In the Laboratory edited by
Topics in Chemical Instrumentation
David Treichel Nebraska Wesleyan University Lincoln, NE 68504
Quantitative Analysis of Non-UV-Absorbing Cations in Soil Samples by High-Performance Capillary Electrophoresis
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An Experiment for Undergraduate Instrumental Analysis Laboratory Jason A. Gruenhagen, Dana Delaware, and Yinfa Ma* Division of Science, Truman State University, Kirksville, MO 63501; *
[email protected] Quantitative analysis of cations has traditionally been performed by using ion chromatography (1, 2), reversedphase high-performance liquid chromatography (RP-HPLC) (3), atomic emission and atomic absorption spectrophotometry (4, 5), potentiometric titration (6 ), and thin-layer chromatography (7). But despite the continuous development of these techniques, there are still situations in which high-performance capillary electrophoresis (HPCE) is the method of choice (8, 9). HPCE is the descendent of many electrophoresis and chromatography techniques. Although the technique was described in 1967 (10), it did not experience significant growth until the 1980s (11). During the last ten years, the applications of HPCE in chemical and biological research have increased exponentially. They include analysis of DNA fragments, proteins, peptides, synthetic polymers, inorganic anions and cations, alkaloids, pesticides, herbicides, and other organic molecules in clinical chemistry, pharmaceutical studies and controls, forensic science, environmental chemistry, and industrial hygiene (12). HPCE has quite a few advantages over gel electrophoresis, HPLC and other conventional techniques: 1. fast, efficient separation of ionic species; 2. fast separations of macromolecules important in analytical biotechnology; 3. ultrasmall amounts of sample used (µL sample volumes and nL injection volumes); 4. ease of automation, and 5. very small amounts of solvent wastes generated compared to HPLC.
It is important for undergraduates to learn the HPCE technique and to know how to apply it for sample analysis. To date, few experiments involving HPCE have been introduced into undergraduate teaching laboratories (13–15). In this paper, we describe how we designed an experiment to separate non-UV–vis-absorbing cations in soil samples by HPCE to (i) help students understand the HPCE concepts being taught in the lecture; (ii) show students how experimental conditions and buffer additives influence the separation; and (iii) explain the principle of indirect detection. The concept of capillary electrophoresis has been clearly described in several books (8, 9) and by Copper et al. in this Journal (16, 17 ). Indirect detection has been discussed in a number of publications and has been proved to be simple, universal, and sensitive (18–23). For determination of cations, a UV-absorbing cation in the running buffer maintains a very high UV absorption background when no analytes are present.
When a non-UV-absorbing cation migrates to the detection window, a negative signal is observed owing to the equal replacement of the UV-absorbing background cation. The area of the negative peak is proportional to the concentration of non-UVabsorbing cations in the sample. However, no previous experiment has been designed for undergraduate students to determine the non-UV-absorbing cations in a real sample by using HPCE with indirect detection. Our analysis of soil samples nicely incorporates indirect detection into an experiment suitable for undergraduate teaching laboratories. Because soil is a complex mixture of mineral and organic substances, its analysis presents a challenge to prospective soil scientists and undergraduates alike. Soil analysis involves more than 60 of the naturally occurring chemical elements (24). Some of these are necessary for the nutrition and growth of plants, but others are toxic. Cations are frequently quantified to assess soil quality, because to apply the optimal amount of the correct fertilizer, a farmer needs to know the concentration of cations such as Ca2+, Mg2+, K+, Na+, NH4+, Mn2+, and Zn2+ in the soil. The importance of these cations is discussed in most introductory soil textbooks (25, 26 ). This experiment uses HPCE to analyze a group of nonUV-absorbing cations in soil samples. It is well suited for an instrumental analysis class. Analyzing a real soil sample is very attractive to students and makes them understand the challenge of routine analysis of real samples. Through this experiment, students can learn quite a few important things: how HPCE works, principles of indirect detection, peak identification for electropherograms, standard addition for quantitation, the effect of buffer additives on separation (in this case crown ether), preparation of real samples, etc. We have performed this experiment for more than two years, with excellent results. All the students achieved good separation and over 95% of them obtained satisfactory data (within ±8% deviation from professor’s results). Their responses are very positive: students claim to have learned important techniques and concepts through the experiment, especially the influences of buffer additives to the cation separation, and some are so excited about their data that they are planning to do undergraduate research using CE techniques. Experimental Procedures
Reagents, Standards, and Materials All chemicals were of analytical reagent or ACS grade unless stated otherwise. Deionized (DI) water was prepared
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with a Milli-Q system (Millipore, Bedford, MA). Imidazole and hydroxypropylmethylcellulose (HMC) (100 cP) were purchased from Sigma (St. Louis, MO). 18-Crown-6 was purchased from Aldrich (Milwaukee, WI). All standard nitrate salts of cations were purchased from Fisher Scientific (Fairlawn, NJ). Soil samples were obtained locally. All glassware and plasticware was washed carefully with 20% nitric acid and DI water. Standard stock solutions of cations were prepared by dissolving ammonium nitrate, potassium nitrate, sodium nitrate, calcium nitrate, magnesium nitrate, manganese nitrate, and zinc nitrate in DI water to give solutions containing 20.0 mM Ca2+, 3.0 mM Mg2+, 5.0 mM K+, 12.0 mM Na+, 2.0 mM NH4+, 2.0 mM Mn2+, and 5.0 mM Zn2+.
Preparation of Background Electrolyte The background electrolyte (BGE) was prepared by dissolving 0.425 g of imidazole and 0.066 g of 18-crown-6 ether in 200 mL of DI water in a 300-mL beaker; 0.75 g of HMC (100 cP) was then dissolved in the imidazole–18-crown-6 ether solution in a manner to avoid clumping.W This solution was adjusted to pH 7.2 with 5% H2SO4, then to pH 6.02 with 1.0 M HCl. Finally, the solution was diluted to 250 mL to prepare a solution containing 20 mM imidazole, 1.0 mM 18-crown-6, and 0.3% HMC with a pH of 6.02. This BGE solution was filtered and degassed before use. Preparation of Soil Samples One to two grams of soil sample was weighed accurately and placed into a 100-mL beaker. Any organic-bound cations were released by addition of concentrated nitric acid. After evaporation of the nitric acid, the cations were extracted with
Figure 1. Electropherogram of cation standards by HPCE with indirect photometric detection. Conditions: 57 cm × 75 mm i.d. pretreated column (50 cm to detector); 10-s vacuum injection; 17-kV separation voltage; scan range: 190–300 nm; analysis wavelength 214 nm; running buffer: 20 mM imidazole + 1.0 mM 18-crown-6 + 0.3% HMC (100 cP), pH = 6.02. Peaks: A = NH4+ (0.6 mM); B = K+ (1.67 mM); C = Na+ (4 mM); D = Ca2+ (6.67 mM); E = Mg2+ (1.0 mM); F = Mn2+ (0.6 mM); G = Zn2+ (1.6 mM).
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four 3-mL portions of 0.01 N HCl. The combined extracts were centrifuged at 3300 rpm (ca. 1400 × g) for 5 min to remove any remaining sediment. The supernatant was then filtered through a 0.45-µm syringe filter into a 25-mL volumetric flask. The solution was diluted to the mark with 0.01 N HCl. At this point, the solution was ready for injection into the HPCE column for analysis.
Instrument A P/ACE 5500 capillary electrophoresis system (Beckman Instruments, Fullerton, CA) with a photodiode array was used. A positive high voltage was applied to the capillary by maintaining the injection end at a positive high potential while the cathodic end was held at ground potential. A capillary 57 cm long with 75 µm inside diameter (Polymicro Technologies, Inc., Phoenix, AZ) was used for separation. It was pretreated with 1.0 M sodium hydroxide for 30 min and then rinsed with water. The outer polymer coating was burned off 7 cm from the cathodic end to form the detection window. HPCE Separation The capillary for separation was kept at 15 °C. The standards and samples were injected for 10 s by the vacuuminjection method. The separation was performed at 17 kV for about 10 min. The analysis wavelength was set at 214 nm. Data were collected and analyzed with an IBM PC with P/ACE 5000 series software (Beckman Instruments, Fullerton, CA). Identification of Cation Peaks in Standards and Samples Even though injection of each standard could have been used for peak identification, the standard addition technique was utilized because of the small variations in migration times from run to run. Since it would take quite a while for students
Figure 2. Separation of cation standards by using BGE running buffer without 1.0 mM 18-crown-6. Experimental conditions and peak identifications are the same as in Fig. 1. The concentration of each cation is 2.50 mM.
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In the Laboratory
to identify all the peaks, they identified two of the cations and the information for the remaining peak identities was obtained from the instructor.
Quantitation of Cations in Soil Samples In our experiments, both the calibration curve method and the standard addition method worked well. To establish a calibration curve, students prepared a standard mixture of cations with a series of different concentrations that covered the concentrations of cations in the real soil sample, and then integrated all the peak areas. For the standard addition method, students first ran one pure unknown sample, then added a known volume and known concentration of the cations to the unknown sample and ran a second sample. By integrating all the peak areas from both electropherograms, the concentration could be calculated from the equation presented in the lecture. Since the indirect UV detection technique was used, all the peaks were negative. Therefore, the students selected for negative peaks during the peak integration process.
sample-dissolving process should be performed in a hood. Results and Discussion
This experiment uses high voltage to separate cations. However, most commercial instruments have a protection system that will disable the high voltage when the separation chamber is opened. Because strong acids (HNO3 and HCl) are used, the
An electropherogram for a standard cation mixture is shown in Figure 1. The peak identification was performed by spiking the standard solution with one of the cation standards for each run. All seven cations were well separated and detected. It should be noted that minor changes in the pH and ion strength of BGE buffer will influence the separation and detection signals significantly. This is due to the change of replacement ratio and is very common in indirect photometric detection (23). Therefore, the BGE buffer should be prepared as described. The concentration of each cation in Figure 1 was adjusted so that its concentration roughly matched the concentration in the soil sample analyzed. This made the quantification of cation concentrations in the soil easier. Even though the Zn2+ concentration was greater than those of NH4+, Mg2+, and Mn2+, the Zn2+ peak was very small. This was due to the absorbance of Zn2+ in the UV region, which made the vacancy peak smaller. Therefore, if Zn2+ is the major species of interest, the wavelength of the detection would need to be changed to one at which Zn2+ does not absorb. The function of 1.0 mM 18-crown-6 was to separate NH4+ and K+. Figure 2 shows the separation of the seven cations in a BGE without 18-crown-6. The NH4+ and K+ coelute without separation. If the electropherogram is enlarged to show only this peak (Fig. 3), NH4+ and K+ are partially separated. These two cations could not be separated by simply adjusting the pH or ionic strength of the BGE without 18-crown-6. Therefore, it is important that the BGE contain 1.0 mM 18-crown-6. It appears that 18-crown-6 interacts with only K+ and not with NH4+. Although the mechanism of interaction between
Figure 3. Expanded separation of NH4+ and K+ in BGE running buffer in absence of 1.0 mM 18-crown-6. Other experimental conditions and peak identifications are the same as in Fig. 1.
Figure 4. Electropherogram of a typical soil sample extract with 8-fold dilution. Experimental conditions and peak identifications are the same as in Fig. 1.
Determining Cation Concentration in Soil Samples by FAA To validate the HPCE method for the determination of cations in the soil samples, soil samples were also analyzed by the atomic absorption method. The instrument used was a SpectrAA-200 atomic absorption spectrophotometer (Varian Techtron, Mulgrave, Australia). Hazards
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18-crown-6 and K+ is not clear, it is possible that K+ falls into the cup of the crown ether and therefore the migration of K+ is delayed. An electropherogram obtained from a soil sample is shown in Figure 4. The concentrations of cations in the sample are listed in Table 1. The data from the FAA method are listed for comparison. The two techniques were comparable, and the FAA method validated the accuracy of HPCE for determining the cation concentration in soil samples. In summary, an HPCE method for analysis of cations in soil samples has been developed, and the experiment has been incorporated into the chemistry curriculum at Truman State University. Undergraduates in our Chemistry 322 (Instrumental Analysis) class acquire excellent results. The experiment could be divided into two labs. During the first, students could get used to the instrument, separate the standard cations, and identify the peaks. During the second lab, they could begin with the soil sample treatment and separate and quantify the cations in the soil samples. If time allows, students can process the data on the computer. Generally, our undergraduates work in pairs. Acknowledgments The project was partially supported by the National Science Foundation’s Division of Undergraduate Education (NSF-ILI grant DUE- 9651437) and internal faculty research grant awarded to Yinfa Ma from Truman State University. We sincerely thank Honglan Shi and Sarah Delaware for editing assistance. W
Supplemental Material
The student handout and notes for the instructor are available in this issue of JCE Online. Literature Cited 1. Caprioli, R.; Torcini, S. J. Chromatogr. 1993, 640, 365–369. 2. LeGras, C. A. A. Analyst 1993, 118, 1035–1041. 3. Marina, M. L.; Andres, P.; Diez-Masa, J. C. Chromatographia 1993, 35, 621–626. 4. Jackson, M. L. Soil Chemical Analysis; Prentice Hall: Englewood Cliffs, NJ, 1965; pp 429–485. 5. Robinson, J. W. Atomic Absorption Spectroscopy, Dekker: New York, 1966. 6. Serin, S.; Gok, Y.; Karabocek, S.; Gultekin, N. Analyst 1994, 119, 1629–1631. 7. Ma, Y.; Yeung, E. S. Mikrochim. Acta 1988, 3, 327–332. 8. Camilleri, P. Capillary Electrophoresis: Theory and Practice; CRC Press: Boca Raton, FL, 1993.
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Table 1. Cation Concentrations in the Soil Sample Soil Concentration/ppma
Ion
by HPCEb +
NH4
241.73
K+
699.07
by FAAc NA. 697.90
Na +
195.50
171.08
Ca2+
10,054.79
11,021.59
Mg2+
1,042.02
1,017.28
Mn2+
676.16
698.30
Zn2+
NDd
NDd
a Values are the average of 3 measurements. The SD for HPCE ranges from 0.051 to 0.093 ppm. bSee Fig. 1 for conditions. cConditions set by Varian software. d Since the zinc signal was small and overlapped with an identified peak, the students did not quantify the zinc level in the soil sample.
9. Landers, J. P. Handbook of Capillary Electrophoresis; CRC Press: Boca Raton, FL, 1994. 10. Hjerten, J. Chromatogr. Rev. 1967, 9, 122. 11. Jorgenson, J.; Lukacs, K. D. Science 1983, 222, 266–272. 12. Beal, S. C. Anal. Chem. 1998, 70 (12), 279R–300R. 13. Thompson, L.; Veening, H.; Strein, T. G. J. Chem. Educ. 1997, 74, 1117–1121. 14. Janusa, M. A.; Andermann, L. J.; Kliebert, N. M.; Nannie, M. H. J. Chem. Educ. 1998, 75, 1463–1465. 15. Boyce, M. J. Chem. Educ. 1999, 76, 815–819. 16. Copper, C. L. J. Chem. Educ. 1998, 75, 343–347. 17. Copper, C. L.; Whitaker, K. W. J. Chem. Educ. 1998, 74, 347–351. 18. Kuhr, W. G.; Yeung, E. S. Anal. Chem. 1988, 60, 2462–2468. 19. Yeung, E. S. Acc. Chem. Res. 1989, 22, 125–130. 20. Zhang, R.; Cooper, C. L.; Ma, Y. Anal. Chem. 1993, 65, 704–706. 21. Ma, Y.; Zhang, R. J. Chromatogr. 1992, 625, 341–348. 22. Xue, Q.; Yeung, E. S. J. Chromatogr. 1994, 661, 287–295. 23. Salimi-Moosavi, H.; Cassidy, R. M. Anal. Chem. 1995, 67, 1067–1073. 24. Gieseking, J. E. Soil Components—Inorganic Components, Vol. 2; Springer: New York, 1975. 25. Miller, R. W.; Donahue, R. L. Soils—An Introduction to Soils and Plant Growth, 6th ed.; Prentice Hall: Englewood Cliffs, NJ, 1990. 26. Winegardner, D. L. An Introduction to Soils for Environmental Professionals; Lewis: New York, 1996. 27. Ma, Y.; Koutny, L. B.; Yeung, E. S. Anal. Chem. 1989, 61, 1931–1934.
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