Articles

LCGC North America, September 1, 2019

Click the title above to open the LCGC North America September 2019 issue, Volume 37, Number 9, in an interactive PDF format.

Vol 37 No 9 LCGC North America September 2019 Regular Issue PDF

Carefully diluting a sample with weak solvent can mitigate the impact of sample solvent on peak shape in both reversed-phase and HILIC separations, but we need to understand how the choice of sample diluent can affect analyte recovery.

The sample solvent can have a big impact on peak shape in both reversed-phase and HILIC separations, especially when large volumes are injected. Diluting the sample with weak solvent can be an effective solution to mitigate this problem, but we have to be careful to not lose analytes of interest to precipitation or phase separation. 

In the August 2019 installment of "LC Troubleshooting," I wrote about one of the hot topics that was discussed by several speakers at the The HPLC Meeting in Milan, Italy, in June-coupling of ion-exchange separations to mass spectrometric detection (1). This month, I'd like to discuss another topic addressed at the meeting. Anne Mack gave a talk that highlighted some things to consider when deciding whether to use reversed-phase or hydrophilic interaction (HILIC) liquid chromatography, especially when some of the components of the sample mixture at hand are hydrophilic. These considerations included the relative retention of the analytes of interest under reversed-phase and HILIC conditions, the effect of the sample diluent and injection volume and peak shape, and the effect of the sample diluent on apparent recovery of the analyte. The recovery aspect nicely complements the "LC Troubleshooting" article I wrote in January 2019 on the effects of the diluent on peak shape (2), so I've asked Anne to join me in writing this month's installment of "LC Troubleshooting."

	Liquid chromatography (LC) is an incredibly versatile analytical tool, enabling quantitative and qualitative analysis of diverse molecule types, ranging from highly hydrophilic and water soluble (for example, inorganic metal ions) to highly lipophilic molecules that are soluble in organic solvents (for example, lipids). The reversed-phase mode of separation is arguably the most versatile single mode of LC separation, and can provide retention and separation of analytes covering a wide range of water solubility. In an extreme case, it could separate molecules as hydrophilic as small organic acids (like succinic acid [3]), and as lipophilic as fat soluble vitamins (like vitamin A [4]). However, a practical problem we run into quickly is that molecules as different in properties as these will not be highly soluble in the same sample solvent. In the worst case, choosing an inappropriate sample solvent or diluent will lead to inaccurate results, particularly for quantitation, because the analytes of interest will not be soluble in the sample solvent, and the sample that is injected into the LC instrument will not be representative of the analytes actually present in the sample vial.

	The problem described here is not new by any means. But, as we push the limits of analytical methods in the reach for more speed, simplicity, and sensitivity, we inevitably run into conditions that will "break" the method, leading to inaccurate results. Therefore, we have to be careful not to break our methods, and a deeper understanding of where the limits lie and the basis for them puts us in a better position to be successful in the long run.

Review: Effects of Sample Diluent and Injection Volume on Peak Shape

	As users of LC, it is common to encounter situations where the samples presented to us from some prior process or step contain a sample solvent that is quite different from the mobile phase associated with the LC method used for the analysis of the sample. For example, when using solid-phase extraction (SPE) to preconcentrate low-concentration analytes from water samples, the last step in the SPE process typically involves elution of the analytes of interest from the SPE adsorbent using an organic solvent like methanol. Frequently, such extracts are subsequently analyzed by reversed-phase LC, with solvent gradient elution that starts with a water-rich mobile phase. On the other hand, we might be interested in analysis of the water-soluble components from a urine sample. In this case, the sample solvent is water, while the HILIC separation will start with a mobile phase containing a high fraction of acetonitrile on the order of 90%. These mismatches between the solvent composition of the sample and the mobile phase of the LC method can lead to trouble. In January of this year, we wrote about how the combination of the volume of sample that is injected and the sample solvent composition can have a dramatic effect on peak shape. As a reminder of the data discussed in that installment, Figure 1 shows examples of bad results obtained under reversed-phase and HILIC conditions, and how much better separations can be obtained simply changing the sample solvent. In the case of the reversed-phase separation, terrible peak shape is observed (Figure 1a) for some simple alkylphenones when the sample solvent contains 20% more acetonitrile than the starting mobile phase used for the solvent gradient elution program. However, simply changing the sample solvent to contain 20% less acetonitrile than the starting mobile phase in the gradient leads to a much nicer separation (Figure 1b). Similar effects can be observed in HILIC separations as well. Figure 1c shows the terrible peak shapes that are observed when a completely aqueous sample is injected into a HILIC column when the mobile phase contains 85% acetonitrile. However, this separation can also be improved dramatically by simply changing the sample solvent composition to contain 95% acetonitrile, as shown in Figure 1d.


	Figure 1: Examples of the effect of sample solvent on reversed-phase-LC (a,b) and HILIC (c,d) separations. The analyte mixture for the reversed-phase-LC separation was a series of alkylphenones, and a solvent gradient was used for elution starting at 50% acetonitrile. The analytes for the HILIC separation were cytidine and guanosine, and the mobile phase was isocratic with 90:10 acetonotrile:water buffer. Complete chromatographic conditions are given in reference (2). Figures are (a) sample solvent 70:30 acetonitrile:water, (b) sample solvent 30:70 acetonitrile:water, (c) sample solvent 0:100 acetonitrile:water, and (d) sample solvent 95:5 acetonitrile:water.

	Recognizing the importance of the sample solvent composition, particularly when the injection volume is large relative to the column volume, several different groups have developed a number of approaches to address this issue. The simplest approach is to adjust the sample solvent composition offline-that is, by addition of "weak solvent" (for example, water in reversed-phase, or acetonitrile in HILIC separations) as part of the sample preparation process prior to the LC separation. For example, in the case of the SPE extract in methanol described above, one could simply dilute this extract with some amount of water prior to analysis.

	Other approaches described over the years include:

		Online dilution of the sample with weak solvent in the autosampler needle; this is sometimes referred to as "sandwich injection" (5).

		Online dilution of the sample with weak solvent by deliberately adding a mixer between the sample injection point and the column inlet (6).

		Online dilution of the sample with weak solvent during injection into the LC column-this is sometimes referred to as 

, and requires an auxiliary pump to deliver the diluent (7).

		Online dilution of the sample with weak solvent during injection by splitting the mobile phase flow path to achieve inline mixing of the sample and diluent (8).

	While these approaches have been used with conventional one-dimensional LC (1D-LC), this issue is also very important in 2D-LC, and a variety of approaches have also been developed to address the problem in the context of 2D-LC specifically (9–12).

	As with most challenges in chromatography, there is no perfect solution to this sample solvent matrix issue, and all of these approaches have advantages and disadvantages. Common to all of them, however, is the question, "How much dilution is enough?" This is obviously a very practical question that must be confronted in method development. Unfortunately, there is no universal answer that is suitable for all types of separation and analyte. Most often, the answer for a particular situation is determined by experiment, and typically the primary focus of such experiments is the effect of the sample solvent on peak shape. In this installment we address a secondary, but still important concern-analyte recovery.

Effect of Sample Diluent on Analyte Recovery: HILIC Separations

	As discussed above, one straightforward approach to address the effect of the sample solvent mismatch relative to the mobile phase is to simply dilute the sample with weak solvent, mix, and inject. However, we have to be careful that when the diluent is added the sample solution remains homogeneous. Two mechanisms that can result in a heterogeneous solution are: 1) precipitation of some analytes or matrix components such that solids and liquids are present in the sample; and 2) phase separation of some analytes or matrix components such that two or more liquid phases are present after adding the diluent. Both of these outcomes are undesirable, because the material sampled from these heterogeneous solutions will not be representative of the entire sample, and will lead to inaccurate quantitative results. For samples that contain analytes having similar physico-chemical properties, these outcomes can easily be avoided by doing a few scouting experiments to see when or if precipitation or phase separation occurs. However, avoiding these outcomes can be more challenging when the components of the sample are more diverse in terms of their solubilities in the sample solvent, particularly when the diluent is added.

	Figure 2 shows an example of this problem in the context of use of a HILIC method for the analysis of a sample containing water soluble vitamins. For nicotinic acid (Figure 2a), the observed peak area is nominally same whether the sample is prepared in a matrix of 100% water or 90:10 acetonitrile/water. However, for thiamine and cyanocobalamin (Figures 2b and 2c, respectively), the observed peak areas in the 90:10 acetonitrile/water matrix are roughly 50% of the area observed when the sample is prepared in 100% water and all other conditions are the same. This suggests that thiamine and cyanocobalamin are not fully soluble in 90:10 acetonitrile/water, and some of the analyte precipitates from the sample matrix before it is sampled for analysis. In a case like this, the effect of the diluent on analyte recovery must be studied, and taken into consideration when deciding what sample solvent composition will be used for the final method.


	Figure 2: Comparison of peak areas obtained from HILIC separations for injected from samples prepared in either water (blue) or 90:10 acetonitrile:water (red). Chromatographic conditions: Solvent A: water, Solvent B: acetonitrile; gradient elution from 95–65% B in 10 min., with 10 mM ammonium acetate (pH unadjusted) throughout; Temperature, 25 °C; Flow rate, 0.5 mL/min.; Column, 100 mm × 2.1 mm i.d. Poroshell 120 HILIC-OH5 (2.7-µm); Injection volume, 0.5 µL; Detection by triple quadrupole mass spectrometry.

Effect of Sample Diluent on Analyte Recovery: Reversed-Phase Separations

	The sample diluent can adversely affect analyte recovery in reversed-phase-LC separations as well. To illustrate this effect, we prepared an analyte mixture of benzylalcohol, butyrophenone, and octanophenone-all at 1 mg/mL-in different sample solvents ranging from 10:90 acetonitrile/water to 100% acetonitrile. The predicted water solubilities for these compounds are 15, 0.34, and 0.0037 mg/mL, respectively (

). Based on this, we would expect to see consistent peak areas over the full range of sample solvent compositions for benzyl alcohol. On the other hand, we would expect to see consistent peak areas for the lipophilic butyro- and octanophenones in the samples with high levels of acetonitrile, but lower areas in the water-rich samples, because of the low water solubilities of these molecules. Figure 3a shows the absolute peak areas measured for the three compounds at each solvent composition; Figure 3b shows the same data, but normalized to the area observed with the sample prepared in 100% acetonitrile. As expected, we see consistent peak areas for benzyl alcohol across the entire range of sample solvents. Note that the areas for intermediate sample solvent mixtures are slightly greater than 100%. This is most likely due to the volume contraction of acetonitrile-water mixtures, given the way the samples were prepared (for example, for the 50% acetonitrile sample, 500 µL of water was added to 500 µL of acetonitrile, and this results in a mixture that has a volume of about 970 µL). However, for butyro- and octanophenone, we see that consistent peak areas are observed down to 40% acetonitrile, but then there is a precipitous decrease in the peak area, with the decrease more dramatic for octanophenone than for butyrophenone. This results from the formation of two phases in the samples with 30% acetonitrile or less because of the very low water solubilities of these compounds. Given that their densities are lower than that of water, there is probably a top layer of the solution in the HPLC vial that is enriched in these compounds, but not sampled by the autosampler needle, which samples from well below the liquid surface.


	Figure 3: Effect of sample solvent on peak area observed for neutral compounds of varying lipophilicity under reversed-phase-LC conditions. Peak areas are plotted as (a) absolute values or as (b) a percentage of the area observed when the sample solvent is 100% acetonitrile. Chromatographic conditions: Solvent A: water, Solvent B: acetonitrile; Gradient: 10-100-100% B for 0-3.5-4.0 min; Temperature, 40 °C; Flow rate, 1.0 mL/min.; Column, 50 mm × 4.6 mm i.d. Zorbax SB-C18 (5-µm); Injection volume, 1.0 µL; Detection by ultraviolet (UV) absorption at 210 nm. The balance of the sample solvent was water, and the concentration of each analyte was 1 mg/mL.

	In the everyday practice of LC, it is common to encounter samples where the sample solvent composition is very different from the mobile phase used in the LC separation. When injecting volumes of these samples that are large relative to the volume of the column itself, this can lead to poor peak shape. Several approaches have been developed to overcome this challenge, most of which involve dilution of the sample with weak solvent prior to or during the injection step. While this can be an effective remedy for poor peak shapes, we must be careful to avoid analyte precipitation or phase separation of the sample as a result of adding too much of the weak solvent diluent. The level of diluent that is considered too much is compound dependent, but can be determined experimentally through simple screening experiments.

	(4) S. Bäurer, W. Guo, S. Polnick, and M. Lämmerhofer, 

 167–180 (2019). doi:10.1007/s10337-018-3615-0.

	(5) D.R. Stoll, D.C. Harmes, G.O. Staples, O.G. Potter, C.T. Dammann, D. Guillarme, and A. Beck, 

 5923–5929 (2018). doi:10.1021/acs.analchem.8b00776.

	(6) Z. Breitbach, C. Randstrom, J. Chang, M. Lesslie, G. Webster, and D. Stoll, 

	(7) U.D. Neue, C.B. Mazza, J.Y. Cavanaugh, Z. Lu, and T.E. Wheat, 

 S121–S127 (2003). doi:10.1007/BF02492093.

	(8) A. McKay, M. Perkins, and J.A. Field, 

	(9) Y. Oda, N. Asakawa, T. Kajima, Y. Yoshida, and T. Sato, 

 411–418 (1991). doi:10.1016/S0021-9673(01)96013-3.

	(10) C. Doneanu, A. Xenopoulos, K. Fadgen, J. Murphy, St.J. Skilton, H. Prentice, M. Stapels, and W. Chen, 

 24–44 (2012). doi:10.4161/mabs.4.1.18748.

	(11) D.R. Stoll, K. Shoykhet, P. Petersson, and S. Buckenmaier, 

9260–9267 (2017). doi:10.1021/acs.analchem.7b02046.

10251-10257 (2019). doi:10.1021/acs.analchem.9b02391.

 is an Application Scientist at Agilent Technologies, in Wilmington, Delaware.

 is the editor of "LC Troubleshooting." Stoll is a professor and co-chair of chemistry at Gustavus Adolphus College in St. Peter, Minnesota. His primary research focus is on the development of 2D-LC for both targeted and untargeted analyses. He has authored or coauthored more than 60 peer-reviewed publications and four book chapters in separation science and more than 100 conference presentations. He is also a member of LCGC's editorial advisory board. Direct correspondence to: 

LC Troubleshooting

Effects of Sample Diluent on Analyte Recovery in Reversed-Phase and HILIC Separations

Most manufacturers ship gas chromatographs with a small collection of consumable parts and accessories, such as extra ferrules, inlet liners, or septa, and a few instrument-specific tools. In earlier times, some convenience items might have been included as well, like a bubble flow meter or a small set of tools, but now it's rare to find such things in the shipping boxes. This edition of "GC Connections" lists a number of essential items that should be on hand in every GC laboratory, their function, and how to use them effectively.  

	Every profession has its specialized tools. Those used in chromatography often are just as specialized as those used in computer repair or automotive work. Many of the tools and accessories that gas chromatographers keep on hand for installing, maintaining, and repairing their chromatographs are also found in plumbers', carpenters', and home toolkits. Wrenches, screwdrivers, pliers, and metal tubing cutters are some easily recognized examples. Other items such as dental instruments or paper correction fluid are familiar, but their use in the laboratory environment may not be immediately obvious. Still others, like a column flow meter, septum nut wrench, or a specialized fused-silica column cutter, aren't found outside the laboratory at all.

	Here then is an updated list of today's tools and accessories, with some information on their use and significance. One or more specialty manufacturers offer many of the chromatography-specific items; just look at their catalogs or on-line offerings. I scanned through several, and gleaned some new items that I have included here.

	Source of Gas for Measuring Unretained Peak Times

	A butane lighter is a convenient source of hydrocarbon gas for measuring an approximate unretained peak time. Butane is effectively unretained at temperatures above 75 °C on liquid-phase coated columns with phase ratios above 50. Columns at low temperatures or with lower phase ratios (thick stationary films) may retain butane, and separate the traces of ethane and propane present in the butane fuel. Use the earliest observable peak for the best estimate of unretained peak time. Natural gas is mostly methane; if your laboratory has a supply of natural gas (mine doesn't), it makes a good substitute for a lighter, and is less retained than butane-just be sure to turn off the gas once you've filled a syringe with it. A lecture bottle of methane with a suitable pressure regulator is another excellent source of the unretained substance. Concentrations in the low percent range work well. At least one chromatography supplier offers a prefilled methane bottle with a low-pressure regulator for this purpose.

	Hydrocarbons won't work for unretained peak time measurements with an electron-capture detector (ECD), because it doesn't respond. Instead, try loading the syringe with a puff from a pressurized can of dust remover such as Dust-Off, which contains 100% 1,1-difluoroethane. Other halocarbons will work as well; check the label to be sure there is an ECD-sensitive compound present.

	Hydrogen or helium-whichever is not the same as the carrier gas-makes good unretained peak markers for porous polymer or molecular sieve columns that retain hydrocarbons strongly, plus these two gases should be readily available in most GC laboratories. Flame ionization detection (FID) will not respond to hydrogen or helium, but other detection methods such as thermal conductivity (TCD), pulsed discharge (PDD), or helium ionization (HID), the latter with hydrogen as the unretained substance, should respond well.

	Remember to use the same inlet pressure and oven temperature to remeasure unretained peak times for any specific column; that is, one with the same dimensions, film thickness, and stationary phase for comparative purposes.

	Polymer caps for capillary columns establish an airtight seal for storage, so that air, moisture, and airborne contaminants cannot enter. I have used septa for this purpose, but there's a risk of breaking the column and puncturing a finger instead; this only happened to me once. Another approach is to seal the end of the column by melting it. This is not recommended, because it requires a hydrogen torch to achieve a high enough temperature, and it will cause stationary phase combustion by-products to migrate a significant distance into the column.

	Use white correction fluid to mark the measured position on a column that corresponds to the correct column penetration depth into an inlet or detector. After inserting the column into the nut and ferrule and making a fresh cut on the column end, measure the depth and apply a small dab of correction fluid. For data integrity reasons, some regulated laboratories' policies don't permit correction fluid on site, although in the fully electronic laboratory this is moot. A septum into which a slot has been cut may be slid onto the column below the nut and ferrule to act as a positioning aid. Remove the septum before heating the column oven. A positioning gauge (see "Ruler" later in this column) is a good alternative. I also have used a black permanent marker for this purpose; just be careful not to get it on your fingers or clothing.

	The best fused-silica column cutting tool is the one that holds the column in an adjustable chuck, and cuts with a diamond chip as the operator rotates a thumb wheel. This tool also has a magnifying glass on the opposite end for inspecting the fresh cut for squareness and lack of burrs or hanging polyimide coating. A pen-like tool with a sapphire tip or ceramic scoring wafers or scribes that make a sharp cut on the column so that it may be broken cleanly in two are the best inexpensive alternatives. In any case, a fresh cut should be made and inspected just before placing the column into the inlet or detector, after sliding on the nut and ferrule.

	Diagonal cutters are used only for cutting electrical wires. They should never be used as a substitute for a tubing cutter. Don't even think about threatening your fused-silica column with one!

	A large plumber's tubing cutter for 1/4-inch metal tubing, and a smaller one for 1/8-inch tubing, are used extensively during instrument installation and gas-supply setup. The correct 1/8-inch size cutters are available from some chromatography supply companies; don't try to use the mini cutters found in home supply stores-they're only good for those weekend plumbing projects. Keep a supply of new cutter blades on hand. These disks wear out rapidly, and a dull blade will distort the tubing diameter, and make it difficult to slide on the back and front ferrules. Power cutters with high-speed rotary abrasive cutting wheels are also available. These are mandatory for cutting 1/16-inch tubing for low dead-volume gas valving applications, and they work well for any tubing up to 1/4-inch diameter. After cutting and squaring the tubing (see "Tubing Reamer" and "Deburring Tools" later in this article), be sure to clean out the ends with a solvent, so that loose particles cannot get into the instrument components, or interfere with the passage of peaks through column ends.

	A plastic dental mirror with a front-silvered surface makes it easy to examine the underside of an inlet fitting in the oven, or to check other inaccessible areas for loose or missing parts. The mirror also can be used to detect the flame in a flame ionization or flame photometric detector by observing condensation of emitted water vapor on the cool mirror surface. A shiny wrench is a good substitute for the mirror in this case.

	A dental pick is very handy for removing septa from septum nuts, and debris such as bits of graphite ferrule from fittings.

	I have a box of plastic eyedrops and they are in frequent demand. I use them to place small drops of isopropanol onto fittings for leak checks. Some types have rough volume indications more like a pipette, and I have used them to make a crude dilution of qualitative test mixtures when accuracy wasn't required.

	An assortment of needle files can be used to pick out ferrules from fittings, as well as to remove burrs and shape the ends of metal tubing before it is connected to a fitting. Don't forget to clean off all traces of metal before connecting.

	A flexible 2-foot magnetic pickup comes in handy when you drop a small part inside the instrument. Another similar tool has a three-jawed "claw" operated by a plunger, and it will pick up non-magnetic items, too.

	An electronic flowmeter is an expensive investment, but I believe that it will pay for itself many times over with improved accuracy and precision over bargain-priced bubble flowmeters. I prefer the type of electronic meter that senses flow directly, and that allows the operator to select the type of gas in use, such as air, helium, or hydrogen. The option to calculate split ratios from the measured split vent and column flows is a handy feature, but many GCs now include this capability in their software.

	If you use bubble flowmeters, keep two sizes on hand. The large size is good for measuring FID air or inlet split vent flows up to several hundred milliliters per minute. The smaller size is better for packed-column or hydrogen flame-gas flows in the 10 to 50 mL/min range. Don't try to use a bubble flowmeter to measure capillary column flows below 10 mL/min. The carrier gas will diffuse out of the bubble, and you will get a low reading. Measure the unretained peak time instead, and calculate the flow rate from it. Note that this calculated flow rate or the rate displayed by electronic pressure control will only be as accurate as the column dimensions the operator uses.

	Manufacturer-specific flow adapters allow a flow meter to connect to the exact dimensions of a detector's exit. Short pieces of flexible tubing in various diameters are handy for connecting a flow meter to different size tubing (see "Tubing, Plastic and Rubber" later in this article).

	This item is useful for those who must install glass wool in inlet liners, or for the hardy few who pack their own columns and use glass wool to hold in the packing. These days I find little use for it, because I use prepacked deactivated inlet liners instead.

	Polymeric nitrile or vinyl lint-free gloves are essential for column installation and detector maintenance, especially for high-sensitivity detectors such as mass spectrometric, electron capture, and helium ionization types. Handling the column inlet without gloves opens the door to contamination from finger oils and dirt, while cleaning the rods of a quadrupole mass-selective detector (MSD) without gloves is asking for trouble. I recall a particular MSD that exhibited poor sensitivity. A tear-down revealed a huge thumb print on the photomultiplier tube window, perhaps deposited there by a technician who had recently finished lunch.

	I keep several sets of hexagonal key wrenches and drivers in the toolbox. I have sets of the traditional right-angle Allen keys in both imperial and metric sizes, as well as a collection of hex screwdrivers of both types. Although used less often, a set of star wrenches is indispensable as well. These also come in spline and security variations, some of which I've accumulated over the years. Many of these are used in laboratory instruments instead of the more traditional Phillips screw head, so they are essential for performing maintenance.

	High-temperature string is a useful item for use in a GC oven to restring capillary columns, attach column connectors to column cages, or hold the column in the GC oven. The string is made of materials that can withstand temperatures of up to 400 °C, or higher.

	A tapered high-temperature silicone rubber tool on a metal holder does a good job of grabbing glass inlet liners and removing them without cracking or chipping the liner top. Most GC instrument manufacturers will supply specific tools and instructions for a particular inlet option.

	An electronic leak detector is expensive, but it indispensable for finding small leaks around hot fittings or inlets on which a liquid cannot be used (see "Leak-Checking Solution" later in the article). The most sensitive type of leak detector uses a small pump to pull air from a probe through a thermal-conductivity cell. The presence of carrier gas or hydrogen changes the thermal conductivity and causes a change in the detector's readout compared to a reference air flow. Sensitivity for nitrogen carrier is limited. I also have a small handheld battery-powered leak detector that has a series of light-emitting diodes, which indicate the detected leak rate. This detector is great for carrying around in a laboratory like mine with lots of instruments and little clear bench space.

	In my toolbox, the only acceptable leak-checking solution is a small bottle of pure isopropanol with an eyedropper. Other solutions may contain material such as surfactants that can leak into the gas-supply lines or columns and cause ghost peaks or other contamination.

	A small magnifier is used to examine freshly-made column or tubing cuts for burrs or uneven edges.

	For each GC system, there are always some specialized tools. These are used, for example, to open up a split–splitless inlet, or to remove the inlet liner. Perhaps a special wrench is required for FID system flame jet replacement. Whatever the case, keep all such tools with their instrument; you will need them eventually. Some of the chromatography suppliers offer their own versions of these tools, which often are more useful than the freebie ones that come with the instruments.

	Kits consisting of a collection of the most often replaced parts and supplies are a good starting point with a new instrument. Available for inlets and detectors, these packages will get used almost immediately upon commissioning. After a while, I find it convenient to re-order only those items that are used frequently instead of purchasing additional kits. The kits commonly include inlet liners, ferrules, flame jets, gauges, cleaning items, adapters, and common tools for their removal and replacement.

	A mini-flashlight is very handy for inspecting the interior of inlets and detectors for obstructions, as well as for illuminating the oven interior. I prefer the type with the bulb on a flexible gooseneck. A small short LED flashlight, the kind that you might get as giveaways at conference booths, also makes an excellent GC oven light. No one yet has built a GC oven with a light that comes on when the door is opened. A flashlight also makes a good dropped small part finder, to locate ferrules, for example. Just place the flashlight horizontally on the laboratory floor, sweep it around, and look for the shadow.

	An artist's paintbrush with handle is handy to clean out debris from small areas inside detectors or inlets. Just watch that a fiber from the brush does not dislodge. It can also be used to apply leak-checking solution to fittings, although I don't recommended this practice, due to potential contamination of the gas stream with the leak-checking solution.

	Jumbo-size paper clips with smooth sides are convenient for blocking off inlet or detector fittings for testing purposes. Unbend the clip and attach it to the fitting with a nut and 1-mm i.d. graphite-vespel ferrule. With the column connection blocked off, you can pressure-check an inlet. A detector check can be run in this manner without column influences on noise or stability.

	A small pin vise and a set of drills can be used in an emergency to drill out a used ferrule, or to enlarge one that is too small to fit a column. Sometimes the small drills can help to remove a ferrule that is stuck in a fitting, or to remove debris from inside fittings or tubing ends.

	I keep some small needlenose pliers, a pair of larger multigrip pliers, and one pair of locking pliers in my toolkit. The larger gripping pliers are useful for holding a straight length of 1/8- or 1/4-inch metal tubing while cutting it, although I take care not to grip the tubing anywhere near a location where a connection is to be made, because the scratches from the pliers would make it impossible to get a good seal.

	Glass press-fit connectors make it easy to repair a broken column temporarily (until a replacement can be installed). These are available in many sizes to connect fused-silica tubing of the same or different diameter. They also connect a column with a retention gap. One manufacturer offers a vacuum–melting device that makes near-perfect connections.

	I have a conventional 0–60 psig pressure gauge with a syringe needle attached that I can insert into an inlet through the septum. Once in a while, I need to check the inlet pressure this way, instead of relying on the instrument's gauges or electronic pressure readouts.

	PTFE tape is used sparingly on tanks and interconnecting fittings where threads form the seal. Use two layers of tape, not more, and wrap them around the threads in the direction the nut tightens, so that the tape will be drawn into the fitting instead of getting pushed out. PTFE tape is never used in swage-type ferrule-sealed fittings, where it will only cause a leak, nor is it used at the high-pressure supply cylinder connection. Several types of this tape are available; be sure to select the right one.

	A rinsing reservoir may be useful for reviving bonded-phase capillary columns contaminated by soluble but non-volatile sample residues. Solvent rinsing is intended for use only after bake-outs and trimming of about 0.5 m of the inlet end plus the exit portion that enters the detector heated zone have failed to restore performance. Use only the solvents that the column manufacturer recommends, and rinse only from the detector end to the inlet end to avoid depositing contaminants deeper into the column. Check and revalidate column performance after rinsing.

	A small metal ruler measures the correct column penetration depth into an inlet or detector. Don't use a plastic ruler, because it might melt in contact with heated inlets or detectors. For convenience, make marks on the ruler that correspond to the correct inlet and detector depths. Several manufacturers offer capillary column installation gauges with the appropriate markings.

	A good sharp pair of scissors comes in handy for opening packages of ferrules, or for making paper stars out of waste paper that's waiting to be recycled while watching for peaks to be eluted. Scissors are never to be used to cut fused-silica columns (but you can believe that I've seen someone try it).

	I have three Phillips-head screwdrivers in the toolbox: large, medium, and small. The small one is part of a set of jeweler's screwdrivers with rotating handles. These are general purpose items that any laboratory should have on hand.

	I also keep three slotted-head screwdrivers. The small one is useful for securing electrical connections to screw-type terminals.

	This is a specialized tool for removing septa that become stuck due to heat-adherence to the inside of the septum area of an inlet. I have also used a dental pick for this purpose. Avoid using a flat-bladed screwdriver or a knife, because these may scratch the metal surfaces that seal to the septum.

	A static pad is a grounded, conductive plastic sheet onto which it is safe to place electronic components that must be protected from damaging electrostatic discharge. Any circuit boards removed from an instrument should be placed on a grounded static pad, or in a static-proof bag.

	A grounded static wrist strap prevents the technician from imparting a potentially harmful static discharge into instrumentation or components. Always wear one when working inside an instrument or removing components, and in all cases be quite sure that the instrument power has been removed while the instrument itself remains grounded.

	A digital stopwatch times bubbles in a bubble flowmeter, and also times an unretained peak. It's often more convenient to use a stopwatch when setting up an instrument than to operate the chromatography data system for each test injection. Select a stopwatch with readout to 0.01 second. Some GC systems include a stopwatch function on the display that includes flow, split ratio, and linear velocity calculations. These days, I just use my smart phone's stopwatch and timer functions, and then its calculator to find flow rates or average linear velocities. Good phone applications are available with additional chromatography functions.

	Recently I have used polyester swabs, mostly intended for cleanroom use, for cleaning out inlets and detector bodies. Take care that the solvents used are compatible with the swab material. Methylene chloride, for example, will dissolve polyester readily. I don't like to use cotton-tipped swabs on a wooden stick. Not only will the cotton release fibers, but the wood may release higher molecular weight compounds into cleaning solvents.

	I keep two manual syringes for setup purposes. One 10-µL gas-tight syringe is for injecting methane or butane to measure the unretained peak time and ascertain that the flame is lit and carrier gas is flowing. The other is for making liquid test-mixture injections as part of a column checkout. Sample syringes are kept separately.

	Syringe cleaning wires may be used in an emergency to clear septum particles or other debris from syringe needles. I recommend discarding stubborn contaminated syringes; take steps to keep the syringe clean instead.

	Column and detector test mixtures verify column performance and detector sensitivity. Keep a fresh vial of each type on hand, and put them in the chemical refrigerator if the label so directs. Column test mixtures are available for polar and for non-polar capillary columns, and there are test mixtures for each detector type. Some manufacturers provide a detector test mix that combines components for testing several different detectors. Once opened, test mixtures can be kept for a while in septum-sealed vials. Their lifetime is limited, because of gradual evaporation. If you keep test mix in a vial, remove the vial cap rather than puncture the septum when withdrawing liquid for injection. Some laboratories find it more convenient to keep dilute test mixtures on hand, because these are more easily disposed of than the concentrated mixes. Many laboratories have their own qualification and validation standards, of course, but the manufacturer's mixtures allow easy comparison to the factory test results, and may be used for a factory-level checkout.

GC Connections

Although manufacturers ship gas chromatographs with a collection of consumable parts and accessories, a number of other essential items should be on hand in every GC laboratory. What items are needed and how can they be used most effectively?

Essential GC Accessories

	Glycosylation is the most common monoclonal antibody (mAb) post-translational modification. It can affect the safety, half-life, and efficacy of mAbs. For example, fucosylation has a direct, negative impact on the overall efficacy of a mAb, by means of a reduction in its ability to perform antibody-dependent cellular cytotoxicity (ADCC), whereas the addition of N-acetylneuramic acid (NANA) generates acidic residues, and typically leads to an immunogenic response.

	Glycosylation occurs in two types, O- and N-linked. O-linked glycans are attached through an oxygen atom on a serine (Ser) or threonine (Thr) residue. This type of glycosylation is rare in human or murine immunoglobulin G (IgG) proteins. N-glycosylation can occur whenever the following consensus sequences occur: Asn-XXX-Ser or Asn-XXX-Thr (where XXX is any amino acid, except for proline [Pro]). The N-glycosylation site in mAbs is on Asn-297 in the fragment crystallizable (Fc) region. Glycan structure in mAbs can be highly heterogeneous, varying widely between different cell lines, clones, and production conditions. Therefore, it is important to be able to accurately characterize mAb glycoforms, including during biosimilar development.

	At the peptide level, reversed-phase high performance liquid chromatography (HPLC) analysis of the differing glycoforms produces a chromatogram in which the glycoforms are represented as several unresolved peaks, rather than a single peak. Further examination of this peak cluster allows fine details relating to the glycosylation profile to be extracted. It is worth noting that, even though reversed-phase LC is not an ideal technique for glycan separation, its high resolving power permits partial resolution of some isomers, and, at the peptide level, a near full approximation of the glycoprofile can be observed. As expected, an increase in the number of polar glycan units reduces retention in reversed-phase LC.

	Hydrophilic-interaction chromatography (HILIC) methodology is particularly applicable to the analysis of glycans, due to their highly polar nature. Typically, glycans are enzymatically released from the mAb, labeled with a fluorotag (2-aminobenzamide (2-AB) or procainamide), and then analyzed via a HILIC separation followed by fluorescence detection. It should be noted that 2-AB and procainamide are both chromotags and fluorotags, indicating that they can be used in conjunction with ultraviolet (UV) and fluorescence detection. However, fluorescence detection adds the benefit of increased sensitivity, due to fewer compounds fluorescing. When employing mass spectrometric (MS) detection, as is often the case during development and characterization, one may be tempted to forego the labeling step, as the unlabeled glycans readily ionize in negative mode electrospray ionization via the numerous terminal hydroxyl groups. However, performing the labeling is always recommended, as the unlabeled, or free, glycan can exist in two anomeric forms, an alpha and a beta, and this can lead to peak splitting. The benefits of labeling glycans is not merely from a detection perspective; the labeled glycans are also more easily retained and separated.

	Mass spectrometry is a powerful tool for the characterization of glycoproteins, and is often used in conjunction with other methodologies to allow full characterization. Glycosylation can be studied at both the intact protein and peptide levels, with MS methods employed depending on the information required and the known properties of the molecules. Intact antibodies or antibody fragments can be rapidly screened using electrospray ionization-quadrupole time-of-flight mass spectrometry (ESI-QTOF) instruments. The high resolution and extended mass range of these instruments allow glycan distribution, relative quantitation of specific glycoforms, and the degree of glycosylation to be assessed. Detection of major glycoforms is routinely performed; however, analysis of minor glycoforms is still challenging at the intact protein level.

	Analysis of glycosylation at the peptide level provides information such as glycosylation site and site-specific distribution of different glycoforms when multiple glycosylation sites are present. This approach also allows analysis of peptide sequences and other post-translational modifications. If glycosylation occurs at other sites other than the Fc region site, specific glycoforms can be analyzed at the glycopeptide level. Antigen-binding fragments (Fab) and Fc-specific glycan profiles can also be analyzed at the antibody fragment or released glycan level.

	ESI and matrix-assisted laser desorption–ionization (MALDI)-MS are both employed for profiling and structural characterization of glycans. A common phenomenon in MALDI is the loss of sialic acids; however, this can be avoided by permethylation or carboxyl group derivatization. ESI-MS of glycans is either carried out using an infusion or coupled to a separation technique, such as HILIC or capillary zone electrophoresis (CZE).

The Essentials

Glycosylation is the most common monoclonal antibody (mAb) posttranslational modification. What is essential to know about it?

An Introduction to Glycan Analysis

 on Good Documentation Guidelines are two sources for good documentation practice (GDocP). Are they similar, or are they complementary, and should we consider any other sources of advice?

	In the first seven parts of "Data Integrity Focus," we have focused on a few elements of data integrity (1–7). In this part, we will look at an essential requirement for data integrity: good documentation practice (GDocP). As it states in the principle of EU Good Manufacturing Practice (GMP) Chapter 4 on documentation:

Good documentation constitutes an essential part of the quality assurance system and is key to operating in compliance with GMP requirements (8).

	This statement can also be applied to good laboratory practice (GLP), or any other quality system, such as ISO 17025 or ISO 9001. Although this is a great statement, what does it mean in practice? What is good documentation, as opposed to bad documentation? In this part, we will look at the regulations and sources of advice on this key subject for data integrity.

	The principle of Chapter 4 describes the main types of documents that are possible in a regulated organization (8):

There are two primary types of documentation used to manage and record GMP compliance: instructions (directions, requirements) and records/reports. 

	As shown in Figure 1, the execution of an instruction results in the generation of records or a report. Underneath both instructions and records or reports, the types of document are described in more detail. It is the existence of accurate and reliable records that demonstrate that instructions have been followed, and hence the work is compliance with GMP regulations.


	Figure 1: Principles of good documentation practice from EU GMP, Chapter 4 (8). 

	Chapter 4 (8) goes into more detail about records and reports:

Records: Provide evidence of various actions taken to demonstrate compliance with instructions, e.g. activities, events, investigations, and in the case of manufactured batches a history of each batch of product, including its distribution.

Certificates of Analysis: Provide a summary of testing results on samples of products or materials together with the evaluation for compliance to a stated specification.

Reports: Document the conduct of particular exercises, projects or investigations, together with results, conclusions and recommendations.

	Therefore, from Chapter 4 (8), we can draw up some basic requirements for good documentation practices for any quality system:

 Have an instruction to document a repetitive task such as an analytical method or an SOP.

Follow the instruction and if there are any deviations from the procedure document them.

Generate a record to show that the instruction was followed.

	This chapter (8) also notes that "the term 'written' means recorded, or documented on media from which data may be rendered in a human readable form." In Chapter 4 speak, a document (instruction or records) does not exist only on paper; it can be any medium, as long as it can be converted into a readable format.

	Chapter 4 also has a section called "Good Documentation Practices," that include clauses 4.7 to 4.9 as shown in Table I. As you can see, the requirements are remarkably short and brief, and need to be interpreted by each regulated organization and laboratory. The focus of the regulatory requirements in Table I is generally on paper, and there is no specific mention of a computerized system, because the applicable regulations are found in EU GMP Annex 11 (9). There are other sections in Chapter 4 on the generation and control of documentation (clauses 4.1–4.6), records retention (clauses 4.10–4.12), and the expected types of documents (clauses 4.13–4.32) (8). This is a good source of information on what is expected in GMP documentation compared with the 21 

	Given the focus of this article is GDocP, you can see that there are not many "Attributable, Legible, Contemporaneous, Original, Accurate" (ALCOA) principles shown in Table I. Clause 4.7 equates to legible, 4.8 to contemporaneous, and 4.9. in part to accurate. We could conclude that Chapter 4 does not comply with the ALCOA principles! That is why Chapter 4, along with Annex 11, is being updated to enhance data integrity requirements (11).

	Having mentioned ALCOA in the previous paragraph, I need to explain what this means for those that do not know. ALCOA was developed by an FDA GLP inspector as an acronym for five requirements for data quality and is now used for data integrity. The four additional criteria were added by the European Medicines Agency on a GCP guidance on electronic source data (12) and called ALCOA+. The nine criteria are:

		Attributable: Who acquired the data or performed an action and when?

		Legible: Can you read and understand the entry or data?

		Contemporaneous: Is work is documented at the time of the activity?

		Original: Is there a printout or observation, or a certified copy thereof?

		Accurate: Are there no errors or editing without documented amendments?

		Complete: Are all data, including any repeat or reanalysis performed on the sample, available?

		Consistent: Are all elements of a chromatographic analysis, such as the sequence of events follow on and dates or time stamped in expected sequence, done in the same way over time, especially so as to be fair or accurate?

		Enduring: Are all data recorded in official media (either paper or a computerised system) , and not on the back of envelopes, cigarette packets, post-it notes, body parts or the sleeves of a laboratory coat?

		Available: Are all data easily retrieved for review, audit or inspection over the lifetime of the record?

 Chapter <1029> on Good Documentation Guidelines was issued in 2014 for public comment. The timing of this is significant, as it was issued before the publication of the tsunami of regulatory and industry guidance on data integrity (13–18) since the start of 2015. The structure of the final version of the general chapter released in 2018 (19) is shown in Figure 2. The high-level structure looks fine, but the listing order of the different types of GMP documents looks as if it has been drawn at random from a grab bag. Figure 2 attempts to put an order to the document, but it is not the same order as in the published version of 


	Figure 2: Structure of USP Chapter <1029> on good documentation guidelines (19).

	The focus of our discussion here is on the scope, principles of good documentation and data collection, and reporting sections. Also, you must bear in mind the numbering of the 

 general chapters: Those between <1> and <999> are mandatory, whereas those between <1000> and <1999> are informational or strong guidance. Hence 

 <1029> is an informational general chapter.

	Overall there are some good points in this general chapter, such as:

		This is the only guidance to mention how to document decimals and also refers to the rounding rules in the USP General Notices, an essential element of data integrity.

		The date format suggested is European: day, month, and year; an excellent approach to harmonization.

		There is a section on second person review of data, but there are also some gaps as can be seen below.

		There is no specific mention at all of the ALCOA+ criteria for data integrity. "Legible" and "accurate" are mentioned specifically, and "attributable" and "contemporaneous" are mentioned indirectly. This, in my view, is a failure; as Food and Drug Administration (FDA), World Health Organization (WHO), Pharmaceutical Inspection Cooperation Scheme (PIC/S), and Medicines and Healthcare Products Regulatory Agency (MHRA) guidances (14–16,20) all mention ALCOA or ALOCA+ specifically, why does 

 <1029> not use the term? The general chapter omits the ALCOA+ criteria complete and consistent in the principles of good documentation, although it does mention 

 in the discussion of purpose and in second person review.

		In the whole document, there is no mention of hybrid systems. Paper and electronic records are covered, but there is no explicit mention of hybrid systems. According to the WHO, hybrid systems are to be discouraged and should be replaced (15). According to me, hybrid systems are the worst possible option, because a laboratory must manage and synchronize signed paper printouts with the electronic records in the computer system (21).

		There is the possibility that predefined correction codes can be used for changes to data, but it does not mention that a procedure needs to define and maintain these codes.

		There is no discussion of the review of audit trails in hybrid and electronic systems

		Substitution of a true, complete, or verified copy of a record implies that the original can be discarded, because there is no explicit requirement for the original to be available for the second person review to check the copying process.

		Although it states that controls should be in place to protect record integrity, 

 <1029> does not differentiate between procedural and technical controls. For computerized systems, technical controls are preferred over procedural controls; this needs to be made clearer in the chapter.

		There is no specific mention of the need to control blank forms used in laboratory analysis, which is a regulatory expectation in a number of guidance documents (14,16,20). Also, there is a bullet point in 

<1029> that states that notebooks, data sheets, and worksheets should be traceable, but this is not specific enough, and can be confusing as to its meaning. A major regulatory concern is the use of blank forms; why is this not mentioned explicitly? For a detailed discussion on the implications of using master templates and blank forms, please read the "Questions of Quality" column by Burgess and McDowall (22), or a recent publication (21).

 <1029> mentions that computerized systems should comply with the requirements of 21 

 11 on electronic records and electronic signatures (23), there is a paragraph at the end of data collection and recording section that should state that print outs from computerized systems should be linked to the data and metadata the generated them.

 is not defined, and is used incorrectly. 

applies to all data and records collected during a GMP activity, not just initial observations. Please read several of my articles on this subject for further explanation (24,25), including Part 4 of this series (4).

	All in all, these gaps make the value of 

 <1029> doubtful. Therefore, we need to find better information. This comes from the WHO (15) and PIC/S (16) guidance documents, that we will consider now.

	WHO Guidance on Good Data and Record Management Practices

	The guide that offers the most comprehensive discussion of ALCOA and good documentation practices for both paper and electronic records is the WHO guidance (15). Appendix 1, comprising nearly half of the whole document, is entitled "Expectations," and examples of special risk management considerations for the implementation of ALCOA or ALCOA+ principles in paper-based and electronic systems are provided. For each of the five ALCOA requirements (attributable, legible, contemporaneous, original and accurate), there is a:

		table with the expectations for paper and electronic records

		discussion of any special risk factors to consider.

	This is an excellent guidance document, and if a reader wants to understand the meaning and scope of ALCOA, this is the place to start. The definitions for the five ALCOA criteria are given in Table II, and expand the ones outlined earlier in this article.

	The document jumps through hoops as it attempts to expand ALCOA to be ALCOA+, but still keep the five main criteria, as it expands 

 to include traceable and permanent (the latter equivalent to enduring).

	Good Practices for Data Management and Integrity (PIC/S PI-041)

	The PIC/S guidance document PI-041 is still a draft, with the third version issued at the end of November 2018 for comment (16), and therefore the content can change when the final version is issued. This is a very comprehensive guidance document, and to understand the good documentation and record keeping practices there are the following sections:

		Section 8 covers paper records and the regulatory requirements for this medium.

		Section 9 considers the data integrity requirements for computerized systems. Hybrid system requirements are specifically mentioned in section 9.8.

	Despite its draft status and the possibility of change following industry comments, it is worth reading to get a comprehensive view of data integrity. When the final version is issued, PI-041 will be one of the best regulatory data integrity guidance documents.

Feature Article

Are EU GMP Chapter 4 and USP Chapter  on Good Documentation Guidelines similar or complementary? Should we consider any other sources of advice?

Data Integrity Focus, Part VIII: What is Good Documentation Practice (GDocP)?

In on-site environmental applications, representative sampling and proper replication are essential to properly characterize a given site. For these reasons, recent work in thin-film solid-phase microextraction (TF-SPME), an extension of fiber-based SPME, has focused on the development of unique holders and customized samplers that have been tailored for distinct sampling environments. Here, we explore the latest developments and designs of high-surface-area thin-film samplers for on-site environmental analysis using TF-SPME.


	Figure 1: Compilation of various thin-film solid phase microextraction (TF-SPME) sampler formats including: (a) carbon mesh supported TF-SPME membrane on six-replicate drill sampler (7), (b) TF-SPME in-bottle grab sampler (6), (c) TF-SPME time weighted average sampler with hydrophilic lipophilic balance-particle-polyacrylonitrile (HLB-PAN) coated thin film blade (8), (d) TF-SPME coated bolt sampler in sampling (open) and storage (closed) position with HLB–PAN coated bolts (10). 

	We all know the saying, "Garbage in equals garbage out." With the proliferation of ultrahigh-pressure liquid chromatography (LC) systems and ever more sensitive mass spectrometry detectors, this adage that is more commonly thrown to the wayside in favor of simpler sample introduction approaches such as dilute-and-shoot. It can be easy to forget that where sampling and sample preparation precede instrumental analysis, any errors experienced during these crucial steps will affect the results, possibly leading an analyst to draw a false conclusion regarding his or her work. This holds especially true for on-site environmental applications, where representative sampling and proper replication are essential to satisfactorily characterize a given site (1,2). For these reasons, recent work in thin-film solid phase microextraction (TF-SPME), an extension of fiber based SPME, has focused on the development of unique holders and customized samplers that have been tailored for distinct sampling environments (3,4). These include more rigid carbon fabric supported membranes to give physical stability for high river flows and agitation (5), a six-replicate TF-SPME membrane holder that attaches to a specialized sampling case or standard power drill (6,7), in-bottle TF-SPME samplers for surface water grab sampling (6), broad spectrum sorbents for the simultaneous extraction of polar and nonpolar analytes (7), time-weighted average (TWA) membrane holders for long duration TWA river sampling (8), and even highly durable thin-film-coated bolts for the sampling of extreme environments (9,10). These devices are all shown in Figure 1. Much like fiber-based SPME, TF-SPME employs the fundamentals of thermodynamic equilibrium to drive the passive diffusion of small organic molecules, defined by the diffusion coefficient (

), across a static boundary layer of thickness (δ) that separates the free flowing sample matrix of analyte concentration (

) from the open sorbent bed. However, unlike an SPME fiber, thin-film devices are designed to have a total surface area (

) that is much larger than the sorbent thickness, resulting in extraction rates (dn/dt) that are orders of magnitude faster (11,12). This relationship is described mathematically in equation 1 below. Increased extraction kinetics are important to maximize the sensitivity of on-site environmental sampling approaches where one may be constrained to only a few minutes to collect each sample.

	Appropriately, it was shown experimentally that when short, 15-min pre-equilibrium extractions were performed from water that had been spiked with various pesticides, a 40 mm × 4.8 mm (L × W) carbon fabric supported divinylbenzene/polydimethylsiloxane (DVB/PDMS) TF-SPME membrane could extract 21.2, 19.8, 18.5, 18,4, 26.8, and 23.7 times the amount of 2,4 dichlorophenol, 2,4,6 trichlorophenol, phorate D10, fonofos, chloropyrifos, and parathion than a comparable 65 µm DVB/PDMS SPME fiber respectively (5).This result was inline with what was expected fundamentally (equation 1), as the aforementioned TF-SPME membranes had a surface area that was approximately 25 times greater than that of the corresponding SPME fiber. Moreover, when directly compared to standardized liquid–liquid extraction, adopting a slightly modified US-EPA 8270 methodology, these sensitive membranes were shown to give MLOD values that were anywhere from 4 to 200 times lower than those achievable using LLE at a Standards Council of Canada accredited laboratory, while maintaining a similar level of accuracy in a double-blind split comparison (13). In fact, this benchtop TF-SPME methodology also required far less sample with all analytes being extracted simultaneously by directly immersing the membrane on a cotter pin into 30 mL of spiked surface water at a pH of 2.5, with 10% NaCl and a stir rate of 900 rpm. This was in sharp contrast to the LLE approach, which required three sequential 50 mL dichloromethane extractions from 800 mL of spiked surface water at acidic, neutral, and basic conditions. With these results in mind, similarly designed membrane holders, shown in Figure 2, were included with the recent commercial release (Gerstel US) of the carbon mesh supported TF-SPME membrane (14).


	Figure 2: In-vial thin-film solid phase microextraction (TF-SPME) membrane holders including: (a) prototype cotter pin TF-SPME holder pushed through septa used for inter-laboratory study (13), (b) comparable commercial membrane holder for direct immersion extraction from 40 mL vial, (c) TF-SPME holder set-up for 10 mL headspace extraction (14), (d) TF-SPME holder set-up for 10 mL direct immersion extractions, (e) TF-SPME holder for 40 mL vial, (f) TF-SPME holder for 10 and 20 mL vials.

	To further improve the applicability of TF-SPME related methods, subsequent research focussed on the development of more novel TF-SPME holders to allow for unique sampling opportunities. In a recently published study, Piri-Moghadam and associates compared an in-bottle TF-SPME methodology to on-site sampling approaches with and without hand portable gas chromatography-mass spectrometry (GC–MS) instrumentation (6). This in-bottle approach was devised in a manner such that the sampling process was not a major departure from conventional grab sampling. When applying the bottle sampling approach, a technician simply needs to place the TF-SPME–equipped PTFE adapter, shown in Figure 1b, and spike an appropriate internal standard into the 800 mL glass bottle when collecting a surface water sample. Following sampling, these bottles were then allowed to sit on an orbital shaker for three days pending analysis by use of a Gerstel thermal desorption unit and cooling injection system (TDU-CIS4) equipped Agilent 6890-5973N GC–MS instrument. As shown in Table I, this in-bottle approach gave the lowest limits of quantitation observed in the study with MLOQ values typically ranging between 1–10 ppt for most analytes. A more novel approach also explored the use of the custom six-replicate TF-SPME membrane holder shown in Figure 1a to perform on-site extraction directly from river water. These membranes were then either analyzed directly on-site using a hand-portable Tridion-9 GC–MS instrument, or transported back to the laboratory for desorption and analysis using the aforementioned TDU-CIS4 equipped Agilent 6890-5973n benchtop GC–MS instrument (6). Although the targeted, banned pesticides could not be detected at appreciable levels from Ontario Rivers during the study, a comparison of the MLOQ values for a selection of these compounds indicated that the entirely on-site TF-SPME methodology could provide quantitative limits similar to those of conventional benchtop LLE and SPME. The comparison of these methods and associated MLODs are listed in Table I.

	Following the optimization and validation of these carbon mesh supported TF-SPME membranes and their various specialized holders, research efforts were shifted toward the development of more sensitive sorbents, particularly those with a broader affinity for polar analytes. Appropriately, hydrophilic lipophilic balance (HLB) particles, based on a co-polymerization of N-vinylpyrrolidone onto a divinylbenzene backbone, made for a logical choice to compare to the now validated DVB/PDMS carbon mesh supported membranes (7). In this study, three types of HLB particles, differing in their particle and pore size, were compared to the established DVB-based thin-film membranes. From the three HLB particles tested, one came from a commercial source, while the other two were synthesized in-house using either a precipitation, or suspension polymerization methodology. All particles were suspended in PDMS, spread onto the carbon fabric support and cut to 20 mm × 4.8 mm to fit the TDU-CIS4 system as before.

	It was observed that the commercial HLB particles, those prepared by suspension polymerization, and the DVB particles had similar specific surface areas (727–800 m

/g), but demonstrated greatly different extraction capabilities for the targeted McReynolds mixture. To better understand this result, complete characterization of the particles using Fourier-transform infrared spectroscopy, elemental analysis and Brunauer-Emmett-Teller analysis were performed, indicating that despite having a similar proportion of the N-vinylpyrrolidone moiety, the diameter of the pores and total pore volume for the sorbents were substantially different. HLB particles synthesized by suspension polymerization and obtained from commercial source had pore sizes of 80 and 71 angstroms (Å) and pore volumes of 1.30 and 0.64 mL/g, respectively, making them generally mesoporous. In the meantime, the DVB particles had the largest pores, with a diameter of 400 Å and volume 1.54 mL/g. Finally the smallest pores were observed in those particles prepared by precipitation polymerization with an average pore diameter of 13 Å and volume of 0.20 mL/g. Surprisingly, despite having the smallest pore volume, the HLB particles synthesized by precipitation polymerization gave the best results, extracting nearly double the amount of the polar standards than the DVB/PDMS membranes (Table II). This result was attributed to the fact that the precipitation polymerized HLB particles had a generally microporous structure stemming from the use of acetonitrile as a porogen. Moreover, the smaller 1.33 µm particle diameter could have also played an important role considering the spread distribution and presence of a higher number of particles on the membranes. Unsurprisingly, as shown in Table II, it was found that all of the HLB sorbents tested did in fact outperform the established DVB particles for all of the McReynolds standards tested. This increase in the affinity for semipolar analytes can be attributed to the presence of the N-vinylpyrrolidone group. These groups provide sites for hydrogen bonding, and allow the hydrophilic functionalities to interact more strongly with the sorbent. Although not as pronounced, it was somewhat surprising to also see slightly increased performance offered by the HLB particles for the octane standard added to the McReynolds mixture. With a LogP value of 5.01, one would expect octane to be primarily extracted by the nonpolar interactions offered by the divinylbenzene. It is also important to note that instrumental runs were fully randomized during the experiment, ruling out the possibility of detector drift during experimentation. Speculatively, it is possible that this result could also be explained by the much smaller pore diameter present in all of the HLB materials tested.

	Interestingly, it was also found that the physical characteristics of the carbon mesh support allowed for an innovative calibration approach for converting raw instrument signal to tangible amounts such as nanograms. This calibration approach, called 

, allowed for liquid standard to be introduced to the GC–MS instrument, without having to remove the TDU-CIS4 system or install a liquid injection syringe onto the autosampler. Instead, it was possible to manually spike a few microliters of a methanolic McReynolds standard directly onto the edge of the TF-SPME membrane, that then wicked the solution into the carbon mesh support. Once analyzed, it was found that an exceptional correlation to response (

 ≥ 0.996) for all analytes could be achieved while most of the volatile methanol solvent was purged from the CIS4 during analyte reconstitution, never reaching the GC column. Moreover, it was also found that once extracted onto the TF-SPME membranes and stored within the Gerstel thermal desorption tubes on the autosampler rack, all of the McReynolds standards were stable on-membrane for up to 24 h. In fact, only the photosensitive pyridine was shown to deplete when this test was further lengthened to 120 h.

	Having shown favorable results for polar VOC analysis, these HLB/PDMS membranes were then applied for an untargeted, on-site determination of chlorination byproducts from a private hot tub. For this proof of concept application the HLB/PDMS thin films prepared using the precipitation polymerized HLB were cut to 40 mm × 4.8 mm (L × W) for analysis on the Tridion-9 portable GC–MS system. As shown in Figure 3, sampling was accomplished by placing four membranes onto the six-replicate TF-SPME holder, which was connected to a sampling case (PAS Technologies GmbH), allowing for extractions to be performed at 2000 rpm for 10 min from the 39.5 °C, pH 7.2–7.4 hot-tub water. Immediately after sampling, the membranes were removed from the drill, and dabbed dry with a Kimwipe before being placed in the 3.5-in. thermal desorption tubes. These tubes were then inserted into the SPS-3 module and desorbed at 250 

min of helium to transfer the desorbed analytes to a 19 gauge Tenax/Carboxen needle trap device (NTD). This NTD could then be placed directly into the Tridion-9 portable GC–MS instrument for desorption and analysis of the compounds transferred from the TF-SPME membranes. In total, six different chlorination by products were identified from the hot tub water. This identification was performed by matching with the NIST 2011 mass spectral database, followed by preliminary verification by use of a standard n-alkanes linear retention index plot. These compounds included chloroform, bromodichloromethane, dichloroacetonitrile, chlorobenzene, benzonitrile, and benzyl chloride, all of which were known disinfectant by-products.

In on-site environmental applications, representative sampling and proper replication are essential. For these reasons, recent work in thin-film solidphase microextraction has focused on the development of unique holders and customized samplers that are tailored for distinct sampling environments. Here, we explore the latest developments.

LCGC North America-09-01-2019

Vol 37 No 9 LCGC North America September 2019 Regular Issue PDF

Effects of Sample Diluent on Analyte Recovery in Reversed-Phase and HILIC Separations

Essential GC Accessories

An Introduction to Glycan Analysis

Data Integrity Focus, Part VIII: What is Good Documentation Practice (GDocP)?

Frontiers of Sampling: Design of High Surface Area Thin-Film Samplers for On-site Environmental Analysis

Shodex Rapid SUGAR Series: Fast Fermentation Monitoring and Sourdough Cultures

Determination of Triphenylmethane Dyes from Aquaculture Samples