How can we tell if a liquid chromatography method is working properly? How can a compendial method be adjusted to speed up the run time?

 A liquid chromatography (LC) method from the 

) for the assay of a drug product was set up in the laboratory of one of the authors (F.O.). Although all of the details of the method cannot be shared because of its proprietary nature, the key elements follow. A 250 mm × 4.6 mm L1 column packed with 5-μm particles was used with a mobile phase of 50:50 acetonitrile–water and a flow rate of 0.6 mL/min. A refractive-index detector was used, with both the detector and column maintained at a temperature of 55 °C. System suitability requirements included a minimum resolution between the active ingredient and a related compound, a relative standard deviation (RSD) of <2% for the peak area of replicate injections of the active ingredient, and a 

 tailing factor (TF) of <2. When the method was run, the resolution requirements were met; TF = 1.13 was observed, and RSD = 0.2% was determined (see Table I). It can be seen that the system suitability requirements were passed, and because the method was operated according to the monograph specifications, no further testing was required before placing the method into routine use.

Table I: System suitability test results			

Although the method proved suitable for use, let's examine it a little more closely to see how it fits into our expectations for a "good" isocratic method. One system suitability parameter that often is included in methods such as this is a minimum column plate number requirement. The plate number can be calculated easily from a chromatogram, such as that of Figure 1. The plate number, 

 is the peak width at half its height, both in the same units; these values are included in the data system report generated for each chromatogram. For the last peak listed in Table I, 

 = 12,300 (as usual, numbers are rounded for simplicity, so if you try to repeat the calculations presented here, you may get slightly different values). 

Figure 1: Chromatogram for system suitability test injection 1.			

What does the calculated plate number tell us? As has been described in previous 

 discussions, for real samples operating under real conditions, we can estimate a reasonable plate number for any column as 

 is the column length (in millimeters) and 

 is the particle diameter (in micrometers). The present column is a 250 mm × 4.6 mm column packed with 5-μm particles, so the predicted value of 

 is within about 20% of this predicted value, the column is in good condition. So, although the column was new, the plate number is at the low end of its expected performance range. In the present case, we'll see in a moment that the retention factor is quite small, which makes the peaks more susceptible to extracolumn band broadening because of injection problems or extracolumn volume. We suspect that this is the reason for the lower-than-expected plate number in the present example. 

Another parameter that can be used to evaluate the quality of an LC method is the retention factor, 

. As a general rule, for the best isocratic methods, we would like to have 2 < 

 < 20 usually is acceptable (see the discussion associated with Figure 1 in reference 1 for more information on this). We calculate 

 is the column dead time, usually identified as the first peak in the chromatogram or "solvent front." We can see in Figure 1 that the retention time of the first peak in the chromatogram is 4.0 min. It is a good idea to double-check that this is where the 

 peak is expected. For 4.6-mm i.d. columns we can estimate 

 is the length of the column (in millimeters) and 

 is in milliliters. Therefore, in the present case 

 = 2.5 mL/0.6 mL/min = 4.2 min. The estimate should be within ≈ 10% of the observed value, so the observed 

 = (6.2 – 4.0)/4.0 = 0.6. This is much less than even the minimum desired 

 = 1. Does this mean that the method is bad or that a bad method was approved and included in the 

Let's take a look at the column that is used for the method, which 

 classifies as L1. L1 columns are defined as "octadecylsilane (C18) chemically bonded to porous silica . . . micro-particles" (p. 982 in reference 2). This reflects the belief many years ago that all C18 columns are the same or very similar. Of course, today we recognize that this is not true, and it is not difficult to find two C18 columns that differ by a factor of two in the retention of any particular compound. Although it is permissible to use any L1 column with the present method, it would be much more appropriate to use the same brand of column that was used when the method was developed for the 

 monograph. Fortunately, the USP website (3) contains a list of which columns are associated with monograph methods. When we consulted the database, we found that the recommended column was a 250 mm × 4.6 mm Inertsil ODS-3 column packed with 5-μm particles (GL Sciences). From data we have available (for example, Table 7.2 in reference 4), we observed that the column that was used to generate the data of Figure 1 and Table I gives 

 values of about 60% of the Inertsil column. If this same retention ratio is true for the analyte of interest, the 

 value would increase from ~0.6 to ~1.0, which would move it into a more reasonable range. This means that the lower-than-desired retention factor observed with the method could be attributed to the choice of a different L1 column than the one used for the original method, and not a poorly designed method. 

 methods is that they do not need to be validated if the method can be set up and shown to pass the system suitability requirements. This is an obvious savings of time and money, so it is understandable that most users of 

 methods are reluctant to make any changes. However, in Chapter 621 of the 

 (2), there is a list of adjustments that can be made to existing methods that do not require revalidation, providing, of course, that system suitability still passes. If we consider the allowable changes and the simplicity of the present method, it is likely that we can shorten the run time without compromising the results. 

First, let's consider the column size. The method designates a 250 mm × 4.6 mm column. This suggests to us that the method is quite old — 250-mm columns rarely are used with methods today. The 

 allows a change in column length of ±70%, so a change to a 150-mm-long column would be a reduction of only 40% in length, well within the allowance. This adjustment would reduce the overall run time from 10 min to 6 min and the retention of the analyte from 6.2 min to 3.7 min. Because resolution changes only with the square root of the plate number (and thus square root of the column length change), the shorter column is unlikely to compromise resolution in the current method, where the analyte of interest is well resolved from the impurity. 

 also allows adjustment of the flow rate by as much as ±50%, meaning that the 0.6-mL/min flow rate could be increased to 0.9 mL/min. A more careful reading of the actual method states that the flow rate is "about 0.6 mL per minute," so it would be acceptable to run the method at 1 mL/min with the adjustment. This would further reduce the run time from 6 min to 3.6 min and the retention time from 3.7 min to 2.3 min. Again, resolution is unlikely to be affected by the flow rate change because for 5-μm and smaller particle sizes a change of flow rate by a factor of two will result in only a small change in the plate number for most methods. 

Two other minor changes are suggested by an examination of the chromatogram of Figure 1. First, the end-integration marker (vertical bar below baseline at ~7 min) appears to be a bit later than is necessary. This could be adjusted to force the peak to end closer to where the peak returns to baseline — for example, at 6.5 min. Second, the negative peak between 8 and 9 min appears to be the last peak in the chromatogram. After this peak has been eluted, there is no need to continue the run, so the run time could be reduced from 10 min to 9 min. This 10% reduction in run time would also apply to the corresponding run-time changes for a shorter column or higher flow rate, or both. 

If the changes in column length or flow rate were made, care should be taken to make sure that system suitability still passed. Special attention should be paid to the minimum resolution requirements to separate the active ingredient from the related compound. Providing that these adjustments still result in acceptable method performance, the changes should be documented, along with a comparison of the method performance (for example, precision, accuracy, resolution, and tailing factor) before and after the change. This documentation will be useful if, during a regulatory inspection, the validity of the changes is questioned. 

We observed that the LC method could be set up and would pass the system suitability requirements. Although technically this is all that needs to be demonstrated to permit use of a compendial method, it is a good idea to check some performance factors that are not required, but can be used to help assure us that the method is operating properly. 

The column plate number is easy to determine, and often it can be calculated automatically by the chromatographic data system. We used an estimate of 

 to determine that the observed plate number is reasonable; so the column is in good condition. 

We also calculated the retention factor to see if the retention time of the analyte of interest was in a range that is desired for a robust LC method. We found that, although the retention time seemed to be large, the 

 value was less than desired. In some methods, this would raise a red flag, but there are several factors that reduce our concern about the low 

 value in the present application. First, the chromatogram is very simple ­— there is only one peak that we are interested in, and as long as it is separated adequately from a particular related compound, there is little concern. Second, there is very little disturbance in the baseline at 

, suggesting that there is little likelihood that unretained materials will interfere with the peak of interest. Third, the repeatability of peak areas and retention times was excellent, which tells us that even though the 

 value is less than desired, the method performs acceptably for the present application. It also is likely that the low 

 value contributes to the slightly lower than expected plate number for the column. The smaller the 

 value, the more likely that extracolumn effects, such as plumbing and injection conditions, will cause peak broadening. Even so, the plate number is acceptable and peak tailing is minimal, so there is little reason for concern. 

This method is a good example of a case where the perfect method is not necessary to get acceptable results. Any changes to the method other than the change in column length and flow rate discussed here would be likely to require some level of method revalidation. Validation is a time-consuming and expensive process, so any additional reduction in method run time would be offset by these extra costs. This is a good example of the maxim, "better is the enemy of good enough." 

United States Pharmacopeia 34–National Formulary 29

 (United States Pharmacopeial Convention, Rockville, MD, 2011).			

http://www.uspchromcolumns.com/chrom/display?cmd=index

(4) L.R. Snyder, J.W. Dolan, D.H. Marchand, and P.W. Carr, 

is a senior HPLC analyst at Profarma in Tirane, Albania. 

"LC Troubleshooting" Editor John Dolan has been writing "LC Troubleshooting" for LCGC for more than 25 years. One of the industry's most respected professionals, John is currently the Vice President of and a principal instructor for LC Resources, Walnut Creek, California. He is also a member of LCGC's editorial advisory board. Direct correspondence about this column via e-mail to 

Articles

LC Troubleshooting

How can we tell if an LC method is working properly?

Evaluation of an LC Method

Aliquid chromatography (LC) method from the 

 = 2.5 mL/0.6 mL/min = 4.2 min. The estimate should be within ˜≈ 10% of the observed value, so the observed 

In part I of this series, we discussed various advantages and practical considerations of performing elevated-temperature separations (1). Here in part II, we continue our discussion of this topic, exploring some basic thermodynamic relationships, including the derivation and use of the van 't Hoff equation. 

We believe that an improved understanding of basic thermodynamics, including the van 't Hoff relationship, will better help chromatographers perform elevated-temperature liquid chromatography (LC). Therefore, here in part II of this three-part series, we introduce some of the basic thermodynamics of liquid chromatography (LC), leading to the van 't Hoff equation. In part III, we will present a more detailed discussion of the van 't Hoff equation, exploring its usefulness and relevance using various examples from the literature. 

) of an analyte in a chromatographic separation is the change in enthalpy that occurs when it moves from the mobile phase to the stationary phase. Enthalpy is defined as 

 is the volume. Thus, for the nearly constant pressures and volumes that are present in analyte transfer in LC, Δ

. In addition, at constant pressure, which is also closely approximated in LC, Δ

 is the heat transferred in the process. Thus, the enthalpy of transfer provides a measure of the difference between the bonding interactions (for example, London dispersion, dipole-induced dipole, dipole-dipole, and hydrogen bonding) of the analyte with the mobile phase compared to its interactions with the stationary phase. The presence of retention in a chromatographic separation implies that the analyte has at least some affinity for the stationary phase. This further implies that, in general, the enthalpy of transfer of the analyte from the mobile phase to the stationary phase is favorable, which is signified by a negative Δ

) of the analyte is the change in entropy that occurs when the analyte moves from the mobile phase to the stationary phase. Entropy can be viewed as a measure of the change in the randomness, mobility, or ability to diffuse of the analyte or, explained more formally, as the change in the number of available states of a system. In general, as an analyte moves from the mobile to the stationary phase its mobility is restricted, which causes the entropy of transfer to be unfavorable. This is signified by a negative Δ

 value. (Unlike changes in enthalpy, positive values of Δ

The explanation presented above for the change in enthalpy or entropy in analyte transfer from the mobile to the stationary phase is admittedly simplistic. Nevertheless, in many cases it will capture a significant fraction of the thermodynamics of the problem, and at the very least provide a starting point for thinking about the problem. Of course, to fully understand these parameters, one would need to consider all interactions in the system, including those of the solvent with itself, and any that may come as a result of the analyte replacing solvent molecules that may be adsorbed on the stationary phase. 

) predicts whether an overall process is favored (indicated by a negative value of Δ

) or disfavored (signified by a positive value of Δ

 has a contribution from the change in enthalpy, Δ

, a contribution from the change in entropy, Δ

, and it depends on the absolute temperate in kelvins, 

 in a typical reversed-phase separation (negative and favored for Δ

) interact with temperature in the Gibbs equation (equation 1). This relationship helps us understand the typically observed phenomenon of the retention factor, 

, decreasing with temperature. As illustrated in Figure 1, at lower temperatures Δ

 is negative — lower temperatures favor retention. However, as the temperature increases, the "– 

 to become less negative (again, because Δ

 is negative in our case) and at some point, Δ

 becomes positive. That is, as temperature increases, adsorption (retention) is increasingly disfavored.  

Figure 1: ÎG0 plotted as a function of temperature in ÎG0 = ÎH0 â TÎS0 for ÎH0 negative (favored) and constant, and also ÎS0 negative (unfavored) and constant, which are good approximations for a typical reversed-phase separation. At lower temperatures, ÎG0 is negative (favored), which leads to higher values of the equilibrium constant, K, and larger values of the retention factor, k. At higher temperatures, ÎG0 is positive (unfavored), which leads to lower values of K and smaller values of k.			

This same analysis provides an even more straightforward explanation for retention in gas chromatography (GC), in which decreases in retention occur with increases in temperature. Δ

 will definitely be negative (favored) for analyte adsorption or partitioning into the stationary phase from the mobile phase in GC because one is essentially comparing no chemical bonding interactions in the gas phase to, at a minimum, London dispersion forces upon adsorption. Furthermore, there will be a substantial decrease in the entropy of the system upon adsorption or partitioning because of the high entropies of gas-phase species compared to the lower entropies of the same species in a condensed phase. 

Therefore, a relatively simple thermodynamic analysis helps us understand why retention in some of the most important forms of chromatography will, in general, decrease with temperature. Of course, there are some underlying assumptions here, one of which is that Δ

 are independent of temperature, and another is that the stationary phase and mobile phase remain unchanged with temperature. These assumptions, which in many cases are reasonable, are discussed in greater detail below and in part III of this series.  

, for the transfer of an analyte from the mobile phase to the stationary phase is  

 is the concentration of the analyte in the stationary phase and [A]

 is the concentration of the analyte in the mobile phase. 

 is not easily measured in a chromatographic separation. However, using the definition of concentration as moles solute per unit volume, 

 can be expressed in terms of the retention factor, 

 (see below), which is an easily measured parameter, as follows:  

 are the number of moles of the analyte A in the stationary phase and mobile phase, respectively, and 

 are the volumes of the stationary and mobile phases, respectively. 

, is the ratio of the time the analyte spends in the stationary phase divided by the time it spends in the mobile phase:  

 is the retention time of the analyte and 

 is the mobile phase hold-up time or dead time. (A variety of approaches can be taken to determine the thermodynamic dead time of a column, which include the disturbance peak [unretained components in the analyte mixture] or the use of isotopically labeled eluent components [2,3]).  

Basic chromatographic theory also teaches us that during a separation, 

 equals the number of moles of analyte in the stationary phase divided by the number of moles of analyte in the mobile phase:  

The ratio of the volume of the mobile phase to the volume of the stationary phase is known as the 

So equations 3, 5, and 6 can be combined to obtain the following relationship for 

. As noted, this equation is important because it relates 

 to something we can measure easily — the retention factor, 

How Gibbs Free Energy Relates to the Equilibrium Constant

Through equation 8, we see that a change in Δ

 has a direct impact on the equilibrium constant, and therefore retention factor: If Δ

The van 't Hoff equation is closely tied to the thermodynamics of the transfer of the analyte from the mobile phase to the stationary phase. To obtain the van 't Hoff equation, we first combine equations 1 and 8:  

Next, we substitute equation 7 into equation 9:  

Finally, with a little more algebra we obtain the van 't Hoff equation, as it is typically seen in LC:  

Now, we can compare equation 12 to the basic equation for a straight line:  

, and then take the slope and intercept of this line, we can directly obtain the enthalpy of transfer of the analyte from the mobile phase to the stationary phase as Δ

, and the entropy of transfer from the intercept of the line, provided we know the phase ratio, β, where we are assuming the plot produces a straight line, which it often does. 

So, in a straightforward way we have derived the van 't Hoff equation, which allows us to gather information about two key thermodynamic parameters through some direct chromatographic measurements. That is, we simply measure the retention factor of an analyte in two or more different chromatograms at different temperatures under otherwise identical conditions, and then plot and analyze ln 

Figure 2 is a mock (synthesized data) example of a van 't Hoff plot for two different hypothetical analytes. The slopes of the two fit lines are different in that the steepest slope (analyte 2) corresponds to the largest value of Δ

 comes from the curve with the largest intercept (analyte 1). Note that because 1/

-axis, temperature increases from right to left and not left to right, which is typical. Consistent with the plot of Gibbs free energy (Figure 1), the  van 't Hoff plot (Figure 2) shows the usual reversed-phase high performance liquid chromatography (HPLC) behavior of an increase in temperature resulting in a decrease in retention. With this information, separation conditions may be chosen to optimize resolution and analysis time.  

Figure 2: Plot of ln k vs. 1/T (van ’t Hoff plots), for two analytes (mock and synthesized data). Analytes 1 and 2 correspond to the circles (top) and squares (bottom), respectively.			

Prelude to the Next Installment on High-Temperature LC 

In our next installment of this series on elevated temperatures in LC, we will discuss the van 't Hoff equation in greater detail, including: the van 't Hoff equation and retention mapping (4,5), the thermodynamics of linear van 't Hoff plots (6), nonlinearities in van 't Hoff plots because of phase transitions (7–9), irregularities in van 't Hoff plots because of pH effects (10,11), confirmation of the linearity of van 't Hoff plots and evaluating changes in entropy (12), as well as some concerns about the van 't Hoff equation (13,14). 

In this installment, we introduced some of the basic thermodynamics of liquid chromatography that lead to the van 't Hoff equation. The next installment in this series will describe examples from the literature and important considerations for the van 't Hoff equation. 

The authors explore some basic thermodynamic relationships.

Elevated Temperatures in Liquid Chromatography, Part II: Basic Thermodynamics of Elevated Temperature LC, Including the van 't Hoff Relationship

The first observations of electrophoresis were made in 1807 by Ferdinand Frederic Reuss. Arne Tiselius (1902–1971) introduced electrophoresis as an analytical technique in 1937 (1) and was rewarded with a Nobel Prize for it in 1948 (2). One of his PhD students was Stellan Hjertén (1928–), who further developed the technique in open tube electrophoresis. His thesis was published in English in 1967 in 

 (3) and he has been rewarded many times over the years for his pioneering work, most recently he was given the 2011 CASSS Award for Separation Science (4). Other pioneers in the field include Virtanen (5) and Mikkers and Everaerts (6) who performed electrophoresis in 200-μm PTFE and glass tubing. In 1981, Jorgenson and Lukacs made the important step to apply 75-μm fused-silica capillaries (7); afterward capillary electrophoresis (CE) really took off. 

CE can be described as an automated, analytical version of the conventional electrophoresis techniques. The two sidebars in this article explain the separation principles of CE. The main advantages of performing electrophoresis in a capillary are the magnificent efficiencies and the automation possibilities. Because of the small diameters of the capillary, typically in the 20–100 μm inner diameter range, the Joule heat dissipation is very efficient. Consequently, high voltages, usually up to 30,000 V, can be applied before excessive Joule heating becomes an issue. Applying high voltages results in fast separations with very little band broadening.  

Usually, the capillary is made from fused silica, so on-column UV detection is feasible. Therefore, time-consuming staining and destaining procedures as seen with slab-gel electrophoresis are no longer needed. The combination of the small-inner-diameter capillary and on-column UV detection means that automated equipment has been developed with peaks resulting in electropherograms, simplifying quantitative analysis. Small bands show as efficient peaks with high plate numbers. 

Electrophoresis separates analytes that differ in charge-to-size ratio, so charge is a prerequisite for separation by CE. Several solutions have been created to induce charge and size differences in a capillary, as well as to add extra separation mechanisms. Table I lists some of the most frequently used modes of CE. 

Table I: The different modes of capillary electrophoresis			

CE is applicable over a wide range of analytes. Anything from small anions and cations to chiral separations, large proteins, DNA, cell organelles, and even complete cells and viruses have been analyzed with CE. The small scale of CE makes it a very green technique. Typically, a 50-cm long, 50-μm i.d. capillary has a capillary volume of 1 μL. Most often the separation medium is aqueous. So, the consumption and cost of chemicals are very low.  

The small scale of the techniques also means that only a few nanoliters of sample are injected. This is advantageous for application areas in which sample size is an issue, such as bioanalysis or the discovery phase of drug development. 

Electrophoresis is a fundamentally different separation technique from chromatography. Chromatographic separations are based on partition differences of the analytes between the stationary and mobile phase. Electrophoresis is based on differences in migration of charged particles in an electric field. Consequently, chromatography and electrophoresis are complementary tools in the analytical chemist's toolbox. Of course, there are separation problems that can be solved with both techniques. But, even so, there are problems for which one technique proves superior. The art of good analytical science is to select the tool that uses the strength and thus robustness of a technique rather than apply it at the limit of its capabilities and, therefore, have it be intrinsically less robust. 

Initially, fused silica was used as a capillary column because of its inert nature, UV transparency, and low cost. However, "inert" is not a very apt word in this case. The use of fused silica for CE results in the so-called electro(end)osmotic flow (EOF, see sidebar). As can be understood from the origin of it, the EOF is sensitive to anything influencing the capillary wall and the protonation or deprotonation of the silanol groups. Thus, for good precision it is important to understand and control the EOF process well. Early on, we understood the importance of caustic rinsing procedures to condition new fused-silica capillaries. Presequence and prerun conditioning procedures have been (and should be) the point of attention for all method developers because these procedures are method specific. Besides conditioning of the wall, there can be other reasons for preconditioning steps: 					

refreshing the separation medium to prevent buffer depletion; 

preventing carryover from highly concentrated sample components; 

and flushing out late-migrating components that are of no interest.

Table II lists examples of potential conditioning steps (8). The simplest step, rinsing with the separation buffer, should always be part of the prerun conditioning. Other steps might be added as needed. 

Table II: Possible preconditioning steps			

Still, a good conditioning scheme is sometimes not sufficient to control the stability, magnitude, and direction of the EOF and cannot always sufficiently suppress analyte-wall interactions. In light of this, coating procedures have become more and more common, especially adsorbed coatings. These can be divided into two groups, static adsorbed and dynamic coatings. Examples of the static adsorbed coatings are polymer coatings such as the cationic polybrene and the anionic dextran sulfate and poly(vinylsulfonic acid). These coatings are very stable and effective as double- and triple-layer coatings, also called successive multiple ionic polymer layer (SMIL) coatings (see reference 9 for a review). If properly applied, these coatings are stable for many runs and the coating polymer does not need to be present in the background electrolytes (BGE). This gives more flexibility to optimize the BGE independent of the coating. SMIL coatings are being applied in many commercial kits, such as CEofix (Analis), and also for many CE–mass spectrometry (MS) analyses of proteins. 

The other group of adsorbed coatings are the so called dynamic coatings. These have to be present in the BGE for an effective, repetitive, and reproducible effect. Dynamic coatings can be ionic surfactants such as cetyl trimethylammonium bromide (CTAB), monoamines such as triethanolamine, diamines such as putrescine, or polyamines such as spermine. To demonstrate how well-used these coatings are, here are two examples from the pharmacopeias: Triethanolamine is being used in the pharmacopeial method for the enantiomeric purity of 

-ropivacaine (Figure 1 [10,11]) and putrescine is being used in the pharmacopeial method for the separation of erythropoietin (EPO) isoforms. 

Figure 1: The enantiomeric purity determination of (S)-ropivacaine. Capillary: 80.5 (72.0) cm Ã 50 Î¼m fused silica; background electrolyte (BGE): 100 mM phosphoric acid, 88 mM triethanolamine, and 10 mM DM-Î²-CD, freshly prepared and filtered over 0.45-Î¼m pore size PTFE filters; BGE pH: 3.0; applied voltage: 30 kV (current 42 Î¼A) with initial ramping of 500 V/s; temperature: 30 Â°C; injection: 50 mbar Ã 5 s; detection: UV absorbance at 206 nm with 4 nm bandwidth; prerun conditioning: 1 min flush of water, 4 min of 0.1 M sodium hydroxide, 1 min of water, and 4 min of BGE (before the start of a sequence the capillary is rinsed for 10 min with 0.1 M sodium hydroxide or for 30 min for a new capillary); capillary storage: stored dry after a rinse for 10 min with sodium hydroxide, 10 min with water, and 10 min from empty vials; sample: 2 mg/mL ropivacaine hydrochloride solution for injection, directly injected without further sample preparation. Quantification by internal normalization of corrected peak areas. The %RSD for the amount of (R)-ropivacaine in this sample was 8%. Sample preparation of other ropivacaine products: higher concentrations of solution for injection are diluted with water to 2 mg/mL. Ropivacine HCl substance is dissolved in water to 2 mg/mL. 			

As discussed, anything influencing the capillary wall can affect precision and reproducibility. Therefore, it is also important to realize that a capillary has a history and to stick with one application per capillary. Furthermore, after method development the capillary has probably "seen" separation buffer components or rinsing solutions that did not make it to the final method. So after method development, it is a good practice to recheck the final optimized conditions on a fresh capillary. 

There are other details, such as the cutting of the capillary, that are part of good CE working practices that are important to get sharp, reproducible peaks. Although not difficult (12) and easy to check, improper cutting often gives rise to loss of efficiency and resolution (see Figure 2 ). Even a broken outlet might result in sloping baselines or tailing peaks. Remove the outer polyimide coating a few millimeters from the inlet and outlet of the capillary. This will reduce carryover and give better precision. This is important especially when using solvents that make the polyimide swell, such as acetonitrile. Removing the polyimide from the inlet and outlet is not advisable for some coated capillaries because it might damage the inner coating. 

Figure 2: Sample zone and peak distortion by badly cut or broken capillary: (a) Conditioned capillary filled with BGE; (b) inlet BGE vial replaced by sample vial; (c) injection of sample; (d) inlet BGE vial back in place, migration ongoing; (e) blue: peak shape that results from the distorted zone shape; red: peak shape if the capillary inlet had been straight and the sample zone normal.			

Do the Pharmaceutical and Biotech Industries Really Use CE?

So, is CE really being used in industry? An academic friend recently remarked that industry must be using CE less than before since the number of scientific publications from industry is declining. Others have made similar remarks. Other rumors are that CE is not reproducible and robust, so it cannot be used for quality control (QC). Additionally, people think that CE methods cannot be validated, and even that regulatory bodies such as the United States Food and Drug Agency (FDA) do not like CE. Are these comments really supported by the facts?  

The facts are that there are many successful methods during all stages of development within industry, although this cannot be fully proven by publications. There are adopted methods in use including applications from the pharmacopeias such as the 

) (for a selection of these methods, see Table III; for a review see references 13 and 14). Of course, to get as far as this, these methods have to be successfully validated. Experiences from the field and sometimes reports in peer-reviewed publications show that for CE, similar robustness and reproducibility is possible, as with other techniques and methods such as high performance liquid chromatography (HPLC). There are several successful intercompany studies using multiple modes of CE (15–21). The recent increase in sales of instruments show that CE is still a growing market. 

Table III: CE tests in the European Pharmacopoeia (Ph. Eur.) and the US Pharmacopeia (USP)			

Furthermore, there is a good knowledge base within the regulatory bodies of CE techniques. Representatives actively participate in symposia and are part of organizing committees such as for CE in the Biotechnology & Pharmaceutical Industries (CEPharm). This CASSS-run meeting has seen a steady attendance from industry, vendors, and regulatory bodies, and had its 14th meeting this year. 

If one submits CE methods to regulatory bodies, they receive the same attention as other submitted methods. Just as with other methods, one must justify the need for the method and provide the thorough information needed for reviewing the submission, irrespective of the technology used. 

Applications from the Pharmaceutical Industry

CE is being used by the pharmaceutical and biotech industry in all phases of drug development: not only for identity, assay, and purity analysis of the active pharmaceutical ingredient (API), but also for other types of applications such as physical-chemical characterization (p

, logP, and logD determinations and drug–protein binding), analysis of counterions, formulation development support, conformation of results obtained by other techniques, and troubleshooting. Because of the proprietary nature of the drug substances being investigated in the pharmaceutical industry, many of these applications never get published. Here are some examples that illustrate the use of the different CE modes. 

There are many similar methods that are successfully being applied for the determination of drug countercations and counteranions. Mostly, these methods use indirect UV detection. An excellent book chapter on ion analysis was written by De l'Escaille and Falmagne (22). Figure 3 shows an example of the determination of bromide as potential impurity in HCl drug salts with CE and direct UV detection (23). Indirect UV detection has limited loadability and linearity and is less suitable for determining the presence of a large excess of chloride because electromigration dispersion of the highly concentrated chloride peak may impair the resolution. The direct-UV bromide determination was optimized with respect to the resolution of the chloride and the bromide peaks by statistical experimental design using a multivariate optimization program. A sample matrix containing a high concentration of an ion such as chloride may cause antistacking. However, if the mobility of the chloride ion is lower than the mobility of the bromide ion, the high chloride concentration may contribute to a sample self-stacking effect. Furthermore, the buffer co-ion must be carefully chosen with respect to its mobility to achieve an efficient separation system (23). 

Figure 3: Determination of bromide in hydrochloride drug salts using CE with direct UV detection. Capillary: 50 Î¼m i.d., effective length 41.5 cm, fused silica, extended light path (bubble factor 3); background electrolyte: 60% acetonitrile and 100 mM methanesulfonic acid adjusted to pH 1.3 with triethanolamine; sample: local anesthetic HCl salt containing 0.3% (w/w) bromide; voltage: -15 kV; temperature: 20 Â°C.			

An example from the biotech industry is the charge heterogeneity determination of proteins such as monoclonal antibodies. Yan He of Pfizer (24) published a rapid analysis on short, dynamically coated fused-silica capillary. The pH, concentration of the separation buffer (?-amino caproic acid), concentration of the triethylenetetramine (TETA) dynamic coating, the capillary internal diameter, and the applied field strength were investigated and optimized. He also evaluated the effects of between-run flushing of the capillary and the data acquisition rate. Compared with other existing methods for charge variants analysis, this method had several advantages including a short run time, rapid capillary conditioning and simple sample preparation. At the MicroScale Bioseparations (MSB 2012) meeting in Geneva, Switzerland, Bernd Moritz of Roche showed that the same methodology on a longer capillary results in improved resolution of the isoforms, at the cost of longer analysis times (25). 

				The pharmacopeial test method for identity and assay of erythropoietin (EPO) is also a capillary zone electrophoresis (CZE) method with a dynamically coated capillary, in this case with putrescine (1,4-butanediamine). Long rinsing procedures are required, but when optimally conditioned, the method shows highly resolved and efficient peaks for the EPO isoforms (26). In the pharmacopeial identification and charge variants test method for somatropin, somatropin concentrated solution, and somatropin for injection, deamidated charged variants migrate after the main peak (27). 			

Micellar Electrokinetic Chromatography and Microemulsion Electrokinetic Chromatography

In micellar electrokinetic chromatography (MEKC), micelles are added to the separation solution to separate uncharged analytes. Uncharged analytes partition between the more lipophilic inner part of the micelles and the hydrophilic aqueous buffer. So part of the time they move with the micelles, and part of the time with the EOF. Their migration order will depend on the strength of the interaction with the micelles: the more lipohilic the analyte, the stronger the interaction. That means that uncharged analytes will show an effective mobility that is between the mobility of the micelles and the mobility of the EOF. Charged analytes will experience both this chromatographic effect and the electrophoretic forces, and their net mobility is harder to predict. 

An elegant example of a MEKC method was published by K. Persson-Stubberud (28,29). She developed, validated, and submitted for regulatory approval one method for the quantification of ibuprofen and codeine and the determination of degradation products in commercial tablets. Factorial design was used for both method development as well as robustness testing. 

If the analytes of interest are more lipophilic, using microemulsions in microemulsion electrokinetic chromatography (MEEKC) might be a better solution than using micelles as additives for optimal separation. An interesting approach in problem solving using MEEKC was nicely demonstrated by McEvoy (30). Suppositories containing paracetamol were dissolved and analyzed with the same microemulsion, making sample preparation simpler. 

CE is a strong tool for chiral separations since the chiral selector is dissolved in the separation buffer. If the dynamic complexation with the chiral selector differs between the enantiomers or if their complexes differ in mobility, they can be separated. Because the chiral selector is in the separation solution, a whole series of chiral selectors can be screened in a short time. On top of that, the amount of chiral selector and its concentration in the buffer can vary. Variation of the chiral selector concentration is an important parameter in method optimization.  

Although one can fairly successfully predict interaction of a compound with a selector, it is still virtually impossible to predict enantiomeric separation; for example, see Figure 4 and references 31 and 32. Consequently, it is advisable to test as many selectors as possible in a screening experiment. The most frequently used chiral selectors are cyclodextrins, as they are UV-transparent and a wide variety of substituted cyclodextrins are commercially available. Figure 4 shows the enantiomeric purity separation of 

-ropivacaine, a method that has been taken up in the pharmacopeias. Under exactly the same conditions (BGE: 100 mM phosphoric acid, 90 mM triethanolamine, 10 mM DM-β-CD), some of the enantiomer pairs are very well separated (for example, resolution of 7.5 between DLDD and LDLL), and some are not separated at all, such as DLDL and LDLD. All chiral forms of the tetrapeptide showed association complexes with the cyclodextrin, as was observed by migration time shifts for all after adding the cyclodextrin to the buffer. The main difference was the association differences for the enantiomer pairs (23). 

Figure 4: The separation of the 16 stereoisomers of the tetrapeptide Tyr-Arg-Phe-Phe-NH2, presented as eight enantiomer pairs.			

Within the biotech industry, conventional isoelectric focusing (IEF) has been replaced by capillary isoelectric focusing (cIEF) in many cases. cIEF is either performed on a short coated capillary with whole-column detection, on a so-called imaging instrument (icIEF), or on a traditional CE instrument. In both cases, the sample is mixed with ampholyte and the capillary is filled with this mixture. When the voltage is applied, the ampholyte forms a pH gradient through the capillary and the proteins migrate in this pH gradient until they reach the pH where their net charge is zero — that is, where the pH equals the isoelectric point (pI). The main advantage of performing IEF in a capillary is its speed and quantitative capabilities. There are generic methods and standard kits available. Traditional ampholytes are being used, although some have better UV properties than others. The major parameters for method optimization are in the sample prep procedure, that is, desalting and the amount of urea needed. 

Yan He compared cIEF with CZE for monoclonal IgG (24) and found that overall, icIEF showed better separation of acidic isoforms, whereas CZE showed better separation of basic isoforms. 

In capillary gel electrophoresis (CGE), a noncrosslinked linear polymer is used to form the sieving gel. Again, the instrument automation and improved quantitation properties make the capillary format interesting. A huge application area is the analysis of DNA, although the capillary form of sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) for proteins receives more attention in the pharma industry.

 also is being used for the capillary format of SDS-PAGE. There are commercial kits available for molecular weight determinations of proteins and for the charge heterogeneity determination of monoclonal antibodies (19). With UV detection, Coomassie blue sensitivity is achieved, and with laser-induced fluorescence (LIF) detection silver staining sensitivity is possible. All of the CGE applications for DNA, RNA, and proteins are also available on chip-sized analyzers. 

Column Watch

A look at what influences the use of CE in the pharma industry.

Revival of Capillary Electrophoretic Techniques in the Pharmaceutical Industry

Gases for gas chromatography (GC) have become a hot topic in recent months, primarily because of concerns over short supplies of helium. Already one of the top discussion items, hydrogen as a carrier gas has garnered much of the attention as chromatographers' awareness of these issues continue to expand. Just last month (October 2012), on-line web seminars from Agilent and CHROMacademy were dedicated to conversion from helium to hydrogen carrier gas. A web search on "helium hydrogen carrier" yields guidelines and instructions from every major GC manufacturer and supplier as well as myriad topical threads on all of the GC blogs, boards, and discussion groups. Go to Pittcon, Analytica, the Eastern Analytical Symposium (EAS), or any other conference where GC is on the agenda and the helium issue will be featured prominently. 

Among all the discourse, I have noticed that the essential related topic of good practices for deployment of carrier gases and, for that matter, all gases used in GC, is largely missing. Although much of this good advice is easy enough to find in instrument installation guides and supplier catalogs, the connection between obtaining the information and putting it into practice is often missed in many laboratories. Most laboratories will install gas filters in-line, but many will fail to obtain the right type of regulator, make the gas connections correctly, or maintain the filters and check the regulators on a regular schedule. 

Questions That Should Be Asked Frequently 

Here, then are some guidelines and recommendations about GC gases, in a question-and-answer format. This is not an exhaustive list, but rather it covers some of the more frequently asked questions as well as some that are not asked as often as they should be. The list starts at the gas source and moves onward to the instrument. Questions about the instrument internals are not addressed because of space limitations. 

Table I: Recommended carrier gas purity*			

What Are the Recommended Gas Purities for Carrier and Detector Gases?

The exact requirements for gas purity should follow the instrument manufacturer's guidelines as found in their site preparation and installation manuals. If that information is not available, then Tables I and II will serve as a general guideline. The gas purities are stated as percent levels rather than referring to supplier-specific names, which can be ambiguous or inconsistent. 

Table II: Recommended detector support gas purity*			

The purity of a gas when it reaches the back of the instrument depends on the supply quality, regulators, filters, fittings, and connecting tubing. Filters will clean up minor contamination, but they are not intended to take gas to a higher purity level. Most of the time the purity of cylinder gas is as labeled on the bottle, but occasionally a contaminated cylinder may make it to delivery. Although it is bad practice on the part of cylinder users, a cylinder might be left open to the atmosphere for hours when empty and removed from service. If not cleaned up by evacuation and baking before filling and then filled with gas to 2450 psig (166 mPa), such a cylinder will contain approximately 6000 ppm of air, which degrades the gas purity to 99.4%. Although it is extremely unlikely to arrive in a cylinder at the receiving dock, this level of contamination represents a conceivable upper limit. When placed in service, the resulting onslaught of oxygen, water, and possibly hydrocarbons will completely exhaust a high-capacity gas filter before the contaminated tank is empty.  

This potential for contamination is an excellent reason to use indicating filters on all gas supply lines. The indicator will change color as the filter reaches capacity. As long as the color change is noticed, a new filter is installed, and a pure gas supply is restored, the GC instrument will be spared the indignity of gas contamination and resultant high detector background, irregular baselines, and accompanying loss of signal-to-noise and repeatability. 

It is possible, but expensive, to order purity analyses of individual cylinders. This step is only significant when it is difficult to observe the filters or replace the cylinder, such as at remote unattended locations. 

How Often Should the Gas Delivery System Be Checked?

Given the slight chance of a contaminated cylinder, check indicating filters every time a tank is replaced or at least once every two weeks if using a gas generator. That's also a good time to check the gas lines for unusual bends or kinks. Check for leaks any time a fitting is disconnected and reconnected, and of course on all new connections. 

It's also a good idea to check regulator function at least once a year. The easiest way to do this is to establish a flow of 500 mL/min or greater on each gas cylinder. Note the pressure output, then turn the tank valve off. Wait until the outlet pressure starts to drop off, and then turn the tank valve back on. The same pressure should be reestablished. Next, turn the regulator pressure knob down and back up while observing how smoothly the knob moves, how the output pressure gauge reacts, and that the knob is not screwed in all the way when the original pressure is restored. Don't forget to set the flow back to normal at the gas chromatograph. 

Check that the correct cylinders are connected through to the correct fittings on the instrument. Some cylinders, such as helium, argon, and nitrogen, use the same gas fitting (at least in the United States); hydrogen and high-level combustible gas mixtures also may share a fitting type. Thus, it is possible to connect the wrong cylinder. Chromatographers should not rely on the uniqueness of cylinder fittings to ensure the correct gas type. 

What Kinds of Regulators Should Be Used for GC?

Generally, dual-stage high-purity regulators with stainless steel diaphragms are the correct choice in all cases. For economy, less expensive dual-stage regulators may be used for detector air, hydrogen, and make-up gas if separate from the carrier supply. The added cost of a dual-stage compared to a single-stage regulator does not justify the potentially poor pressure regulation of a single-stage regulator as the cylinder pressure decays. 

A regulator outlet valve is a handy accessory that I like to order on all regulators. However, in laboratories where the downstream connections remain in place permanently, the outlet valve can be omitted. 

Always dedicate a regulator to its intended use. Never change the cylinder fitting on a regulator to switch it from inert-gas to detector-gas service or the other way around. Making the cylinder fitting connection correctly so that it will maintain high pressures is best left to the regulator manufacturer. 

What Types of Fittings and Tubing Are Required?

The fittings and ferrules used in any GC installation should all be new, as should the tubing. The fittings should be of a suitable type for high-purity gas lines, such as those that are available from instrument company and supplier catalogs. I always like to order some spare fittings to have on hand for the usual problem connection that refuses to seal, as well as for later on when the gas setup needs some modifications. 

Only metal tubing should be used for GC gases, with one exception. Copper tubing is the easiest to install, while stainless steel tubing is more robust and generally can be more organized visually. Either type of tubing must be precleaned before installation, and can be ordered that way as "GC" or "Chromatography" cleaned tubing. "Refrigerator" designated tubing is not suitable. For installations with tanks or gas generators dedicated to one or two instruments, ⅛-in. or 3-mm o.d. tubing is appropriate. If multiple instruments share a tank or generator then consider using ¼-in. or 6-mm o.d. tubing up to the point where the flow path splits to the individual instruments.  

Polymer tubing is appropriate only in one case. If the GC system uses air-actuated sampling valves then that air supply can be connected with polymer tubing. But if the air tank is shared with a flame ionization detection (FID) system — which is not such a good idea, but it is done — then metal tubing is required throughout. Polymer tubing may allow air and airborne contaminants as well as monomers from the plastic into the gas stream. This is unacceptable for carrier and detector gases, even for FID air. 

A nice touch when installing tubing is to mark either end of each tube with differently colored electrical tape. This makes it clear which tube goes to which gas inlet and regulator and avoids some of the more hazardous mix-ups such as swapping FID hydrogen and air. (Yes, I have seen that situation and the result of igniting the flame was, well, exciting!) 

What Is the Right Way to Make Gas Connections?

Three gas connection types are encountered in a GC system. Occasionally chromatographers encounter other fitting types, but these three are the most common. First is the high-pressure tank fitting, identified by a letter and number designation such as CGA-580, DIN 477-6, or BS-341-3 for inert gases. With few exceptions no additional sealing is required — just assemble the fitting to the cylinder and tighten to seal. Most regulator fittings used in GC have a torque specification of 40–60 ft-lb (54–81 Nm). Some types of high pressure fittings, notably for liquid CO

 in the United States, require a plastic washer that is usually suppled with each tank. If a washer is present then assume it must be used, and the tightening torque must be reduced by half. If not in place, the absence of a washer will immediately be evident upon opening the cylinder valve! Never use polyfluorocarbon tape or liquid sealant anywhere on a high-pressure tank fitting; this will just make it leak. If either side of the fitting is scratched or deformed then replace the fitting. 

The second type of fitting found in GC systems is the pipe-thread type. This fitting involves matching internally and externally threaded sides with no washers or ferrules. After making sure the threads are clear of old sealing tape, wrap two layers — no more — of polyfluorocarbon pipe sealing tape (available from instrument suppliers) onto the externally threaded side of the fitting. Holding the exit end of a right-hand threaded fitting toward you, smoothly wrap the tape in a clockwise direction while stretching it slightly. Then thread the taped piece into the internally threaded side and tighten. 

A number of frequently asked questions about gases and their delivery to a gas chromatography instrument are addressed.

LCGC North America-11-01-2012

Evaluation of an LC Method

Evaluation of an LC Method

Elevated Temperatures in Liquid Chromatography, Part II: Basic Thermodynamics of Elevated Temperature LC, Including the van 't Hoff Relationship

Revival of Capillary Electrophoretic Techniques in the Pharmaceutical Industry

Gases Q & A

Assay for Parts-Per-Million Levels of Azide in Drug Substances

Quadrupole Mass Analyzers and MS-MS Analysis

Phyllis Brown: A Story of Serendipity, Perseverance, Vision, and Chromatography

Vol 30 No 11 LCGC North America November 2012 Regular Issue PDF