A liquid chromatography tandem mass spectrometry (LC–MS–MS) “dilute and shoot” method for the determination of 331 (toxic) secondary metabolites of fungi and bacteria has recently been optimized and validated for different food matrices (1) according to the guidelines established in the Directorate General for Health and Consumer Affairs of the European Commission (SANCO) document No. 12495/2011. This article will provide useful tips for smooth validation of multi-analyte LC–MS–MS methods and summarizes important validation outcomes for 295 analytes, including over 200 mycotoxins. The second part of this article focuses on the performance of the method in proficiency testing with an emphasis on difficult matrices. Z-scores obtained with this method were between -2 and 2 in 368 out of 408 cases for maize, wheat, triticale, bran, nuts, baby food, raisins, figs, coffee, liquorice, and hot pepper. From these results it can be concluded that quantitative determination of mycotoxins by LC–MS–MS based on a “dilute and shoot” approach is also feasible in the case of complex matrices such as such as pepper, coffee, or liquorice. 

	Spoilage of food and feed is still a global problem, leading to enormous annual losses of several hundred million tonnes. In relation to that, microscopic filamentous fungi are considered as one of the most feared agents because of mycotoxin production (2). Mycotoxins are low-molecular-weight, secondary metabolites of mold, and are toxic to animals and humans even in low concentrations (3). The European Union (and many other countries) has therefore established maximum levels for the most important mycotoxins in food (4). Such regulations can be laid down or amended only based on long-term occurrence monitoring and availability of toxicological data followed by exposure risk assessment. The European Food Safety Authority (EFSA) organizes a continuous call for data collection about the occurrence of already regulated (aflatoxins, ochratoxin A, deoxynivalenol, HT-2 toxin, and T-2 toxin) and so-far unregulated mycotoxins (nivalenol and ergot alkaloids) (5). Further collection of accurate data is also needed for enniatins, beauvericin, sterigmatocystin, 

 toxins, citrinin, and phomopsins for human and animal exposure assessment (6). 

	The increasing requirements for accurate and reliable occurrence data for more than one toxin has led to great progress in the development of highly selective, sensitive, and accurate LC–MS–MS methods for multi-mycotoxin determination in food and feed (7). A range of methods has been published on the identification and accurate quantification of single or chemically related mycotoxins in several matrices (8). However, the number of validated multi-analyte methods covering mycotoxins of different classes is limited for several reasons, beginning with sample preparation. The extraction solvent used has to solve a broad range of chemically unrelated toxins, and sophisticated cleanups have to be omitted to avoid discrimination of some compounds. Unspecific extraction and the reduction of sample cleanup to a minimum can result in suppression or enhancement of analyte responses during the ionization process. So-called matrix effects are a major challenge in successful development of reliable, quantitative multi-analyte methods. Therefore, considerable efforts to control matrix effects should be performed to obtain accurate results. However, common approaches for counteracting matrix effects (unspecific QuEChERS clean-up, standard addition, matrix-matched standard addition, SIDA [stable isotope dilution assay]) are not feasible for LC–MS–MS methods covering more than 200 mycotoxins. In this case, injection of diluted extracts on a highly sensitive LC–MS–MS instrument has been shown to be the most effective approach (1). 

	Performing in-house validation is necessary for reliable quantification at a high level of trueness, preferably according to international guidelines. However, validation of each matrix that should be analyzed for around 300 substances is costly and time-consuming. The SANCO document for development of multi-analyte methods in pesticides residue analysis recommends that at least one representative commodity from each commodity group should be validated and evidence for fitness of purpose should be provided (9). This approach, which was successfully applied in pesticide residues analysis, has also been followed for the validation of the developed multi-mycotoxin LC–MS–MS method (1). 

	Another difficulty in quality assurance of multiâmycotoxin LC–MS–MS data is the limited or even non-availability of certified reference materials containing multiple structurally unrelated mycotoxins. Therefore, participation in proficiency testing is the only option to prove the long-term performance of the method.

	The objective of this article is: (i) to provide useful tips for smooth validation of multi-analyte LC–MS–MS methods; (ii) to summarize the important validation outcomes; and (iii) to present the long-term performance of this “dilute and shoot” method in the proficiency testing with a special focus on “difficult” matrices. 

Methanol, acetonitrile, ammonium acetate (all LC–MS grade quality), and glacial acetic acid (p.a.) were purchased from Sigma Aldrich. A Purelab Ultra system (ELGA LabWater) was used for further purification of reverse osmosis water.

	Standards of fungal and bacterial metabolites were obtained either as gifts from various research groups or from the following commercial sources: Romer Labs Inc., Sigma-Aldrich, Iris Biotech GmbH, Axxora Europe, and LGC Promochem GmbH.

Method Validation According to the SANCO Document:

 Detailed information about method validation is provided in (1). Briefly, four different model matrices were chosen for spiking experiments - apple puree (high water-containing matrix), hazelnuts (high fat content), maize (high starch content), and green pepper (complex or “difficult” matrix) (9). Blank ground samples of 0.50 g were spiked with solutions containing all analytes and left overnight in darkness. Samples were then extracted with 2 mL of acetonitrile/water/acetic acid 79:20:1, v/v/v for 90 min on a rotary shaker (GFL) and subsequently centrifuged for 2 min at 3000 rpm. The extracts were transferred into glass vials using Pasteur pipettes, and 350 μL aliquots were diluted with the same volume of acetonitrile/water/acetic acid 20:79:1, v/v/v. Finally, 5 μL of the diluted extract was injected into a LC–MS–MS system without further pre-treatment. Spiking experiments were performed at four levels in five repetitions of each, resulting in relative concentrations of 1:3:10:30 of the final diluted extracts. External and matrixâmatched calibrations were prepared by dilution of the final working solution with acetonitrile/water/acetic acid (49.5/49.5/1, v/v/v) and diluted blank extracts of each model matrices, respectively, at the levels corresponding to spiked samples. In addition, one calibration point below and one above the spiking concentration range were prepared in anticipation of the expected matrix induced signal suppression and enhancement of target analytes.

	The peaks were integrated and linear, 1/x weighted, calibration curves were constructed from the data obtained from the analysis of each sample type (spiked sample, neat solvent standard, spiked extract) using MultiQuant 2.0.2 software (Sciex) to evaluate the linearity of the method. Further data evaluation was performed in Microsoft Excel 2007. All the other performance characteristics of the method (recovery, apparent recovery, repeatability, and matrix effects) were evaluated at each spiking level for all model matrices (1).

Detection and quantification was performed with a QTrap 5500 MS/MS system (Sciex) equipped with a TurboV electrospray ionization (ESI) source and a 1290 series UHPLC system (Agilent Technologies). Chromatographic separation was performed at 25 °C on a 150 × 4.6 mm, 5-μm Gemini C18-column equipped with a C18 security guard cartridge, 4 × 3 mm i.d. (Phenomenex). Elution was carried out in binary gradient mode. Both mobile phases contained 5 mM ammonium acetate and were composed of methanol/water/acetic acid 10:89:1 (v/v/v; eluent A) and 97:2:1 (v/v/v; eluent B), respectively. After an initial time of 2 min at 100% A, the proportion of B was increased linearly to 50% within 3 min. Further linear increase of B to 100% within 9 min was followed by a hold-time of 4 min at 100% B and 2.5 min column re-equilibration at 100% A. The flow rate was 1000 μL/min. ESI–MS–MS was performed in the scheduled multiple reaction monitoring (sMRM) mode both in positive and negative polarities in two separate chromatographic runs. The sMRM detection window of each analyte was set to the respective retention time ±27 s and ±42 s in positive and in negative mode, respectively. The target scan time was set to 1 s. The settings of the ESI source were as follows: source temperature: 550 °C; curtain gas: 30 psi; ion source gas 1 (sheath gas): 80 psi; ion source gas 2 (drying gas): 80 psi; ion-spray voltage: −4500 V and +5500 V, respectively; collision gas: (nitrogen) medium. Confirmatory identification was obtained through the acquisition of two sMRMs per analyte (with the exception of moniliformin and 3-nitropropionic acid that each â¨exhibit only one fragment ion) (1), which yields 4.0 identification points according to commission decision 2002/657/EC (10).

 The trueness of the method is shown by participation in proficiency tests organized by Bipea (Gennevilliers, France) and by the Institute of Science of Food Production of the National Research Council of Italy (ISPA-CNR, Bari, Italy) (15). 

Useful Tips for the Experimental Set-up of the Validation Process and Validation Data Evaluation: 

As no guidelines and recommendations suitable for validation of multiâmycotoxin methods (or multi-analyte methods in general) have been laid down so far, the recommendations provided by SANCO for pesticides have been followed (9). However, slight amendments were necessary to improve validation outcomes, minimize working time for sample preparation, and lower validation costs.

 SANCO suggests validation at two concentration levels using five repetitions, one level close to limit of quantification (LOQ) and one higher. As the final working mixture of analytical standards contained 331 analytes at various concentrations, we decided to validate four levels, each at five repetitions. Such experimental setup amendment prevented a potential non-linearity of the method for some analytes and discrimination of analytes with high LOQs. 

The whole sample preparation procedure can be miniaturized for validation purposes to decrease the amount of standards needed for spiking. For example, we used 0.5 g of blank sample material plus 2 mL of extraction solvent for validation, but in routine analysis and proficiency testing, 5 g of sample was extracted with 20 mL of extraction solvent instead. 

 High sugar content matrices (for example, raisins) cannot be miniaturized to 0.5 g because of inhomogeneity (1).

This was evaluated based on apparent recoveries instead of extraction recovery because we use neat-solvent calibration and spiked samples in daily routine analysis rather than matrix-matched standards. The reason for this is that it is very difficult to find a blank matrix for all analytes involved in the method. Apparent recovery expresses both extraction efficiency and matrix effects, and therefore it contains more relevant and “real-life” information (1). In routine analysis, data are also corrected for apparent recovery.

As SANCO provides no recommendations about the acceptable range for matrix effects, the range of 90–110% was considered to have no effect. Matrix effect evaluation should be performed using two approaches: “traditional slope” and “one calibration point”. “Traditional slope” evaluation is commonly used during validation. The slope obtained from matrixâmatched standards is compared to the one obtained from neat solvent standards. However, evaluation of matrix effects by comparison of each calibration point instead of the slope better reflects the real extent of matrix effect across the whole concentration range. Therefore, we recommend using both approaches during validation (1).

Evaluation of LOQ according to SANCO is highly recommended for multi-analyte methods. LOQ is estimated as the lowest spiking level (LL) that allows reliable detection of all five replicates meeting the performance criteria of RSD < 20% and mean recovery of 70–120% at both SRM transitions. LOQs obtained by the LL approach did not differ significantly from LOQs calculated from signal-to-noise in our experience (1).

 Following the guidelines laid down by SANCO for pesticide analysis, convenient validation data of our method were obtained for 295 out of 331 analytes. The remainder of the compounds could not be reliably validated for several reasons: (i) non-availability of analytical standards; (ii) instability of an analyte in the final working solution; and (iii) low concentration of analytical standard for spiking. Validation for some analytes was done only semi-quantitatively; a fungal extract was used for spiking instead of analytical standards that were not commercially available (1). 

	The distribution of apparent recoveries and matrix effects throughout the set of 295 analytes in apple puree, hazelnuts, maize, and green pepper at the lowest and the highest spiking levels is displayed in Figure 1 and 2, respectively. The highest validation level corresponds to a 1:10 dilution of the final working solution of the analytical standards. As the lowest validated level, the lowest spiking level reliably detectable at all five repetitions (RSD < 20) on both SRM transitions was considered. As expected, green pepper was the most difficult matrix. Only 20% of analytes met acceptable apparent recoveries of 70–120%, as recommended by SANCO (9). Moreover, 25% of the compounds were not detected in green pepper at any level lower than the highest validated level. With regards to the other matrices, more than 50% of compounds were in the acceptable range of 70–120%. For the analytes out of this range, either high matrix suppression or enhancement or â¨low extraction efficiency was observed. More detailed information is provided in (1).

	Any influence of matrix effects was observed for all matrices, albeit to a varying extent, which was strongly dependent on the analyte/matrix combination (see Figure 2). Both matrix suppression and enhancement were observed. In general, the lowest matrix effects were obtained in apple puree and the highest in green pepper. Only 10% of analytes were not affected by matrix effects in green pepper. 

	Method precision was proven by repeatability of five repetitions at the highest and lowest level. Less than 5% of compounds were above the RSD of 20% in apple puree, maize, and hazelnuts. In green pepper, the RSD was not below 20% for 11% of all validated analytes. 

To check for the trueness of the results obtained by our “dilute and shoot” LC–MS–MS approach, we have been participating in a number of laboratory proficiency testing (PT) schemes organized by various institutions (Bipea, CODAâCERVA, FAPAS) since 2010. Regular PT participation enables method drawbacks to be highlighted, helps to detect if something is out of control (for example, spoilage of a standard), and assists in the improvement of the overall method performance. Furthermore, analysis of uncommon and difficult matrices such as raisins or spices adds information about method accuracy in complex or even non-validated matrices. Generally, the spectrum of analytical methods used by participating laboratories varies from ELISA through conventional HPLC coupled to fluorescence detection (FLD) to advanced LC–MS–MS or LC–high resolution mass spectrometry (HRMS)-based methods. Therefore, each successful PT participation has proved that the developed multi-mycotoxin method can reliably compete with other methods in terms of accuracy. A few examples of PTs are discussed below. 

	A total of 408 z-scores have been calculated out of the reported data within our continuous PT participation, and 368 values were in the acceptable range of -2 < z < 2. Only 23 z-scores were considered as questionable (between -3 and 3) and 17 were unacceptable (z < -3, z > 3). Figure 3 summarizes the z-scores of our multi-mycotoxin LC–MS–MS method in Bipea PT (11), where multiple mycotoxins were determined in maize, grains, nuts, baby food, and “difficult” matrices. The distribution shows that the majority of outliers (z-scores) were calculated for baby food, which exhibited concentrations very near the LOQ, and difficult matrices.

	The performance of the method in some “difficult” matrices is presented in Table 1. Although, matrices such as raisins, liquorice, or figs have never been fully validated, most of the obtained results were within the acceptable z-score range (-2 < z < 2). However, the z-scores for aflatoxins in hot pepper (02-1031) were out of the acceptable range. The reported results were evaluated on the external neat-solvent calibration curve and corrected on the apparent recovery obtained from spiking of the validated results of green pepper (12). Another very complex matrix was raisins. The sample of raisins with identification number of 03-3131 was not successfully analyzed for aflatoxins. In this case the results reported by laboratories were dispersed widely. During the homogeneity testing and checks between bags, some differences for aflatoxins G1 were found (13). Obviously, most of the participants did not homogenize the whole obtained batch. The assigned value was not calculated from the data of participating laboratories, but the reference laboratory measurements instead. It should be noted that reported results for aflatoxins in raisins (04-3131) analyzed later were in agreement with other laboratories, yielding acceptable z-scores (14). 

	In 2014, ISPA-CNR organized a proficiency test on mycotoxins (15). It was aimed at laboratories willing to test the trueness of their multiâmycotoxin methods. Therefore, mainly LC–MS–MS methods were used for the quantitative determination of deoxynivalenol, zearalenone, HT-2 and T-2 toxins, fumonisins, aflatoxins and ochratoxin A in maize and deoxynivalenol, zearalenone, HT-2 and T-2 toxins, and ochratoxin A in wheat within this proficiency testing scheme. The performance of our method is summarized in Table 2. The developed LC–MS–MS multiâmycotoxin method was successful, yielding z-scores within the range of -2 < z < 2 (15). Concerning aflatoxin B2 and aflatoxin G2 in maize, no statistical evaluation was reported because of a lack of sufficient data. 

	This article presents information on the background and the personal experiences which have been gained during the validation of a “dilute and shoot” multi-analyte LC–MS–MS method for the determination of 331 (toxic) secondary metabolites of fungi and bacteria in food. In addition, guidance on proper method validation of a multi-analyte LC–MS–MS method is provided. 

	The performance of our “dilute and shoot” method in proficiency tests has been satisfactory. The routine participation in proficiency testing has proved that the developed and validated LC–MS–MS method is able to provide accurate results for regulated mycotoxins in the case of “difficult” matrices such as raisins, figs, or liquorice.

	(1) A. Malachová, M. Sulyok, E. Beltrán, F. Berthiller, and R. Krska, 

	(2) http://www.mycotoxins.org/node/56 (accessed on 15 July 2015).

	(4) Commission regulation (EC) No. 1881/2006 http://eur-lex.europa.eu/LexUriServ/LexUriServ.do?uri=OJ:L:2006:364:0005:0024:EN:PDF (accessed on 15 July 2015).

	(5) EFSA. http://www.efsa.europa.eu/en/search.htm?search=call&text=call (accessed on 15 July 2015).

	(6) CEN mandates, Call for tender for project leaders for the development of standardized methods for the analysis of mycotoxins in food (available online since 10 May 2013) file:///C:/Users/ataml/Downloads/Call_520_CENwebsite_2013_05_10.pdf

	(8) P. Zöllner and B. Mayer-Helm, â¨

	(9) Document SANCO/12495/2011. http://ec.europa.eu/food/plant/plant protection products/guidance documents/docs/qualcontrol en.pdf (accessed on 10 December 2013).

	(10) 2002/657/EC. http://eur-lex.europa.eu/LexUriServ/LexUriServ.do?uri=OJ:L:2002:221:0008:0036:EN:PDF (accessed on 15 July 2015).

	(11) Bipea, Provider of Proficiency Testing Programs. http://www.bipea.org/â¨content/pt-programs (accessed on â¨15 July 2015)

	(12) Bipea, Interlaboratory comparisons report on hot pepper (02-1031), 31a-133-Mycotoxins-Aflatoxins and Ochratoxins, 12 April 2013.

	(13) Bipea, Interlaboratory comparisons report on raisins (03-3131), 31a-134-Mycotoxins-Aflatoxins and Ochratoxins, 24 July 2013.

	(14) Bipea, Interlaboratory comparisons report on raisins (04-3131), 31a-140-Mycotoxins-Dried Fruits, Spices and Other Products, 16 October 2014.

	(15) A. De Girolamo, B. Ciasca, J. Stroka, S. Bratinova, A. Visconti, and V.M.T. Lattanzio, Report of the 2014 Proficiency Test for LC-MS(MS) multi-mycotoxin methods, June 2015. http://www.ispacnr.it/?p=479

 studied analytical chemistry at the Department of Food Analysis and Nutrition, Faculty of Food and Biochemical Technology of the University of Chemistry and Technology (UCT Prague), Czech Republic. She received her PhD degree in 2012. She currently works as a postdoc in the Christian Doppler Laboratory for Mycotoxin Metabolism and the Center for Analytical Chemistry at the Department of Agrobiotechnology (IFA-Tulln) of the University of Natural Resources and Life Sciences, Vienna (BOKU). She is an expert in LC–MS, including high resolution MS. Her main field of expertise is in development, optimization, and validation of LC–MS-based methods for multiple analytes or methods for analysis of complex matrices. 

 is Head of the Laboratory for Multi-Toxin Screening at the Department of Agrobiotechnology (IFA-Tulln) at BOKU. His publication from 2006 on a validated LC–MS–MS based method for the determination of 39 mycotoxins in wheat and maize is the most often cited original paper in the field of mycotoxins from the past 10 years. This method has been continuously expanded to become the world’s most comprehensive approach for the quantitative determination of mycotoxins and fungal metabolites in food, feed, and damp indoor environments. The analytical quality of the method is continuously monitored by participation in proficiency tests and multi-toxin inter-laboratory comparison studies with the aim to prove that the accuracy of the LC–MS–MS-based “dilute and shoot” approach may compete with conventional methods. Dr. Sulyok has (co-)authored more than 100 SCI publications (h-factor 27).

obtained an International PhD degree at University Jaume I (Castellon, Spain) in April 2014. His research was based on the development of analytical methods for the determination of natural toxins in food and environmental samples by liquid chromatography coupled to tandem mass spectrometry with triple quadrupole analyzers. He has published around 15 peer-reviewed articles in international journals and 20 communications in workshops and congresses. He has performed pre-doctoral and post-doctoral research at the Center for Analytical Chemistry-IFA (Tulln, Austria). Current research is mainly focused on the application of LC–MS for the development of analytical methods for quantification and confirmation of different compounds (toxins, hormones, pesticides) in different matrices at very low levels.

is an associate professor at the University of Natural Resources and Life Sciences, Vienna (BOKU) and Head of the Christian Doppler Laboratory for Mycotoxin Metabolism. In 2006 he received the Brigitte-Gedek-Award from the German Society of Mycotoxin Research for his PhD thesis on masked mycotoxins. He continued his research in the laboratory of Rudi Krska at the IFA-Tulln, as well as some months abroad at the Danish Technical University and the Food Research Division of Health Canada in Ottawa. As an expert in the areas of (modified) mycotoxins and LC–MS, his scientific output includes about 100 SCI publications.

Articles

Cover Story

Multi-Toxin Determination in Food – The Power of "Dilute and Shoot" Approaches in LC–MS–MS

Are your solvent costs too high? Here are some ideas on how to decrease the amount of mobile phase you use.

	 	In today’s laboratories, expense reduction has become a standard part of doing business. In some cases, the amount of money we spend on solvents for the mobile phase in liquid chromatography (LC) operations can be a significant part of the cost of analysis. In other cases, it can be less of a concern. For this month’s “LC Troubleshooting” discussion, I’d like to look at some of the options that may be appropriate ways to reduce the cost of mobileâphase solvents in your laboratory. There are three major ways to approach the problem: we can recycle all or part of the mobile phase and use it again; we can recover solvents through distillation of the waste mobile phase; and we can reduce the amount of mobile phase that we send through the column. I’ll consider each of these techniques below. Sometimes a combination of techniques may be used for further gains in cost reduction. Be aware that not all of these techniques may be available for your application. For example, direct recycling will work only for isocratic separations and not gradients.	 	

	 	In my community, we have at-curb recycling for part of our household waste. This reduces the amount of waste piling up in the landfill and recovers certain materials, such as aluminium, steel, paper, and some plastics, for reuse. We are also encouraged to reduce the amount of waste we generate by reusing some materials and using less of others. When I return beverage containers to the grocery store, my deposit on these is returned. In some cases, the whole recycling process reduces my expenses, whereas in others it is done because it is the right thing to do from an environmental stewardship standpoint.	 	When we consider the mobile phase used in LC operations, it may be possible to recycle all or some of the components, but even if it is possible, the economics of the process may not make practical sense. If your method is isocratic, the mobile-phase composition is constant, and you can directly recycle your mobile phase in many cases. This does not work for gradient methods, because the waste bottle will contain an “average” mixture of the mobile phase that is not directly useful.	 	The simplest way to recycle the mobile phase is to direct the waste line from the detector directly back into the reservoir. This seems like a rash decision and many workers are reluctant to do this, even though it can be done successfully for many methods. “But”, you ask, “aren’t you contaminating the mobile phase with sample components that are eluted from the column?” Indeed you are, but let’s consider what is happening. Under isocratic conditions, the mobile-phase composition is constant, and even if you have a retained component of the mobile phase, it will not generate a peak in the chromatogram. For example, most ion-pairing reagents are chosen because they stick strongly to the column and build up a charge on the column packing. Under the steady state conditions of isocratic flow, you don’t see a peak for the ionâpairing reagent in the chromatogram. However, if you were to inject a sample that contained that same ionâpairing reagent - for example, hexane sulphonate - into a mobile phase that did not contain that compound, you might see a peak for the reagent if the detector were set to detect it. In other words, if a compound is introduced to the column at a steady state, its detector response will be constant and no peaks will be observed. This is exactly what happens if you direct the waste stream back into the reservoir.	 	There are a few things to be aware of to minimize potential problems with direct recycling of mobile phase. First, you would like the mobile-phase composition to be constant and homogeneous. If you are continually adding sample waste components, the composition will not be truly constant, but if the dilution factor is sufficient, it will be constant from a practical standpoint. For example, if you have 1 L of mobile phase and you add 1 mL/min of waste to it while pumping 1 mL/min out of the reservoir, the volume will stay the same, but the composition will change very slowly over time. In this example, the already-diluted sample that is eluted from the column would be diluted 1000-fold more as it entered the reservoir. The background signal will gradually increase as more and more analyte is added to the reservoir, but this will generally not be noticeable over a few days’ time. To keep the solution as homogeneous as possible, place the reservoir on a stir plate and add a stir bar to keep it well mixed. Just as you need to be careful not to use a single preparation of mobile phase for too long without replacement so as to avoid evaporation, microbial growth, or other problems, you can’t use recycled mobile phase forever, either. You’ll have to determine how long you can use the mobile phase, but I’d suggest a maximum of 1–2 weeks. Generally, I prefer to make fresh buffers at least once a week even without recycling, and recycling will certainly not extend your current expiration policies.	 	Direct recycling generally will be most successful when your analytes generate a large signal-to-noise ratio. It is most likely that any problems related to recycling will appear first for trace levels of analytes, such as with an impurity method, so direct recycling may not be practical for these methods.	
			 	 	Because direct recycling gradually contaminates the mobile phase, an alternate recycling technique can be used, which I’ll call 

. If you consider a normal day’s LC operation in the laboratory, you put the mobile phase on the system and allow the column to equilibrate. Then you run a system suitability test, followed by some calibration standards, and finally run your samples for analysis, perhaps with occasional standards or check samples added. Of the total time of operation, for only a small fraction of the time is there a sample component being eluted from the column. If the fractions of the mobile phase that contain analytes and unwanted peaks, as well as the material eluted at the column void (solvent front), could be directed to waste, the remaining mobile-phase stream should be pure. If you put a two-way valve after the detector, you could watch the chromatogram, and when peaks were being eluted, you could direct the waste stream to a waste container; when no peaks were being eluted, you could direct the waste stream back to the sample reservoir. If you do this manually, of course, the labour costs will quickly eclipse any savings realized by reducing solvent consumption. What you need instead is a device that looks at the detector output and determines if a peak is present or not - this is what your data system does. By using a similar device to detect peaks and control the switching valve, you could have an automated recycler to replace the person running a manually controlled valve. Such a device is available from Spectrum Chromatography, called the S-3 HPLC Solvent Recycler. (I’m sure similar devices are available from other sources, as well.) The manufacturer claims that if you are able to recycle 25 L of high performance liquid chromatography (HPLC)-grade acetonitrile, you will pay for the device. If you use it, I’d suggest taking the same precautions I mentioned above for direct recycling: use a stir bar to keep the reservoir contents homogeneous, and don’t try to extend the expiration date of your mobile phase beyond current limits.	 	As an alternate technique, you could use an automated switching valve and control it with the timed events output from your LC system. If you diverted the timed section of the chromatogram at the solvent front and perhaps where large peaks eluted, you would get the major advantages of fractional recycling without having to buy a specialized instrument. However, the commercial recyclers don’t cost much more than an automated switching valve, so it may not be worth your time to design your own device.	 	

	 	When I started doing LC analyses in a pesticide residue laboratory in graduate school, our department distilled all our solvents to increase their purity. Today most of us simply purchase high-purity, HPLC-grade solvents. However, you could use distillation to recover the solvents from the waste mobile phase, perhaps with a spinning-band distillation column to get added purity. This would allow you to recover the organic solvent from an aqueous–organic mobile-phase waste stream that also contained buffers or other additives. Because distillation separates the organic solvent for recovery, it will work just as well for isocratic and gradient LC waste streams.	 	At least one company, B/R Instrument Corp., offers several different distillation products to allow you to recover mobile-phase solvents. These recovered solvents can then be used to prepare fresh mobile phases. If I were to use one of these devices, I would like to be confident that it produces sufficiently pure solvents for LC analysis, especially if you are performing trace analysis with gradient elution. I’m sure that the noted supplier (or others) would be happy to discuss this concern with you.	 	

	 	Perhaps the easiest way to conserve solvent is to reduce how much you use. You can make most of the changes discussed below and remain within the guidelines that the pharmacopoeias recommend. This means that all you need to do is verify that the new conditions give the same results as the original, document these results, and proceed with analysis. Your company may require additional procedures or documentation when such changes are made.	 	As a general rule, if you keep the column chemistry, column plate number, and mobile-phase composition constant, you should obtain the same separation. Special attention needs to be paid to gradient methods, which may require compensating changes in the conditions, and for cases when instrument extra-column volume plays a significant role in band width. Next let’s look at three possible approaches to reduce mobile-phase use.	 	

Perhaps the simplest way to reduce solvent consumption is to reduce the column diameter. If the flow rate is adjusted for the change in column crossâsectional area, you should get the same separation and retention times with both isocratic and gradient methods. Take, for example, a method running on the most popular column, a 150 mm × 4.6 mm column, operated at a flow rate of 2.0 mL/min. We can generally reduce the column diameter to other popular sizes of 3.0 or 2.1 mm i.d. without excessive extracolumn band broadening. For the 3.0-mm column, the reduction in crossâsectional area is (3.0/4.6)

 = 0.4. To keep the same linear velocity of the mobile phase through the column, we need to reduce the flow rate by this same factor, so (2.0 mL/min × 0.4) = 0.8 mL/min. (As usual for these discussions, I’ve rounded or truncated calculated values, so your results may vary slightly if you repeat my calculations. Also, to avoid clutter I’ve not included units in some of the examples where they should be obvious.) This should give the same retention times as the original conditions, but with approximately 60% reduction in solvent used. In a similar manner we can calculate the effect of changing to a 2.1-mm i.d. column: (2.1/4.6)

 = 0.2. Apply this to the flow rate, (2.0 mL/min × 0.2) = 0.4 mL/min, and you’ll reduce your solvent consumption by 80%. You could use even smaller diameter columns, but if you reduce the diameter below 2.1 mm, it is likely that you’ll have excessive band broadening because of extra-column effects unless you modify the LC instrument.	 	

A reduction in column packing particle diameter (

) will increase the column plate number inversely to the change in particle diameter. This, then, will allow you to use a shorter column for the same plate number. Take the example above for a 150-mm-long column packed with 5-µm 

 particles. If we change to 3-µm particles, this will increase the plate number by 5/3 = 1.67-fold. For a constant plate number, we can reduce the column length by the same factor: (150 mm/1.67) = 90 mm. Because 90-mm columns aren’t commonly available, we’d round this length to 100 mm. This tells us that the 100âmm column packed with 3-µm 

 particles would have approximately the same plate number as the 150-mm, 5-µm column. But because the length has been reduced by one-third, isocratic retention times would be reduced by one-third also, as would the method run time, so we’d save one-third of the solvent costs. In this example, the smaller particles will increase the pressure by (5/3)

 = 2.8-fold and the shorter length will reduce the pressure by 100/150 = 0.67âfold, resulting in an overall pressure increase of (2.8 × 0.67) = 1.8-fold. Many methods operate at pressures where this increase would still be within the instrument’s pressure capabilities. You could lower the flow rate to reduce the pressure; this would increase the retention times, but would not change the amount of solvent used per run. For example, reducing the flow rate 1.8-fold would mean operating at (2.0 mL/min /1.8) = 1.1 mL/min. A flow rate of 1.1 mL/min with a 100 mm column would give retention times of (0.67 × 1.8) = 1.2-fold larger than the original method, or a 20% increase in run time. An alternative would be to run at 1.5 mL/min for the same retention times and run time, but a 30% increase in pressure, which perhaps would be more acceptable. As discussed below, flow-rate changes beyond those used to generate constant linear velocity are appropriate for isocratic methods, but not for gradient methods without additional method adjustments.	 			 	 	

Column Length and Diameter Plus Particle Size: 

By combining a change in column diameter with a change in the length and particle size, we can further reduce solvent consumption. Let’s consider that the original isocratic method used a 150 mm × 4.6 mm, 5-µm 

 column operated at 2 mL/min and a run time of 10 min. This would use (2 mL/min × 10 min) = 20 mL of solvent per run. Reducing the column diameter to 2.1 mm was compensated for by reducing the flow rate to 0.4 mL/min for a solvent volume of (0.4 mL/min × 10 min) = 4 mL of solvent per run. If we combine this diameter change with the length and particle size change to a 100-mm, 3-µm column, we saw that this would reduce run times by 0.67. This would also reduce the total solvent used per run to (4 mL × 0.67) = 2.7 mL/run. We would now be using (2.7/20) ≈ 15% of the original solvent. I’ll leave you to calculate the change in pressure for this new condition.	 	The above changes in length, diameter, particle size, and flow rate will not change the separation for isocratic methods, but more care needs to be taken for gradient methods. For gradients, it is necessary to keep the number of column volumes of solvent constant for each gradient segment when the conditions are adjusted, using the following equation:	 	constant = (

 is the time for each gradient segment (in minutes), 

 is the flow rate (in millilitres per minute), and 

 is the column volume (in millilitres). A reduction in 

 is proportional to the change in column length times the cross-sectional area (or diameter squared). This means that for two conditions (with subscripts 1 = original and 2 = new), we need to have	 	(

)  [2]	 	For any given set of changes, as proposed above, we can determine the necessary gradient time adjustment by solving equation 2 for the new gradient time, 

LC Troubleshooting

Solvent costs too high? Here are some ideas on how to decrease the amount of mobile phase you use.

How to Reduce Mobile-Phase Consumption

’s e-learning tutorial on solid-phase extraction at CHROMacademy.com

	 	If you use solid-phase extraction (SPE) in your work, its proper implementation is key to the performance of your analyses. Variability in the quality of SPE protocols come from some common issues, misunderstandings, and ignorance of the mechanisms that are in play.	 	

Know Your Sorbent and Analyte Chemistry 

	 	Mixed-mode separations are the key to cleaner extracts. SPE sorbent selection follows the same rules as high performance liquid chromatography (HPLC) in terms of matching the hydrophobicity of the analyte, but more-polar sorbents, those capable of hydrogen bonding and those capable of electrostatic interaction (ion exchange) with the analyte, are increasingly more selective, and therefore result in increasingly cleaner extracts. It is always beneficial to know something of the analyte structure, such as logD (LogP) and p

 values, which are all valuable in designing an optimized extraction protocol. Polar or ionizable functional groups can be used to improve selectivity through the use of more-polar sorbents or ion-exchange sorbents, and higher logD (P) values will indicate the possibility of using a less hydrophobic stationary phase, which again can help to improve selectivity, especially when the matrix is less polar.	 	

Condition and Equilibrate to Stop Breakthrough

	 	With silica sorbents, conditioning ensures that the phase is activated or wetted and will have maximum capacity for the analyte. This conditioning is typically achieved by washing with methanol, followed by water. Equilibration matches the sorbent environment with that of the sample solution in terms of eluotropic and ionic strength. To stop analyte breakthrough and help achieve minimum elution volumes, match the equilibrating solvent as closely as possible to the eluotropic strength of the sample matrix or solvent.	 	When using ion-exchange sorbents, ensure that the counterion associated with the sorbent can be easily displaced by the analyte (hydrogen ions may typically be used in this regard), and adjust the pH of the conditioning solvent to 2 pH units above the sorbent p

 of weak cation exchanger or 2 pH units below the p

Be Brave with Wash and Elution Solvent Strength

	 	It is easy to be over cautious during the sorbent washing protocol step to avoid loss of analyte; however, this conservative approach will result in a dirtier extract than necessary. It is important to properly understand the relationship between the analyte and the stationary phase to optimize the strength of the wash solvent and remove as many interferents as possible while still retaining the analytes.	 	Wash solvent optimization is typically done by titrating (increasing in a stepwise fashion) the eluotropic or ionic strength, by adjusting the pH to suppress the ionization of the stationary phase or analyte, or increasing the strength of the counterion to disrupt analyte or sorbent interactions. One should aim to use the strongest wash solvent possible while still retaining the analyte.	 	Exactly the opposite can be said of the elution step in most SPE protocols - the temptation is to be over cautious and use a very strong elution solvent, in fear of not eluting the analyte. Again, a titration of the elution solvent will identify the weakest solvent strength (or ionic strength or pH) to elute the analyte, but leave any interferents behind on the sorbent. One caveat here is that we should balance the elution protocol solvent strength with the volume of solvent required to elute the analytes - that is, weaker solvents may require larger elution volumes, which may compromise the sensitivity of the analysis.	 	

	 	The capacity of silica-based phases is around 5–10% of the sorbent mass (per analyte), and polymers show around 20–25% capacity - so this information should be enough for you to judge the sorbent mass that you need. Also, remember that narrower longer beds tend to suffer less breakthrough but need larger elution volumes.	 	

	 	Hydrogen bonding and electrostatic mechanisms are point-to-point and analytes take longer to orient themselves with the surface moieties that cause retention, which means that we need to process the solvents at a slower speed. Typical processing speeds are around 1 mL/min; however, when using pointâtoâpoint retention mechanisms, one may need to drop the rate to as little as 100 µL/ min to ensure useful recovery.	 	

	 	Soak steps involve holding a solvent stationary within the sorbent to allow processes that are slow to equilibrate or when loading into a dried sorbent bed or to allow the solvent to penetrate the fine pore structure of silica-based sorbents. Typically used during solvent loading, elution, or when switching between immiscible solvents, soak steps lasting 1–5 min can really help to improve recovery and reproducibility.	 	

	 	Drying steps are typically used when switching between organic and aqueous solvents that are less miscible. The application of high levels of vacuum for up to 2 min after a protocol step is used to drive all remaining solvent from the sorbent matrix and helps to avoid very slow equilibration in instances where, for example, residual aqueous solvents in the fine pore structure of the sorbent exclude an organic solvent used in the next protocol step. Again, these steps typically improve recovery and reproducibility.

The Essentials

An excerpt from LCGC’s e-learning tutorial on solid-phase extraction at CHROMacademy.com

The Most Common Mistakes in Solid-Phase Extraction

In gas chromatography (GC) split injection systems, as much as 99% or more of the injected sample never enters the column: It is released downstream of the inlet where it can encounter and possibly perturb precision gas control devices. The split-vent trap acts to prevent or at least moderate such effects. This month’s instalment addresses the operation and maintenance of split-vent traps.

	 	A plethora of gas chromatography (GC) injection techniques confront chromatographers in their daily work, and the related sample injection systems can be rather complex. Split, splitless, direct, programmedâtemperature, headspace, thermal desorption, and more all serve the purpose of transferring analyte species of interest into the separation column, and each does so in ways uniquely suited to the nature of the sample and its matrix. Beyond simply moving sample molecules from outside the column to inside, injection techniques share some additional requirements. First, they should transfer sample accurately and repeatably. Of course, these two terms don’t have significant meaning until they are cast in the light of analytical needs that define accuracy and repeatability. Another desirable trait is less-frequent maintenance and tolerance of a larger number of injections without a loss of performance. Although unavoidable in some extreme cases, no one wants to disassemble an inlet every few samples.	 	The primary consideration for achieving the best possible injection performance is a close match between the sample, the injection technique, and the column. The analysis of traces of methylene chloride in polycarbonate beads, for example, is much better matched to headspace sampling than to direct or splitless injection. Headspace sampling leaves polymer residues behind in a disposable vial, while whole-sample vaporizing techniques such as direct, split, or splitless injection would rapidly contaminate the inlet and inlet liner, necessitating frequent maintenance. On the other hand, determination of polynuclear aromatic hydrocarbons (PAHs) is not well-suited to headspace sampling because of the high temperatures needed to volatilize the target compounds. Instead, the higher temperature ranges of vaporizing injection makes it a much better choice. But then vaporized sample components that migrate beyond the inlet may cause other problems.	 	

	 	Split injection helps match convenient and easy-to-dispense liquid sample sizes to the sample size and carrierâgas flow-rate requirements of open-tubular (capillary) columns. Liquid sample sizes of 0.5 µL or larger are best for handling by syringe; smaller injected sample volumes tend to compromise accuracy and repeatability because of evaporative and droplet-size effects. However, if the sample is relatively concentrated and it is all injected into the column, the total amount of solute may cause peak distortion because of overloading of the stationary phase. Split injection can reduce a too-large solute amount down to a volume the column can handle. Sample dilution is an alternative, but it may not be desirable as an extra sample preparation step or a source of possible contaminants. Split injection also reduces the amount of solvent in the column and avoids very large solvent peaks that might interfere with the separation of early-eluted solutes.	 	

	 	Figure 1 illustrates the basics of split injection with back-pressure regulated pneumatic control. A constant mass-flow controller provides carrier gas to the split inlet at flow rates from as little as 10 mL/min up to 500 mL/min or greater. This split flow enters the inlet at point A and then flows into the inlet liner. A small amount of carrier-gas flow is diverted upwards, across the septum, and out the septum purge vent B. The main carrier-gas flow passes through the inlet liner and past the tip of the column at the split point C. Some of the carrier gas flows into the column entrance, while the rest continues out of the inlet liner and exits the inlet at split exit D. The internal seal isolates the incoming and outgoing carrierâgas flows.	 	The split ratio, 

, is the ratio of the flow through the inlet liner to the flow rate of the column, 

 . Both flows must be expressed at the same temperature and pressure, normally at standard conditions of 0 °C or 20 °C and 101 kPa. The liner flow is equal to the total flow through the inlet, 

  [1]	 	The split ratio would be 100:1, for example, with a column flow rate of 1.5 mL/min, a total split flow of 152.0 mL/min, and a split-vent flow of 2.0 mL/min.	
			 	 	The split ratio refers to the ratio of the flows in equation 1 under static conditions and not to the actual fractional amount of sample that enters the column. The injection process itself is far from static. Upon entering the inlet liner, solvent and sample undergo rapid heating and expansion, which causes a pressure and flow impulse to develop and propagate along the liner in both directions. Note that this is not the “pressure pulse” of pressureâpulsed split–splitless injection, a technique that purposely increases the inlet pressure setpoint before injection and then reduces it back to normal after injection is complete. This higher initial inlet pressure can help narrow the injection pulse and control some of the effects of vaporizing injection.	 	Column flow momentarily increases as the pressure impulse encounters the column entrance, but because of the relatively high restriction of the column and low restriction of the inlet liner and other inlet internal passages, the split flow will not react proportionately and the split ratio will deviate away from its initial quiescent value. The amount of each solute that enters the column is a complex function not only of the solvent, inlet temperature, injection amount, and inlet flow rates, but also of the liner packing and configuration, solute volatility, and injection speed. For these reasons, the split ratio should be taken as an approximation, and actual solute transfer into the column should be determined by careful calibration, along with recalibration upon modification of any aspects of the injection.	 	

	 	After leaving the inlet at the split exit D, carrier gas passes through a split-vent trap, then through a back-pressure regulator, and finally is vented to the surrounding air. The split-vent trap serves two main functions. First, it acts as a buffer between the inlet and the backâpressure regulator that reduces the size of the injection pressure impulse that reaches the regulator. Second, it traps and holds lessâvolatile sample components that could otherwise contaminate the regulator and affect its operation permanently. An additional benefit is the trapping of toxic substances so they are not released into the laboratory atmosphere. An optional external split-vent trap, installed after the back-pressure regulator, at the external split-vent exit connection, can help ensure that toxics do not leave the instrument. Such external traps are useful if solvent exposure is a concern as well.	 	The injection pressure pulse reaches the back-pressure regulator quickly, while sample splitting is still occurring inside the inlet. The regulator will sense the pulse and open itself up a bit to relieve the increased pressure. As the pressure pulse subsides, the regulator will head back to its previous static position. These fluctuations add to the carrier flow perturbations caused by injection and can increase uncertainty in the injected solute amounts as well as change the retention times of early-eluted solutes.	 	By including the extra volume of the split-vent trap in the flow path, the extent of the pressure impulse is reduced, and it is spread out over a longer period. Fluctuations in the regulator are mediated and the inlet pneumatic system is made more stable. Using larger diameter tubing between the split exit D and the split-vent trap also helps mediate the pressure impulse.	 	The initial injection pressure impulse propagates rapidly from the split-vent exit, at the speed of sound through the carrier gas and tubing. Shortly afterwards, moving at the split-vent flow rate, sample components and solvent start to move out of the inlet. In the absence of the splitâvent trap, these would reach the backâpressure regulator within about 1 s after the start of injection and would further perturb its operation. Solvent vapour has a higher viscosity than carrier gas, and its presence at the back-pressure regulator would cause the regulator to open even further in compensation.	 	The split-vent trap entrains solvent and sample components that are vented from the inlet, keeping them from entering the regulator immediately. With a split ratio of 100:1, this amounts to 99% of the total sample amount. Solvent will eventually break through the trap and be released slowly into the backâpressure regulator; over time most solvents will be purged from the trap. Less-volatile solutes will tend to accumulate, so that the backâpressure regulator is protected from the solutes condensing inside.	
			 	 	

	 	Many chromatographers tend to neglect their split-vent traps, which are hidden underneath instrument panels and are easy to forget while troubleshooting or maintaining inlet systems. If normal maintenance procedures of replacing the inlet liner, internal seals, and septum do not remedy problems such as peak tailing, poor accuracy, and poor repeatability, or inability to achieve high split ratios, then the split-vent trap or connecting tubing may be 	at fault.	 	Before replacing the split-vent trap, carefully check the split-vent and septum purge flow rates, preferably with an accurate digital flowmeter, and make sure there are no leaks around the inlet seals and septum. Leak checking is best done with an electronic leak sniffer and hydrogen or helium carrier gas. Using a surfactant solution on a cooled-off inlet will also find major leaks, but may then create a major headache because the solution can be drawn into the inlet by capillary action back through a leak.	 	A blocked or restricted septum purge is a sure sign of a repeatedly overloaded inlet and the accompanying back-flash of sample up into the septum area. This problem is always accompanied by corresponding downstream contamination in the direction of the split-vent trap. While most common with splitless injection, low split ratios can yield the same effects. It may be necessary to replace the inlet nut assembly in this case, and if so be sure to replace the split exit tubing and split-vent trap as well. In extreme cases it may be necessary to remove the inlet entirely and solvent clean it.	 	If the septum purge flow is normal, meaning about 2–5 mL/min, then the problem may be a routine buildâup of contamination in the split-vent trap. The trap may last for up to a year if injections are primarily clean volatile samples with little to no high-boiling residues. Samples that contain high levels of nonvolatiles, on the other hand, could necessitate trap replacement as often as monthly.	 	With a replacement trap or trap cartridge on hand, proceed to remove the existing split-vent trap by following the instrument manufacturer’s procedures. With the trap removed, inspect the incoming tubing, the connections, and the trap itself for any visible residue. If found, such residue indicates the connecting tubing back to the inlet also needs replacement. This may be best performed with a service visit by an authorized technician. Consider modifying sample preparation in such cases so that as little high-boiling material as possible is injected.	 	Be sure to leak-check the new connections and verify the correct split-vent flow rate, then try again to achieve good chromatographic accuracy and repeatability.	 	Other injection systems also have vent lines that conduct sample components to the outside. Many headspace samplers, for example, vent a large portion of the sample. Thermal desorption systems usually have at least one split point as well. I remember one site visit where several headspace samplers were in use. Every one of them had a blocked vent exit tube, as evidenced by a complete lack of flow when the vent valve was activated manually. The sample was not particularly dirty, but the material under analysis, which needed an elevated thermostating temperature for good headspace results, had been condensing in the final short, unheated, section of vent line tubing for an extended period.	 	

	 	Split injection vent traps are a necessary adjunct to high-performance gas chromatography with openâtubular columns. These traps protect carrierâgas regulation components in the short term from performanceâdegrading perturbations because of injection, and in the long term from accumulation of sample residues that can also affect performance. Split-vent traps also help prevent nonvolatile toxics from entering the laboratory atmosphere.	 	Often, however, split-vent traps are not serviced and maintained as regularly as they should be. Residues can build up rapidly when analyzing many samples with heavy components, and contamination can sometimes extend back into split-vent connecting tubing. Careful attention to all of the inlet components, including splitâvent and septum-purge elements, can significantly improve results and reliability.	 	“GC Connections” editor 

 is a Senior Scientist at Serveron Corporation in Beaverton, Oregon, USA, and a member of 

’s editorial advisory board. Direct correspondence about this column should be addressed to “GC Connections”, 

, Hinderton Point, Lloyd Drive, Ellesmere Port, Cheshire, CH65 9HQ, UK, or email the editor-inâchief, Alasdair Matheson, at amatheson@advanstar.com

GC Connections

In GC split injection systems as much as 99% or more of the injected sample never enters the column: It is released downstream of the inlet where it can encounter and possibly perturb precision gas control devices. The split-vent trap acts to prevent or at least moderate such effects. This month’s instalment addresses the operation and maintenance of split-vent traps.

Split-Vent Traps: GC’s Dirty Secret

If sample preparation is the most time and labour intense step in the analytical process, and uses the largest amount of solvents, it stands to reason that sample preparation may present the most significant safety risks in the analytical laboratory. While most laboratory workers receive significant safety training, we may become numb to the prospect of accidents or get into the mindset that accidents only happen to other people. Given some recent, significant safety incidents, this month we step back and take a quick refresher on safety concerns to be aware of during our sample preparation activities.

	 	When I was first approached about taking on the “Sample Prep Perspectives” column, I sought the advice of “LC Troubleshooting” columnist John Dolan. I asked, “How do you come up with fresh ideas each month?” John replied that he often gets e-mail queries from readers that provide the basis for some of his instalments. That’s finally happened to me. Earlier this summer, someone from a large chemical company wrote me about guidance concerning solvent safety and their new safety programme.	 	Safety issues are simultaneously overlooked and are seeing a resurgence of interest. Wellâpublicized laboratory accidents are leading university and research institutes to place even greater scrutiny on safety. Yet, at the same time, a trusted colleague working in a top-tier analytical department confided to me the complete lack of safety concerns at their new workplace. 	 	Since solvent use is a major contributor to safety concerns with extractions and sample preparation, this month we take a look at these considerations. Note that we will not be looking at alternative solvents and replacements - that’s a topic for another column. Here we will focus on safety aspects with traditional molecular solvents. As one would expect, there is a wealth of information on solvent safety considerations in the literature and on the internet, but finding this information in a concise format is somewhat more difficult. One important resource, though not oriented toward analytical use, is the European Solvents Industry Group (www.esig.org), including the presentation of a series of “Best Practice Guides”.	 	

			 	It is unquestioned that extractions generally use the largest amounts of solvents in analytical methods. Anastas (1) has pointed out that waste solvents from Environmental Protection Agency (EPA) laboratories tend to be ignitable, corrosive, or toxic. Furthermore, 86% of EPA laboratory wastes are Resource Conservation and Recovery Act (RCRA) F-listed, or multicode, solvents, including acetone, ethyl acetate, methyl isobutylketone, methanol, toluene, and dichloromethane. Some concern over the nature of organic solvents has been noted. For example, a study of papers in the journal 

 (2) found that between 1997 and 2012, the use of chloroform and n-hexane significantly declined, but the use of ether and dichloromethane remained somewhat constant (see Figure 1). In reporting these results, I only considered solvents of interest for analytical extractions and recognize that solvent considerations for synthetic organic processes are different from analytical extractions. The analytical community has broadly explored the use of alternative solvents like supercritical fluids and ionic liquids, but given the use of regulatory and other standard methods, the widespread use of molecular solvents still predominates. The solvents often used in analytical extractions are typically flammable and possess a diversity of toxicological concerns, including skin irritation or defatting, nervous system depression, reproductive effects, and other acute or chronic effects.	 	Regarding the flammability of organic solvents, flash points can be used to predict the risk associated with solvents (flash points greater than 27–57 °C are suggested for laboratory solvents), though data is missing with mixed solvents. Liaw and colleagues (3) determined that for binary mixtures the flash point is intermediate between that of the component solvents, though in most cases, phase equilibrium has not been achieved. The liquids themselves do not burn, but rather the solvent vapours are flammable. Meanwhile, Klein, King, and Kosior (4) investigated the accumulation and clearance of solvent vapours in the laboratory. They found that chemical vapour pressure and density, room temperature, and direction and velocity of room air currents influenced contaminant levels. While no definitive statement could be made, room air change per hour rates of 6–8 are generally recommended.	 			 	 	Keeping in mind that safety considerations with extraction solvents depend on both the property and nature of the solvents and the exposure to the solvents, important guidelines can be compiled as follows:	 	• â¨Laboratory guidelines should be documented in standard operating procedures (SOP) or similar means.	• â¨Consult the supplier’s safety data sheet for specific concerns on a given solvent. Ethers and some nonaromatic unsaturated cyclic hydrocarbons may form peroxides, which can be explosive. Do not allow these solvents to be evaporated to dryness. (Note: Petroleum ether is a distillate cut; it is not an ether.)	• â¨Restrict laboratory access to only trained staff with a reason to be there. All laboratory staff should be trained on appropriate safety considerations regarding working with solvents.	• â¨Handle solvents inside fume hoods whenever applicable. Maintain appropriate air flow within the fume hoods.	â¨• Clean up solvent spills immediately. Approved spill kits should be readily available in the laboratory.	• â¨Replace lids on solvent containers after use.	â¨• Store solvents in approved containers, using secondary containment as needed, and dispose of empty containers according to local policy. Generally, solvents should be stored below eye level.	• â¨Keep personal protective equipment clean and replace at recommended intervals and as appropriate. Glove material should be appropriate for solvent used (see Figure 2).	• â¨Ignition sources, including open flames and static discharge sources, should be well isolated from solvent use areas.	• â¨Do not use solvents to wash hands or skin.	â¨• Solvent bottles should be dated when they are opened.	• â¨Waste solvent disposal should follow your organizations guidelines and applicable regulations.	 	

	 	Please note a few more remarks about the solvent compatibility of gloves: 	 	â¨• The resistance of the materials will be dependent on the thickness of the gloves, temperature, and many other environmental factors.	• â¨The recommendations given are based on laboratory testing with pure chemicals. Check with the glove manufacturer for specific applications.	• â¨Glove manufacturers have databases with test results of their gloves against many chemicals. This information is available from your supplier; make sure that you check your specific product and application.	 	Gaber and colleagues (5) created a free software program, HPLC-EAT, that evaluates the environment, health, and safety of liquid chromatography mobile phases. The program is available for download in the supplementary material at: http://pubs.rsc.org/en/Content/ArticleLanding/2011/GC/c0gc00667j#!divAbstract. Presumably, this information could also be applicable to extraction solvents. In addition to the advice provided above, standard safety precautions not exclusive to working with solvents should be followed. These include the use of safety glasses (goggles), regular inspection of eyeâwash stations, and so on. When developing laboratory safety guides for your laboratory, consult similar guides that may be publically available. Those developed by the University of Wisconsin and Yale University have been recommended to me. Specific extraction techniques may have additional safety considerations, such as those involving the use of glassware, heat, high pressure, or sharp needles, and should also be addressed.	 	

	 	Solvents represent probably the largest source of safety concerns in sample preparation procedures. While laboratory supervisors and workers receive training in the proper use of solvents, the results of this training are not always put into practice. Guidelines for developing safe practices were presented here that can be used to assist organizations in developing, or reinforcing, a culture of laboratory safety.	 	

	(2) C.P. Ashcroft, P. Dunn, J. Hayler, and A.S. Wells, 

	(3) H.-J. Liaw, V. Gerbaud, C.-C. Chen, and C.-M. Shu, 

	(4) R.C. Klein, C. King, and A. Kosior, 

	(5) Y. Gaber, U. Tonvall, M.A. Kumar, M.A. Amin, and R. Hatti-Kaul, 

	 	“Sample Prep Perspectives” editor 

 is an Associate Research Professor at South Dakota State University, USA. His research interests include green chemistry, alternative solvents, sample preparation, high resolution chromatography, and bioprocessing in supercritical fluids. He earned his PhD in 1990 at Brigham Young University under the direction of Milton L. Lee. Direct correspondence about this column should be addressed to “Sample Prep Perspectives”, 

, Hinderton Point, Lloyd Drive, Ellesmere Port, CH65 9HQ, UK, or email the editorâin-chief, Alasdair Matheson, at amatheson@advanstar.com

Sample Preparation Perspectives

Given some recent, significant safety incidents, this month we step back and take a quick refresher on safety concerns to be aware of during our sample preparation activities.

Let’s Do It Right the First Time: The Importance of Solvent Safety Considerations

The 40th International Symposium on Capillary Chromatography (ISCC) and 13th GC×GC Symposium

40th International Symposium on Capillary Chromatography (ISCC)

. The meeting will focus on recent advances in the fields of pressure and electrodriven microcolumn separations, and comprehensive two-dimensional gas chromatography (2D GC). Particular emphasis will be placed on combining capillary chromatography with various forms of mass spectrometry (MS) - from unit-mass to highâresolution and from single to hybrid analyzers. 	 	The 

 scientific programme will be a rich one, characterized by contributions from leading scientists reporting on the latest most exciting developments; keynote lectures from promising young researchers; active poster sessions; discussion sessions; workshop seminars presenting the most recent novelties in scientific instrumentation; and a world-class GC×GC course. Researchers in all areas relevant to the subjects of the symposia are invited to submit abstracts. As is traditional for the Riva meetings, the majority of presentations will be in a poster format and the Scientific Committee will select contributions for oral presentations.	 	As always, many awards will be presented at both the ISCC and GC×GC events, recognizing excellence in both established and young scientists, in oral and poster presentations. Exhibitors and sponsors remain a fundamental part of the meeting and the organisers nvite companies to participate by reserving booth space and becoming a sponsor.	 	Last but not least, the traditional “Riva” social programme will return with one or two events each day: Cocktails, welcome reception, concert, wine and cheese evening, and of course, the disco night! Please visit the web site (

Young Scientists and Researchers Environmental and Food Analysis SIG Meeting

The Auditorium, BAT Research & Development, Southampton, UK

.	 	The aim of this one-day meeting is to highlight and focus on the research and innovation of young graduate scientists and researchers in areas relating to the applications of chromatography–mass spectrometry to environmental and food matrix analysis.	 	The meeting is open to all organizations and institutions with interests in environmental and food mass spectrometry using chromatography and mass spectrometry across the spectrum of instrumentation. Please note that although the focus is on the activities of young scientists, presentation submissions are welcome from experienced scientists working in similar areas, so that there is a blend of the established with the new. It is hoped that invited keynote speakers to the sessions will be representative of both groups. Confirmed Postgraduate and Young Scientist Speakers include: Ms Sally Gamble, Department of Security and Crime Science, University College London, UK; Dr Benjamin Savareear, University of Liege, Belgium; Aneela Arshad, Polymer Science, Manchester Metropolitan University, UK; Kate Sidwick, Keele University, UK; Russel Cole, University of Portsmouth, UK; and Glenn Castle, University of Portsmouth, UK. The organizers would appreciate your support and attendance.	 	Website: 

Departments

The 40th International Symposium on Capillary Chromatography (ISCC) and the 13th GC×GC Symposium will once again be held in Riva del Garda, Italy, from 29 May to 3 June 2016.

Issue PDF

Click the title above to open the LCGC Europe October 2015 regular issue, Vol 28, No 10, in an interactive PDF format.
