Separation of Stereoisomers

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Chapter 10

Separation of Stereoisomers 10.1. 10.2. 10.3. 10.4.

10.5.

10.6.

10.7. 10.8. 10.9.

Introduction Enantioselectivity and Absolute Configuration Separation of Enantiomers Chiral Stationary Phases 10.4.1. Cyclodextrin Derivatives 10.4.1.1. Gas Chromatography 10.4.1.2. Liquid Chromatography 10.4.2. Poly(saccharide) Derivatives 10.4.2.1. Liquid Chromatography 10.4.2.2. Supercritical Fluid Chromatography 10.4.3. Macrocyclic Glycopeptides (Antibiotics) 10.4.4. Proteins 10.4.5. Low-Mass Synthetic Selectors 10.4.5.1. Gas Chromatography 10.4.5.2. Liquid Chromatography Chiral Mobile Phase Additives 10.5.1. Liquid Chromatography 10.5.2. Thin-Layer Chromatography 10.5.3. Capillary Electrophoresis Complexation Chromatography 10.6.1. Silver Ion Chromatography 10.6.2. Ligand-Exchange Chromatography 10.6.3. Enantioselective Metal Complexation Gas Chromatography . . . Separation of Enantiomers as Covalent Diastereomer Derivatives Liquid-Crystalline Stationary Phases References

794 797 800 802 803 803 808 809 811 812 813 815 817 817 818 821 822 824 825 830 830 832 833 834 837 839

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The Essence of Chromatography

10.1 INTRODUCTION Isomers are compounds that have the same molecular formula (constitution) but different structures (configuration). Structural (or configurational) isomers such as n-butanol and diethyl ether, for example, often differ appreciably in their physical and chemical properties and, in general, are expected to be relatively easy to separate using conventional chromatographic techniques. Stereoisomers, on the other hand, differ only in the spatial configuration of substituent groups within a molecule (same atomic bond order) and usually have similar physical and chemical properties. Their separation in conventional chromatographic systems can be difficult or impossible without resorting to specific conditions Several different classes of stereoisomers can be distinguished [1-4]. Conformational isomers (conformers) can be interconverted by rotation about single bonds and correspond to different internal energy minima, for example, the chair and boat conformations of cycloalkanes. These energy minima are generally too small for distinction in chromatographic systems and single peaks are observed. Configurational isomers have significant energy barriers to interconversion and exist as distinct forms that are stable to typical separation conditions. Configurational isomers include geometric isomers, enantiomers, and diastereomers. Geometric isomers owe their existence to hindered rotation about double bonds. The isomers differ in the position of atoms (or groups) relative to a reference plane: in the c/5-isomer (or Z form) the atoms are on the same side; in the rran^s-isomer (or E form) they are on opposite sides. Geometric isomers have different physical and chemical properties and can usually be separated, if at times with difficulty, in conventional chromatographic systems. Enantiomers are stereoisomers that are non-superposable mirror images of each other. Enantiomers are chiral molecules (see Table 10.1 for a description of common terms used to describe different properties of stereoisomers [5-7]). Common examples of enantiomers are molecules containing tetrahedral carbon, silicon, sulfur, or phosphorus atoms bearing four different substituents (or three different substituents and a lone pair of electrons for Group V and VI elements), unsymmetrical sulfoxides, and substituted aziridines. Chirality may also result from the helicity of a macromolecule, such as a protein or polymer. Molecules in which there is a permanent and rigid twist in the planes of atom connectivities display chirality. So do compounds in which a pair of rings is joined at a single common (spiro) atom, when the rings are substituted so as to distinguish between their two faces. Molecules in which two ring systems are joined by a single bond are chiral, if steric effects hinder complete rotation about this bond and the two ring systems are differently substituted, so planes of symmetry are absent. Some examples of the different types of enantiomers are presented in Figure 10.1. Enantiomers have identical physical and chemical properties except for their ability to rotate the plane of polarized light to equal extents but in opposite directions. Diastereomers (or diastereoisomers) are stereoisomers that are not mirror images of each other. Diastereomers include molecules containing more than one asymmetric (chiral) center and geometric isomers. They have different physical and chemical

Separation of Stereoisomers

795

Table 10.1 Common terms to describe properties of stereoisomers Term Absolute Configuration

Achiral Anomers Atropisomers

Chirality

Chiral Additive Chiral Selector Diastereomers

Enantiomer Entiomer Excess

Racemate Stereoisomers

Description The spatial arrangement of the atoms of a chiral molecular entity (or group) and its stereochemical description. The stereochemical arrangement about a particular chiral center is normally assigned using the (/?,5)-nomenclature according to the Cahn-Ingold-Prelog rules [7]. The Fischer convention (D,L) based on glyceraldehyde as a reference is considered obsolete. It continues to be used in naming amino acids and carbohydrates, where trivial names are traditional. An object that is superposable on its mirror image Diastereomers of glycosides, hemiacetals, or related cyclic forms of sugars, differing in configuration only at C-1 of an aldose, C-2 of a 2-ketose, etc. Result when complete rotation about a single bond in a molecule is prevented by the bulk of non-identical neighboring substituents, so that a pair of enantiomers is formed. The geometric property of a rigid object of being non-superposable on its mirror image; such an object has no symmetry elements, such as a mirror plane, center of inversion, or a rotating-reflection axis. A chiral selector added as a component of a mobile phase or electrophoretic medium. The chiral component of the separation system capable of interacting selectively with the enantiomers to be separated. Stereoisomers not related as mirror images. They are characterized by differences in physical properties, and by differences in chemical behavior towards achiral, as well as chiral reagents. One of a pair of molecular entities which are mirror images of each other and non-superposable. The proportion of one enantiomer in a given mixture of both enantiomers usually expressed as a percent ee = 100(XR - xs) / (XR + Xs) where XR and xs ^ ^ the mole or weight fraction of the R- and S-enantiomers, respectively, and ee the enantiomer excess. An equimolar mixture of enantiomers. It does not exhibit optical activity. Isomers with an identical constitution but different spatial arrangement of atoms.

properties and can be separated using conventional chromatographic techniques, if at times with difficulty. The formation of diastereomers is the basis of the separation of enantiomers. The diastereomers can be formed by direct interaction with a chiral phase (formation of transient diastereomer association complexes), or after chemical transformation by reaction with a single enantiomer derivatizing reagent. Enantiomers have identical chemical properties in relation to their reactions with achiral reagents. Their physical properties are identical (e.g. solubility, partition coefficients, boiling points, etc.) So why the interest in enantiomer composition? This arises from the fact that in a chiral environment enantiomers behave as different compounds. The natural world is constructed of chiral systems that employ structure recognition mechanisms as a regulatory function [1,4,8]. The single enantiomers of racemic drugs exhibit differences in their bioavailability, distribution, metabolism, and excretion. It is often the case that one enantiomer is the more active isomer for a given

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The Essence of Chromatography

"°2S„.H

I mirror

"'

^COgH

Figure 10.1. Examples of enantiomers. A, an asymmetric carbon atom with four different substituent groups; B, a spiro compound (pair of rings containing different substituents joined at a single [spiro] atom); C, cummulenes (two or more successive double bonds with a rigid twist in the planes of atom connectivities); and D, a molecule in which rotation about a single bond is prevented by the bulk of non-identical neighboring substituents.

action, while the other enantiomer might be even active in a different way, contributing to side-effects, displaying toxicity, or acting as an antagonist. Regulatory authorities now require separate evaluation of single enantiomers throughout the drug discovery process even for products eventually marketed as racemates. Beyond pharmaceutical science, stereochemistry is recognized as a central component of the agrochemical and flavor and fragrance industries. In large part, the interest in the separation of enantiomers has to be seen as unavoidable, to gain an understanding of the properties of biologically active compounds. The success in developing stereoselective separation mechanisms has resulted in new applications exploiting the characteristic properties of enantiomers. For example, in archeology, the measurement of the degree of racemization of specific amino acids is used to date human remains [9]. Enantiomer ratios are used to establish the authenticity and quality of essential oils and flavors [10,11]. Enantiomer labeling, an application of the standard addition technique, is used for the accurate quantification

Separation of Stereoisomers

797

of enantiomers in complex matrices, in a manner similar to isotopic labeling without the need for mass selective detection [12]. In biological samples, the stability and conversion rates of enantiomers can be quite different, affecting the utility of enantiomer labeling for some sample types and conditions [13]. 10.2 ENANTIOSELECTIVITY AND ABSOLUTE CONFIGURATION The enantioselectivity of a chromatographic system is defined as the preferential interaction with the chiral selector of one enantiomer over the other. It is usually determined as the ratio of the retention factors of two enantiomers in a chiral chromatographic or electrophoretic system. The ratio of the retention factors is equivalent to the chromatographic separation factor (a). It is related to the difference in free energy of the reversible diastereomer association complexes formed by each enantiomer with the chiral selector (section 1.4.4). The separation factor provides a useful practical guide to the suitability of a chromatographic system for the separation of enantiomers. The true enantioselectivity and the chromatographic selectivity, however, are not one and the same thing, and this affects the calculation and interpretation of thermodynamic relationships for enantiomer-chiral selector interactions. The discrepancy arises from the fact that the chromatographic retention factor is composed of contributions from nonspecific interactions of the enantiomers with the chromatographic system, which are the same for both enantiomers, as well as specific enantiomer-chiral selector interactions, which are not identical when a separation is obtained [14-16]. The true enantioselectivity will almost always be greater than the ratio of the retention factors and the temperature dependence of the chromatographic selectivity factor, commonly used to evaluate thermodynamic contributions to the retention mechanism, will not be the same as the temperature dependence of the true enantioselectivity. In enantioselective complexation gas chromatography the retention increment can be used to quantitatively differentiate between non-specific and chiral interactions [16], but in other cases isolation of the two contributions to retention is difficult. Since enantiomers have identical physical and chemical properties, their separation requires a mechanism that recognizes the difference in their shape. A suitable mechanism for chromatography is provided by the formation of reversible transient diastereomer association complexes with a suitable chiral selector. To achieve a useful separation the association complexes must differ in stability resulting from a sterically controlled preference for the fit of one enantiomer over the other with the chiral selector. In addition, the kinetic properties of the formation/dissociation of the complex must be fast on the chromatographic time scale to minimize band broadening and achieve useful resolution. Enantioselectivity based on the formation of transient diastereomer complexes is commonly rationalized assuming a three-point interaction model [14,17,18]. Accordingly, enantioselectivity requires a minimum of three simultaneous interactions between the chiral selector and at least one of the enantiomers, where at least one of these interactions is stereochemically dependent. The points of interactions

The Essence of Chromatography

^

ENANTIOMER

CHIRAL SELECTOR

Figure 10.2. Stereoselective formation of diastereomer association complexes between two enantiomers and a chiral selector according to the three-point interaction model.

usually occur by hydrogen bonding, charge transfer (TI-TT), dipole-type, hydrophobic or electrostatic interactions between substituents of the chiral selector and the enantiomers. For example, in Figure 10.2, enantiomer I forms a three point interaction with the chiral selector represented by the dotted vertical lines while enantiomer II can form only a two point interaction because substituent C is in the wrong configuration to interact with the complementary group of the chiral selector. Enantiomer I, therefore, forms a more stable complex with the chiral selector than enantiomer II, and if the difference in stability of the complexes is sufficient then the two enantiomers will be separated with enantiomer II being less retained. In the modern interpretation of the three-point rule, all the pointto-point interactions do not have to be attractive. Some of them may be repulsive in nature, for example, resulting from steric crowding at the chiral center. If the chiral selector resides in the stationary phase and all interactions are assumed to be attractive, as in Figure 10.2, then the interactions between groups A and A' and B and B^ determine the retention of the weakly bound enantiomer. The third interaction between groups C and C is responsible for the retention difference and enantioselectivity. In an ideal situation, one would prefer to see the shortest possible retention for the first enantiomer and the largest possible retention for the second enantiomer. This implies that the ideal chiral selector should produce the lowest possible binding energy due to the interactions between groups A and A^ and B and B' and the highest possible binding energy for the sterically controlled interaction between groups C and C^ When interactions between groups A and A' and/or B and B' are significantly stronger than between groups C and C retention is increased without a concomitant change in enantioselectivity. Of course the strength of the individual interactions can

Separation of Stereoisomers

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and do vary significantly, and in practice are generally unknown, so the three-point model provides only a convenient framework for explaining enantioselectivity without providing a quantitative tool for its prediction. The three-point interaction model is successful at explaining enantioselectivity for well-defined chiral selectors, but does not give an accurate description of the chiral recognition mechanism for chiral polymers, also widely used in chromatography to separate enantiomers. An expansion of the original three-point interaction model into a model of molecular chiral recognition can account for these differences [18]. In the model of molecular chiral recognition, binding of the enantiomers to a region of the chiral selector need not involve formation of three simultaneous interactions between the enantiomer and chiral selector. The initial binding of the enantiomers is assumed to result in the formation of diastereomer complexes of equivalent stability. In a subsequent step, the enantiomers and the chiral selector conformationally adjust to each other to maximize the interactions and, thereby, the stability of each of the diastereomer complexes. The conformational adjustment might be driven by several factors other than single point interactions. For example, differences in the steric fit in a topographical feature of the stationary phase, such as a ravine or cavity, and the interdiction of solvent molecules in the binding process providing for the possibility of multipoint interactions in a structured solvent environment. It is the stereochemical dependence of this conformational adjustment step for the initially bound diastereomer complexes that is responsible for the difference in stability of the complexes, and therefore, separation of the enantiomers. The probability of separating a pair of enantiomers in a particular system is not easy to predict from their structure. Lack of knowledge of the geometry of the solute accessible binding structures in the chiral stationary phases, and the multiple types of cavities involved, has to date hindered a detailed understanding of the chiral discrimination process at the molecular level. The success rate in separating enantiomers is quite high in practice owing to the large number of chiral stationary phases available, and the high efficiency of chromatographic systems, that provide adequate separations with very small separation factors. The selection of the initial separation conditions remains an empirical process, generally made with scant regard for the chiral recognition mechanism. Commercial databases, such as CHIRBASE and CHIRSOURCE, containing about 61,000 separations from some 40,000 bibliographic entries (2001) can provide a useful starting point [19,20]. These databases are searchable by either two- or three-dimensional structures and substructures, indicating separation conditions suitable for related compounds when the input structure is absent from the database. NMR, and to a lesser extent X-ray crystallography, has been used to provide direct evidence for the binding of enantiomers to chiral selectors and for optimizing chiral selectors in the development of new stationary phases [8,21,22]. Quantitative structure enantiomer-retention relationships (QSERRs) have been developed to predict enantioselectivity but with only limited success [8,23]. These models provide some insight into the general importance of different binding interactions at the chiral selector, but have not proven useful for identification of

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The Essence of Chromatography

chiral selectors for particular separations. The main problem is identifying suitable molecular descriptors for the enantiomers that adequately reflect their chemical and structural properties at the detail required for useful predictions. Atomistic chiral recognition models based on molecular mechanics may provide a suitable approach for predicting enantioselectivity in the future, but so far have provided qualitative information only, suitable for establishing the elution order of a limited set of closely related compounds [24,25]. Chromatographic methods are suitable for the determination of the absolute configuration of enantiomers when only limited sample amounts are available. The sample need not be pure, if the separation system is able to separate the enantiomers of interest from interfering compounds. The determination of absolute configuration is straightforward when authentic standards for the single enantiomers are available. The enantiomer configuration is identified by co-elution, and should be confirmed by the reversal of the elution order on the same stationary phase with the opposite configuration of the chiral selector, when available. In principle, the absolute configuration of chiral samples can be determined in the absence of single enantiomers after calibration of the separation system with related reference compounds of known stereochemistry. Assignment of absolute configuration by analogy, however, is suspect, if sometimes adopted for pragmatic reasons. As a minimum, comparisons should be made at identical temperatures for gas chromatography and mobile phase compositions for other methods. The elution order of enantiomers is strongly influenced by temperature in gas chromatography and an inversion of the elution order is possible at temperatures either side of the isoelution temperature (determined in a van't Hoff plot as the temperature at which the enthalpy and entropy contribution cancel each other) [16]. In liquid chromatography temperature and mobile phase composition can affect the elution order through changes in the conformation of the stationary phase or competition at the chiral selector-binding site [26,27].

10.3 SEPARATION OF ENANTIOMERS There are two general approaches for the separation of enantiomers [1-4,28-32]. The direct method is based on the formation of transient diastereomer association complexes with a chiral selector immobilized in the stationary phase, or added to the mobile phase. The former approach requires the use of special stationary phases (section 10.4) while the later uses conventional stationary phases with special additives included in the mobile phase (section 10.5). When preparative applications are contemplated the use of immobilized chiral selectors is the more common approach. Method selection also depends on the choice of the separation mode. Table 10.2. While chiral stationary phases are the only choice for gas chromatography [16,28,33-38], and are used almost exclusively for supercritical fluid chromatography [39-43] and capillary electrochromatography [44-47], they also dominate the practice of liquid chromatography

801

Separation of Stereoisomers Table 10.2 Common methods for separating enantiomers by formation of transient diastereomer complexes Technique Gas Chromatography

Selector Stationary phase

Liquid Chromatography

Stationary phase

Mobile phase

Thin-Layer Chromatography

Stationary phase Mobile phase

Supercritical Fluid Chromatography

Stationary phase

Capillary Electrophoresis Mobile phase

Capillary Electrochromatography

Stationary phase

Type Amino Acid Derivatives Metal Chelates Cyclodextrin Derivatives Amino Acid Derivatives Low-Mass Synthetic Selectors Poly(saccharide) Derivatives Cyclodextrin Derivatives Glycopeptides Metal Chelates Proteins Helical Polymers Cyclodextrin Derivatives Metal Chelates Amino Acid Derivatives Proteins Metal Chelates Poly(saccharide) Derivatives Cyclodextrin Derivatives Proteins Amino Acid Derivatives Polysaccharide Derivatives Cyclodextrin Derivatives Glycopeptides Low-Mass Synthetic Selectors Metal Chelates Cyclodextrin Derivatives Chiral Surfactants (MEKC) Poly(saccharides) Glycopeptides Proteins Metal Chelates Poly(saccharide) Derivatives Cyclodextrin Derivatives Glycopeptides

[4,5,28-32,47,48]. Mobile phase chiral additives are dominant in capillary electrophoresis [49-57], and are preferred for thin-layer chromatography [58-60]. These techniques consume relatively modest amounts of mobile phase allowing expensive chiral additives to be used economically. Other general considerations in choosing a separation system are the volatility and solubility of the enantiomers and the efficiency of the chromatographic system. For thermally stable and volatile enantiomers, gas chromatography is the preferred technique, because its superior efficiency results in the baseline separation of enantiomers with very small separation factors. For water-soluble enantiomers of low volatility, liquid chromatography is the most important technique, owing to the availability of a wide range of stationary phases with complementary selectivity.

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The Essence of Chromatography

Liquid chromatography also provides easy scale-up for preparative applications. When only analysis is required, capillary electrophoresis provides an alternative choice, that is gaining in popularity because of its higher efficiency, although liquid chromatographic methods are considered more robust and more reliable for the quantification of minor enantiomers. For enantiomers of low water solubility and low volatility normal-phase liquid chromatography, supercritical fluid chromatography, and non-aqueous capillary electrophoresis are generally used. Traditionally, normal-phase liquid chromatography is the dominant technique for the separation of these enantiomers, but supercritical fluid chromatography is gaining increasing support because it offers higher efficiency and faster separations. Thin-layer chromatography provides an inexpensive and flexible approach for the separation of enantiomers, but is limited by low separation efficiency. Capillary electrochromatography offers higher efficiency than liquid chromatography with the same stationary phases, and has a higher loading capacity than capillary electrophoresis. It has considerable potential for enantiomer separations that may be realized more fully with on-going improvements in column and instrument technology (section 8.4). The second, and less popular approach for the chromatographic separation of enantiomers, is the indirect method (section 10.7). Separation is possible using conventional chromatographic systems after reaction of the enantiomers with a single enantiomer derivatizing reagent to form a pair of covalently bonded diastereomers. This method requires that the enantiomer contain a suitable functional group for the reaction. Indirect methods generally require more effort for validation than direct methods because of concerns over additional sources of error (enantiomer purity and stability of the derivatizing reagent, differences in reaction rates for the two enantiomers, different detector responses for the diastereomers, etc.) The direct method can be applied to enantiomers lacking reactive functional groups, and is not limited by the need for reagents of high enantiomeric purity. Although the extent of resolution on a chiral stationary phase will depend on the enantiomeric purity of the phase, separations are still possible with phases exhibiting reasonable enantiomer excess. However, there is no universal chiral phase for the separation of all enantiomers, and therefore, both the direct and indirect method of separation has their uses for specific applications.

10.4 CHIRAL STATIONARY PHASES Over 100 stationary phases with immobilized chiral selectors are commercially available with further stationary phases described in the literature. Any attempt at a general classification into a reasonable number of groups with similar properties is not easy. Probably the most widely adopted classification scheme was proposed by Wainer, and is loosely based on the chiral recognition mechanism. Table 10.3 [61]. In the following sections, a different approach has been adopted based on the type of chiral selector. Coverage is incomplete, but includes all of the widely used stationary phases.

Separation of Stereoisomers

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Table 10.3 Classification of immobilized chiral selectors (chiral stationary phases) by enantioselectivity mechanism Type I

II

III IV

V

Description Diastereomer complexes formed by attractive interactions (e.g. hydrogen bonding, TT-TT, dipole-type interactions) between enantiomers and chiral selector. Examples: low-mass synthetic selectors, amino acid derivatives, glycopeptides Primary mechanism of diastereomer complex formation is through attractive interactions (Type I) but inclusion complexes also play a role. Example: poly (saccharide) derivatives. Formation of inclusion complexes is important but secondary attractive interactions also play a role. Examples: cyclodextrin derivatives, helical polymers and crown ethers Formation of diastereomer ternary complexes involving a transition metal ion and a single enantiomer ligand (usually an amino acid) Examples: Cu(II)-L-hydroxyproline, Mn(II) /7i5[3-heptafluorobutanoyl)-(lR)-camphorate] Formation of diastereomer complexes by a combination of hydrophobic, electrostatic, and hydrogenbonding interactions with a protein. Examples: bovine and human serum albumin, aj-acid glycoprotein, enzymes

10.4.1 Cyclodextrin Derivatives Cyclodextrins are natural macrocyclic oligosacharides containing six (a-), seven (p-), or eight (y-) D-glucose monomers in a chair conformation connected via a-(l,4)-linkages, Figure 10.3 [3,28-30,33,62-67]. The glucose rings are arranged in the shape of a hollow truncated cone with a relatively hydrophobic cavity and a polar outer surface where the hydroxyl groups are located. The larger opening of the cone is surrounded by the secondary (C-2 and C-3) hydroxyl groups, while the primary (C-6) hydroxyl groups are located at the smaller end of the cone. The primary (C-6) hydroxyl groups are free to rotate and can partially block the smaller entrance to the cavity. The restricted conformational freedom and orientation of the secondary hydroxyl groups encircling the opposite end of the cavity are thought to play an important role in the enantioselectivity of cyclodextrins. Derivatizing the outer rim hydroxyls with various functional groups offers a simple mechanism to modify the properties of the natural cyclodextrins, expanding their application range for enantiomer separations, and to optimize their physical properties for use as stationary phases. A wide range of derivatized cyclodextrins is currently used as stationary phases for gas, liquid, and supercritical fluid chromatography, and as mobile phase additives in capillary electrophoresis and thin-layer, and liquid chromatography (section 10.5). 10.4.1.1 Gas Chromatography Initial attempts to use cyclodextrins and their derivatives as chiral stationary phases in gas chromatography met with limited success owing to their unfavorable melting points or decomposition temperatures [28,30,33-36]. This changed with the discovery that some peralklylated derivatives (e.g. pentyl), partially alkylated derivatives, and mixed alkylated and acylated derivatives were viscous liquids that could be coated on deactivated glass surfaces. Table 10.4 [68-73]. In the absence of oxygen contamination of the

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The Essence of Chromatography

primary hydroxy!

secondaiy hydroxy!

MW972

MWU35

MW1297

Figure 10.3. Structures of natural cyclodextrins indicating position of hydroxyl groups and cone dimensions. a-Cyclodextrin (n = 0), p-cyclodextrin (n = 1) and y-cyclodextrin (n = 3).

carrier gas these phases have good thermal stabiUty, and can be used at temperatures up to about 200°C. Some 85% of enantiomers separated by gas chromatography on derivatized cyclodextrins can be separated on one of three derivatized cyclodextrins: octakis(3-0-trifluoroacetyl-2,6-di-0-n-pentyl)-y-cyclodextrin; heptakis(di-O-methyl)P-cyclodextrin; and heptakis(2,6-di-0-n-pentyl)-P-cyclodextrin. Octakis(3-0-butryl-2,6di-0-n-pentyl)-Y-cyclodextrinis also stated to have a broad application range. It is generally accepted that enantioselectivity results from either inclusion formation and/or surface interactions occurring to different extents for the various cyclodextrin derivatives [73,74]. Although one phase may provide a better separation than others may for a particular enantiomer, several phases will often providing some separation for the

Separation of Stereoisomers

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Table 10.4 Some cyclodextrin derivatives used as chiral selectors in gas chromatography (Stationary phases commercially available on open tubular columns) Cyclodextrin derivative (i) Viscous liquids Hexalds(2,3,6-tri-(9-n-pentyl)-a-cyclodextrin Hexakis(3-0-acetyl-2,6-di-0-n-pentyl)-a-cyclodextrin Hexakis(3-0-trifluoroacetyl-2,6-di-0-n-pentyl)-a-cyclodextrin Hexakis(2,6-di-0-n-pentyl)-a-cyclodextrin Hexa]ds(0-(5-2-hydroxypropyl)-per-0-methyl)-a-cyclodextrin (mixture) Heptakis(2,3,6-tri-0-n-pentyl)-p-cyclodextrin Heptakis(2,3,6-tri-0-methyl)-P-cyclodextrin Heptakis(3-0-acetyl-2,6-di-0-n-pentyl)-P-cyclodextrin Heptakis(3-0-trifluoroacetyl-2,6-di-0-n-pentyl)-p-cyclodextrin Heptalds(di-0-methyl)-P-cyclodextrin (mixture) Heptalds(2,6-di-0-n-pentyl)-P-cyclodextrin Heptakis(0-(5-2-hydroxypropyl)-per-0-methyl)-P-cyclodextrin (mixture) Octalds(3-0-trifluoroacetyl-2,6-di-0-n-pentyl)-Y-cyclodextrin Octakis(3-0-propyl-2,6-di-0-n-pentyl)-Y-cyclodextrin Octakis(3-0-butryl-2,6-di-0-n-pentyl)-Y-cyclodextrin Octakis(2,6-di-0-n-pentyl)-y-cyclodextrin Octakis(0-(5-2-hydroxypropyl)-per-0-methyl)-y-cyclodextrin (mixture)

MAOT' 180 200 180 200 200 200 230 200 200 230 200 200 180 200 200 200 200

(ii) Cyclodextrin derivatives dissolved in a poly(siloxane) Hexakis(2,3,6-tri-0-methyl)-a-cyclodextrin 250 Hexakis(2,3-0-dimethyl-6-0-f-butyldimethylsilyl)-a-cyclodextrin 260 Hexalds(2,3-0-diacetyl-6-0-f-butyldimethylsilyl)-a-cyclodextrin 230 P-Cyclodextrin 230 Heptakis(2,3,6-tri-0-methyl)-^-cyclodextrin 250 Heptakis(3-0-n-pentyl-0-2,6-dimethyl)-p-cyclodextrin 250 Heptakis(2,3-0-dimethyl-6-0-^butyldimethylsilyl)-P-cyclodextrin 260 Heptakis(2,3-0-diethyl-6-0-r-butyldimethylsilyl)-P-cyclodextrin 230 Heptakis(2,3-0-dipropyl-6-0-r-butyldimethylsilyl)-p-cyclodextrin 230 Heptakis(2,3-0-diacetyl-6-0-f-butyldimethylsilyl)-P-cyclodextrin 230 Octakis(2,3,6-tri-0-methyl)-Y-cyclodextrin 250 Octalds(2,3-0-dimethyl-6-0-^butyldimethylsilyl)-Y-cyclodextrin 260 Octalds(2,3-0-diacetyl-6-0-f-butyldimethylsilyl)-Y-cyclodextrin 230 ^Highest temperature (°C) for isothermal operation. ^Most common poly(siloxane) solvents are poly(cyanopropylphenyldimethylsiloxane) with 14% cyanopropylphenylsiloxane monomer or poly(dimethyldiphenylsiloxane) with 20 to 35 % diphenylsiloxane monomer. Mixing ratio for the cyclodextrin derivative is 5-50% (w/w).

same enantiomers. So far no simple method has emerged to identify the optimum stationary phase for a separation from enantiomer structure. As well as separation of the natural enantiomers, achiral derivatives can be used to enhance resolution (e.g. separation of enantiomeric amines and alcohols as trifluoroacetyl derivatives) and to allow separation at lower temperatures, which are generally more favorable for enhancing enantioselectivity.

806

The Essence of Chromatography

The general use of cyclodextrin derivatives as stationary phases has dechned in recent years in favor of cyclodextrin derivatives either dissolved in, or chemically bonded to, a poly(siloxane) stationary phase. These phases generally possess higher thermal and mechanical stability, as well as higher efficiency. A wider range of cyclodextrin derivatives can be employed, since melting point and phase transition are no longer considerations for solutions. Table 10.4 [30,33-38]. The cyclodextrin derivatives are typically dissolved in moderately polar viscous poly(siloxane) stationary phases at a concentration of 5-50 % (w/w). The coating characteristics of the column resemble those of the bulk solvent, which can be varied to reduce retention and elution temperatures [75]. Both retention and enantioselectivity is influenced by the choice of poly(siloxane) solvent and the concentration of cyclodextrin derivative. It is generally observed that the enantioselectivity does not increase linearly with the concentration of the cyclodextrin derivative, and optimum values are often found in the low to intermediate concentration range (< 30% w/w for ^-cyclodextrin derivatives). In general terms, the ring size and substituents bound through the C-2, C-3 and C-6 hydroxyl groups significatly influence the physical properties and enantioselectivity of the cyclodextrin derivatives [76-81]. Alkyl and acetyl substituents at C-2 and C-3 contribute significantly to the enantioselectivity of the cyclodextrin derivatives. There is some evidence that the substituent at C-2 is more important when the chiral recognition mechanism involves interactions at the outer surface of the cyclodextrin and the substituent at C-3 when inclusion dominates the chiral recognition mechanism. These considerations are based largely on the relative orientation of the C-2 and C-3 substituents at the larger rim of the cyclodextrins. Substituents at the C-6 position play a significant role in determining the physical and chemical properties of the cyclodextrin derivative (melting point, solubility, etc.) They may also influence enantioselectivity by blocking the entrance at the smaller rim, and by modifying the conformation of the cyclodextrin. The a- and ^-cyclodextrins with 6-0-r-butyldimethlsilyl ether (or 6-0-thexyldimethylsilyl ether) groups with methyl or acetyl groups at C-2 and C-3 are among the most effective cyclodextrin derivatives for the separation of different enantiomer types [82-90]. Improvements in column technology have been achieved by incorporating the cyclodextrin into a poly(dimethylsiloxane) polymer that is suitable for thermal immobilization (section 2.3.4). Cyclodextrin derivatives with a monooctamethylene spacer arm attached to either the C-6 or C-2 hydroxyl group of a single glucose monomer can be incorporated into a poly(dimethylmethylhydrosiloxane) polymer by the hydrosilylation reaction. Figure 10.4 [91-96]. Some columns of this type are commercially available [e.g. heptakis(2,3,6-tri-0-methyl)-P-cyclodextrin (Chirasil-Dex), heptakis(30-trifluoroacetyl-2,6-di-0-n-pentyl)-P-cyclodextrinandoctakis(3-(9-butryl-2,6-di-0-npentyl)-y-cyclodextrin (Chirasil-y-Dex)], although columns containing dissolved selectors remain the most widely used. The advantages of immobilization are: an improvement in efficiency, inertness, and stability; compatibility with all injection techniques; the possibility of preparing columns with a higher concentration of the chiral selector; and the use of a non-polar poly(dimethylsiloxane) matrix (in which cyclodextrin deriva-

807

Separation of Stereoisomers

A

B

CH3

CH3

HgC

0-Si-OI CH3 , (CH2)8

I

(H3CO)6

ir

I

O

—o

)?-CD

1

r

(OCHa), (OCH3), Figure 10.4. Structures of immobilized poly(siloxane) chiral stationary phases containing (A) a cyclodextrin derivative (Chirasil-Dex) and (B) a metal complex (Chirasil-Nickel).

tives have poor solubility) resulting in lower elution temperatures for polar enantiomers. These columns are also suitable for applications using open tubular column supercritical fluid chromatography, liquid chromatography, and capillary electrochromatography [38,93-96]. The rationalization of chiral recognition involving cyclodextrin derivatives is difficult, since almost all classes of enantiomers are susceptible to separation on a certain clylodextrin derivative, often with no logical dependence on molecular shape, size, or functionality. The high efficiency of open tubular columns allows small selectivity factors to be exploited to yield a useful separation. This contributes just as much to the success of the cyclodextrin phases in gas chromatography as does their enantioselectivity, which can be poor, and yet an acceptable separation obtained. The ability of cyclodextrins to discriminate between enantiomers of unfunctionalized analytes, such as hydrocarbons, implies that the formation of inclusion complexes with the chiral cavity of the cyclodextrin derivatives is important. Variations in the separation properties with cavity size tend to support this general assumption. It is also clear that enantiomer separations occur under conditions where inclusion complexation is unlikely implying that formation of an inclusion complex is not a prerequisite for enantiomer discrimination. Cyclodextrin derivatives have been spectacularly successful in separating a wide range of low molecular mass alcohols, amines, amino acids, epoxides, carboxylic acids, esters, lactones, ethers, haloalkanes, and hydrocarbon enantiomers. Many of these compounds are difficult or impossible to separate by other means, and are economically important

808

The Essence of Chromatography

in evaluating the properties of essential oils, food additives, flavors, and chiral synthetic building blocks for more complex molecules [30,33,37,38]. 10.4.1.2 Liquid Chromatography Cyclodextrins and their derivatives can be covalently bonded to silica gel via a spacer arm using any of three general approaches [30,63,97-99]. The silica surface is first reacted with a reagent containing a spacer arm with a terminal functional group that can react with a free hydroxy 1 group of the cyclodextrin; the cyclodextrin is substituted with a spacer arm containing a terminal functional group that reacts with the silanol groups of the silica surface; or both the cyclodextrin and the silica support are functionalized with substituents that react to form the spacer arm. A wide range of chemistries is employed for these reactions, and in general, the approach used for the preparation of commercially available columns has not been disclosed. Due to the spatial requirements of the bulky cyclodextrins, surface coverage is typically relatively low ( ^ 80-120 |xmol/g). In general, unreacted silanol groups and surplus surface bound groups used for the immobilization of the cyclodextrins, influence the enantioselectivity of the column packing. Immobilized cyclodextrins and their derivatives are used in three general modes for liquid chromatography [28-30,32,63,66,99-102]. Cyclodextrins are used primarily in the reversed-phase or polar organic solvent mode. Acylated a-, p- and y-cyclodextrins and 2-hydroxypropyl, 1-naphthylethyl carbamate, 3,5-dimethylphenyl carbamate and 4-toluyl derivatives of p-cyclodextrin were introduced for normal-phase separations originally, but are now used for reversed-phase separations as well. Inclusion complex formation is believed to be significant only when the native or derivatized cyclodextrins are used in the reversed-phase mode. Only if the less polar (preferably aromatic) part of the analyte fits tightly into the cyclodextrin cavity and functional groups close to the stereogenic center interact with the chiral exterior (hydroxyl groups of the glucose units), can enantiomer separations be expected. The tighter the fit of the guest molecule the better the separation. Small molecules that fit loosely into the cavity are not resolved. In general, analytes with substituted phenyl, naphthyl, and biphenyl rings can be separated on p-cyclodextrin and analytes with 3 to 5 rings in their structure, on y-cyclodextrin. Retention and enantioselectivity depends on the pH, ionic strength, type of organic modifier and its concentration, and temperature. The pH and ionic strength of the mobile phase has most effect on enantiomers containing ionizable functional groups, is difficult to predict, and small changes can have a significant influence on enantioselectivity. Typical buffer concentrations are 10-200 mM of triethylamine phosphate, ammonium nitrate, ammonium acetate, or sodium citrate. Methanol and acetonitrile are the most commonly used organic modifiers, but ethanol and propan-1-ol are more efficient at displacing strongly retained analytes. Reaction of the cyclodextrin with single enantiomer reagents introduces new chiral selector sites into the cyclodextrin suitable for enantiomer separations in the normalphase mode. Inclusion complex formation is assumed to play a minor role (the least polar component of the mobile phase occupies the cavity thereby preventing inclusion

Separation of Stereoisomers

809

complex formation). Instead, enantioselectivity results from the stereoselective binding to the external functional groups of the (partially) derivatized cyclodextrin by hydrogen bonding, dipole-type and n-n interactions. Since inclusion complex formation is not important, cavity size is not a significant factor for stationary phase selection, while the choice of derivative is. Separations are usually performed with hexane containing various amounts of ethanol or propan-2-ol as the mobile phase. The normal-phase mode is suitable for the separation of enantiomers of low water solubility, or watersoluble enantiomers after derivatization. The separation mechanism is similar to the poly (saccharide) derivatives (section 10.4.2), which often provide better resolution in the normal-phase mode. In the polar organic solvent mode, the mobile phase is usually acetonitrile with small amounts of hydrogen-bonding modifiers, such as methanol, acetic acid, and triethylamine. Inclusion complex formation is suppressed by acetonitrile while at the same time promoting hydrogen bonding between hydroxyl groups on the cyclodextrin rim with suitable enantiomers. At least two functional groups must be present in the analyte for effective separation, and at least one of these functional groups must be on or near the stereogenic center for the analyte. The chiral recognition mechanism is thought to involve interactions at the rim of the cyclodextrin cavity, organizing the analytes as a cover over the entrance to the cavity. Hydrogen bonding and dipole-type interactions, together with steric factors, are all thought to be important for this interaction. Most of the enantiomers separated in the polar organic solvent mode contain amine groups, while a few have carboxylic acid or phenol groups. Separations are usually optimized by varying the amount of acetic acid and triethylamine, or the ratio of acetic acid to triethylamine, such that the combined acid/base concentration is in the range 0.002 to 2.5% (v/v), in a mobile phase containing 85-100% acetonitrile and 15-0% methanol. Addition of methanol reduces the retention time of strongly retained analytes. Crown ethers, such as 18-crown-6 and its derivatives and calixarenes also form hostguest complexes. Crown ether derivatives are suitable for the separation of enantiomers containing a primary or secondary amine group near a stereogenic center using reversedphase liquid [103-105] or gas [106] chromatography. Currently, a crown ether stationary phase containing binaphthyl atropisomers as chiral units with an 18-crown-6 backbone is commercially available for liquid chromatography (Crownpak CR). This phase binds ammonium ions stereoselectively by inclusion complex formation and multiple hydrogen bonding interactions between the ammonium ion and the oxygens of the crown ether. Enantioselectivity results from steric interactions at the binding site. The application range is limited by the specific nature of the chiral recognition mechanism. 10.4.2 Poly(saccharide) Derivatives The ester and carbamate derivatives of cellulose and amylose are among the most successful and versatile chiral stationary phases for liquid and supercritical fluid chromatography [1,4,28,107-109]. These phases are prepared by reaction of the poly(saccharide) with an acid chloride (ester derivative) or phenylisocyanate (carbamate

810

The Essence of Chromatography

Cellulose

o

R~

o

"0

~O"~ .ok

~~

R-{.o
R=

)-{

~

-O-C~

CH:!

Figure 10.5. General structures of the cellulose and amylose derivatives. Additional R groups are identified in Table 10.5.

derivative). The derivatized poly(saccharides) are then coated from solution onto a wide pore 3-aminopropylsiloxane-bonded silica support by solvent evaporation. The general structure of the cellulose and amylose derivatives is shown in Figure 10.5. Commercially available columns are summarized in Table 10.5. Because these stationary phases are physically coated, the solubility or swelling of the stationary phase restricts the range of solvents that can be used as a mobile phase. To improve the stability of the stationary phases, a number of methods have been described to immobilize the poly(saccharide) derivatives on silica-based supports, including coupling reactions with bifunctional reagents, radical crosslinking, or reaction of the terminal group of the poly(saccharide) with the amino terminal group of the support [108-110]. Immobilization was found to significantly alter the enantioselectivity of the stationary phase, often, but not always, leading to lower resolution. These phases have been little used in practice. Microcrystalline cellulose triacetate, which has a different physical form and enantioselectivity to the precipitated cellulose ester stationary phases, is widely used in medium-pressure liquid chromatography for preparativescale separations, as a relatively low cost and versatile chiral stationary phase. The helical structure of the microcrystalline cellulose phases contributes significantly to their enantioselectivity by providing a cavity for inclusion, which is absent in the precipitated cellulose phases. In the latter case, hydrogen bonding, rc-rr, and dipoletype interactions at the polymer surface dominate the chiral recognition mechanism.

811

Separation of Stereoisomers Table 10.5 Chiral stationary phases prepared from silica-based coated poly(saccharide) derivatives Poly(saccharide) (i) Esters Cellulose

Derivative

Designation*

triacetate tribenzoate tris (4-methylbenzoate) tricinnamate

Chiralcel Chiralcel Chiralcel Chiralcel

OA OB OJ OK

(ii) Carbamates Cellulose

Chiralcel OC tris(phenyl carbamate) Chiralcel OD tris(3,5-dimethylphenyl carbamate) tris(4-methylphenyl carbamate) Chiralcel OG Chiralcel OF tris(4-chlorophenyl carbamate) Chiralpak AD Amylose tris(3,5-dimethylphenyl carbamate) tris(l-phenylethyl carbamate) Chiralpak AS * Trademark of Daicel Chemical Industries. -H indicates the poly(saccharide) derivative is coated on a 5 ixm particle support (usual particle size 10 |xm) and -R that the poly(saccharide) derivative is intended for use in reversed-phase liquid chromatography.

A wide range of alcohols, carbonyl compounds (particularly if the carbonyl group is close to the stereogenic center), lactones, phosphorus compounds, aromatic compounds, and sulfoxides have been successfully resolved on the different poly(saccharide) derivatives. Non-polar mobile phases are commonly used, although not exclusively, and the selectivity of the various cellulose polymers changes with the character of the ester or carbamate groups used to derivatize the polymer. 10.4.2.1 Liquid Chromatography The popularity of the poly(saccharide) derivatives as chiral stationary phases is explained by the high success rate in resolving low molecular mass enantiomers. It has been estimated that more than 85% of all diversely structured enantiomers can be separated on poly(saccharide) chiral stationary phases, and of these, about 80% can be separated on just four stationary phases. These are cellulose tris(3,5-dimethylphenyl carbamate), cellulose tris(4-methylbenzoate), amylose tris(3,5-dimethylphenyl carbamate), and amylose tris(l-phenylethyl carbamate). Typically, n-hexane and propan-2-ol or ethanol mixtures are used as the mobile phase [111]. Both the type and concentration of aliphatic alcohols can affect enantioselectivity. Further mobile phase optimization is restricted to solvents compatible with the stationary phase, such as ethers and acetonitrile, as binary or ternary solvent mixtures, but generally not chloroform, dichloromethane, ethyl acetate, or tetrahydrofuran. Small volumes of acidic (e.g. trifluoroacetic acid) or basic (n-butylamine, diethylamine) additives may be added to the mobile phase to minimize band broadening and peak tailing [112]. These additives, however, may be difficult to remove from the column by solvent rinsing to restore it to its original condition. The derivatized cellulose and amylose stationary phases also shows high enantioselectivity in the reversed-phase mode [113-115]. The poly (saccharide) derivatives have

812

The Essence of Chromatography

no ionic sites, and to obtain adequate retention of ionized compounds, it is necessary to maintain all analytes in a neutral form using ion suppression or ion-pair formation. For basic analytes, pH selection is restricted by the solubility of the silica support (pH < 7-9), and ion-pair reagents, such as perchlorate or phosphorous hexafluoride, are commonly used. The enantioselectivity often depends on the concentration and identity of the ion-pair reagent. The chiral recognition mechanism of poly(saccharide) derivatives appears to depend on the conformation of the polymer chain and the structure of the substituents introduced by derivatization. Cellulose and amy lose are glucose polymers with a different connection between the monomer units (1,4-a or 1,4-p link) resulting in a helical structure with different enantioselectivity. A reversal of the elution order of enantiomers on cellulose and amylose derivatives is not unusual. The chiral recognition mechanism for carbamate derivatives is influenced by the substituents on the phenyl group, since the substituents modify the polarity of the carbamate group. The hydrogen bond donor and acceptor sites of the carbamate are arranged close to the core of the helical polymer axis of the glucose units forming helical grooves. These can direct the interaction and selective insertion of enantiomers with complementary binding sites, by directional hydrogen bonding and dipole-type interactions. In addition, the aromatic groups of the enantiomers may undergo TT-TT interactions with the phenyl groups of the polymer, which are located on the exterior of the groove. Polymerization of methacrylates and (meth)acrylamides, in the presence of a chiral anion catalyst results in the formation of a polymer with a helical structure, if the ester side chains contain a bulky group [116]. These polymers can be physically coated onto macroporous silica for liquid chromatography. Enantioselectivity in this case results from insertion and fitting of the analyte into the helical cavity, similar to the poly(saccharide) derivatives. Aromatic compounds, and molecules with a rigid nonplanar structure, are often well resolved on this phase. The triphenylmethyl methacrylate polymers are normally used with eluents containing methanol, or mixtures of hexane and 2-propanol. The polymers are soluble in several common solvents (e.g. chlorinated hydrocarbons, tetrahydrofuran, etc.) resulting in poor column stability. This together with the fact that there are few indications that separations performed on these columns cannot be replicated on other phases, has resulted in dechning use. 10.4.2.2 Supercritical Fluid Chromatography The lower mobile phase viscosity and higher solute diffusivity in supercritical fluids often provide improved resolution and shorter separation times for the separation of enantiomers when compared with liquid chromatography. Rapid column equilibration after changes in the separation parameters in supercritical fluid chromatography shortens method development time. The ease of mobile phase removal makes supercritical fluid chromatography attractive for preparative-scale separations. The derivatized poly(saccharide) phases are stable under supercritical fluid chromatography conditions, even though the polymers are coated and not bonded to the silica support [28,39-43]. A large number of enantiomers have been separated on cellulose and amylose tris(3,5-

Separation of Stereoisomers

813

Table 10.6 General properties of glycopeptide chiral stationary phases Property Molecular weight Stereogenic centers Macrocycles Sugar groups Aromatic rings Hydroxyl groups Amide groups Amine groups pH stability range

Ristocetin A 2066 38 4 6 7 21 6 2

Teicoplanin 1877 23 4 3 7 15 7 1 3.8-6.5

Vancomycin 1449 18 3 2 5 9 7 2 4-7

dimethylphenyl carbamates) at 30°C with carbon dioxide-methanol containing 0.1% (v/v) triethylamine and 0.1% (v/v) trifluoroacetic acid mobile phases, usually with composition programming [117-121]. The glycopeptide stationary phases (section 10.4.3) were only slightly less successful than the poly(saccharide) derivatives and demonstrated a similar application range [117,122,123]. Other phases used successfully include immobilized cyclodextrins and chemically bonded low-mass synthetic selectors (Pirkle phases). Chiral stationary phases based on proteins, ligand exchange, and crown ethers are either unstable or ineffective in supercritical fluid chromatography. Most enantiomer separations depend, at least in part, on multiple polar interactions between the enantiomers and chiral selector to form diastereomer complexes. These complexes are often too stable for the enantiomers to be eluted by carbon dioxide in the absence of polar modifiers. Optimum resolution is usually observed at low temperatures, frequently subcritical. These conditions typically result in increased peak separations accompanied by increased band broadening. The observed change in resolution depends on which factor is dominant. Mobile phase composition and temperature are the two most important parameters for optimizing resolution in the minimum separation time; pressure is often less important as the modified mobile phases are not very compressible. 10.4.3 Macrocyclic Glycopeptides (Antibiotics) The macrocyclic glycopeptides are a relatively new class of chiral selector with favorable chromatographic properties and broad applicability [124-129]. So far, eight glycopeptides (vancomycin, teicoplanin, thiostreption, rifamycin B, kanamycin, streptomycin, fradiomycin and ristocetin A) have been used for the separation of enantiomers by either liquid chromatography or capillary-electromigration separation techniques. Vancomycin, teicoplanin, and to a lesser extent ristocetin A, Table 10.6 and Figure 10.6, are generally used for liquid chromatography [130]. The macrocyclic glycopeptides can be bonded covalently to silica gel by multiple linkages involving the hydroxyl and amine groups of the glycopeptides, in such a way as to ensure their stability, while retaining their enantioselectivity [129]. All the glycopeptides contain an aglycone portion, made up of fused macrocyclic rings forming a characteristic "basket"

814

The Essence of Chromatography

H3C NH

NH

A ,

HO HO

^^~-NH. / ^ O

B Figure 10.6. Structures of the macrocyclic glycopeptide chiral selectors vancomycin (A) and teicoplanin (B).

shape, with carbohydrate groups attached to the aglycone basket. In addition, each glycopeptide has a large number of stereogenic centers and various functional groups, promoting multiple interactions with different enantiomers. The application range of the macrocyclic glycopeptides is similar to protein-based stationary phases (section 10.4.4), with the advantage that the glycopeptide columns are more robust and efficient, as well as possessing a higher loading capacity. The macrocyclic glycopeptides can be used in the normal-phase, reversed-phase, and polar organic solvent modes with few problems in changing from one mode to another. The enantioselectivity for each mode is generally different, increasing the versatility of the glycopeptide chiral selectors. If the compound has more than one functional group capable of interacting with the stationary phase, and at least one of those groups is on or near the stereogenic center, then the polar organic solvent mode should provide a good starting point. Methanol is the preferred solvent, with small amounts of acetic acid and triethylamine used to control retention and enantioselectivity. The total concentration of acid and base within the range 0.001 to 1 % (v/v) is used to control retention, and the ratio of acid to base, to control enantioselectivity. If the total concentration of acid and base required for elution exceed 1 % (v/v), then separation in the reversed-phase mode is indicated. If the total concentration of acid and base to achieve acceptable retention is less than 0.001 % (v/v), then separation in the normal-phase mode is indicated. Hexanealiphatic alcohols (e.g. ethanol) are commonly used for normal-phase separations, and mixtures of organic solvent and buffer for the reversed-phase mode. The most suitable buffers for reversed-phase separations are ammonium nitrate, triethylamineacetic acid, and sodium citrate. The pH range should be restricted to the stable operating range for the stationary phase. Table 10.6. Separations in the reversed-phase mode usually benefit from the presence of buffer (10-100 mM), even for neutral compounds. Enantioselectivity is usually highest in the reversed-phase mode if ionizable racemates

Separation of Stereoisomers

815

are separated in their neutral form. Tetrahydrofuran is the preferred organic solvent modifier for vancomycin, while for teicoplanin, methanol seems more versatile. Other factors affecting enantioselectivity include the buffer type, and ionic strength. The excellent enantioselectivity of the macrocyclic glycopeptides is based on distinctive shape parameters related to multiple stereogenic centers, and to multiple binding sites that are readily available close to the stereogenic centers. Enantiomers are probably bound to the pocket of the glycopeptide by partial inclusion, particularly in the reversed-phase mode. Anionic or cationic sites on the glycopeptide can interact strongly with oppositely charged enantiomers. Carboxylic acids enter into strong electrostatic interactions with an ammonium center located in the aglycone basket of teicoplanin [126]. Close to this site are located a hydrophobic cavity, as well as additional hydrogen bonding and dipolar groups associated with the aglycone peptide bonds, and the pendent carbohydrate groups. These features provide a facile method for the separation of amino acid and peptide enantiomers. Hydrogen-bonding, dipoletype, and TT-TC interactions with the aromatic groups of the peptide side chains are favored in the normal-phase mode, and are also likely to contribute to a lesser extent to enantioselectivity in the reversed-phase mode. The availability of a large number of enantioselective binding sites provides the basis for broad applicability, but complicates the rational understanding of the chiral recognition mechanism, and the ability to predict absolute configurations from elution order. Given the large number of macrocyclic glycopeptides available for study, further examples of this class of chiral selector are likely to become available in the future. 10.4.4 Proteins Proteins are composed of chiral monomers, organized in different sequences, providing a huge structural diversity. Specific folding (secondary and tertiary structures) and post-translational modification (e.g. glycosidation and sialinic acid modification) add to this diversity, creating chiral selectors with numerous binding sites of different stereoselectivity. Protein-based stationary phases are considered indispensable tools for analyzing stereoselective phenomena of biological significance (e.g. drug-protein binding studies), which are not discussed here. Their general use for analysis is in decline, however, since more rugged and efficient chiral stationary phases with a higher sample capacity are now available and suitable for the separation of enantiomers formally only possible on protein phases. The low sample capacity of protein phases can be a problem, even for analytical applications, and precludes their use for preparativescale separations. The complexity of protein structures results in a limited mechanistic understanding of the separation process. Typical proteins used for separating enantiomer include albumins (bovine and human serum), glycoproteins (ai-acid glycoprotein, ovoglycoprotein, avidin), and enzymes (cellobiohydrolase I, pepsin). Table 10.7 [28-32,131-134]. Proteins can be physically adsorbed on a suitable support for chromatography, but are usually prepared by covalently bonding the protein to a suitable support. Proteins can be bound to wide pore.

The Essence of Chromatography

816

Table 10.7 Protein-based stationary phases for liquid chromatography Protein (i) Albumins Bovine serum Human serum (ii) Glycoproteins ai-Acid glycoprotein Ovoglycoprotein Avidin (Hi) Enzymes Cellobiohydrolase I Pepsin

Molecular mass (kDa)

Carbohydrate content (%)

Isoelectric point

66 65

0 0

4.7 4.7

41 30 66

45 25 7

2.7 4.1 10.0

64 34.6

6 0

3.9 <1

chemically bonded silica gels (e.g. 3-aininopropylsiloxane-bonded silica) by coupling either the side-chain or terminal amino groups, or terminal carboxylic acid groups, to the chemically bonded support. The chiral recognition properties of adsorbed or bound proteins are often different to those of the protein in solution because of blocking of functional groups and/or conformational changes. To minimize denaturation, proteinbound phases are suitable for use with aqueous mobile phases in the pH range 3-8. The a 1-acid glycoprotein is built up of a single peptide chain containing 183 amino acids [132-134]. Five poly (saccharide) groups are linked to the peptide chain via the aspargine residues. The poly (saccharide) groups contain 14 sialic acid residues, giving the protein an acidic character with an isoelectric point of 2.7. At least two different stereogenic-binding sites have been identified for a i-acid glycoprotein. The main binding site is most likely a hydrophobic pocket, formed by an enrichment of hydrophobic amino acid residues, and containing numerous ionizable groups, both acidic and basic, allowing for the possibility of ionic binding of enantiomers. Separations are usually performed at pH 4-7, where the protein has a negative charge, and binds cationic enantiomers by electrostatic interactions. Neutral and acidic enantiomers are bound mainly by hydrophobic, hydrogen bonding, and dipole-type interactions. In addition, cationic enantiomers can be bound by adsorption of ionpairs with the buffer as a source of counterions. Enantioselectivity generally depends on pH, type and concentration of organic modifier, buffer type and its ionic strength, and temperature. For cationic enantiomers, mobile phase pH is often the most critical parameter with enantioselectivity generally increasing at higher pH and retention with an increase in buffer concentration. For hydrophobic compounds, an organic modifier, such as propan-2-ol or acetonitrile, is required to optimize retention and enantioselectivity. The adsorption of organic solvent by the protein induces reversible changes in its conformation leading to changes in enantioselectivity in addition to those induced by competition with the enantiomers for adsorption at the binding site. The a 1-acid glycoprotein and ovoglycoprotein stationary phases have the broadest application range of the protein-based stationary phases. The albumins may provide improved resolution of acidic and zwitterionic enantiomers. Cellobiohydrolase I is

Separation of Stereoisomers

817

sometimes preferred for the separation of basic compounds, but is usually less effective than a 1-acid glycoprotein and the albumin stationary phases, for the separation of neutral compounds. 10.4.5 Low-Mass Synthetic Selectors Low-mass synthetic selectors are represented by a large number of chiral stationary phases developed from concepts of a rational model of the chiral recognition mechanism. They usually contain a single or small number of chiral centers near substituent groups that enter into attractive or steric interactions with the enantiomers. Separations are often rationalized using the three-point rule based on hydrogen bonding, 71-Tt, and dipole-type interactions between complementary features of the chiral selector and enantiomers. The chiral selectors are relatively small and suitable for immobilization on silica gel or glass surfaces by standard chemical procedures. This results in efficient and durable phases with a comparatively high bonding density of the chiral selector. The chiral selectors are generally available in both single-enantiomer forms, facilitating reversal of the elution order for the quantification of the minor enantiomer in samples of high enantiomer purity. 10.4.5.1 Gas Chromatography The development of amino acid ester, dipeptide, and diamide chiral stationary phases provided the first indication that enantiomers could be separated by gas chromatography (or any chromatographic technique) on stationary phases designed to promote multiple attractive interactions with an enantiomer [35,38,135]. Real interest and progress in chiral separations resulted from the preparation of diamide phases, grafted onto a poly(siloxane) backbone, suitable for coating on open tubular columns [136-138]. Stationary phases of this type can be prepared by a number of different methods, for example, from poly(cyanopropylmethyldimethylsiloxanes) by hydrolysis of the cyano group and subsequent coupling of the carboxylic acid group with L-valine-f-butylamine or L-valine-1-phenylethylamide. Trifluoroethyl ester-functionalized poly(siloxanes) provide a suitable substrate for the introduction of a variety of selectors (e.g. L-valine^butylamide, L-1-naphthylethylamide, L-^leucine-^butylamide) by nucleophilic displacement. Methods have also been developed for immobilization of the poly(siloxane) stationary phases at relatively low temperatures to minimize racemization of the chiral selector. Separation of each chiral center by several dimethylsiloxane or methylphenylsiloxane monomers (about 7) seems essential for optimum resolution of enantiomers and good thermal stability [139]. The only commercially available example of this class of chiral selector is the poly(methylsiloxane) phase containing L-valine-f-butylamide (Chirasil-Val), which has been used to separate a wide range of racemic amino acid derivatives, amino alcohols, amines, hydroxy ketones, 2-hydroxy acids and their esters, 3-hydroxy acids, lactones, and sulfoxides [140,141]. It permitted for the first time, a single separation of all the enantiomers of the common protein amino acids after conversion to A^, O, ^-pentafluoropropanoate isopropyl ester derivatives. Figure 10.7

818

The Essence of Chromatography

CH.

^3

—Si—0-4-Si—0(CH2)3

\CH3

/n

C -0

I

NH CHCH (CHo)o CONHCICHjlj

-87«»C

120* U0« (cmpcraturr program

160* l,*Cim\T\

Figure 10.7. Separation of the enantiomers of the common protein amino acids by gas chromatography after derivatization on a 20 m x 0.25 mm I. D. open tubular column coated with Chirasil-Val. (From Ref. [142]; ©Elsevier).

[9,12,35,140-143]. This led to the development of the enantiomer labeling technique for quantitative amino acid analysis. The common protein amino acids are also fully separated on the cyclodextrin stationary phase (Chirasil-y-Dex) [143]. The L-valine-1phenylethylamide-containing stationary phase provides a facile method for the determination of the configuration of monosaccharides, as well as resolving enantiomers with neutral oxygen-containing functional groups [140]. Enantiomer separations by hydrogen bonding chiral stationary phases, such as Chirasil-Val, generally requires derivatization of the analyte in order to increase volatility and/or introduce suitable functional groups for additional hydrogen bonding association. 10.4.5.2 Liquid Chromatography Following the pioneering work of Pirkle, a large number of chiral stationary phases based on low-mass synthetic selectors covalently bonded to silica gel for liquid chromatography have been described [21,144-148]. These phases have undergone evolutionary improvements or replacement to the extent that the number of stationary

Separation of Stereoisomers

819

Table 10.8 Covalently bonded synthetic low-molecular mass chiral selector stationary phases for liquid chromatography Chiral selector (i) n-electron acceptor stationary phases Dimethyl-A/^-3,5-dinitrobenzoyl-a-amino-2,2-dimethyl-4-pentenylphosphonate 3,5-Dinitrobenzoyl-3-amino-3-phenyl-2-( 1,1 -dimethylethyl)propanoate N-(3,5-Dinitrobenzoyl)leucine amide A/^-(3,5-Dinitrobenzoyl)phenylglycine amide A^-(3,5-Dinitrobenzoyl)tyrosine butylamide A^-(3,5-Dinitrophenyl)valine urea

Column type

ChyRoSine A CHIREX 3010

(ii) 7i-electron donor stationary phases A/^-(l-Naphthyl)leucine ester {iV-l-[(l-Naphthyl)ethyl]amido}valine amide {A^-l-[(l-Naphthyl)ethyl]amido}indoline-2-carboxyhc acid amide

CHIREX 3014 CHIREX 3022

(Hi) 71-electron acceptor/n-electron donor stationary phases l-(3,5-Dinitrobenzamido)-l,2,3,4-tetrahydrophenantlirene Ar-(3,5-Dinitrobenzoyl)-1,2-diphenyl-1.2-diaminoethane 7V-(3,5-Dinitrobenzoyl)-l,2-diaminocyclohexane A^-(3,5-Dinitrobenzoyl)-( 1 -naphthyl)glycine amide

Whelk-O 1 ULMO DACH DNB CHIREX 3005

a-Burke 2 p-GEM 1

phases in current use is only a small fraction of those described. These surviving phases tend to have a broader application base than those that went before them, and form the basis of this discussion. Some examples of the chiral stationary phases belonging to this group are summarized in Table 10.8 and Figure 10.8. The Pirkle-concept phases are designed to operate using attractive hydrogen bonding, n-n interactions, and dipole type interactions between the chiral selector and the enantiomers. If need be, analytes lacking the complementary functional groups for attractive interactions with the chiral selector can be derivatized to enhance their interactions with the chiral selector. Compounds containing amine or carboxylic acid groups close to the chiral center are usually derivatized first. Amines are converted to amides or carbamates and carboxylic acids to amides, anilides, or esters. The most commonly used derivatizing reagents contain either a naphthyl moiety or a 3,5-dinitrobenzoyl or 3,5-dinitrophenyl moiety to ensure maximum TT-TC interactions between the analyte and the chiral selector. The chiral selectors belonging to the Pirkle-concept stationary phases posses either a strong electron-deficient aromatic group (jt-acid) and/or an electron-rich aromatic moiety (iT-base) for face-to-face and/or face-to-edge TT-TT interactions with complementary sites within the enantiomers [148]. In addition, these directing 7t-jt interactions must be favorably supported by strong and directional hydrogen bonding and/or dipole-type interactions. These require hydrogen donor-acceptor groups (e.g. amide, carbamate, urea, sulfonamide, hydroxyl, etc.) readily accessible and close to the stereogenic center of the chiral selector. Bulky groups or rigid networks close to the sterogenic center of the chiral selector may enhance enantioselectivity by steric interactions. These stationary phases are primarily used in the normal-phase mode

820

The Essence of Chromatography

O2N

Si(o—(SioJ D

CONH{CH2)3CH3

NO2

O2N,

E

O2N

Figure 10.8. Synthetic low-mass chiral stationary phases. (A) DNP-phenylglycine; (B) ChyRoSine-A; (C) P-GEM 1; (D) Whelk-0 I; and (E) ULMO.

to enhance attractive interactions, and to allow the strength of these interactions to be optimized using competitive organic solvents in the mobile phase [149]. In most cases, enantioselectivity in the reversed-phase mode is lower than in the normal-phase mode because of the dominant solvation properties of water with respect to enantiomer binding interactions with the chiral selector [114]. Many of the chiral stationary phases have been developed by systematically applying the principle of reciprocity to the enantiomer binding interactions. A number of diverse racemates are analyzed on a chiral stationary phase containing as chiral selector the immobilized target molecule for the enantipmers of which a new selector is desired. The racemate showing the highest enantioselectivity in this system is selected, and

Separation of Stereoisomers

821

a chiral stationary phase prepared from one of its enantiomers. If the enantioselectivity is not adversely affected by the method of immobilizing the selector, then it is expected that a new stationary phase with superior properties for the separation of the target compound (and often-related compounds) will be obtained. To make the screening process more efficient combinatorial libraries of potential selectors can be used for target identification [150]. The latest generation of Pirkle-concept phases contains both 7t-acceptor and 7t-donor substituent close to the stereogenic center, Table 10.8. These phases have broad applicability to many compound classes, and are able to separate most enantiomers separated on the single TC-acceptor or TT-donor phases [29,144,151]. In the Whelk-0 1 phase, Figure 10.8, the Tt-acceptor and K-donor aromatic systems are held perpendicular to each other and form a flexible cleft in which the analyte is held by simultaneous face-to-face and face-to-edge interactions. Additionally, hydrogen bonding to the amide group and steric interactions occur near the stereogenic center. In general, racemates with an aromatic group and a hydrogen bond acceptor group near a chiral center, axis, or plane can be separated into enantiomers. The synthetic low-mass chiral selectors, developed for liquid chromatography, are useful for supercritical fluid chromatography as well [28,39-43]. Only the WhelkO chiral selector has been intentionally modified for use in packed column supercritical fluid chromatography by incorporating the selector as a side chain in a poly(dimethylsiloxane) matrix immobilized on silica gel (PolyWhelk-O) [152]. The PolyWhelk-0 chiral stationary phase is more robust to high temperatures and mobile phase additives than its liquid chromatographic analog, and possesses similar enantioselectivity. Separations by supercritical fluid chromatography on synthetic low-mass chiral selectors generally follow those observed in normal-phase liquid chromatography, but are usually achieved in a shorter time, and with higher efficiency. In many cases, enantioselectivity can be enhanced using subcritical operating conditions. The scope of chiral separation methods is expanded by coupling columns of different enantioselectivity, facilitated by the low viscosity of supercritical fluids. An example is given in Figure 10.9, for the simultaneous separation of acidic and basic drugs on three coupled columns, each containing a different chiral stationary phase [122]. None of the individual coupled columns was able to separate all the enantiomers in the mixture.

10.5 CHIRAL MOBILE PHASE ADDITIVES The chiral selectors used as mobile phase additives are generally the same as those employed for the synthesis of chiral stationary phases. Mobile phase additives offer greater flexibility in identifying suitable chiral selectors for a particular application. It is generally faster to screen additives than columns for a separation, and more economical to include a wider range of potential selectors in the screening process. Conventional stationary phases of higher efficiency than chiral stationary phases are used for the separation, improving the prospects for favorable resolution. The

822

The Essence of Chromatography

5

J 10

Ul 15

n 20 Min

Figure 10.9. Separation of racemic drugs belonging to different structural types on series coupled Chiralpak AD, Chiralcel OD and Chirex 3022 columns with carbon dioxide-methanol (containing 0.5% triethylamine and 0.5% trifluoroacetic acid) as mobile phase using composition programming. Peaks: 1 = ibuprofen; 2 = fenoprofen; 3 = clenbuterol; 4 = propranolol; and 5 = lorazepam. (From Ref. [122]; ©Preston Publications).

main requirement for a mobile phase additive is that, the chiral selector should be soluble in suitable mobile phases, and be available as a single enantiomer or in high enantiomeric excess. The main limitation is that many chiral selectors interfere in the response of common chromatographic detectors, either causing sample detectability problems or poor baseline stability. This limits the use of some chiral selector and detector combinations, or necessitates the use of special techniques to separate the chiral selector and enantiomer before detection. This is easier to achieve in capillary electrophoresis, where chiral mobile phase additives represent the main approach for the separation of enantiomers, than for column liquid chromatography, where chiral mobile phase additives are used to a limited extent. The same problems exist for thin-layer chromatography, but in this case, the absence of a wide range of chiral stationary phases results in mobile phase additives being the commonly used approach for enantiomer separations. 10.5.1 Liquid Chromatography The most important applications of chiral mobile phase additives in liquid chromatography are chiral ion-pair chromatography (section 4.3.3) and inclusion complex formation with cyclodextrins and similar chiral selectors. Other chiral mobile phase additives have been used only occasionally, and with modest success [1,3,32,153]. Multifunctional ionpair reagents, such as 10-camphorsulfonic acid [154], derivatives of tartatic acid (e.g. din-butyltartrate) [155,156], peptides (e.g. N-carbobenzoxyglycine-L-proline) [157-161],

Separation of Stereoisomers

823

and cinchona alkaloids (e.g. quinine) [28] can form diastereomer ion pairs with racemic amines and carboxylic acids. For enantioselectivity, the counterion and enantiomer must enter into multipoint interactions where the stability of the ion-pair complex is different for each enantiomer. This requires the availability of additional hydrogen bonding, dipole-type, or steric interactions, close to the stereogenic center, to supplement the basic electrostatic interaction holding the ion-pair complex together. Suitable counterions, therefore, have an ionizable functional group close to their stereogenic center, additional binding sites in the vicinity of the stereogenic center, and a rigid structure to enhance stability differences in the ion-pair complexes. Ion-pair formation is favored in low-polarity solvents. Most applications of chiral ion-pair chromatography, therefore, employ organic solvents as the mobile phase with small amounts of an acid or base to promote sample ionization, and silica-based spacer bonded propanediol or porous graphitic carbon as stationary phases. The planar surface, high non-specific retention, and absence of functional groups, makes porous graphitic carbon (section 4.2.5) a particularly effective stationary phase for chiral ion-pair chromatography. The strong retention of the ion-pair reagent and/or their complexes allow more polar mobile phases to be used, including mobile phases containing water, as well as expanding the range of suitable chiral selectors that can be used as ion-pair reagents. It seems to facilitate separations by allowing greater flexibility in the choice of separation conditions and enhanced discrimination for the adsorption of diastereomer complexes. Cyclodextrins and their derivatives are generally used as chiral mobile phase additives in the reversed-phase mode to promote inclusion complex formation, and to provide adequate solubility for the cyclodextrins. Native cyclodextrins and cyclodextrins with ionized functional groups (e.g. sulfated cyclodextrins) are thought to discriminate between enantiomers by differences in the stability of their inclusion complexes formed in the mobile phase [162-164]. Interactions between the cyclodextrin or enantiomercyclodextrin complex and typical (achiral) siloxane-bonded stationary phases are believed to be minimal. Alkylated cyclodextrins (e.g. methyl, hydroxypropyl), on the other hand, are believed to be strongly adsorbed by the stationary phase, with enantioselectivity resulting from the difference in stability of the diastereomer complexes formed with the adsorbed cyclodextrin [165]. In which case, enantioselectivity will depend on the concentration of the cyclodextrin in the mobile phase, the choice of achiral stationary phase (since this controls the non-selective retention of the enantiomers and surface coverage by the adsorbed cyclodextrin), temperature, and the type and concentration of organic modifier. Mixtures of cyclodextrin derivatives, with one cyclodextrin primarily forming complexes in the mobile phase and the second forming a dynamic stationary phase, can result in enhanced enantioselectivity in some instances [165]. This is likely to be the case, if one enantiomer binds more strongly with the cyclodextrin inmiobilized on the stationary phase, and the second enantiomer with the cyclodextrin with minimal interactions with the stationary phase. Buffers, if used at all, are used to suppress ionization of the analyte, or to improve peak shapes resulting from undesirable interactions with stationary phase silanol groups. Beyond the general control of retention, the role of organic modifier in enantiomer separations is not well defined. It is hypoth-

824

The Essence of Chromatography

esized that the type of organic modifier can affect enantioselectivity by: (1) reducing the hydrophobic driving force for inclusion complex formation; (2) by destabilizing the analyte-cyclodextrin complex by enhancing dispersion interactions between the analyte and mobile phase; (3) by competing with the analyte for occupation of the cyclodextrin cavity; (4) by undergoing inclusion complex formation simultaneously with the analyte; and (5) by disturbing the stoichiometric equilibrium between water and the complex [166,167]. Chiral ion-pair reagents and cyclodextrin derivatives have also been used to separate enantiomers using packed column supercritical fluid chromatography [168,169]. 10.5.2 Thin-Layer Chromatography There are few chiral stationary phases available for the separation of enantiomers by thin-layer chromatography [58-60]. Layers prepared with microcrystalline cellulose and its acetate and benzoyl derivatives, are the most versatile and suitable for the separation of polar racemates with multiple hydrogen bonding and dipole interaction sites close to the stereogenic center [60,170-172]. They are used mainly in the reversed-phase mode with water-aliphatic alcohol mixtures as the mobile phase. The mobile phase composition affects enantioselectivity through competition with enantiomers for polar interactions at the binding sites, and by altering the conformation of the stationary phase due to solvent regulated swelling, which is believed to control inclusion complex formation. Several racemates have been separated on silica gel or silica-based spacer bonded propanediol layers impregnated with chiral selectors, such as D-galacturonic acid, N-carbobenzoxy-L-amino acids, 10-camphorsulfonic acid, etc., using low-polarity mobile phases [58,59]. Only octadecylsiloxane-bonded silica layers, impregnated with a copper (II) complex of N-(2-hydroxydodecyl)-4-hydroxyproline, and used in the reversed-phase mode for ligand-exchange chromatography, are used to any significant extent (section 10.6.2). Chiral mobile phase additives provide a more versatile and cost-effective approach for enantiomer separations in thin-layer chromatography. Typically, chemically bonded layers with cyclodextrin and its derivatives, bovine serum albumin, or macrocyclic glycopeptides are used as chiral additives in the reversed-phase mode [59,60,172-178]. For P- and y-cyclodextrins and their derivatives, a 0.1 to 0.5 M aqueous methanol or acetonitrile solution of the chiral selector is used as the mobile phase. Bovine serum albumin is generally used at concentrations of 1 -8 % (w/v) in an aqueous acetate buffer of pH 5 to 7 or in a 0.5 M acetic acid solution, in either case with from 3-40 % (v/v) propan-2-ol (or another aliphatic alcohol), added to control retention. Enantioselectivity usually increases with an increase in concentration of the chiral selector, and may be non existent at low concentrations of the chiral selector. The low efficiency and short migration distances typical of thin-layer chromatography limit useful separations to those with relatively large enantioselectivity factors. Absorption by the chiral selector can cause baseline instability and reduced sample detectability for quantitative measurements by scanning densitometry. The chiral sepa-

Separation of Stereoisomers

825

ration mechanisms employed in thin-layer chromatography are not specific to this technique, and any separation achieved by thin-layer chromatography, is also possible by column liquid chromatography. 10.5.3 Capillary Electrophoresis The general use of capillary-electromigration separation techniques for enantiomer separations has grown significantly in the last few years [51-56]. The low-mobile phase consumption facilitates the economic use of even expensive chiral mobile phase additives. This together with the high separation efficiency of capillary-electromigration separation techniques enables successful conditions to be found for the separation of most enantiomeric mixtures. Capillary electrophoresis remains the dominant technique for enantiomer separations, but the frequency of use of capillary electrochromatography is increasing. At equilibrium, the effective mobility of an enantiomer in the presence of a chiral selector in capillary electrophoresis is the sum of the electrophoretic mobility of each species containing the enantiomer, weighted by the mole fraction of each species. Assuming for a given separation the enantiomers exist as either the free enantiomer with a mobility, |XR, or the enantiomer-chiral selector complex, |XC,R, then the effective mobility for the enantiomer is given by M>eff,R = (^iR + KR[CS]^Jlc,R) / (1 + K R [ C S ] )

(10.1)

where KR is the formation constant for the enantiomer-chiral selector complex and [CS] the chiral selector concentration. The equation has been written for the R configuration of the enantiomer (an identical equation can be written for the S configuration). The general requirement for a separation of enantiomers by capillary electrophoresis is that |Xeff,R 7^ |Xeff,s- This requirement is met, if the complex formation constants or the mobility of the enantiomer-chiral selector complexes, are not identical (KR ^ Ks or |xc,R ^ |J^c,s)- When it can be reasonably assumed that differences in the enantiomer-chiral selector complex mobility are small (i.e. |XC,R ^ |xc,s = M^c) the enantioselectivity, A|jLeff,R,s, is expressed by |Xeff,R,s = {(M^o - [ic)( KR - Ks)[CS]} / {1 + [CS]( KR + Ks) + KRKS[CS]2}

(10.2)

where |xo is the mobility of the analyte in the absence of the chiral selector. To maximize the enantioselectivity, a large difference in mobility between the analyte and the analyte-chiral selector complex is desirable (i.e. the vector terms |xo and |xc should have opposite signs corresponding to migration of the free and complexed form of the analyte in opposite directions). The enantioselectivity is enhanced by a significant difference in the affinity of each enantiomer for the chiral selector (choose a chiral selector that maximizes the difference between KR and Ks). In addition, for any chosen chiral selector, there is an optimum concentration of the chiral selector for maximum

826

The Essence of Chromatography

enantioselectivity ([CS]opt = 1 / V K R K S ) , which is unique for each analyte. Thus, when screening several racemates it is unUkely there will be a single chiral selector concentration that is optimum for the separation of all enantiomers. The analyte as well as the chiral selector may be neutral, anionic, cationic, or zwitterionic. In addition, buffer additives (other than the chiral selector) and the nature of the capillary wall (coated or uncoated), which may both affect the electroosmotic flow, the charge of the analyte, and the chiral selector determine the enantioselectivity and migration time in capillary electrophoresis. Several models have been proposed to describe the influence of the chiral selector concentration, pH, electroosmotic flow, buffer type, and charge type for the chiral selector and enantiomer, multiple binding equilibria, and organic solvent modifier on enantioselectivity determined as a difference in mobility for the Rand S-enantiomers [49-52]. As well as these mobility difference models, Vigh and coworkers introduced the chiral charged resolving agent migration model (CHARM) to optimize separation conditions for ionized chiral selectors, where the enantioselectivity is expressed as the ratio of the effective mobility of the R- and S-enantiomers [178,179]. All these models are complex, but provide reasonable agreement with experimental data when the full range of experimental parameters is known. They are most useful for simulating the influence of experimental parameters on enantioselectivity and migration time. The above models provide an explanation for the observed reversal of the enantiomer migration order with the same chiral selector when different experimental conditions are employed [49,180]. The migration order is determined by the interaction of several parameters. Variation of these parameters is an effective means of changing the migration order so that a desired enantiomer (e.g. a minor component) is migrated before the opposite enantiomer of higher concentration. The enantiomer migration order is affected by: (1) the enantiomer-chiral selector formation constants; (2) the direction and magnitude of the free analyte mobility; (3) the direction and magnitude of the chiral selector mobility; (4) the direction and magnitude of the enantiomer-chiral selector complex mobility; (5) the direction and magnitude of the electroosmotic flow; and (6) the concentration of the chiral selector. These parameters, in turn, can be modified by subsets of experimental parameters providing a flexible, if some times confusing, number of options. Most mobile phase chiral additives for capillary electrophoresis are available only in a single-enantiomer form. In which case, change of the enantiomer migration order cannot be accomplished by a change of the configuration of the chiral selector. For general use in capillary electrophoresis, a mobile phase chiral additive should possess the following properties: (1) a high degree of enantioselectivity for a wide range of analyte structures; (2) solubility and stability in all common electrolyte solutions; (3) transparency in the low wavelength UV region; (4) favorable kinetics for the enantiomer-chiral selector binding process to minimize additional band broadening; (5) enantioselectivity independent of pH and other buffer parameters; and (6) availability in high purity at a reasonable cost. These properties are reasonably represented by the cyclodextrins and their neutral and ionic derivatives, which account for about two thirds

Separation of Stereoisomers

827

Table 10.9 Charged cyclodextrin chiral mobile phase additives for capillary electrophoresis Charged cyclodextrin a-,P-,y-Cyclodextrin phosphate Carboxymethyl P-,y-cyclodextrin Carboxyethyl p-cyclodextrin Succinyl p-cyclodextrin Sulfobutyl ether p-cyclodextrin Sulfated p-cyclodextrin Sulfated y-cyclodextrin Hepta-6-sulfato-P-cyclodextrin Heptakis(2,3-dimethyl-6-sulfato)-P-cyclodextrin Heptakis(2-0-methyl-3,6-di-0-sulfato) )-p-cyclodextrin Octakis(2,3-diacetyl-6-sulfato)-y-cyclodextrin Aminomethyl-^-cyclodextrin Heptakis(6-methoxyethylamine-6-deoxy)-P-cyclodextrin 2-Hydroxypropyltrimethylammonium- P -cyclodextrin Mono-(6-delta-glutamylamino-6-deoxy)-p-cyclodextrin Heptalds(2-A^,A/^-dimethylcarbamoyl)-P-cyclodextrin

Number of ionic groups 6 3.2, 3.5 6 3.5 3.5 7-11 7 7 14 8 1-2 7

of all published enantiomer separations by capillary electrophoresis [53-56,63,66,181184]. Other chiral mobile phase additives include chiral surfactants [55,185-187], proteins [188,189], macrocyclic glycopeptides [190], poly(saccharides) [191,192], and metal chelates (section 10.6.2). Nonaqueous capillary electrophoresis (section 8.2.6) has been explored to a limited extent for the separation of enantiomers with low water solubility or stability, and to modify the makeup of enantiomer-chiral selector interactions at the binding site compared with those in aqueous solution [193-195]. Cyclodextrins and their neutral derivatives, particularly ^-cyclodextrin and its alkyl and hydroxyalkyl derivatives, are widely used for the separation of enantiomers in capillary electrophoresis [22,51,53,55,66,181]. The low solubility of p-cyclodextrin, in particular, limits separation possibilities for enantiomers that interact weakly with the cyclodextrin, because the optimum cyclodextrin concentration may be beyond the solubility limit. The solubility of P-cyclodextrin in aqueous electrolytes is less than 20 mM, while typical optimum concentrations of cyclodextrins for the separation of enantiomers span the range 1-100 mM (usually < 50 mM). The solubility of the derivatized ^-cyclodextrins is about an order of magnitude higher than for the parent cyclodextrin, allowing the full concentration range for optimum enantioselectivity to be explored. Charged cyclodextrins. Table 10.9, are required for the separation of neutral enantiomers, and are also beneficial for ionized enantiomers, especially those of opposite charge to the cyclodextrin. For ionized enantiomers, this provides a means of maximizing the difference in electrophoretic mobility between the free and complexed forms of the enantiomer. General reactions used for the preparation of charged cyclodextrins produce randomly substituted products containing many different species that vary both in degree and in position of substitution. The range of

828

The Essence of Chromatography

formation constants for the different enantiomer-cyclodextrin complexes may enhance the enantioselectivity. On the other hand, different degrees of substitution may be responsible for peak tailing, because of the formation of several inclusion complexes, and electromigration dispersion phenomena (section 8.2.5.1). Furthermore, variation in the number of ionic substituents affects the optimum chiral selector concentration, as well as ionic strength of the electrolyte solution. To obtain more rugged separation conditions, a number of single isomer cyclodextrins, which are fully sulfated on the primary hydroxyl groups and uniformly substituted on the secondary hydroxyl groups, have been developed. Table 10.9 [55,183,195-198]. Cyclodextrins with strong electrolyte functional groups are generally preferred, because then the pH of the electrolyte solution can be adjusted freely according to the needs of the analyte, without effecting the charge of the cyclodextrin. To increase enantiomer mobility differences, several parameters, such as type, number of substituents and cyclodextrin concentration, pH, ionic strength, composition of the electrolyte solution, and temperature must be carefully optimized [29,63,199,200]. Basic compounds are usually separated at acid pH, where enantiomers are positively charged, and the electroosmotic flow is minimized. High pH is more appropriate for the separation of acids. There are many examples, where using a cyclodextrin together with another mobile phase additive enhances the separation of enantiomers [51,53,56,201-203]. The second additive need not be chiral (e.g. sodium dodecyl sulfate), or a second chiral selector, such as a chiral surfactant or another cyclodextrin, is used. In most cases a charged cyclodextrin is used in conjunction with a neutral cyclodextrin. Achiral selectors can be used with neutral cyclodextrins to create electrophoretic mobility differences, where neutral cyclodextrins are used with neutral analytes, or to improve enantioselectivity when either the analyte or the cyclodextrin carries a charge. Regardless of the enantioselectivity of complex formation, a neutral chiral selector cannot separate neutral enantiomers in the absence of a mobility difference between the free and complexed form of the analyte. The use of a second charged selector means that the important competition process is not that between the free and complexed forms of the analyte, but between different complexed forms. For neutral analytes the charged cyclodextrin shows no enantioselectivity itself, in some cases, but provides a mechanism whereby the enantioselectivity of the neutral cyclodextrin can be expressed. In other cases, with charged analytes, both cyclodextrins can contribute to enantioselectivity. Neutral cyclodextrins are also used in micellar electrokinetic chromatography with achiral surfactants to modify their enantioselectivity, particularly for the separation of hydrophobic analytes [53,55,185-187]. Enantioselectivity in this case results from differences in the distribution of enantiomers between the micellar pseudostationary phase and the cyclodextrin, as well as from the different migration velocities of the cyclodextrin and micelles. Neutral enantiomers can be separated based on differences in their equilibrium constants between the electrolyte solution and a charged chiral surfactant micellar phase, if the micelle has a different electrophoretic mobility to the free enantiomers. Suitable chiral surfactants include the bile salts (section 8.3.3), long alkyl-chain amino acid derivatives (e.g. sodium N-dodecanoyl-

829

Separation of Stereoisomers

DEXTRAN SULFATE

CHaOSOa*

NHS03*

OS03"

NMS03'

CHONDROITIN SULFATE A CH2OH >-—-O,

"V^l

•O3SO

NHCOCH3

Figure 10.10. Poly(saccharide) chiral mobile phase additives for capillary electrophoresis. Heparin (molecular mass 7,000-20,000 with 2-3 sulfate groups per disaccharide), Chondroitin sulfate A (molecular mass 30,000-50,000 with 0.2-0.3 sulfate groups per disaccharide) and Dextran sulfate (molecular mass 7,300 with 6 sulfate groups per disaccharide).

L-valinate, N-dodecoxycarbonylvaline), polymer surfactants (e.g. poly [sodium Nundecenyl-L-valinate]), and n-alkyl-P-glucopyranosides. An important property of the glycosidic surfactants is that they can be charged in situ through complexation with borate anions, which allows the surfactant charge density to be adjusted to optimize enantioselectivity [204]. Many ionic poly(saccharides), such as heparin, chondroitin sulfates, dextran sulfate, and natural poly(saccharides), such as dextran, dextrin, pullulan, and their charged derivatives have been used as mobile phase additives for the separation of different enantiomers. Figure 10.10 [191,192,205,206]. Dextrins were found to have a wide application range, thought to be due in part to their helical structures. Enantiomerchiral selector complexes seem to be weaker than for cyclodextrins, and it has not been demonstrated that enantiomer separations obtained by the poly(saccharide) chiral selectors cannot be obtained using cyclodextrins. Natural poly (saccharides) are typically complex mixtures of homologues and isomers, with a composition that can vary for different sources, resulting in differences in enantioselectivity. The same macrocyclic glycopeptides used as immobilized chiral selectors for liquid chromatography (section 10.4.3) are also suitable chiral mobile phase additives for capillary electrophoresis [129]. All the macrocyclic glycopeptides are unstable in aqueous organic solvent mixtures, particularly at pH <4 or pH >9. This does not

830

The Essence of Chromatography

generally prevent their use for capillary electrophoresis, but should be taken into account when selecting operating and storage conditions. It is believed that the amine groups of the glycopeptide are the primary sites for interaction. Higher enantioselectivity is generally exhibited for acidic compounds, or compounds containing an anionic group (e.g. carboxylic acid, phosphate, or sulfate group). There seems to be an enhancement in enantioselectivity when the acid group is either a- or P- to the stereogenic center. This is especially so for enantiomers which also contain a carbonyl group, an aromatic ring, or an amide nitrogen at the a-, P-, or y-position to the stereogenic center. The choice of macrocyclic glycopeptide and its concentration, pH of the electrolyte solution, and concentration of organic modifier, are the primary parameters used to control enantioselectivity. The preferred organic modifier is propan-2-ol. In addition, proteins, such as serum albumins and ai-acid glycoprotein (section 10.4.4), are used as chiral mobile phase additives in capillary electrophoresis [188,189]. Depending on the pH, the proteins are either negatively or positively charged, and suitable for the separation of neutral or ionized enantiomers. A general feature of the use of macrocyclic glycopeptides and proteins as chiral mobile phase additives, is problems associated with adsorption of the chiral selector on the capillary wall (loss of resolution), and high background UV absorption (diminished sample detectability and poor baseline stability). A solution to these problems is provided by the partial-filling technique [53,207-210]. The buffer containing the chiral selector is used to fill the separation capillary up to a position below the detection region. The sample is introduced, and the capillary inserted into the electrolyte solution without the chiral additive, for the separation. The separation conditions are chosen such that the chiral selector does not migrate past the detector. In this way the enantiomers migrate through the separation zone and are then detected in a zone, which does not contain the chiral selector. Changing either the chiral selector concentration or the length of the separation zone is used to optimize separations. 10.6 COMPLEXATION CHROMATOGRAPHY The rapid and reversible formation of charge-transfer complexes between metal ions and organic compounds that act as electron donors can be used to adjust selectivity in chromatography and electrophoresis [211]. Such coordinate interactions are influenced by subtle differences in the composition or stereochemistry of the donor ligand, owing to the sensitivity of the chemical bond towards electronic, steric, and strain effects. A number of challenging separations of stereoisomers and isotopomers has been achieved by complexation chromatography that would be difficult or impossible by other chromatographic methods. 10.6.1 Silver Ion Chromatography Silver ion (or argentation) chromatography is based on the characteristic property of unsaturated organic compounds to form transient charge-transfer complexes with

Separation of Stereoisomers

831

Table 10.10 General considerations for predicting the elution order of unsaturated compounds in silver ion chromatography • Unsaturated aliphatic and alicyclic compounds form more stable complexes than do aromatic compounds. • Methylene-interrupted dienes bind more strongly to silver ions than conjugated dienes, and monoenes bind more strongly than monoynes. • For methylene-interrupted dienes, the stability of complexes increases with the number of double bonds. • ci^-Double bond isomers usually form more stable complexes than trans-douhlo bond isomers. • Stability of complexes usually decreases with increasing chain length of the aliphatic substrate. • Stability of complexes usually decreases with increasing number of substituents attached to the double bond, and increases when deuterium or tritium replaces hydrogen.

transition metals in general, and with silver ions in particular [211-213]. Suitable electron donors are organic compounds containing 7t-electrons in various kinds of double and triple bonds, or containing heteroatoms (e.g. N, O or S) with electron lone pairs. For olefins, the silver ion is believed to act as an electron acceptor forming a a-type bond between the occupied 2p orbitals of the olefinic double bond and empty 5 s and 5p orbitals of the silver ion, and a 7t-acceptor backbond between the occupied 4d orbitals and the free antibonding 2p 7t* orbitals of the olefinic double bond [213,214]. The stability of these complexes is sensitive to small changes in the bond electron density and steric factors affording separations of olefinic compounds according to the number, geometry, and position of the double bonds. Interactions of silver ions with the electron lone pairs of oxygen of the carbonyl group were implicated in the separation of triacylglycerols [214]. In general, conjugated dienes form weaker complexes with silver than isolated double bonds, and double bonds with substituents differing in size and position can frequently be separated based on differences in their complex stability constants. Table 10.10. The stability of the complexes increases at lower temperatures, and the low maximum temperature for useful separations effectively limits gas chromatography to the separation of volatile olefins [211]. Silver ion chromatography is used primarily in liquid [213,215-218], thin-layer [212,213,216,219], and supercritical fluid chromatography [220-222] for the separation of fatty acid derivatives and triacylglycerols, and to a lesser extent, terpenes, sterols, carotenoids, and pheromones. Silver ion chromatography is widely used on its own, or as a preliminary simplification step, to elucidate the structures of fats, oils and lipids [213]. It is possible to fractionate animal or fish oils into fractions with zero to six double bonds. Species with the same total number of double bonds can be separated based on the difference in the number of double bonds in individual acyl residues. Silver-loaded layers for thin-layer chromatography are usually prepared by slurrying silica gel with an aqueous solution of silver nitrate (1-2 % w/v), or by immersing precoated silica gel layers in a 0.5 % (w/v) solution of methanolic silver nitrate

832

The Essence of Chromatography

for five minutes [212,213,219]. Columns for liquid chromatography are prepared by loading a solution of silver nitrate onto a prepacked silica gel column, or a silicabased, benzenesulfonic acid, cation-exchange column [212,213]. The silver-loaded, cation-exchange column is the preferred approach, and results in more stable retention and minimal contamination of collected fractions with silver salts. Separations are usually performed in the normal-phase mode with chlorinated hydrocarbon solvents containing small amounts of acetonitrile or ethyl acetate for the separation of fatty acids. Hexane containing small amounts of acetonitrile, ethyl acetate, acetone, or acetic acid is used for the separation of derivatized fatty acids and triacylglycerols [213,215,217,218]. Acetonitrile is a particularly effective solvent for eluting compounds containing several double bonds from silver-loaded, cation-exchange columns. Gradient elution is also commonly used for the fraction of complex mixtures. Reversed-phase liquid chromatography is usually the method of choice for analyzing molecular species from complex lipids (e.g. phosphatidylcholines) [213]. In this case, silver nitrate is used as a mobile phase additive. This results in a decrease in retention due to an increase in the hydrophilic character of the complex compared to the parent analyte. Varying the concentration of silver nitrate in the mobile phase enables the retention of the complexed species to be changed over a wide range. Since silver nitrate solutions are corrosive and stain clothing, this approach is not as popular as those based on silver-loaded, cation-exchange columns. So far, applications of silver-loaded packings in supercritical fluid chromatography have been limited to the analysis of lipids using silica-based, silver-loaded, cation-exchange sorbents, silvercomplexed, dicyanobiphenyl-substitutedpoly(dimethylsiloxane) encapsulated sorbents, or silver-complexed 8-quinolinol bonded sorbents [42,220,221]. All phases provide a good separation of fatty acid methyl esters according to their degree of unsaturation, but the silver-loaded, cation-exchange sorbents require the use of organic solvent modifiers, restricting detector options to the UV absorption or evaporative lightscattering detectors. In general, separations by supercritical fluid chromatography show small selectivity differences compared with similar separations by normal-phase liquid chromatography. Both techniques are valuable for the separation of triacylglycerols. 10.6.2 Ligand-Exchange Chromatography Ligand-exchange chromatography is used to resolve enantiomers of amino acids and their derivatives, and some a-hydroxycarboxylic acids, and compounds with an imide structure capable of forming bidentate complexes with transition metal ions, such as copper, zinc, and nickel [17,223-225]. Almost all practical examples employ copper (II) amino acid complexes as the chiral selector in liquid or thin-layer chromatography. Based on its high enantioselectivity, ligand-exchange chromatography was at one time the principal method for separating amino acid enantiomers. Relatively poor column efficiency and effective competition from other methods (e.g. cyclodextrin and macrocyclic glycopeptide stationary phases), however, has resulted in a marked decline in its use in recent years. A limited amount of work indicates that ligand-exchange

Separation of Stereoisomers

833

chromatography is suitable for the separation of amino acids and their derivatives by capillary electrophoresis [226]. Ligand-exchange chromatography is also the basis for the separation of oligosaccharides on metal-loaded, cation-exchangers (section 4.3.7). There are three general approaches for the separation of enantiomers by ligandexchange chromatography. The chiral selector, a Cu (II) amino acid complex, can be bonded to a silica support by a spacer arm connected to the amino acid at a position that does not interfere in the chelation mechanism [223,225,227]. The Cu (II) amino acid complex can be physically adsorbed on an octadecylsiloxane-bonded silica [225,228] or porous graphitic carbon [229,230] sorbent by attaching a hydrophobic group to a position on the amino acid that does not interfere in the chelation mechanism. Or, the copper (II) amino acid complex can be used as a mobile phase additive [223,225]. The highest enantioselectivity for amino acids and their derivatives is usually obtained using a single enantiomer of proline, 4-hydroxyproline, or histidine methyl ester as the chelating ligand. Separations are performed in the reversed-phase mode with the enantioselectivity optimized by varying the copper (II) concentration, pH, and the type and concentration of organic modifier. In chiral ligand-exchange chromatography, there is no direct contact between the chiral selector and the analyte. Their interaction is mediated by a metal ion, which simultaneously binds the chiral selector and the enantiomers to be separated. Equilibration of the complex with an entiomeric mixture capable of complexing with the metal, results in one of the previously chelated amino acids being displaced, forming a mixture of diastereomeric complexes. These complexes are separated based on the difference of stability constants for metal complexes forming part of the stationary phase, or adsorbed onto the stationary phase. Formation of diastereomeric complexes in the mobile phase results in a separation based on differences in the solubility of the complexes in the mobile phase and their adsorption by the stationary phase. Slow ligand-exchange processes are the primary cause of poor chromatographic efficiency. 10.6.3 Enantioselective Metal Complexation Gas Chromatography Metal (II) bis[3-(trifluoroacetyl)-(lR)-camphorate] and bis[3-(heptafluorobutanoyl)(IR)-camphorate] of nickel, cobalt, and manganese dissolved in a noncordinating solvent, such as poly(dimethylsiloxane), or incorporated into the structure of a poly(dimethylsiloxane). Figure 10.4, have emerged as highly selective stationary phases for the separation of a variety of stereoisomers that includes hydrocarbons and oxygen, nitrogen, and sulfur-containing electron-donor analytes (e.g. cyclic ethers, aziridines, thiranes, ketones, and aliphatic alcohols) by gas chromatography [16,35,38]. These complexes are also suitable for the determination of the enantiomer composition of volatile insect pheromones. Although the camphorates are the most widely evaluated and the only metal terpeneketonate complexes available as immobilized stationary phases, a number of other terpeneketonates have been evaluated as complexing ligands, as well as some further metals [16]. Selectivity results from the difference in stability constants for the fast and reversible chemical equilibrium between the

834

The Essence of Chromatography

metal coordination compound and analytes. Thermal instability of the coordination complexes, and small stability constants at higher temperatures, restrict separations to volatile compounds that can be separated at temperatures less than about 125°C. Many earlier applications of metal coordination compounds, can now be achieved using cyclodextrin derivatives (section 10.4.1.1). Supercritical fluid chromatography employing immobilized metal coordination compounds is suitable for the separation of thermally or configurationally labile analytes of lower volatility than required for gas chromatography [ 16,231 ]. 10.7 SEPARATION OF ENANTIOMERS AS COVALENT DIASTEREOMER DERIVATIVES Enantiomers containing a reactive functional group can be derivatized with a singleenantiomer reagent to produce a mixture of diastereomers [1,2,28,29,232-2361. By itself, this does not guarantee that a separation will be obtained, but it does make possible a separation in the absence of a chiral selector, using conventional separation systems. A useful separation depends on the difference in physical properties of the diastereomers, the extent to which these differences influence the relative distribution of the diastereomers between phases in the chromatographic system, and the separation efficiency of the chromatographic system. In contemporary practice, methods based on the formation of covalent diastereomer derivatives have continued to decline in popularity, as a wider range of options for direct separation by chiral chromatography and electrophoresis became available. However, formation of covalent diastereomer derivatives remains a viable option for many compounds, and sometimes it is the only option available. Several factors need to be considered in the selection of reagents and reaction conditions for the formation of diastereomer derivatives. The analyte must contain at least one functional group capable of reaction with the derivatizing reagent. A wide range of reagents is available for reaction with amines, hydroxyl (alcohol and phenol groups), and carboxylic acids, with a smaller number for aldehydes, epoxides and thiols, Table 10.11. The most widely used reactions employ relatively mild conditions, and short reaction times, to minimize racemization of the chiral compounds. The diastereomer transition states should have similar conversion rates for the enantiomers (absence of kinetic resolution) and similar stability in the reaction medium. The derivatizing reagent must be available as a single enantiomer in high enantiomer purity, and have an acceptable shelf life. To detect the presence of 0.5% of a minor enantiomer in a mixture of enantiomers requires a reagent of at least 99.9% enantiomer purity. It is advantageous if the derivatizing reagent is available in both the R- and S-form, to allow elution order reversal, to facilitate the detection of minor enantiomer components. Derivatization with suitable reagents also provides a method to improve detection properties of the enantiomers. This applies particularly to UV absorption properties, or the introduction of a fluorophore for fluorescence detection. It should be noted, however, that the UV

Separation of Stereoisomers

835

Table 10.11 Reagents for the formation of covalent diastereomer derivatives (i) Primary and secondary amines N-Trifluoroacetylprolyl chloride 1 -(4-Nitrophenylsulfonyl)prolyl chloride a-Methoxy-a-(trifluoromethyl)phenylacetyl chloride 2-Methyl-2P-naphthyl-l,3-benzodioxole-4-carboxylic acid chloride Menthyl chloroformate l-(9-Fluorenyl)methyl chloroformate (FMOC) 2-(6-Methoxy-2-naphthyl)-l-propyl chloroformate (NAP-C) 1 -Phenylethylisocyanate l-(l-Naphthylethyl) isocyanate 2-(6-Methoxy-2-naphthyl)ethyl isothiocyanate (NAP-IT) 1,3-Diacetoxy-1 -(4-nitrophenyl)-2-propyl isothiocyanate (DANI) (2,3,4,6-tetra-0-acetyl)-p-glucopyrasonyl isothiocyanate (GITC) 4-(3-isothiocyanatopyrrolidin-1 -yl)-7-(N,N-dimethylaminosulfonyl)-2,1,3-benzoxadiazole (DBD-PyNCS) N-[(2-isothiocyanato)cyclohexyl)pivalinoyl amide (PDITC) l-Fluoro-2,4-dinitrophenyl-5-alaninamide (Marfey's reagent) (ii) Alcohols and Phenols rraw5-Chrysanthemoyl acid a-Methoxy-a-(trifluoromethyl)phenylacetic acid (Mosher's reagent) a-Methoxy-a-(trifluoromethyl)propionic acid (MTPA) 2-rer^Butyl-2-methyl-1,3-benzodioxolo-4-carboxylic acid 2-Phenylpropionyl chloride 1 -Phenyethylisocyanate l-(l-Naphthylethyl) isocyanate Menthyl chloroformate l-(9-fluorenyl)ethyl chloroformate (FLEC) (Hi) Carboxylic acids 2-Butanol 3-Methyl-2-butanol Menthyl alcohol 2-Octanol 1-Phenylethanol a-Methyl-4-nitrobenzylamine 1 -(1 - Anthryl)ethylamine l-(4-Dansylaminophenyl)ethylamine 2-[4-(l-Aminoethyl)-phenyl]-6-methoxybenzoxazole (iv) Thiols N-[(2-isothiocyanato)-cyclohexyl)-pivalinoyl amide (PDITC) 4-(3-isothiocyanatopyrrolidin-1 -yl)-7-(N,N-dimethylaminosulfonyl)-2,1,3-benzoxadiazole (DBD-PyNCS)

and fluorescence properties of each diastereomer derivative are not necessarily identical, and should be checked by calibration. The separation of diastereomers is often enhanced when the stereogenic centers of the enantiomer and the reagent are in close proximity in the derivative, and when the reagent is comprised of conformationally immobile groups, or contains bulky groups, attached directly to the stereogenic center. The most likely sources of error in establishing the enantiomer composition of a mixture are the presence of enantiomeric impurities in the derivatizing reagent.

836

The Essence of Chromatography

racemization during the derivatization reaction, and different rates of reaction for individual enantiomers. In general, methods based on the formation of covalent diastereomer derivatives require more effort for validation than direct methods, owing to the larger number of factors that affect the results. If the reaction is to be used for preparative purposes, easy conversion of the diastereomers into the parent enantiomers is also a necessary condition. Currently, several hundred reagents are available for the preparation of diastereomer derivatives, and this list continues to grow. In consequence, any type of comprehensive coverage is impossible here. As well as the nature of the reactive group, cost, reactivity, stability, and availability are important selection criteria, since in advance of trial experiments, success cannot be guaranteed for analytes not studied previously. For trace analysis, the choice of a more selective and sensitive detector for the analysis may minimize the number of options to reagents with suitable characteristic detector properties. The nature of the reactive group defines the possible application range for each reagent. For amines activated carboxylic acids (e.g acid chlorides), chloroformates (forming carbamate derivatives), isocyanates (forming urea derivatives), isothiocyanates (forming thiourea derivatives) are popular choices [234,237-247]. Marfey's reagent (Table 10.11) is widely used to derivatize peptides and for other apphcations. The nucleophilic attack of the amine group on the C-F bond activated by the two nitro groups on the aromatic ring results in a smooth reaction to form aniline derivatives with good UV detectability [248,249]. The naproxen reagents (NAP-C and NAPIT) have strong chromophores, but weak fluorescence. A number of reagents indicated in Table 10.11 and suitable for fluorescence detection. The reaction of amino acids with o-phthalaldehyde and a single enantiomer thiol (e.g. N-butyrylcysteine) result in the formation of highly fluorescent isoindole derivatives [234]. a-Methoxy-a(trifluoromethyl)phenacetyl chloride and a-methoxy-a-(trifluoromethyl)propionic acid form stable derivatives with a wide range of amines for gas and liquid chromatography [234,237]. For alcohols, activated carboxylic acids or acyl nitriles (forming ester derivatives), chloroformates (forming carbonate derivatives) and isocyanates (forming carbamate derivatives) are widely used [234,250,251]. Dicyclohexylcarbodiimide can be used as the coupling agent for carboxylic acid reagents, but in situ transformation of the acid to the acid chloride is more widespread. Acid chlorides, anhydrides, and acyl cyanides usually require strictly anhydrous conditions and a catalytic amount of an organic base (e.g. N,N-dimethylaminopyridine] for quantitative reaction. Because of the lower reactivity of hydroxyl groups with isocyanates and chloroformates, a catalyst (organic base) is usually also required. The most frequently used approaches for derivatizing carboxylic acids are esterification with a variety of single-enantiomer alcohols, or formation of amides with single-enantiomer amines [234,252]. The formation of amide derivatives requires activation of the carboxylic acid by formation of the acid chloride with thionyl chloride, mixed anhydrides with chloroformates, N-acylimidazoles with 1,1-carbonyldiimidazole or N-acylureas with dicyclohexylcarbodiimide. Esterification reactions generally re-

Separation of Stereoisomers

837

quire harsh conditions, and this should be considered if either the enantiomer or the diastereomer derivative is conformationally labile or unstable. For liquid chromatography, both the reversed-phase and normal-phase mode is used to separate diastereomer derivatives. Normal-phase chromatography is particularly effective for separating diastereomers of moderate polarity in the absence of strong hydrogen bond acid/base functional groups. The requirements for supercritical fluid chromatography are similar to normal-phase liquid chromatography. For gas chromatography, the volatility and thermal stability of the derivatives is an important factor in selecting suitable reagents. The formation of diastereomer derivatives for enantiomer separations by capillary electrophoresis is not established as well as it is for chromatography [253-257]. Derivative formation in capillary electrophoresis has frequently been performed to improve sample detectability, especially in combination with laser-induced fluorescence detection. The use of chiral mobile phase additives is the favored approach for the separation of enantiomers in capillary electrophoresis (section 10.5.3). 10.8 LIQUID-CRYSTALLINE STATIONARY PHASES Liquid crystalline stationary phases are used for the separation of positional and geometric isomers of rigid molecules, such benzene derivatives, terpenes, polycyclic aromatic compounds, poly chlorinated biphenyls, and steroids [258-261]. Nearly all practical applications result from their use in gas chromatography, while rather limited success has been demonstrated for liquid and supercritical fluid chromatography. Liquid crystals exhibit the mechanical properties of a liquid while retaining some of the anisotropic properties of the solid state. This preservation of order, permits shape selectivity, while the liquid properties result in acceptable chromatographic efficiency. Several hundred liquid-crystalline phases of different chemical types have been used in gas chromatography. They all have in common a markedly elongated, rigid, rod-like structure, and generally have polar terminal groups. The most common types are Schiff bases, esters, azo, and azoxy compounds [258-263]. At their melting point, these compounds are transformed from a solid crystalline state to ordered smectic or nematic liquid states. The smectic configuration is more ordered than the nematic configuration, and occurs at a lower temperature for compounds that exhibit both phase transitions. At the clearing temperature, the liquid-crystalline state is transformed into an isotropic liquid. The temperature range bordered by the solid melting point and the clearing temperature defines the liquid-crystalline phase region. Supercooling coated liquid-crystalline phases allows some phases to be used below their solid melting point, if adequate column efficiency is maintained. The phase loading and support type can also alter the separation characteristics of liquid-crystalline phases. The thin film of liquid phase in immediate contact with the support has properties influenced by interactions with the support surface. As the film thickness builds up, these surface forces dissipate quickly. Most monomeric liquid-crystalline phases provide limited opportunities for use in open tubular columns because of poor column efficiency, poor coating charac-

838

The Essence of Chromatography

(CH3)3SiO-4-;

-Si(CH3)3

y(^

R =-(CH2)3-0-
^002-1

V0CH3

DB-5 MS DBT

2-Me 4.Me \ /

3-Me 1-Me

UUi

M ' " •! ' • " ! " " I

27

28

29

30

31

32

33

' " M

34

" " I ' ' "

35

36

I "

37

' 'I • ' "

38

I • ' '

39

SB-Smectic DBT

• I " " I' 9 10

11

12

13 14 15 Tiwm (min)

16

17

18

19

Figure 10.11. Structure of the SB-smectic side-chain Hquid-crystalline poly(siloxane) stationary phase. The poly(methylsiloxane) backbone is substituted with about 50 % 3-[4'-(4-methoxyphenoxycarboxyl)biphenyl-4-yloxy]propyl groups. Separation of dibenzothiophene (DBT) and four methyl and three ethyl isomes on a poly(dimethyldiphenylsiloxane) stationary phase containing 5 mol % phenyl (DB-5 MS) and the SB-smectic side-chain liquid-crystalline poly(siloxane) stationary phase. (Fromref. [268]; ©Elsevier).

teristics, poor thermal stability, high column bleed, and limited temperature operating ranges. These shortcomings were largely overcome by the development of sidechain liquid-crystalUne poly(siloxane) phases [264-267]. The biphenylcarboxylate ester poly(siloxane). Figure 10.11, is the most widely used liquid-crystalline stationary phase. It can be immobiUzed on open tubular columns, and has a useful temperature range from about 100 to 250°C isothermal (270°C temperature programmed). This phase has favorable properties for the separation of alkyl substituted, and ring heteroatom substituted, isomers of poly cyclic aromatic compounds [268].

Separation of Stereoisomers

839

The general elution order of geometric and positional isomers of rigid molecules, on liquid-crystalline stationary phases in gas chromatography, correlates with their lengthto-breadth ratio and planarity [268,269]. The length-to-breadth ratio and thickness are calculated as the dimensions of a box, drawn to enclose the atoms of the molecule. Retention normally increases with the length-to-breadth ratio with differences in planarity (thickness), vapor pressure, and polarity being important in some cases, possibly leading to inversion of the predicted order. Long and planar molecules fit better into the ordered structure of the liquid-crystalline phase, whereas nonlinear and nonplanar molecules do not permeate so easily between the liquid crystal molecules, and are less retained. The elution order of polycyclic aromatic compounds on the SB-smectic phase (Figure 10.11) is strongly correlated with the reversed-phase liquid chromatographic retention of the same compounds on polymeric octadecylsiloxanebonded silica sorbents [269]. No single phase is likely to separate all components of a complex mixture of isomers. A liquid-crystalline phase is a useful component of a multicolumn strategy for gas chromatography, because of its complementary separation mechanism to phases that separate isomers based on vapor pressure or polarity differences. Liquid-crystalline coated and chemically bonded phases have demonstrated limited shape selectivity in liquid chromatography [261,270-274] and modest success in supercritical fluid chromatography [261,275,276]. The low bonding density of siloxanebonded phases probably contributes to the loss of long-distance order in the bonded phase structure. Shape-selectivity of these phases is generally similar to polymeric octadecylsiloxane-bonded phases (section 4.2.2.1) indicating that the contribution of the liquid-crystalline substituent is of limited importance. As for gas chromatography, side-chain polymeric liquid-crystalline stationary phases show greater retention of the liquid-crystalline order, and better prospects for shape-selective separations. Useful applications have still to be demonstrated for these phases, however.

10.9 REFERENCES [1] W. J. Lough (Ed.), Chiral Liquid Chromatography, Blackie, Glasgow, 1989. [2] I. W. Wainer (Ed.), Drug Stereochemistry: Analytical Methods in Pharmacology, Dekker, New York, NY, 1993. [3] G. Subramanian (Ed.), A Practical Approach to Chiral Separations by Liquid Chromatography, WileyVCH, Weinheim, 1994. [4] P. Schreier, A. Bemreuther and M. Huffer, Analysis of Chiral Organic Molecules - Methodology and Applications, Walter de Gruyter, Berlin, 1995. [5] G. P. Moss, Pure & Appl. Chem. 68 (1996) 2193. [6] V. A. Davankov, Pure & Appl. Chem. 69 (1997) 1469. [7] R. S. Cahn, C. Ingold and V. Prelog, Angew. Chem. Intemat. Edit. 5 (1966) 385. [8] N. M. Maier, P Franco and W. Lindner, J. Chromatogr. A 906 (2001) 3. [9] E. R. Waite, M. J. Collins, S. Ritz-Timme, H. W. Schutz, C, Cattano and H. I. M. Borrman, Forensic Sci.Int. 103(1999)113. [10] A. Mosandl, Food Rev. Intemat. 11 (1995) 597.

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The Essence of Chromatography

[11] W. A. Konig, C. Fricke, Y. Saritas, B. Momeni and G. Hohenfeld, J. High Resolut. Chromatogr. 20 (1997)55. [12] M. Juza, H. Jalcubetz, H. Hettesheimer and V. Schurig, J. Chromatogr. B 735 (1999) 93. [13] J. Ducharme, C. Fernandez, F. Gimenez and R. Farinotti, J. Chromatogr. B 686 (1996) 65. [14] W. H. Pirlde, D. W. House and J. M. Finn, J. Chromatogr. 192 (1980) 143. [15] G. Gotmar, T. Fomstedt and G. Guiochon, Chirality 12 (2000) 558. [16] V. Schurig, J. Chromatogr. A (2002) in press. [17] V. A. Davankov, J. Chromatogr. A 666 (1994) 55. [18] T. D. Booth and I. W. Wainer, J. Chromatogr. A 737 (1996) 157. [19] R Piras, C. Roussel and J. Pierrot-Sanders, J. Chromatogr. A 906 (2001) 301. [20] B. Koppenhoefer, R. Graf, H. Holzschuh, A. Nothdurft, U. Trettin, R Piras and C. Roussel, J. Chromatogr. A 666 (1994) 557. [21] C. J. Welch, J. Chromatogr. A 666 (1994) 3. [22] M. Wedig, S. Laug, T. Christians, M. Thunhorst and U. Holzgrabe, J. Pharm. Biomed. Anal. 27 (2002)531. [23] J. R Wolbach, D. K. Lloyd and I. W. Wainer, J. Chromatogr. A 914 (2001) 299. [24] K. B. Lipkowitz, J. Chromatogr. A 906 (2001) 417. [25] K. B. Lipkowitz, G. Pearl, B. Coner and M. A. Peterson, J. Am. Chem. Soc. 119 (1997) 600. [26] M. Okamoto, J. Pharm. Biomed. Anal. 27 (2002) 401. [27] B.-A. Persson and S. Andersson, J. Chromatogr. A 906 (2001) 195. [28] I. D. Wilson, E. R. Adlard, M. Cooke and C. F. Poole (Eds.), Encyclopedia of Separation Science, Academic Press, London, vol. 5, 2000. [29] K. Valko (Ed.), Separation Methods in Drug Synthesis and Purification, Elsevier, Amsterdam, 2000. [30] T. E. Beesley and R. R W. Scott, Chiral Chromatography, Wiley, Chichester, 1998. [31] S. AUenmark, Chromatographic Enantioseparations, Methods and Applications, Ellis Horwood, Chichester, 1991. [32] A. M. Krstulovic (Ed.), Chiral Separations by HPLC. Applications to Pharmaceutical Compounds, Ellis Horwood, Chichester, 1989. [33] W, A. Konig, Gas Chromatographic Enantiomer Separation with Modified Cyclodextrins, Huthig, Heidelberg, 1992. [34] W A. Konig, Trends Anal. Chem. 12 (1993) 130. [35] V. Schurig, J. Chromatogr. A 666 (1994) 111. [36] Z. Juvancz and R Petersson, J. Microcol. Sep. 8 (1996) 99. [37] W Vetter and W. Schurig, J. Chromatogr. A 774 (1997) 143. [38] V. Schurig, J. Chromatogr. A 906 (2001) 275. [39] K. Anton and C. Berger, Supercritical Fluid Chromatography with Packed Columns: Techniques and Applications, Dekker, New York, 1997. [40] K. L. WiUiams and L. C. Sander, J. Chromatogr. A 785 (1997) 149. [41] K. L. Williams and L. C. Sanders and S. A. Wise, J. Pharm. Biomed. Anal. 15 (1997) 1789. [42] C. E Poole, J. Biochem. Biophys. Methods 43 (2000) 3. [43] G. Terfloth, J. Chromatogr. A 906 (2001) 301. [44] D. Wisuba and V. Schurig, J. Chromatogr. A 875 (2000) 255. [45] S. Fanah, R Catarcini, G. Blaschke and B. Chankvetadze, Electrophoresis 22 (2001) 3131. [46] M. Lammerhofer, F. Svec, J. M. J. Frechet and W Lindner, Trends Anal. Chem. 19 (2000) 676. [47] B. Chankvetadze, J. Sep. Sci. 24 (2001) 691. [48] J. Haginaka, J. Pharm. Biomed. Anal. 27 (2002) 357. [49] G. K. E. Scriba, J. Pharm. Biomed. Anal. 27 (2002) 373. [50] B. Chankvetadze and G. Blaschke, J. Chromatogr. A 906 (2001) 309. [51] S Wren, Chromatographia 54 (2001) S-1 [52] A. Rizzi, Electrophoresis 22 (2001) 3079. [53] A. Amini, Electrophoresis 22 (2001) 3107.

Separation of Stereoisomers [54] [55] [56] [57] [58] [59] [60] [61] [62] [63] [64] [65] [66] [67] [68] [69] [70] [71] [72] [73] [74] [75] [76] [77] [78] [79] [80] [81] [82] [83] [84] [85] [86] [87] [88] [89] [90] [91] [92] [93] [94] [95]

841

B. Chakvetadze, Trends Anal. Chem. 18 (1999) 485. G. Gubitz and M. G. Schmid, J. Chromatogr. A 792 (1997) 179. B. Chankvetadze, Capillary Electrophoresis in Chiral Analysis, Wiley, Chichester, 1997. S. FinaH, J. Chromatogr. A 735 (1996) 77. J. Sherma and B. Fried (Eds.), Handbook of Thin-Layer Chromatography, Dekker, New York, 1997. L. Lepri, J. Planar Chromatogr. 10 (1997) 321. Sz. Nyiredy (Ed.), Planar Chromatography. A Retrospective View for the Third Millennium, Springer, Budapest, 2001. I. W. Wainer, Trends Anal. Chem. 6 (1987) 125. J. L. Atwood, J. E. D. Davies, D. D. MacNicol and F. Vogtle (Eds.), Comprehensive Supramolecular Chemistry, Pergamon, Oxford, vol. 3, 1996. F. Bressolle, M. Audran, T. N. Pham and J. J. Vallon, J. Chromatogr. B 687 (1996) 303. J. SzejtH, Chem. Rev. 98 (1998) 1743. E. Schneiderman and A. M. Stalcup, J. Chromatogr. B 745 (2000) 83. S. Fanah, J. Chromatogr. A 875 (2000) 89. Z. Juvanez and J. Szejth, Trends Anal. Chem. 21 (2002) 379. W. A. Konig, R. Krebber, P Evers and G. Bruhn, J. High Resolut. Chromatogr. 13 (1990) 328. D. W. Armstrong, W. Li and C.-D. Chang, Anal. Chem. 62 (1990) 914. D. W. Armstrong, W. Li, A. M. Stalcup, H. V. Secor, R. R. Izac and J. I. Seeman, Anal. Chim. Acta 234 (1990) 365. L. Lindquist and P E. Jansson, J. Chromatogr. A 767 (1997) 325. D. Q. Xiao, Y. Ling, R. N. Fu, J. L. Gu, Z. T. Zhao, R. J. Dai, B. Q. Che and A. Q. Luo, Chromatographia 47 (1998) 557. J. L. Anderson, J. Ding, R. D. McCuUa, W. S. Jenks and D. W. Armstrong, J. Chromatogr. A 946 (2002) 197. A. Berthod, W. Li, D. W. Armstrong, Anal. Chem. 64 (1992) 873. A. Dietrich, B. Maas and A. Mosandl, J. High Resolut. Chromatogr. 18 (1995) 152. I. Spanik, P. Oswald, J. Krupcik, E. Benicka, P. Sandra and D. W. Armstrong, J. Sep. Sci. 25 (2002) 45. T. Beck, J. M. Liepe, J. Nandzik, S. Rohn and A. Mosandl, J. High Resolut. Chromatogr. 23 (2000) 569. B. Maas, A. Dietrich and A. Mosandl, J. Microcol. Sep. 8 (1996) 47. T. Beier and H. D. Holtje, J. Chromatogr. B 708 (1995) 1. R. Reinhardt, M. Richter, P. P Mager, P. Hennig and W. Engelwald, Chromatographia 43 (1996) 187. R. Cardinael, E. Ndzie, S. Petit, G. Coquerel, V. Combret and J. C. Combret, J. High Resolut. Chromatogr. 20 (1997) 560. C. Bicchi, A. Damato, V. Manzin, A. GaUi and M. GaUi, J. High Resolut. Chromatogr. 18 (1995) 295. C. Bicchi, C. BruneUi, G. Cravotto, P Rubiolo and M. Galh, J. Sep. Sci. 25 (2002) 125. C. Bicchi, G. Cravotto, A. D'Amato, P Rubiolo, A. Galh and M. GalH, J. Microcol. Sep. 11 (1999) 487. T. Beck, J. Nandzik and A. Mosandl, J. Microcol. Sep. 12 (2000) 482. E. Miranda, F. Sanchez, J. Sanz, M. I. Jimenez and I. Martinez-Castro, J. High Resolut. Chromatogr. 21(1998) 225. B. E. Kim, S. H. Lee, K. S. Park, K. P Lee and J. H. Park, J. High Resolut. Chromatogr. 20 (1997) 208. B. Maas, A. Dietrich, D. Bartschat and A. Mosandl, J. Chromatogr. Sci. 33 (1995) 223. W. Vetter, U. Klobes, B. Luckas and G. Hottinger, J. Chromatogr. A 846 (1999) 375. W. Vetter and V. Schurig, J. Chromatogr. A 774 (1997) 143. Y Tang, Y Zhou and D. W. Armstrong, J. Chromatogr. A 666 (1994) 147. M. Jung and V. Schurig, J. High Resolut. Chromatogr. 16 (1993) 289. J. Donnecke, W. A. Konig, O. Gyllenhaal, J. Vessman and C. Schulze, J. High Resolut. Chromatogr. 17 (1994) 779. J. Donnecke, C. Paul, W. A. Konig, L. A. Svensson, O. Gyllenhaal and J. Vessman, J. Microcol. Sep. 8(1996) 495. V. Schurig, M. Jung, S. Mayer, M. Fluck, S. Negura and H. Jakubetz, J. Chromatogr. A 694 (1995) 119.

842 [96] [97] [98] [99] [100] [101] [102] [103] [104] [105] [106] [107] [108] [109] [110] [Ill] [112] [113] [114] [115] [116] [117] [118] [119] [120] [121] [122] [123] [124] [125] [126] [127] [128] [129] [130] [131] [132] [133] [134] [135] [136] [137] [138] [139] [140]

The Essence of Chromatography H. Grosenick and V. Schurig, J. Chromatogr. A 761 (1997) 181. H. Dittmann and W. A. Konig, J. High Resolut. Chromatogr. 23 (2000) 583. T. AraJd, S. Tsunoi and M. TanaJca, Anal. Chim. Acta 410 (2000) 37. L. Chen, L. F. Zhang, C. B. Ching and S. C. Ng, J. Chromatogr. A 950 (2002) 65. S. M. Han, Biomed. Chromatogr. 11 (1997) 259. S. C. Chang, G. L. Reid, S. Chen, C. D. Chang and D. W. Armstrong, Trends Anal. Chem. 12 (1993) 144. A. Berthod, S. Chang and D. W. Armstrong, Anal. Chem. 64 (1992) 395. R. J. Steffeck, Y. Zelechonok and K. H. Gahm, J. Chromatogr. A 947 (2002) 301. R. A. Thompson, Z. H. Ge, N. Grinberg, D. EUison and R Tway, Anal. Chem. 67 (1995) 1580. M. H. Hyun, S. C. Han, B. H. Lipshutz, Y.-J. Shin and C. J. Welch, J. Chromatogr. A 959 (2002) 75. X. C. Zhou, H. Yan, Y. Y Chen, C. Y Wu and X. R. Lu, J. Chromatogr. A 753 (1996) 269. K. Oguni, H. Oda and A. Ichida, J. Chromatogr. A 694 (1995) 91. Y Okamoto and E. Yashima, Angew. Chem. Intemat. Edit. 37 (1998) 1021. E. Yashima, J. Chromatogr. A 906 (2001) 105. R Franco, A. Senso, L. Oliveros and C. Minguillon, J. Chromatogr. A 906 (2001) 155. C. Perrin, V. A. Vu, N. Maththijs, M. Maftouch, D. L. Massart and Y Vander Heyden, J. Chromatogr. A 947 (2002) 69. Y K. Ye, B. Lord and R. W. Stringham, J. Chromatogr. A 945 (2002) 139 K. Tachibana and A. Ohnishi, J. Chromatogr. A 906 (2001) 127. L. Y Tang, D. Xiang and J. A. Blackwell, Enantiomer 5 (2000) 345. Y K. Ye, B. S. Lord, L. Yin and R. W. Stringham, J. Chromatogr. A 945 (2002) 147. T. Nakano, J. Chromatogr. A 906 (2001) 205. A. Medvedovici, R Sandra, L. Toriba and F. David, 785 (1997) 159. A. van Overbeke, R Sandra and A. Medvedovici, W. Baeyens and H. Y Aboulenein, Chirality 9 (1997) 126. S. Svensson, A. Karlsson, O. Gyllenhaal and J. Vessman, Chromatographia 51 (2000) 283 K. W. Phinney, Anal. Chem. 72 (2000) 204A. M. Garzotti and M. Hamdan, J. Chromatogr. B 770 (2002) 53. A. Kot, R Sandra and V. Venema, J. Chromatogr. Sci. 32 (1994) 439. J. Donnecke, L. A. Svensson, O. Gyllenhaal, K. E. Karlsson and J. Vessman, J. Microcol. Sep. 11 (1999)521. S. Chen, Y Zhou, C. Bagwill and J. R. Chen, Anal. Chem. 66 (1994) 1473. D. W. Armstrong, Y Liu and K. H. Ekborg-Ott, Chirahty 7 (1995) 474. A. Berthod, X. Liu, C. Bagwill and D. W. Armstrong, J. Chromatogr. A 731 (1996) 123. A. Berthod, T. L. Xiao, Y Liu, W. S. Jenks and D. W. Armstrong, J. Chromatogr. A 955 (2002) 53. H. Y Aboul-Enein and I. Ali, Chromatographia 52 (2000) 679. T. J. Ward and A. B. Farris, J. Chromatogr. A 906 (2001) 73. K. H. Ekborg-Ott, J. R Kullman, X. D. Wang, K. Gahm, L. F He and D. W. Armstrong, Chirality 10 (1998) 627. S. G. Allenmark and S. Andersson, J. Chromatogr. A 666 (1994) 167. J. Hermansson, Trends Anal. Chem. 9 (1989) 251. J. Hermansson and A. Grahn, J. Chromatogr. A 694 (1995) 57. J. Haginaka, J. Chromatogr. A 906 (2001) 253. B. Feibush, Chrirality 10 (1998) 382. I. Abe, T. Nishiyama and H. Frank, J. High Resolut. Chromatogr. 17 (1994) 9. I. Abe, K. Terada and T. Nakahara, J. High Resolut. Chromatogr. 19 (1996) 91. I. Abe, K. Terada, T. Nakahara and H. Frank, J. High Resolut. Chromatogr. 21 (1998) 592. B. Koppenhoefer, U. Muhleck and K. Lohmiller, J. Chromatogr. A 699 (1995) 215. W. A. Konig, The Practice of Enantiomer Separation by Capillary Gas Chromatography, Huethig, Heidelberg, 1987.

Separation of Stereoisomers [141] [142] [143] [144] [145] [146] [147] [148] [149] [150] [151] [152] [153] [154] [155] [156] [157] [158] [159] [160] [161] [162] [163] [164] [165] [166] [167] [168] [169] [170] [171] [172] [173] [174] [175] [176] [177] [178] [179] [180] [181] [182] [183] [184] [185] [186] [187] [188] [189]

843

B. Koppenhoefer and E. Bayer, Chromatographia 19 (1989) 123. H. Frank, G. Nicholson and E. Bayer, J. Chromatogr. 167 (1978) 187. V. Schurig, M. Juza, M. Preschel, G. J. Nicholson and E. Bayer, Enantiomer 4 (1999) 297. C. J. Welch, Adv. Chromatogr. 35 (1995) 171. F. Gasparrini, D. Misiti and C. Villani, J. Chromatogr. A 906 (2001) 35. N. Oi, H. Kitahara and E Aoki, J. Chromatogr. A 694 (1995) 129. N. M. Maier and G. Uray, J. Chromatogr. A 740 (1996) 11. M. H. Hyun, J. J. Ryoo and W. H. Pirlde, J. Chromatogr. A 886 (2000) 47. W. Golkiewicz and B. Polak, Anal. Chem. 43 (1998) 591. C. J. Welch, M. N. Protopopova and G. Bhat, Enantiomer 3 (1998) 471. W. H. Pirkle, C. J. Welch and A. J. Zych J. Chromatogr. 648 (1993) 101. G. J. Terfloth, W. H. Pirlde, K. G. Lynam and E. C. Nicolas, J. Chromatogr. A 705 (1995) 185. M. Hedeland, R. Isaksson and C. Pettersson, J. Chromatogr. A 807 (1998) 297. M. Josefsson, B. Carlsson and B. Norlander, J. Chromatogr. A 684 (1994) 23. E. Heldin, K. J. Lindner, C. Pettersson, W Lindner and R. Rao, Chromatographia 32 (1991) 407. L. I. IVlonser and G. M. Greenway, Analyst 122 (1997) 719. A. Karlsson and C. Pettersson, J. Chromatogr. 543 (1991) 287. A. Karlsson and C. Pettersson, Chirality 4 (1992) 323. N. H. Huynh, A. Karlsson and C. Pettersson, J. Chromatogr. A 705 (1995) 275. A. Karlsson and C. Charron, J. Chromatogr. A 732 (1996) 245. A. Karlsson and G. Karlsson, Chirality 9 (1997) 650. C. Roussel and A. Favrou, J. Chromatogr. A 704 (1995) 67. E. Ameyibor and J. T. Stewart, J. Chromatogr. B 703 (1997) 273. P K. Owens, A. F Fell, M. W Coleman and J. C. Berridge, J. Chromatogr. A 797 (1998) 187. A. Bielejewska, R. Nowakowski, K. Duszczyk and D.Sybilska, J. Chromatogr. A 840 (1999) 159. K. A. Connors, Chem. Revs. 97 (1997) 1325. A. Bielejewska, K. Duszczyk and D. Sybilska, J. Chromatogr. A 931 (2001) 81. A. Salvador, B. Herbreteau and M. Dreux, Chromatographia 53 (2001) 207. A. Salvador, B. Herbreteau, M. Dreux, M. Karlsson and O. Gyllenhaal, J. Chromatogr. A 929 (2001) 101. ' L. Lepri, A. CincineUi and M. Del Bubba, J. Planar Chromatogr. 12 (1999) 298. L. Lepri, M. Del Bubba, A. CincineUi and I. Boddi, J. Planar Chromatogr. 13 (2000) 384. L. Lepri, M. Del Bubba, A. CincineUi and I. Boddi, J. Planar Chromatogr. 14 (2001) 134. L. Lepri, M. Del Bubba, V. Coas and A. CincineUi, J. Liq. Chromatogr. & Rel. Technol. 22 (1999) 105. L. Lepri, V. Coas, M. Del Bubba and A. CincineUi, J. Planar Chromatogr. 12 (1999) 221. D. W Armstrong and Z. Zhou, J. Liq. Chromatogr. 17 (1994) 1695. A. M. Tivert and A. Backman, J. Planar Chromatogr. 6 (1993) 216. Y. Bereznitski, R. Thompson, E. O'NeiU and N. Grinsberg, J. AOAC Int. 84 (2001) 1242. W Zhu and G. Vigh, Electrophoresis 21 (2000) 2016. B. A. WiUiams and G. Vigh, J. Chromatogr. A 777 (1997) 295. B. Chankvetadze, G. Schulte and G. Blaschke, Enantiomer 2 (1997) 157. S. Finah, J. Chromatogr. A 792 (1997) 227. B. Chankvetadze, G. Endresz and G. Blaschke, Chem. Soc. Rev. 25 (1996) 141. M. R. Hadley, M. Decrette, G. Guillore, C. Rosini, M. L DonnoU, S. Superchi and A. J. Hutt, J. Sep. Sci. 24 (2001) 766. J. Zukowski, J. De Biasi and A. Berthod, J. Chromatogr. A 948 (2002) 331. H. Nishi, J. Chromatogr. A 735 (1996) 57. K. Otsuka and S. Terabe, J. Chromatogr. A 875 (2000) 163. H. H. Yarabe, E. BiUiot and I. M. Warner, J. Chromatogr. A 875 (2000) 179. J. Haginaka, J. Chromatogr. A 875 (2000) 235. D. K. Lloyd, S. Li and P Ryan, J. Chromatogr. A 694 (1995) 285.

844 [190] [191] [192] [193] [194] [195] [196] [197] [198] [199] [200] [201] [202] [203] [204] [205] [206] [207] [208] [209] [210] [211] [212] [213] [214] [215] [216] [217] [218] [219] [220] [221] [222] [223] [224] [225] [226] [227] [228] [229] [230] [231] [232] [233] [234] [235]

The Essence of Chromatography T. J. Ward and T. M. Oswald, J. Chromatogr. A 792 (1997) 309. H. Nishi and Y. Kuwahara, J. Biochem. Biophys. Methods 48 (2001) 89. H. Nishi and Y Kuwahara, J. Pharm. Biomed. Anal. 27 (2002) 577. F. Wang and M. G. Khaledi, J. Chromatogr. A 875 (2000) 277. M.-L. Riekkola, S. K. Wiedmer, I. E. Valko and H. Siren, J. Chromatogr. A 792 (1997) 13. W. Zhu and G. Vigh, J. Chromatogr. A 892 (2000) 499. W. Zhu and G. Vigh, J. Microcol. Sep. 12 (2000) 167. T. Christians and U. Hoizgrabe, J. Chromatogr. A 911 (2001) 249. D. K. Maynard and G. Vigh, Electrophoresis 22 (2001) 3152. L. Zhou, R. Thompson, S. Song, D. EUison and J. M. Wyvratt, J. Pharm. Biomed. Appl. 27 (2002) 541. M. Fillet, R Hubert and J. Crommen, Electrophoresis 19 (1998) 2834. M. Fillet, R Hubert and J. Crommen, J. Chromatogr. A 875 (2000) 123. I. S. Lurie, J. Chromatogr. A 792 (1997) 297. A. M. Abushoffa, M. Fillet, R Hubert and J. Crommen, J. Chromatogr. A 948 (2002) 321. M. Ju and Z. El Rassi, Electrophoresis 20 (1999) 2766. H. Nishi, J. Chromatogr. A 792 (1997) 327. R. Gotti, R. Pomponio and V. Cavrini, Chromatographia 52 (2000) 273. Y Tanaka and S. Terabe, J. Chromatogr. A 694 (1995) 277. E. Oliveri, R. Sebastiano, A. Citterio, C. Gelfi and R G. Righetti, J. Chromatogr. A 894 (2000) 273 and 281. Y Tanaka, K. Otsuka and S. Terabe, J. Chromatogr. A 875 (2000) 323. M. Lammerhofer, E. Zarbi and W. Lindner, J. Chromatogr. A 892 (2000) 509. D. Cagniant (Ed.), Complexation Chromatography, Marcel Dekker, New York, NY, 1991. I. D. Wilson, E. R. Adlard, M. Cooke and C. F. Poole (Eds.), Encyclopedia of Separation Science, Academic Press, London, v. 9, 2000. G. Dobson, W. W. Christie and B. Nikolova-Damyanova, J. Chromatogr. B 671 (1995) 197. B. Nikolova-Damyanova, W. W. Christie and B. Herslof, J. Chromatogr. A 749 (1996) 47. R. O. Adlof, J. Chromatogr. A 764 (1997) 337. M. Buchgraber and F. Ulberth, J. AOAC Intemat. 84 (2001) 1490. R Fevrier, A. Binet, L. Dufosse, R. Gree and F. Yvergnaux, J. Chromatogr. A 923 (2001) 53. R. E Cross and H. A. Widman, J. Sep. Sci. 25 (2002) 239 and 245. B. Nikolova-Damyanova, W. W. Christie and B. Herslof, J. Planar Chromatogr. 7 (1994) 382. M. Demirbuker and L. G. Blomberg, J. Chromatogr. 550 (1991) 765. L. G. Blomberg, M. Demirbuker and R E. Andersson, J. Am. Oil Chem. Soc. 70 (1993) 939. Y Shen, S. L. Reese, B. E. Rossiter and M. L. Lee, J. Microcol. Sep. 7 (1995) 279. V. A. Davankov, J. D. Navratil and H. F Walton, Ligand Exchange Chromatography, CRC Press, Boca Raton, FL, 1988. V. A. Davankov, Enantiomer 5 (2000) 209. A. Kurganov, J. Chromatogr. A 906 (2001) 51. M. G. Schmid, N. Grobuschek, O. Lecnik and G. Gubitz, J. Biochem. Biophys. Methods 48 (2001) 143. G. Golavema, R. Corradini, A. Dossena, E. Chiavaro, R. Marchelli, F. Dallavalle and G. Folesani, J. Chromatogr. A 829 (1998) 101. M. RemeUi, P Fomasari and P Puhdori, J. Chromatogr. A 761 (1997) 79. Q. H. Wan, P N. Shaw, M. C. Davies and D. A. Barrett, J. Chromatogr. A 765 (1997) 187. Q. H. Wan, P N. Shaw, M. C. Davies and D. A. Barrett, J. Chromatogr. A 786 (1997) 249. V. Schurig and M. Fluck, J. Biochem. Biophys. Methods 43 (2000) 223. H. Lingeman and W. J. M. Underberg (Ed.), Detection-Oriented Derivatization Techniques in Liquid Chromatography, Marcel Dekker, New York, NY, 1990. K. Blau and J. Halker (Eds.), Handbook of Derivatives for Chromatography, Wiley, Chichester, 1993. S. Gorog and M. Gazdag, J. Chromatogr. B 659 (1994) 51. T. Toyo'oka, Biomed. Chromatogr. 10 (1996) 265.

Separation of Stereoisomers [236] [237] [238] [239] [240] [241] [242] [243] [244] [245] [246] [247] [248] [249] [250] [251] [252] [253] [254] [255] [256] [257] [258] [259] [260] [261] [262] [263] [264] [265] [266] [267] [268] [269] [270] [271] [272] [273] [274] [275] [276]

845

T. Toyo'oka (Ed.), Modern Derivatization Methods for Separation Sciences, Wiley, Chichester, 1999. F. Yasuhara, S. Yamaguchi, M. Takeda, Y Ochiai and S. Miyano, J. Chromatogr. A 694 (1995) 227. R. Buschges, H. Linde, E. Mutschler and H. Spahnl-Langguth, J. Chromatogr. A 725 (1996) 323. O. P. Kleidemigg and W. Lindner, Chromatographia 44 (1997) 465. D. R. Jin, K. Nagakura, S. Murofushi, T. miyahara and T. Toyo'oka, J. Chromatogr. A 822 (1998) 215. R. Herraez-Hemandez, R Campins-Falco and L. A. Tortajada-Genaro, Analyst 123 (1998) 2131. D. R. Jin and T. Toyo'oka, Analyst 123 (1998) 1271. M. Peter and F. Fulop, J. Liq. Chromatogr. & Rel. Technol. 23 (2000) 2459. M. Peter, A. Peter and F Fulop, J. Chromatogr. A 871 (2000) 115. K. H. Kim, H. J. Kim, J. H. Kim, J. H. Lee and S. C. Lee, J. Pharm. Biomed. Anal. 25 (2001) 947. M. Peter, A. Gyeresi and F. Fulop, J. Chromatogr. A 910 (2001) 247. K. H. Kim, J. H. Lee, M. Y Ko, K. S. Shin, J. S. Kang, W. C. Mar and J. R. Youm, Chromatographia 55 (2002) 81. K. Harada, A. Matsui, Y Shimizu, R. Ikemoto and K. Fujii, J. Chromatogr. A 921 (2001) 187. M. Bruckner and M. Leitenberger, Chromatographia 42 (1996) 683. Y Zhou, P Luan, L. Liu and Z. P Sun, J. Chromatogr. B 659 (1994) 109. B. Fransson and U. Ragnarsson, J. Chromatogr. A 827 (1998) 31. T. Arai, J. Chromatogr. B 717 (1998) 295. H. Wan and L. G. Blomberg, J. Chromatogr. A 875 (2000) 43. L. G. Blomberg and H. Wan, Electrophoresis 21 (2000) 1940. J. C. M. Waterval, H. Lingeman, A. Bult and W J. M. Underberg, Electrophoresis 21 (2000) 4029. O. P Kleidemigg and W Lindner, J. Chromatogr. A795 (1998) 251. G. Thorsen, A. Engstrom and B. Josefsson, J. Chromatogr. A 786 (1997) 347. G. M. Janini, Adv. Chromatogr. 17 (1979) 231. Z. Witkiewicz, J. Chromatogr. 466 (1989) 37. T. J. Betts, J. Chromatogr. 641 (1993) 189 F. Gritti and G. Felix, Chromatographia 55 (2002) 523. F. Perez, P. Berdague, J. Courtieu, J. P. Boyle, S. Boudah and M. H. Guermouche, J. High Resolut. Chromatogr. 20 (1997) 379. F. Ammar-Khodja, S. Guermouche, M. H. Guermouche, P. Berdague and J. P. Boyle, Chromatographia 30 (1999) 338. N. Nishioka, B. A. Jones, B. J. Tarbet, J. S. Tarbet, J. S. Bradshaw and M. L. Lee, J. Chromatogr. 357 (1986) 79. J. S. Bradshaw, C. M. Schregenberger, H.-C. Chang, K. E. Markides and M. L. Lee, J. Chromatogr. 358 (1986) 95. K. P Naikwadi and P P Wadgonkar, J. Chromatogr. A 811 (1998) 97. G.-P Chang-Chien, J. Chromatogr. A 808 (1998) 201. S. G. Mossner, M. J. Lopez de Alda, L. C. Sander, M. L. Lee and S. A. Wise, J. Chromatogr. A 841 (1999) 207. L. C. Sander, M. Schneider, S. A. Wise and C. Woolley, J. Microcol. Sep. 6 (1994) 115. J. J. Pesek, M. T. Matyska, E. J. WiUiamsen, R. Tarn and Z. X. Wang, J. Liq. Chromatogr. & Rel. Technol. 21 (1998) 2747. J. J. Pesek, M. T. Matyska and S. Muley, Chromatographia 52 (2000) 439. O. Ferroukhi, N. Atik, S. Guermouche, M. H. Guermouche, P. Berdagne, P. Judenstein and J. P. Bayle, Chromatographia 52 (2000) 564. F. Gritti, G. Felix, M. F. Achard and E Hardouin, J. Chromatogr. A 922 (2001) 51. F. Gritti, S. Sourigues, G. Felix, M. F. Achard and F. Hardouin, Chromatographia 55 (2002) 149. Y Shen, J, S. Bradshaw and M. L. Lee, Chromatographia 43 (1996) 53. F. Gritti, G. Felix, M. F. Achard and F. Hardouin, Chromatographia 53 (2001) 201.