Ion-Exchange

Ion-exchange chromatography relies on the exchange of ions between charged groups bound to a stationary phase and ions of opposite charge in the mobile phase (see Figure 13-3). Depending on the charge of the stationary phase, components binding to the column may be cations (positively charged) or anions (negatively charged). Cation-exchange solid phases contain covalently bound, negatively charged functional groups. Examples include strongly acidic groups, such as sulfonate ions, or weakly acidic groups, such as carboxyl or carboxymethyl. Anion-exchange solid phases have strongly basic quaternary amines, such as triethylaminoethyl groups, or weakly basic groups, such as aminoethyl (AE) and diethylaminoethyl (DEAE) groups. Separations are performed under conditions where analytes are charged and bind to the ion-exchange stationary phase. Solutes are eluted with a solution containing competing counterions, such as sodium for cation exchange and chloride for anion exchange, and, in some cases, by adjustment of pH to decrease the charge of bound ions or the ion-exchange surface. Important factors in performance of ion-exchange chromatography include (1) type of stationary phase ionic group, (2) charge density, (3) stationary phase matrix, (4) type and concentration of ions in the mobile phase, and (5) mobile phase pH. Many stationary phases exhibit mixed mode retention by a combination of ion exchange and adsorptive binding. As an example, ion-exchange resins used for amino acid analysis are able to separate amino acids with virtually the same charge because of variable adsorptive interactions of different amino acids with the stationary phase. Some solid-phase extraction media are designed to have mixed hydrophobic–ion-exchange retention, offering improved selectivity versus extraction media designed predominantly for hydrophobic or ion-exchange extractions.

Ion-exchange chromatography has a number of clinical applications, including analyses of amino acids and hemoglobins. It is also used to analyze small inorganic and organic ions with conductivity detection, a technique termed ion chromatography. Water purification is an important preparative application of ion-exchange chromatography consists of using mixed-bed resins with both cation and anion exchangers. Ions are removed by exchange of hydrogen ions for other cations and hydroxyl ions for other anions. The hydrogen ions and hydroxyl ions displaced by salt ions form water.

Adsorption

The basis of separation by adsorption chromatography is the difference between adsorption and desorption of solutes at the surface of a solid particle (see Figure 13-3). Electrostatic, hydrogen-bonding, and dispersive (van der Waals) interactions are the physical forces that control this type of chromatography.^17,28 As solvent molecules compete for binding to the stationary phase, the strength of binding to the stationary phase depends on the properties of the stationary phase and of the mobile phase. In general, for a polar stationary phase, elution tracks the polarity of the components in a mixture: less polar ones elute first, followed by those of increasing polarity.⁷

In GC, this mode is used to separate low molecular weight compounds such as methyl, ethyl, isopropyl alcohols from compounds that are normally gases at room temperature. It uses particles of support, such as “molecular sieves,” alumina, and styrene-divinylbenzene copolymers. In LC, three types of adsorbents are generally used: (1) nonpolar, (2) acid polar, and (3) basic polar. Nonpolar adsorbents include charcoal and polystyrene-divinylbenzene. The main acidic polar adsorbent is silica gel, the surface silanol (SiOH) groups of which adsorb basic substances. Alumina is the main basic adsorbent for retaining acidic substances. Florisil has also been used as a basic adsorbent when catalytic decomposition of the analyte is observed with alumina.

Hydrophilic interaction liquid chromatography (HILIC) is a type of adsorption chromatography that is used to separate highly polar compounds that form hydrogen bonds with a stationary phase having a relatively polar surface.8,26,41 Examples of stationary phases used for HILIC include (1) unmodified silica, (2) silica with bonded zwitterionic groups, (3) hydroxyl-rich compounds, and (4) amide-rich compounds. Initial binding of solutes to the stationary phase is performed in relatively hydrophobic solvent, and elution is attained with solvents of increasing polarity. A limitation of HILIC for clinical laboratory use is that biological fluids are highly polar and include a substantial quantity of salts that also interact with polar stationary phases. Therefore, the specimen may need to be extracted or modified by the addition of a less polar solvent such as acetonitrile to promote binding to the stationary phase. Interest is growing in the application of HILIC as a separation technique linked to mass spectrometry, but most separations in clinical laboratories have relied on reversed-phase or ion-exchange separations.

Partition Chromatography

The differential distribution of solutes between two immiscible liquids is the basis for separation by partition chromatography (see Figure 13-3).^46,55,56 Operationally, one of the immiscible liquids serves as the stationary phase. To prepare this phase, a thin film of liquid is adsorbed or chemically bonded onto the surface of support particles or onto the inner wall of a capillary column.

In reversed-phase, partition chromatography, the (1) stationary phase is nonpolar, (2) mobile phase is relatively polar, and (3) hydrophobic molecules are preferentially retained. It is the most widely used technique for solid-phase extraction and liquid chromatography in the clinical laboratory. An example of separation of several antidepressant drugs by reversed-phase chromatography is shown in Figure 13-2. Examples of stationary phases for reversed-phase adsorption include silica with bonded hydrophobic groups such as octadecyl (C18), octyl (C8), phenyl, or butyl (C4) and polystyrene-divinylbenzene. For solid-phase extractions, similar materials are used including copolymers of styrene with more hydrophilic monomers that form stationary phases with balanced hydrophobic-hydrophilic character or with charged groups that offer mixed-mode extraction with both hydrophobic and ion-exchange retention.^6,20,55 The retention characteristics of silica-based stationary phases depend on (1) the nature of bonded phases, (2) the amount of bonded phase (often expressed as % carbon load), (3) the surface area of the particles, (4) the pore size of the particles, and (5) the quantity of accessible silanol groups on silica particles that depend on surface treatments such as “end-capping,” where such groups are replaced by trimethylsilyl groups. For reversed-phase separations, solutions are applied in relatively polar solvent such as water or water with a low concentration of organic solvent such as methanol or acetonitrile. Reversed-phase separations are often used with aqueous specimens such as blood and urine, because most of the solvent (water) and inorganic salts pass through the column with minimal retention. The partitioning of solutes in reversed-phase separations usually is favored by adjustment of pH to minimize the charge of solutes, such as decreasing the pH so that organic acids are in uncharged forms, or adding ion-pairing agents that yield less polar ion pairs. Prior to the introduction of a sample, hydrophobic stationary phases may require conditioning or “wetting” with a solvent such as methanol before equilibrating with a more polar solvent. Some stationary phases, such as C18-silica, may undergo “phase collapse” of the bonded phase if run in completely aqueous solvent, which probably represents the aliphatic bonded phase folding down onto the surface and having decreased access to solvent. During a solid-phase extraction, drawing air into the column can similarly require reapplication of organic solvent to wet surfaces. Use of balanced hydrophobic-hydrophilic phases for extractions reduces the need for wetting of the stationary phase and avoids problems of phase collapse with loading of aqueous solutions.^19,55 Polymeric stationary phases usually have higher capacity than bonded silica stationary phases, because the entire particle serves as the stationary phase.

Elution of compounds in reversed-phase separations relies on competition of molecules in the mobile phase with molecules bound to the stationary phase. Elution is hastened by changing the solvent strength with a linear or stepwise gradient, or by changing the pH to increase the polarity of solutes. The strength of solvents, sometimes referred to as the eluotropic series, for reversed-phase separations generally follows this order:

More hydrophobic solvents generally are stronger eluents for reversed-phase separations. For normal-phase chromatography (polar stationary phase, nonpolar mobile phase), the eluotropic series is reversed. In practice, it is also possible to change the solvent strength by using mixtures of solvents and gradients of changing proportions of solvents during an analysis (Figure 13-4).⁹ It is also possible to modify the polarity of aqueous solutions by changing the salt concentration. This characteristic is employed in a variation of reversed-phase chromatography termed hydrophobic interaction chromatography, which has been applied mainly for separation of proteins. Binding to a weakly hydrophobic stationary phase containing interaction groups such as phenyl or butyl groups is promoted by loading specimens in a high-salt concentration, and elution is achieved by lowering the salt concentration rather than by adding an organic solvent.

A wide range of solid phases for reversed-phase separations have slightly different specificities, and some have mixed modes of separation.6,16,39 Some solid phases such as porous graphite or fluorinated hydrocarbons or hydrophobic stationary phases with embedded polar groups offer different selectivities than traditional C18-silica. Stationary phases bonded on silica have weak cation-exchange and normal-phase properties, depending on the quantity of accessible hydroxyl groups on the silica. Basic compounds may exhibit peak tailing caused by ion exchange with the silica. Ion-exchange properties of silica are minimized by chemically modifying the surface to block hydroxyls and by operating at low pH. Silica tends to dissolve slowly at pH > 7, so that silica particles usually require separations below pH 8, unless the silica is stabilized by surface treatment.⁵ Other types of stationary phases such as polystyrene or porous graphite are stable at high pH and allow separations over a broader pH range.

Because of its wide diversity of stationary phases with differing selectivities and varying ability to change solvent conditions and ion-pairing agents, reversed-phase chromatography is a very versatile technique, and separations often achieve high efficiency and, sharp peaks. Compounds representing the greatest challenge for reversed-phase separations are small ions and highly polar compounds such as sugars or amino acids, which tend to be weakly retained by reversed-phase media, and basic compounds, which may exhibit peak tailing as the result of interactions with silica. Derivatization of some compounds such as amino acids has been employed to improve reversed-phase retention and to add chromophores or fluorophores to improve detection.⁴⁸ Separation of large peptides or proteins requires stationary phases with larger pore sizes than are routinely used for separation of small molecules.^3,32,63

Chiral Separations

Many biological molecules occur as specific stereoisomeric forms, such as L-amino acids, and in some cases, only a particular stereoisomer of a therapeutic drug may be active. In general, the two mirror-image forms of a compound or enantiomers have very similar physical properties and do not separate in most chromatographic analyses. Separation of enantiomers generally requires use of a stationary phase that has an enantiomeric component that will interact with molecules in a stereospecific manner. For example, chiral stationary phases (CSPs) that contain groups such as derivatized L-amino acids or stereospecific carbohydrates have been used to separate enantiomers.52,62A,72

Size Exclusion Chromatography

Size exclusion chromatography, also known as (1) gel filtration, (2) gel permeation, (3) steric exclusion, (4) molecular exclusion, or (5) molecular sieve chromatography, is a technique that separates molecules or other particles based on size (Figure 13-5). In this technique, the stationary phase consists of porous particles with an inert surface designed to have minimal adsorption of components. Materials used for stationary phases include cross-linked dextran, polyacrylamide, agarose, polystyrene, and porous glass with an inert surface coating. Beads of these materials are porous and have pore sizes that allow small molecules to enter. Molecules too large to enter the pores remain entirely in the mobile phase and are rapidly eluted from the column. Molecules that are intermediate in size have access to various fractions of the pore volume and elute between large and small molecules according to the following relation:

where V_r is the retention volume, V₀ is the void volume between particles, K is the fraction of the pore volume accessible to the molecule, and V_i is the volume within the support particles.

By calibrating a size-exclusion column with molecules of known size, it is possible to estimate the molecular size of proteins or other molecules.⁶⁴ However, this is a relatively low-resolution technique, and separation of components requires substantial proportional differences in molecular weight. The diameter of globular proteins changes only in proportion to the cube root of molecular weight, so that an eightfold difference in molecular weight is required to yield a twofold change in diameter.⁶⁴ Linear polymers, such as DNA, and proteins denatured with sodium dodecylsulfate or guanidine hydrochloride have a much larger diameter for the same molecular weight, and diameter changes approximately in proportion to the square root of molecular weight, allowing better resolution of linear molecules relative to molecular weight than of globular molecules. Therefore, analysis of molecules under denaturing conditions provides the ability to resolve molecules with smaller differences in molecular weight.

Size exclusion chromatography allows the separation of molecules under physiologic salt conditions; this is useful for identifying intact complexes of molecules such as lipoproteins, antibody-antigen complexes, and other binding proteins and their ligands under mild nondenaturing conditions. Also, it is used as a rapid preparative technique for salt exchange or to remove small molecules from large molecules using small centrifuge or gravity flow columns.

Other Size Exclusion Separations

Separations by size exclusion also are performed with ultrafiltration membranes or particles with defined pore sizes. Ultrafiltration is employed in the clinical laboratory as a preparative technique to separate small molecules such as amino acids or drugs, which pass through the membrane in the ultrafiltrate, from proteins, which remain in the retentate fraction. Ultrafiltration and equilibrium dialysis serve as means of separating free from protein-bound components, such as for measurement of free phenytoin or thyroxine concentrations. Ultrafiltration also is used as a technique for concentrating proteins in dilute solutions, such as urine, for analysis by electrophoresis or other techniques. Ultrafiltration is useful mainly for separating molecules or particles that have large differences in size, typically more than a 10-fold difference in molecular weight.

Affinity Chromatography

Specific molecular interactions are used as the basis for affinity chromatography (Figure 13-6).^25,57 Examples include (1) interactions of antibody with antigen, (2) enzyme with substrate, (3) aptamer with ligand, (4) receptor with ligand, and (5) lectin with sugar. The stationary phase for affinity chromatography is prepared by immobilizing one component of an interaction pair and using it to capture the other component of the interaction pair. Orientation of the capture molecule and its accessibility are important elements in the function of capture molecules. Using a spacer to provide distance from the surface of the stationary phase may improve access and function of the capture molecule. An alternative approach to developing affinity matrices without immobilized ligands is to form molecularly imprinted surfaces on the capture phase.³⁸ The shape and structure of pores formed by molecular imprinting yield selective binding of molecules that have a complementary structure.

To perform affinity chromatography, a solution is applied to the stationary phase under conditions that allow specific binding to the capture molecule, and nonbound components are removed by extensive washing of the stationary phase. Specifically bound components are eluted by adding competing ligand or changing conditions such as pH or ionic strength, so as to eliminate binding to the stationary phase. Affinity separations may be performed batchwise with sedimentation of the stationary phase, or by column chromatography. Chromatographic elution often is seen with stepwise change of solvent to allow elution of desired components in a single fraction.

The power of affinity chromatography lies in its selectivity. Affinity chromatography is widely applied to purifying antibodies and other proteins used in clinical assays. Lectin affinity chromatography offers a means of assessing changes in the oligosaccharide structures bound to glycoproteins. Immobilized phenylboronic acid has been used to separate glycated hemoglobin from unmodified hemoglobin. Most heterogenous assays in the clinical laboratory are based on the selective extraction or capture of a specific target molecule onto a solid phase with elution of all other molecules (see Chapter 16). Immunodepletion or immunopartitioning of high-abundance proteins has been used as a means of preparing biological specimens for proteomic analysis to assist in the detection of lower-abundance components.⁵³

t_r(A) = retention time for solute A

t_r(B) = retention time for solute B

w(A) = peak width (units of time) measured at base for solute A

w(B) = peak width (units of time) measured at base for solute B

Retention Factor

The retention factor k′ is a measure of the time a solute resides in the stationary phase relative to the time it resides in the mobile phase. Mathematically, it is the ratio of the adjusted retention volume(v_r) or retention time (t_r) to the void volume (v₀) or hold-up time (t₀) (the time for unretained components to elute from the column).

A k′ of 0 indicates no binding between a solute and the stationary phase. Greater partitioning into the stationary phase results in longer retention times and increased k′. In practice, it is desirable to have a k′ between 1 and 10 for optimal separations that do not take an excessive length of time.

In planar chromatography, such as paper and thin-layer chromatography, all separation of solutes must occur within the distance traveled by the mobile phase. Thus, a solute’s migration is expressed by its R_f value, which is calculated as follows:

Therefore, the greater the solute affinity for the stationary phase, the smaller the R_f value.

Efficiency (N)

Ideally, a component moving through a chromatographic column migrates as a very narrow band. However, nonuniformity of flow, diffusion, and mass transfer effects between particles and the mobile phase produce band broadening. Chromatographic efficiency is considered to be highest when band broadening is minimized. Column efficiency is commonly expressed as N, the number of theoretical plates. For a peak forming a Gaussian curve, , where t_r is retention time and σ is the standard deviation of the curve measured as time. A practical way to calculate N is based on the measurement of peak width () at half the maximal peak height; represents 2.354 σ. Therefore,

An increase in N represents improved chromatographic efficiency and sharper peaks. Efficiency is often expressed as the number of theoretical plates per column length (N/L). The efficiency and the number of theoretical plates for a column are directly related to the column length, but usually a tradeoff of longer analysis times for longer columns occurs. Column efficiency also is expressed as the height equivalent of a theoretical plate (HETP):

In theory, a theoretical plate is equivalent to the length of a column necessary to allow one equilibration of the solute to occur between stationary and mobile phases. To increase the efficiency of a column, the number of theoretical plates is increased. In practice, this typically is accomplished by increasing the length of the column.

The flow rate has a major effect on chromatographic efficiency as described by the van Deemter equation:

where

Decreasing values for HETP represent improved efficiency, in that the number of theoretical plates for a particular column length is increased. The van Deemter equation predicts that optimization of flow rates involves a balance between peak broadening at low flow rates from diffusion and at high flow rates from increased mass transfer effects between mobile and stationary phases. It also predicts that, for a particular chromatographic system, an optimal flow rate yields the best efficiency. Empirically, the effects of flow rate on efficiency are tested by measuring peak widths at different flow rates. In practice, there usually is no interest in defining the complete van Deemter curve, but rather in identifying the highest flow rate that does not result in excessive peak broadening. The usual goal is to identify conditions for the most rapid analysis time that still yields adequate separations.

Factors that affect chromatographic efficiency are listed in Box 13-1. In practice, efficiency is improved by using (1) smaller particles, (2) nonporous particles, or (3) a coated capillary surface as the stationary phase to minimize mass transfer effects (the C constant in the van Deemter equation). This allows the use of higher mobile phase flow rates without peak broadening, resulting in faster separations. The limiting factor in reducing particle size is the increased resistance to flow and the need to operate at higher pressures.

Selectivity

Selectivity of a separation of two components reflects variable partitioning of different components between the two phases of an extraction or between the stationary and mobile phases of a chromatographic system. The selectivity factor (α) for two components A and B is the ratio of retention factors for two components:

with the later eluting component B in the numerator. If two components have the same retention times, then α = 1. Separations of components generally seek to identify conditions in which there are moderate differences in retention times of components. This is achieved through appropriate selection of mobile phase, stationary phase, and analytical conditions. A large selectivity coefficient is desirable in extractions where analysts seek to selectively extract one component. However, a large selectivity component in chromatography may result in the need to change the mobile phase or temperature during an analysis to achieve elution of both components within a practical length of time. An α of 1.1 or greater for two components usually represents adequate chromatographic separation.

Peak Capacity

Peak capacity is the theoretical maximum of the number of components that are able to be separated in a single chromatographic analysis20,51,54; it assumes a continuous distribution of peaks separated by four standard deviations. In practice, the number of components that are resolved will be lower than expected, because retention times of components are not evenly distributed. The peak capacity, therefore, serves as an upper limit for the number of components that are able to be resolved and as a performance measure for a chromatographic method. The peak capacity of high-performance liquid chromatographic (HPLC) separation usually is limited to several hundred peaks. Some gains in peak capacity are achieved by (1) increasing column length, (2) elution with solvent gradients, (3) extended run times, or (4) increasing chromatographic efficiency. However, the greatest gain in peak capacity is achieved by using two-dimensional or multidimensional chromatography, where the different dimensions usually are performed sequentially (with the second dimension representing analyses of fractions collected from the first dimension).^20,58 Peak capacities of greater than 10,000 have been achieved for two-dimensional HPLC separations, but this requires prolonged analysis time for each sample to perform multiple runs in the second dimension. Linking of chromatographic methods to other analytical methods or detectors, such as mass spectrometers, with the capacity to resolve or detect multiple components serves as an alternative approach to overcome the fundamental limitation of chromatographic peak capacity without extending analysis time. This enables the practical analysis of up to thousands of components in a single specimen, such as for metabolomic²² or proteomic analysis.⁴²