Small changes in retention time of a liquid chromatography (LC) method are normal. At what point is a problem suggested? Retention time shifts can be frustrating when you can't figure out if a shift is something you caused or if there is another reason for it.

It is interesting how problems tend to cluster — in the last few weeks I've received several questions related to retention time shifts with liquid chromatography (LC) methods. Some of these correspond to the preparation of new batches of mobile phase, some are from one day to another, some are from within a batch of samples, and some result from a change in instruments. In this month's "LC Troubleshooting," I will discuss some of the factors that influence small changes in retention. 

Most LC methods run on modern instrumentation with a good quality column will have quite stable retention times. I generally expect to see run-to-run variation in the second decimal place of the retention time, such as ±0.02–0.05 min. However, the historical behaviour of the method should be used to determine what is normal for that method. For example, with large biological molecules, such as proteins, that use shallow gradients, the variability can be an order of magnitude or more larger. 

One of the most common causes of shifts in retention time in reversed-phase LC separations is a minor change in the concentration of the organic solvent, usually methanol or acetonitrile. This can happen from a minor error in formulation or a change in the mobile-phase composition if one solvent evaporates over time. 

For small molecules (arbitrarily <1000 Da), we can use the "Rule of Three" to estimate the effect of a change in organic solvent, or %B. This rule states that the retention factor, 

, changes approximately threefold for a 10% change in %B. This rule derives from plots, such as that in Figure 1, where 

 is plotted versus %B on a semi-log plot. The red line in Figure 1 represents the retention behaviour for a 500 Da analyte. Retention changes from ~29 min at 40% B to ~4 min at 60%. For practical purposes, this relationship can be considered linear and represented in the standard 

 is the (theoretical) retention in 100% water, 

 is the slope of the plot, and Φ is the %B as a decimal (0.5 = 50%). Values of 

 can be determined from two experimental runs using equation 1. Empirical observations indicate that 

where MW is the molecular weight (Da). Thus for a 500 Da analyte, 

 ≈˜ 5 as an average for sub-1000 Da molecules, we can then estimate how 

  = 3.16 ≈˜ 3. This is the basis of the Rule of Three. 

As an example, our 500 Da analyte in Figure 1 has 

 = 5 at 50% B. We can convert this to retention time, 

 is the column dead time (also abbreviated 

). If we assume a 150 mm × 4.6 mm column operated at 1 mL/min, 

 = 1.5(1 + 5) = 9.0 min. The Rule of Three would suggest that 

 ˜≈ 3 × 5 = 15 for a change to 40% B, which would correspond to 

 = 28.7 min — close enough for a rule of thumb. 

Mobile-phase formulation errors should not be in the 10% region if you are at all careful, so what happens for smaller changes? We can use equations 3 and 5 to determine the effect of a 0.1%, 0.5%, and 1% error in formulating our 50% B mobile phase for our "average" 

 = 5 small molecule. I'll leave the calculations to you, but the results summarized in Table 1 show that these correspond to retention shifts of ≈˜ 0.1, 0.4, and 0.9 min, respectively (I have rounded values for display, so if you try to repeat my calculations, your results may vary somewhat). So, you can see that it takes only a minor error in mobile-phase formulation to shift retention times enough to notice the change. 

 Table 1: Effect of small errors in mobile-phase composition on the retention for analytes of different S values. 			

With larger molecules, the problem is magnified, because 

 increases markedly with an increase in molecular weight. This makes the log 

 versus %B plots steeper, as is seen for 5000 Da (blue) and 50,000 Da (green) compounds, such as peptides or proteins, respectively. Steeper plots mean that these compounds are much more sensitive to minor changes in the mobile-phase composition. Our Rule of Three for <1000 Da samples becomes the Rule of 60 at 5000 Da, and the Rule of 400 at 50,000 Da. This extreme sensitivity of the retention of large molecules to small changes in %B means that isocratic separation is not practical for most separations of these analytes. 

Linear plots, as in Figure 1, are also observed if you plot log 

 versus reciprocal temperature (in kelvin [K], where K = °C + 273). This means that small changes in column temperature can also affect retention times. A rule of thumb for small molecules is that retention changes by ˜≈2% for each 1 °C change in temperature. In Figure 2, you can see that a 10 °C change in temperature changes the retention of the last peak from approximately 12 min to 10 min, as expected. Most workers know that retention can shift with a change in temperature, but many overlook the fact that relative retention can also change. In the (specially selected) sample of weak acids and bases of Figure 2, there are three reversals in retention for a 10 °C change in temperature. Smaller changes in temperature will have smaller, but noticeable effects on the appearance of the chromatogram. Such problems can be exacerbated by two common factors. Operation with the column at ambient temperature can result in several degrees change in column temperature during a single day, with subsequent retention shifts. In my observation, the temperature of the mobile phase within the column can vary several degrees from the column oven setting and if the oven is not properly calibrated, or of different design, switching from one column oven to another can give similar temperature differences. Usually, temperature differences cause shifts in retention times of all peaks to earlier or later times, and small adjustments in the temperature setting can correct for instrument-to-instrument differences. Because of the wide variation in ambient temperatures that are possible, I strongly recommend that you always use a column oven. 

Figure 1: Plots of log k versus %B for compounds of 500 Da (red), 5000 Da (blue), and 50 kDa (green). See text for details.			

A change in the mobile-phase pH can have a dramatic effect on the appearance of the chromatogram when ionizable compounds are present, but it makes no difference for neutral samples. The behaviour of retention with a change in pH will vary depending on how close the mobile-phase pH is to the p

 of the analyte, and in any case, is not a linear relationship, so a general rule of thumb for the effect of pH on retention is not possible. The lower chromatogram of Figure 2 illustrates how sensitive retention can be to changes in pH. A change in pH of 0.2 units gives similar retention changes as a 10 °C change in temperature for this sample of weak acids and bases. Typical laboratory practice of adjusting the mobile-phase pH with the help of a pH meter can result in errors of ±0.1 pH units, so special care needs to be taken if the separation is particularly pH sensitive, as is that of Figure 2. To minimize pH-related problems, use buffers within ±1 unit of their p

, make buffers by blending equimolar portions of acid and base, ensure that the buffer is sufficiently concentrated (usually 20–25 mM is adequate), and make sure to use a column oven. 

Figure 2: Effect of a change in column temperature of 10 Â°C or 0.2 pH units for a sample of weak acids and bases. Data from reference 1; see text for details.			

Any change in flow rate of the pump will directly affect retention for isocratic separations, whereas the effect on gradients can be a bit more complicated. I did an informal on-line survey of the specifications of seven instrument models from three major LC manufacturers. This shows that the typical specifications for flow accuracy are ±1% and for precision are ±0.07% relative standard deviation (RSD) or ±0.02 min, whichever is greater.

This means that it would not be unusual for the retention to vary by 0.1 min (~1% of our 

 = 9 min sample used in the examples above) between instruments. Between-run variation of 0.02 min should be considered reasonable. With quaternary, low-pressure mixing systems, flow will not affect the mobile-phase composition; specifications for such systems are for maximum retention variations of ±0.02–0.04 min because of proportioning errors. However, with binary high-pressure mixing systems, flow rate will affect the solvent proportions, and thus retention.  

Any pump malfunction, such as problems with check valves, pump seals, leaks, or bubbles, will also be reflected in analyte retention times. Because such changes will only reduce the flow rate, increases in retention would be observed when these problems exist. 

Sometimes the instrument can be working within its specifications and you have taken care to control the column temperature and carefully formulate the mobile phase, but retention time variations are still not acceptable. An example was given in an earlier "LC Troubleshooting" column (2) where a freshly serviced instrument (new check valves and pump seals) still resulted in retention times that differed by up to 1 min between runs, as can be seen at the top of Figure 3. The sample was a peptide sample run with a very shallow gradient (0.17%/min). The instrument was specified to generate mobile phase mixtures within ±0.1%, but it can be seen that this variation was more than half of the gradient change per minute. Although the system was working as specified, the demands of the method were too great. In this case, instead of using 100% buffer as solvent A and 100% acetonitrile as the solvent B, the solvents were blended so A comprised 10:90 buffer–acetonitrile and B comprised 30:70 buffer–acetonitrile. The gradient programmed into the instrument was modified to give the same actual gradient as the original, and consecutive injections varied by <0.1 min in retention. By premixing the mobile phase, the effective precision of the instrument was increased from ±0.1% to ±0.02%, which allowed the method to be run acceptably. This problem was partly related to demanding more of the LC system than it was capable of and the extreme sensitivity of the peptide sample to small changes in the mobile phase, as discussed at the beginning of this article. 

Figure 3: Run-to-run retention variation for three consecutive injections of a peptide sample separated with a 0.17%/min gradient. Mobile phases: A = buffer and B = acetonitrile (top); A = 10:90 bufferâacetonitrile and B = 30:70 bufferâacetonitrile (bottom). Data from reference 2, see text for details.			

All of the above discussion centres around normal variations that can be expected with a well-maintained instrument and reasonably careful laboratory practice. Small errors in method conditions are inevitable, so LC methods should be designed to tolerate them, and areas of specific sensitivity should be noted in the method document. These areas are addressed by the International Conference on Harmonization in two places, a document about method validation (3) and one on quality by design (QbD) (4). When a method is validated, its robustness should be demonstrated. Robustness is the tolerance of the method to small, but normally expected, changes in operational variables. This might encompass ±1% change in %B, ±2 °C, ±0.2 pH units, and so forth. QbD principles state that quality should be designed into a method, in particular its sensitivity to changes in operational variables. This means that during method development, experiments should be run to determine how each variable (for example, %B) influences the separation, as well as how interactions that may occur when changes in more than one variable take place. Based on such studies, a design space can be established. The 

 is a multidimensional space bounded by the limits of each of the operational variables. As long as the method is operated within the design space, it should work properly. Therefore, a method can include instructions about how large a change in each variable can be made, while allowing the method to produce acceptable results. Part of this information could include tolerances in retention time or relative retention. Armed with this kind of information, it would be much easier to know when an observed change in retention was a problem and when it was normal. In addition, if an unacceptable change occurred, the QbD information in the method would describe how to adjust the method to bring it back into compliance.   

We've seen that even small changes in the operational variables of a method can result in noticeable changes in retention time. Such changes in retention need to be evaluated to determine if they have any negative impact on the quality of the analytical results, and if so, corrective action can be taken. The observation that most methods operate without problems of excessive retention variation is an indicator of generally good laboratory practice and reliable instrumentation. So keep up the good work, but be watchful for potential problems. 

has been writing "LC Troubleshooting" for 

 for more than 30 years. One of the industry's most respected professionals, John is currently the Vice President of and a principal instructor for LC Resources in Walnut Creek, California, USA. He is also a member of 

's editorial advisory board. Direct correspondence about this column via e-mail to 

. To contact the editor-in-chief, Alasdair Matheson, please email 

(2) D.H. Marchand, P-L Zhu, and J.W. Dolan, 

(3) International Conference on Harmonization, 

Validation of Analytical Procedures: Text and Methodology, Q2(R1)

(4) International Conference on Harmonization, 

Articles

LC Troubleshooting

A fast, sensitive and accurate quantitative method was developed using HPLC–MS–MS for analysis of phthalate metabolites in urine samples.

How Much Retention Time Variation is Normal?

World of Technology & Science (WOTS) 2014 

 World of Technology & Science 2014 (WOTS)

 exhibition, formally known as the HET Instrument Exhibition, will be held between 

Jaarbeurs Exhibition and Conference Centre, Utrecht, The Netherlands,

 organized by two collaborating Dutch associations, FHI (Federation of Technology Branches) and FEDA. 

(PHOTO CREDIT: MARIA SWÃRD/GETTY IMAGES)			

is the instrument exhibition that is split into four different themed "worlds" that include the World of Automation, World of Electronics, World of Laboratory, and World of Motion & Drives.The "Laboratory World" will introduce visitors to new technologies, with around 150 companies expected to showcase their latest developments in analytical instruments, optical instruments, and laboratory equipment. Information about laboratory applications of technology will be given in the conference programme and through interactive projects. X-peditions are interactive guides that lead visitors through six to eight booths centred around different themes, for example "Safety in the Lab X-pedition".  

conference programme will offer 20 seminars covering topics including pharmaceuticals, food automation, factory automation and industrial efficiency. One particularly interesting seminar, "Insights into the Daily Challenges of Chromatography", will focus on the challenges and trends in sample preparation, ultrahigh-pressure liquid chromatography (UHPLC), liquid chromatography (LC), gas chromatography (GC), hydrogen as a carrier gas, multi-dimensional separation, and data processing. Johan Paul from Flanders Materials Centre (Ghent, Belgium) will present the optimal approach to automating sample preparation for analysis. All too often, automated sample preparation is seen as just a translation of a manual procedure to a classical autosampler when a more targeted approach could give better results. David Benanou of Veolia (Paris, France) will evaluate the use of hydrogen delivered by a gas generator as a carrier gas for gas chromatography–mass spectrometry (GC–MS) in sludge analysis. Several parameters will be explored, including the effect of the diffusivity of hydrogen on a vacuum system, the eventual variation of the fragmentation pattern of the steroids, and the change in detection capacity of the system. Hans-Gerd Janssen, Professor of Analytical Chemistry at the University of Amsterdam (Amsterdam, The Netherlands) and researcher at Unilever Research, will present the principles of multidimensional chromatography, with attention to several systems, including LC×LC, GC×GC, and LC×GC. This seminar will be held on the morning of Thursday 2nd October and will be organized by Avans+, Bruker, Da Vinci Laboratory Solutions, Interscience, JSB Joint Analytical Systems, and Waters Corporation.  

 for more information about the exhibition and free registration. 

30th International Symposium on Chromatography (ISC 2014)  

				 Salzburg Congress, Salzburg, Austria 			

 Wolfgang Buchberger, Michael Laemmerhofer, and Wolfgang Lindner 

5th International Conference on Polyolefin Characterization (ICPC) 

				 The Westin Valencia Hotel in Valencia, Spain 		

International Symposium on GPC/SEC and Related Techniques 

				 The Westin Grand Frankfurt, Germany 		

 Waters Corporation and PSS Polymer Standards Service GmbH 

The 17th International Symposium on Field- and Flow-based Separations (FFF 2014) 

				 The University of Utah, Salt Lake City, Utah, USA 			

 The Organizing and Scientific Programme Committees headed by Bruce Gale 

Events

World of Technology & Science (WOTS) 2014

In high performance liquid chromatography (HPLC) column development, the bonded-phase silica is often only investigated to determine the suitability of a packed column for a particular separation class and not much thought is given to optimization of the base silica to make the overall packing material more suitable for the assay. Here, we examine general-purpose standardized tests to determine the quality of bulk silica before and after bonding. Base silica features such as mechanical strength, acid and alkaline durability, loadability, and overload characteristics are a few of the parameters evaluated that have an impact on the final separation and purification characteristics for a difficult test analyte such as insulin.

Despite improvements in polymeric particles, the introduction of organic–inorganic hybrids and investigations of other inorganic oxides (for example, titania and zirconia), totally porous and superficially porous silica gels still dominate as the favoured base material in high performance liquid chromatography (HPLC) (1). The properties of silica, outlined in Table 1, make it an ideal packing base material for bonded-phase chromatography, which is by far the most popular approach for analytical and preparative separations. For example, more than 90% of chromatographers use reversed-phase liquid chromatography (LC) in their daily work (1). Even though spherical silica has been around for several decades, manufacturers are still improving silica particles to allow them to be used for even more demanding separations and purifications.  

Table 1: List of silica particle parameters.			

Although the actual number of analytical applications using bonded silica columns exceeds the number of preparative applications, for industrial purifications using porous silica-based packing materials, the amount of silica gel consumed worldwide greatly exceeds the amount of silica used to pack HPLC and ultrahigh-pressure liquid chromatography (UHPLC) analytical columns. In addition, although the properties of the packings used in preparative chromatography share similarities with the optimum properties of analytical chromatography, there are some properties that are more demanding when it comes to large-scale purifications. The purpose of this instalment is to discuss the properties of silica gel that must be improved further and then carefully controlled to meet the demanding applications in large-scale industrial purifications. For our test example, the purification of insulin was chosen since it is the most important large-scale process purification using bonded-phase silica gel chromatography. 

Diabetes is an incurable, life-threatening disease. The increase in individuals who are and will be affected by this disease correlates with the worldwide rise in obesity and the growing number of people with access to modern medications. The number of diabetes patients is projected to balloon to a staggering 500 million in the coming years. 

The most widely used medication for diabetes is insulin. This peculiar biomolecule comprises 51 amino acids with a molecular weight of a little less than 6 kDa. Insulin is considered either one of the biggest peptides or the smallest of the protein molecules. By its nature, the insulin molecule has a strong tendency to form dimers and further undergo aggregation-coupled misfolding (oligomerization) to form a cross-β assembly. This process is referred to as 

and can complicate the manufacturing process as well as its chromatographic purification.  

Insulin manufacturing is a standalone industry these days. Human insulin or its slightly modified variations are cloned into microorganisms. After the expression, insulin is harvested and must be rigorously purified not only from the fermentation broth residues but also from the misshappen isoform impurities. Such difficult separations can be performed using process-scale HPLC with huge dynamic-axial compression columns packed with silica-based reversed-phase stationary phases. Interestingly, the insulin manufacturing industry gobbles up a huge part of the total acetonitrile consumption of the world and almost half of the total spherical reversed-phase silica globally produced. 

The biggest technological challenge for insulin manufacturers is the insulin molecules fibrillating on top of the chromatographic column, forming a tough, chewing gum-like layer that blocks the solvent flow and increases the column back pressure. Currently, the solution, though not preferable for silica gel-based packings, is to use a pH 13 sodium hydroxide solution to dissolve and remove this layer blocking the column. This procedure combined with the generally higher than average pressure of the process can damage the silica-based stationary phases badly. Thus, there is a need to develop special stationary phases to address these challenges. 

Development of an Optimized Packing for Purification of Insulin

For small-scale (analytical) HPLC analysis of a given peptide (for example, the insulin molecule) typically needed at the discovery stage of a new drug development process, the most important parameter is the selectivity (separation factor) of the stationary phase used. However, when the very same peptide goes several years later into large-scale production, a whole set of new parameters become very important to make the stationary phase used in the chromatographic purification steps the best choice. 

Standardized testing protocols for general chromatographic bonded phase packings abound. For example, the Tanaka test (2), the Engelhardt test (3–5), the Neue test (6), and the National Institute of Standards and Technology (NIST) and The United States Pharmacopeia (USP) test (7) among others (8,9) are generally suited for the evaluation of the separation characteristics of the bonded-phase silica packed into analytical columns and not specifically for the bare silica used in their production. 

Few standardized tests have been developed for the base silica used for the preparation of chromatographic silica, especially when used for process-scale separation columns. Tests on the bulk silica before bonding can reveal underlying characteristics that may rule out a material before having it go through the bonding and characterization process. Part of the development process described here was the characterization of the base silica for attributes considered as the most important for large-scale purification. Several commercially available silicas and newly developed silicas were put through the testing process to understand the most important characteristics for large-scale insulin and peptide production. Research and development (R&D) chemists often attempt to optimize one or two packing parameters such as efficiency or resolution; however, we found that there are six important criteria for an optimized chromatography material for the preparative purification of insulin and other peptide products. Here, we cover each of these tests to determine how one might find the most suitable packings for this important application. Table 2 provides a list of the characteristics of the silicas tested in these experiments.  

Table 2: Bonded silicas and base silicas used in this study.			

Nowadays, dynamic axial compression columns are widely used for preparative and process separations. Since this type of column configuration puts additional stress on a packed bed, special attention has to be paid to the mechanical stability of the silica used. The mechanical strength of the base silica is further challenged by the high pressure of operation of the packed columns, quick pressure buildup during startup, and when the high pH washing step (described next) causes additional stress on the base particle.  

Figure 1: X-ray fluorescence sample compressor used for mechanical stability test.			

Two methods of testing the mechanical strength of the bare silica were used. These methods can be used by nearly everyone since the testing is straightforward and easily repeatable. Initially, the particle size distribution of the base silica gel is measured by a Coulter counter. Next, the silica particles are tightly packed into the ring of an X-ray fluorescence (XRF) spectrometer sample compressor (see Figure 1). The sample is pressed at 500 bar for 1 min. The ring is unpacked and the particle size distribution is measured again by the Coulter counter method (Figure 2). The value represented in the small table at the bottom of Figure 2 is the difference between the smaller particle percentage before and after the compression test. In this case, there was very little difference between the two tested silicas. 

Figure 2: Test 1, mechanical strength of base silicas. Plot shows the particle size distribution for silica SP-100-8-PK.			

A second method to measure the mechanical strength of the particle is the repeated packing and unpacking of the same silica into the same column configuration. Changes in bed length or back pressure with subsequent packings can indicate that the particles are becoming disturbed by the process of repeated packing. In this experiment, the slurry packing method was used. The packing pressure was 9.6 MPa using a mobile phase of isopropanol and flow rate of 150 mL/min. Table 3 shows the results of the back pressure changes for 1, 5, 10, 15, and 20 cycles of repeated packing and unpacking. For both silicas, back pressure (Table 3) and bed length (results not shown) were unaffected by the packing process and, thus, both tested silicas have very similar mechanical strengths under these conditions. Although not shown, scanning electron micrographs of the unpacked silica particles after the initial, 10th, and 20th cycles also revealed that no fines were being created.  

Table 3: Test 2: change in back pressure with repeated dynamic axial compression packing and unpacking.			

 As indicated above, in the production of insulin the use of sodium hydroxide solution is a requirement for cleaning in place (CIP) of the process column to get rid of fibrillation by-products. This requirement places severe constraints on the silica-base material as well as the bonded packing material. 

Silica gel itself is soluble in water; its solubility increases with temperature and pH, as depicted in Figure 3 (10). At low pH values, the base silica gel itself is fairly stable to dissolution. At neutral pH and at high pH, dissolution of the underlying silica gel through attack by the hydroxide ion causes a loss of column performance. The degradation process is enhanced by higher temperatures. Often, a column void is formed as the base silica gel is solubilized resulting in abnormally high pressure buildup (because of particulates building up on the column outlet frit), peak doublets, peak tailing, and other undesirable characteristics. To suppress the dissolution of silica in typically used mobile phases, we must prevent the hydroxide ion in the mobile phase from reaching the underlying silica.  

Figure 3: Solubility of silica gel in water as a function of pH.			

There are several methods to increase the base stability of silica gel and bonded silica gel. One approach to accomplish this blockage is to coat the silica with a polymer that can protect the underlying silica against dissolution, provided that the mobile phase doesn't penetrate the coating and begin the dissolution process underneath the polymer shell. The polymer coating may also lead to a loss of chromatographic efficiency since diffusion in polymeric layers is generally poorer than in silica monolayer bonded phases. For high-pH work, another approach that also delays the dissolution of silica is to cover the silica gel base with a high density of chemically bonded phase. With the bonded phase coverage, the rate of silica dissolution is determined by the bonding characteristics, type of bond, type of silica, presence of certain salts and buffers, and the buffer concentration, mobile phase pH, and other experimental parameters (11,12). Hybrid silica–organic materials offer yet another approach to prevent dissolution of the packing, especially at basic pH values (13).  

Another approach to increasing the alkaline durability of the base silica is to fabricate the silica to be more stable itself. Unfortunately, the procedures to make this happen are usually highly proprietary and are mostly treated by the silica companies as trade secrets. The manner of preparation, the specific surface area, the gel type, the gel time, the temperature, and the pH of the gelation step are all known procedures that affect silica solubility. Figure 4 shows the results of a study of the alkaline durability of four types of bare silica gels prepared by different methods. The tests were conducted using a 250 mm × 4.6 mm packed column with a 0.05 N sodium hydroxide solution running through it, a temperature of 25 °C, and the HPLC upper pressure limit set to 400 kg/cm

. The pressure was recorded as a function of the number of column volumes (one column volume equals 4.2 mL) that was passed through the column. For two of the silicas, less than 50 column volumes of the sodium hydroxide solution passed through the column before the pressure became catastrophic and flow through the column could no longer be achieved at the upper pressure limit. Two of the silicas showed respectable alkaline resistance and were clearly superior to the others.  

A side note: It was found in these tests that a bare silica with stronger alkaline resistance also makes a stronger bonded-phase packing material. 

Figure 4: Alkaline durability of base silicas.			

 Clearly, the addition of a bonded phase should help in lessening the attack of hydroxide ion on the base silica. Depending on the type of bonding and the bonding density (or coverage), the underlying surface is more protected than the bare silica used in the previous testing. In this set of experiments, C8 siloxane phases were studied on the earlier base silicas. The same flow experiment that was conducted with the bare silica was repeated, but instead a stronger solution of 0.1 N sodium hydroxide was used to accelerate the degradation test. The conditions for the experiment were as follows: 

Mobile phase:  30:70 (v/v) 0.1 N sodium hydroxide–ethanol 

The remaining experimental conditions were the same as outlined for the bare silica studies above. 

Figure 5 shows the experimental results obtained with C8 bonded phases. By covering the base silica with bonded phase, the stability of all silicas were improved over the bare silica lifetime (compare Figure 5 to Figure 4). In fact, the bonded silicas designated as SP-200-10-C8-BIO and SP-100-10-C8-PK showed remarkable stability in such a caustic environment. Undoubtedly, this improvement was a result of improved base silica stability coupled with improved bonding phase chemistry. 

Figure 5: Plot of the relative pressure drop showing the alkaline durability of C8 bonded silicas.			

 refers to the amount of sample which can be injected onto a column without overload. It may be expressed as a milligram of sample per gram of packing basis. 

 is defined as the sample mass injected where the column efficiency falls by 10% from its normal value. Higher loadability will result in better process economy and better productivity at production scale. However, loadability is only one part of the overall preparative requirement. Loadability combined with adequate separation selectivity (covered next) and optimized retention time will lead to the highest possible productivity. Loadability is governed by the surface area of the packing material or the bonded phase coverage. 

One approach to determine loadability is to use breakthrough volume or mass. Here, a known concentration of target analyte is continuously pumped through a packed column and absorbance (or other solute parameter) is monitored at the column exit. Conditions are chosen so that the compound of interest is totally retained by the column packing. After the active sites on the column are saturated, the compound breaks through and the absorbance jumps from the stable baseline; the total volume of liquid passed through the column can then be used to calculate the total mass on that compound that was retained on the column while the liquid was being pumped through.  

To measure relative phase loadability, several packings were subjected to the following conditions: 

Sample concentration: 10 mg/mL in 10:90:0.5 (v/v/v) acetonitrile–water–trifluoroacetic acid 

Figure 6 shows the breakthrough profiles for the compound of interest (insulin) on the four different bonded-phase silicas. Each of the bonded silicas has its own breakthrough value and one (SP-100-10-C8-HP) has a somewhat higher capacity than the others with the SP-200-10-C8-BIO being of much lower capacity, which is undoubtedly related to the lower surface area of the wider pore starting silica gel. 

Figure 6: Determination of bonded-phase breakthrough volumes (loadability). See the text for experimental conditions.			

 Separation selectivity (separation factor) is one aspect of preparative chromatography that affects compound purity. In both analytical and preparative work, selectivity is an important part of the resolution equation. It is imperative in the purification of a drug to be injected into a human body that the compound be of the highest purity possible. For insulin, a closely eluted impurity, des-amido insulin, is used to monitor the overall purity of insulin. In fact, according to the USP, the relative amount of des-amido insulin must not exceed 3% of the total amount of insulin plus des-amido insulin (14). Thus, a high separation factor between the pure insulin peak and the des-amido impurity peak indicates the quality of separation. The easiest and most reproducible way is to use standard pure insulin (reagent-grade) and create an artificial impurity by simply letting the water-dissolved insulin sit on the workbench overnight, which results in the formation of the des-amido form. This binary model of insulin–insulin impurity is an ideal model for measuring separation selectivity.  

To measure the separation selectivity, the following conditions were used: 

				 Column dimensions: 250 mm × 4.6 mm 

				 Mobile phase: 30:70:0.1 (v/v/v) acetonitrile–water–trifluoroacetic acid 

Column Watch

This year's symposium was held in New Orleans, Louisiana, USA. In this review, Ron Majors covers HPLC and UHPLC column and sample preparation highlights, summarizes the awards presented, and reviews the overall liquid-phase chromatographic trends.

Standardized Testing of Silica as a Base Material for Difficult Bonded-Phase Preparative Applications

The seven faux pas discussed here are common practices in high performance liquid chromatography (HPLC) that may no longer make sense because of improved technologies or other changes in modern HPLC. Therefore, careful consideration should be given to alternative approaches.

 is French in origin and literally means "false step", or "misstep". Although in English the phrase typically refers to an embarrassing social blunder rather than to a technical error, I am borrowing this expression, and stretching its meaning a bit, as a way to highlight some common practices in high performance liquid chromatography (HPLC) — legacy practices that have been in use for many years — that may no longer make sense because of improved technologies or other reasons in modern HPLC. In this instalment, I highlight seven such questionable practices and show why they should not be followed without first considering some alternative approaches:  									

						 Using columns packed with 5-µm particles

						 Using 4.6-mm i.d. columns at 1 mL/min

						 Preparing fresh reference standard solutions with every assay

						 Using stainless steel ferrules for column connections

As each topic is discussed, I will offer rationales and references to support why the practice is being called into question — and in many cases, I will offer exceptions to the rule.  

Using HPLC Columns Packed with 5-µm Particles

For many years, the "standard" HPLC analytical column has been a 250 mm × 4.6 mm column packed with 5-µm particles that could deliver ~20,000 plates (1,2). In a survey on column trends published in 2012, Ron Majors reported that the "standard" workhorse column was likely to be shifting towards a 150 mm × 4.6 mm column packed with 3-µm particles, which offers comparable efficiencies with a 40% or more reduction in analysis time (3). This trend is not surprising for experienced practitioners because plate count (

) is proportional to the ratio of column length (

. The 3-µm particle has proven to be as reliable as its 5-µm counterpart and offers higher performance (lower plate height [

]) while maintaining compatibility with conventional HPLC equipment (1,2). As ultrahigh-pressure liquid chromatography (UHPLC) and liquid chromatography–mass spectrometry (LC–MS) systems are becoming standard equipment in many laboratories, the "standard" column particle sizes are expected to trend downwards to sub-3-µm or sub-2-µm particles (4). Another emerging trend is the increased acceptance of superficially porous packings, which have improved particle morphology and substantially higher performance (~20–30%) compared with totally porous materials (5).  

 Although columns packed with 3-µm particles are generally superior to those packed with 5-µm materials, they do use finer inlet frits (0.5 µm versus 2 µm), which are more prone to clogging from "dirtier" samples. Another exception to this rule may come from specialized columns (for example, size-exclusion chromatography columns for protein analysis) where higher pressure and higher temperature from frictional heating may lead to increased on-column protein aggregation (6). 

The "standard" internal diameter for analytical HPLC columns has been 4.6 mm since the early 1970s and remains that way today (2,3). The reason for the popularity of this diameter might be the wide availability of stock 316 seamless, stainless steel ¼-in. o.d. tubing (which has a wall thickness of 0.035 in., yielding an internal diameter of 4.6 mm). With column hardware of this size, analysts typically used a 1.0-mL/min flow rate (

) as a default (a nice round number which is also the optimum flow rate for columns packed with 5-µm particles).  

Figure 1 shows the van Deemter curves of three columns (50 mm × 2.1 mm) packed with 5.0-, 3.5-, and 1.7-mm C18 particles (7). Note that the minimum plate height (

), is proportional to particle diameter (

 Figure 1: Van Deemter curves for 5-, 3.5-, and 1.7-Âµm C18 particles. Column: 50 Ã 2.1 mm; mobile phase: 50:50 (v/v) acetonitrileâwater; detection: UV absorbance at 210 nm; temperature: 45 Â°C; analyte: benzylphenone. Adapted with permission from reference 7.			

If we accept the new "default" particle size to be 3-µm (or 3.5-µm) as indicated above, then 

 would be ~3.0 mm/s or 0.4 mL/min for 2.1-mm i.d. columns (equivalent to ~2.0 mL/min for 4.6-mm i.d. columns). A 2.0 mL/min flow rate is less "green", and a 1-L reservoir of mobile phase would last for only ~8 h. However, if one selects a 3.0-mm i.d. column, then the optimum flow rate would be ~0.8 mL/min using geometrical scaling calculations (8,9) as shown below. 

The use of 3.0-mm columns reduces solvent consumption while conserving sample (smaller sample volume or concentration) and is also highly compatible with conventional HPLC systems typically used in a quality control (QC) setting. Column efficiency and the effect of extracolumn band broadening are usually not adversely impacted if a modern HPLC instrument with intermediate dispersion is used (variances < 20 µL

 ) (1,2,5). Many laboratories (including my own) have switched from 4.6 mm to 3.0 mm i.d. as a "preferred" column internal diameter for these reasons (10). The use of 2.1-mm i.d. columns is logically the next step, but requires low-dispersion UHPLC equipment. 

The use of smaller internal diameter columns can be advantageous to generate higher resolution under gradient conditions. A useful equation in this regard (1,2,11) is shown below. 

 or retention factor under gradient conditions; 

* value typically means higher resolution by allowing greater volumes of lower strength mobile phase to be pumped through the column (1,2). Equation 2 shows that 

* or resolution. So, the use of 3-mm columns is preferable in this regard versus 4.6-mm i.d. columns. Along the same lines, one useful application of this equation is the use of small columns (for example, 50 mm × 2.1 mm), short ballistic broad gradients (

 = 1 or 2 min) at high flow rates of 1.0–1.5 mL/min for quick, in-process testing or high-throughput screening analyses (7,12). 

It is a common practice to filter mobile phases through 0.5-µm membrane filters (nylon, polytetrafluoroethylene [PTFE], or cellulose acetate), using a vacuum to draw the solvent across the membrane to remove any particulate matter that may cause problems in the HPLC system or column (1,2). However, this may be redundant since all HPLC-grade solvents and water from purification systems have already been sufficiently prefiltered. An extra filtration step can actually be counterproductive by increasing the chances for chemical contamination, which can lead to more "ghost" peaks in the blank and sample chromatograms (2). By using high-purity reagents (such as ammonium formate at 99.995% purity), many laboratories have been operating well for years without a single vacuum filtration apparatus. Instead, they rely mainly on an annual preventive maintenance programme in which the internal filters in the HPLC equipment (in the purge valve and pump compartment) and the 10-µm solvent inlet filters in the mobile phase reservoirs are replaced.  

Some situations where mobile-phase filtering is still highly recommended (1,2) are when ion-pairing reagents, low purity buffers (for example, citrates and phosphate), or high salt content mobile phases (6 M sodium chloride) are used, particularly for UHPLC applications using columns packed with sub-3-µm particles (which are more prone to clogging problems). Another situation where extra filtration is warranted is when the quality of the water from purification systems is insufficient because of the quality of the source water or other reasons.  

For the analysis of acidic and basic analytes (that is, most pharmaceuticals), it is imperative to acidify or basify the mobile phase to control the ionization state of the analytes to maintain good peak shape (2). Simple mobile phases such as 0.1% (v/v) acid in water (for example, trifluoroacetic acid, formic acid, acetic acid, and phosphoric acid) may be sufficient in most cases and can be prepared quickly by pipetting 1 mL of the acid into 1 L of water. Similarly, 0.1% (v/v) ammonia in water works well for many high-pH-compatible columns. Additional filtration of these simple mobile phases is not generally needed as per the previous section. So, buffers have little utility if one uses a low silanophilic activity column with either low- or high-pH mobile phases, and for the injections of low sample mass prepared in mobile phase A as a diluent. 

 Buffered mobile phases may still be needed for critical purity analyses of complex molecules (for example, compounds with multiple ionizable nitrogen atoms and p

) and very complex mixtures to maintain tight selectivity of critical pairs, particularly in mid-pH mobile phases.  

In other cases, some highly basic analytes (such as amines) tend to display poor peak shapes using low ionic strength mobile phases (for example, 0.1% formic acid or acetic acid). In these applications, a new generation of columns designed for use with low ionic strength mobile phases (such as charged surface hybrids [CSH]) (4) appears to offer a more convenient alternative. Figure 2 shows comparative chromatograms of a test mix of seven drugs using columns packed with regular C18 versus CSH C18 with 0.1% formic acid in water. Note that considerable peak tailing is observed with basic analytes (imipramine and amitriptyline) on the regular C18 bonded phase (Figure 2[a]) with 0.1% formic acid (though less peak tailing are typically observed when used with a higher ionic strength buffer such as 20 mM ammonium formate at pH 3.7). However, the CSH columns used for the separations shown in Figures 2(b–d) produce excellent peak shapes for basic analytes because of the presence of positive surface charges. 

Figure 2: Comparative chromatograms on the separations of seven acidic, basic, and neutral drugs using (a) regular C18 and (bâd) charged surface hybrid C18, phenyl-hexyl, and fluoro-phenyl columns (all 50 mm Ã 2.1 mm, 1.7-Âµm). Mobile-phase A: 0.1% formic acid in water; mobile-phase B: acetonitrile; gradient: 5â95% B in 5 min; flow rate: 0.5 mL/min; detection: UV absorbance at 254 nm. Peaks: 1 = 1-pyrenesulphonic acid, 2 = flavone, 3 = imipramine, 4 = fenoprofen, 5 = amitriptyline, 6 = diclofenac, 7 = octanophenone. Adapted with permission from reference 17. 			

Preparing Fresh Reference Standard Solutions with Every Assay

It is a standard practice in pharmaceutical QC laboratories to prepare two reference standard solutions (or three in some European countries) by weighing out accurately 20–100 mg of a highly purified reference standard in separate volumetric flasks. These solutions are then analyzed to ensure their response factors are comparable (that is, < ±2%) (2,13). This is an excellent QC practice, albeit time-consuming and problematic for hygroscopic materials. 

A potentially workable idea is to prepare a set of reference standard solutions and save them for later use. For instance, one can prepare hundreds of HPLC vials of a qualified reference standard solution and keep them refrigerated or frozen for later assays. For regulatory testing, the stability (shelf-life) of these solutions in the chosen storage conditions must be verified and documented first.  

This idea is not really new as standard reference materials from the National Institute of Standards and Technology (NIST) and other commercial manufacturers have been available in pre-prepared, batch-processed ampoules or vials for many years. Interestingly, the use of pre-prepared standards is not widely practiced in pharmaceutical laboratories except for highly stable compounds, although many pharmaceuticals may have sufficient solution stability at low temperatures and under proper storage conditions (for example, filled vials in sealed freezer bags to prevent loss of water). By moving to this idea, more consistent potency assay data may be generated for long-term stability studies because all standard solutions would be derived from a single stock solution. A note of caution is needed here when operating under a regulatory environment where supporting documentation or validation is needed before any significant change of established procedures is implemented. 

Figure 3: A picture and a drawing of an HPLC vial containing an extra layer of liquid near the neck of the vial. This extra layer of liquid is easily formed by inverting the filled vial and is physically stable. Adapted with permission from reference 14.			

In 2011, we encountered some strange sporadic HPLC assay results that took us several weeks to resolve. The episode was reported by my research associate with a molecular biology degree who was trained to shake sample vials before placing them into the autosampler to ensure sample homogeneity. For two weeks, we were unable to explain to our formulator colleagues why some of their new prototype tablets appeared to be losing 20% of potency, for some sample vials. 

Fortunately, this story had a happy ending after the root cause was found. We eventually discovered that inverting sample vials could result in an extra film of liquid extract adhering to the vial septum because of the surface tension of the diluent. Ordinarily, this would not be a problem except that we were using a new model of UHPLC autosampler which drew an air gap from the headspace near the vial septum to segment the sample plug in the sampling needle. This resulted in an additional sample volume of ~0.6 µL (default air gap volume) being injected sporadically. After reporting this to the UHPLC manufacturer, the autosampler firmware was changed to draw the air gap above the vial, and we managed to publish a nice article in John Dolan's "LC Troubleshooting" column in

The lessons learned from this experience were to shake the HPLC vial with caution, watch out for the extra liquid film underneath the vial cap, and always keep your eyes open during troubleshooting situations. 

 Frozen samples that have been "thawed" will require vortexing or shaking to ensure sample homogeneity (15). 

Using Stainless Steel Ferrules for Column Connections

Service engineers from instrument companies typically install HPLC systems with stainless steel fittings and ferrules (for example, Swagelok, Waters, Parker-Hannifin, Rheodyne, Valco, and so on). While these kinds of fittings work well for most internal or external fluidic connections, they are not ideal for column connections that require frequent changes for two primary reasons: Firstly, stainless steel ferrules will only fit perfectly with the endfitting port into which they are originally "swaged", making preswaged connections noninterchangeable with different columns. Secondly, preswaged stainless fittings can reseal only 5–10 times. This is particularly significant with columns from some manufacturers that have very different endfittings and insert depths from other column brands. The simple solution is to use finger-tight polyetheretherketone (PEEK) fittings (Figure 4[a]), which are inexpensive and can seal well to 5000 psi for many different column types (16).  

Finding good fittings for UHPLC was problematic in the early days (from 2004 to 2006) until the availability of improved products in the marketplace increased. The ones we found most useful are shown in Figures 4(b)–4(f). Most of these fittings are rated to 20,000 psi by just finger-tightening (Figures 4[c], 4[d], and 4[e]) or finger tightening augmented by a quarter turn with a wrench (Figures 4[b] and 4[f]). Although these UHPLC fittings allow columns to be changed more easily and reliably (>10 make–break cycles), they also tend to be quite expensive. 

Figure 4: Modern HPLC and UHPLC fittings for column connections: (a) IDEX Health & Science (IH&S) one-piece PEEK fitting for HPLC (5000 psi); (b) IH&S VHP-320 (wrench tight to 25,000 psi); (c) IH&S-Upchurch VHP-320 with winged nut; (d) IH&S VHP-3200 finger-tight fitting (25,000 psi); (e) Thermo Scientific Viper (20,000 psi); and (f) Optimize Technologies OPTI-LOK (hand-tight to 9000 psi and wrench tight to 20,000 psi). 			

In this column instalment, I have highlighted seven common "faux pas" or inefficient practices that should not be followed without considering alternative approaches that can offer improved chromatographic results, lower operational costs, or faster run times. Some of these alternatives may have already been adopted by experienced chromatographers, although most are probably not widely practiced. Others (not filtering mobile phases, not using buffered mobile phases, and not preparing two fresh standard solutions with every assay) may seem controversial, but can yield significant benefits and simplification if applied properly. Instead of working harder, the analytical scientist should work smarter, by questioning these outdated traditional practices, which in the context of modern HPLC can be considered "faux pas".  

Perspectives in Modern HPLC

Safe and effective gas cylinder setup and use requires the right pressure regulators, fittings, tubing, and filters as well as careful attention to procedure. This month's instalment reviews how to get the best results from laboratory gas cylinders, how pressure regulators work, and what filters should be installed in gas lines.

Seven Common Faux Pas in Modern HPLC (Slideshow)

Green analytical chemistry is a widely recognized concept that has led to the development of new analytical methods with reduced environmental impact and minimized analyst occupational exposure. Achievements include the development of microextraction, ultrasound-assisted extraction (UAE), and microwave-assisted extraction (MAE) techniques. Research towards greener separation processes focuses on the elimination of toxic solvents in liquid chromatography (LC) and the reduction of separation time. The recent developments in the labelling and categorization of analytical procedures are also presented. The National Environmental Methods Index (NEMI), Eco-Scale, and grouping with multivariate statistics are discussed together with their advantages and limitations. This article presents the most recent progress in the development of greener sample preparation and chromatographic separation techniques. 

Green analytical chemistry is now a widely recognized term and is defined in the literature as clean, benign, or environmentally friendly analytical chemistry (1). Green analytical chemistry means performing reliable analysis while applying quality assurance/quality control (QA/QC) and minimizing any environmental impacts resulting from the analysis. When one thinks of chromatographic procedures that meet the standards of green analytical chemistry principles, green chromatography comes to mind first (2). Chromatographic procedures are usually a series of operations that involve sample collection, transportation, preparation, proper chromatographic separation, and analysis. Greening chromatography concerns all stages of analysis, but the steps with the greatest environmental impact are sample preparation and the chromatographic separation. These stages are considered to be the most environmentally unfriendly and therefore have a lot of potential for "greening" (3). 

Green chromatography originates from the 12 green chemistry principles developed to minimize the environmental impact of chemical synthesis. When compared to chemical synthesis methods, the amount of pollutant waste products generated during chromatographic analysis appears small. However, it has been estimated that one liquid chromatograph can generate one litre of liquid waste daily (2). Three years ago, a series of three articles were published in

addressing the different aspects of green chromatography. The first article provided an introduction to green chromatography, describing its origin and the potential of green liquid chromatography (LC) (4); the second article proved that gas chromatography (GC) is often beneficial compared to LC (5); and the third article reviewed the importance of sample preparation before chromatographic analysis and how green chromatography principles could be applied (6). This article summarizes the achievements and developments over the last three years that have advanced the field of green chromatography. 

In a chromatographic analysis, sample preparation has the most detrimental effect on the environment and the question of whether or not it is really needed should always be asked. Unfortunately in most cases the answer is "yes" but there are chromatographic procedures that can omit this procedural step, as reviewed in reference 7. Direct procedures can be used to analyze samples with simple matrices such as water, petroleum products, and alcoholic beverages. Such approaches reduce environmental impact as well as saving money. 

Figure 1 presents the options available to implement green sample preparation including miniaturization (8), solvent elimination (9), and application of extraction assisting or enhancing agents (such as ultrasounds or microwaves) (10). It can be difficult to automate microextraction techniques; however, advances have been made including the development of dispersive liquid–liquid microextraction (DLLME); liquid-phase microextraction (LPME), and single drop microextraction (SDME) (11).  

 Figure 1: Solutions in sample preparation that meet green chromatography requirements.			

In recent years, a lot of work has focused on new solutions and applications that improve the "greenness" of sample preparation techniques. One technique that has been intensively worked on is DLLME. DLLME utilizes a tiny amount of nonpolar solvent (usually below 100 µL) and approximately 1 mL of less toxic dispersive solvent. Procedures based on DLLME have been proposed to determine UV filters (12), antibiotics (13), polycyclic aromatic hydrocarbons (PAHs) (14), or oestrogens (15) in water samples; and pesticides in honey, after sample dissolution in water (16). 

Performing DLLME extraction can be assisted with ultrasounds that eliminate or reduce the need for disperser solvent. The application of dispersive solvent increases the solubility of the extraction solvent. Without the dispersive solvent, extraction solvent volume can be reduced, because less of it is dissolved in water; and despite low initial solvent volume it is easy to recover enough extraction solvent volume to handle. 

For example, ultrasound-assisted extraction with 13 µL of chlorobenzene can be applied to determine siloxanes in water samples (17). In LC analysis, ionic liquids are considered as green solvents and can be applied as extraction agents in DLLME.  

An alternative solution to DLLME is solidification of floating organic drop (SFO). The extraction solvents used in SFO are higher alcohols such as nonanol, decanol, or undecanol. After extraction, the sample is subjected to a lower temperature and the solvent solidifies and can be easily mechanically separated from the sample matrix. The technique can be used in sample preparation before GC and LC analysis. It is suitable for the extraction of compounds with log k

 in the range 3–7 (18). For example: Triazines in water samples can be determined using 10 µL of 1-undecanol and 100 µL of acetonitrile as dispersive solvent (19). A similar procedure was designed for the determination of amphetamines in urine samples (20), and for the determination of pesticides in vegetables and fruits after extraction with acetone (21). In 2013, Dong 

. applied DLLME to the analysis of soil samples (22). The authors extracted phytocides from soil with a water–methanol mixture and evaporated to dryness before the residue was redissolved in methanol. The samples were subjected to ionic liquid-based ultrasound-assisted DLLME before high performance liquid chromatography (HPLC) analysis. 

A derivatization step can be easily combined with DLLME. It is particularly practical when the derivatizing agent easily hydrolyzes in the water sample (23). In 2014, Casado 

. performed acetylation with simultaneous extraction of benzotriazoles on a mixture of microvolumes of acetic anhydride, acetonitrile, and toluene (24). In 2013, Farajzadeh 

. applied butylchloroformate as a derivatizing agent in DLLME for the determination of amantadine, a pharmaceutical prescribed for influenza treatment, in human plasma samples (25). 

The main trends in solid-phase microextraction (SPME) are automation, 

 applications, and the development of new coatings (non-bonded, bonded, or cross-linked polymers) (26). Polydimethylsiloxane (PDMS) has traditionally been used as a SPME fibre coating, but fibres based on polyacrylate or polyglycol polymers are now available for the extraction of polar analytes. The support for coatings has traditionally been fused-silica, but recently metal-based fibres have been developed for increased durability and improved mechanical properties. Increasing the coating film thickness to increase fibre capacity and therefore sensitivity is another trend; however, it can increase the extraction time required to obtain equilibration (27). 

Ionic liquids have recently been demonstrated as fibre coatings, opening up the possibility to design and modify coatings to obtain greater selectivity towards analytes (28). In 2012, Feng 

. applied a polymerized ionic liquid 1-vinyl-3-octylimidazolium bromide as a coating in SPME to isolate alcohols from water (even methanol) and produced good recoveries (29). In 2012, Ho 

. extracted chlorophenols from water with 1-butyl-3-methylimidazolium hexafluorophosphate-coated SPME fibre (30). 

Novel applications of SPME include the determination of contaminants or bioactive compounds directly in tissues of living organisms, or in their habitat, as demonstrated by Bessonneau 

 SPME sampling has also been developed to determine emerging contaminants in fish (32), and volatile halogenated organic compounds in trees (33), insects, higher animals, and even humans (34). 

Stir-bar sorbent extraction (SBSE) is another rapidly developing technique, where the majority of applications use PDMS coatings. Recently, alternative polar coatings have been introduced and intensively investigated for the extraction of analytes of a wide range of polarities including polyacrylate and polyglycol coatings (35). For the extraction of chlorophenols and chloroanizoles from wine samples, ethylene glycol with silicone in the form of a copolymer was found to be more effective than a polyacrylate coating (36). This is because it allows for simultaneous extraction and preconcentration of both polar phenols and nonpolar chloroanisoles. Another coating for the extraction of polar compounds from water samples is methacrylic acid and divinyl benzene copolymer (37). This SBSE coating gives much better recoveries (around 100% for the most of investigated compounds) for polar pharmaceuticals than commercial PDMS coating-based stir bar (less than 20%).  

The stir bar has also been applied as a passive sampler for the 

 extraction of nitroorganic pesticides (38). Such an approach seems to be a promising technique for the collection of integrated samples. The 

exposition and sample preparation steps are both straightforward. Selection of the most appropriate stir bar coating can result in sampling of polar analytes, which are usually not collected with more traditional passive samplers. 

Why should you be concerned about mobile-phase degassing - it's all done automatically, isn't it?

LCGC Europe-08-01-2014

How Much Retention Time Variation is Normal?

World of Technology & Science (WOTS) 2014

Standardized Testing of Silica as a Base Material for Difficult Bonded-Phase Preparative Applications

Seven Common Faux Pas in Modern HPLC (Slideshow)

Developments in Green Chromatography

Vol 27 No 8 LCGC Europe August 2014 Regular Issue PDF