"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 John.Dolan@LCResources.com

When you want to adjust a United States Pharmacopeia (USP) method for a different size column or to meet system suitability criteria that fail, how much of a change can you make without revalidating the method?

	A colleague recently asked me to write an update in “LC Troubleshooting” discussing the current method adjustment guidelines from the United States Pharmacopeial Convention. This is a topic that is high on the interest scale whenever I talk to scientists doing liquid chromatography (LC) in the pharmaceutical industry. The guidelines in the present discussion are contained in Chapter 621 (abbreviated by the United States Pharmacopeia [USP] and here as <621>) of the USP (1). The USP is updated at least twice a year, so it is wise to check the most recent edition for any changes. Here I’m using United States Pharmacopeia40–National Formulary 35 (USP 40–NF 35) (1), which became official on 1 May 2017.

	First, let me address a common misconception that I frequently encounter. USP <621> has become a de facto standard as rules for adjusting chromatographic methods, and as such, many users think this means all methods. In fact, <621> applies only to monograph methods published in the USP. This means that it does not apply to methods that you’ve developed and validated in your laboratory, obtained from the scientific literature, or inherited from somewhere else. With all that said, most of the guidelines make sense as a basis of method adjustment for many other types of methods. For example, you might adapt them as a basis for your own standard operating procedure (SOP) that you will use to govern how you deal with method adjustment in your laboratory-but at that point, they are your guidelines, not the United States Pharmacopeial Convention’s. Finally, in the present discussion, interpretation of the guidelines is based on my opinion, not an official opinion by the United States Pharmacopeial Convention or anyone else.

	One key to reliable operation of any LC method is to have a good system suitability test. This test will help to verify that the entire method is working well enough to produce analytical results with acceptable precision and accuracy. Typically, characteristics such as retention time, column efficiency, resolution, peak tailing, detector response, precision, and accuracy are assessed to some degree during system suitability testing. So it is not surprising that the USP has a strong focus on system suitability (1):

Adjustments to the specified chromatographic system may be necessary in order to meet system suitability requirements. Adjustments to chromatographic systems performed in order to comply with system suitability requirements are not to be made in order to compensate for column failure or system malfunctions. Adjustments are permitted only when . . . adjustments or column change yields a chromatogram that meets all the system suitability requirements specified in the official procedure. 

	The way I read this is that if the adjustment is within the recommended range and after the adjustment, the system suitability passes, it is considered an adjustment. All I need to do is document the adjustment (and meet any of my company’s internal requirements) to continue using the method. A revalidation is not required. If I go beyond the adjustment limits, it is considered a method modification or change and will require some level of revalidation.

	Next, let’s go through the various adjustments listed in USP <621>. Some of these adjustments can be made for either isocratic or gradient methods, whereas others are not universal. I have summarized the adjustments in Table 1. Table 1 is best interpreted in conjunction with the discussion below, which considers the nuances of the adjustment. Although I cannot find a specific statement to this effect, the variations in Table 1 make sense for reversed-phase separations, and I assume that that is the intent.

	Table 1 shows that the allowed adjustment range for the pH of the mobile phase buffer is ±0.2 units. At first glance, this seems like a reasonable allowance. After all, most laboratories consider normal laboratory variation of ±0.05–0.1 pH units when a pH meter is used. So twice the normal variation should be fine. For example, a method that calls for a nominal pH of 2.5 could be adjusted over the range 2.3 ≤ pH ≤ 2.7. Be careful about applying these guidelines without paying close attention-I have a chromatogram in my collection in which a well-resolved peak pair degrades to two barely distinguishable bumps with a pH change of 0.1 units. For other cautions about pH adjustment refer to the recent “LC Troubleshooting” instalment on pH (2).

	The USP guidance allows a ±10% change in buffer concentration. I have no problem with this, but I doubt if you’ll see any change in a reversed-phase method with this small a change, unless the method has either insufficient buffer now or is near the saturation point. Recall the discussion of orthogonal leverage associated with table 1 of last month’s instalment (3) where it was shown that a twofold change in buffer concentration was unlikely to change selectivity under reversed-phase conditions. I suspect that a method with a buffer concentration of 25 mM could be adjusted over a range of 10–50 mM without significant change in the chromatogram. The ±10% restriction doesn’t make much sense to me for reversed-phase LC. However, buffer concentration can be a significant player when ionic interactions are important, such as in ion-exchange chromatography or hydrophilic-interaction chromatography (HILIC), so we can’t conclude that buffer concentration will never change a chromatographic separation.

	The instructions in Table 1 for mobileâphase composition may seem a bit confusing: ±30% relative, but not more than ±10% absolute change in the minor components of the mobile phase. A couple of examples should clarify things. First consider a mobile phase of 50:50 A–B, where A is the aqueous portion (buffer or water) and B is the organic (generally acetonitrile or methanol)- 30% of 50% is 15%, but this is more than 10%, so we are limited to a 10% change in solvent concentration. Thus, we’d be allowed to go from 40:60 A–B to 60:40 A–B. This is an easy calculation, but the change seems a bit extreme to me-when was the last time you had a method for which the mobile phase could be changed by ±10% acetonitrile and still work? Remember the “Rule of Three”, from the recent discussion on retention (4), where a 10% change in mobileâphase organic can change the retention factor (or retention time for well-retained peaks) by about threefold. So be careful when applying the USP guidelines for mobileâphase adjustment. 

	What about a case where there is a large difference in concentrations of A and B; for example, 5% buffer and 95% acetonitrile? In this case, 30% of the buffer concentration is 1.5%, well below the ±10% limit. Now the allowable range would be 3.5:96.5 to 6.5:93.5 A–B. This seems like a reasonable allowance. 

	The calculations get a bit more complicated for a ternary mobile phase: for example, with a mobile phase consisting of 35:5:60 A–B–C, where C is a second organic solvent. In this example, 30% of 35% is 10.5%, so we would be limited to a 10% change in A. We would be allowed the same 1.5% adjustment calculated above for B. The allowable adjustment would be any combination of 35 ± 10% of A, 5 ± 1.5% of B, and the remainder C. You can see that a considerable amount of variation is allowed-once again, be cautious with such changes.

	The guideline for detector wavelength is a bit puzzling. There is no allowance for changing the detector wavelength, but if a second detector is out of calibration by up to 3 nm, it can be used. So if I don’t like the wavelength, I find a detector that isn’t working properly and use it? No, no, no! This adjustment note is a dinosaur from the days when detector calibration was a regular problem. Most of today’s ultraviolet (UV) detectors do an automatic calibration check when they are turned on and often self-calibrate as part of the process. I haven’t seen a UV detector calibration problem for 20 years, and that was probably with a detector that had been dropped or otherwise abused.

	One of the areas where the USP has made the biggest changes to improve user flexibility is in the allowed columnârelated changes. As recently as 2012 (USP 35–NF 30), the allowed variations were quite limited. For example, you could change the column length, L, by +70%, which seems quite generous. You could change from a 150-mm column to a 250-mm one (250/150 = +67%) or from 150 mm to 100 mm (-33%) or 50 mm (-67%), which seems fine. You also could reduce the particle size, dp, by 50%, but were not allowed to increase it. So you could go from a 5-µm dp particle to a 3-µm one (-40%), but not much smaller. The big flaw in these recommendations is that they ignore the influence of column length or particle size on the column plate number, and thus resolution. They also would not allow scaling a method from a 5-µm particle column to an ultrahighâpressure LC (UHPLC) column with ≤2-µm particles or to scale up a UHPLC method to perhaps a more robust 5-µm particle for routine work. So, although these allowances were used for many years and did allow some very practical changes (such as the transition from older 250-mm, 10-µm columns to the more standard 150-mm, 5-µm columns that are so popular today), they weren’t as appropriate for today’s laboratory environment.

	Now, however, there is a more flexible allowance-and one that has a much sounder scientific basis. The central focus is on keeping the column plate number, and thus resolution, fairly constant. Because the plate number is a function of the length of the column divided by the particle diameter, the L/dp ratio is the key factor here. The column length and particle diameter can be changed as long as L/dp is constant; an allowed variation in this result is from -25% to +50%, which makes sense because there are a limited number of discrete column lengths and particle sizes commercially available.

	Table 2 comes from the current USP version (1) and contains examples of some of the changes that can be made. Many of the USP monograph methods are rather old and specify a 250 mm × 4.6 mm, 10-µm dp column (L/dp = 25,000). You can easily update the method to a 150 mm × 4.6 mm, 5-µm dp column (L/dp = 30,000) for an increase in L/dp of 20%, which is within the limits. Alternatively, if you favour smaller particles, you could use a 100 mm × 4.6 mm, 3-µm column (L/dp = 33,300; note that I’m rounding numbers for presentation), a 33% increase. You could even move the method to UHPLC and use a 50-mm column packed with 1.7-µm particles (L/dp = 29,400) and still stay within the limits. All these columns give approximately the same plate number and thus the same separation. This assumes all the columns have the same chemistry (same bonded phase, likely from the same manufacturer, and same brand of packing). However, don’t be too worried about chemistry changes, because to be acceptable, the method still has to pass system suitability; any unacceptable chemistry changes will cause system suitability to fail.

	The L/dp approach works very well if the same particle type is used, most commonly totally porous particles (TPPs). However, the technique can fall apart when switching from TPPs to superficially porous particles (SPPs) that are becoming increasingly popular. SPPs typically give plate numbers corresponding to much smaller particles, so the particle size can be misleading if the L/dp approach is used. For example, a 2.7-µm dp SPP column has the back pressure of a ~3-µm dp TPP column, but a plate number that is more like that of a ~1.8-µm dp TPP column. So in the above example, a 50-mm, 2.7-µm SPP column would appear to have an L/dp = 18,500, which is a drop of 26% if we’d started with the 250-mm, 10-µm column, but nearly 40% lower than the 150-mm, 5-µm column. It doesn’t make scientific sense to reject the SPP column based on the L/dp result; instead there is an optional allowance that the plate number, N, should be constant within the same -25% to +50% range. Now the SPP column would be an acceptable substitute (again, assuming system suitability passes).

	The column diameter, dc, can be changed as long as the flow rate, F, is adjusted so that the linear velocity of the mobile phase stays the same. In theory, the linear velocity for maximum column efficiency increases inversely with a change in particle diameter. When linear velocity is adjusted for particle diameter, it is best to refer to the reduced velocity. Most of us don’t worry about this additional adjustment, but it is included in equation 1 (1), which will make the required adjustments straightforward. In addition to the changes allowed according to equation 1, the flow rate may be adjusted by up to ±50%.

	F2 = F1 × [(dc22 × dp1) / (dc12 × dp2)]         [1]

	where the subscripts identify the variables for the original column, 1, and the new column, 2.

	Examples in Table 2 show the flow rate relative to an initial flow rate with a 150 mm × 4.6 mm, 5-µm column. So you can see that equation 1 would suggest that you operate a 250 mm × 4.6 mm, 10-µm column at half the flow rate of the 150-mm column. If we assumed an initial flow rate of 1.0 mL/min, we would lower the flow rate to 0.5 mL/min to have the same reduced velocity. However, the combination of the lower flow rate and longer column may make the run unacceptably long, so most of us would keep the flow rate at 1 mL/min (allowed in the additional adjustment of ±50%) for a faster run and reasonable pressure. Table 2 also gives estimates of the pressure and run time for the new column relative to the original one.

There is additional fine print in the USP discussion for changes in particle diameter that allows further adjustment of the flow rate when moving from traditional LC conditions (≥3 µm dp) to UHPLC (<3 µm dp) or vice versa if the plate number doesn’t drop by more than 20%. This allows for flexibility with UHPLC that might be otherwise restricted, as long as system suitability passes.

	A final note here is that adjustments to column dimensions, particle size, and flow rate are allowed only for isocratic methods. These changes are specifically not allowed for gradient methods. Although there are techniques for properly adjusting gradient methods for such changes, they are not included in the current version of the USP, so you’ll have to do some amount of revalidation if you decide to adjust gradient methods.

	Injection volume can be increased or decreased as long as method performance does not suffer. Be watchful for excessive band broadening and retention time shifts if you increase the injection volume significantly. Make sure you have sufficient signal to provide acceptable precision and accuracy if you reduce the injection volume. A few injections made above and below the proposed new injection volume should help you demonstrate (and document) the robustness and suitability of the change.

	Column temperature can be changed by ±10 °C. Remember, however, that isocratic retention times are reduced by ~2% for each 1 °C change in temperature. Selectivity can change with temperature changes, especially if ionizable compounds are present in the sample. So be careful if you change the column temperature that you do not compromise the separation of critical peaks in your sample.

	We’ve seen that the USP offers reasonable guidelines for the adjustment of LC methods. These allowances apply to isocratic methods, but may be prohibited or not recommended for gradient methods. I’ll circle back to what I stated at the beginning: USP guidelines are intended only for the adjustment of monograph methods included in the USP. The guidelines are not assumed to apply to non-USP methods, so they don’t give you permission to change other methods without complying with the appropriate regulations. Lastly, many companies have their own interpretation of the USP and other regulatory documents, so be sure to consult your internal SOPs and other regulatory guidance when you decide to adjust an LC method. Oh, yes, remember when you make adjustments or changes, “if it isn’t documented, it didn’t happen”, so keep good records.

		General Chapter <621> “Chromatography” in United States Pharmacopeia 40 National Formulary 35 (USP 40–NF 35, United States Pharmacopeial Convention, Rockville, Maryland, 2017), pp. 508–520.

 has been writing “LC Troubleshooting” for LCGC for more than 30 years. One of the industry’s most respected professionals, John is currently a principal instructor for LC Resources in McMinnville, Oregon, USA. He is also a member of LCGC Asia Pacific’s editorial advisory board. Direct correspondence about this column via e-mail to LCGCedit@ubm.com

LC Troubleshooting

Method Adjustment the USP Way

 Department of Instrumental Analysis and Environmental Chemistry, IQOG-CISC, Madrid, Spain,
	

 Agilent Technologies, Wilmington, Delaware, USA

The quantitative extraction and subsequent purification of trace contaminants from (semi-)solid environmental and food matrices of regular size (that is, a few grams) is still recognized as a challenging task, typically accomplished through relatively complex off-line multistep treatment procedures. When these conventional sample preparation procedures are applied to the treatment of size-limited samples (of less than 1 g), the difficulties increase. This review discusses the different analytical strategies that can be adopted to overcome (or at least reduce) these difficulties when chromatographic techniques are involved for final instrumental determination.

	The analysis of minor organic components in solid and semisolid environmental samples and foodstuffs is typically regarded as a challenging task. This consideration derives from the usually complex and heterogeneous nature of these matrices and the variety of mechanisms governing the retention of trace analytes on them, which range from adsorption or absorption to integration of the target compound into the matrix structure. Whatever the strength of the retention, accurate determination of residual organic analytes by direct sample analysis is in most cases virtually impossible, making some level of sample preparation before instrumental compound determination mandatory. The required level of sample pretreatment (understood here as analytical operations performed for analyte extraction, purification, and concentration) will depend on the selectivity and sensitivity offered by the technique selected for final separation and detection of the analyte and, indeed, on the criteria of the analyst. Nowadays, the analytical determination of residues of legacy and emerging organic micropollutants of medium and high volatility is usually accomplished by gas chromatography (GC) coupled to highly selective and sensitive detectors. Although detectors, including microelectron capture detectors (µECDs), nitrogen or phosphorous detectors (NDs or PDs), and many others, are still used in many control laboratories, the benchmark for these determinations lies with the modern mass spectrometry (MS)-based detectors. These instruments allow the simultaneous accurate determination of a significantly large number of pollutants in relatively complex extracts within a single chromatographic run. The enhanced selectivity offered by the modern hyphenated and hybrid MS detectors improves the signal-to-noise (S/N) ratios and reduces interferences. The former ultimately results in lower limits of detection (LODs) and allows accurate analyte determination at concentration levels much lower than expected only a decade ago. 

	Conventional sample preparation protocols used for the analysis of trace organic pollutants in (semi-)solid matrices involve the exhaustive (and, in most instances, nonselective) extraction of the target compounds from the matrix structure. This is followed by the thorough purification of the obtained complex extract and the final concentration of the analytes of interest before instrumental analysis. In general, integration among these several treatment steps, and particularly their automation, is very limited. This means that the methods to be considered are limited in terms of analysis response and sample throughput, as well as prone to analyte loss or contamination because of the constant manual manipulation of the sample and extracts. The enhanced detectability provided by modern GC–MS systems used for these types of determinations means that, in most instances, only a small fraction of, typically, 1–2 µL out of the final 500–1000âµL concentrated extract, is used for analyte instrumental determination. In principle, this means that accurate analyte detection and quantification should be possible, even if the initial amount of sample is reduced from the typical 2–20 g to a representativeâsize sample of a few grams (0.5–5 g). In addition, the volume of the final purified extract can be proportionally reduced (for example, to 150–300 µL) and large-volume injection (LVI) can be used to ensure that a similar amount of analyte to that involved in the conventional largeâscale procedure is injected into the instrument.

	This principle is the base for a plethora of widely accepted miniaturized analytical techniques developed during the last two decades for the treatment of liquid and viscous samples. Some of these techniques, such as miniaturized on-line solidâphase extraction (SPE), solid-phase microextraction (SPME), and stir-bar sorptive extraction (SBSE), have been implemented in control and routine laboratories and have demonstrated their performance for the analysis of both major and trace components. Other liquid-phase-based microextraction techniques, in particular hollow fibreâprotected solvent microextraction, in both its two or three-phase formats, and dispersive liquid–liquid microextraction (DLLME), are rapidly increasing in popularity and practical application range. In contrast, the application of such an approach for the treatment of (semi-)solid samples has been much more limited in the specialized literature, where in many instances it has been limited to the academic demonstration of the proof of concept, and it is essentially nonexistent in commercial laboratories. Several reasons are responsible for this different level of development and acceptance, and the most relevant are identified and discussed in this article. Whatever the reasons, the most immediate consequence is that, even today, sample preparation procedures used in most laboratories for the treatment of (semi-)solid samples are well established and validated protocols that are applied as described for conventional-size samples, irrespective of the actual initial sample amount. In our opinion, this approach is probably far from being the best option from both a practical and an analytical point of view, and the development of more sizeâsuited procedures would be preferable. 

	This article discusses the advantages and shortcomings of miniaturization for the treatment of size-limited solid and (semi-)solid samples. The practicality of this approach will be discussed on the base of representative application examples dealing with the analysis of minor (that is, trace) organic pollutants because of the greater difficulty typically associated with this type of determination. The focus will be on those pollutants with medium and high volatility that will involve GC-based systems for instrumental determination, although gaseous compounds will not be considered because their extraction is typically accomplished using different extraction techniques. Nevertheless, if relevant, examples involving other types of separation techniques, such as liquid chromatography (LC), will also be considered. In all cases, applications and techniques dealing with the analysis of real-life (that is, nonspiked) samples will be preferred.

Miniaturized Sample Preparation: Advantages and Limitations

	As previously indicated, the miniaturized treatment of liquid and viscous (including slurry-like) matrices can now be considered an achieved goal. The widespread use and implementation of a number of these well-accepted and established extraction techniques in both research and control laboratories supports this statement. The success of techniques such as SPME, on-line SPE, SBSE, and the more recently introduced DLLME or disposable pipette extraction (DPX, a miniaturized version of dispersiveâSPE), relies, at least partially, on the several advantages derived from their miniaturized nature. Most of these techniques allow a significant reduction (and in some cases, virtual elimination) of the use of organic solvents and reagents compared with the corresponding largeâscale counterparts. In many instances, they also allow the completion of the sample preparation protocol within a shorter time frame by significantly reducing (or avoiding) the need for a concentration step in between treatment steps. The elimination of this need has also facilitated the hyphenation and integration among the several treatments and techniques involved in the analytical process. Hyphenated or not, the miniaturization of the analytical processes has been recognized as a mandatory step when trying to set up (at least partially) integrated and (semi-)automated systems contributing to increased sample throughput and minimized waste generation and energy consumption. Moreover, these methods have demonstrated equally satisfactory analytical results to conventional (that is, large-scale) methods, even for complex determinations, such as the analysis of trace micropollutants in influents of wastewater treatment plants and river waters (1). 

	Equivalent application examples described in the literature for the miniaturized treatment of solid samples reported similar benefits to those stated from liquid matrices, viz. reduced solvent and reagents consumption, improved capability of coupling between the extraction step and the subsequent treatments, the possibility to yield readyâforâanalysis extracts, less waste generation, and improved analytical response as a result of the shortened analytical time (2). As for liquid samples, experimental results have demonstrated that the reported miniaturized protocols can achieve equally satisfactory analytical performances to their equivalent large-scale protocols, even for difficult determinations, such as residues of polyaromatic hydrocarbons (PAHs) in sediments, polychlorinated biphenyls (PCBs) in fatty animal tissues, or pesticides in single insects (see Table 1).

Despite the many positive features and advantages shared by the miniaturized treatment of liquid and solid samples, the sharp differences between the levels of development and general acceptance reached by liquid samples compared to solid samples makes it clear that significant differences exist between these two application areas. Several aspects can be considered in this respect, but probably the most relevant would be: (i) the natural reticence of researchers to work with small heterogeneous samples, and (ii) the lack of appropriate commercial miniaturized instrumentation required to implement some of these methods. Regarding the former aspect, we should accept that the heterogeneity of some (semi-)solid matrices can make the collection of a representative subsample difficult. In such cases, we tend to assume that the larger the subsample analyzed, the more representative the analytical result. However, that is not necessarily the case. The representativeness of the subsample analyzed can only be guaranteed by the application of appropriate sampling protocols, which is out of the scope of this discussion. Whatever the sampling protocol applied, and assuming that a representative subsample is available for analysis, protocols for the treatment of (semi-)solid environmental and foodstuffs need to include a number of sample pretreatments prior to analyte extraction. In the case of trace pollutants, these pretreatment operations must include agglomerates disaggregation, sample chopping and mixing, water and humidity removal (for example, by freeze drying in the case of foodstuffs), grinding of the dried sample, and, in some cases, sieving to an appropriate sample size particle (for example, in the case of soils and sediments). The aim is to increase the homogeneity of the sample subjected to analysis and to ensure proper contact among the matrix components and the extraction solvents and reagents during the extraction step. When done properly, the enhanced homogeneity should also enable smaller fractions that are equally representative of the selected subsample to be defined. In other words, this approach should make the analysis of smaller fractions of the initial sample possible without compromising the representativeness of the final results. For obvious reasons, the final size of these smaller fractions is not expected to be the same for all matrices. The minimum amount of sample required for the analysis is determined by the heterogeneity of the original sample, the nature and effectiveness of the pretreatment operations performed, and the sensitivity provided by the instrumental step, all carefully determined by the analyst. Nevertheless, such considerations do not apply in size-limited samples, that is, in those in which the initial amount of sample available is determined by the sample nature (for example, the analysis of single insects) or the sampling procedure (biopsies). In these cases, direct application of the conventional methodologies results in an extremely high analyte dilution, which significantly increases the risk of analyte loss or contamination during sample treatment. Another undesirable consequence is the significant increase of the blank values when compared to those of the target analyte-something that can negatively affect the accuracy of the determination. In our opinion, for these types of samples, the analytical method should preferably be adapted to the sample characteristics, which makes miniaturization one of the better-if not the best-analytical alternatives. 

	A second identifiable aspect hampering progress in miniaturized preparation of solid matrices is the limited commercial miniaturized instrumentation adapted for such methods and, in particular, for sample extraction. While some of the miniaturized techniques used for the treatment of liquids can be applied for analyte purification, the number of instruments really adapted for the extraction of small-size solid samples is rather limited. In this context, strategies for the in-house scaling-down of the methods and the setting-up of homemade size-adapted instruments have been adopted with satisfactory results. The advantages and shortcomings of these different analytical strategies will be discussed next through illustrative application studies.

Analytical Techniques for the Extraction of Size-Limited Samples

	When considering what techniques or strategies to use for size-limited samples, one has to take into account what techniques are easily scaled down in the laboratory from methods that are used for samples without size limitations, as well as what technologies are commercially available that can be used. Table 1 shows an overview of selected applications involving a variety of miniaturized methods used for the treatment of size-limited solid samples.

	Manual Methods that Can be Scaled Down: Soxhlet extraction has been used for many years, and is often used for large samples with corresponding copious amounts of solvents because this configuration is so common. It has been reported that Soxhlet was used without any modification for the treatment of small-size samples (11), despite the final analyte dilution. However, one can purchase much smaller glassware to perform Soxhlet extractions on samples of less than a gram in size (3). With smaller samples, the long times typically required for extractions can be shortened. As for any other method development and validation, optimization should preferably be performed using standard reference materials to ensure that the correct solvent and time have been used for the target analytes and matrix.

	Another more traditional technique for the extraction of solid samples, solid–liquid extraction (SLE), can be easily miniaturized and used for small samples. Glass vials can be loaded with the weighed samples and the appropriate amount of extraction solvent and then shaken either by hand or a mechanical type of shaker to ensure good interaction of the sample and solvent. The samples can also be heated on hot plates or water baths to temperatures above ambient but below the boiling point of the extraction solvent to speed up the extraction process. The nonextracted sample can be separated from the liquid by either centrifugation or filtration to obtain the liquid extract for subsequent sample processing prior to analysis. Despite the difficulty of automating the technique because of the latter steps (19), SLE is still a common technique used in laboratories for its simplicity. In recent years, its use in combination with alternative, green and more selective solvents, such as ionic liquids (ILs) and deep eutectic solvents (DES), has helped to solve additional shortcomings associated with its use with conventional volatile organic solvents, such as limited selectivity. As a typical example, Helalat-Nezhad et al. (4) reported on the feasibility of choline chlorideâoxalic (ChClâOx) to completely dissolve marine biological samples (fish and macroalgae samples). The dissolved extract (5 mL) was then percolated through a SiO2 column for selective retention of biogenic material, while the targeted PAHs were quantitatively eluted with 5 mL of cyclohexane. The authors reported recoveries in the 72–110% range, and LODs of 0.50–3.08 ng/g using LC–fluorescence (FL) detection and only 100 mg of sample for analysis. Interestingly, the extraction was completed at 55 ºC in 30 min at atmospheric pressure, which is an extraction time in the range of those used with other enhanced extraction techniques, despite the less drastic extraction conditions. 

	Matrix solid-phase dispersion (MSPD) is a technique that is often used for solid samples of more conventional sizes. In this technique, the solid or semisolid sample is mixed with an appropriate solid-phase sorbent to disperse the sample. This dispersed sample is then loaded into an SPE-type cartridge and subsequently treated like an SPE cartridge. The sorbent in the cartridge is eluted with the appropriate organic solvent to obtain a liquid extract containing the target analytes and essentially free of (nonstructurally related) interferences. This process can be easily miniaturized by adjusting the amount of solid sorbent and the volume of the cartridge used. The addition of a cosorbent on the base of the MSPD column to retain coeluted (structurally related) matrix components (5) or the use of highly selective sorbents, like molecular imprinted polymers (MIPs) (6), can result in readyâforâanalysis extracts, thereby minimizing extract manipulation and improving analytical response times. Despite its many positive features, MSPD is difficult to automate and it can easily fail at efficiently extracting analytes from various classes at the same time, except if a careful optimization of the extraction conditions is performed (7,13) (Figure 1).

Ultrasonic extraction (USE) is another technique frequently used for conventionally sized samples (10–30 g) that can be scaled down to work with sizeâlimited samples. USE is typically performed one of two ways: either extraction in an ultrasonic bath or extraction with an ultrasonic tipped probe. To perform USE in a bath, the solid sample is placed in an appropriately sized glass vial to contain the sample and solvent and then placed in the bath for a preselected extraction time (8,9). Afterwards, the solid and solvent are separated using either centrifugation or filtration. In the dynamic mode, the sample is typically placed in a refillable column, which is immersed in the ultrasonic bath. The extraction solvent flows continuously through the sample and is transferred either to a vial (11) or on-line to the instrumental analysis (21). If a sonic probe is used, it is placed directly in the vial containing the sample and extraction solvent. This direct contact between the tip and the sample results in a very efficient extraction and shortened extraction times (that is, a few minutes or even seconds suffice for matrix disaggregation and analyte extraction) (10). However, because of the large amount of heat dissipation, the sample often needs to be cooled, and volatile and labile analytes can easily be lost. Whatever the USE format selected, subsequent concentration and purification of the extracts before separation+detection is most frequently required because of the nonselective extraction process. A variety of techniques ranging from classical open columns (11) to disposable pipettes (10) or stir bars followed by analyte back extraction (when combined with LC [8]) or thermal desorption (with GC [9]) have been used. Approaches involving on-line systems are based on dynamic USE and are scarce in the literature (21). 

	Methods with Automation that Can be Scaled Down: The discussed methods can be easily adapted to small samples, but they are, in general, manual and can involve a lot of hands-on manipulation. While this may be fine for a few samples, for laboratories that have high sample loads, methods that use at least some automation would be advantageous.

	Microwave assisted extraction (MAE) is often used for the extraction of environmental and food-related samples of conventional sizes. One advantage of MAE is the rapid heating caused by the microwave radiation. Some solvents do not absorb much energy and the vessel that contains the sample is constructed of materials that absorb the microwave energy and cause the sample to heat up. Alternatively, the extraction solvent can be mixed with a polar solvent. However, this can negatively affect the efficiency of the extraction process (13). In general terms, MAE can be performed in two basic ways: open vessel or closed vessel. Open vessel MAE is not performed as often as closed vessel because organic solvents can be heated to temperatures above their boiling points very quickly, and that could cause sample loss. There have been some reports of using focused microwaves along with Soxhlet-like glassware for open vessel MAE so that the boiling solvent is captured via the condenser (22). However, to the best of our knowledge, the technique has not been applied for the analysis of trace organic pollutants in small-size samples. The use of closed vessels appears to be the more common approach used for MAE. An advantage of this approach is that the use of closed vessels allows extractions to take place above the boiling point of the solvents chosen. Operation at these elevated temperatures and pressures (50–150 ºC; 100–500 psi) facilitates the rapid extraction of solid samples and minimizes the amount of solvent used. In addition, multiple samples can be extracted simultaneously in single batches to improve the overall productivity (up to 50 in a batch). Different vessel sizes are available from commercial vendors to accommodate smaller sample sizes (1 g or less). There are also systems that can extract one sample at a time and are equipped with autosampler-type robotics that accommodate many samples for unattended operation. After the extraction, the sample vessels are allowed to cool and the contents are filtered to separate the extracted solid sample from the solvent containing the extracted compounds. Depending on the complexity of the sample, subsequent (off-line) cleanup of the obtained extracts before instrumental analysis can become mandatory, making the entire sample preparation process more labour-intensive and time-consuming. Different analytical strategies have been explored in the literature to solve this type of shortcoming and to improve the selectivity of the extraction step. The development by Guo and Lee was an imaginative and interesting example (12). They proposed the combined use of MAE for the extraction of PAHs from soil with in-cell solvent bar microextraction (SBME) for simultaneous purification and selective preconcentration of the target analytes. In this study, 1 g of soil was extracted for 20 min using water at 60 ºC as a green and inexpensive solvent. The minimum volume of solvent recommended by the instrument manufacturer-10 mL-was enough to immerse the sample completely and to ensure that the solvent bar could move and tumble freely into the extraction vessel to promote appropriate analyte enrichment during the extraction step. The results obtained by MAE–SBME were similar to or slightly better than those obtained with conventional MAE followed by analyte purification with SPE. However, in the former case, analyte extraction, purification, and preconcentration were accomplished in a single step. This application example clearly illustrates the potential of innovative approaches, such as MAE–SBME, for the simple, fast, and efficient preparation of complex matrices.

	Supercritical fluid extraction (SFE) has been used for the extraction of solid samples for several decades. In SFE, carbon dioxide, with added organic modifiers like methanol, is used as the extraction solvent. The extraction takes place at elevated temperatures and pressures (35–100 ºC; 2000–10,000 psi). The solvent (supercritical fluid) flows through the extraction cell containing the solid sample. As with MAE, these conditions facilitate rapid extractions, and small volumes of organic solvents are consumed, especially with small samples. SFE can be used in two basic configurations for the preparation of size-limited samples: off-line and on-line. In off-line SFE, the system is a standâalone instrument that can extract samples individually or in parallel. The extracts are collected in organic solvents directly or on solid supports that are subsequently washed with organic solvents. These extracts can then be analyzed via separate chromatographic instrumentation. In on-line SFE, the effluent from the extraction vessel is directed to an analyte trap or concentrator column. Once the extraction has been completed, the flow path is changed via a switching valve, so that the trap or concentrator column is in-line with the chromatographic separation column. Instrumental analysis is then performed on the extracted and collected sample. Either of these configurations can be adapted to accommodate small samples for extraction, as demonstrated by Cuong et al. (14) in a recent example dealing with the SFE of pesticides of varying polarity from hair collected from nonexposed humans. The optimized method allowed SFE of nonpolar organochlorine pesticides (OCPs), such as hexachlorobenzene, lindane, butapon, p,p′-DDE, and p,p′-DDT, with pure CO2 (recoveries in the 51–92% range; standard deviation [SD], 7–17). However, the quantitative SFE of more polar organophosphorus pesticides (OPPs), such as dichlorvos, 2,4-D, and chlorofos, required modification of supercritical CO2 with 5% methanol (recoveries, 71–81%; SD, 7–20). When combined with a brief static SFE step for previous selective removal of endogenous material, the proposed dynamic SFE method resulted in clean chromatograms and allowed accurate analyte determination at the µg/g level. 

	Pressurized liquid extraction (PLE) is also known as accelerated solvent extraction (ASE) and pressurized solvent extraction (PSE). PLE is similar to SFE, except organic solvents or solvent mixtures are used as the extraction fluid, and no carbon dioxide is used in the extraction. Elevated temperatures and pressures are used (40–200 ºC and 1500–2000 psi) to facilitate rapid extractions with small solvent volumes. The solvent flows through the extraction cell containing the solid sample, and the extracts are collected in glass vials or bottles. There are some unique advantages to SFE and PLE as compared to MAE because of the flow through mode of operation. The sample extracts are filtered as they are collected because the extraction cells contain fritted materials in SFE and PLE. With the flow through operation, sorbents can be placed in the flow path and unwanted interferences can be trapped as the samples are extracted. Sample extracts that are filtered and contain fewer interferences can therefore be easily produced with SFE and PLE. When the sorbents are packed in the PLE extraction cell, the technique is called selectiveâPLE (s-PLE). This latter arrangement (see Figure 2 for a typical example for the analysis of persistent organic pollutants [POPs] in feedstuffs) has been shown to contribute to minimized sample manipulation, sorbent, and solvent consumption, and to significantly shorten the analytical times (16).

While PLE is often performed on larger samples (5–100 g), commercial instrumentation is available with extraction cells as small as 1 mL. These will work for small samples (1 g or less), but the results might not be as satisfactory as with laboratory or in-house built instruments because of dead volume and other considerations (3,16). More importantly, this type of miniaturized system is now the only feasible method for on-line coupling between PLE and the instrumental analytical step (23). 

	Dynamic PLE has generally been adopted in on-line coupling of subcritical water extraction (SWE, a particular type of PLE) with subsequent analyte separation+detection typically done by LC (18). Unlike conventional PLE, green water-based solvents are used for SWE. The technique also involves much higher extraction temperatures (typically, 100–300 ºC), making subsequent depressurization and cooling down of the cell eluent mandatory before analyte trapping. These extra elements complicate the setting up of SWE devices and could be responsible for the limited progress observed in this field, despite its remarkable analytical features.

Desirable Future Developments and Conclusion

	As previously stated, there have been many reports in the literature on the use of laboratory-built instruments either directly coupled to GC–MS or LC–MS or in a stand-alone configuration for the extraction and cleanup of sizeâlimited solid and semisolid samples. These are useful for sizeâlimited samples across a broad spectrum of matrices and analyte types. However, there are few commercially available instruments with these capabilities. In this sense, probably the most significant addition is the recent introduction of an SFE–SFC system that offers extraction vessels as small as 0.2- and 0.5-mL, and is ideally suitable for the extraction of milligram quantities of solid samples followed by the determination of target analytes by SFC–UV or SFC–MS.

	We are not aware of other commercially available instruments. While small volume PLE has been successfully demonstrated in the literature (3,16,23), the smallest volume of commercial extraction cells available for this technology is 1 mL, which can still be too big for certain applications. The same can be said of the other techniques covered in previous sections. The authors encourage commercial vendors to carefully consider the development and commercialization of sample preparation and extraction instrumentation that has been optimized for use with samples of less than 1 g in size. This is important as concern for the environment grows alongside the drive for more efficient and more “green” laboratory practices.

		M. Kock-Schulmeyer, M. Villagrasa, M. Lopez de Alda, R. Cespedes-Sanchez, F. Ventura, and D. Barcelo, 

		L. Ramos, J.J. Vreuls, and U.A.T. Brinkman, 

		Z. Helalat-Nezhad, K. Ghanemi, and M. Fallah-Mehrjardi, J. Chromatogr. A 

		M. Llompart, M. Celeiro, J.P. Lamas, L. Sanchez-Prado, M. Lores, and C. Garcia-Jares, 

		Y. Wen, L. Chen, J. Li, Y. Ma, S. Xu, Z. Zhang, Z. Niu, and J. Chho, 

		E.M. Kristenson, S. Shahmiri, C.J. Slooten, J.J. Vreuls, and U.A.T. Brinkman, 

		P. Tolgyessy, B. Vrana, and K. Silharova, 

		M. Pena-Abaurrea, V.S. Garcia de la Torre, and L. Ramos, 

		C. Domeño, M. Blasco, C. Sánchez, and C. Nerín, 

		L. Haroune, R. Cassoulet, M.P. Lafontaine, M. Belisle, D. Garant, F. Pelletier, H. Cabana, and J.P. Bellenger, 

		L.P. Cuong, M.I. Evgen’ev, and F.M. Gumerov, 

		Y. Ren, J.J. Lian, H.X. Xue, J.M. Chen, and T.T. Cheng, 

		M. Pena-Abaurrea, J.J. Ramos, M.J. Gonzalez, and L. Ramos,

		T. Hyotylainen, K. Hartonen, S. Syndjoki, and M.L. Riekkola, 

		K. Kuosmanen, T. Hyötyläinen, K. Hartonen, and M.L. Riekkola, 

		E.M. Kristenson, E.G.J. Haverkate, C.J. Slooten, L. Ramos, R.J.J. Vreuls, and U.A.Th. Brinkman, 

		C. Sanchez, M. Ericsson, H. Carlsson, A. Colmsjö, and E. Dyremark, 

		F. Preigo-Capte and M.G. Luque de Castr, 

		Y.S. Zhou, W.M. Liu, J.H. Zhao, and Y.F. Guan, 

 is currently a Senior Scientific Researcher of the Scientific Research Council (CSIC, Madrid) of Spain at the Department of Instrumental Analysis and Environmental Chemistry, in the Institute of Organic Chemistry. Her research interests include the development of novel sample preparation methods for the determination of trace pollutants in environmental and food samples (including their scaling down and miniaturization) and the evaluation of new chromatographic techniques, especially GC×GC-based approaches, for unravelling the composition of complex mixtures of organic microcontaminants. 

 is currently an R&D Manager in the Agilent CrossLabs Group (ACG) of Agilent Technologies in Wilmington, Delaware, USA. He has been involved in the development of sample preparation technologies for many years, and he was a member of the R&D team that developed accelerated solvent extraction (ASE) at Dionex Corporation (now Thermo Scientific). His research interests include LC column development, GC column development, and the development of new approaches for improving overall analytical performance through innovative sample preparation technologies.

Cover Story

Extraction of Micropollutants From Size-Limited Solid Samples

An excerpt from LCGC’s e-learning tutorial on high performance liquid chromatography (HPLC) methods at CHROMacademy.com

	In chromatography, “like dissolves like”; that is, nonpolar analytes interact well with nonpolar stationary phases and vice versa. Increased and improved interactions with the stationary phase lead to higher distribution constant (kd) values and generally improved separations. Neutral or ion-suppressed analytes are less polar than ionized analytes and will therefore have improved retention on nonpolar reversed-phase type stationary phases, while in their ionized forms they will exhibit decreased retention. The mobile-phase pH can have a dramatic effect on the ionization state of ionizable analytes, so it must be fixed by buffers to maintain the analyte in the desired state.

	Knowledge of the functional group chemistry and the associated pKa of an analyte will allow for tuning of the mobileâphase pH. The pKa gives an indication of the strength of the acid or the base; however, it cannot be decided from the pKa alone if the molecule is an acid or a base-the functional groups contained within the molecule must also be known. For example, the pKa of aspirin and diazepam are very similar (3.5 and 3.3, respectively). Aspirin is a weak acid because it contains carboxylic acid groups, and diazepam is a weak base and contains basic nitrogen functional groups. The pH will affect the extent of analyte ionization, which will in turn influence elution and retention properties. For an acidic analyte, in a buffered solution, the addition of an acid will lower the pH and the analyte will become less ionized. The change in degree of ionization happens over a limited pH range because of pH and pKa being logarithmic: 1 pH unit away from the pKa, the extent of ionization is ~90%; at 2 pH units away from the pKa, the extent of ionization is ~99%; and at 3 pH units it is 99.9% (Figure 1). At a pH equal to the pKa an ionizable molecule will be 50% ionized and 50% non-ionized, which can lead to poor peak shape. 

Unlike ionized acids, which are eluted rapidly from the column when charged, protonated bases may have long retention times and poor peak shape because of interaction with silanol species on the silica surface. Separations of basic compounds are not usually carried out under ionâsuppression conditions because the increase in pH to produce the neutral species would damage traditional silica columns (although hybrid columns can be used at extremes of pH). Traditionally, the analysis of weak bases is carried out at low pH so that surface silanol species are non-ionized (pKa 3.5–4.5), which results in improved peak shape.

	Mobile-phase pH is controlled using a buffer consisting of a weak acid or base in cosolution with its conjugate acid or base. Buffers are only reliable within 1 pH unit either side of their pKa, and their concentration must be adequate but not excessive: below 10 mM buffers will have very little buffering capacity; above 50 mM, there is a high risk of precipitation of the salt in the presence of high organic concentration, hence, concentrations of 25–50 mM are typical. For liquid chromatography–mass spectrometry (LC–MS) applications the buffer must be volatile. It is good practice to prepare buffers daily because pH can change on standing due to ingress of carbon dioxide. 

	Sacrificial bases or ion-pair reagents can be used to improve peak shape or retention of basic analytes. Sacrificial bases are sterically small, highly surface active species (for example, triethylamine) that preferentially interact with surface silanol groups. They are added to the mobile phase in sufficient concentration (10–100 mM) to ensure full surface coverage. Modern, high purity, low-silanol packing materials negate the need for sacrificial bases. 

	Ion-pair reagents can be used when other methods such as ion suppression have not been successful. Samples containing both anionic and cationic components have one type “masked” by the ion-pair reagent and the other suppressed by pH. This technique is useful if analyte pKa values are not similar. For example, tetrabutylammonium phosphate at pH 7.5 forms a strong ion pair with acids and pH suppresses weak base ions. Ion-pair reagents do have disadvantages, including long equilibration times (>100 column volumes); difficult removal from columns (it is recommended that a guard column or dedicated column is used); irreversible modification of the stationary phase, which drastically reduces lifetime; neutral analytes precluded from the stationary phase at very high ion pair concentrations, which results in decreased retention; and interference from ultraviolet (UV) activity of some ion-pair reagents at the analytical wavelengths being used (such as trifluoroacetic acid, 210 nm). Finally, ionâpair reagents are not suitable for LC–MS work because they suppress ion formation and reduce sensitivity. Alternative approaches, such as hydrophilic-interaction chromatography (HILIC) and mixed-mode chromatography, can be used for LC–MS. Another alternative is the use of modified column chemistries that have counterions built in as part of the stationary phase; these embedded functional groups can be activated by changing the mobileâphase pH, without the need for additional chemical modifiers.

The Essentials

Critical Evaluation of HPLC Methods: Working with Ionizable Analytes

In this instalment, John Hinshaw presents an annual review of new developments in the field of gas chromatography (GC) seen at Pittcon 2017 and other venues in the past year.

	For its 69th session 5–9 March 2017, the Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy (Pittcon) headed north to McCormick Place in Chicago, Illinois, USA. In sharp contrast to the many previous subzero sessions held in Chicago, the weather was unusually mild, so much so that I was able to enjoy a Sunday morning run along the Lakefront Trail. I passed by the 17th Annual Chicago Polar Plunge where hundreds of bewildered swimmers encountered water colder than the air. Attendance was up slightly compared to last year with more than 14,000 registered participants, 22% of whom came from 89 countries outside the United States. The exposition hosted 78 exhibitors from 37 countries, 141 of which displayed their products for the first time.

	The technical programme continued to be a strong part of the conference: more than 2000 sessions were presented in 72 symposia, 89 oral sessions, 19 contributed sessions, six workshops, 56 poster sessions, and 15 awards. Among these were the 2017 LCGC Lifetime Achievement in Chromatography Award presented to Professor Pat Sandra (Research Institute for Chromatography), and the LCGC Emerging Leader in Chromatography Award presented to Dr. Deirdre Cabooter (University of Leuven). 

	Pittcon 2018 will meet in Orlando, Florida, 26 February to 1 March at the Orange County Convention Center, where participants likely, but not necessarily, will enjoy even balmier weather. The 2019 session is scheduled for 17–21 March in Philadelphia, Pennsylvania.

	This annual “GC Connections” instalment reviews gas chromatography (GC) instrumentation, columns, and accessories shown at this year’s Pittcon or introduced during the previous year. For a review of new products in other areas of chromatography, columns, and related accessories, please see the “Column Watch” and “Perspectives in Modern HPLC” column instalments from the April 2017 issue of LCGC Europe (1,2) as well as the “Sample Prep Perspectives” column in the May issue of LCGC North America (3).

The information presented here is based on manufacturers’ replies to questionnaires, as well as on additional information from manufacturers’ press releases, websites, and product literature about the past year’s products, and not upon actual use or experience of the author. Every effort has been made to collect accurate information, but because of the preliminary nature of some of the material LCGC Asia Pacific cannot be responsible for errors or omissions. This column instalment cannot be considered to be a complete record of all new GC products introduced this year at Pittcon or elsewhere because not all manufacturers chose to respond to the questionnaire, nor is all of the submitted information necessarily included here because of the limited available space and the editors’ judgment as to its suitability.

	As the gas–solid variety of GC attains its 70th anniversary in 2017, this past year had a number of notable advances in GC technology that clearly demonstrate its ongoing viability. Far from those early university experiments, some of the newest developments help remove a number of significant obstacles in routine GC while other advances deliver even higher performance to the gas chromatography–mass spectrometry (GC–MS) realm. The most significant development in GC instrumentation this year has to be Agilent’s new Intuvo 9000 GC system, designed for reduced time spent on routine tasks with minimized maintenance risks. The core Intuvo developments are outlined in the following tables. Laboratories that adopt the system will deploy columns in instrumentâspecific directly heated modules, but this should prove not to be a significant barrier to the multiyear cycle of instrument upgrades and replacements. Ellutia’s 500-Series GC systems implemented a hybrid airâoven column heating system, which supports resistively heated as well as conventionally heated columns. Qmicro displayed application-specific versions of their micro-GC platform, which is based on microelectromechanical systems (MEMS) for the inlets and detectors plus direct column heating. All three of these systems make use of customized column formats. 

	For GC–MS, the Exactive GC Orbitrap GC-MS system from ThermoâFisher Scientific, the GCMS-QP2020 GC–MS system and GCMS-TQ8050 tripleâquadrupole GC–MS system from Shimadzu, and the Pegasus BT GC–timeâofâflight MS system from LECO represent the latest advances in GC–MS technologies. Two new applicationâspecific offerings from Agilent in this area, the SureTarget GC/MS Water Pollutants Screener and the GC/MS Arsine Phosphine Analyzer, plus the EPA 8270D analyzer kit from Thermo Fisher Scientific, all emphasize the ongoing march towards routine application of highâperformance GC–MS for standardized methods.

	Two optical spectroscopy GC detectors were announced during the past year. Continuing their advancements in vacuum ultraviolet spectroscopic detection for GC, VUV Analytics expanded the
	wavelength range of its VGA-101 detector as well as enabling series coupling with some conventional GC detectors. From Dani Instruments, the DiscovIR–GC Solid Phase GC–FTIR detector produces solid-phase transmission infrared (IR) spectra of eluted GC components.

	A number of sampling accessories round out the newest instrumental offerings, including the XR Series of thermal desorption instruments and HiSorb extraction probes from Markes, the 6000 Series Pyroprobe pyrolyzers from CDS Analytical, and the Master MTAS robotic autosampler platform from Dani, all of which advance their respective sampling tasks with new or improved capabilities. 

	The zone of general GC accessories was not left out this year, either. The ADM flowmeter from Agilent has some new calibration capabilities that minimize downtime. New Thermolite septa and Topaz inlet liners from Restek are designed to extend trace-level analysis capabilities. Restek also now offers an electron ionization (EI) filament replacement for a number of Agilent MS detectors. There was a large crop of new gas generators, too: VICI DBS entered this segment with several offerings for GC systems, Proton OnSite showed a new G-Series family of hydrogen generators, and Peak Scientific has a new nitrogen generator.

	In GC columns, Agilent and Phenomenex introduced some new polar columns that offer increased stability and upper temperature limits as well as application-specific selectivity tailoring. Agilent’s new Intuvo System modular columns are being made available in a variety of common dimensions and stationary phases; the modules feature a quick-connect planar cage that does not use conventional ferrules.

	2016–2017 was another very active year in GC that again emphasized the pivotal role that small-molecule and volatile component analyses fulfil in the fields of separation science. As we spin around to Pittcon 2018, I expect to be pleasantly surprised by more new developments and innovations.

	I would like to thank the manufacturers and distributors that kindly furnished the requested information, which allowed a timely report on new product introductions over the past year. For those manufacturers who did not receive a “New Products” questionnaire this year and would like to receive one and be considered for early inclusion into the 2018 new GC and related product introductions review, as well as the other related review articles to be published in LCGC, please send the name of the primary company contact plus the mailing and e-mail addresses to Laura Bush, Editorial Director, LCGC and Spectroscopy, UBM Americas, 485 Rte. 1 South, Bldg. F, Suite 210, Iselin, NJ 08830, USA, Attn: 2018 New Chromatography Products. The questionnaire will be sent out later in 2017.

 is a Senior Scientist at Serveron Corporation in Beaverton, Oregon, USA, and a member of LCGC Asia Pacific’s editorial advisory board. Direct correspondence about this column to the author via e-mail: lcgcedit@lcgcmag.com

GC Connections

New Gas Chromatography Products for 2016–2017

Click the title above to open the LCGC Asia Pacific June 2017 regular issue, Vol 20, No 2, in an interactive PDF format. 

LCGC Asia Pacific-06-01-2017