Research: Science & Education Topics in Chemical Instrumentation
Capillary Electrophoresis: Part II. Applications Christine L. Copper United States Naval Academy, Annapolis, MD 21402 Kylen W. Whitaker University of Tennessee, Knoxville, TN 37996
Since its advent, capillary electrophoresis (CE), also known as capillary zone electrophoresis (CZE) or high performance capillary electrophoresis (HPCE), has been used to solve many chemical separation problems. Isolation and identification of components in samples ranging from human cells to shale oil have been achieved using CE (1, 2). A comprehensive review of the theoretical and experimental background of CE was presented in Part I of this article (3). CE is an extremely efficient separation technique. It typically employs capillaries 50 to 100 cm long with inner diameters of 25 to 100 µm. The capillary is filled with an aqueous buffer solution and suspended between two buffer reservoirs. A high voltage (typically 10 to 40 kV) is applied across the capillary causing solvent flow by a phenomenon known as electroosmosis. Injected ionic solutes are separated on the basis of their different rates of ion migration in the applied field. However, all neutral solutes migrate with a velocity equal to the electroosmotic velocity of the aqueous buffer and there-
fore are not separated without the addition of reagents that will selectively alter their mobilities. Separations of Ionizable Solutes The nature of CE renders it most useful for separations of charged solutes. Because their anions and cations will have opposing mobilities in an applied electric field, they are easily separated. Furthermore, components (with different massto-charge ratios) in a mixture of anions (or a mixture of cations) can be completely separated using CE. A striking example of one such separation is seen in Figure 1. This electropherogram, produced by Jones and Jandik, shows separate peaks for 36 different anions (4). Note the impressively brief analysis times realized, which are also possible in many other CE separations. By contrast, attempts to resolve the same 36 anions via ion chromatography yielded peaks for only three anions within a three-minute analysis period (4). Continued on page 348
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Figure 1. The power of CE is evidenced in this electropherogram showing the separation of 36 anions. Peak identification: 1 = S 2 O 3 2᎑ ; 2 = Br ᎑ ; 3 = Cl ᎑ ; 4 = SO 4 2᎑ ; 5 = NO 2᎑ ; 6 = NO 3᎑ ; 7 = MoO 4 2᎑ ; 8 = N 3᎑ ; 9 = WO 4 2᎑ ; 10 = monofluorophosphate; 11 = ClO 3 ᎑ ; 12 = citrate; 13 = F ᎑ ; 14 = HCOO ᎑ ; 15 = PO 4 3᎑ ; 16 = PO33᎑ ; 17 = ClO2᎑; 18 = glutarate; 19 = o -phthalate; 20 = galactartrate; 21 = CO32᎑; 22 = CH3COO᎑; 23 = chloroacetate; 24 = ethanesulfate; 25 = propionate; 26 = propanesulfonate; 27 = DL-aspartate; 28 = crotonate; 29 = butyrate; 30 = butanesulfonate; 31 = valerate; 32 = benzoate; 33 = L-glutamate; 34 = pentasulfonate; 35 = D-gluconate; and 36 = D-galacturonate. (Figure reprinted from J. Chromatogr. 1992, 608, 385–393; Jones, W. R.; Jandik, P.; Various Approaches to Analysis of Difficult Sample Matrices of Anions Using Capillary Ion Electrophoresis; with kind permission from Elsevier Science NL, Sara Burgerhartstraat 25, 1055 KV Amsterdam, The Netherlands.)
Figure 2. Moderate-field (500 V/cm) electropherograms of restriction enzyme (Hin fI) digests of DNA from two bacterial plasmids. Fragments elute in order of increasing size. (A) Plasmid pBR322: fragment sizes range from 154 base pairs (bp) (peak 1) to 1631 bp (peak 7). (B) Plasmid pBR328: sizes range from 145 bp (peak 1) to 660 bp (peak 8). (Figure reprinted from Rapid Separation of DNA Restriction Digests Using Size Selective Capillary Electrophoresis with Application to DNA Fingerprinting; B. K. Clark, et al.; J. Microcolumn Sep. 1994, 6, 503–513; by permission of John Wiley & Sons.)
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In the example above, the solutes studied were of reasonably low molecular weight with few charged sites per ion and showed little or no affinity for the negatively charged inner (silica) surface of the capillary. Such interactions would be a detriment to solute resolution as they contribute to band broadening. The possibility of electrostatic interactions between solutes and the capillary wall increases when larger, ionizable molecules such as proteins, which are known to interact strongly, are studied (5). Often, unmodified fused silica capillaries provide poor, if any, resolution of protein samples. For these compounds, there is a need to minimize solute– wall interactions, and numerous methods of modifying the inside silica surface have been developed. Probably, the most widely used column-coating technique, first reported by Hjerten, is the addition of a layer of non-cross-linked polyacrylamide (6). Using his method, it is possible to coat the capillary wall with a well-defined monomolecular neutral layer of polymer covalently bound to the silica surface. This polyacrylamide coating suppresses the adsorption of ionized solutes and eliminates electroosmotic flow. The termination of these two events allows resolution of previously inseparable proteins. Nevertheless, the elimination of electroosmotic flow, while helpful in some instances, can also increase analysis times. Numerous other materials have been used to coat CE columns. Most often, the purpose of these coatings is to decrease or eliminate adhesion of solutes to the capillary wall. However, some coatings have been developed to reverse the direction of electroosmotic flow or even to enhance resolution by providing unique selectivity via solute–coating interactions (7). A variation of CE that is applicable to the separation of proteins is called affinity capillary electrophoresis (ACE). ACE employs charged receptor ligands as separation buffer additives. These ligands demonstrate very specific interactions with certain injected solutes (most often proteins or peptides). Basically, the electrophoretic mobility of a solute molecule is altered upon binding with the charged ligand. The magnitude of the mobility change gives rise to ligand–solute complex stoichiometry and equilibrium constants (8). A rapidly growing application of ACE is identification of specific solutes in large mixtures of chemical variants that bind with a particular receptor molecule. Many times, the large mixtures of variants, called combinatorial collections or libraries, comprise drug candidates and the receptor molecule is of biological importance. Using ACE, many drug candidates can be quickly studied to determine which ones have a favorable interaction with the biological site and warrant further study (9, 10). Another variation of CE applicable to the separation of biopolymers such as proteins and DNA is capillary gel electrophoresis. This technique evolved from slab gel methods traditionally used to study DNA samples. Capillary gel electrophoretic analyses are faster than those employing slabs because the capillary can more efficiently dissipate heat, allowing for the use of higher applied voltages and thus faster separations. However, a problem common to these two separation methods is the difficulty of reproducibly manufacturing stable gels. Furthermore, Joule heating can degrade the integrity of these gels (11).
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To circumvent the aforementioned problems, soluble polymers have been employed in place of gels for DNA separations. Methylcellulose, for example, has been dissolved in the CE separation buffer (11). A range of pore sizes between methylcellulose molecules results, depending on the concentration of polymer used. In this approach, therefore, the separation mechanism is size selectivity. Figure 2 shows the ability to separate several fragments of DNA using methylcellulose-filled capillary columns. Fragment patterns, such as those seen in Figure 2, can be used to identify DNA digests by comparing migration times, number of fragments, and relative peak heights (11). This digest identification, known as fingerprinting, is also aided by high efficiencies and short analysis time (similar to those in the figure). Another advantage of using soluble polymers in CE separation buffers for fingerprinting is the ability to quickly and easily replace the polymer solution before each injection. This remedies problems of sample carry-over that can occur in gelfilled columns. Separations of Neutral Solutes Packed capillary columns have found utility in CE separations of neutrals (12, 13). Dubbed capillary electrochromatography (CEC), this variant typically uses reversed-phase silica packing materials. The separation mechanism is comparable to that of high-performance liquid chromatography (HPLC) in that solutes are partitioned between mobile and stationary phases. However, the CEC mobile phase is transported electrokinetically, thus eliminating the use of highpressure pumps and numerous fittings. This leads to the advantages of experimental simplicity and increased efficiency due to a pluglike (rather than parabolic) flow profile (3). Furthermore, CEC does not have a finite elution window as micellar electrokinetic capillary chromatography (MECC) and cyclodextrin distribution capillary electrophoresis (CDCE) do (see below), allowing for higher peak capacities. CEC columns can be slurry-packed with pressure or can be packed electrokinetically, as some packing materials will migrate in an applied electric field. Recently, some of the obstacles encountered with these two packing methods have been overcome by creating polymer-based stationary phases within the capillary. Polymers used for this purpose must possess both hydrophobic and ionic groups in order for retention of solutes and electroosmotic flow, respectively, to be possible (14). Surfactants are frequently added to CE separation buffers to aid in separating neutral compounds by providing micelles into which solute molecules can selectively partition. Most surfactants contain a polar and/or ionic head group and a nonpolar hydrocarbon tail (15). When employed at concentrations above the critical micelle concentration (cmc), surfactant monomers will aggregate to form assemblies known as micelles. Normal micelles are formed when the hydrophobic tails point inward and the polar head groups become the surface of these aggregates. When micelles are used in CE separations, the technique is often dubbed micellar electrokinetic capillary chromatography (MECC). MECC was first reported by Terabe et al. in 1984 when they separated several phenol derivatives using a sodium dodecyl sulfate (SDS) micellar system (16). In this system,
which is typical of all MECC systems, the micelles represent a secondary phase and have a mobility that is a combination of their intrinsic electrophoretic mobility and the electroosmotic flow of the separation buffer. In cases where negatively charged surfactants such as SDS are utilized, the mobility of the micelles is opposite to the electroosmotic flow. However, this mobility is overcome by the stronger electroosmotic flow and the micellar phase is ultimately transported in the same direction as electroosmotic flow. The result of these unequal mobilities is a two-phase system comprising an electrophoretically transported separation buffer and an electrophoretically retarded micellar phase (15 ). Resolution in MECC is a function of efficiency, selectivity, and system retention. One distinct advantage of MECC is the ease with which resolution can be manipulated by simple adjustment of the composition of the separation buffer (15 ). Both the primary (bulk solution) and secondary (micellar) phases can be rapidly changed to provide changes in selectivity (17 ). For example, different types of surfactants can be used as in Figure 3; and organic modifiers can extend the MECC elution range by altering the charge at the capillary wall, thus slowing electroosmotic flow. Cyclodextrins (CDs) can also be added to MECC separation buffers to provide desired selectivity.
Figure 3. Separation of ingredients of a cold medicine by MECC employing (A) SDS (0.1 M) and bile salts, (B) 0.1 M sodium cholate, and (C) 0.05 M sodium deoxycholate. Peaks: 1, caffeine; 2, acetaminophen; 3, sulpyrin; 4, trimetoquinol hydrochloride; 5, guaifenesin; 6, naproxen; 7, ethenzamide; 8, phenacetin; 9, isopropylantipyrine; 10, noscapine; 11, chlorpheniramine maleate; 12, tipepidine hibenzate; 13, dibucaine hydrochloride; and 14, triprolidine hydrochloride. (Figure reprinted from J. Chromatogr. 1990, 498, 313– 323; Terabe, S., et al. Separation and Determination of the Ingredients of a Cold Medicine by Micellar Electrokinetic Chromatography with Bile Salts; with kind permission from Elsevier Science NL, Sara Burgerhartstraat 25, 1055 KV Amsterdam, The Netherlands.)
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Figure 4. Separations of PAHs (structures shown) with 6 mM carboxymethyl-β-CD and 1 mM β-CD. Figure reprinted from J. Chromatogr. 1990, 498, 313–323; Terabe, S. et al. Separation and Determination of the Ingredients of a Cold Medicine by Micelllar Electrokinetic Chromatography with Bile Salts; with kind permission from Elsevier Science NL, Sara Burgerhartstraat 25, 1055 KV Amsterdam, The Netherlands.)
Figure 5. (A) Schematic of a cross microchip. This design was used to obtain the data. (B) High-speed electrophoretic separation of rhodamine B (Rb) and disodium fluorescein (Fl). (Figure reprinted from Jacobson, S. C.; Moore, A. W.; Ramsey, J. M. Anal. Chem. 1995, 67, 2059; and Jacobson, S. C., et al. Anal. Chem. 1994, 66, 1114; by kind permission of the American Chemical Society.)
CDs are a product of the enzymatic digestion of starch. They have the shape of a truncated cone with a hydrophobic cavity and a hydrophilic exterior. The most common CDs are comprised of six (α-CD), seven (β-CD), or eight (γ-CD) glucopyranose units. Inclusion complex formation between solutes and the CD’s cavity is dependent upon the geometry, size, and physicochemical properties of the solutes. Hydrophobic interactions predominate in the cavity. However, these interactions can act in concert with polar or hydrogen bonding interactions that occur with hydroxyl groups on the outer lip of the CD cavity (18). Modified CDs (including neutral and ionizable derivatives) have also been used as separation buffer additives (19, 20). In CD-MECC, neutral CDs are added to the MECC separation buffer to selectively reduce solute retention times (relative to unmodified MECC). Since the CDs have no charge and therefore no intrinsic mobility, they travel at the electroosmotic velocity of the separation buffer. This results in a three-phase system in which the separation buffer and CD phase are moving at the rate of electroosmotic flow and the electrophoretically retarded micellar system travels more slowly. Unequal partitioning of neutral solutes among these phases results in their separation. CD-MECC is especially powerful in separations of highly hydrophobic compounds
such as polycyclic aromatic hydrocarbons (PAHs), which tend to totally associate with the micellar phase, thus co-eluting at tm, in unmodified MECC (see Fig. 4) (21). Cyclodextrin modified CE (CD-CE) is used to separate solutes that are ionized at the pH of the separation buffer. Generally, charged solutes can be separated in unmodified CE because their intrinsic mobilities are quite different. However, some charged solutes have similar mobilities, which makes resolution difficult. In these cases, CDs are added to the separation buffer to alter selected solutes’ mobilities through inclusion complex formation. CDs can also be employed as chiral selectors in a CD-CE separation buffer (22). Cyclodextrin distribution CE (CDCE) employs combinations of charged and neutral CDs in the separation buffer to separate neutral compounds (23). The combination of these two types of CDs gives a system that behaves similarly to CD-MECC, as there are three phases moving at two different velocities. However, in CDCE, the charged CD phase is more selective than the micellar phase in CD-MECC, as seen in Figure 4. Moreover, many types of charged CDs (each of which can have various degrees of substitution) are readily available, suggesting that numerous and unique selectivities in CDCE are possible (24).
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Capillary Electrophoresis on a Chip Recently, CE separations have been performed in channels that are micromachined into planar, insulating substrates (as opposed to using fused silica capillary tubing) (25, 26). Typically, a CE separation using one of these devices is carried out in a channel with a cross at its origin (Fig. 5A). The cross acts as a valve and allows for the introduction of sample or separation buffer into the channel in which the separation will take place. High voltage is applied to induce electroosmotic flow, thus providing a transportation or separation mechanism. These microfabricated devices can be used to perform separations using only picoliters of sample, with total analysis times in the range of seconds rather than minutes (25). Separations of binary mixtures have even been performed in less than 150 milliseconds (Fig. 5B) (26). In addition, multiple functional elements can be integrated into a monolithic device—for example, filters, mixers, reactors, valves, pumps, separators, and detectors. Consequently, an entire analysis from sample in to information out can be performed using a single chip. Discussion CE has been proved to be a highly versatile, low-cost separation technique. With simple modification to the capillary or through simply changing running buffers, a wide variety of solutes can be separated. CE can be used with a variety of detection schemes (UV-vis, fluorescence, electrochemical, and mass spectrometric), making it adaptable to many situations. It is hoped that some problems with reproducibility and insufficiently low limits of detection for some analytes will be solved as more research is performed. Sample preconcentration and fluorescence-based detection methods are some of the ways that limits of detection in CE can be enhanced. Improvements in capillary coating procedures and CE instrumentation have already helped ease problems with reproducibility of retention times. In the near future CE can be expected to compete with HPLC and other separation techniques for use in routine separations of analytes. Acknowledgments We are grateful for the funding provided by the United States Naval Academy Research Council under Office of Naval Research Grant N0001496WR20008 and by the Division of Chemical Sciences, United States Department of En-
ergy, under Grant DE-FG02-96ER14609. We also wish to acknowledge Stephen Jacobson of Oak Ridge National Laboratory for his gracious contribution of the planar CE figures in this article. Literature Cited 1. Ewing, A. G.; Mesaros, J. M.; Gavin, P. F. Anal. Chem. 1994, 66, 527A–537A. 2. Copper, C. L.; Staller, T. D.; Sepaniak, M. J. Polycyclic Aromat. Compd. 1993, 3, 121. 3. Copper, C. L. J. Chem. Educ. 1998, 75, 343. 4. Belen’kii, B. G.; Belov, Y. V.; Kasalainen, G. E. J. Anal. Chem. 1996, 51, 753–769. 5. Schoneich, C.; Huhmer, A. F.; Rabel, S. R.; Stobaugh, J. F.; Jois, S. D.; Larive, C. K.; Siahaan, T. J.; Squier, T. C.; Bigelow, D. J.; Williams, T. D. Anal. Chem. 1995, 67, 155R–181R. 6. Hjerten, S. J. Chromatogr. 1985, 347, 191–198. 7. St. Claire, R. L. Anal. Chem. 1996, 68, 569R–586R. 8. Chu, Y. H.; Lees, W. J.; Stassinopoulos, A.; Walsh, C. T. Biochemistry 1994, 33, 10616–10621. 9. Borman, S. Chem. Eng. News 1996, 74(7), 29-73. 10. Carson, S.; Cohen, A. S.; Belenkii, A.; Ruiz-Martinez, M. C.; Berken, J.; Karger, B. L. Anal. Chem. 1993, 65, 3219. 11. Clark, B. K.; Nickles, C. L.; Morton, K. C.; Kovac, J.; Sepaniak, M. J. J. Microcolumn Sep. 1994, 6, 503–513. 12. Yan, C.; Dadoo, R.; Zhau, H.; Rakestraw, D. J.; Zare, R. N. Anal. Chem. 1995, 67, 2026–2029. 13. Whitaker, K. W.; Sepaniak, M. J. Electrophoresis 1994, 15, 1341– 1345. 14. H. Lino, J. L.; Chen, N.; Ericson, C.; Hjertén, S. Anal. Chem. 1996, 68, 3468–3472. 15. Sepaniak, M. J.; Powell, A. C.; Swaile, D. F.; Cole, R. O. In Capillary Electrophoresis: Theory and Practice, Grossman, P. D.; Colburn, J. C., Eds.; Academic: New York, 1992, Chapter 6. 16. Terabe, S.; Otsuka, K.; Ichikawa, K.; Tsuchiya, A.; Ando, T. Anal. Chem. 1984, 56, 111. 17. Cole, R. O.; Sepaniak, M. J.; Hinze, W. L.; Gorse, J.; Oldiges, K. J. Chromatogr. 1991, 557, 113. 18. Bereuter, T. L. LC-GC 1994, 12, 748. 19. Ward, T. Anal. Chem. 1994, 66, 633A. 20. Novotony, M.; Soini, H.; Stefansson, M. Anal. Chem. 1994, 66, 646A. 21. Copper, C. L.; Sepaniak, M. J. Anal.Chem. 1994, 66, 147. 22. Belder, D.; Schomburg, G. J. Chromatogr. 1994, 666, 351–365. 23. Sepaniak, M. J.; Copper, C. L.; Whitaker, K. W.; Anigbogu, V. C. Anal. Chem. 1995, 67, 2037. 24. Whitaker, K. W.; Copper, C. L.; Sepaniak, M. J. J. Microlumn Sep. 1996, 8, 461–468. 25. Effenhauser, C. S.; Manz, A.; Widmer, H. M. Anal. Chem. 1993, 65, 2637. 26. Jacobson, S. C.; Hergenroder, R.; Koutny, L. B.; Ramsey, J. M. Anal. Chem. 1994, 66, 1114.
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