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Vol 32 No 9 LCGC Europe September 2019 Regular Issue PDF

Metabolomics, the comprehensive analysis of all metabolites and intermediate products of reactions present within a biological system, is a promising field to enable precision medicine. Clinical metabolomics faces two main challenges at the bioanalytical level. The first is the need for high resolution to obtain maximum metabolome coverage. This is exemplified by the latest version of the Human Metabolome Database (HMDB), which reports more than 110,000 metabolites and endogenous compounds. The second is the high-throughput needed to enable the analysis of a large number of samples typically encountered in large-scale cohort studies. Reversed-phase liquid chromatography (LC) at regular or ultrahigh pressures combined with high-resolution mass spectrometry (HRMS) has long been considered the “gold standard” in metabolomics. However, these conventional reversed-phase LC–MS approaches are no longer sufficient to analyze the vast variety of polar compounds, as well as discriminate closely related compounds such as isomers or enantiomers. This review article discusses the novel separation and detection strategies that are considered promising in clinical metabolomics to enhance the metabolome coverage. It includes hydrophilic interaction chromatography (HILIC), supercritical fluid chromatography (SFC), multidimensional LC approaches, as well as ion-mobility mass spectrometry (IM-MS) and data-independent acquisition (DIA) analysis methods.

Metabolomics, first formally introduced in the early 2000s and described as the comprehensive analysis of all metabolites present within a biological system, has attracted a growing interest over the last decade in clinical research. Together with other “omics” approaches, such as genomics and proteomics, metabolomics plays a key role in the implementation of personalized medicine. Two approaches are typically considered in metabolomics. In targeted metabolomics, known metabolites from given biochemical pathway(s) are measured in a quantitative manner. Untargeted approaches, on the other hand, focus on the global and unbiased analysis of the highest number of compounds included in the metabolome leading to qualitative and semiquantitative information (relative differences between populations). Both approaches have been increasingly used over the last couple of years in personalized medicine and drug discovery, in the aim of finding new metabolite biomarker candidates for earlier and more accurate diagnosis, improve the prognosis and staging of diseases, and increase the global understanding of pathophysiological processes via the discovery of novel biomolecular pathways (1,2).

	In 2017, the fourth version of the Human Metabolome Database (HMDB 4.0) covered more than 110,000 fully annotated metabolites. This is a threefold increase compared with the previous release of HMDB 3.0 in 2013 (3). The human metabolome is very complex and comprises a large diversity of compounds, including amino acids, organic acids, nucleosides, lipids, small peptides, carbohydrates, biogenic amines, hormones, vitamins, and minerals. Moreover, xenobiotics such as drugs, cosmetics, contaminants, pollutants, and their respective phase-I and phase-II metabolites are also part of this metabolome. The (ideally) comprehensive analysis of the metabolome is therefore linked to several analytical challenges due to (i) the large differences in physicochemical properties (polarity, solubility, pKa values, molecular mass), (ii) the broad dynamic range needed to analyze both trace compounds and highly abundant metabolites (up to nine orders of magnitude difference), and (iii) the presence of multiple isomers with structural similarities but significant differences in their biological activities (lipid-based signalling molecules) (4).

	Overall, this complexity highlights the need for stateâofâtheâart analytical approaches capable of tackling such challenges and enabling a qualitative and quantitative assessment of the metabolome. This should be done with the highest possible metabolic coverage via high resolving power and selectivity. In this context, metabolomics has strongly benefitted from the latest developments in the fields of both chromatography and mass spectrometry (MS) over the last two decades. The use of reversedâphase liquid chromatography (LC) columns equipped with sub-2-µm fully porous particles (ultrahighâpressure liquid chromatography, UHPLC) or sub-3-µm superficially porous particles (core–shell technology) are now considered wellâestablished methods in metabolomics owing to the dramatic improvements in resolution and throughput obtained with such phases compared with conventional high performance liquid chromatography (HPLC) (5–7). On the other hand, recent liquid-based chromatographic and MS innovations, notably within hydrophilic interaction chromatography (HILIC), supercritical fluid chromatography (SFC), multidimensional LC, ion-mobility mass spectrometry (IM-MS), and data-independent acquisition (DIA) approaches, are not widely used in metabolomics, despite the significant improvement in metabolite coverage expected with such techniques. Therefore, in this review article, the latest developments in the aboveâmentioned fields of chromatography and MS are discussed with a focus on their ability to increase the metabolome coverage.

Improvement of Metabolic Coverage: Chromatographic Innovations

	Hydrophilic Interaction Chromatography: Reversed-phase LC-based methods have long prevailed in metabolomics because of the large variety of column chemistries available, the ease of use, and retention time reproducibility. However, a large number of polar or ionizable metabolites, such as amino acids, small organic acids, nucleosides, phosphate derivatives, or saccharides, are not well-retained using reversed-phase LC. Still, many of these polar metabolites play an essential role in multiple (patho)physiological processes, showing the need for alternative approaches. HILIC, a technique first proposed by Alpert in 1990 (8), is well-suited for the analysis of polar compounds. Retention is based on a multimodal separation mechanism between a polar stationary phase and a relatively hydrophobic mobile phase composed of an aqueous–organic mixture with a high organic proportion. With a concentration of 5–40% of water in the eluent, a water-enriched layer is formed at the surface of the stationary phase, facilitating analyte partitioning between this stagnant phase and the bulk mobile phase. The exact mechanisms involved in retention and separation are not fully understood but mostly rely on hydrophilic partitioning, dipole-dipole interaction, hydrogen bonds, and electrostatic interactions (depending on the stationary phase chemistry) (5,9).

	A large diversity of phase chemistries based on silica or polymer material modified with polar functional groups, for example, aminopropyl and amine, amide, diol, triazole, sulfobetaine, phosphorylcholine, hydroxyethyl, and sulfoethyl are nowadays commercially available for HILIC analysis. Whereas for reversed-phase LC analyte retention can be easily predicted, helping to facilitate method development, this remains difficult in HILIC. The chromatographic selectivity is also strongly dependent on the stationary phase chemistry and composition of the mobile phase, as illustrated in Figure 1. A careful and extensive screening of different conditions during method development using a large set of representative metabolites is therefore recommended to obtain an adequate metabolite coverage. The help of modern computer-assisted method development strategies, such as the predictive elution window shifting and stretching (PEWS

) approach (10), could be useful here to speed up method development. Numerous studies comparing the different stationary phases for metabolomics applications showed that diol, amide, and zwitterionic phases usually give the best results in terms of metabolite coverage, therefore representing a good starting point in the method development process (11,12). Small organic acids, sugar phosphates, and nucleosides are difficult to analyze with reversed-phase LC. Using HILIC mode these compounds are better retained, especially with polymeric zwitterionic phases, which allows analysis at high pH (pH 9–10) thanks to the polymeric nature of the stationary phase (13). Adding phosphate at micromolar concentrations to the mobile phase has also shown to further improve the peak shape and sensitivity when analyzing such metabolites with a zwitterionic phase (14). Next to the stationary phase chemistry, the composition of the mobile phase strongly influences the selectivity and quality of the separation. Acetonitrile is the optimal organic solvent because it is water-soluble and aprotic. Protic solvents such as methanol, isopropanol, and ethanol are not recommended due to competition with water for the solvation of the stationary phase, which may lead to lower analyte retention (15,16). In HILIC, the high proportion of acetonitrile in the mobile phase decreases its viscosity compared with the mobile phase mixtures used in reversed-phase LC, which offers additional advantages such as the possibility to use longer columns (leading to higher efficiencies), a higher electrospray ionization (ESI) sensitivity, and higher volatility (15). The buffer composition (that is, salts concentration and pH) has a strong impact on both selectivity (Figure 1) and retention time reproducibility. The buffer concentration (commonly ≤50 mM to avoid salt precipitation in acetonitrile) influences the thickness of the water layer and thus the hydrophilic interaction, and plays an essential role in electrostatic interactions. Ammonium formate and acetate buffers are commonly used because of their MS compatibility. They also give better peak shapes than the corresponding acid solutions (16). Trifluoroacetic acid is not recommended in HILIC–MS because it leads to strong ion suppression in the range of compounds studied. Finally, an adequate and repeatable buffer pH is crucial in HILIC to ensure reproducible analyses. Changes in buffer pH will lead to a higher retention variability, showing the importance of repeatable procedures when preparing the buffer solutions.

	Despite all the above mentioned advantages and the improved metabolite coverage that can potentially be obtained using the technique, HILIC remains sparsely used in metabolomics, mostly confined to untargeted studies (16,17). The complex mechanism of HILIC separation, the longer equilibration times, the attention required to ensure reproducibly prepared mobile-phase buffers, and the challenges in finding an adequate sample injection solvent might explain why this technique has not been widely adopted yet. However, there are now numerous excellent reviews available discussing these challenges, offering solutions and providing guidelines for state-ofâthe-art HILIC analysis (1,9,15,16,18). This will hopefully foster the use of HILIC in routine metabolomics.

Although not new-the use of fluids in their supercritical state was first reported in the 1960s-SFC has shown a spectacular comeback in the last decade. This is due to the introduction of a new generation of instruments capable of performing robust, reproducible, reliable, and quantitative analysis. Similar to what has been observed in conventional LC, these new instruments have also fostered the development of columns packed with subâ2âµm fully porous (ultrahighâperformance SFC, UHPSFC) and sub-3-µm superficially porous particles specially designed for SFC analysis. Moreover, the new source designs recently developed for interfacing SFC with MS have also strongly contributed to developing the use of SFC in bioanalysis, including metabolomics (19,20). The metamorphosis of the technique has transformed UHPSFC–MS into a very competitive separation approach, complementary to UHPLC–MS, as underlined in the first ever inter-laboratory study. Between the 19 participating laboratories, similar or even better repeatability and reproducibility using SFC was shown for the determination of impurities in pharmaceutical formulations compared with conventional LC methods (21).

	Supercritical fluids have unique properties that take advantage of both gas and liquids, with viscosity and diffusivity very close to those of a gas, while their density and solvating power is close to those of a liquid. Overall, these inherent characteristics enable high separation efficiency at high mobile phase velocity with a low back pressure generated, and good solvation and fast transportation of the analytes (22). Carbon dioxide has been considered the solvent of choice as a result of the low critical temperature and critical pressure (31 °C and 74 bar, respectively) as well as its low toxicity, low flammability, and environmentally friendly properties. However, the low polarity of pure CO

 limits its application to the analysis of rather nonpolar or hydrophobic compounds such as lipids. The addition of a miscible co-solvent (referred to as 

, typically methanol) to the mobile phase is an adequate strategy to enable the retention of polar compounds. The use of an organic coâsolvent influences the solvating power of the mobile phase, its hydrogen-bonding donor and acceptor properties, its density, the interaction between analytes and mobile phase, as well as the adsorption of analytes on the stationary phase (20,22). Yet, adding a modifier to this supercritical fluid increases the critical temperature and pressure of the fluid. In current applications, the pressure is commonly maintained over its critical point while the temperature is below its critical value. In this case, the fluid is in a subcritical state, showing a chromatographic behaviour close to LC.

	The addition of acids (formic acid, citric acid), bases (trimethylamine, isopropylamine), or salts (ammonium acetate, ammonium fluoride) at low concentrations in the modifier also increases the range of compounds that can be analyzed using SFC, especially ionizable compounds such as polyacids, aliphatic amines, and other polar metabolites (20,22). These additives also increase the separation efficiency and peak shape by acting as ionâpairing agents and by covering active sites on the stationary phase, leading to less tailing and better elution of polar compounds. The latest trend in SFC is the use of water as an additive in a CO

–methanol mobile phase to improve peak shape (at a proportion of 1–5%, miscible in the mobile phase) or to enable the elution of very polar compounds (up to 30%, forming a ternary mixture) (20,22).

A number of SFC-specific stationary phases have been developed and commercialized in the past years, including 2-ethylpyridine, 4-ethylpyridine, pyridine amide, amino phenyl, 2-picolylamine, diethylamine, diol, and 1-aminoanthrocene. In addition to SFC-specific columns, reversed-phase LC- and HILICâtype stationary phases (ethyleneâbridged silica, C18, fluorophenyl, amide) can also be used in SFC.

	Most of the SFC-specific columns are also available in sub-2-µm format. However, the extracolumn band broadening of the state-of-the-art UHPSFC instruments currently on the market are still higher than the corresponding values obtained on UHPLC systems (namely, 85 µL

), hindering the use of typical UHPLC column dimensions (50 mm × 2.1 mm, 1.7-µm) with these systems. On the other hand, 4.6-mm internal diameter (i.d.) columns require flow rates above the system limits (22). Therefore, most of the current state-of-the-art SFC applications are performed using 100 mm × 3.0 mm columns packed with sub-2-µm fully porous and sub-3-µm superficially porous particles, which represents an adequate compromise and can lead to excellent kinetic performance with a low pressure drop, as illustrated in Figure 2 (23).

Modern UHPSFC–MS analysis has recently started to gain more attention from the metabolomics community, not only in the fields of lipidomics but also as a complementary technique to UHPLC–MS to increase the metabolome coverage. Multiple metabolite classes, including amino acids, bile acids, cannabinoids, fatty acids, saccharides, steroids, and tocopherols, have been successfully analyzed using UHPSFC–MS (24). A good example of the potential of UHPSFC was described by Holcapek and co-workers, who demonstrated the comprehensive and quantitative analysis of different lipid classes (25). Figure 3 shows the chromatogram obtained for the analysis of lipid internal standards with UHPLC–HILIC–MS (Figure 3[a]) and with UHPSFC (Figure 3[b]), both coupled to a quadrupole–travelling-wave ion mobility–time-of-flight (TOF) mass analyzer. Both HILIC and UHPSFC enable the separation of lipid classes without the typical overlap that is seen when using conventional reversedâphase LC approaches. Nonpolar lipids (cholesterol esters and triglycerides) as well as species with one hydroxyl group (ceramides, diglycerides, monoglycerides, and cholesterol) show poor retention in HILIC and elute in the void volume (Figure 3[a]). On the other hand, all lipids are retained using UHPSFC (Figure 3[b]). Whereas the positional isomers 1,2-DG/1,3-DG and 1,2-MG/1,3-MG are well resolved in UHPSFC (Figure 3[b]), the positional isomers of the more polar lysolipids 1-LPG/2-LPG, 1-LPE/2-LPE, and 1-LPC/2-LPC are well resolved using HILIC (Figure 3[a]) (25). This example illustrates the complementary nature of the different chromatographic modes, where no single analytical technique currently enables a comprehensive coverage of the metabolome.

	Despite the excellent performance that can be reached with modern UHPSFC–MS, it remains little used in metabolomics-and only by a limited number of research groups. This reluctance might be explained by the large diversity of stationary phases currently available, together with the flexibility offered in the composition of the mobile phase (modifier, additives, gradient composition). Indeed, the method development step a priori may be seen as very cumbersome and time-consuming. However, it can be facilitated by using column classification maps to help select adequate column chemistries (26) and by using method optimization work (27).

	Overall, there is no doubt that the multiple advantages of modern SFC will foster its use in metabolomics in the coming years as a complementary chromatographic approach to expand the metabolome coverage. The advantages of SFC in metabolomics include (i) its application range versatility, with a large range of metabolites with very diverse physicochemical properties that can be analyzed within a single run (as shown in reference 27), (ii) the sample compatibility with the mobile phase used in SFC, (iii) the excellent sensitivity of UHPSFC–MS, comparable or superior to that of UHPLC–MS, and (iv) the flexibility offered with the state-of-the-art instruments, which allows for both UHPLC and UHPSFC analysis within one single system and an unlimited combination of solvent and stationary phases. SFC technology has faced the same reluctance as HILIC a decade earlier, but both techniques are promised to rise further in the field of metabolomics.

Multidimensional Chromatographic Separations: 

A straightforward approach to increase the metabolome coverage of very complex samples or closely related metabolites is to add another separation dimension to provide additional selectivity. Similar to what has been observed in the field of SFC, onâline two-dimensional liquid chromatography (2D-LC) is far from being a new concept but has seen a significant breakthrough in the last couple of years thanks to significant advances in theory and instrumentation. In on-line 2D-LC, two individual LC separations are combined, typically using a four-port duo valve or 10âport valve with two sampling loops, or connected to special valves with multiple sample parking loops.

	Capturing all peaks-or a large number of fractions-from the first dimension into the second dimension is referred to as comprehensive 2D-LC (also called LC×LC), while multiple heart-cutting 2D-LC (also called LC-LC) is used when one or few distinct fractions are collected from the first dimension and are subjected to a high-resolution analysis in the second dimension. Selective comprehensive 2D-LC (sLC×LC) is an intermediate approach where a series of fractions across one or more regions in the first dimension chromatogram are transferred to the second dimension (28,29). The comprehensive LC×LC approach appears particularly interesting in untargeted metabolomics, where hundreds of features can be profiled during one single analysis.

	A large diversity of chromatographic modes can be combined in 2D-LC, including reversed-phase LC, HILIC, normal-phase LC, ion-exchange chromatography (IEX), ion-pairing chromatography, and porous graphitized carbon (PGC) columns. Different stationary phase chemistries and mobile-phase compositions can be employed, aiming for the highest orthogonality of separation between the two dimensions. The selection of the two separation dimensions depends on the analytes, the compatibility and miscibility of the mobile-phase solvents, the compatibility with the detector, and the selection of a faster technique (that is, based on UHPLC conditions) for the second dimension (4,28).

	With the recent advent of state-of-the-art instruments for 2D-LC analysis, the number of experimental parameters that can be optimized during method development has dramatically increased. Indeed, setting up a complete 2D-LC method requires optimization of multiple parameters, including column dimension, stationary phase, particle sizes, mobile-phase composition, gradient conditions, sample loop volume, injection volumes, flow rates, and modulation times. This can lead to a rather cumbersome and lengthy method development. Moreover, state-of-the-art 2D-LC analyses usually require a dedicated instrument (even though one-dimensional [1D]-LC systems can be upgraded to 2D-LC with only minor investment). Finally, hyphenating 2D-LC to MS adds another challenge, since the insertion of an additional LC dimension may induce a significant dilution of the effluent injected to the MS system (28,30).

	Overall, this might explain the reluctance in using this technique in metabolomics, despite the remarkable promises 2D-LC holds in significantly expanding the metabolome coverage. This reluctance is similar to the one observed for HILIC and SFC in metabolomics, where inexperienced users are struggling to get reproducible data. Moreover, they might lack sufficient theoretical and practical knowledge to get the best out of those techniques. However, 2D-LC is currently a very dynamic field and a number of excellent guidelines and tutorials have recently been published by experts in the field, guiding the inexperienced user through this method development (28–31). The recent developments in instrumentation, including the use of active-modulation techniques to alleviate the MS detector sensitivity problems and minimize effects from poorly compatible mobile phases, software tools to support method development, as well as continuous improvements in the algorithm available for processing 2D chromatograms, will certainly foster its use in clinical metabolomics. Most of the applications reported so far have been mostly based on heart-cutting approaches and proof-of-concept studies rather than clinical applications. However, the results presented highlighted the potential of 2D-LC in metabolomics, showing for instance, a twofold increased coverage of intracellular energy metabolites using a combination of reversed-phase LC with PGC (32) and the acquisition of both metabolomic and lipidomic information in a single analysis using heart-cutting 2D-LC (33).

 Improvement of Metabolic Coverage: MS Developments

	Ion-Mobility Mass Spectrometry: Among all recent developments discussed here, IM-MS is probably the one that has already been largely accepted by the metabolomics community even though it remains a relatively young technique. IM-MS adds an orthogonal separation dimension between chromatographic separation and MS detection without impacting the analysis time. IM-MS separation occurs in a timescale of milliseconds, which makes this technique fully compatible with both fast LC and high-throughput MS approaches (especially TOF mass analyzers, which offer fast duty cycles) (34).

	IM-MS is a gas-phase technique separating ions driven through an ion mobility cell under an electric field in the presence of an inert buffer gas. Ions are separated according to their mobility or drift time, which is intrinsically linked to their size, shape, and charge. Assuming that the experimental parameters (for example, drift-tube length, gas pressure, temperature, electric field) are constant, the ion drift time is proportional to the rotationally averaged collision crossâsection (CCS) value, which represents the effective area involved in the interaction between an ion and the gas present in the ion mobility cell. The CSS value is not only highly reproducible but also unique for each analyte, and reflects its chemical structure and three-dimensional configuration. This shows the power of IM-MS in metabolomics, especially in untargeted metabolomics, where CCS values can be used in addition to conventional parameters typically reported in libraries (retention time, mass-to-charge ratio, fragmentation pattern) for metabolite characterization and to increase the confidence in metabolite identification (4,35).

Different IM-MS technologies are currently commercially available, namely, (i) drift-tube ion-mobility spectrometry (DTIMS), (ii) travelling-wave ion-mobility spectrometry (TWIMS), (iii) field-asymmetric ion-mobility (FAIMS), also called differential-mobility spectrometry (DMS), (iv) differential mobility analyzer (DMA), and (v) confinementâandâselectiveârelease ion mobility, also called trapped ion mobility spectrometry (TIMS). They differ amongst each other in terms of applied electric field and state of the buffer gas. DTIMS and TWIMS belong to the timeâdispersive methods, where all ions drift along the same pathway and have a different drift time. FAIMS and DMA are spaceâdispersive methods that separate ions following different drift paths, based on their mobility difference. In TIMS, the ions are first trapped in a pressurized region before being selectively released based on their mobility differences. By using DTIMS instruments, CCS values can be directly derived from the drift time while other approaches require the use of calibrants with known CCS values to calculate the CCS value from the drift time of an unknown (35,36).

	IM-MS is able to improve the metabolome coverage by enhancing the selectivity and resolution between metabolites, but one of its major impactful applications probably lies in the field of lipidomics. Indeed, lipid analysis remains exceptionally challenging because of their structural diversity and the multiple lipid isomers that can be present in a biological sample. Contrary to conventional MS/MS approaches, IM-MS enables the discrimination between lipid isomers that differ only in the position of the acyl chain or the double bond, or with a different double bond geometry. An example is shown in Figure 4 with the two lipids 5S,12S-diHETE and LTB4 , both of which arise from different pathways and have different biological activities. 5S,12SâdiHETE and LTB4 are diastereomers and geometrical isomers. Therefore, they show identical mass spectra and similar retention behaviour using conventional LC–MS/MS analysis. However, adding IM-MS (in this example DMS) enables a baseline separation of these two compounds using two different compensation voltages (37).

	Beside lipid analysis, IM-MS has also demonstrated its usefulness for the analysis of polar metabolites in various body fluids. Most of these applications were untargeted, as discussed in references 34 and 36.

	Despite its promising contribution to improve the metabolome coverage and metabolite annotation using the CCS value, IM-MS still faces important challenges linked to data interpretation. Indeed, in an LC–IM-MS workflow, the potential in-source fragments, dimers, and adducts will also be separated in the ion mobility cell. A correct regrouping and assignment of these signal features adds another layer of complexity, which is currently not completely tackled by the software available, especially in untargeted metabolomics workflows (35).

Another MS-based strategy used to improve the metabolic coverage is dataâindependent acquisition (DIA), which allows for the detection and identification of lower abundant metabolites otherwise not recorded with conventional data-dependent acquisition methods. DIA approaches are not new but they have gained more attention since the advent of SWATH-MS approaches. In DIA, precursors selection windows are defined in the first quadrupole (MS1) of a tandem mass spectrometer; all ions are then fragmented in the collision cell and collected into a composite spectrum in the third quadrupole (MS2). Several DIA techniques have been reported so far, including MS

), all ion fragmentation (AIF), MSX, and SWATH (38). In MS

 and AIF, all coeluted precursor ions in the whole selected mass range are fragmented to acquire MS2 spectra. MSE alternatively acquires the full MS1 scan with low collision energy (full MS spectrum) and MS2 scan from all precursor ions with high collision energy (MS/MS spectrum). In AIF, all precursor ions are transmitted into a higher energy collisional dissociation cell for fragmentation. Both AIF and MS

 acquisitions generate highly complex multiplexed MS2 spectra. SWATH, which stands for S

equential Window Acquisition of all THeoretical fragment ion spectra

 and was first described in 2012, has been developed to reduce this data complexity by using a narrow isolation window (39). In SWATH-based DIA techniques, implemented on quadrupole-time-of-flight (QTOF) or less frequently Q-Orbital trap instruments, all precursors ions are sequentially fragmented in a serial of quadrupole isolation windows (Q1 windows). The complete “snapshots” of all metabolite ions and their product ions in MS2 are recorded through the whole chromatogram. The full mass range can be covered in one cycle depending on the selected MS1 scan range and the width of the isolation window. The SWATH windows can be both fixed (typically 25 Da) or variable (that is, the window width is not uniform), and are selected depending on the selectivity required and the cycle time (as short as possible if combined with UHPLC). The complexity of the multiplexed MS2 spectra is therefore decreased by reducing the number of simultaneously fragmented precursor ions, which also improves the overall quantitative performance (37,38).

	SWATH-MS is now widely used in proteomics and has emerged as a powerful technique in other clinical applications because of its reproducibility, speed, compound coverage, and quantitation accuracy. The great performance observed in proteomics fields has also attracted the attention of the metabolomics community looking to expand the information gathered on the metabolome within a single run. A number of metabolomics and lipidomics applications of LC–SWATH-MS have already been reported in the literature. For example, UHPLC–SWATH-MS was used to investigate the changes in the urinary metabolome of rat models upon administration of vinpocetine. Information on both drug metabolism and endogenous metabolite expression changes were gathered, with the simultaneous detection of 28 drug metabolites as well as altered endogenous compounds (40). Using a combination of SWATH-MS and selected reaction monitoring (SRM), Zha 

developed a two-step workflow to discover potential biomarkers for colorectal cancer. In this method, SWATH-MS was first used to acquire the MS2 spectra for all metabolites in one pooled biological sample. In the second step, a large set of SRM transitions was acquired, targeting both known and unknown compounds (around 1000–2000 metabolites). This approach increased the coverage in targeted metabolomics analysis, where more than 1300 metabolite were profiled in one run in colorectal cancer tissues (41).

	Further developments of SWATH technology are still required, particularly in the data analysis pipeline. Indeed, in a DIA-based dataset, the direct connections between precursor and product ions are missing, rendering the metabolite identification very challenging. Chromatographic ion profiles can be used to reconstruct these connections, but coelution and co-fragmentation of precursor ions makes it complicated. Several software tools have been recently developed to overcome the challenges related to DIA-based data analysis (39). The open-source software MS-DIAL, for example, uses a mathematical deconvolution of fragment ions to extract the original spectra and reconstruct the link between precursor and product ion, allowing for compound identification, annotation, and quantitation. It also implements additional functions typically used in untargeted data processing, namely, peak alignment, filtering, and missing value interpolation (42).

	Overall, SWATH-MS represents a great tool to expand the metabolome coverage and obtain both qualitative and quantitative information within a single run. The complexity of the generated data remains a challenge since the reconstructed spectral quality impacts both the confidence in metabolite annotation and quantitation accuracy. The addition of IM-MS in LC–SWATH-MS workflows might help to decrease the spectral complexity by adding an additional separation of the precursor ion to help facilitate the spectral deconvolution, as well as providing CCS values to help in metabolite identification, but also increase the need for adequate data processing software tools.

	The last decade has seen a tremendous amount of technological developments in liquid-phase chromatography and MS techniques, developments initially for other applications but showing a considerable potential in metabolomics. Modern clinical metabolomics applications rely on two essential aspects, namely, high-throughput analysis and comprehensive metabolome coverage. The latter is crucial in the quest for the Holy Grail, that is, the discovery of new biomarkers that could ultimately lead to a better understanding of (patho)physiological conditions, an earlier disease diagnosis, a better prognosis evaluation, and an individualized prediction of treatment response. The more  comprehensive the metabolome coverage is, the higher the chances are of finding specific metabolites or metabolite fingerprints.

	The chromatographic and mass spectrometric innovations presented here have also largely demonstrated their relevance in expanding the metabolome coverage. Most of those techniques, however, are still in their infancy in the field of clinical metabolomics and are rarely used for large-scale studies, where reversed-phase LC–MS and gas chromatography (GC)–MS remain the gold standard techniques. Very few studies have reported the use of HILIC–MS for the analysis of hundreds of samples, while the robustness of SFC and 2D-LC needs to be further investigated, as well as the potential of these two novel techniques in largeâscale metabolomics applications. One of the main obvious reasons is the lack of practical background knowledge of non-experienced users, who struggle to get repeatable and reproducible results. In this context, leading experts in these fields and professors play a crucial role and are strongly encouraged to share their knowledge with the younger generation of scientists. Moreover, further technological improvements are needed to ensure the batchâto-batch reproducibility of SFC and HILIC chromatographic columns, which currently remains a clear bottleneck in metabolomics. As an example, acceptable repeatabilities can be obtained in HILIC with >1000 injections (depending on the stationary phase chemistry) of pretreated biological samples using standard procedures (including adequate column reâequilibration time). However, it is much more difficult to reach such repeatability when using HILIC columns from different batches, rendering the use of HILIC in large-scale studies much more challenging than reversed-phase LC.

	Increasing the metabolome coverage does not stop at discriminating metabolites with close physicochemical properties. An important aspect often overlooked in clinical metabolomics is the distinction of optical isomers (stereoisomers such as enantiomers). An excellent example is 2-hydroxyglutarate (2-HG), the first oncometabolite (cancerâcausing metabolite) ever reported. Both D- and L- stereoisomers of hydroxyglutaric acid are normal endogenous metabolites found in human body fluids. D-2âHG-not L-2-HG-is produced in the presence of gainâof-function mutations of isocitrate dehydrogenase, causing a cascading effect in the cell that leads to genetic perturbations and malignant transformation. Typical routine analytical techniques only measure 2-HG, which strictly speaking corresponds to the sum of both D- and L-forms. Since the endogenous serum levels of L-2-HG have shown to be equal or even exceed the levels of D-2-HG in healthy individuals, it is essential to use stateâof-the-art analytical techniques to discriminate between the two stereoisomers (43). Some of the advanced techniques discussed here, mostly SFC, are applicable to chiral analysis and are therefore expected to play a crucial role in next-generation metabolomics.

	Overall, despite the technological improvements within each of the discussed techniques, none of the stateâof-theâart analytical techniques is currently capable of exhaustively assessing the metabolome. SFC will certainly become a gold standard chromatographic technique complementary to reversed-phase LC and HILIC because of the versatility and flexibility offered (convergence chromatography) and the experimental conditions, where a large diversity of metabolites can be analyzed without strong variations of the operating parameters. Moreover, the higher throughput obtained with UHPSFC is also a clear advantage in clinical metabolomics, notably with the next generation of instruments, allowing for higher back pressure to be generated, which is still a limitation in the instruments currently on the market.

	The future of metabolomics probably relies on the combination of different separation dimensions in an on-line format, as demonstrated with 2D-LC approaches. The first 2D-LC-SFC application has been reported in the literature for simultaneous achiral-chiral analysis of pharmaceutical compounds (44), a multidimensional approach that might be further investigated for metabolomics-based applications. The combination of multidimensional LC with IM-MS has a promising future in metabolomics, showing the remarkable advantage of improving the metabolome coverage while keeping similar throughput. Alternative approaches based on miniaturization of conventional LC techniques and the use of micro-pillar array columns instead of columns packed with porous particles will also probably help to further expand the metabolome coverage, as already shown in lipidomics where structural lipid isomers were chromatographically baseline resolved using micro-pillar array columns (45).

	One should also keep in mind the challenges associated with a substantial improvement of the metabolome coverage. First, the development of cutting-edge analytical instruments should not forget the importance of sample preparation, which should be as simple and generic as possible while providing sufficient clean-up to lower the occurrence of matrix effects. Moreover, an increased number of metabolites in quantitative targeted metabolomics means an increased number of internal standards, which raises the overall costs. Finally, enhancing the number of metabolite features measured in a studied population requests a much higher number of samples and subjects included in the study design to keep a sufficient statistical power, which in turn substantially increases the costs and the number of samples analyzed. A compromise between all aspects is definitely needed to achieve successful results in the field of clinical metabolomics.

		I. Kohler, A. Verhoeven, R.J. Derks, and M. Giera, 

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		P. Rainville, G. Theodoridis, R.S. Plumb, and I.D. Wilson, 

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J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 

		J.L. Spalding, F.J. Naser, N.G. Mahieu, S.L. Johnson, and G.J. Patti, 

		D.Q. Tang, L. Zou, X.X. Yin, and C.N. Ong, 

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J. Chromatogr. B Analyt. Technol. Biomed. Life Sci.

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		A. Grand-Guillaume Perrenoud, J.L. Veuthey, and D. Guillarme, 

		M. Lisa, E. Cifkova, M. Khalikova, M. Ovcacikova, and M. Holcapek, 

		C. West, E. Lemasson, S. Bertin, P. Hennig, and E. Lesellier, 

		B.W.J. Pirok, A.F.G. Gargano, and P.J. Schoenmakers, 

		B.W.J. Pirok, D.R. Stoll, and P. Schoenmakers, 

		K. Ortmayr, S. Hann, and G. Koellensperger, 

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 (2019) https://doi.org/10.1016/j.trac.2018.10.014.	

		H. Zha, Y. Cai, Y. Yin, Z. Wang, K. Li, and Z.J. Zhu, 

Cover Story

This review article discusses the novel separation and detection strategies that are considered promising in clinical metabolomics to enhance the metabolome coverage. It includes hydrophilic interaction chromatography (HILIC), supercritical fluid chromatography (SFC), multidimensional LC approaches, as well as ion-mobility mass spectrometry (IM-MS) and data-independent acquisition (DIA) analysis methods.

Clinical Metabolomics: Expanding the Metabolome Coverage Using Advanced Analytical Techniques

The sample solvent can have a big impact on peak shape in both reversed-phase and hydrophilic interaction liquid chromatography (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 instalment of “LC Troubleshooting”, I wrote about one of the hot topics that was discussed by several speakers at the HPLC meeting in Milan, Italy, in June-coupling of ion-exchange separations to mass spectrometric (MS) 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 reversedâphase or hydrophilic interaction liquid chromatography, (HILIC) 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 instalment 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 instalment, 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 1[a]) 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 1[b]). Similar effects can be observed in HILIC separations as well. Figure 1(c) 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 1(d).

	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:

		On-line dilution of the sample with weak solvent in the autosampler needle; this is sometimes referred to as “sandwich injection” (5).

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

		On-line dilution of the sample with weak solvent during injection into the LC column-this is sometimes referred to as at-column dilution, and requires an auxiliary pump to deliver the diluent (7).

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

	While these approaches have been used with conventional oneâdimensional LC (1D-LC), this issue is also very important in twoâdimensional (2D)â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 instalment 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 with similar physicochemical 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 2[a]) the observed peak area is nominally the same whether the sample is prepared in a matrix of 100% water, or 90:10 acetonitrile–water. However, for thiamine and cyanocobalamin (Figures 2[b] and 2[c], 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.

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 (www.chemicalize.com). 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 waterârich samples, because of the low water solubilities of these molecules. Figure 3(a) shows the absolute peak areas measured for the three compounds at each solvent composition; Figure 3(b) 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.

	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.

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

 is the editor of “LC Troubleshooting”. Stoll is a professor and co-chair of chemistry at Gustavus Adolphus College in St. Peter, Minnesota, USA. His primary research focus is on the development of 2Dâ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

Mind the Diluent: 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 gas chromatography (GC) laboratory, their function, and how to use them effectively.

	Every profession has its specialized tools. Those used in chromatography are often 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 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 chromatographyâspecific items; just look at their catalogues or onâ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 liquidâ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 lowâ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 gas chromatography (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 have also 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 1/4â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 frontâ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 vapour 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 bargainâ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 GC systems 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 millilitres 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 and 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 rightâ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 batteryâ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 are often 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 fibre 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 nonvolatile 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 jeweller’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 heatâ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 a readout to 0.01 s. 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 smartphone’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 fibres but the wood may also release higher molecular weight compounds into cleaning solvents.

GC Connections

Essential GC Accessories

The 9th International Symposium on Recent Advances in Food Analysis (RAFA 2019)

	The RAFA 2019 symposium will provide an overview of contemporary trends in analytical and bioanalytical strategies in food quality and safety control and will discuss challenges and novel approaches in food and natural products analysis. The following areas related to recent and emerging issues will be addressed within the RAFA 2019 sessions: mycotoxins, marine, and plant toxins; migrants from food contact materials; processing contaminants; pesticide and veterinary drug residues; industrial contaminants; allergens, antinutrients; metals, metalloids, and speciation; healthy nutrients and vitamins; flavourâsignificant compounds; quality assurance/quality control (QA/QC), chemometrics, and big data handling; authenticity and food fraud; omics in food analysis; food forensics; microplastics in food; novel food bioactives and supplements; organic crops and foodstuffs; human biomonitoring; portable on-site food analysis.

	The conference programme will be accompanied by several satellite events alongside workshops, seminars, and tutorials on novel analytical strategies, including workshops on “Vibrational Spectroscopy and Chemometrics for the Monitoring of Food and Feed Products and Contaminants Detection”, “Human Biomonitoring in Food Quality and Safety”, “Microplastics in the Food Chain”, and “Portable Food Analysis and Citizen Science”. Other sessions include “Data Quality and Smart Data Handling in Food Analysis” and “METROFOODâRI: Workshop on Metrology in Food and Nutrition”, among many others. 

	An open day from the EU-China-Safe, Delivering an Effective, Resilient, and Sustainable EU-China Food Safety Partnership, will focus on the latest developments and strategies in food safety and authenticity control and demonstrate the best practices for a joint EU-China food safety control system. Showcasing the FoodSmartphone and PhasmaFood projects, RAFA Smart Lab will demonstrate smart analyzers and applications for on-site testing of food quality and safety. Another session will discuss the collaboration challenges within the EU framework programme for research and innovation.

	Vendor seminars provide the opportunity for leading companies to introduce recent instrumentation and analytical strategies for advanced food quality and safety control. An exhibition of modern instruments, laboratory equipment, reference materials, and consumables used in food analysis will be an important part of the symposium.

	A platform for young scientists will be offered to present their scientific work, including Student Travel Grants and the prestigious RAFA Poster Award and sponsored poster award(s).

	In summary, RAFA 2019 will offer a high-quality scientific programme with top-quality presentations followed by stimulating discussions, a series of satellite events, a large state-of-the-art exhibition, and an attractive social programme. Scientific contributions will be presented by leading scientists through keynote lectures and by contributed oral and poster presentations. The RAFA 2019 programme will be tailored to provide a lot of opportunities for networking as well as exploration of the latest results of the food analysis community.
	
	

Events

The 9th International Symposium on Recent Advances in Food Analysis (RAFA 2019) will take place in Prague, Czech Republic, on 5–8 November 2019.

LCGC Europe-09-01-2019

Vol 32 No 9 LCGC Europe September 2019 Regular Issue PDF

Clinical Metabolomics: Expanding the Metabolome Coverage Using Advanced Analytical Techniques

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

Essential GC Accessories

The 9th International Symposium on Recent Advances in Food Analysis (RAFA 2019)