 sec/nm, 1 sec integration time) with a Stokes shift of 15 nm (optimized for the visible fluorescent dye tracers). Other reported data used 1.25 mm slits (2.13 nm bandpass). The absorption maximum of Tinopal was determined on a Lamda 9 (Perkin-Elmer, Norwalk, CT) spectrometer.

RESULTS AND DISCUSSION

Migration Studies

Fluorescent dyes were injected into wells at the RCRA site, and monitoring wells located at the nearby Superfund site were sampled regularly by means of water samples and by receptors placed in the well. Tinopal was found to have an absorption maximum at 334 nm in DI water. The fluorescence emission maximum of Tinopal for excitation at 354 nm was found to be 427 nm. The Raman band of water at an excitation of 354 nm had an emission maximum at 403 nm, and therefore this band contributes substantially to the background signal.

The "standard" protocol involved spectrofluorimetry with synchronous scans. The concentrations of the dyes in the groundwater were so low that only dyes adsorbed onto pads could be detected (in the eluting solvent system). Preconcentration using SPE is also considered later in this work. Based on 1 g charcoal extracted with 10 mL of the extracting solvent, results are reported as ppt in the extractant. Table 1 gives results by spectrofluorimetry, CE/LIF, and corrected migration times for Tinopal in environmental samples compared with a standard run on that day. The agreement between spectrofluorimetry and CE/LIF is fairly good considering that the former is estimated by peak height against a fairly high background level. An example of this is Figure 1 for a standard and a sample.

Table 1. Quantitation of Tinopal in Groundwater Samples (receptors).

Sample
ppt Spectro-fluorimetry
ppt CE/LIF
MT (min) Tinopal
Corrected MT Tinopal
Tinopal Standard MT
% error MT Tinopal
1
70.0
196.
5.252
5.233
5.182
-0.98
2
70.0
-- b
--
--
--
--
3
100.
171.
5.062
6.006
5.979
-0.45
4
N/Da
88.3
5.190
5.387
5.384
-0.05
5
N/D
112.
5.404
5.386
5.384
-0.05
6
N/D
59.0
5.631
5.325
5.332
0.13

a N/D = not detected
b Not run by CE/LIF

Image of chromatograph - contact brumley.william@epa.gov for further information

Figure 1. Spectrofluorimetry results using a synchronous scan with a difference of 15 nm for Tinopal recovered from groundwater (receptor) and designated sample 1. Inset enlarges view around the vertical marker.

The power of coupling a separation with fluorescence detection is an obvious advantage of CE/LIF.

Separation Conditions

A free-zone CE separation using a 40 mM borate buffer was chosen for Tinopal. With this system, Tinopal gave a migration time of about 5.4 min. Figure 2 reveals a typical response for injections of 1 X 10-8 M and 1 X 10-9 M standards, each having been prepared with internal standard at 1 X 10-7 M and buffered at 10% of the running buffer concentration.

Tinopal

Image  of molecular line structure - contact brumley.william@epa.gov for further information

Image of chromatograph - contact brumley.william@epa.gov for further information

Image of chromatograph - contact brumley.william@epa.gov for further information

Figure 2. CE/LIF electropherograms for Tinopal standards at (a) 1.0 X 10-8 M and (b) 1.0 X 10-9 M.

Two impurities in the internal standard that migrated earlier than Tinopal further bracketed the tinopal response, yielding great certainty about its migration time (or corrected migration time).

Detection Levels Achieved in DI Water and Environmental Waters

With a molecular weight of 562, Tinopal at 1.00 X 10-9 M corresponds to 562 ppt. Typical quantitations are illustrated for each of six separate 1-g samplings of the "pads" or receptors and are given in Table 1 (sample 2 was not run by CE/LIF). The responses from samples 4 and 6 are shown in Figure 3.

Image of chromatograph - contact brumley.william@epa.gov for further information1

Image of chromatograph - contact brumley.william@epa.gov for further information

Figure 3. Samples 4 and 6 quantitated at (a) 88.3 ppt and (b) 59.0 ppt.

Tinopal was clearly detected in the samples down to the detection limit of the technique represented by 6. The separation has eliminated the fluorescent background evident in the spectrofluorimetric results, and this enabled the CE/LIF technique to detect tinopal below the screening level of spectrofluorimetry. This level probably represents a background level of Tinopal in groundwater.

The detection of tracer dye helped to establish that groundwater was migrating from one site (RCRA) to the other (Superfund) based on the arrival of Tinopal in one of the monitoring wells at the Superfund site. The findings of spectrofluorimetry and CE/LIF were in qualitative agreement. The greater specificity provided by CE/LIF affords increased confidence in the findings relative to spectrofluorimetry.

The SPE results are summarized as 61.1% 9.4% recovery at the 10-10 M level for standards from 100-mL spiked water samples. Further results for a spiked water sample at 10-12 M were qualitatively similar with a recovery of 45.9%. This would potentially extend the detection of Tinopal in environmental waters to below the ppt level (ca. 0.5 ppt). It would be feasible to achieve greater preconcentration with an even larger sample (1 L) and with smaller final volumes of isolate to inject. Levels of 1 to 10 ppq would be approachable, but were not attempted here because of the tendency of Tinopal to adsorb to glass surfaces at trace levels. No attempt to determine Tinopal in the collected water samples (as opposed to the receptors) was made in view of the long storage time for the samples. Previous studies have indicated that low Tinopal concentrations decrease by greater than 90% after two days in water solution.6

Variations in Migration Times

Under free-zone electrophoresis, variations in migration times (MTs) of target ions are principally due to variations in the EO flow. Corrections to this variation were carried out12 and corrected migration times for Tinopal are given in Table 1 along with the quantitations. Usually, agreement is within 0.3% for predicted versus observed MTs. Variations outside this range may be due to a number of factors or could indicate that the peak assignment is questionable.

Advantages/Disadvantages of CE/LIF Versus Spectrofluorimetry

CE/LIF offers greater specificity in determining dyes and eliminating background signal levels. The ease of sample handling is very similar for the two techniques. Availability of a CE instrument and proper laser excitation sources may be current drawbacks for some laboratories. Consideration of using CE/LIF as a confirmatory technique for spectrofluorimetry is recommended. Spectrofluorimetry is capable of high throughput with about 5 min per sample required compared with about 10 min per sample for CE. An advantage of commercial CE instrumentation is the ability to use an autosampler to run samples unattended. Spectrofluorimetry is a straightforward technique that can be used with minimal training and is recommended for screening purposes.

CONCLUSIONS

CE/LIF detection offers a sensitive, specific approach to groundwater migration studies involving fluorescent tracer dyes. The analytical problem of detecting fluorescent tracer compounds (ionic analytes) in water samples is perfectly matched to separations involving free-zone CE. The detection limits afforded by CE/LIF are as good or better than those of traditional spectrofluorimetry, while providing increased specificity because of the separation based on ion mobilities. Future studies of groundwater migration should be planned with this capability in mind. Choice of tracer dyes can be based on the availability of lasers matched to the absorption maximum of appropriate dyes.

NOTICE

The U.S. Environmental Protection Agency (EPA), through its Office of Research and Development, funded and performed the research described here. This work has been subjected to the Agency's peer review and has been approved as an EPA publication. The U.S. Government has the right to retain a non-exclusive, royalty-free license in and to any copyright covering this article. Mention of trade names or commercial products does not constitute endorsement or recommendation for use.

REFERENCES

  1. Smart, P. L., Laidlaw, I.M.S. An evaluation of some fluorescent dyes for water tracing. Water Resources Res. 1977; 13:15-33.
  2. Davis, S. N., Campbel, D. J., Bentley, H. W., Flynn, T. J., Ground Water Tracers, 1985, Natl. Water Well Assoc., Worthington, OH; Cooperative agreement CR-810036, U.S. Environmental Protection Agency, Robert S. Kerr Environmental Research Laboratory, Ada, OK 74820.
  3. Aulenbach, D. B., Bull, J. H., Middlesworth, B.C. Use of tracers to confirm ground-water flow. Ground Water 1978; 16:149-157.
  4. Laane, R.W.P.M., Manuels, M. W., Staal, W. A procedure for enriching and cleaning up rhodamine B and rhodamine WT in natural waters using a Sep-Pak C18 cartridge. Water Res. 1984; 18:163-165.
  5. Van Soest, R.E.J., Chervet, J. P., Ursem, M., Suijlen, J. M. Analysis of fluorescent water tracers using on-line preconcentration in micro HPLC. LCGC International 1996; September: 586-593.
  6. Jones, T. L., Master's Thesis: The detection and evaluation of fluorescent brighteners as organic tracers in alkaline groundwaters. Department of Chemistry, University of Nevada-Las Vegas, Las Vegas, NV, December, 1991.
  7. Timperman, A. T., Khatib, K., Sweedler, J. V. Wavelength-Resolved fluorescence detection in capillary electrophoresis. Anal. Chem. 1995; 67:139-144.
  8. Chem, D., Dovichi, N. J. Single-Molecule detection in capillary electrophoresis: molecular shot noise as a fundamental limit to chemical analysis. Anal. Chem. 1996; 68: 690-696.
  9. Brumley, W. C. Environmental applications of capillary electrophoresis for organic pollutant determination. LCGC 1995; 13:556-568.
  10. Brumley, W. C. Techniques for handling environmental samples with potential for capillary electrophoresis. J. Chromatogr. Sci. 1995; 33:670-685 (1995).
  11. Nie, S., Dadoo, R., Zare, R. N. Ultrasensitive fluorescence detection of polycyclic aromatic hydrocarbons in capillary electrophoresis. Anal. Chem. 1993; 65:3571-3575.
  12. Brumley, W. C., Brownrigg, C. M. Electrophoretic behavior of aromatic-containing organic acids and the determination of selected compounds in water and soil by capillary electrophoresis. J. Chromatogr. 1993; 646:377-389.

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Author: William C. Brumley


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