B

Figure 2.9 HPLC chromatograms of a single component showing (A) an example of "tailing" and (fl) the profile obtained when a column is overloaded.

can be used, the conversion of these to traditional concentration units can be carried out easily after a calibration curve has been constructed.

Finally, the retention time (or volume) provided by a chromatogram can be used to identify an unknown compound. For example, comparison of the retention time of the unknown to the retention time of a series of standards (i.e., known compounds) is often sufficient to identify the unknown. However, a word of caution. Since any two compounds may coelute merely by coincidence, it is often necessary to apply criteria other than retention time before feeling certain about the identity of an unknown. Sometimes recourse to other methods, such as spectral analysis, is required to obtain a more definitive identification.

Enzymes themselves are often of use in identification of an unknown. Figure 2.10 shows a compound that had been tentatively identified as inosine 5'-phosphate (IMP) on the basis of its retention time. This conclusion was subjected to further testing using the enzyme 5'-nucleotidase, with the expectation that if the compound was IMP, the enzyme would catalyze the removal of the phosphate and the formation of inosine. The chromatogram obtained following the addition of the enzyme and incubation for about 20 minutes is shown in Figure 2.10. The chromatogram now shows in addition to the IMP, which is reduced in amount, a new peak with the correct elution time expected for inosine, the amount of which increases with incubation time (Fig. 2.10). These data add credibility to the claim that the starting material was IMP.

Retention Time (min)

Retention Time (min)

Figure 2.10 The use of enzymes to identify an unknown compound. The compound tentatively identified as IMP, based on its retention time of 2 minutes (chromatogram obtained at zero reaction time), was incubated with a commercially available preparation of 5'-nucleotidase. Samples of the incubation mixture were removed and analyzed by HPLC. The chromatograms, obtained at 10 and 20 minutes of reaction time, showed a reduction in the area of the IMP peak and an increase in the area of the inosine (Ino) peak, confirming that the original peak was IMP.

Figure 2.10 The use of enzymes to identify an unknown compound. The compound tentatively identified as IMP, based on its retention time of 2 minutes (chromatogram obtained at zero reaction time), was incubated with a commercially available preparation of 5'-nucleotidase. Samples of the incubation mixture were removed and analyzed by HPLC. The chromatograms, obtained at 10 and 20 minutes of reaction time, showed a reduction in the area of the IMP peak and an increase in the area of the inosine (Ino) peak, confirming that the original peak was IMP.

2.7 SELECTION OF THE STATIONARY PHASE: SOME HELP FROM AN UNDERSTANDING OF THE PROCESS OF SEPARATION

While the selection of a stationary phase to be used in the analytical column may appear complex, the decision can be greatly simplified by considering the three basic methods of separation currently in use. These are gel filtration or size-exclusion separation, reversed-phase or hydrophobic separation, and ion-exchange separation. In general, each type of separation uses a different kind of packing material, and since each type of separation exploits a different property of the molecules, the choice of packing really comes down to which property of the molecules would be most useful in achieving the separation.

For example, to use size-exclusion chromatography, the compounds to be separated must differ in size, shape, or both, while to use solubility or charge, the compounds must differ in polarity or net charge, respectively (Table 2.2).

TABLE 2.2 Selection of the Stationary Phase

Property

Separation mode

Size, shape Solubility, polarity Charge, polarity

Gel filtration Reversed-phase Ion exchange

While deciding on a stationary phase using these as the only criteria clearly represents an oversimplification, and the reader is referjed to specific works for more details, this approach can go a long way toward facilitating the selection of the analytical column.

2.7.1 Gel Filtration Chromatography

To understand gel filtration chromatography, imagine an analytical column packed with beads as shown in Figure 2.11A If a single bead were examined

Figure 2.11 Gel filtration chromatography applied to a sample containing two compounds that differ in size larger (□) and smaller (•). (A) The two compounds are loaded onto a column packed with beads, which, when viewed at the ultrastructural level, would appear as depicted in (B). The larger compound passes between the beads, while the smaller enters the crevices of each bead (C). The chromatographic profile (D) illustrates that the larger molecule will emerge from the column in less time than the smaller.

Figure 2.11 Gel filtration chromatography applied to a sample containing two compounds that differ in size larger (□) and smaller (•). (A) The two compounds are loaded onto a column packed with beads, which, when viewed at the ultrastructural level, would appear as depicted in (B). The larger compound passes between the beads, while the smaller enters the crevices of each bead (C). The chromatographic profile (D) illustrates that the larger molecule will emerge from the column in less time than the smaller.

by scanning electron microscopy, its image might be represented by Figure 2.115. Each bead would be seen to have an irregular surface through and around which the mobile phase can enter and exit, in effect making the interior of the bead accessible to the mobile phase.

What about the sample? Imagine a sample to be composed of two types of compound, the molecules of which can be classed as "larger" and "smaller," as in Figure 2.11. Now imagine that because of these differences, the smaller molecules can follow the mobile phase as it meanders through and around the irregularities of the beads (Fig. 2.11C), while the larger cannot. Thus, the larger molecules will be excluded from taking the longer path, and as a result of this exclusion the path, or distance D, followed by the larger molecules through the column will be short, and the larger or excluded molecules will exit the column first. The volume of solvent requires for these molecules to emerge is spoken of as the included volume. The smaller molecule will follow a longer path and will emerge later. Because, in the ideal case, none of the molecules will interact with beads, the difference in times of emergence reflects the additional distance traversed by the smaller compound. Of course, altering the size of the irregularities of the beads will alter the size of the compounds excluded and therefore change the operating range of the analytical column.

2.7.2 Reversed-Phase Chromatography

We have seen that chromatography requires two phases: one solid and localized to the analytical column, the other mobile—the eluent or buffer that flows around and through the packing. The packing used by early workers was made of a material that was basically polar, while the mobile phases were nonpolar organic solvents. This arrangement of a polar stationary phase and a nonpolar mobile phase is, by virtue of tradition, referred to as normalphase liquid chromatography. Fortunately, the type of liquid chromatography performed when the situation is reversed [i.e., the analytical columns are packed with a stationary phase that is nonpolar and eluted with polar (aqueous) buffers] was not referred to as "abnormal." Instead, since the phases have been reversed, this type of chromatography came to be known as reversed phase. More recently the term "hydrophobic" has been suggested.

In describing the underlying mechanism of operation of reversed-phase chromatography, it is convenient to focus again on the compounds in the sample and their movement through the analytical column. However, unlike gel filtration, where the order of elution of the compounds is determined by their path or the distance, traveled, in reversed-phase chromatography all the compounds in the sample travel the same path. In this case it is the rate at which they move through the column packing that determines the order of elution. Thus, a molecule that moves at a slower rate is said to be retained, and its time of elution is referred to as its retention time.

To understand the operation of the reverse phase, it is useful to consider the illustration in Figure 2.12, where oil droplets (or beads) are suspended in a column filled with an aqueous buffer. The water represents the mobile phase,

SAMPLE

RETENTION TIME

Figure 2.12 Reversed-phase HPLC of a sample composed of two compounds, one polar, the other nonpolar. The column packing (stationary phase) is symbolized by spheres and labeled "Oil" and the mobile phase as wavy lines labeled "Water." The polar molecules are shown remaining in the mobile phase (water), while the nonpolar molecules "enter" the stationary (oil) phase. Finally, the chromatographic profile illustrates that in this case the polar molecule will not be retained and will emerge with a shorter retention time than the nonpolar molecule.

RETENTION TIME

Figure 2.12 Reversed-phase HPLC of a sample composed of two compounds, one polar, the other nonpolar. The column packing (stationary phase) is symbolized by spheres and labeled "Oil" and the mobile phase as wavy lines labeled "Water." The polar molecules are shown remaining in the mobile phase (water), while the nonpolar molecules "enter" the stationary (oil) phase. Finally, the chromatographic profile illustrates that in this case the polar molecule will not be retained and will emerge with a shorter retention time than the nonpolar molecule.

while the oil represents the stationary phase in the analytical column. We load the sample, in this case molecules of compounds A and B, onto the surface of the column, and the question becomes whether the compounds will remain in the water (mobile phase) as they flow through the column or whether they will enter the beads of oil. Of course, the rate of progress through the column of a compound that remains in the water is effectively the flow rate of the mobile phase. There will be no interaction, and this compound will not be retained. It will emerge soon after loading and will have a short retention time.

In contrast, by entering the oil, a compound leaves the mobile phase and interacts with the beads: Its rate of passage through the system is in effect slowed. The compound will be retained, and it will have a longer retention time than a compound that does not interact.

A great deal of time and effort has been spent in trying to predict the retention time of compounds in the reversed-phase system. While some rules have emerged and some generalizations have been made, to date the best approach remains a few trial runs.

The most useful parameters to consider when developing a feel for the operation of this type of chromatography are polarity and the related parameter solubility. Values of both parameters have been published for many of the compounds used in biological systems.

In what follows, I introduce the notion of polarity and, after differentiating it from the net charge of a molecule, use it to explain the retention time for some classes of compounds.

Polarity should not be confused with any net charge a molecule might have. For example, in some cases, highly polar molecules contain no net charge. Polarity is a result of an electrical asymmetry that is due primarily to distribution of electrons. A case in point is the water molecule (Fig. 2.13). The polarity of a molecule of water is measurable when the two positive hydrogen atoms are localized on one side of the oxygen, resulting in negative and positive sides to the molecule, and the value is often expressed as a function of its dielectric constant: the greater the dielectric constant, the more polar the

Figure 2.13 A representation of a water molecule to illustrate polarity. The positions of each of the two positively charged protons is shown as ( + ). The position of the negative charge of the oxygen atom is shown as (-). The asymmetric distribution of the positive and negative charges produces the polarity.

HOH MOLECULE

Figure 2.13 A representation of a water molecule to illustrate polarity. The positions of each of the two positively charged protons is shown as ( + ). The position of the negative charge of the oxygen atom is shown as (-). The asymmetric distribution of the positive and negative charges produces the polarity.

molecule. For example, the dielectric constant of water is about 81, while less polar molecules like alcohols have lower values. Thus, if retention time can be predicted from polarity or dielectric constant, such tables might prove useful.

Polar molecules are generally more soluble in water than nonpolar molecules, and therefore solubility values can also be useful in predicting retention times. For example, some amino acids (e.g., glycine, alanine, and others containing nonpolar side chains) are not very soluble in water; thus, on reversed-phase columns washed with only aqueous buffers, such compounds would be expected to interact with the nonpolar packing and be retained. Elution would be promoted by increasing the organic composition of the elution buffer.

Similarly, a comparison of the solubilities of some nucleobases in carbon tetrachloride and water will show that adenine is more soluble than guanine in organic solvents. Therefore, adenine will more likely enter the oil phase of the analytical column and be retained longer than guanine. Additional data obtained by measuring the distribution (or solubility) of a compound in either octanol or water show that adenosine is about 10 times more soluble than inosine in octanol. Again based on these findings, we might expect adenosine to have a longer retention time than inosine, and in fact it does.

Similarly, consider the compounds adenosine and ATP. As is well known to most biologists, ATP has greater solubility than adenosine in aqueous buffers. This knowledge, therefore, can be of value in predicting the behavior of such compounds in a reversed-phase system. Figure 2.14 shows the elution sequence of ATP and adenosine on a reversed-phased column eluted with an aqueous buffer: ATP elutes significantly before adenosine. In fact, such a short retention time suggests that ATP has great difficulty interacting with the nonpolar stationary phase.

0 6 12 ELUTION TIME Imin)

Figure 2.14 The separation of ATP and adenosine by reversed-phase HPLC. The prepacked column was Qg (/¿Bondapak), and the mobile phase was a 10 mM potassium phosphate buffer (pH 5.5) containing 20% methanol. The column was eluted isocrati-cally and monitored at 254 nra. The flow rate was 2 mL/min.

Adenosine, however, being less soluble (or more nonpolar) than ATP, will enter the stationary phase, and this is reflected in a longer retention time (Fig. 2.14). However, if a more nonpolar mobile phase were used to elute the column, the adenosine would remain longer in the mobile phase, hence its retention time would shorten. Thus, if a mobile phase was being used in which adenosine had as short a retention time as ATP, the separation of the two would be encouraged by reducing the amount of organic solvent in the mobile phase and causing the retention time of the adenosine to increase relative to that of ATP.

An interesting and useful variant of reversed-phase HPLC is called ion-paired reversed-phase HPLC. In such a system the analytical columns are packed with the same material, but a compound such as tetrabutylammonium is added to the mobile phase. The separation of ATP and adenosine on such a system is shown in Figure 2.15. A comparison of this profile to that shown for the same compounds in Figure 2.14 immediately highlights the change in the elution sequence. Whereas without ion pairing, the order is ATP followed by adenosine, with ion pairing the order is adenosine followed by ATP.

An explanation for the difference in retention times can be developed if one imagines the tetrabutylammonium compound, which is positively charged, paired with the negatively charged ATP molecule. While this pairing will, in fact, reduce the net charge, the reduction in net charge will also reduce the polarity of the ATP molecule. Since the short retention time initially was a result of the polarity, any reduction in polarity would be expected to increase retention time. Thus, coming full circle, the effect of the tetrabutylammonium salt on retention times might be explained by its effect (reduction) on polarity.

In a series of experiments designed to explore further the role of polarity in affecting retention time in reversed-phase chromatography, we developed chemical procedures for the condensation of molecules of known polarity,

ELUTION TIME (min)

ELUTION TIME (min)

Figure 2.15 Separation of ATP and ADO on ion-paired, reversed-phase HPLC. The column was Qg (/¿Bondapak), and the mobile phase was 65 mM potassium phosphate (pH 3.7) with 5% methanol and 1 mM «-tetrabutylammonium phosphate. The column was eluted isocratically, and the eluent was monitored at 254 nm.

expecting, for example, the joining two polar molecules to produce a relatively nonpolar molecule. In our first experiment we coupled the very polar nucleoside monophosphate AMP to lysine, an amino acid with a very polar side chain. The behavior of the two starting compounds in reversed-phase HPLC is shown in Figure 2.16/1. Both have relatively short retention times, consistent with their polar character. However, when the retention time of the conjugate was determined, it was found to be longer than that of either of the starting compounds (Fig. 2.16A). Thus, the combination of two polar compounds can produce a compound more nonpolar than either of the parent compounds.

Similar experiments were undertaken joining AMP to a dipeptide hippuryl-lysine. This particular dipeptide was used because a comparison of the retention times of lysine and hippuryllysine revealed that the addition of the hip-puric acid to the lysine reduced the polarity of the latter. However, a determination of the retention times of the dipeptide and AMP on a reversed-phase column (Fig. 2.165) reveals both to be polar. Nevertheless, their conjugate has a longer retention time than either of the starting materials. Note, however, that the decrease in polarity of this conjugate is very much less than what was observed following the summation of the AMP aiid lysine (Fig. 2.16/1).

Finally, the AMP was coupled to the tetrapeptide tuftsin, which has the amino acid sequence Thr-Lys-Pro-Arg. Based on its extremely long retention time on a reversed-phase column, the tuftsin can be considered a nonpolar molecule, a conclusion supported by its rather low solubility in aqueous systems and the requirement for 40% methanol to elute it from the column. When AMP is condensed onto the tuftsin, usually a single AMP per molecule of tuftsin, the polarity of the tuftsin is significantly decreased, as indicated by the decrease in the retention time of the conjugate. As shown in Figure 2.16C, the conjugate has a retention time much closer to that of AMP than to that of tuftsin. This finding suggests that the addition of the polar AMP to the nonpolar tuftsin decreases the overall electrical asymmetry but does not eliminate it completely. Thus, while the combination of two polar molecules can produce a nonpolar molecule, the combination of a polar with the nonpolar molecule can produce a molecule more polar than its nonpolar parent.

It should be noted that reversed-phase HPLC has been used to deduce polarity as, for example, in a study of cAMP and its analogs, as well as to predict partition coefficients and lipid solubility.

2.7.3 Ion-Exchange Chromatography

In ion-exchange chromatography, as in reversed-phase HPLC, the rate at which a molecule moves through the analytical column and its interaction with the packing determine the order of elution of given compounds. In this case, both the number and the magnitude of the charge influence interactions. The principles and method of operation in ion-exchange HPLC are similar to those of the more conventional ion-exchange systems.

Retention time (min)

Retention time (min)

Figure 2.16 Effects of polarity on retention time. HPLC chromatography carried out on reversed-phase Qg (/xBondapak) column. (A) Chromatographic profiles of AMP, lysine, and the lysyl-AMP conjugate obtained using a mobile phase of 65 m M potassium phosphate (pH 3.6) and 2% acetonitrile. (B ) Profiles of AMP, hippuryllysine, and the hippuryllysyl-AMP conjugate eluted with a mobile phase of 10 mM potassium acetate (pH 7.2) containing 1% acetonitrile. (C) Chromatograms obtained with AMP, tuftsin, and tuftsin-AMP. Compounds were eluted with a mobile phase of 65 mM potassium phosphate (pH 3.6) and 2% acetonitrile for AMP and tuftsin-AMP. Tuftsin was eluted by 20% acetonitrile. Detection was at 230 nm for lysine and 254 nm for all others.

Retention time (min)

Figure 2.16 Effects of polarity on retention time. HPLC chromatography carried out on reversed-phase Qg (/xBondapak) column. (A) Chromatographic profiles of AMP, lysine, and the lysyl-AMP conjugate obtained using a mobile phase of 65 m M potassium phosphate (pH 3.6) and 2% acetonitrile. (B ) Profiles of AMP, hippuryllysine, and the hippuryllysyl-AMP conjugate eluted with a mobile phase of 10 mM potassium acetate (pH 7.2) containing 1% acetonitrile. (C) Chromatograms obtained with AMP, tuftsin, and tuftsin-AMP. Compounds were eluted with a mobile phase of 65 mM potassium phosphate (pH 3.6) and 2% acetonitrile for AMP and tuftsin-AMP. Tuftsin was eluted by 20% acetonitrile. Detection was at 230 nm for lysine and 254 nm for all others.

In general, the support (stationary) phase carries either a positive or a negative charge. During equilibration of the column with the eluent, a counter-ion is introduced. The molecules to be separated must also be charged, and when the sample is loaded, they bind to the fixed charges of the column packing and displace the counterion. Elution of the bound molecules is brought about by a second counterion, which is usually introduced as salt onto the packing by adding it to the elution buffer. The ability of the counterions (salts) to displace bound molecules relies on the difference in their affinities for the fixed charges of the stationary phase.

The interaction between the fixed charges of the stationary phase and the compounds adenosine, AMP, ADP, and ATP with zero, one, two, and three charges, respectively, is shown schematically in Figure 2.17. In anionic-

VOLUME

Figure 2.17 Representation of operation of ion-exchange chromatography. Top: Column functional groups, or fixed charges, are represented by serrated-edge circles carrying a positive charge, shown "fixed" to a lattice. The compounds to be fractionated are ATP, ADP, AMP, and Ado. The first three are shown bound to the fixed charges, while the Ado is shown unbound. The introduction of the counterion, a chloride, is shown displacing the bound molecules from the fixed charges. Bottom: The order of elution as a function of the NaCl molarity, which is represented by the dashed diagonal line. The relative elution position (volume) of each of the four compounds is shown.

VOLUME

Figure 2.17 Representation of operation of ion-exchange chromatography. Top: Column functional groups, or fixed charges, are represented by serrated-edge circles carrying a positive charge, shown "fixed" to a lattice. The compounds to be fractionated are ATP, ADP, AMP, and Ado. The first three are shown bound to the fixed charges, while the Ado is shown unbound. The introduction of the counterion, a chloride, is shown displacing the bound molecules from the fixed charges. Bottom: The order of elution as a function of the NaCl molarity, which is represented by the dashed diagonal line. The relative elution position (volume) of each of the four compounds is shown.

exchange chromatography, adenosine, with no charge, is not retained; it will be eluted in the absence of the addition of any salt. Thus, as represented in Figure 2.17, adenosine (Ado) will have a short retention time.

Increasing concentrations of chloride will be required to displace and elute in series AMP, ADP, and even higher concentrations of ATP (Fig. 2.17). Thus the elution order will be Ado, AMP, ADP, and ATP, with Ado having the shortest retention volume of the four (see Fig. 2.17). Such an elution order is consistent with the explanation that increasing the number of charges of a molecule increases its interaction with the stationary phase packing, thereby reducing its flow rate through the analytical column.

A consideration of the effectiveness of the different salt cations (or anions) in displacing or exchanging bound molecules requires a discussion of the magnitude of the charge. Thus, molecules with one charge are not all equal when it comes to interacting with the fixed charges of the column packing. As a first approximation, the strength of the interaction may be considered in terms of the number of water molecules between or surrounding the salt ion; this is its hydrated radius.

For example, as represented in Fig. 2.18, cesium has only a few water molecules surrounding it compared to the lithium ion, which has many more (Fig. 2.18). One might think of these water molecules as a shield, with their elimination required for any interaction to take place between the ion and the packing. Clearly, it takes less energy to eliminate one molecule than several, and therefore it is not surprising that cesium is more effective than lithium at displacing molecules such as ATP bound to ion exchangers (see Fig. 2.18). This effectiveness is seen operationally in terms of the concentration of the counterion required to elute the bound sample. It is also not surprising that affinities can be affected by modifications that alter the water content of the system—for example, by increasing salt concentrations, temperature, or the organic solvent content of the mobile phase. However, on an ion-exchange HPLC column, the AMP is eluted at a lower salt concentration than cAMP, as illustrated in Figure 2.19. This difference in affinity might be explained by the model already described in which relative affinity is a function of the

Figure 2.18 Effect of radius of hydration on distance between counterion and fixed charge of ion-exchange stationary phase. Cesium, with smaller radius of hydration, is shown with one water molecule (small circle) between it and fixed charge of the bead. Lithium is shown with three water molecules.

Figure 2.19 Separation of several nucleosides on ion-exchange HPLC was carried out on an ion-exchange column (AX-100) eluted isocratically with a mobile phase of 0.1 M sodium phosphate buffer (pH 7.3) containing 0.8 M sodium acetate. The column was monitored at 254 nm. A standard solution containing approximately 2 nmol each of adenosine, AMP, cAMP, ADP, and ATP was loaded onto the column.

distance between the mobile ion and the fixed ion. In this model, AMP, with less affinity, would have a greater distance between it and the fixed charge on the bead than cAMP would have. In both cases, as illustrated schematically in Figure 2.20A, the space would be occupied by water molecules. The net effect of this difference would be a lower concentration of the chloride required to displace the AMP, which would be eluted before cAMP, as illustrated in Figure 2.20B.

Figure 2.19 Separation of several nucleosides on ion-exchange HPLC was carried out on an ion-exchange column (AX-100) eluted isocratically with a mobile phase of 0.1 M sodium phosphate buffer (pH 7.3) containing 0.8 M sodium acetate. The column was monitored at 254 nm. A standard solution containing approximately 2 nmol each of adenosine, AMP, cAMP, ADP, and ATP was loaded onto the column.

exchange beads and the cAMP and AMP molecules.

2.8 COMPOSITION AND PREPARATION OF THE MOBILE PHASE

A mobile phase containing salt and organic modifiers is commonly used to elute samples from a reversed-phase column. The salt is added to suppress ionic effects that could alter separation. However, with samples from enzymatic reactions, which are often at pH values different from that of the mobile phase, a buffer should be added to the mobile phase, as well. The buffering capacity of the mobile phase should be in excess of that in the incubation mixture. This excess will ensure that when the sample is injected, its pH will equilibrate to that of the mobile phase. These buffers should be made with distilled, deionized water that has been degassed to remove any trapped air. Degassing prevents bubble formation, which otherwise occurs in the pump head, particularly at high pressures. Bubbles are one of the major causes of variations in pump pressure, which, in turn, can produce artifacts in the chromatographic profile. Degassing, accomplished by vacuum aspiration, should be carried out with constant stirring until only a few bubbles are formed. Usually about 20 min/L is adequate.

Phosphate in concentration ranges of 10 to 100 mM can be used not only for buffering but also for ion suppression. However, some thought should be given to later uses contemplated for the compounds purified by the HPLC. For example, if after purification the component will be examined by phosphorus NMR, a mobile phase containing phosphate obviously should not be used. In addition, the use of phosphate in ion-exchange mobile phases can lead to high background absorbance in the UV range as a consequence of impurities in the phosphate buffers. Although methods have been described to reduce this background (see General References), where possible the phosphate should be eliminated. The pH can always be adjusted with KOH. With such bases, however, the choice of cation (e.g., K or Na), should be made with the composition of the sample buffer in mind. For example, if the buffer contains sodium dodecyl sulfate (SDS), potassium, which will precipitate SDS, should be avoided. Also, since halides such as chloride can cause corrosion of stainless steel tubing, they should be avoided. Finally, if the purified components are to be concentrated by an evaporation procedure, a buffer with volatile components should be chosen.

To reduce retention time of nucleotides on reversed-phase columns, organic modifiers such as methanol or acetonitrile may be added to the buffer. Acetoni-trile is usually more effective than methanol in the sense that less is required to elute a given nucleotide with a specific retention time.

Once the buffer (mobile phase) of the appropriate composition has been made and the pH adjusted, it should be filtered through a 0.45 (im filter to remove particles that may clog the column head. Following filtration, the buffer may be used for about 2 to 3 days if stored at room temperature although the pH should be checked and precautions taken to keep the organic modifer from evaporating during storage.

At the conclusion of each work day it is advisable to wash the salt buffer thoroughly from the both the pump heads and the column. A 0.02% sodium azide solution prepared with degassed water should be used to wash the system free of salts. About 15 minutes of washing time is adequate with a flow rate of 2 mL/min. The sodium azide is used to control bacterial growth. A reversed-phase column should then be washed with and stored in a methanol-water solution (80:20).

Washing removes material from the guard column and the top of the analytical column, and when this material passes through the detector, it often causes changes in optical density. Therefore, the recorder should be left on during the washing to ensure complete removal of this debris. The recorder will return to a baseline reading when the washing is complete.

Note that water and methanol differ in viscosity. Thus the change in viscosity of the solutions accompanying a switch from one solvent to the other will produce an increase in back pressure that may be significant in magnitude; such changes are not worrisome, however. Also, a sodium azide solution moving through a detector set at 254 nm will usually produce an increase in optical density. Again these are normal changes and should not cause concern.

2.9 COLUMN MAINTENANCE

When reversed-phase columns are used for the analysis of enzymatic reactions, many of the components of the reaction may become bound to the packing material. As a result, the debris may alter the retention time, chromatographic profiles, or both of subsequently injected molecules. Types of column malfunction include peak splitting or the appearance of a shoulder; loss of baseline resolution, broadening of peaks, particularly at their base, or both; and an increase in back pressure. To some extent, all these symptoms may be traced to material that adhered to the column and was not removed during the methanol wash.

If divalent metals are suspected as the cause of the peak splitting, washing the column with 100 mL of 10 mM EDTA in 10 mM phosphate at pH 5.5 may help eliminate the problem by removing the metal.

Many components that bind can be eluted by changing the pH of the mobile phase. Thus, a wash consisting of 200 mL total volume, at 2 mL/min, of 100 mM phosphate solution ranging in pH from 2 to 8 is frequently useful. Finally, if an increase in back pressure is suspected to be a result of contamination from bound protein, washing the column with at least 100 mL of 6 M urea in 20 mM phosphate (pH 7.8) may eliminate the problem. Again, the return of back pressure to normal values can be taken as a sign of success of any one of these steps.

We have found urea to be a poor wash solution on columns packed with reversed-phase packings. Dimethyl sulfoxide (DMSO) has been useful in some cases.

In addition, a gradient progressing from 100% methanol through a series of less polar, more organic solvents such as carbon tetrachloride will serve to remove other reversibly bound contaminants. A reverse gradient should be used to reequilibrate the column to standard conditions.

Note that following any of the maintenance procedures listed above, re-equilibrated the column must be to the original mobile phase, probably involving more time than would be needed for the normal change from water to the methanol solution routinely used to prepare the column for overnight storage.

As an illustration, consider the problem of the contamination of a reversed-phase column with a very sticky dextran sulfate material that had been added as an activator for an enzyme reaction. The compounds AMP, ADP, and ATP were being separated using a mobile phase containing phosphate buffer, acetonitrile, and tetrabutylammonium ion. The separation usually obtained is shown in Figure 2.21 A. However, in the presence of dextran sulfate the separation was less than adequate (Fig. 2.21.8). The column was regenerated by first washing with 6 M urea. Analysis of the sample produced is shown in Figure 2.21 C. Some improvement in the separation is evident. Next the column was washed with toluene. The result, (Fig. 2.21D) shows complete restoration of the separation capabilities of the packing. Additional details may be obtained by consulting the General References at the end of this chapter.

2.10 MONITORING COLUMN PERFORMANCE

In general, new columns should be calibrated in the laboratory in which they will be used, and a standard mobile phase and a standard series of compounds should be available for this purpose. The resolution obtained under the standard conditions of the laboratory at the start of the useful life of the column should be recorded, together with the date of the analysis. All should be part of the record for that column. At the first sign of problems with this column, its performance should be checked against these records, using the same mobile phase and standards.

2.11 SUMMARY AND CONCLUSIONS

Chromatography involves the separation of classes or groups of molecules. Two phases are usually required: one is stationary or solid, and the other mobile—eluent or buffer.

If the solid phase is in the form of particles or beads, it is usually packed into a tube or column and the buffer or mobile phase is pulled through the packing by gravity or forced through with a pump.

The time required for a given compound to emerge from the column is a function of the packing material and the interactions between the compound and the packing. Transition time is affected by the distance the compound travels or the rate at which it travels a fixed distance.

AFTER WASH WITH 6M UREA

AFTER WASH WITH TOLUENE

Figure 2.21 The effect of washing procedures on removal of debris (dextran sulfate) as measured by separation of nucleosides. Separations were carried out on a reversed-phase (Cig) column with a mobile phase of 65 mM potassium phosphate (pH 3.6), 2% acetonitrile, and 1 mM tetra-n-butylammonium phosphate. The flow rate was 2 mL/ min; the column was eluted isocratically and monitored at 254 nm. (A) Separation routinely achieved with AMP, ADP, and ATP. (B), Separation observed after clogging the column with 10 mM dextran sulfate. (C) The separation observed after washing the column with 6 M urea. (D) The separation obtained after washing the column with toluene.

Other things being equal, resolution is enhanced by using smaller particles of the packing material. However, smaller particles result in tighter packing, which in turn requires higher pressures to push through the solvents. The combination of small particles with better pumps led to improved performance and the method called high performance liquid chromatography.

The basic equipment required for an HPLC system includes a solvent reservoir, a pump, an injector, an analytical column, a detector, and a recorder. The analysis of the sample is displayed as a chromatogram, with detector deflection presented usually as a function of time after loading the sample. By virtue of the shape of the curves, the distance between them, and their area, it is possible to determine whether the volume of the sample is too large, as well as the number of different compounds present and the amount of each compound occurring in the sample. From an understanding of the process of separation, it is possible to select the appropriate stationary phase. Separation by gel filtration requires compounds of different sizes and shapes, while reversed-phase HPLC will separate molecules that have different polarities. In contrast, ion-exchange HPLC separates molecules with different charges.

The mobile phase contains salts and organic modifiers. A buffer is also required with enzymatic reactions to ensure a constant pH during the separation step.

Column maintenance will often require washing the column with chelators, dénaturants, organics, or salt solutions of high concentration, all designed to remove column-bound debris that is not removed by routine washing.

GENERAL REFERENCES

General references for HPLC

Brown PR (1973) High Pressure Liquid Chromatography: Biochemical and Biomedical Applications. Academic Press, New York.

Hearn MTW, Ed. (1985) Ion-Pair Chromatography: Theory and Biological and Pharmaceutical Applications. Dekker, New York.

Henschen A, Hupe K, Lottspeich F, Voelter W (1985) High Performance Liquid Chromatography in Biochemistry, VCH Publishers, Deerfield Beach, FL.

Krstulovic AM, Brown PR (1982) Reversed-Phase High-Performance Liquid Chromatography: Theory, Practice and Biomedical Application. Wiley-Interscience, New York.

Regnier FE (1983) Science 222:245.

Snyder LR, Kirkland JJ (1979) Introduction to Modern Liquid Chromatography. Wiley, New York.

Polarity and solubility

Cohn EJ, Edsall JT (1939) Proteins, Amino Acids and Peptides. Hafner, New York. Cullis PM, Wolfenden R (1981) Biochemistry 20:3024.

Greenstein JP, Winitz M (1961). Chemistry of the Amino Acids, Vol. 1. Wiley, New York.

Hansch C, Leo AJ (1979) Substituent Constants for Correlations Analysis in Chemistry and Biology. Wiley, New York. Kolassa N, Pfleger K, Rummel W (1970) Eur J Pharm 9:265. Nahum A, and Horvath C (1980) J Chromatog. 192:315. Plaut, GWE, Kuby SA, Lardy HA (1950) J Biol Chem 184:243.

Reversed-phase chromatography

Hacky JE, Young AM (1984) J Liquid Chromatog, 7:675.

Hammers WE, Meurs GJ, DeLigny CL (1982) J Chromatog, 247:1.

Hancock WS, Ed. (1984) Handbook ofHPLC for Separation of Amino Acids, Peptides and Proteins, Vols 1 and II. CRC Press, Boca Raton, FL.

Krstulovic AM, Brown PR (1982) Reversed-Phase High-Performance Liquid Chromatography: Theory, Practice and Biomedical Applications, Wiley-Interscience, New York.

Perrone P, Brown PR (1985) Ion-pair chromatography of nucleic acid derivatives. In Ion-Pair Chromatography, MTW Hearn, Ed. Dekker, New York.

Rossomando EF, Hadjimichael J (1986) Int J Biochem 18:481.

Care and maintenance of columns

Runser DJ (1981) Maintaining and Trouble Shooting HPLC Systems. Wiley, New York.

Detectors

Henderson RJ, Jr, Griffin CA (1984) J Chromotogr 298:231. Ion-exchange chromatography

Jahngen JH, Rossomando EF (1983) Anal Biochem 130:406.

Regnier F (1984) High-performance ion-exchange chromatography, In Methods in Enzymology, Vol. 104, WB Jakoby, Ed., p. 170. Academic Press, Orlando FL.

Preparation of mobile phase

Karkas JD, Germershauser J, Liou R (1981) J Chromatogr 214:267.

Plunkett W, Hug V, Keating MJ, Chubb S (1980) Cancer Res 40:588.

Pumps: operation and troubleshooting

Dolan JW, Berry VV (1983) LC Mag 1:470

CHAPTER 3

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